No title

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 100 Development in Drosophilu 103 VII. Cytoplasmic Incompatibility VIII. Speculative Models of Sperm Function in the Fertilized Egg 107 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 118 121 Influence of the Ionic Environment on Spermatozoa 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 Mutational Analysis of mHLH Function 182 Early Activation of the Myogenic Program 188 MEF2 Family of Transcription Factors 194 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

xii

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

1

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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 r,

< < < a < <4< Inverted repeat -5’a m (>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

314 amino acids

79

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

Poschiavinus 129

Block 3

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

32

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).

3a

D. Poccia and P.Collas

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 (Des~ ~ ~ 2 B complex from extracts of sev et al., 1989). A purified ~ 3 4 kinase-cyclin 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 2 structure and an acrosome (or a rudimentary acrosome; composed of a 9 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, calciuminduced calcium release; DIDS, 4,4’-diisothiocyanatostiIbene-2,2’disulfonic acid; E,, Nemst potassium 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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4. Ion Channels: Elements in Gamete Signaling

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Fig. 2 Schematic igram of the sea urchin sperm acrosome reaction. Egg ctor (FSP) binding to the sperm receptor ( I ) opens a Ca2+ channel (2) which inactivates in a few seconds. This channel is sensitive to dihydropyridines (DHPs), verapamil (VER), and trifluoperazine (TFP). The activation of this channel initiates an increase in [Ca2+],(A) and somehow modulates the opening of a second Caz+ channel (3) that is pH,-dependent and induces a sustained increase in the [Car+],. At the same time, or immediately after the first Ca2+ channel (2) opens, a K + channel (4) is activated. hyperpolarizing the cell (B, dashed circle) and activating a voltage-dependent Na+/H+ exchange ( 5 ) that increases pH, (C). This latter change in pH, is linked to the opening of the second Ca2+ channel (3) and to a large depolarization. The FSP-induced hyperpolarization (-A,!?,,,) and the increases in [Caz+], and pH, activate the sperm adenylyl cyclase ( 6 ) that modulates another K+ channel (7). Possibly CI-- channels (8) are involved in setting the resting membrane potential. Single channel records are from sperm membranes incorporated into planar lipid bilayers and may correspond to the ones causing [Ca*+],increases (A, D) and E M changes (B). FSP addition is indicated by a gray arrow.

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described recently (Labarca et a / . , 1996). Sea urchin sperm adenylyl cyclase does not undergo a classical regulation (Garbers, 1989). It has been described as strictly dependent on [Ca2+Ii and perhaps calmodulin (Watkins et al., 1978; Garbers and Kopf, 1980; Bookbinder et al., 1990). The authors of this chapter have found conditions where adenylyl cyclase is stimulated by membrane potential independently of [Ca2+], and pHi; therefore, it could be another piece of the puzzle (Beltrin et a l . , 1995). The high-conductance, low selectivity for Ca2+ and substate complexity of the channel from sea urchin and mouse spermatozoa recorded by the authors are properties closer to those displayed by the ryanodine receptor (RyR) (Berridge, 1993) than L, N, T, and P Ca2+ channels (Snutch and Reiner, 1992). There is high homology in the pore structure domain between the RyR and the InsP, receptor, and both channels are very sensitive to Ca2+ and pH (Berridge, 1993). Thus, these factors, plus InsP,, may regulate the second Ca2+ channel operated during the AR, and one or more may link it to the first channel.

B. The Starfish Acrosome Reaction Starfish eggs are surrounded by an egg jelly layer which contains a mixture of biologically active compounds that regulate sperm function. It increases Ca2+ uptake, modulates [CAMP],raises pH, in a Na+-dependent fashion (Tubb er al., 1979; Matsui et al., 1986a, C; Hoshi et al., 1990, 1991), and induces the degradation of sperm histones (Amano et al., 1992a, b). In seawater containing > I 0 mM CaCl,, and at pH > 8.0, under nonphysiological conditions, a high molecular weight fucose sulfated glycoprotein (ARIS) from egg jelly was found to induce AR species-specifically (Ikadai and Hoshi, 1981a, b; Matsui et al., 1986a). Later, it was found that a cofactor (CoARIS), which by itself does not trigger AR, is necessary to achieve this reaction in normal seawater (Matsui et a/., 1986b). CoARIS is not species-specific and belongs to a family of sulfated steroidal saponins that freely diffuse from the outer layer of the egg (Nishiyama et a l . , 1987a, b). Some of these saponins are cofactors of ARIS (Nishiyama et al., 1987a) and some agglutinate spermatozoa (Uno and Hoshi, 1978). The carbohydrate and sulfate moieties of ARIS are believed to be responsible for the biological activity and species-specificity (Matsui et a l . , 1986a, b; Okinaga et al., 1992). ARIS or CoARIS alone can induce Ca2+ uptake and increases in [CAMP], but cannot trigger AR (Ikadai and Hoshi, 1981a, b; Matsui et al., 1986a). In contrast to egg jelly, ARIS and CoARIS together induce AR without increasing pHi (Hoshi er a l . , 1990). The lack of requirement for a pH, increase under these conditions establishes a difference between starfish and sea urchin spermatozoa. Addition of crude preparations of CoARIS and sperm activating peptides, fraction M8, to ARIS, enhances its potency to induce AR more greatly than

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when pure CoARIS is used. M8 increases pH, and sperm motility, and in so doing it may augment biological activity (reviewed in Hoshi et al., 1990). As in the case of sea urchin sperm, the coordinated action of multiple stimuli involved in activation of motility, respiration, chemotaxis, and AR optimizes the response of this cell under physiological conditions. Maitotoxin, a marine toxin which appears to activate Ca2+ channels, induces the AR in starfish Asrerina pectinferu spermatozoa only when fraction M8 is present (Amano et al., 1993a). This effect depends on external Ca2+ and is blocked by the Ca2+ channel blocker verapamil. Maitotoxin alone stimulates histone degradation in a verapamil-insensitive manner. Suspension of sperm in Na+-free seawater induces a Ca2+-dependent and Ca2+ channel blocker-insensitive histone degradation without triggering AR. However, 10-30 mM Na+ seawater induce a Ca2+-dependent, spontaneous AR which is blocked by diltiazem or by increasing K+ (30 mM KCl). These results suggest the participation of Ca2+ channels in the starfish AR (Amano et a / . , 1993a, b), as it probably occurs in all species, and also that K+ channels may play a yet to be defined role in sperm function.

C. The Mammalian Sperm Acrosome Reaction

It is generally agreed that the zona pellucida (ZP) is the main mediator of the sperm AR in mammals (Salig, 1990; Wassarman, 1990a,b; Bleil, 1991; Florman and Babcock, 1991; Ward and Kopf, 1993). In spite of this, other factors associated with the female reproductive tract may participate in triggering the AR at different stages of the sperm path, to ensure that only the fittest, acrosome-intact spermatozoa cross the ZP and fertilize the egg (reviewed in Kopf and Gerton, 1990). As described in section IIB, murine ZP consists mainly of three sulfated glycoproteins: ZP1, ZP2, and ZP3. ZP1 is a 200-kDa dimer that maintains the three-dimensional structure of ZP. ZP2 ( I20 kDa) appears to mediate the binding of reacted sperm to ZP (Bleil and Wassarman, 1980b, 1988). ZP3 (83 kDa) displays most of the sperm binding and AR-inducing activity of unfertilized eggs (Bleil and Wassarman, 1980a, 1988). The sperm-binding region of ZP3 is likely to be the highly glycosylated carboxy-terminal half of the polypeptide (Rosiere and Wassarman, 1992). Both protein and carbohydrate regions of ZP3 appear to be involved in its AR-inducing activity (Florman et al., 1984; Wassarman, 1990a). It is thought that a specific receptor(s) for ZP3, found on the plasma membrane of the acrosome-intact sperm overlying the acrosome, mediates its binding and induction of AR. After binding to ZP3 and undergoing the AR, sperm-ZP interactions remain throegh secondary contacts between ZP2 and its receptors located on the inner acrosomal membrane (reviewed in Wassarman, 1990a,b;

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Myles, 1993). Following ZP penetration, acrosome-reacted sperm traverse the perivitelline space, bind to, and then fuse at, distinct post-acrosomal regions of the sperm head (Yanagimachi, 1988; Wassarman, 1990). Shortly after spermegg fusion, the egg cortical granule reaction occurs and the enzymes that modify ZP are released. ZP2 is converted to ZP2, and ZP3 to ZP3,, resulting in the block to polyspermy. In this manner, acrosome-intact sperm cannot bind to the modified ZP3,, nor can acrosome-reacted cells maintain binding with ZP2, (Wassarman, 1988, 1990a, b). Recombinant cDNA experiments indicate that the ZP3 from mouse is a member of a highly conserved peptide family representing the physiological agonist of the acrosomal exocytosis in mammals. Its unique function is suggested by the limited amino acid sequence homology existing between ZP3 and other examined proteins (Ringuette et al., 1988). The gamete interactions required to attain AR are species-specific. This suggests the presence of specialized sperm receptors. Several putative receptors have been described and it remains to be seen which of them fulfills all the requirements of a bona fide receptor or, if several receptors are involved, how their action is coordinated. Binding studies with W - Z P 3 have shown the existence of specific and discrete binding sites in appropriate regions of mature spermatozoa (Bleil and Wassarman, 1986; Mortillo and Wassarman, 1991). Labeled ZP3 or glycopeptides that are active in binding and are derived from it, specifically crosslink to a 56-kDa protein (sp56) on acrosome-intact mouse spermatozoa and not to ZP2 (Bleil and Wassarman, 1990).The purified native protein behaves like a 110-kDa homodimer or homotrimer. lmmunohistochemical and immunoblotting studies with monoclonal antibodies recently revealed that sp56 is a peripheral membrane protein located on the outer surface of the sperm head plasma membrane, precisely where sperm bind ZP3 (Cheng et al., 1994). These results indicate that sp56 is involved in sperm-egg recognition in the mouse. It has been postulated that a sperm surface P-galactosyltransferase (GalTase) binds to 0-linked oligosaccharide residues on ZP3 (Shur and Hall, 1982; Shur, 1993). Cell-cell and cell-extracellular matrix interactions have been proposed to be mediated by such an enzyme in other systems (reviewed in Shur, 1993). In virro, mouse sperm GalTase glycosylates ZP3, but not ZP1 and ZP2, while a milk GalTase has no activity toward ZP proteins. However, it remains unclear if the specific activities of both enzymes are comparable (Miller et al., 1992). The enzyme from acrosome-reacted sperm does not react with ZP3, and ZP from fertilized eggs is no longer a substrate for galactosylation. Different results regarding the effect on its sperm-binding capacity of removal of enzymatic N-acetylglucosamine from ZP3 have been reported (Bleil and Wassarman, 1988; Miller et al., 1992). On the other hand, a 95-kDa sperm protein has been identified as a ZP3 receptor, using anti-phosphotyrosine antibodies (Saling, 1990). These antibodies identify proteins of 52, 75, and 95 kDa and bind to the acrosomal region of the

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sperm head (Leyton and Saling, 1989a). 1251-ZP3binds to a 95-kDa sperm protein recognized by the anti-phosphotyrosine antibodies. This protein is phoset al., 1992). Its phosphorylaphorylated in the presence of [ Y - ~ ~ P I A(Leyton TP tion increases when ZP is added and is reduced by the tyrosine kinase inhibitor tyrphostin RG-50864, which inhibits AR. It remains to be seen if this 95-kDa sperm receptor for ZP3 is a tyrosine kinase. Recently, a 95-kDa phosphotyrosine containing protein from epididymal mouse sperm that is phosphorylated in a CAMP-dependent manner during capacitation was reported (Duncan and Fraser, 1993). Why are there several ZP3 sperm receptor candidates? Work from several groups has indicated that multiple concerted and cooperative interactions between ZP3 and sperm surface, possibly involving receptor aggregation, may be needed to achieve the signal transduction events that result in the AR (Bleil and Wassarman, 1983; Kopf et al., 1989; Leyton and Saling, 1989b; Ward and Kopf, 1993). The next key question is: how do these receptors convey information to initiate signal transduction in mammalian spermatozoa? The ZP-induced AR is inhibited by Pertussis toxin (PTX), a specific inactivator of the Gi class of heterotrimeric G proteins, in mouse, bovine, and human spermatozoa (Endo et al., 1987; Florman et al., 1989; Lee et al., 1992). PTX ADP ribosylates the aisubunits of Gi. Multiple species of G proteins, such as Gi, and G,, which is PTX-insensitive, have been detected in mammalian sperm. G , has three subtypes (Gil, Giz, and Gi3),all of which have been found in mouse sperm (Glassner et al., 1991). The ai subunit of Gi is known to regulate adenylyl cyclase, phospholipase C, and various ion channels; nonetheless, the specificity of the subtypes of aiis not known (Katziro et al., 1991). PTX blocks the ZP3, or ZP-induced AR, but does not inhibit sperm-ZP binding. In contrast, spontaneous AR, or that induced by A23187 or progesterone, is insensitive to PTX (Endo et al., 1988; Lee et al., 1992; Tesarik et al., 1993). Using isolated mouse sperm plasma membranes, it has been shown that the Gi subtypes activated by solubilized ZP are Gi, and Gi,. These results suggest that ZP preferentially activates Gil and G , in sperm (Ward et al., 1994). Which of the receptor candidates mentioned earlier can activate these Gi proteins and what their targets are still need to be determined. Interestingly, mRNAs encoding for members of the superfamily of G protein-coupled receptors have been found in pachytene spermatocytes and round spermatids (Meyerhof et al., 1991; Parmentier et al., 1992). Are these proteins present in mature sperm? If so, what would their function be? Hopefully, these provocative questions will be answered in the near future. It would be expected that, as in many somatic cells, the physiologically relevant exocytosis in mammalian sperm mediated by Gi involves changes in second messengers and in plasma membrane ion permeability. It has been pointed out that alterations in the phospholipid and CAMP metabolism may participate in the induction of the AR by ZP and by nonphysiological agents like

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the Ca2+ ionophore A23187. Due to space limitations, no further discussion will be given to the latter (reviewed in Thomas and Meizel, 1989; Kopf and Gerton, 1990; Fraser and Monks, 1990; Florman and Babcock, 1991; Roldan and Harrison, 1993; Roldan and Fragio, 1994). Biologically active phorbol diesters and diacylglycerols modify the time course of the ZP-triggered AR (Lee et al., 1987). The activation of protein kinase C (PKC) was reported to enhance the AR in human capacitated spermatozoa in the absence of external Ca2+. Type-specific PKC antibodies revealed the presence of PKCa and PKCpII in the equatorial segment, while PKCpI and PKCr were found in the principal piece of the tail (Rotem et al., 1992). In this direction, an epidermal growth factor (EGF) receptor has been detected in the head of bull sperm. Binding of EGF induces 30% AR in capacitated sperm. This response is blocked by staurosporin, a PKC inhibitor, suggesting its participation in the EGF-induced AR (Lax er al., 1994). Clearly, much work is needed to understand the delicately choreographed participation of many sperm receptors and the role of phospholipid metabolism during mammalian AR. A raise in internal Ca*+ is an essential step in the ZP3 signaling path leading to the AR. External Ca2+ is required for the physiological AR (Yanagimachi, 1988) and ZP induces [Ca2+], increases that precede exocytosis in single sperm loaded with fluorescent ion indicator dyes (Florman et al., 1989; Storey et al., 1992). Bovine spermatozoa can also be loaded with pH,-sensitive dyes (Babcock and Pfeiffer, 1987). Solubilized ZP activates a transduction mechanism in mature sperm and not in immature sperm (Florman and First, 1988; Florman et al., 1989), resulting in a fast increase in pH, and [Ca2+], that leads to acrosomal exocytosis. These changes are linked to the AR and are somehow mediated by G, proteins, since both are inhibited by PTX (Florman et al., 1989, 1992). Lately it was proposed that a regulatory factor, isolated from bovine seminal fluids and identified as caltrin, couples ZP signal transduction to the mechanism that regulates [Ca2+], (Clark et al., 1993). Caltrin, a peptide isolated from bovine seminal fluids, had been shown to inhibit 45Ca2+ uptake by sperm and to have bactericidal activity (Rufo et al., 1982; Lardy and San Agustin, 1989). Addition of this peptide to cauda epididymal sperm efficiently couples ZP stimulus and the cell machinery that regulates [Ca2+1, (Clark et al., 1993). Evidence has been provided for the presence of voltage-dependent Ca2+ channels in the plasma membrane of mammalian sperm. Elevation of external K+ in ram (Babcock and Pfeiffer, 1987) and bull (Florman and Babcock, 1991) sperm induces a dihydropyridine-, benzothiazepine-, and phenylalkylamine-sensitive increase in [Ca2+],, which depends on extracellular Ca2+ and pH. High-affinity binding sites ( K , ca. 0.4 p M ) for the L-type, voltage-dependent Ca2+ channel antagonist PN200-110 were detected in sperm membranes from both of these species. Several inorganic divalent cations like Co2+ and Ni2+ and dihydropyridine antagonists equally block acrosomal exocytosis triggered by a combined elevation of pH, and a K+-induced depolarization (in mouse, bull, and ram

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spermatozoa) or by ZP3 in bull sperm. Depolarizing conditions that appear to open Ca2+ channels bypass the inhibition of the ZP3-induced exocytosis produced by pertussis toxin. The resting membrane potential of bull spermatozoa, determined with DiS-C,-(5), as was done in sea urchin sperm (GonzBlezMartinez and Darszon, 1987), was estimated at around -40 mV (Florman et al., 1992). These experiments indicate that opening of sperm voltage-dependent Ca2+ channels is enough to trigger the AR when pH, is increased and that the activation of these channels, which involves a pertussis toxin-sensitive G protein, is a required step in the ZP3 signal transduction pathway (Florman et d., 1992). In addition, the results strongly suggest the presence of K+ channels in the mammalian sperm plasma membrane, whose regulation remains to be studied. If Ca2+ channels need a depolarization to open, how does ZP depolarize sperm? If not, do second messengers activate the channel, maybe by altering its phosphorylation state? So far, no ZP-induced membrane potential changes have been reported. There is solid evidence today that ion channels from various cell types are regulated directly or indirectly by G proteins (Sterweis and Pang, 1990; Brown and Birmbaumer, 1990; Hille, 1992). It is difficult at this time to propose the G protein mode of channel regulation, since the type of Ca2+ channels involved has not been established. The fact that micromolar dihydropyridines block AR and the increase in [Ca2+],is not enough to identify the channels as L-type, since micromolars of this antagonist can also block T-type Ca2+ channels (Likvano et a / . , 1994). As in the sea urchin sperm, pH, and [Ca2*li appear to be intimately related and finely tuned in mammalian spermatozoa. The mechanisms that regulate and link these two key parameters are probably also critical for spermatogenesis and spontaneous AR and remain to be established. In human spermatozoa, progesterone and other progestins have been shown to induce large, extracellular-dependent increases in [Ca2+], which result in AR (Thomas and Meizel, 1989; Blackmore et al., 1990; Baldi et al., 1991). As mentioned earlier, this response is insensitive to PTX and to Ltype Ca2+ channel blockers, implying a different transduction path and Ca2+ uptake mechanism (Foresta et al., 1993). Since the progesterone-induced increase in [Ca2+], is associated with a membrane depolarization that is even larger in the absence of external Ca2+, it probably involves a Ca2+ channel permeable to monovalent cations (Foresta et al., 1993). These results suggest that, as in the sea urchin sperm, at least two different Ca2+ channels are present in mammalian spermatozoa. Progesterone metabolites have been shown to enhance the interaction of y-aminobutyric acid (GABA) with the GABA receptor in central nervous system neurons. The GABA receptor is a multisubunit protein containing a C1- channel (DeLorey and Olsen, 1992), and it has been detected in boar and ram sperm (Erdo and Werkele, 1990). It has been proposed that the fast, progestin-induced human sperm responses may involve steroid interaction with a sperm steroid

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receptor/Cl- channel complex similar to the GABA,/Cl- channel complex (Wistrom and Meizel, 1993). Regardless of the wealth of information indicating the fundamental participation of ion channels in mammalian sperm physiology, little is known about them (Darszon et a l . , 1994). Only a few reports have been published on single channel activity from mammalian sperm. Cationic channels, with a single channel conductance of 130 pS in 0.1 M NaCl, were recorded upon addition of a partially purified, 110-kDa human sperm plasma membrane protein to a preformed lipid bilayer. Apparently the channel was formed from functionally aggregated triplets, albeit no follow-up on this work has been published (Young et al., 1988). Two types of Ca2+ channels having single channel conductances of 10-20 pS and 50-60 pS were described in tip-dip bilayers formed from liposomes containing boar sperm plasma membrane. The smaller channels were partially blocked by unknown concentrations of nitrendipine and verapamil and completely blocked by 0.5 mM La3+ (Cox and Peterson, 1989). In another report, a nonselective, voltage-independent cation channel from cauda epididymal or ejaculated boar sperm plasma membranes, incorporated into planar lipid bilayers, was presented. Monovalent and divalent cations permeate the channel, which is blocked by verapamil (1 mM) and 50 pkt of either nitrendipine or ruthenium red (Cox et a l . , 1991). A recent abstract from this group using planar bilayers indicates that a Ba2+-permeable, 10-20 pS channel is blocked by micromolar dihydropyridines, and, although the voltage-dependence was not defined, it was identified as an L-type Ca*+ which could mediate Ca*+ uptake during the AR (Tiwari-Woodruff et al., 1994). The newly cloned cyclic nucleotide-gated channel from bovine testis is apparently the first sperm channel to be cloned and expressed into Xenopus oocytes (Weyand et al., 1994). The channel has 78% identical residues as the cone photoreceptor (for review see Yau, 1994); in oocytes at 60 mV it displays a single channel conductance of 20 pS, allows Ca2+ and monovalent cations through, and has a much higher affinity for cGMP (>lOO-fold) than for CAMP. It was not possible to detect cGMP-induced increases in [Ca2+], in fura-2-loaded sperm; however, such changes were observed in 10% of vesicles thought to be sperm cytoplasmic droplets. Small cGMP-induced currents associated with single channel transitions of > 10 pS were detected in these vesicles after swelling sperm in a similar fashion to sea urchin sperm (Babcock et a l . , 1992). Similar cGMPinduced currents were recorded in 7/150 attempts from inside-out patches of plasma membrane of human and bovine sperm. What region of the sperm plasma membrane was patch-clamped is unclear; further work is needed to demonstrate that the cloned channel is the one that was recorded. The authors of this chapter have initiated the characterization of the ion channels present in isolated mouse sperm plasma membranes in planar bilayers and have detected a high-conductance Ca2+ channel resembling the one from S. purpurarus sperm (see Fig. 3). It is attractive to consider that this Ca2+ channel

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I

C2+

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CI -

v

Fig. 3 Schematic diagram of the mammalian sperm acrosome reaction. Sperm AR is initiated when egg ZP3 binds to several possible receptors which may have to aggregate. Only the three main ones are shown: 95-kDa receptor protein ( l ) , sp56 (2), and GalTase (3). The receptors could then activate G proteins, sensitive (Gi) or insensitive (G?)to PTX, that directly or indirectly regulate Ca2+ and K+ channels and maybe phospholipases C (PLC), D (PLD), and Az (PLA,), which in turn could activate protein kinase C (PKC). Changes in E M and in (Ca2+],may participate coordinately in regulating pH, and adenylyl cyclase. How the possible second messengers are interrelated, who their targets are, and how their responses are organized is unclear. The bottom shows a novel strategy in the study of sperm ion channels. It consists of the addition of sperm cells to a preformed black lipid membrane (BLM) under fusogenic conditions. This eventually leads to the interaction between sperm and the lipid bilayer and to ion channel transfer to the lipid bilayer. The right part shows a current record of the large conductance CaZ+ channel obtained from mouse sperm at -60 and f60mV membrane potential.

may be important in sperm physiology, since it is present in such diverse species (BeltrAn et al., 1994). In addition, in this laboratory it has been possible to successfully incorporate ion channels to lipid bilayers directly using sea urchin

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and mouse spermatozoa (Beltrin et al., 1994; see Fig. 3). The high-conductance Ca2+ channel, several K+ channels, and a smaller, voltage-dependent 10 pS Ca2+ channel resembling the one described by Tiwari-Woodruff et al. (1994) have been recorded with this strategy. This approach opens new avenues for exploring cell-cell interactions, such as sperm-egg fusion, at the single channel level.

VI. Do Ion Channels Turn the Egg On? The answer to this question is definitely yes: one way or another, ion channels are at the heart of egg activation by sperm. The egg is much larger than the sperm, and because of this it is much easier to utilize electrophysiological techniques to study its permeability changes and its ion channels, and much is known about them (Whitaker and Steinhardt, 1982; Nuccitelli et al., 1989). This is not the field of the authors; the reader is referred to excellent reviews dealing with the mechanisms involved in sperm-egg fusion and egg activation (Nuccitelli et al., 1989; Jaffe, 1990; Nuccitelli, 1991; Shen, 1992; Whitaker and Swann, 1993; Myles, 1993; Foltz and Lennarz, 1993; Miyazaki et al., 1993; Swann et al., 1994). However, a brief summary of this area will be given, mainly as an excuse to emphasize the role of sperm components-possibly its ion channels-in this fundamental event. Studies on the fertilization in sea urchin eggs have contributed significantly to establishing the crucial role of intracellular Ca2+ homeostasis as a regulator of non-excitable cells (Steinhardt and Epel, 1974; Whitaker and Steinhardt, 1982; Steinhardt et al., 1974, 1977; Jaffe, 1980, 1983; Ponie et al., 1985). Sperm triggers a CaZ+ explosion at fertilization in deuterostome eggs (Jaffe, 1980, 1983). A Ca2+ wave is initiated where sperm and egg interact and travels at about 5-10 p,m/sec to its antipode (Gilkey et al., 1978; Jaffe, 1983; Eisen e t a l . , 1984; Busa and Nucitelli, 1985; Miyazaki et al., 1986; Swann and Whitaker, 1986; Yoshimoto et al., 1987; Kubota et al., 1987; McCulloh and Chambers, 1991). These transient (60-120 sec) increases in [Ca2+li(from -0.1 to -0.5-2 M ) cause cortical granule exocytosis, preventing polyspermy in echinoderms, frogs, and mammals (Epel, 1978; Jaffe, 1985; Yanagimachi, 1988). Fertilized eggs from hamster (Igusa and Miyazaki, 1986), mouse (Kline and Kline, 1992), pig (Sun et al., 1992), and cow (Fissore et al., 1992) are also able to undergo multiple transient increases in [Ca2+],. The explosive increase in [Ca2+], may also signal cell cycle progression (Whitaker and Patel, 1990; Ciapa et al., 1994) and other important aspects of cell function (Vitulo and Ozil, 1992; Tombes et al., 1992). Ca2+ transients and [Ca2+Ii oscillations are a common feature of somatic cell responses to a broad range of extra- and intracellular signals (Berridge, 1991, 1993). Both of them result from a finely tuned balance between regulated release

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and reuptake of Ca*+ from intracellular stores (reviewed in Tsien and Tsien, 1990; Berridge, 1991). It is generally accepted that the sites of Ca2+ release and reuptake are elements of the endoplasmic reticulum (Terasaki and Sardet, 1991; Berridge, 1993). Mainly two types of Ca2+ channels belonging to the same family are responsible for releasing this divalent: the inositol 1,4,5-trisphosphate receptor (InsP, receptor), modulated by InsP, (Ross et al., 1989; Furuichi et al., 1989; Mignery et al., 1989); and the ryanodine receptor (RyR) (Smith et al., 1988; Lai et al., 1988; Fleischer and lnui, 1989), modulated by Ca2+ itself (Endo, 1977; Lai et al., 1988; Takeshima et al., 1989) and, directly or indirectly, through calmodulin by a newly discovered second messenger, cyclic-ADP ribose (cADPr) (Lee et al., 1989; Galione et al., 1991; Lee et al., 1994). The InsP, receptor participates in InsP,-induced Ca2+ release (IICR), and the RyR in Ca*+induced Ca2+ release (CICR). Positive feedback can also occur as a result of Ca2+ stimulation of InsP, production (Whitaker and Irvine, 1984; Meyer and Stryer, 1988) and InsP,-induced increase in the InsP, receptor sensitivity to Ca2+ (Finch et al., 1991; Bezprovzanny et al., 1991; Missiaen et al., 1991). Basically, two mechanisms for sperm to turn the egg on can be envisaged (Epel, 1989; Jaffe, 1990; Turner and Jaffe, 1989; Whitaker and Swann, 1993): (1) Sperm may interact with receptor sites to alert the egg of its arrival and prepare its entry; (2) The sperm may fuse with the egg and deliver its activating message, which could include its baggage of second messenger-synthesizing enzymes, soluble factors, and/or its ion transport systems. Of course, a combination of the two might operate. Both mechanisms require specific and successful sperm-egg interactions. The recent molecular characterization of some of the actors involved in gamete recognition and fusion has contributed significantly to a better understanding of fertilization (reviewed in Foltz and Lennarz, 1993; Myles, 1993). In the sea urchin egg there is a 350-kDa membrane glycoprotein which functions as a sperm receptor (Foltz and Lennarz, 1990, 1992; Foltz et al., 1993). A 70-kDa fragment of this sperm receptor binds species-specifically to acrosome-reacted sperm. Antibodies against this fragment bind to the 350-kDa receptor, block fertilization, and have been used to clone it. The species-specific sperm-binding domain of the receptor structurally resembles the peptide-binding groove of hsp70 and major histocompatibility class 1 proteins. The sequence in this region is speciesspecific, while the cytoplasmic domain is conserved among echinoderms and is not homologous to any other sequences (Foltz et al., 1993). Although the sperm receptor does not appear to be a G protein-linked or tyrosine kinase receptor, it could transduce signals. In sea urchin eggs it has been shown that protein tyrosine kinase activity rapidly increases upon sperm binding (Ciapa and Epel, 1991). Recent results indicate that the sperm receptor is phosphorylated in response to sperm binding or addition of bindin, the complementary molecule on the sperm (Glabe and Vacquier, 1977; Vacquier and Moy, 1977; Glabe et al., 1991; Glabe and Clark, 1991; Lopez et ul., 1993), by a tyrosine

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kinase present in the egg surface (Abassi and Foltz, 1994). In somatic cells, receptor-mediated tyrosine kinase activation has been shown to release Ca2+ (Schlessinger and Ulrich, 1992). The biological active form of the sperm receptor appears to be a disulfide-linked multimer of 350-kDa subunits (Ohlendieck et al., 1994). Preliminary electrophysiological experiments indicate that sperm fusion to artificial black lipid bilayers is significantly enhanced by the addition of an homogeneous sperm receptor (K. Ohlendieck, W. J. Lennarz, A. Litvano, and A. Darszon, unpublished results). Thus, the sperm receptor may mediate gamete fusion. In guinea pig, a sperm membrane glycoprotein heterodimer (PH30-aP) is involved in sperm-egg fusion. PH30-a contains a short (22 amino acids) stretch, common to viral fusion peptides, which is relatively hydrophobic and modeled as a sided a helix (Blobel et al., 1992). This motif is thought to promote viral-host cell fusion (White, 1992). The N-terminal90 amino acids of the mature PH30-P have a high homology with the desintegrins, a family of ligands that binds to the integrin class of surface adhesion molecules. These ligands contain a tripeptide of arginine, glycine, and aspartic acid (RGD) in their binding domain. PH30-P instead has TDE (Blobel et al., 1992), and peptides that include this tripeptide prevent sperm-egg fusion (Myles, 1993). PH30 has the characteristics and location to participate in sperm-egg fusion; the integrin-like receptor in eggs remains to be discovered (Blobel et al., 1992). Once sperm and egg interact using their specific receptors, the beginning of the [Ca*+], transient can take from seconds in hamster eggs (Miyazaki and Igusa, 1981) to minutes in the mouse (Jaffe et al., 1983). This lag time, which could suggest an enzymatic reaction or diffusion limitations, is determined by the sperm (Igusa et al., 1983). In line with the first mechanism, where sperm interacts with a receptor, the finding that InsP, accumulates in eggs (Turner et al., 1984; Whitaker and Irvine, 1984) gave birth to the idea that a G protein could mediate egg activation regulating the polyphosphoinositide (PPI) metabolism. Injection of InsP, initiates the Ca2+ wave and egg activation in sea urchin (Whitaker and Irvine, 1984), frog (Busa et al., 1985), mouse (Kurasawa et al., 1989; Kline and Kline, 1992; Swann, 1992), and hamster (Miyazaki, 1988; Cran et al., 1988). These eggs are activated by GTPyS, a guanine nucleotide analog that turns on G proteins. However, GDPPS, an inhibitory analog, does not block the fertilization Ca2+ transient in sea urchin eggs, while heparin, an InsP, receptor antagonist, blocks the GTPyS-induced Ca2+ transient but not the one triggered by sperm. Diacylglycerol production is inhibited in sea urchin eggs injected before insemination with a Ca2+ chelator, indicating the PPI generation results from the [Ca2+Ii increase and is not caused by it (Crossley et al., 1991). These results cast some doubts about the role of G proteins in sea urchin eggs. In contrast, GDPPS blocks the sperm-induced Ca2+ transient but not the one induced by InsP, in hamster eggs, suggesting the participation of a G protein in fertilization (Miyazaki, 1988). In spite of this, down regulation of the G protein

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response with phorbol ester treatment of hamster eggs inhibits the Ca2+ transients triggered by GTPyS but leaves those induced at fertilization unaffected (Swann et al., 1989; Miyazaki et al., 1990). Even though there is solid evidence for the presence of the G protein-PPI-signal transduction system in eggs and its capacity to activate them, the amount it contributes during fertilization for each egg is unclear (reviewed in Whitaker and Swann, 1993; Miyazaki et al., 1993; Swann et al., 1994). According to the second mechanism, eggs activate after sperm-egg fusion occurs. In the sea urchin egg, a fast inward current, probably due to sperm ion channels, is the first detectable signal at fertilization, and it indicates spem-egg electrical continuity (Dale et al., 1978). It occurs before the Ca2+ wave and coincides with the moment at which the electrical block to polyspermy (Jaffe, 1976; Gould-Somero and Jaffe, 1984) operates (Shen and Steinhardt, 1984). It has been elegantly established that sperm-egg fusion precedes the Ca2+ transient (Hinckley et al., 1986; Longo et a l . , 1990; McCulloh and Chambers, 1992). Early on it was proposed that Ca2+ carried by the fertilizing sperm triggered the Ca2+ wave (Jaffe, 1980, 1983). Working against this idea (known as the “Ca2+ bomb”) is the fact that sperm incorporation into eggs is enhanced by treating them either with Ca2+ channel blockers (McCulloh et al., 1989) or with Ca2+ chelators to prevent [Ca2+],increases (Swann et al., 1992). On the other hand, injection of Ca2+ into hamster and frog eggs produces normal regenerative Ca2+ waves (Igusa and Miyazaki, 1983; Busa and Nuccitelli, 1985; Jaffe, 1990), indicating the presence of CICR, but not in sea urchin eggs (Swann and Whitaker, 1986). It is known that Ca2+ and InsP, are elevated in sea urchin and mammalian acrosome-reacted sperm (reviewed in Iwasa et al., 1990; Florman and Babcock, 1991; Darszon et al., 1994), and cGMP levels may be high in reacted sea urchin sperm (Garbers, 1989). Injection of cGMP activates the Ca2+ wave in sea urchin eggs (Whalley et al., 1992). At the present time it is not known if cADPr, the most potent sea urchin egg activating factor thus described (Clapper et al., 1987; Lee e f al., 1989; Galione et al., 1991), is present in sea urchin sperm. Recently it was shown that cGMP can stimulate production of cADPr in sea urchin eggs (Galione et al., 1993). When sperm fuse with the egg they incorporate all their second messengers, enzymes, and ion transport systems. At the time of fusion drastic changes occur for both sperm and egg. In principle, sperm arrive at the egg depolarized and with an alkaline pH,; fusion first hyperpolarizes sperm to the egg potential and then the egg depolarizes. These membrane potential changes will modulate the sperm voltage-dependent ion channels, now part of the egg, and those of the egg itself, possibly leading to changes in the local concentrations of Ca2+, CAMP, cGMP, and H+, among others. For the time being it is difficult to propose a defined mechanism where one or several of these changes could participate in egg activation; however, they deserve further consideration. In Urechis, a marine worm, a protein isolated from the sperm acrosomal granule causes electrical

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responses in oocytes with the same form, amplitude, and ion dependence as the fertilization potentials induced by sperm (Gould and Stephano, 1987). In the sea urchin, sperm could deliver into the egg, in addition to Ca2+ (Jaffe, 1990) and InsP, (Iwasa et al., 1990), cGMP (Whaley et al., 1992). The sperm guanylyl cyclase might be activated by the decreased egg pH, (Suzuki et al., 1984; Ward et al., 1985a, b; Vacquier and Moy, 1986; Ward et al., 1986; Bentley et al., 1986b). The local increase in cGMP, sustained enzymatically until the pHi of the egg increases, would stimulate synthesis of cADPr and activate the egg. On the other hand, a protein factor that induces the characteristic periodic Ca2+ transients in hamster eggs has been isolated from the sperm cytosol (Swann and Whitaker, 1990; Swann, 1990). Ca2+, InsP,, or GTPyS cannot do this (Swann and Whitaker, 1994). Could the soluble sperm factor be a specific kinase for sensitizing egg CICR or IICR (See Miyazaki et al., 1993)? Which of the Ca2+ release mechanisms operate at fertilization? Even though the Ca2+ transients found in different eggs apparently share many characteristics, including the modes of positive feedback described in section VI, most probably the mechanisms that generate them are distinct. InsP,R has been detected immunohistochemically in hamster eggs (Miyazaki et al., 1992) and in Xenupus oocytes (Parys et al., 1992) and functionally in frog, mouse, hamster, and sea urchin (Miyazaki et al., 1992; Swann et al., 1994). RyR has been detected using antibodies in sea urchin eggs (McPherson et al., 1992) and functionally in sea urchin and mouse eggs. Thus, the frog egg responds through InsP,-mediated Ca2+ release, IICR (Galione et al., 1993). The sea urchin egg is versatile; both InsP,R and RyR are functionally expressed (Fujiwara et al., 1990) and either InsP, or cADPr triggers the Ca2+ wave. The egg uses both CICR and IICR (Galione et al., 1993; Lee et al., 1993). The reason why Ca2+ injection is unable to induce the Ca2+ wave in sea urchin eggs is a bit mysterious. Mammalian eggs behave more like the frog egg; the fertilization Ca2+ wave is triggered exclusively by IICR (Miyazaki et al., 1992, 1993), although RyR-mediated CICR is present in the mouse egg (Swann, 1992). Finally, the fundamental questions about fertilization remain open. Is a G protein-mediated activation of PLC involved in egg signal transduction or can tyrosine kinases do it? Is fusion and a cytoplasmic messenger the answer for all eggs? What is the cytoplasmic messenger? Do sperm membrane components significantly contribute to the transduction machinery? How exactly does the Ca2+ wave, once started, propagate across the egg?

VII. Concluding Remarks Life is a wonder and fertilization its beginning. Among other bimolecules, ion channels lie at the heart of this process. These are exciting times for ion transport mechanisms; the powerful strategies of Molecular Biology, combined with new

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enhanced tools to study ion fluxes at the single molecule level, promise to uncover their masterful regulation during gamete signaling. Looking through another glass, these are sad times in the world, although wild, and perhaps idealistic, it is at least worth hoping that the marvelous mystery of nature and man’s curiosity to approach it will defeat his greed and allow him to look within himself and recognize his place as another simple creature in the dance of the universe.

Acknowledgments This work was supported by grants from CONACyT, DGAPA, and an International Research Scholar Award to A. D. from the Howard Hughes Medical Institute. The authors thank Lucia de De La Tone, Irma Vargas. and Otilia Zapata for technical help, and Pedro Labarca for discussion and ideas, and for his and Celia Santi’s work on the CAMP-regulated K channel. Felipe Espinoza’s suggestions are acknowledged.

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Molecular Embryology of Skeletal Myogenesis ludith M . Venuti and Peter Cserjesi Department of Anatomy and Cell Biology Columbia College of Physicians and Surgeons New York, New York 10032

I. Introduction 11. MyoD Family of Myogenic Basic HLH Factors (mHLHs) A. Cloning of mHLH Transcription Factors B . Functional and Structural Properties of mHLHs C. Regulation of mHLHs 111. Developmental Expression of mHLHs A. Somitogenesis B , Temporal and Spatial Patterns of mHLH Gene Expression C . Subdomains of mHLH Expression in Somites IV. mHLH Factors in Invertebrate and Nonmammalian Vertebrates A. Invertebrate mHLH Factors B , Nonmammalian Vertebrate mHLHs V. Mutational Analysis of mHLH Function A. Mutational Analysis of Invertebrate mHLH Genes B . Targeted Mutation of mHLHs in Mice VI. Early Activation of Myogenic Program A. Instructive Cues from Axial Structures B. Migratory versus Myotomal Myoblasts C. Analysis of Regulatory Elements in mHLH Genes VII. MEF2 Family of Transcription Factors A. MEF2 Gene Family of Transcription Factors B. Developmental Expression of MEF2s C. MEF2s as Regulators of Myogenesis VIII. Summary and Conclusions References

1. Introduction The last few decades have witnessed tremendous advances in our understanding of the molecular events regulating myogenic determination and differentiation. During this time we have come full circle from early descriptions of myogenesis in the embryo, through cloning and characterization of myogenic structural and Currenf Topics m Developmenid Biolosy. Vol. 34 Copyright 0 1996 by Academic Press, Inc. All rights of reprcduction in any form reserved

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regulatory genes, and back again to the use of embryos to understand the function of these genes in the context of the developing organism. The reasons for the rapid rate of discoveries are many. The identification and characterization of the MyoD family of myogenic helix-loop-helix transcription factors (mHLHs) are perhaps the most important. However, this was possible only with the discovery that mesodermal progenitors can be converted to the myogenic lineage (Taylor and Jones, 1979) and myogenic cell lines can be maintained in culture indefinitely and induced to differentiate on demand (Yaffe, 1968; Yaffe and Saxel, 1977). Skeletal muscle progenitors, myoblasts, can be cultured in a highly proliferative state for generations yet retain the ability to differentiate in response to an appropriate change in culture conditions (Fig. 1). In proliferating myoblasts, muscle-specific proteins and their RNAs are absent or expressed at very low levels. When cells are induced with appropriate cues, they will fuse to form multinucleate myocytes expressing skeletal muscle contractile protein genes. This dramatic switch in phenotype results in the coordinated activation of a battery of muscle-specific genes (Devlin and Emerson, 1978). Pursuit of common elements in the regulatory regions of muscle-specific structural genes and identification of the transcription factors that bind them were among the first strategies used to identify the regulatory molecules underlying myogenic gene

Mesodermal ProgenitoIs

Determination

Mitogens

Myotube

\

Myofiber

Fig. 1 The myogenic differentiation pathway involves (1) determination of mesodermal precursors as myoblasts, (2) differentiation of myoblasts into multinucleate myotubes, and (3) maturation of myotubes into skeletal myofibers expressing different combinations of muscle structural gene isoforms.

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activation. A second approach sought to identify determination factors involved in the commitment of mesodermal precursors to the skeletal muscle lineage. Convergence of these two approaches led to the identification and characterization of the MyoD family of mHLHs. These transcription factors have the remarkable ability to coordinate the activation of the skeletal muscle differentiation program and convert a number of different cell types to the myogenic lineage. Although many cell types can be converted to muscle by the mHLH factors some cell lineages are refractory. Those that convert appear to have amassed a number of other gene regulatory proteins that are likely to cooperate with the myogenic proteins to switch on muscle-specific genes (Schafer et al., 1990). It has now become apparent that the mHLHs require the activity of other transcription factors and it is through the cooperative interaction of these different regulators that the final myogenic phenotype is attained. Cloning and characterization of a second family of essential myogenic regulatory factors, the muscle enhancer factor 2 (MEF2) family, have generated insight into the cooperative nature of muscle-specific gene regulation and raised the possibility of alternative pathways for determination, regulation, and maintenance of the myogenic phenotype. This chapter focuses on developmental and genetic studies that have provided an understanding of the developmental functions of mHLHs and other myogenic regulatory factors during skeletal muscle determination and differentiation.

II. MyoD Family of Myogenic Basic HLH Factors (mHLHs) A. Cloning of mHLH Transcription Factors

Differences between myoblasts and their mesodermal precursors were thought to center around activation of muscle-specific “determination genes” in myoblasts. Evidence for the existence of a myogenic determination gene stemmed from early studies using genomic DNA transfection where myoblast DNA, but not fibroblast DNA, could induce fibroblasts to convert to stable myogenic lineages (Konieczny and Emerson, 1984; Lassar et al., 1986). Following 5-azacytidine treatment, which alters cytidine methylation patterns, C3HlOT1/2 fibroblasts (lOTl/2 cells) form colonies containing myotubes, adipocytes, chrondrocytes, or mixtures of these cell types that will give rise to stable determined lineages (Taylor and Jones, 1979). Transfection experiments using genomic DNA from 5-azacytidine-converted myoblasts (azamyoblasts), or other myoblasts, suggested that a single gene was responsible for myogenic conversion of the 10T1/2 cells. The first myogenic regulatory factor, MyoD, was identified in a subtractive hybridization screen exploiting the ability of 5-azacytidine to convert 1OT1/2 fibroblasts to myoblasts (Davis et a l . , 1987). MyoD is expressed in myoblasts but not fibroblasts, and has the remarkable ability to convert fibroblasts to the myogenic lineage. Subsequently, three related members of this multigene family

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were identified in mammals. These include myogenin (Edmondson and Olson, 1989; Wright et a/., 1989), Myf5 (Braun et al., 1989a), and MRF4/herculin/ Myf6 (Rhodes and Konieczny, 1989; Miner and Wold, 1990; Braun et a l . , 1990a). Myogenin was initially cloned on the basis of its differential expression during the myoblast-to-myotube transition, whereas MRF4 and Myf5 were identified on the basis of heterologous library screens using MyoD cDNA as a probe. These genes show sequence homology in a basic helix-loop-helix (bHLH) domain. When driven by a viral promoter their transfected cDNAs convert cell lines of various origins to the myogenic phenotype; the transfected cells align, fuse, and express muscle-specific markers in response to appropriate culture conditions (Davis et al., 1987; Weintraub et al., 1989; Choi et al., 1990), demonstrating that all four factors can function as myogenic determination genes. Activation of the array of muscle-specific genes that accompanies myogenic differentiation is thought to occur through common regulatory elements in their promoters and enhancers that bind muscle-specific transcriptional regulatory proteins. Analysis of the regulatory regions of genes activated in myogenic cells led to identification of elements important for muscle-specific gene expression. Among the first cis elements identified were the myocyte enhancer factor elements MEFl and MEF2 in the muscle creatine kinase (MCK) enhancer. Gel retardation and DNA footprinting show MEFl elements bind proteins only from muscle nuclear extracts and deletion or mutation of these elements disrupts MCK activation, establishing MEF 1 elements as important regulatory sites for MCK gene expression (Buskin and Hauschka, 1989). Shortly after cloning, MyoD was shown to bind the MEFl motif in the MCK and myosin light chain (MLC113) enhancers (Lassar et aE., 1989). All members of the mHLH family are now known to recognize and bind to the MEFl DNA motif, commonly called an E-box, which is found in the regulatory regions of numerous muscle-specific genes. A second element shown to be important for MCK enhancer function, MEF2, is an AT-rich element located upstream and downstream of the MEF-1 motif. Deletion of the downstream MEF-2 element substantially reduces the activity of the MCK enhancer (Gossett et al., 1989; Horlick et al., 1990). Mutation of the upstream element silences all MCK enhancer activity (Cserjesi et a l . , 1994). The MEM family of genes has now been cloned and shown to be a second family of important myogenic regulatory factors that can interact directly with the mHLHs to confer muscle specific expression.

B. Functional and Structural Properties of rnHLHs

The mHLH proteins contain an evolutionarily conserved core consisting of a basic region, which is involved in DNA binding, and a helix-loop- helix (HLH)

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domain, through which proteins dimerize. These domains are shared by a large family of transcription factors, the HLH gene family, whose members regulate cell fate determination and proliferation in numerous organisms (Murre et al., 1994). A large number of bHLH proteins bind a DNA element containing the E-box motif, CANNTG. Since site-directed DNA binding by mHLHs is essential for both determination and transcriptional activation of muscle-specific downstream genes (Davis et al., 1990), it has been proposed that specific E-box motifs carry the recognition code for myogenic determination. E-box binding sites for the mHLH regulatory proteins have been identified in the regulatory regions adjacent to most skeletal muscle-specific genes as well as in the regulatory regions of the mHLH genes themselves. All mHLHs can bind to the E-box motif as heterodimers formed with the ubiquitous bHLH transcription factor E l 2 (Braun et al., 1990b; Brennan and Olson, 1990; Lassar et al., 1989, 1991). Transcriptional activation domains reside at both the amino and carboxy portions of the mHLH proteins (Braun et al., 1990b; Weintraub et al., 1991; Schwarz et al., 1992), but how these domains exert an effect on the transcription initiation complex remains unclear. The DNA-binding domains of the mHLH factors are essential for the muscle specificity of transcription. Domain-swapping experiments, in which the basic regions of MyoD or myogenin are replaced by the basic domain of the ubiquitously expressed bHLH E12, show that binding ability is not altered, but the ability of these chimeric proteins to convert 10T1/2 fibroblasts or activate reporter genes is severely affected (Weintraub er al., 1991; Brennan et al., 1991a). Systematic mutagenesis of the bHLH region of MyoD and myogenin has defined two amino acid residues, an alanine and a threonine, that are conserved in the basic domains of all mHLHs, mediate recognition of the E-box, and impart muscle specificity (Davis et al., 1990; Brennan et al., 1991a; Weintraub et a l . , 1991). In transfection experiments the mHLH myogenic determination genes appear equivalent in their ability to convert nonmuscle cells to the skeletal muscle lineage (Weintraub et al., 1989; Choi et al., 1990). The mHLHs have the potential to autoactivate and to cross-activate each other (Thayer et al., 1989; Braun et al., 1990b; Edmondson et al., 1992; Naidu et al., 1995). The ability of each mHLH to convert cells to the myogenic phenotype may be due to its ability to cross-activate other members of the mHLH family. For this reason, they are often present in the same cells or tissues simultaneously, and differential roles in gene regulation are difficult to ascertain. Muscle cell lines express each mHLH in a distinct temporal sequence during differentiation. MyoD and/or Myf5 are expressed in myoblasts prior to differentiation, myogenin is expressed in all cell lines after they are induced to differentiate, while MRF4 is usually not expressed until well after myogenic differentiation is induced in vitro (Montamas et al., 1991). These results suggest that MyoD and Myf5 may have predifferentiation functions and myogenin and MRF4 have differentiation or postdifferentiation functions. Temporal differences in the ex-

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pression patterns of the mHLHs during cellular differentiation in vitro suggest the mHLHs play distinct roles in myogenesis and are not entirely redundant. Differences in transactivation ability of mHLHs also suggest potential differences in their functions in vivo. MRF4 preferentially transactivates the epsilon subunit of the acetylcholine receptor promoter (Sunyer and Merlie, 1993) but is less efficient than other mHLH factors in transactivating MCK and troponin I enhancers (Yutzey et al., 1990; Chakraborty and Olson, 1991). Activation of myogenesis by expression of Myf.5 in 10TlI2 cells is associated with expression of MyoD, but activation of myogenesis with MyoD is not accompanied by MyfS expression (Braun et al., 1989a; Montarras et al., 1991), indicating that MyfS is not a target for MyoD activation but that MyfS may activate MyoD. A difference in the function of MyoD and myogenin in the transcriptional activation of the chicken M X l F gene also has been reported (Fujisawa-Sehara et al., 1992). This gene has two E-boxes in its enhancer; one preferentially responds to MyoD, the other to myogenin (Asakura et al., 1993). In an elegant experiment designed to identify the specific targets of MyoD regulation, a hormone-inducible MyoD expression vector was employed. Hormone-induced MyoD expression led to an upregulation of myogenin but not other muscle genes such as MyoD, MCK, or cardiac a-actin (Hollenberg et al., 1993). This experiment suggests that the primary target of MyoD transcriptional regulation is myogenin. Although the mHLHs bind the same DNA sequence and can convert cells to the myogenic lineage, they demonstrate differences in transactivational potential and gene targets.

C. Regulation of mHLHs In skeletal muscle cells, as in most cells, an antagonism exists between proliferation and differentiation. When muscle cells fully differentiate, they irreversibly withdraw from the cell cycle. Terminal differentiation in cultured myoblasts requires serum deprivation in addition to expression of the mHLHs. Serum components or purified growth factors such as basic fibroblast growth factor (bFGF), transforming growth factor P (TGF-P), or epidermal growth factor (EGF) will prevent overt differentiation and block transcription of muscle genes. The ability of the mHLHs to activate myogenic-specific transcription is negatively regulated by peptide growth factors. Growth factors act at several levels: they block the expression of mHLHs (Vaidya et al., 1989) and prevent existing mHLHs from activating transcription (Li et al., 1992a,b; Brennan et al., 1991b; Martin et a l ., 1992). C2C12 myoblasts constitutively express MyoD and are committed to the myogenic lineage but are prevented from cell cycle withdrawal and differentiation under growth conditions by high concentrations of mitogens. Since determined myoblasts express mHLHs in growth medium but cannot differentiate, suppression of mHLH function must occur during myoblast proliferation. The behavior of mHLHs in

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the presence of TGF-P is reminiscent of the behavior of basic domain mutants that bind DNA but cannot activate muscle-specific transcription (Brennan et al. , 1991b; Martin et al., 1992). The ability of growth factors to suppress myoblast differentiation in the presence of high levels of mHLHs suggests other factors are required for myogenic differentiation. Activation of protein kinases that phosphorylate and inactivate the mHLH factors is one mechanism proposed to explain the effect of growth factors on myogenic activity. Phosphorylation of myogenin and MyoD by protein kinases A and C at a number of residues, including mHLH-specific residues in their basic domains, inactivates their myogenic activity (Li et a l . , 1992a,b). This may not be a general mechanism for all mHLHs, however, since basic FGF apparently inhibits MRF4 activity independently of the phosphorylation status of the threonine residue in the basic domain (Hardy et al., 1993). Another mechanism of growth factor-dependent repression of mHLHs is mediated by the Id (inhibitor of DNA binding) family of HLH proteins. Stable overexpression of Ids in muscle cells in v i m inhibits myogenic differentiation (Jen el al., 1992). Mitogens block terminal differentiation of myogenic cells by promoting expression of the Ids, Ids lack basic domains yet heterodimerize with the E2A gene products, E l 2 and E47, making them inaccessible to the mHLHs for heterodimerization (Benezra et al., 1990). Ids are broadly expressed during proliferation and are downregulated on differentiation and withdrawal from the cell cycle. In the embryo Id transcripts are downregulated in regions where mHLH transcripts are expressed (Wang et al., 1992; Evans and O’Brien, 1993). This mutually exclusive expression of Id and mHLHs during embryogenesis supports a role for Ids as negative regulators of myogenesis and a model in which Id inhibits muscle cell differentiation by associating with E2A proteins, preventing them from complexing with the muscle determination gene products.

111. Developmental Expression of mHLHs A. Somitogenesis

Paraxial mesoderm gives rise to all skeletal muscle in the trunks and limbs of vertebrate embryos (for reviews see Buckingham, 1992; Wachtler and Christ, 1992). The paraxial mesoderm segments into somites, which further compartmentalize into the sclerotome, dermatome, and myotome. These compartments give rise to several mesodermal derivatives including bone, cartilage, dermis, and skeletal muscle. The initial stage in somite differentiation, the epithelial somite, is an epithelial ball (Fig. 2A). In the mouse embryo, the first somite forms on embryonic day 8 (E8), with subsequent somites forming in a rostra1 to caudal progression until E12-E13. As the somites mature, cells in the ventromedial region lose their epithelial organization, become mesenchymal, and form the sclerotome. Sclerotomal cells migrate away from the somite to surround

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. 9

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C. Rostra1

Q

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myotome sclerotome dermatome

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lateral somite

A. -Caudal

fig. 2 Somitogenesis and myogenesis proceed in a rostral to caudal progression in the developing vertebrate embryo. Somites in the rostral portion have compartmentalized into the various somitic derivatives while somites in the tail are just segmenting from the paraxial mesoderm. (A) The initial stage of somite formation is the epithelial somite; the only mHLH factor expressed at this stage is MyfS. (B) Subsequently, somites compartmentalize into the dermamyotome and sclerotome. All the mHLHs are expressed at this stage. (C) As development proceeds cells first migrate away from the sclerotomal region to surround the axial structures and form the axial skeleton. Concurrently cells migrate away from the dermamyotome to form the myotome and migratory myoblasts. mHLHs are expressed in myotomal myoblasts but are not expressed in migratory myoblasts until they reach their final destination.

the notochord and neural tube and form the axial skeleton (Fig. 2B and C). The remaining somitic cells retain their epithelial character and form a bilaminar structure, the dermamyotome (Fig. 2C). All axial and appendicular skeletal muscle develops from the dermamyotome (Christ et al., 1977). Initially cells move from the dorsomedial edge of the dermamyotome, positioning themselves beneath to form the myotome (Fig. 2B). Myoblasts within the myotome immediately begin to express skeletal muscle-specific markers including transcripts for the mHLHs. Cells also delaminate from the ventrolateral edge of the dermamyotome and migrate to the limb and body wall, where they later express skeletal muscle markers (Fig. 2C). Elegant chick-quail transplantation experiments have identified two populations of myogenic cells in the somite (Ordahl and LeDouarin, 1992). Progeny from the medial region of the somite produce muscles of the trunk whereas the lateral region of the somite generates migratory myoblasts that colonize the limb buds and body wall. It is not clear how these two populations of myoblasts are initially determined, but it is likely that external cues or signals that initiate or activate differentiation of the myotomal myoblasts are distinct from those that activate the myoblasts of ventrolateral somite origin.

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6. Temporal and Spatial Patterns of mHLH Gene Expression

Each of the mHLH genes is expressed in a unique temporal and spatial pattern during skeletal muscle development in the mouse (Fig. 3). In situ hybridization with antisense riboprobes shows mHLH transcription occurs sequentially during somitogenesis and myogenesis and that the timing of expression of these transcription factors differs in myoblasts originating from different parts of the somite. Myf5 mRNA is the first mHLH detected in epithelial somites at E8 and is expressed in the embryo until E l 4 (Ott et al., 1991). Although MyfS is not expressed in the unsegmented paraxial mesoderm, it is expressed in somites prior to compartmentalization (Fig. 2A). Early expression of Myf.5 in the uncompartmentalized somite suggests a role in the initial commitment of cells to the myogenic lineage. The first MyfS transcripts are located in the dorsomedial comers of the newly formed somite and later remain associated with the dorsomedial edge of the dermamyotome. Myotomes form at E8.5 concurrent with the initial appearance of myogenin transcripts, which continue to be expressed in the myotomal cells as they differentiate (Sassoon et a l . , 1989). MRF4 mRNA is expressed transiently in the somite between days El0 and 1 1, and is reexpressed

insomites

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fig. 3 The mHLHs display distinct temporal and spatial expression patterns in the developing mouse embryo. (Compiled from Sassoon er a l . , 1989; Bober er al., 1991; Ott et al., 1991; Hinterberger et al., 1991.)

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at El6 throughout the muscle-forming regions of the embryo to become the most abundant mHLH expressed after birth (Bober et al., 1991). Finally, MyoD mRNA appears around E10.5 in both somites and limbs and is present throughout embryonic development (Sassoon er al., 1989). This pattern of mHLH expression recurs as each somite differentiates. Since somites form and mature in a rostral to caudal progression, the paraxial mesoderm in the tail is beginning to segment as more rostral somites are compartmentalizing into the dermatome, myotome, and sclerotome. In the developing limb the temporal pattern of expression is different from that observed in the somite (Fig. 3). As in the somites, Myf5 is the first mHLH to be expressed in the limb and its expression is transient (Ott et al., 1991). Myogenin and MyoD mRNAs are coexpressed in limb myoblasts beginning at day E10.5, in contrast to their pattern in the somite, where they appear sequentially. MRF4 mRNA is not expressed in the limb myoblasts until late in development, first appearing at E l 6 (Bober et al., 1991). The migratory myoblasts leaving the ventrolateral edge of the somite to colonize the limb and body wall (Fig. 2C) do not express mHLHs until they reach their final destination (Sassoon et al., 1989). This suggests that the signals or cues that activate the mHLHs in the medial somite are distinct from those that activate mHLH expression in migratory myoblasts that arise from the lateral somite. The temporal expression pattern of the mHLHs in vivo is not recapitulated exactly in myoblasts differentiating in vitro. All myogenic cell lines express Myf5 and/or MyoD mRNA at low levels in myoblasts and on differentiation upregulate their expression (Braun et al., 1989a). All myogenic cell lines express myogenin after cells differentiate (Wright et al., 1989; Edmondson and Olson, 1989) while MRF4 is expressed several days after myoblasts have fused to form myotubes (Montarras et al., 1991; Miner and Wold, 1990). The expression of Myf5 transcripts in the epithelial somite prior to myoblast differentiation and before other mHLHs are expressed makes it the most likely candidate for a true myogenic “detennination gene.” In contrast, the late expression of MyoD in the somite, after myogenin and MFiF4 are expressed, suggests a postdetermination function for MyoD. That individual mHLHs show distinct temporal patterns of expression during myogenesis in vivo argues they each have distinct functions in the embryo. Different combinations of myogenic factors may be required for the establishment of distinct myogenic lineages that arise from the medial and lateral somite. Individual mHLHs or different combinations thereof may define distinct populations of myoblasts.

C. Subdomains of mHLH Expression in Somites

An analysis of mHLH protein expression during somitogenesis and myogenesis in vivo provides additional understanding of the roles of the individual mHLHs during development. Immunocytochemical analysis shows each of the four

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mammalian mHLH proteins has a distinct expression pattern within developing somites and at least two subdomains of myogenic cells can be identified within the somite (Smith et al., 1994). Myf5 protein is detected at E8 in dorsal anterior cells prior to formation of the dermamyotome, confirming that Myf5 is the first mHLH to be activated in the developing somites of the mouse. Myogenin protein is detected next and MRF4 protein appears approximately 12 hr later. MyoD protein first appears at E9.5 in a few cells in the ventral somite and by El 1.5 expression has spread throughout the dermamyotome, with highest levels of expression in the ventral region. By day 9.5 myogenin is expressed in myocytes throughout the myotome, while MRF4 is expressed in fewer cells and only in a dorsal subdomain. At early stages of somite development, the expression of MyfS and MyoD protein appears to be mutually exclusive, but as somites mature their expression overlaps. In newly compartmentalized somites, myogenin is the only mHLH in the ventral myotome prior to expression of MyoD. This early solo expression suggests myogenin may have a role in early events of myogenesis in cells derived from this region of the somite. In contrast to previous reports that myogenic differentiation occurred in somitic cells in the absence of any detectable mHLH expression (Cusella-De Angelis et al. , 1992), immunohistochemical analysis of mHLH protein expression in the differentiating somites in vivo found no evidence for myogenic differentiation in the absence of mHLH expression (Smith et al., 1994). MHC is always coexpressed with myogenin and occasionally with MRF4, but not always with MyoD or Myf5 (Smith et al., 1994). The reason for the conflicting results is unclear, but resolution is important since a lack of mHLH expression in MHC-expressing somitic cells implies an mHLHindependent myogenic pathway of skeletal muscle differentiation. mHLH protein localization suggests myogenic cells arise from multiple cellular origins and molecular pathways. The distinct temporal and spatial expression patterns of the mHLHs during myogenesis in the mouse embryo argues for a unique role for each factor in the generation of different myogenic lineages. Since MyoD and MyfS are expressed in discrete subdomains of the somite they may play a role in the establishment of the myotomal versus migratory myogenic lineages (Smith et al., 1994). Whether these different expression patterns are truly indicative of different populations of myoblasts is not entirely clear. Further analysis of these somitic cells is needed to determine whether the different subdomains represent myoblasts expressing individual mHLH proteins or different combinations of the mHLHs.

IV. mHLH Factors in Invertebrate

and Nonmammalian Vertebrates A. Invertebrate rnHLH Factors

The importance of the mHLHs in skeletal muscle development is reinforced by the structure and expression of homologs in invertebrates and nonmammalian

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vertebrates. The sequence conservation among these diverse mHLHs in the functionally important bHLH domain suggests they have similar myogenic regulatory functions. While most vertebrates have four mHLH genes, all invertebrates possess a single mHLH gene. The three-exon gene structure conserved among vertebrate mHLH genes is lost in invertebrates. The sequences of invertebrate mHLHs are similar only within the bHLH region. Sequence divergence outside the bHLH prevents classification of invertebrate mHLH factors as specific homologs of any one mammalian mHLH gene. Although the bHLH regions of the mHLHs are conserved across evolution, the specifics of the myogenic pathways in which these mHLHs operate are diverse. Consequently, temporal and spatial patterns of expression of these genes in relation to the myogenic events of an embryo are variable. Nevertheless, analysis of the expression and regulation of these genes has provided further understanding of the functions and hierarchical relationships of the mammalian mHLHs. In Caenorhabditis elegans (Krause et al., 1990), Drosophila (Michelson et a l. , 1990; Paterson et al. , 1991), sea urchins (Venuti et a l . , 1991), and ascidians (Araki et al., 1994) a single mHLH gene has been identified. The C . elegans homolog of the mammalian mHLHs, CeMyoD, is activated and its protein expressed in early multipotential lineages of the 28-cell embryo. These cells give rise to body wall muscle and other cell types. CeMyoD is expressed prior to any overt signs of muscle terminal differentiation in the nucleus of body wall muscle cell precursors; later in development expression becomes restricted to cells undergoing myogenic differentiation. The expression of CeMyoD in multipotential cells that give rise to nonmuscle as well as muscle suggests either negative regulatory mechanisms restrict CeMyoD activity in cells not destined to become muscle or positive coregulators of myogenic commitment are lacking in nonmuscle cells. The homolog of mammalian mHLHs in Drosophila, nautilus, is restricted to muscle pioneer cells that are specified by the mesoderm-specific factors, twist and S59 (Bate et al., 1991; Michelson et al., 1990; Paterson et al., 1991). nautilus is expressed in muscle precursors just prior to cell fusion. It is restricted to the precursors of the body wall muscle and is not expressed in other muscle lineages of the Drosophila embryo. Unlike the myogenic factors in vertebrate muscle cells, the Drosophila mHLH is expressed at much lower levels in differentiated Drosophila muscle (Paterson et al. , 1991). In sea urchins the single mHLH gene identified, SUM- 1, is expressed prior to the overt expression of muscle differentiation markers such as MHC (Venuti el a l . , 1991, 1993). SUM-1 protein is restricted to muscle precursors, but SUM-1 transcripts are detected in cells not destined to form muscle (J. Venuti, unpublished observation). These results are similar to those observed in C . elegans, where expression is not restricted to myogenic precursors (Chen et a l . , 1992, 1994). Together these results suggest that posttranscriptional regulatory mechanisms may be involved in regulating mHLH function in these invertebrate embryos.

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The exact spatial pattern of expression of the ascidian mHLH homolog, AMDl, has not been determined. AMDl is expressed in the early ascidian embryo prior to the onset of expression of overt muscle genes such as MHC, and is restricted to the body wall muscle in the adult (Araki et al., 1994). Analysis of the developmental expression of the invertebrate mHLH genes during development suggests they are involved in determinative events of myogenesis in these diverse organisms. Further analysis of myogenesis in these different species will provide a better understanding of the roles the mHLHs play in myogenic commitment and differentiation.

B. Nonmammalian Vertebrate mHLHs The pattern of expression of the mHLH genes in avian embryos is somewhat different from that observed in the mouse. The avian MyoD homolog is the first mHLH factor to be expressed in the somite followed by expression of myogenin and MyfS. These are first observed in the dorsomedial region of the somite and later are expressed throughout the myotome (de la Brousse and Emerson, 1990; Pownall and Emerson, 1991). This temporal sequence of expression contrasts with that observed in mouse myogenesis, suggesting MyoD and Myf5 might be interchangeable in the earliest events of the somite where both function as “determination factors.” In Xenopus laevis, the temporal pattern of mHLH expression is also different from that observed in other vertebrates. A maternal MyoD homolog (XMyoD) is expressed throughout the embryo during early embryogenesis and is not specifically localized to regions fated to give rise to muscle (Hopwood et al., 1989; Harvey, 1990). The initial broad expression of XMyoD eventually becomes restricted to fully committed muscle cells. Zygotic XMyoD transcription is activated following mesoderm induction and is thought to be the earliest musclespecific response to this event. The broad, early expression of XMyoD would suggest that XMyoD by itself is insufficient to convert cells into muscle. XMyoD appears to be under negative control in the frog embryo and is restricted to the cytoplasm until mesoderm induction, when it is able to enter the nucleus (Rupp et al., 1994). The Xenopus MRF4 homolog appears to accumulate only in the more mature muscle of the frog and Xenopus myogenin does not appear to be expressed at significant levels during development (Jennings, 1992). Lack of myogenin expression during Xenopus development indicates that other mHLHs may substitute for myogenin during embryogenesis in the frog. As in the avian system, the temporal pattern of expression of the different mHLHs in Xenopus is distinct from that observed in mammals. Reversal in temporal expression patterns of mHLHs during development in different organisms implies muscle differentiation programs vary among these organisms and the vertebrate mHLHs may function redundantly. The evolutionary conservation of mHLH genes has made possible the study of

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myogenesis in a diversity of organisms that are accessible to genetic, molecular, and embryological manipulations. Although the different species exhibit variations in fundamental mechanisms of myogenesis, analysis of the mHLH genes in the context of these developing embryos allows dissection of alternative pathways leading to myogenic commitment and differentiation. The absence of restricted expression to muscle progenitors in several nonmammalian species argues for either more complex regulatory interactions in these lower organisms or for a divergence in function from that observed in higher organisms. Evolutionary analysis of the amino acid sequences of the mHLH family members across several species suggests all are derived from a single ancestral gene and that the vertebrate genes were derived by gene duplication (Atchley et a l . , 1994). The ancestral gene split into two lineages early in the evolution of vertebrates; MyoD and MyfS evolved from one lineage and myogenin and MRF4 arose from the other. A proposed consequence of evolution by gene duplication is that different mammalian mHLHs may have preserved a redundancy in function. A common genetic origin predicts functional redundancy between MyoD and MyfS and between myogenin and MRF4. Targeted mutations in the mHLH genes demonstrate these pairs of genes do functionally substitute for each other during myogenesis in viva.

V. Mutational Analysis of mHLH Function A. Mutational Analysis of Invertebrate mHLH Genes

Analysis of mHLH gene function in invertebrates, where a single mHLH gene exists, avoids the complexity that might arise from redundancy among the four mammalian mHLHs. The first genetic analysis of an mHLH was reported for the invertebrate C . elegans (Chen et al., 1992, 1994). Genetic deficiencies spanning the CeMyoD gene locus, hlh-1, were isolated. Mutations of these regions are lethal, but early muscle development proceeds normally. Animals make the correct number of body wall muscles, but sarcomeres are disorganized. Embryos containing these deficiencies exhibited extensive body-wall muscle differentiation as determined by the expression of muscle-specific markers but only weak contractility. Since there is no evidence that maternal messages can support myogenic differentiation (Chen et al., 1992, 1994), zygotic expression of the single mHLH in C. elegans is not essential for determination and differentiation of the muscle precursors. In Drosophila, embryos containing deficiencies deleting the nautilus gene also show disorganized muscle, and express muscle structural genes, but are missing subsets of muscle cells (S. A. Abmayr, personal communication). Neither nautilus nor hlh-1 is required for muscle gene expression or the establishment of the myogenic lineage during embryonic development. These results are difficult to reconcile with our current understanding of

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mHLH function and strongly suggest alternative pathways for rnyogenic determination and differentiation exist.

B. Targeted Mutation of mHLHs in Mice

To ascertain the functions of the mHLH genes during myogenesis in vivo, homologous recombination was used to disrupt the individual mHLH genes in mice. The results of these studies (summarized in Fig. 4) have confirmed and extended our understanding of the importance of these transcription factors in mammalian myogenesis. MyoD was the first mHLH to be mutated by homologous recombination. Homozygous MyoD mutants show no obvious muscle defect; mice were viable and fertile and appeared morphologically indistinguishable from nonmutant littermates (Rudnicki et al., 1992). Histological examination and RNA analyses of muscle structural genes did not reveal overt differences in the skeletal muscle of mutant mice. The only discernible difference between MyoD mutants and controls was an upregulation and maintained expression of MyfS mRNA in skeletal muscle. These results suggest that (1) MyoD may normally repress MyfS expression; (2) MyfS can substitute for MyoD during myogenesis in vivo; and (3) MyoD can be completely eliminated from the myogenic pathway without any

Mice homozygous null for: MyoD

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Fig. 4 Summary of targeted mutations of the mHLHs in mice. X, Stage of myogenic differentiation achieved in these mutant mice. (Adapted from Rudnicki and Jaenisch, 1995.)

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deleterious effects on the development of the embryo. These experiments confirm that the mHLH genes have redundant functions in vivo. Myfs-null mice survive to birth, but in contrast to the MyoD-null mice, they die perinatally due to defective rib development and an inability to respire normally (Braun et al., 1992). The distal portions of the ribs necessary for proper insertion of the diaphragm fail to develop. Skeletal myofibers in the Myfs mutant mice appear normal although myotome formation is delayed in the Myfs mutants until MyoD is expressed at E10.5. Abnormal differentiation of the sclerotome from which the axial skeleton and ribs derive is attributed to this delay in myotome formation. Since Myf5 is the first mHLH factor to be expressed during mouse embryogenesis (Ott et al., 1991) and MyoD is not expressed until later (Fig. 3; Sassoon et al., 1989), MyfS must be required for the normal formation and interactive events of the early myotome. MyoD can substitute for MyfS in myogenesis, but it is not expressed in time for the proper patterning of the somite. The delay in myotome formation also suggests that myogenin, which is expressed earlier than MyoD, cannot substitute for MyfS or MyoD. Since MyfS is expressed in the epithelial somite before it compartmentalizes, it may play a direct role in the establishment of the sclerotomal lineage. Myf5-positive cells have been detected in the presumptive sclerotome of E9.5 forelimb somites (Smith et al., 1994). These results raise the possibility that the rib-deficient phenotype of MyfS-null mice is due to loss of Myf5-expressing cells within the sclerotome. Taken together with the results from the MyoD knockout, in which upregulation of MyfS occurs in the absence of MyoD, it appears that MyoD and Myf5 can functionally substitute for one another and influence each other's expression. When MyfS is absent, MyoD can compensate in myogenesis and vice versa. MyoD, however, is unable to provide the essential function of MyfS that influences sclerotome formation. It will be interesting to see if this function is specific for Myf5 or whether all mHLHs expressed in the epithelial somite can compensate for the loss of Myf5. Expression of other mHLHs under the control of Myf5 gene regulatory elements may resolve this issue. When the myogenin gene was disrupted in mice by homologous recombination, homozygous-null mice displayed a more severe muscle phenotype than had been observed in either the MyoD or Myfs mutants. Mice homozygous null for myogenin die perinatally like the Myfs mutants, but in contrast, they show a severe muscle deficiency. Two independent groups generated myogenin-null mice with similar results (Hasty et al., 1993; Nabeshima et al., 1993). Both studies provide dramatic evidence for an essential role of myogenin in muscle development. These mutant mice show a reduction in differentiated skeletal muscle throughout their bodies and die at birth, presumably due to an inability to respire. The inability to respire at birth is most likely a consequence of the absence of a functional diaphragm (Hasty et al., 1993). Although muscle is deficient in the myogenin-null mice, the muscle-forming regions are correctly formed and are occupied by mononucleate

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cells that stain positively with myoblast markers (Hasty et al., 1993). The myogenin-null mice have a defective rib cage but this defect is distinct from that observed in the My@ mutant. The defect is associated with the terminal portion of the ribs and the sternum, possibly as a consequence of the abnormal development of chest muscle. Although the myogenin-null mice are deficient in skeletal muscle they do possess sparse, apparently differentiated myofibers throughout their bodies. It is not clear whether these myofibers develop stochastically or because myogenin is not required for a particular myogenic lineage. MyoD mRNA and protein expression appears to be unaffected in the mutant mice (Hasty et al., 1993) and transgenics containing MyoD regulatory sequences driving a LacZ reporter are expressed normally in the myogenin-null background (Venuti et al., 1995). Cells isolated from the muscle-forming regions of mutant embryos differentiate in culture, confirming that normal determined myoblasts are present in mutant embryos (Nabeshima et al., 1993; Rawls et al., 1995). These results indicate that MyoD expression is independent of myogenin, and despite normal levels of expression, MyoD cannot functionally replace myogenin. Myoblasts are generated in myogenin mutants but fail to differentiate, supporting the hypothesis that myogenin plays a unique role in terminal differentiation of myoblasts into myotubes. Analysis of myogenin mutants has been extended in a detailed examination of muscle gene expression in the myogenin-null embryos during development (Venuti et al., 1995). It had been suggested that myotomal myoblasts are less affected by the absence of myogenin than migratory myoblasts (Nabeshima et al., 1993). This observation is most likely due to a delay in the differentiation of premuscle masses in the limbs of myogenin mutant embryos (Venuti et al., 1995). Surprisingly, muscle development throughout the body is close to normal (premuscle masses form and muscle differentiation markers are expressed) by the end of the embryonic stage (up to E15.5), but during fetal stages (E16.5 to birth) no further myogenic differentiation is observed in either axial or limb muscles. This suggests that muscle development is normal during primary myogenesis in myogenin mutants, but secondary myogenesis, which normally occurs during fetal stages and accounts for as much as 80% of the skeletal muscle in the neonate, is abnormal. Myogenesis is independent of myogenin during embryonic stages, but requires myogenin for the second wave of myogenic differentiation that occurs late in embryogenesis. Since MRF4 RNA is not expressed in myogenin mutant neonates it has been proposed that MRF4 requires myogenin for its expression. While this may be true for the second wave of MRF4 expression, which occurs during fetal stages beginning at E16, it has not been assessed for the early transient expression of MRF4 in the somite (Fig. 3). MRF4 protein is detected in the residual myofibers of myogenin mutants and these myofibers may have been specified during the early expression of MRF4 in the somite (Rawls et al., 1995). If myogenin and MRW function redundantly, as do MyoD and Myf5, the early expression of

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MRF4 during embryonic stages may be activated independently of myogenin and account for the differentiation of primary muscle fibers observed in myogenin mutant embryos. Conversely, failure of myogenin-dependent MRF4 expression during secondary myogenesis could account for the failure to generate secondary muscle fibers that normally occurs during fetal stages. This hypothesis remains to be tested through careful analysis of MRF4 expression in myogenin mutants and muscle formation in double knockouts of MRF4lmyogenin mutant mice. The late onset of stable h4RF4 expression during embryogenesis and in differentiating cells in culture suggests this mHLH plays a differentiation role in skeletal myogenesis, perhaps in stabilizing or maintaining the differentiated phenotype. The early somite expression (Fig. 3), however, suggests it may also play a role in early events of myogenesis. Several groups have generated mice with targeted mutations in the MRF4 gene with differing results (Braun and Arnold, 1995; Zhang et al., 1995; Patapoutian et al., 1995). The three MRF4 mutant lines have subtle differences most likely due to differences in the targeting constructs. In one MRF4 mutant line, MyfS RNA expression is severely affected and the phenotype is a phenocopy of the Myfs mutant; MyfS expression is reduced, they fail to form the distal portion of their ribs, and die at birth (Braun and Arnold, 1995). Since the two genes are located only 7 kb apart on the chromosome, cis elements important for Myfs gene regulation may have been altered during the generation of this mutant allele. A second mutant line shows a rib defect distinct from that observed in Myfs-null mice, does not have significantly altered MyfS expression, and homozygous mutants are viable (Zhang er al., 1995). Interestingly, myogenin mRNA expression is upregulated fivefold in the adult skeletal muscle of these MRF4null mice relative to wild-type expression, suggesting that myogenin can compensate for MRF4 in adult muscle and shares redundant functions with MRF4. The third mutant MRF4-null line demonstrates a phenotype that is intermediate to the other two lines in severity (Patapoutian et al., 1995). This line possesses rib defects similar to those observed by Zhang and co-workers, but in this MRF4null line rib defects are of sufficient severity to result in inviability. Defects in early myotome formation have also been reported for this MRF4-null line but this defect does not appear to alter subsequent myogenic differentiation. This supports the hypothesis that distinct combinations of mHLHs are responsible for discrete waves of myoblast differentiation and alterations in the expression patterns of the mHLH genes may influence patterning of other somite derivatives, particularly the sclerotome. Myotome differentiation has not been examined to the same extent in the other MRF4-null lines, so it is uncertain whether they exhibit the same myotomal alterations and if this is a common feature of MRF4-null mice. Although myogenin and MRF4 mutant mice display rib defects, these are less severe than that observed in Myfs mutants. Why these different mutations all affect rib differentiation is unclear, particularly since the defects observed are distinct. The observation that different subdomains within the early somite ex-

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press different mHLHs (Smith et al., 1994) suggests that different myogenic lineages are dependent on different combinations of the mHLHs. Alterations in the generation of these subdornains during somite differentiation may affect development of other somite derivatives, particularly the sclerotome. However, the question of exactly how early myogenic development influences the development of the axial skeleton remains unanswered. Combinations of null mutations in the mHLH genes are now being generated to unmask functions that may be obscured when genes share overlapping functions. The ability of Myf5 and MyoD to substitute functionally for one another was tested by interbreeding MyoD and M y f s mutant mice (Rudnicki et al., 1993). Mice deficient for both mHLHs die at birth. They possess the same skeletal defects as M y f i mutants and are totally devoid of skeletal myoblasts and muscle. This deficiency is based on lack of expression of myoblast and muscle differentiation markers, using both biochemical and histological criteria. Since there are few good early myoblast markers other than MyoD and MyfS themselves, it may be premature to state unequivocally that these mice are truly devoid of myoblasts. Myoblasts may be generated and fail to express the tested myoblast markers or fail to survive beyond a certain stage. Despite the failure of myogenic cells to occupy the muscle-forming regions in these embryos, the mice are remarkably normal in all other aspects of development, indicating that myogenesis is not essential for normal overall morphogenesis. The severity of the muscle defect in MyoDIMyf5 mutant mice depends on the M y f s and M y o D gene dosage. MyoD-null mice with a single Myfs allele generate normal muscle and ribs but are inviable. They contain only 50% of the normal complement of skeletal muscle. Mice with a single MyoD allele and no MyJS allele have normal muscle and were viable. This suggests that compensatory mechanisms may regulate the number of muscle cells formed during normal myogenesis. Since MyoD and Myf5 expression is confined to distinct domains within the somite (Smith et al., 1995), the redundancy between MyoD and MyfS implied by these “knockout” studies can be explained by cellular compensatory mechanisms. In this model, MyoD-expressing cells expand to compensate for the absence of Myf5-dependent cells in the MyfS knockouts and vice versa. MyoD is more efficient at compensating for the lack of Myf5 than MyfS is for MyoD. Downstream targets of these two genes may be distinct and/or these mHLHs may be essential for different myogenic lineages with different developmental potentials. Regardless of the mechanisms, these results suggest that either MyoD or Myf5 is required for the determination or survival of the myoblast lineage. The failure to generate myoblasts in the MyfsIMyoD double mutants argues that one of these genes must be present for myoblasts to form/survive and both mHLHs function in myoblast commitment and determination. Since myogenin-null mice possess myoblasts that cannot differentiate, myogenin must lie downstream of MyoD and MyfS in the myogenic pathway and play a role in myogenic differentiation. To investigate further the hypothesis that myogenin is downstream of MyoD

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and MyfS in the genetic hierarchy, or whether myogenin shares overlapping functions with MyoD and Myf5, myogeninlMyoD and myogeninlMyf5-null mice were generated by interbreeding the targeted mouse lines (Rawls et al., 1995). These mice show embryonic and perinatal phenotypes characteristic of the combined defects for mutation of each gene alone and with the number of undifferentiated myoblasts comparable to that seen in myogenin mutant mice. The function of myogenin in vivo does not overlap with those of either MyoD or Myf5. Results from the mHLH knockouts have clarified the hierarchical relationships and functional redundancies among the mHLHs. Either MyoD or MyfS is sufficient for formation or survival of skeletal myoblasts. Myogenin acts later in development and plays an essential role in terminal differentiation of myotubes. Since MRF4 has many features in common with myogenin, it probably acts similarly to myogenin as a differentiation factor although an additional role in early somitogenesis has been proposed (Patapoutian et al., 1995). Interbreeding the different mutant mouse lines confirms the redundant functions of MyoD and MyfS (Rudnicki et al., 1993) and demonstrates that the function of myogenin is restricted to the control of myoblast differentiation and does not overlap with the early functions of MyoD and Myf5 (Rawls et al., 1995). In the myogeninlMyoD and myogeninlMyf5 double knockouts, a subset of muscle fibers still differentiates in vivo and isolated myoblasts can differentiate in vitro, suggesting that two of the four mHLHs (MyoD and MRF4 or MyfS and MRF4) are sufficient to support myogenesis. An analysis of the evolutionary relationships between mHLHs predicts that MRF4 and myogenin may function in a redundant manner similarly to MyoD and MyfS (Atchley et al., 1994). This is in part based on the theory that the mHLH family of transcription factors was derived by gene duplication from a single ancestral gene. MyoD and MyfS arose from a common gene, suggesting they are more closely related and predicting functional redundancy that has, in fact, been substantiated (Rudnicki et al., 1993). Myogenin and M W 4 also arose from a common gene, which also predicts they may have redundant functions. Elevated expression of myogenin in MRF4 mutants (Zhang et al., 1995) supports this hypothesis, but confirmation of this prediction awaits interbreeding of these mutant mouse lines. The results from the mHLH mutational analysis in mice indicate that myogenesis proceeds when there are only two of the four possible mHLH factors present, but only when appropriately paired.

VI. Early Activation of Myogenic Program The specific inductive signals that initiate the myogenic program and specifically activate the mHLH genes have not been identified. Several approaches have been taken to identify the upstream events that activate the mHLH genes and subsequent myogenic differentiation. These include identification and characterization of signals from axial structures required for the development and differentiation

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of somites, examination of the differences between somitic myogenic lineages, and characterization of the cis elements in the mHLH gene regulatory regions that confer correct temporal and spatial expression. For the analysis of inductive cues, the chick embryo has been the experimental model of choice; somites and axial structures can be dissected and cultured in different combinations with relative ease. Analysis of gene regulatory elements relies on myogenic cell lines and on generation of transgenic mouse embryos to identify elements required for correct temporal and spatial expression during myogenic differentiation and development. Ultimately, these two approaches should coalesce to provide a greater understanding of the activation of the myogenic regulatory pathway.

A. instructive Cues from Axial Structures Manipulation of the somite and adjacent structures during somitogenesis and myogenesis indicates that axial structures, the notochord and neural tube, profoundly influence the differentiation of cell types derived from the somite. The somites lie directly adjacent to the axial structures and the first myogenic cells in the somite appear at the dorsomedial edge, closest to the neural tube (Fig. 2B). Removing both the notochord and neural tube leads to a loss of both sclerotomal and myotomal derivatives, but has no influence on differentiation of the limb musculature (Rong et al., 1992). This implies that axial structures influence the determination of the myotomal myoblasts but do not influence the migratory myoblasts. Since myotomal myoblasts arise from the medial somite and migratory myoblasts lineages that migrate away to colonize the limb arise from the lateral somite (Ordahl and Le Douarin, 1992) the axial structures are instructive for the medial half of the somite but do not influence the derivatives of the lateral somite. Implanting an extra notochord adjacent to a somite will cause its ventralization; dermamyotomal derivatives do not form and ectopic vertebrae are generated instead (Pourquie et al., 1993). Similarly, when the dorsal and ventral halves of the somite are exchanged at an early developmental stage, the cells in the transplanted halves will develop appropriately for their new position (Christ et al., 1992); the dorsal halves give rise to myotomal cells and the ventral halves give rise to sclerotomal cells. Cells within the ventral and dorsal compartments of the somite are also responding to inductive signals from the axial structures. Exactly how the axial structures exert their influence on somite myogenesis and/or migratory myoblasts is not well understood. To identify specific signals required for establishment of the myotomal lineage and the activation of mHLHs in the somite, several groups have analyzed the induction of myogenic differentiation in isolated somites by in vitro coculture experiments. These assays are a first step toward identification of the specific source and nature of inductive signals that influence myogenesis in somitic cells. Explants of mature somites cultured in vitro express muscle markers only

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when cocultured with cells from the adjacent neural tube (Kenny-Mobbs and Thorogood, 1987). Similarly, somitic cells from dissociated mouse somites generate more myogenic cells when cultured in the presence of neural tubes than without (Vivarelli and Cossu, 1986). Either the neural tube or the notochord can induce myogenesis in unspecified somites from early chick embryos (Stem and Hauschka, 1995; Buffinger and Stockdale, 1995). Neither segmentation of paraxial mesoderm into somites nor myogenic induction requires direct contact with the axial structures, suggesting diffusible factor(s) are involved. The exact role of the individual mHLHs during myogenesis in vivo is still unclear. Studies of early embryos show that the mHLHs are expressed in somites prior to overt signs of myogenic differentiation. It is not clear if signals from the axial structures are required for initial specification of the myogenic cells or for the differentiation of already determined muscle progenitors. Most studies have focused on the expression of muscle differentiation markers such as actin and MHC to assess the degree of myogenic induction in the cultured somites. To determine the source and nature of the inductive signals that specifically activate the expression of the mHLHs in somites, somites from different stage embryos were cocultured with different combinations of axial structures and the expression of mHLHs monitored (Munsterberg and Lassar, 1995). These experiments show that early myogenesis in somites requires interaction with both the neural tube and floor plate; more mature somites require only interaction with the neural tube in the absence of the floor plate. This supports a model in which two signals are necessary for somite myogenesis in v i m and interaction between the floor plate and neural tube mediates the competence of the somite to respond to a muscle-promoting signal from the neural tube (Munsterberg and Lassar, 1995).

B. Migratory versus Myotomal Myoblasts

Activation of mHLH gene expression is initiated very early in the presumptive myotomal cells (Ott et al., 1991; Pownall and Emerson, 1991; Sassoon et al., 1989); however, myogenic cells migrating from the dermamyotome into the developing limb and body wall do so without expressing the mHLH genes (Sassoon et al., 1989; Cheng et al., 1993; Yee and Rigby, 1993). It is not known if the migratory myoblasts derived from the lateral somite are committed myoblasts or whether their commitment takes place only after reaching their final destination. Axial structures are essential for the generation of the axial myotoma1 myoblast lineage but migratory myoblasts develop in the absence of influence from these structures (Rong et al., 1992). These results suggest that the axial myotomal and migratory myoblast lineages arise by different mechanisms. In situ hybridization and mHLH transgene analyses show the mHLHs are not expressed in the migratory myoblasts that colonize the limb and body wall until they reach their final destination. Since the migratory myoblasts are potentially

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determined as myogenic cells but do not express mHLHs at detectable levels it is not evident how they are specified. A mutant allele of the PAX3 gene in mice, splotch, was shown to produce major disruptions in limb muscle development but not in the muscle derived from the myotome (Franz, 1993; Franz et al., 1993). PAX genes encode a family of transcription factors that contain a paired box, a conserved motif found in several Drosophila pattern formation genes (Gruss and Walther, 1992). PAX genes are expressed in vertebrates in the developing nervous system, somitic mesoderm, and cells derived from somites. PAX3 is expressed in the paraxial mesoderm before, and in the dermamyotome and its myogenic derivatives after, segmentation (Goulding et al., 1991, 1994). Detailed analysis reveals PAX3 expression is higher in the lateral half of the dermamyotome than in the medial half (Williams and Ordahl, 1994) and is later confined to the caudal region and ventrolateral aspect of each somite (Bober et al., 1994). As somites mature, PAX3 expression specifically marks the migratory myoblast lineage that leaves the ventrolateral edge of the dermamyotome to colonize the limb (Williams and Ordahl, 1994; Bober et al., 1994). No other molecular marker defines these migratory myogenic cells. As development proceeds, PAX3 expression is repressed in the dermamyotome concomitant with activation of mHLH expression in myotomal cells (Williams and Ordahl, 1994), but PAX3 transcripts colocalize with Myf5 and MyoD transcripts in cells of the premuscle masses in the limb buds (Bober et al., 1994; Goulding et al., 1994). In splotch mutant mice, PAX3 transcripts are not detected in the region of migrating myoblasts or in the limb; these animals fail to develop limb musculature but not myotomally derived muscle (Bober et al., 1994). The loss of PAX3 expressing migratory cells appears to be the underlying cause for the limb muscle defect in splotch mutant mice. Whether PAX3 plays a role in the specification of the dermamyotomal cells that colonize the limb and body wall or whether it is required for their migration is unclear. Once these cells reach the limb PAX3 expression is downregulated and they upregulate their mHLHs in response to local signals. PAX3 may act upstream of the mHLH factors in the migratory myoblast lineage and prevent their activation or myogenic differentiation until the time is propitious (i.e., they have reached their final destination).

C. Analysis of Regulatory Elements in mHLH Genes

While much has been learned about the mechanisms through which the mHLH factors activate muscle-specific transcription, little is known about the regulatory circuits controlling expression of the mHLH regulators themselves. The temporal and spatial expression patterns of the individual mammalian mHLH genes are distinct during development (Fig. 3), suggesting numerous elements regulate their activation. The regulatory sequences that are required for the initial induction of the

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determination genes MyoD and MyfS and the trans factors involved in their correct spatiotemporal expression have not been identified. Transgenic lines have been generated using regulatory elements from these genes but all the essential cis elements and trans factors have not been defined. Since MyfS is the first of the mHLHs to be expressed in the developing mouse embryo, factors regulating the expression of this mHLH are likely to be involved in the initial activation of the myogenic program. Transgenic analysis of 5.5 kb of the 5' flanking sequence of My@ in transgenic mice showed this region can drive LacZ expression in the visceral arches and craniofacial muscle at early stages but somite staining is not detected until E l 2 (Patapoutian et al., 1993). MyfS mRNA is first detected in the visceral arches and somites at -E8, indicating this Myfs transgene responds to early MyfS activators in the visceral arches but not in somites. Myogenic determination in the nonsomitic head mesoderm is distinct from that of the somite and identification of the somitic-specific enhancer will require further analysis. An enhancer located approximately 18 kb upstream of the human MyoD gene has been shown to direct myotomal expression in transgenic mice (Goldhamer et al., 1992). Further dissection of this enhancer has identified a core enhancer of 258 bp with similarity to the mouse My00 enhancer (Goldhamer et al., 1995). Analysis of this enhancer in transgenic mice has revealed that MyoD expression is controlled by regulatory mechanisms distinct from other characterized musclespecific genes. E-boxes in the core enhancer are not required for MyoD activation. When MyoD-LacZ mice were bred with the myogenin-null mice to yield homozygous myogenin mutants carrying the MyoD transgene, normal patterns of MyoD-LacZ transgene expression were observed (Venuti et al., 1995). This suggests that myogenin is not required for either the activation or maintenance of MyoD. MyoD transgene expression does not show a strict rostral to caudal sequence of activation in the somites. Expression is detected in thoracic somites at the level of the forelimb bud first, then in more rostral somites approximately 12 hr later (Goldhamer et al., 1995). Spatial expression of the MyoD-LacZ transgene within the somites also exhibits a position-dependent pattern of expression; anterior to the forelimb bud transgene expression is prominent in the dorsal somite, posterior to the limb bud it is prominent more ventrally. This pattern of expression is reminiscent of the alterations of LacZ expression in transgenic mice observed when the regulatory elements of the myogenin gene are mutated (see below). Reexamination of MyoD transcript expression in developing embryos confirmed that the MyoD-LacZ expression pattern recapitulates the true endogenous pattern (Faerman et a l . , 1995). This nonuniform expression pattern suggests multiple myotomal cell populations exist and distinct sets of transcription factors operate in these cells to activate MyoD transcription. MRF4 gene regulation has also been analyzed in transgenic animals (Patapoutian et al., 1993). The sequences that regulate early myotomal activation of MRF4 have not been identified; MRF4-LacZ transgenic mice do not recapitulate

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the early transient expression of MRF4 in somites (Bober et al., 1991; Hinterberger et al., 1991). In adult skeletal muscles MRF4 is the predominant mHLH expressed (Rhodes and Konieczny, 1989), suggesting that MRF4 or non-mHLH factors must regulate MRF4 gene expression at this time. Analysis of the regulatory elements of the MRF4 gene in cell culture has shown that the MRF4 promoter is activated in nonmuscle cells by the other mHLHs, but not by its own gene product. A critical element in MRF4 gene regulation identified in cell culture is a MEF2 site. MEF2 and myogenin play a key role in activating the MRF4 gene in cell culture and do so in a synergistic manner (Naidu et al., 1995). MEF2- and mHLH-binding sites also have been identified in the mouse, chicken, and Xenopus MyoD genes, but these sites are utilized in distinct fashions. Mouse myogenin and Xenopus MyoD genes are MEF2 dependent but E-box independent (Buchberger et al., 1994; Cheng et al., 1993, 1995; Leibham et al., 1994). The chicken MyoD gene is independent of both E-boxes and MEF2binding sites (Dechesne et al., 1994). At present, the role of MEF2 or mHLHs in M y f s gene regulation is unknown. In contrast to the other mHLH genes the myogenin gene regulatory region is well defined. In cultured muscle cells, the elements regulating myogenin transcription include binding sites for the mHLHs and MEF2 (Edmondson et al., 1992). In mouse embryos, the myogenin promoter functions in an E-box- and MEF2-dependent fashion (Cheng et al., 1993; Yee and Rigby, 1993). Mutation of the E-box site has no effect on expression in the somites, but expression in the limb bud and visceral arches is delayed. This suggests that myogenin regulators operating through the E-box change both temporally and spatially. When the MEF2 site is mutated in the myogenin-LacZ transgene, limb bud expression is delayed and, strikingly, the pattern of transgene expression is altered in the somites. Mutation of the MEF2 site leads to a pattern of LacZ expression that is reminiscent of the expression seen with MyoD-Lac2 transgenes in cervical and thoracic somites. Expression is reduced in dorsal portions of thoracic somites and becomes more prominent in ventral portions of cervical somites. Mutation of both the E-box and MEF2 sites in the transgene results in complete loss of expression. Positive autoregulation of myogenin gene expression has been proposed as a mechanism of maintaining the myogenic program; however, a myogenin-LacZ transgene is expressed in the correct temporal and spatial pattern in myogenin-null mice. That myogenin is not required to initiate or maintain expression of the myogenin-LacZ transgene argues against a myogenin autoregulatory loop (Cheng et al., 1995). Differential activation of mutant myogenin-LacZ transgenes in somites and limb buds suggests positional regulation of myogenic regulators. MEF2 and E-box mutations in the myogenin promoter reflect steps in the myogenic determination process that differ in the two myogenic lineages. The apparent dependence on MEF2 appears paradoxical since ectopic expression of myogenin leads to activation of MEF2 (Cserjesi and Olson, 1991; Lassar et al., 1992). These

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results suggest that activation of myogenin gene transcription during myogenesis in vivo is mediated by an interplay between mHLH proteins and MEF2. The regulatory regions that are required for the initial induction of the determination genes MyoD and Myfs and the transcriptional factors involved in their correct spatiotemporal expression have not been fully identified. Analysis of the MRF4 and myogenin genes reveals a further level of complexity with the addition of a second family of muscle-specific transcription factors, the MEF2 family.

VII. MEF2 Family of Transcription Factors A. MEF2 Gene Family of Transcription Factors Another family of transcription factors that plays a role in the establishment of the myogenic lineage is the MEF2 (myocyte enhancer factor 2) family of MADS box-containing regulatory proteins (see Olson et al., 1995, for review). Although many skeletal muscle genes are activated by the mHLH family, others lack the DNA regulatory binding site for mHLHs, the E-box. The mHLH transcription factors do not account for all aspects of muscle gene regulation. It has been proposed that mHLHs act through intermediate myogenic regulators to activate muscle-specific genes and that regulatory factors outside the mHLH family also participate in myogenic gene regulation. MEF2 was originally characterized in myogenic cells as a muscle-specific DNA-binding activity that recognizes a conserved A/T-rich element [T(A/T), ATA(AG)] associated with numerous muscle specific genes, including the rat and human MCK enhancers, the rat myosin light-chain (mlc)-1/3 enhancer, and the chicken cardiac mlc-2A promoter (Gossett et al., 1989; Cserjesi et al., 1994). The DNA-binding activity of MEF2 is restricted to muscle cells of the skeletal, smooth, and cardiac lineages and to brain (Gossett et al., 1989; Horlick et al., 1990; Cserjesi and Olson, 1991; Yu et al., 1992; Leifer et al., 1993). MEF2 elements are found in many gene regulatory regions including those of the mHLHs (Edmondson et al., 1992; Wong et al., 1994; Leibham et al., 1994; Naidu et al., 1995). The importance of the MEF2 element for muscle-specific transcription is shown by mutational analysis at this site and the unique ability of tandem copies of MEF2 sites to direct muscle-specific transcription from nonmuscle basal promoters (Cheng et al., 1993; Edmondson et al., 1992; Gossett et al., 1989; Cserjesi et al., 1991; Nakatsuji et al., 1992; Yu et al., 1992). The MEF2 factors have now been cloned from a variety of organisms (Fig. 5). MEF2 is encoded by four human and mouse genes (MEF2A, MEF2B, MEF2C, and MEF2D) (Yu et al., 1992; Breitbart et al., 1993; Martin et al., 1993, 1994; P. Cserjesi, unpublished observations), two Xenopus genes (SLl and SL2) (Chambers et al., 1992; Wong et al., 1994), and single Drosophila (Lilly et al.,

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9.5

In cardiac muscle 11.5

13.5

MEF2A SGZ, RSRFC2

MEF2C

I

u-cardiac actin

MyfS

* MEFZB not detemned Fig. 5 (A) The structures of the different MEF2 family members and (B) temporal and spatial expression of MEF2 transcripts in the mouse embryo. (Compiled from Yu et al., 1992; Breitbart et al.. 1993; Chambers et a l . , 1992; Edmondson et a l . , 1994; Martin et al., 1992, 1994.)

1994; Nguyen et al., 1994) and C . elegans genes (Krause, 1995). MEF2 genes are members of the MADS box family of transcription factors originally termed RSRFs [related to serum response factor (SRF)], based on a shared sequence homology of greater than 80% within the 56-amino acid DNA-binding and dimerization domain of SRF known as the MADS box (Fig. 5). A 29-amino acid region adjacent to the MADS box, the MEF2 domain, is unique to the MEF2 subclass of MADS box proteins. The MADS designation arose from the initial family members, which included transcription factors in yeast (MCM- 1 , involved in mating type regulation), homeotic genes from plants (Agarnous and Deficiens), and SRF (Pollock and Treisman, 1991). Human serum response factor (SRF) is involved in coordinating transcription of the protooncogene c-fos. MCMl is central to the transcriptional control of cell type-specific genes and the pheromone response in yeast (see Shore and Sharrocks, 1995, for a review). The MEF2 proteins constitute a subclass of this family of transcription factors involved in muscle-specific gene regulation (Olson et al., 1995). While MEF2 genes exhibit many properties consistent with their encoding the proteins responsible for tissue-specific MEF2 DNA-binding activity, the transcripts for these genes are widely expressed (Breitbart et al., 1993; Yu et al., 1992; McDermott et al., 1993). Although MEF2 DNA-binding activity has been identified in nonmuscle, nonneural cell types (Pollock and Treisman, 1991; Chambers et al., 1992), immunocytochemical analyses have revealed a strict correlation between MEF2 protein and MEF2 DNA-binding activity in a cell type-restricted distribution, implicating posttranscriptional mechanisms in the

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regulation of MEF2 expression (Yu et al., 1992; Breitbart et al., 1993). Alternative splicing may be important in this process: two alternative MEF2D domains, at least one of which is specifically included during myogenic differentiation, correlate precisely with endogenous MEF2 activity (Fig. 5). Since MEF2 proteins bind DNA as dimers, combinatorial interactions among the different MEF2 proteins and alternative splicing of MEF2 gene products suggest a broad regulatory potential for these proteins. Alternatively spliced variants suggest posttranscriptional regulation plays a role in tissue-specific MEF2 activity (Yu et al., 1992; McDermott et al., 1993).

B. Developmental Expression of MEF2s Analysis of MEF2 gene expression during embryogenesis in a variety of organism confirms a role in muscle development. MEF2s are initially expressed in the early developing cardiac tissue and subsequently in the developing somites in the mouse embryo (Edmondson et al., 1994). MEF2C expression is first detected at E7.5 in the cardiac mesoderm, making MEF2C one of the earliest markers for the cardiac muscle lineage (Fig. 5). Later, MEF2A and MEF2D mRNAs are also detected in the myocardium (Fig. 5). By E9.0, MEF2C is expressed in myotomes, where its expression lags behind the mHLHs MyfS and myogenin (Fig. 5). MEF2A and MEF2D are expressed at a lower level than MEF2C in the myotome at E9.5 and are more broadly expressed throughout the embryo. Expression of all the MEF2 genes is observed in muscle-forming regions within the limbs at E l 1.5 and within muscle fibers throughout the embryo at later developmental stages. After E12.5, MEF2 transcripts are detected at high levels in specific regions of the brain and ultimately in a wide range of tissues. MEF2 gene expression is also observed in smooth muscle before the onset of smooth muscle structural gene expression. In the brain, MEF2A, B, and D show highly localized expression patterns that coincide with gradients of neuronal differentiation (Leifer et al., 1993; Lyons et al., 1995). MEF2C is restricted to skeletal muscle, brain, and spleen in adults. Early expression of MEF2 in the cardiac lineage suggests a potential role in cardiac muscle determination, but in the skeletal muscle lineage, MEF2 expression is initiated after that of MyfS and myogenin. This post-mHLH expression suggests the MEFs are unlikely to play a determination role in skeletal myogenesis; however, since the regulatory regions of mHLHs contain MEF2 sites, the MEF2s may function in a positive-feedback loop to amplify and maintain mHLH expression. It is not clear whether expression of MEF2s is dependent on mHLHs. Normal expression of myogenin-lac2 transgenes in myogenin mutant embryos would argue that activation of MEM expression is not dependent on myogenin (Cheng et al., 1995). In Drosophila a single MEF2 gene, D-MEF2, has been identified. It is first expressed in the presumptive mesodem at the late cellular blastoderm stage and

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continues to be expressed in all mesoderm after invagination (Lilly et al., 1994; Nguyen et al., 1994). Following the dorsal migration of the mesoderm, D-MEF2 expression becomes restricted to the primordia for visceral, cardiac, and somatic muscles. MEF2 expression in Drosophila precedes the expression of the Drusophila mHLH nautilus, implicating D-MEF2 in the upstream regulation of myogenesis in the fly. D-MEF2 functions in early mesoderm differentiation and is required for subsequent cell fate specifications within the different muscle lineages (see below). The Xenupus MEF2 homologs (XMEF2s) accumulate preferentially in forming somites after the appearance of XMyoD transcripts (Chambers et al., 1992; Wong et al., 1994) and are important regulators of XMyoD transcription (Wong et al., 1994; Leibham et al., 1994). A C. elegans MEF2 homolog has also been cloned but it has not been fully characterized (Krause, 1995). The identification of MEF2 family members in these diverse organisms suggests they form a conserved alternative pathway for myogenic gene regulation and perhaps myogenic lineage specification. Because of the multiplicity of MEF2 genes, analysis of the function of the MEF2 factors in vertebrates is likely to be complex. Interbreeding of mice with targeted mutations for the individual MEF2 family members may be required before the hierarchy of regulation can be determined. However, confirmation of the role of MEF2s in myogenesis has come from loss-of-function mutations in the single MEF2 gene of Drosophila, D-MEF2 (Lilly et al., 1995; Bour et al., 1995). Analysis of myoblast-specific markers indicates early specification and differentiation of somatic muscle precursors are not affected in MEF2 mutant flies. However, in the absence of MEF2 these cells are unable to undergo further differentiation to form muscle fibers. In the D-MEF2 mutant embryos, somatic, cardiac, and visceral muscle cells do not differentiate, but myoblasts are normally specified and positioned. D-MEF2 is required for later aspects of differentiation of the three major types of musculature in the Drosophila embryo and these different muscle types share a common myogenic differentiation program controlled by D-MEF2.

C. MEF2s as Regulators of Myogenesis

MEF2 DNA-binding activity is upregulated during the differentiation of established muscle cell lines and can be induced in nonmuscle cells by MyoD and myogenin, suggesting that it lies downstream of the mHLHs (Cserjesi and Olson, 1991; Lassar et al., 1991; Cserjesi et al., 1994). However, several mHLH genes possess essential MEF2-binding sites in their regulatory regions, suggesting MEF2s may function as regulators of the mHLHs. MEF2 sites in the myogenin promoter (Edmondson et al., 1992), MRF4 promoter (Naidu et al., 1995), and MyoD promoter (Wong et a l . , 1994) are shown to be essential for the transcription of these mHLH genes. Mutation of the MEF2 site in transgenes (containing

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the myogenin regulatory region upstream of a LacZ reporter) alters tissue-specific expression during mouse embryonic development (Cheng et al., 1993; Yee and Rigby, 1993). The MEF2 site and E-box in the MRF4 gene function in concert to activate MRF4 gene transcription synergistically in nonmuscle cells coexpressing MEF2 and myogenin proteins (Naidu et al., 1995). The MEF2binding site but not the E-box is necessary for cell type-specific expression and activation of the myogenin gene by MyoD in tissue culture cells (Buchberger et al., 1994). This activation of myogenin is induced in the presence of protein synthesis inhibitors, suggesting myogenin expression requires preexisting MEF2 and a posttranslational mechanism. A regulatory circuit involving direct and indirect feedback loops has been proposed to explain the interactions between the MEF2s and myogenin in myogenin gene regulation (Edmondson et al., 1992). Gene-specific activation by the different MEF2s may depend on combinatorial interactions with other transcriptional regulators. MADS box proteins have the potential to interact with other regulatory proteins to control gene transcription (Shore and Sharrock, 1995). For example, SRFs specifically interact with homeobox-containing genes of the paired family (Pollock and Treisman, 1991). Protein complexes that increase the affinity of myogenin for DNA contain MEF2. Myogenin DNA-binding site selection using cyclic amplification and selection of target sequences (CASTing) with C2C 12 nuclear extracts identifies a predominance of sequences with myogenin sites adjacent to MEF2 consensus sites (Wright et al., 1991; Funk and Wright, 1992). These results suggest an intimate relationship between the MEF2s and mHLHs and that these two families of factors bind cooperatively to regulatory elements in the skeletal muscle genes they activate. Myogenic conversion of nonmuscle cells by MEFs and cooperative interaction between MyoD and MEF2A to activate myogenesis in nonmuscle cells has been reported (Kaushal et al., 1994). Cooperative interaction between MEF2A and MyoD is sequence specific. It requires the MADS domain of MEF2 and the basic domain of MyoD. The bHLH of mHLHs can discriminate between the MADS domain of MEF2s and the MADS domain of SRF. The ability to convert cells to the myogenic phenotype was previously thought to be unique to the mHLHs. This feature is now shared by the MEF2 family and strengthens the hypothesis that positive feedback loops involving cooperative interactions between members of the MEF2 and mHLH gene families reinforce myogenic gene activation and differentiation.

VIII. Summary and Conclusions During myogenesis, mHLH and MEF2 proteins cooperatively activate skeletal muscle genes and physically interact through the basic domain and MADS domain, respectively. Association between MEF2s and mHLHs provides specificity

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for activation of skeletal muscle genes. Thus, skeletal myogenesis is mediated by two distinct families of mutually inducible and interactive muscle transcription factors, either of which can initiate the developmental cascade. The complexity of the mHLH and MEF2 families of regulatory proteins provides the potential for fine-tuning of transcriptional responses as a consequence of combinatonal interactions among multiple isofoms encoded by the four MEF2 genes and the different mHLHs. MEF2 genes may similarly participate in the differentiation pathways of cardiac muscle, smooth muscle, neural, and other cells by inducing the expression of and/or directly associating with cell-specific transcriptional regulators in these lineages. Analysis of the temporal and spatial expression patterns of these muscle regulatory factors during early somitogenesis has provided a new view of muscle cell fate determination, differentiation, and gene expression. Identification of the mHLH and MEF2 families and characterization of their roles in myogenic determination and differentiation have provided not only an understanding of the molecular events of muscle cell fate determination but also a framework to understand determination and differentiation in general. The molecular genetics and embryological approaches employed to unravel the hierarchical interactions of the mHLH gene family in particular, have led to a coalescence between classic embryology and modern molecular biology.

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Miner, J. H., and Wold, B. (1990). Herculin, a fourth member of the MyoD family of myogenic regulatory genes. Proc. Natl. Acad. Sci. U.S.A. 87, 1089-1093. Montarras, D., Chelly, J., Bober, E., Arnold, H. H., Ott, M.-o., Gros, F., and Pinset, C. (1991). Developmental patterns in the expression of Myf-5, MyoD, myogenin, and MRF4 during myogenesis. New Biol. 3, 592-600. Miinsterberg, A., and Lassar, A. B. (1995). Combinatorial signals from the neural tube, floor plate and notochord induce myogenic bHLH gene expression in the somite. Development 121, 651-660. Murre, C., Bain, G., VanDijk, M. A,, Engel, I., Furnari, B. A,, Massari, M. E., Matthews, J. R., Quong, M. W., Rivera, R. R., and Stuvier, M. H. (1994). Structure and function of helix-loop-helix proteins. Biochim. Biophys. Acta Gene Struct. Express. 1218, 129-135. Nabeshima, Y., Hanaoka, K., Hayasaka, M., Esumi, E., Li, S., Nonaka, I., and Nabeshima, Y. (1993). Myogenin gene disruption results in perinatal lethality because of a severe muscle defect. Nature (London) 364,532-535. Naidu, P. S., Ludolph. D. C., To, R. Q., Hinterberger, T. J., and Konieczny, S. F. (1995). Myogenin and MEF2 function synergistically to activate the MRF4 promoter during myogenesis. Mol. Cell. Biol. 15, 2707-2718. Nakatsuji, Y., Hidaka, K., Tsujino, S . , Yamamoto, Y., Mukai, T., Yanagihara, T., Kishimoto, T., and Sakoda, S. (1992). A single MEF-2 site is a major positive regulatory element required for transcription of the muscle specific subunit of the human phosphoglycerate mutase gene in skeletal and cardiac muscle cells. Mol. Cell. Biol. 12, 4384-4390. Nguyen, H. T., Bodmer, R., Abmayr, S. M., McDermott, J. C., and Spoerel, N. A. (1994). D-mef2: A Drosophila mesoderm-specific MADS box-containing gene with a biphasic expression profile during embryogenesis. Proc. Natl. Acad. Sci. U.S.A. 91, 7520-7524. Olson, E. N., and Klein, W. H. (1994). hHLH factors in muscle development: Deadlines and commitments, what to leave in and what to leave out. Genes Dev. 8, 1-8. Olson, E. N., Perry, M., and Schulz, R. A. (1995). Regulation of muscle differentiation by the MEF2 family of MADS box transcription factors. Dev. Biol. 172, 2-14. Ordahl, C. P., and Le Douarin, N. M. (1992). Two myogenic lineages within the developing somite. Development 114, 339-353. Ott, M.-O., Bober, E., Lyons, G . , Arnold, H. H., and Buckingham, M. (199 1). Early expression of the myogenic regulatory gene, Myf-5, in precursor cells of skeletal muscle in the mouse embryo. Development 111, 1097-1 107. Patapoutian, A,, Miner, J. H.,Lyons, G. E., and Wold, B. (1993). Isolated sequences from the linked Myf-5 and MRF4 genes drive distinct patterns of muscle-specific expression in transgenic mice. Developrnenf 118, 61-69. Patapoutian, A , , Yoon, J. K., Miner, J. H., Wang, S . , Stark, K., and Wold, B. (1995). Disruption of the mouse MRF4 gene identifies multiple waves of myogenesis in the myotome. Development 121, 3347-3358. Paterson, B. M., Waldorf, U., Eldridge, J., Dubendorfer, A,, Frasch, M., and Gehring, W. J. (1991). The Drosophila homologue of vertebrate myogenic determination genes encodes a transiently expressed nuclear protein marking primary neurogenic cells. Proc. Natl. Acad. Sci. U.S.A. 88, 3782-3786. Pollock, R., and Treisman, R. (1991). Human SRF-related proteins: DNA-binding properties and potential regulatory targets. Genes Dev. 5 , 2327-2341. Pourquie, O., Coltey, M., Teillet, M. A , , Ordahl, C. P., and Le Dourain, N. M. (1993). Control of dorsoventral patterning of somitic derivatives by notochord and floor plate. Proc. Natl. Acad. Sci. U.S.A. 90, 5242-5246. Pownall, M. E., and Emerson, C. P. (1991). Sequential activation of three myogenic regulatory genes during somite morphogenesis in quail embryos. Dev. Biol. 151, 67-79. Rawls, A,, Moms, J. H., Rudnicki, M. A,, Arnold, H. H., Klein, W. H., and Olson, E. N.

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(1995). Myogenin’s functions do not overlap with those of MyoD or Myf5 during mouse myogenesis. Dev. Biol. 172, 37-50. Rhodes, S. J., and Konieczny, S . F. (1989). Identification of MRF4: A new member of the muscle regulatory gene family. Genes Dev. 4, 2050-2061. Rong, P. M., Teillet, M.-A,, Ziller, C., and Le Dourain, N. M. (1992). The neural tubeinotochord complex is necessary for vertebral but not limb and body wall striated muscle development. Development 115, 657-672. Rudnicki, M. A., Braun, T., Hinuma, S . , and Jaenisch, R. (1992). Inactivation of MyoD in mice leads to up-regulation of the myogenic HLH gene Myf-5 and results in apparently normal muscle development. Cell 71, 383-390. Rudnicki, M. A,, Schnegelsberg, P. N. J., Stead, R. H., Braun, T., Arnold, H. H., and Jaenisch, R. (1993). MyoD or Myf5 is required for the formation of skeletal muscle. Cell 75, 1351-1359. Rudnicki, M. A,, and Jaenisch, R. (1995). The MyoD family of transcription factors and skeletal myogenesis. Bioessays 17, 203-209. Rupp, R. A., Snider, L., and Weintraub, H. (1994). Xenopus embryos regulate the nuclear localization of XMyoD. Genes Dev. 8, 1311-1323. Sassoon, D., Lyons, G., Wright, W. E., Lin, V., Lassar, A., Weintraub, H., and Buckingham, M. (1989). Expression of two myogenic regulatory factors myogenin and MyoD during mouse embryogenesis. Nature (London) 344,303-307. Schwarz, J. J., Chakraborty, T., Martin, J., Zhou, J. M., and Olson, E. N. (1992). The basic region of myogenin cooperates with two transcription activation domains to induce muscle-specific transcription. Mol. Cell. Biol. 12, 266-275. Shafer, 8. W., Blakeley, B. T., Darlington, G. J., and Blau, H. M. (1990). Effect of cell history on response to helix-loop-helix family of myogenic regulators. Nature (London) 344, 454458. Shore, P., and Sharrocks, D. (1995). The MADS-box family of transcription factors. Eur. J . Biochem. 229, 1-13. Smith, T. H., Block, N. E., Rhodes, S. J., Konieczny, S. F., and Miller, J. B. (1993). A unique pattern of expression of the four muscle regulatory proteins, distinguishes somitic from embryonic, fetal, and newborn mouse myogenic cells. Development 117, 1125-1133. Smith, T. H., Kachinsky, A. M., and Miller, J. B. (1994). Somite subdomains, muscle origins, and the four muscle regulatory factor proteins. J. Cell Biol. 127, 95-105. Stem, H. M., and Hauschka, S . D. (1995). Neural tube and notochord promote in vitro myogenesis in single somite explants. Dev. Biol. 167, 87-103. Sunyer, T., and Merlie, J. P. (1993). Cell type- and differentiation-dependent expression from the mouse acetylcholine receptor epsilon-subunit promoter. J. Neurol. Res. 36, 224-234. Taylor, S. M., and Jones, P. A. (1979). Multiple new phenotypes induced in 10T112 and 3T3 cells treated with 5-azacytidine. Cell 17, 771-779. Thayer, M. J., Tapscott, S. J., Davis, R. L., Wright, W. E . , Lassar, A. B . , and Weintraub, H. (1989). Positive autoregulation of the myogenic determination gene, MyoD. Cell 12, 49945003. Vaidya, T. B., Rhodes, S. J . , Taparowsky, E. J., and Konieczny, S. F. (1989). Fibroblast growth factor and transforming growth factor B repress transcription of the myogenic regulatory gene MyoD1. Mol. Cell. Biol. 9 , 3576-3579. Venuti, J. M., Goldberg, L., Chakraborty, T., Olson, E. N., and Klein, W. H. (1991). A myogenic factor from sea urchin embryos capable of programming muscle differentiation in mammalian cells. Proc. Narl. Acad. Sci. U.S.A. 88, 6219-6223. Venuti, J. M., Kozlowski, M. T., Gan,L., and Klein, W. H. (1993). Developmental potential of muscle cell progenitors and the myogenic factor SUM-1 in the sea urchin embryo. Mech. Dev. 41, 3-14.

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Venuti, I. M., Moms, J., Vivian, J., Olson, E. N., and Klein, W. H. (1995). Myogenin is required for late but not early aspects of myogenesis during mouse development. J. Cell Biol. 128, 563-576. Vivarelli, E., and Cossu, G. (1986). Neural control of early myogenic differentiation in cultures of mouse somites. Dev. Biol. 117, 319-325. Wachtler, F., and Christ, B. (1992). The basic embryology of skeletal muscle: The avian model. Semin. Dev. Biol. 3, 217-227. Wang, Y., Benezra, R., and Sassoon, D. A. (1992). Id expression during mouse development: A role in morphogenesis. Dev. Dynam. 94, 222-230. Weintraub, H. (1993). The MyoD family and myogenesis: Redundancy, networks, and thresholds. Cell 75, 1241-1244. Weintraub, H., Tapscott, S. J., Davis, R. L., and Lassar, A. (1989). Activation of muscle-specific genes in pigment, nerve, fat, liver, and fibroblast cell lines by forced expression of MyoD. Proc. Natl. Acad. Sci. U.S.A. 86, 5434-5438. Weintraub, H., Dwarki, V. J., Verma, I., Davis, R., Hollenberg, S., Snider, L., Lassar, A,, and Tapscott, S. .I. (1991). Muscle-specific transcriptional activation by MyoD. Genes Dev. 5 , 1377- 1386. Wentworth, B., Donoghue, M. J., Engert, J., Berglund, E., and Rosenthal, N. (1991). Paired MyoD binding sites regulate myosin light chain gene expression. Proc. Narl. Acad. Sci. U.S.A. 88, 1242-1246. Williams, B. A,, and Ordahl, C. P. (1994). Pax-3 expression in segmental mesoderm marks early stages in myogenic specification. Developmenr 120, 785-796. Wright, W. E., Sassoon, D. A,, and Lin, V. K. (1989). Myogenin, a factor regulating myogenesis, has a domain homologous to MyoD. Cell 56, 607-617. Wright, W. E., Binder, M., and Funk, W. (1991). Cyclic amplification and selection of targets (CASTing) for the myogenin consensus binding site. Mol. Cell. Biol. 11, 4104-4110. Wong, M. W., Pisegna, M., Lu, M. F., Leibham, D., and Perry, M. (1994). Activation of Xenopus MyoD transcription by members of the MEF2 protein family. Dev. Biol. 166, 683-695. Yaffe, D. (1968). Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc. Natl. Acad. Sci. U.S.A. 61, 477-483. Yaffe, D., and Saxel, 0. (1977). Serial passaging and differentiation of myogenic cells isolated from dystrophic mouse muscle. Nature (London) 270, 725-727. Yee, S.-P., and Rigby, W. (1993). The regulation of myogenin gene expression during the embryonic development of the mouse. Genes Dev. 7, 1277-1289. Yu, Y.-T., Breitbart, R. E., Smoot, L. B., Lee, Y., Mahdavi, V., and Nadal Ginard, B. (1992). Human myocyte specific enhancer factor 2 (MEF2) comprises a group of tissue restricted MADS box transcription factors. Genes Dev. 6, 1783-1798. Yutzey, K. E., Rhodes, S. J., and Konieczny, S. F. (1990). Differential transactivation associated with the muscle regulatory factors MyoD1, myogenin, and MRF4. Mol. Cell. Biol. 10, 39343944. Zhang, W., Behringer, R., and Olson, E. N. (1995). Inactivation of the myogenic bHLH gene MRF4 results in up regulation of myogenin and rib anomalies. Genes Dev. 9, 1388-1399.

6 Developmental Programs in Bacteria Richard C. Roberts, Christian D. Mohr, and Lucy Shapiro Department of Developmental Biology Stanford University School of Medicine Stanford, California 94305

I. Introduction: The Concept of Development among Bacteria 11. Some Examples of Development among Bacteria A. Differentiation Events Occurring as Part of the Normal Cell Cycle in Cuulobncter crescentus

B. Differentiation Events Occurring in Response to External Stimuli 111. Control of Cellular Differentiation during the Caulobucter crescentus Cell Cycle A. Temporal and Spatial Control of Chromosome Replication

B. Control of the Timing and Subcellular Location of Transcription C. Targeting of Proteins to Specific Locations within the Cell D. Specific Degradation of Proteins at Discrete Times during the Cell Cycle IV. Conclusions and Future Perspectives References

1. Introduction: The Concept of Development among Bacteria A common misconception about the world of bacteria is that, in contrast to the developmental complexities seen for multicellular eukaryotic organisms, their life cycles are very simple; a ceaseless repetition of growth and binary fission with little variation and no communication between the individual, autonomous cells. While this does typify the growth properties of many bacterial species multiplying exponentially on abundant nutrient sources, this is a condition rarely encountered in nature. Many bacteria respond to the less than optimal growth conditions that are prevalent by initiating alternative regulatory mechanisms for gene expression and growth control. Still others respond to environmental stress by initiating alternative developmental cycles, often quite different from the standard exponential-phase growth. Further, some bacteria progress through simple developmental cycles as an integral part of each cell cycle. These processes of development, whereby one cell type is morphologically transformed into another cell type, abound among different prokaryotic species in their natural environments, with a variety of intracellular mechanisms responsible for altering the cellular architecture. In some instances, the formation of complex multicellular structures accompanies the developmental cycle, whereas others Current Topics 8n Developmental Biology, Vol 34 Copyright Q 1996 by Academic Press, Inc. All rights of reproduction in any form reserved

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are fundamentally cell autonomous. In this chapter we introduce some of the better-studied examples of developmental cycles among bacteria, and then focus on the mechanisms that regulate these cycles. Much of our discussion will be on Cuulobucter crescentus (Fig. l), as an example of a bacterium with an extensively studied developmental cycle that is independent of environmental stress. The ease of synchronization and amenability to genetic analysis of C . crescentus have allowed significant strides to be made in understanding the forces that drive its obligate developmental cycle.

Fig. 1 The bacterium C . crescentus. (A) A predivisional cell, with the stalked pole at the top and the flagellated pole at the bottom. The single polar flagellum, normally many times the length of the cell, has been sheared in sample preparation. (B) The two types of daughter cells produced by division of a predivisional cell: a stalked cell (top) and a flagellated swanner cell (bottom). The electron micrographs, courtesy of J. Maddock (University of Michigan, Ann Arbor, MI), were taken with a Philips 300 transmission electron microscope, using whole-mounted samples of C. crescentus on carbon-coated grids stained with uranyl acetate.

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It. Some Examples of Development among Bacteria A. Differentiation Events Occurring as Part of the Normal Cell Cycle in Caulobacter crescentus

The cell cycle of C. crescentus is shown in Fig. 2 . This bacterium is found in nutrient-poor aquatic environments, both marine and freshwater (reviewed in Poindexter, 1964). To facilitate dispersal and location of nutrient sources, C. crescentus is motile and chemotactically competent during one phase (the

Rg. 2 The cell cycle of C. crescentus. The swarmer cell possesses a single polar flagellum, several polar pili, and is unable to initiate chromosomal replication. During the swarmer to stalked cell transition, the flagellum is shed, and a stalk is synthesized in its place, pili are lost, and chemoreceptors are degraded. DNA replication initiates in the stalked cell, as indicated by the theta structure within the cell. A transcriptional regulatory cascade is activated in the stalked cell that culminates in the synthesis and assembly of a new polar flagellum. The asymmetric predivisional cell has distinct polar morphologies, compartment-specific proteins, and differential chromosomal transcription. Cell division produces two distinct cell types, a replication-competent stalked cell and a motile swarmer cell with a condensed nonreplicating chromosome. The progeny stalked cell behaves like a stem cell, immediately initiating another round of cell division, while the progeny swanner cell delays cell division until it differentiates into a stalked cell.

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swarmer phase) of its life cycle, and is propelled toward nutrient sources by a single, polar flagellum. At the same pole occupied by the flagellum are clustered the chemoreceptors (Alley et al., 1992), pili (Lagenaur and Agabian, 1977), and receptors for several bacteriophages (Shapiro et al., 1971). The swarmer cell is metabolically somewhat quiescent, with replication of its single circular chromosome blocked; even introduced broad-host-rangeplasmids replicate at a reduced rate (Marczynski and Shapiro, 1992). After approximately one-third of the cell cycle (independent of the length of the cycle), the polarly localized chemotaxis machinery, flagellum, and pili are lost, and in their place a stalk is synthesized. The stalk and the associated holdfast material located at its tip allow the bacterium to adhere strongly to a surface; the stalk has also been proposed as a structure that increases the surface area of the bacterium, thereby facilitating nutrient uptake in dilute environments. Differentiation into a stalked cell involves a number of physiological changes, including specific proteolysis of polar components and the initiation of chromosomal DNA replication. These events are accompanied by activating expression of a number of genes, the best studied being the cascade that is ultimately responsible for the synthesis of a new flagellum later in the cell cycle (reviewed in Brun et al., 1994; Gober and Marques, 1995; Stephens et al., 1995b; Jenal et al., 1995). During stalked cell growth, the pole opposite the one bearing the stalk differentiates into a new flagellated pole, with its associated chemotaxis protein complexes, pili, and phage receptors. Prior to the time of cell division, the two halves of the predivisional cell are quite distinct, both morphologically and in their pattern of gene expression. The newly replicated chromosome in the stalked compartment of the predivisional cell is able to initiate DNA replication, whereas the chromosome in the swarmer compartment is not. At cell division, the new swarmer cell disperses to another location, while the stalked cell remains behind to act as a stem cell for subsequent cell cycles. This developmental cycle of C . crescentus is an obligate part of its life cycle. Two major events in the C . crescentus cell cycle have been the focus of research aimed at elucidating the fundamental processes required for development. These include the tight regulation of chromosomal replication and the temporally and spatially regulated biogenesis of the flagellum. As is discussed below, these two processes appear to be linked by a common transcriptional regulator that orchestrates the response of multiple cellular processes to the progression of the cell cycle (Quon et al., 1996). A hallmark of C . crescentus development is that the timing of these events is regulated predominantly by internal signals as an intrinsic part of each cell cycle.

B. Differentiation Events Occurring in Response to External Stimuli

In contrast to the obligate cell cycle of C . crescentus, a large number of bacteria enter developmental programs in response to external stimuli. The most common

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inducer of development is nutritional limitation, which elicits survival responses in a number of bacteria. The nature of the response varies substantially, from the production of endospores by individual Bacillus subtilis cells, to the formation of a multicellular fruiting body and associated spores by Myxococcus xanthus, to the pathway of differentiation of Streptomyces coelicolor vegetative hyphae into spore-bearing hyphae. Other bacteria enter developmental pathways as part of symbiotic relationships to maximize nutrient acquisition; the association of Rhizobium species with leguminous plants being a prime example. Furthermore, non-spore-forming bacteria also respond to starvation conditions by inducing distinct physiological events, perhaps best studied in Escherichia coli on its entry into stationary phase. Each of these developmental processes is briefly addressed in the next sections. The reader is referred to the cited reviews for more in-depth discussions of these different systems.

1. Endospore Formation in Bacillus subtilis upon Starvation When the gram-positive bacterium B . subtilis finds itself in conditions of high cell density and/or starvation, it undergoes processes of differentiation that manifest themselves in several ways. Several different and overlapping regulatory responses are involved, controlling genes for the establishment of a state of competence for DNA uptake (Dubnau, 1993; Grossman, 1995), those involved in the production of peptide antibiotics (Marahiel et al., 1993), and genes whose products synthesize extracellular degradative enzymes used for enhanced liberation of nutrients (Doi, 1991). An additional pathway leads to the terminal differentiation into endospores, diagrammed in Fig. 3 (reviewed in Errington, 1993; Grossman, 1995). This bacterium and its relatives, important not only in the study of processes of bacterial development, but also as sources of industrially important metabolites (Harwood, 1992), have been studied extensively and much is known about the complex networks regulating the developmental processes. As one manner of increasing the genetic diversity (which may enhance the long-term survival of the organisms), B . subtilis at high cell density induces a system to actively take up exogenous DNA, a state termed competence. There appear to be two signals that mediate development of competence, both of which are proposed to report conditions of high cell density, similar to autoinducer sensing in Vibrio (Fuqua et al., 1994). However, both signal molecules appear to be short peptides, rather than the homoserine lactone used by Vibrio. The first signal to be identified corresponds to the last 9-10 amino acids of the comX gene product (Magnuson et al., 1994). Its production is dependent on both c o r d and c o m e ; after synthesis it is exported out of the cell. When this peptide reaches sufficiently high extracellular concentrations (thus reflecting high cell density), it activates the membrane-bound sensor kinase ComP, which phosphorylates the cognate response regulator ComA (reviewed in Dubnau et al., 1994). ComA then activates transcription of other genes involved in a variety of responses, indirectly elevating expression of the comK gene. The comK gene product is

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4

a

E9

Replication completed

a

Asymmetric septation

Germination

Spomlation

I t

a

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8 Mother cell lysis

tI Spore maturation

Forespore engulfment

Fig. 3 Vegetative and sporulation cell cycles of B . subtilis. Top: Reproduction by binary fission of this bacterium, with each cell cycle producing two progeny cells, as indicated by the “(X 2)” notation. Conditions of stress, including starvation (indicated) or accumulation of toxic metabolites, trigger exit from this vegetative cycle to enter the developmental sporulation pathway (bottom). Asymmetric cell division follows a round of DNA replication. After division, the smaller forespore Compartment is engulfed by the larger mother cell compartment. Communicating signals between the two compartments result in differential gene expression, culminating in the maturation of the forespore into an environmentally resistant spore and release by lysis of the mother cell. When this spore encounters favorable nutrient conditions, it germinates to produce a cell that once again enters the vegetative cycle.

another transcription factor that activates synthesis of the machinery responsible for extracellular DNA uptake (van Sinderen et al., 1995).Phosphorylation of the ComA response regulator may also be induced through the sensing of a second extracellular signal peptide, called competence-stimulatingfactor (Solomon et al., 1995). This factor, also a small peptide, accumulates during growth to reach higher levels in late log-phase B . subtilis cultures (Solomon et al., 1995). These conditions, including the presence of the alternate sigma factor UH and gene

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products involved in the SpoOFf SpoOBJSpoOA phosphorelay pathway, play a role in establishing either competence or sporulation. Once exported, accumulating competence-stimulating factor attains sufficient extracellular concentrations for signaling; it is then thought to be transported into the cell through the peptide transport protein SpoOK (or perhaps alternatively by a second, normally cryptic, transporter encoded by the app operon; Koide and Hoch, 1994). In an unknown manner, this peptide then promotes ComA activation and thus development of competence. Under harsh environmental conditions, development of competence is inadequate for immediate survival, so Bacillus initiates a differentiation process that results in production of a metabolically dormant, environmentally resistant form called the endospore (Fig. 3). The process of endospore formation in B . subtilis initiates on the onset of starvation and high cell density, and a series of gene families are transcriptionally activated in a temporal manner by employing a combination of alternate sigma factors (reviewed in Losick and Stragier, 1992; Haldenwang, 1995) and transcriptional activators (Strauch and Hoch, 1993). The initial steps for the activation of the sporulation process also independently play a role in transcriptionally regulating other developmental events in Bacillus (such as competence establishment and protease production), with many of the transcription factors involved demonstrating complex patterns of regulation to coordinate these events (Strauch and Hoch, 1993). Many of these genes are activated in a localized manner, either in the mother cell compartment or the prespore compartment through the use of alternate sigma factors, which change the specificity of RNA polymerase. This process provides a genetically amenable system for addressing questions of both temporal and spatial localization of gene expression and intercellular communication. Accumulation of the earliest gene products important for sporulation is directed by RNA polymerase carrying either the vegetative sigma factor, IT*,or the alternate sigma factor, crH (reviewed in Hoch, 1993). Some but not all of the early genes, which are expressed in vegetative cells at lower levels, depend on mH for properly timed expression at levels adequate for sporulation. Three of these genes, SPOOF, spoOB, and spoOA, form a phosphorelay system that is sensitive to activation at several points, allowing responses to different physiologically generated signals. Ultimately, a phosphate is transferred to the SpoOA protein, which is a transcriptional activator of another set of genes, including several global transcriptional regulators. Activation of SpoOA is sensitive to DNA damage (Ireton and Grossman, 1994b) as well as to the position of the cell in the cell cycle; DNA replication must proceed past a specific point before the onset of SpoOA phosphorylation and sporogenesis may begin (Ireton and Grossman, 1994a; Hauser and Emngton, 1995). SpoOA activation is also responsive to the metabolic state of the cell, through sensing of Krebs cycle intermediates (Ireton et a l . , 1995). Once SpoOA-dependent genes are activated, the next step in spore formation is

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the asymmetric septation of the cell into two compartments: mother cell (the larger) and forespore (the smaller). Prior to division, the cell carries two copies of its circular chromosome; bacterial chromosomes and their associated proteins are referred to as nucleoids. One of the two nucleoids is actively transported into the forespore compartment with the aid of the spoZZZE gene product (Wu and Errington, 1994; Sharpe and Errington, 1995), possibly in a manner analogous to plasmid conjugation (L. J. Wu et al., 1995). Once septation is complete, different sets of genes are transcribed in each compartment, as a consequence of the activation of different sigma factors. In the forespore, uF is initially maintained in an inactive state by association with the anti-sigma factor SpoJIAB. When septation is completed, the SpoIIE protein, localized at the site of septation (Arigoni et al., 1995), dephosphorylates the anti-anti-sigma factor SpoIIAA protein (Duncan et al., 1995), which then preferentially combines with SpoIIAB to release active uF. This equilibrium is also influenced by the ratio of ATP to ADP in the compartment, further favoring activation of UF in the forespore (Alper et al., 1994; Diederich et al., 1994). While aFactivates transcription specifically in the forespore compartment, a different set of genes is transcribed in the mother cell by a distinct sigma factor, crE. This sigma factor is synthesized as p r o d , with an additional 29 N-terminal amino acids that must be removed for its activation. The protease that removes this pro sequence is likely to be the product of the spoZZC gene, located within the same operon encoding pro-aE. However, processing also requires a forespore-specific gene product dependent on ITF for transcription ( Jonas and Haldenwang, 1989; Errington and Illing, 1992). This regulation of genes in one compartment by factors located in the other compartment is referred to as criss-cross regulation (Losick and Stragier, 1992). It appears that the @-dependent, forespore-specificgene is spollR (or csm,which is proposed to be secreted from the forespore via an N-terminal signal sequence (Karow et al., 1995; LondoiioVallejo and Stragier, 1995). An extracellular factor, probably the SpoIIR protein, has been detected with functional activity as the specific signaling molecule (Hofmeister et a l . , 1995). The mechanisms by which this signal molecule activates the SpoIIG protease, and how activation is limited specifically to the mother cell compartment, remain unknown. With the activation of crE and uF,transcription of new sets of genes is initiated in the differentiating compartments. Information is just beginning to emerge about the role of the aF-transcribed genes in the forespore. In the mother cell, another transcription factor, SpoIIID, acts in conjunction with uE.The appearance of SpoIIID is somewhat delayed relative to & activation, and thus serves to establish an additional temporal pattern of &-dependent transcription (Illing and Errington, 1991). The genes under uE control are responsible for driving the next morphological stage of sporulation, the engulfment of the forespore by the mother cell. Once engulfment is complete, another sigma factor, uG, is expressed in the forespore compartment. Expression of aG is dependent on UF as well as on as

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yet unidentified factors from the mother cell compartment (Partridge and Errington, 1993). The structural spore formation genes are under crG control. Unidentified accessory transcription factors are likely to play a role in setting the timing of transcription of these genes, based on the differential timing of their expression (Panzer et al., 1989). At approximately the same time that uG is activated in the forespore, UE in the mother cell is replaced by uK. Like u E , uK is synthesized as inactive pro-uK, with an additional 20 N-terminal amino acids. Removal of the pro sequence is again dependent on a protease, apparently SpoIVFB (Lu et al., 1995), and on a signal from the forespore compartment. Transduction of this signal only requires the &-transcribed gene spoIVB, operating in an unknown manner (Gomez et al., 1995). Genes transcribed by @-containing RNA polymerase primarily include those responsible for spore maturation and release, to complete the process of endospore formation. While the extensive work on B . subtilis development has resulted in an understanding of many of the events and processes regulating its differentiation, there are still interesting questions to be asked. With the advent of sensitive techniques to localize gene products within the developing cell (Arigoni et al., 1995), questions about the role of spatial orientation in the function of the regulatory pathways may fruitfully be pursued. Further genetic and biochemical analyses of the components of the regulatory pathways are likely to reveal additional players as well. Because of the depth of understanding of this organism, Bacillus as a model system will remain at the forefront of investigation of the fundamental processes of bacterial development.

2. Induction of Myxococcus Fruiting Body Development by Starvation Like Bacillus, the gram-negative myxobacteria also respond to starvation by the formation of metabolically dormant, environmentally resistant spores, although the sporulation process is quite distinct (see Fig. 4). The best studied member of this family is M. xanthus. The multicellular developmental process of this organism has been the focus of several reviews (Shimkets, 1990; Dworkin and Kaiser, 1993; Hartzell and Youdarian, 1996), to which the reader is referred for a more detailed treatment. Myxococcus xanthus is a rod-shaped soil bacterium with gliding motility. It produces a variety of extracellular hydrolases, lipases, and proteases that degrade macromolecular organic material, including other cells (Burnham et al., 1984). It is a social bacterium, maintaining close contact with other M . xunthus cells both to maximize efficiency of extracellular degradation and as a basis for swarming by gliding motility. On starvation, the myxobacteria aggregate into fruiting bodies, within which a subpopulation of cells differentiates into spores, a process phenotypically very similar to sporogenesis of the eukaryote Dictyostelium (reviewed in Cotter et al., 1992; Kay, 1994; Firtel, 1995). These

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Fruiting body formation, Spore maturation

Dormant spores

Aggregation

\

Germination

Vegetative cells

Fig. 4 Developmental sporulation of M .xnnthus. This soil bacterium exists vegetatively in a social state, communally taking advantage of their ability to break down extracellular macromolecules, including other bacteria. Division in the vegetative state is by binary fission (not shown). On sensing both stress conditions (including starvation or toxic metabolite accumulation) and high cell density, the cells initiate a program of development. The first step is aggregation of the M . xunrhus into a mound containing approximately 100,000 cells; this mound then differentiates into a fruiting body, with concomitant differentiation of some of the cells into resistant spores localized to the interior of the fruiting body. When favorable conditions are present, the spores germinate into the vegetative cell form. Enclosure of the spores in fruiting bodies allows them to remain associated and germinate together, to facilitate social interaction with neighboring M . xunthus cells immediately after their germination.

myxospores are formed by the differentiation of the entire cell, rather than by endospore formation as seen above for B . subtilis. This developmental process has been shown to be orchestrated by a series of intercellular signaling events, which have been divided into five groups (designated A, B , C , D, and E signals) (Hagen et al., 1978; Downard et al., 1993). Deficiencies in any of these signals

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may be complemented by adding cells that are wild type for that signaling pathway, demonstrating the extracellular nature of these signals. These signals are temporally regulated during the process of aggregation, with A, B, and E signals generated early, while C and D are produced later. Developmentally regulated genes induced during the aggregation phase require different combinations of these signals for activation. How are these intercellular signals activated by starvation? Since amino acids are a major nutrient source for Myxococcus, an early supposition was that intracellular sensing of amino acid starvation could serve as a metabolic signal. In E. coli, amino acid starvation induces the formation of guanosine pentaphosphate or tetraphosphate [abbreviated (p)ppGpp] in the stringent response; indeed, starved M. xanthus cells exhibited increased levels of these molecules (Manoil and Kaiser, 1980). More recently, it has been shown that ectopic synthesis of (p)ppGpp is sufficient to activate developmentally controlled genes in nonstarved cultures, indicating that it does play a role in the initiation of the sporulation pathway (Singer and Kaiser, 1995). However, (p)ppGpp itself does not appear to be sufficient to activate the entire process of sporulation (Singer and Kaiser, 1995), suggesting that other signals are required to sustain development, once initiated. The A signal is composed of low concentrations of six free amino acids or small peptides containing them. They appear to act as a reporter of cell density, important in M . xanthus sporulation, since high concentrations of cells are required for successful fruiting body formation. Genes identified as important for releasing the A-signaling set of amino acids have been shown to encode a novel signal transduction protein (AsgA) with similarity to members of the two-component response regulators (Plamann et al., 1995), as well as a DNA-binding protein (AsgB) (Plamann et al., 1994). These gene products are proposed to sense starvation conditions leading to induction of A signal production. One gene whose transcription requires A signal has been characterized, and found to be expressed from a +-like promoter requiring an upstream-binding transcriptional activator (Keseler and Kaiser, 1995; Gulati et al., 1995), suggesting that the A signal may alter cellular transcription (either directly or indirectly) by activating dependent transcription. Production of the B signal, important early in development, requires a membrane-associated protease, BsgA, with similarities to the Lon protease family (Gill et a/., 1993). The early E signaling requires the esg gene product, an a-keto acid dehydrogenase involved in branched-chain amino acid metabolism (Toal et al., 1995). The mutation defining the D signaling pathway, acting later in development, has been localized to the Myxococcus homolog of translation initiation factor IF3; the M . xanthus IF3 homolog was found to have a C-terminal extension that appears to be developmentally significant (Cheng et al., 1994; Kalman et al., 1994). While the identification of these gene products allows signaling pathway models to be proposed, biochemical mechanisms of extracellular B , D, or E signaling remain unclear at this time.

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Perhaps the best characterized of the intercellular signals is the C signal. Mutations blocking signal production all map to the csgA gene, which encodes a small protein shown by immunological studies to apparently reside at the cell surface (Shimkets and Rafiee, 1990). Purified CsgA protein can be added to csgA cells to rescue development, indicating that it is necessary and sufficient for extracellular signaling (Kim and Kaiser, 1990b). Signaling appears to be transmitted by end-to-end contact between aggregating cells (Kim and Kaiser, 1990a; Sager and Kaiser, 1994), to modulate directly the activities of more than 20 genes whose products are involved in motility, aggregation, and sporulation. However, the product of the csgA gene shares homology with a family of alcohol dehydrogenases, and the possibility has been put forth that it functions enzymatically rather than by direct signaling (Lee and Shimkets, 1994). Further studies on the biochemical mechanism of C signaling should address this possibility. In addition to these intercellular signals, myxobacterial development involves a number of other regulatory pathways, including developmentally dependent ADP ribosylation of proteins (Eastman and Dworkin, 1994), protein phosphorylation by eukaryotic-like serine/threonine kinases (Udo et al., 1995), and the use of alternate sigma factors (Inouye and Inouye, 1993). However, no clear cascade of intracellular signaling events controlling the sporulation process has been demonstrated for M . xunthus; further studies aimed at elucidating both intercellular and intracellular pathways and their interrelation will shed light on the complex process of development in this bacterium.

3. Sporulation of Streptornyces on Starvation Bacteria in the genus Streptornyces exhibit a life cycle quite distinct from most prokaryotes, and indeed more like that of the eukaryotic fungi (Fig. 5; reviewed in Chater, 1993). In vegetative growth on a rich nutrient source, the bacteria grow as a mycelium of branching hyphae. Internal septa that normally segregate nucleoids into discrete cells are rather infrequent, so that hyphal compartments (especially those at the growing tips) contain many nucleoids. When nutrients become limited or inhibitory metabolites or signaling molecules begin to accumulate, then aerial hyphae begin to emerge. These hyphae obtain some of their nutrients by recyling the biomolecules incorporated in the vegetative mycelium. Eventually, the tips of the aerial hyphae undergo further development by multiple septation to give unigenomic compartments. These undergo additional differentiation to form long chains of dormant spores that allow long-term survival and dissemination. At about the time when aerial hyphae begin to grow, the colony starts to produce the antibiotics and other secondary metabolites for which the genus is famous. This temporal link appears to reflect the existence of common regulatory elements between the two processes. Such a supposition is supported by the phenotype of “bald” (bld) mutants, which cannot form aerial hyphae and also are

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Spore maturation

Septation of Hyphae

T

=

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Aerial Hyphae (

t

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Fig. 5 Sporulation pathway of the filamentous bacterium S . coelicolor. In its vegetative state, this bacterium grows as a mycelium of surface or subsurface-associated filamentous hyphae. Septa are rare within the hyphae, resulting in multiple nucleoids within each physical compartment. On exposure to stress (here indicated by starvation], the mycelium produces aerial hyphae. During sporogenesis, the aerial hyphae first assume distinctive morphologies, including branching or curling at the hyphal tips (not shown here), and nucleoids accumulate at these tips. Next, septa segregate the nucleoids into cells that then differentiate into dormant spores. These environmentally resistant spores may then be disseminated to a location with favorable growth conditions, where they germinate and form vegetative mycelia.

blocked in secondary metabolite production. The aerial mycelium deficiency of many of the bld mutants is manifested only under certain nutritional growth conditions, including growth in the presence of glucose or certain other carbon sources. This suggests the existence of complex interactions between developmental and metabolic processes. Analysis of bld mutants has revealed important

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developmental roles for extracellular signaling molecules. For example, in S . coelicolor , several different classes of bld mutations are complemented for growth of aerial hyphae (but interestingly, not for antibiotic production) when grown close to a colony either of a different class of bld mutant or of wild type, implying the existence of a hierarchy of intercellular signal exchange (Willey et al., 1993). It has been suggested that these regulatory interactions culminate in the production of an aerial hyphae-associated small peptide, SapB, which can also partially complement many bld mutants if added extracellularly (Willey et a l . , 1991). The best characterized of the extracellular signaling molecules is the A factor, an excreted y-butyrolactone of Streptomyces griseus. A factor is freely diffusible across membranes, and at sufficiently high extracellular concentrations it activates genes involved in sporulation and antibiotic production by combining with a cytoplasmic transcriptional repressor protein to inactivate it (Miyake et al., 1990; Onaka et al., 1995). Sporulation can be restored to A factor-deficient S. griseus mutants, or accelerated in some wild-type strains, by extra copies of a locus encoding homologs of an ATP-dependent membrane transporter protein and a two-component response regulator protein (Ueda et al., 1993; Ma and Kendall, 1994). The manner in which these operate to restore sporulation is currently being investigated. Translational regulation of development is implied by the discovery that the bZdA gene encodes the only tRNA capable of recognizing the rare UUA codon (Lawlor et al., 1987). The TTA codon appears to be absent from all essential vegetative genes, but is present in some regulatory genes for secondary metabolite production and in a gene that complements the sporulation defect in some bld mutants of S. griseus (Babcock and Kendrick, 1990). Following the successful production of aerial hyphae, additional genes are specifically required for formation of the mature spore chains. Many of these have been identified in S. coelicolor by the study of white colony (whi) mutants, which form an aerial mycelium but fail to develop the gray-brown spore coat pigment. Six whi genes acting early in spore development, at the time of septation of the hyphae destined to become spores, have been identified. The sequence of one of these, whiG, suggests that it encodes an alternate sigma factor (Chater et al., 1989). The closest homologs of a W h i G in other bacteria are involved in motility, and do not function in sporulation. It is not yet clear whether any of the other whi genes depend directly on a W h i G for transcription, although two other awhiG-dependentpromoters for genes of unknown function have been described (Tan and Chater, 1993). Most of these early whi genes have now been cloned (K. F. Chater, personal communication), so their regulatory interplay should soon be clarified. whi genes acting late in development have also been described, mutants of which complete prespore hyphal septation but fail to complete spore wall thick-

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ening and/or spore pigment biosynthesis. Notably, one of these (called sigF) also appears to encode an alternate sigma factor, the closest homologs of which are the three B . subtilis sigma factors UB, uF, and crG; two of these ( a F and aG) are responsible for the forespore developmental program (PotGckovB et al., 1995). This may suggest that the later stages of sporulation in bacilli and streptomycetes share common regulatory properties. A number of additional alternate sigma factors have been identified in various species of Streptomyces, which demonstrate differing temporal patterns of expression (Kormanec and Farkasovsky, 1993; Marcos et al., 1995). This diversity of identified Streptomyces sigma factors also suggests that a regulatory cascade similar to that in B . subtilis may yet be elucidated. Spatial localization of sporulation in Streptomyces raises other questions, such as how nutrients are supplied to aerial hyphal tips and whether the metabolic genes involved are subject to developmental regulation. At least in the cases of polyketide biosynthetic enzymes needed to make the spore pigment and certain antibiotics, and of branching enzymes needed to make glycogen, spatially localized enzyme isoforms are encoded by distinct genes (Yu and Hopwood, 1995; Bruton et al., 1995). Perhaps surprisingly, however, the products of some cell division genes (such as ftsz) that might be predicted to be differentially used in sporulation appear to function both in vegetative and sporulation-specific septation (McCormick et al., 1994). Further analysis of the regulation of the sporulation-activated expression of ftsZ and of spore pigment and glycogen metabolismrelated genes may provide insight into the control of spatially localized gene expression and the regulatory roles and interplay of the early whi genes in this process. Mutational analysis has identified a number of Streptomyces genes important in the process of sporulation. Those genes identified at a molecular level thus far have provided an intriguing glimpse into the process of differentiation in this bacterium. Additional analysis of these mutants should help characterize the signaling cascades responsible for regulating the complex developmental processes involved in Streptomyces aerial hyphae production and sporulation, and how these relate to the developmental induction of secondary metabolite production.

4. The Symbiotic Relationship between Rhizobium and Legumes In their free-living state, members of the gram-negative Rhizobium genus have growth cycles similar to those seen for E . coli. However, these bacteria are able to establish symbiotic relationships with members of the plant family Leguminosae (Fabaceae) (Fig. 6 ) , with variable host range seen for the different Rhizobium species (more specifically reviewed in Long and Staskawicz, 1993; Downie, 1994; Van Rhijn and Vanderleyden, 1995). This symbiosis represents a

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Cortical

Attraction

Binding, root hair curling

Bacteroid release

Infection thread formation

Fig. 6 Nodulation of legume roots by Rhizobium. Diagrammed is a cross-section of a portion of a legume root, with a root hair, the epidermal layer and the cortical layer of cells indicated. Arrracrion: The plant excretes flavonoids and other compounds into the surrounding soil where a strain of Rhizobium, capable of recognizing a specific Aavonoid, responds by associating with the root hair. Binding, roof hair curling: The flavonoids induce expression of the nod family of genes within the bacterial cell, culminating in the synthesis and export of a Nod factor signaling molecule (a lipooligosaccharide). The Nod factor induces physiological changes in the root tissue that prepare it for bacterial invasion. Infection threadformation: One effect of the intercellular signaling is to promote the formation of an infection thread, composed of plant cell wall material, within which the bacteria multiply. Bacteroid release: The infection thread penetrates to the cortical layer of cells within the root, where the Rhizobiurn cells are release, enclosed in a plant cell-derived membrane layer. These bacteria differentiate both metabolically (most significant is the induction of genes responsible for nitrogen fixation) and structurally to form bacteroids. Cortical cells, primed by the Nod factor to differentiate into cells prepared for nodulation, become filled with bacteroids to form the functional root nodule (not shown).

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developmental pathway unique from those discussed above, since it requires signaling from an entirely different organism to initiate and sustain the differentiation process. In the soil, free-living Rhizobium is motile and is chemotactically attracted to legume roots both by nutritional gradients (Caetano-Anollts et al., 1992) as well as by flavonoids, specific signaling molecules excreted from the roots and received by the bacteria (Phillips, 1992). Minor chemical differences between flavonoids excreted by different plants assist in determining the specificity of the plant-bacterial interaction (Phillips, 1992). The flavonoid interacts with the product of the nodD gene, which encodes a transcriptional activator of the LysR family (reviewed in Schell, 1993). The activated NodD protein is able to bind to a conserved sequence upstream of other nod genes to activate their transcription (Fisher and Long, 1993). These induced nod gene products then contribute to the synthesis of a signal sent back to the plant: a lipooligosaccharide (Nod factor), with a chitin-like backbone linked to a fatty acid. One group of nod genes conserved broadly among the rhizobia is responsible for synthesis of the basic lipooligosaccharide backbone, while others that are specific for a given Rhizobium species are responsible for modification of this basic backbone to impart plant host specificity (Denarie et al., 1992; Fisher and Long, 1992). The exported lipooligosaccharide acts to induce specific morphological changes in the plant in preparation for symbiosis establishment, especially those that allow bacterial entry into the root. To invade the plant, different rhizobia use different strategies. Entry can occur by penetration of the cell surface of a morphologically altered root hair; alternatively the bacteria may invade directly through the primary root wall, either between the outer epithelial cells or at a site of cell wall damage (Brewin, 1991). Invasion is facilitated by carbohydrates found on the surface of the bacterium, including lipopolysaccharides, exopolysaccharides, and neutral p- 1,2-glucans (reviewed in Gray et al., 1992). It is not clear how these molecules serve in the signaling of the plant to induce bacterial uptake. Once the bacterial cell has entered the plant, it is surrounded by newly synthesized plant cell wall material; in this environment the bacterium continues to grow, forming an infection thread that extends into the root. On reaching cortical cells primed for symbiosis, the bacterial cells are released, encased in a plant cytoplasmic membrane vesicle. Up to this point, the Rhizobium cell, while expressing a family of nodulationspecific genes, has not been observed to undergo dramatic alterations in its physiology. Once within the plant cortical cell, however, the bacterium develops into a bacteroid. The heme-containing sensor kinase FixL (de Philip et al., 1990; Gilles-Gonzalez et al., 1991) detects the plant cell-mediated establishment of a suitable microaerobic environment and initiates a signaling cascade to turn on the genes responsible for nitrogen fixation (nf genes). FixL passes its phosphate signal to the response regulator FixJ (Weinstein et al., 1992; Reyrat et al., 1994), which in turn activates NifA, the global activator of the nfgene family, and

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FixK, controlling a separate activation cascade (reviewed in Fischer, 1994). Other bacterial regulatory genes are also likely to play a role in the establishment of symbiosis, as indicated by mutations in the bacA gene, encoding a membrane protein, which block bacteroid maturation (Glazebrook el al., 1993). Sweeping changes also occur in many other aspects of the bacterium; it becomes substantially enlarged and may change its shape, it alters its metabolism to limit the intracellular oxygen concentration, and it modifies its transport mechanisms to export the fixed nitrogen in exchange for carbohydrate from the plant cell (reviewed in Werner, 1992). It has been proposed that the bacteroid represents a terminally differentiated state, although this has not been clearly demonstrated. Owing to the difficulty in manipulating these intracellular symbiotic bacteroids, much remains to be examined as to the specific symbiotic relationship and the steps in the developmental process responsible for bacteroid formation once inside the plant.

5. Adaptation of Escherichia coli to Stationary Phase Thus far, we have distinguished between developmental processes employed by bacteria and their normal, "vegetative" pattern of growth. Many bacteria, including the enteric gram-negative E . coli, do not produce physiologically obvious differentiated forms (such as environmentally resistant spores or the bacteroids seen in Rhizobium nodulation of legumes), so they have been generally considered to lack a differentiated cell type. However, it has been demonstrated that entry of E. coli, and many other bacteria, into stationary growth phase involves a variety of physical and metabolic changes, in several ways akin to spore formation. Thus developmental components of the life cycle appear to be the rule rather than the exception even among the bacteria. In most natural settings, lack of nutrients forces a bacterium to spend most of its time in a metabolically repressed stationary phase. When the required nutrients become available, the cell must be able to assimilate them rapidly and use them to maximize growth. At some point, one or more nutrients again become limiting, and the cell must now make the transition to stationary phase to allow survival until nutrients are again available. On sensing nutrient deprivation, including carbon or nitrogen starvation (reviewed in Matin, 1991), cellular stress, or phosphate limitation (Spira et al., 1995), the cell synthesizes the messenger molecule (p)ppGpp as part of the stringent response (reviewed in Cashel and Rudd, 1987). (p)ppGpp then serves to attenuate transcription of rRNA operons and other genes by a direct interaction with FWA polymerase (Reddy et al., 1995). This signaling molecule also acts to inhibit new rounds of DNA replication and promotes reductive cell division, thus distributing the multiple nucleoids that may exist in rapidly growing bacteria to individual cells (Schreiber et al . , 1995). These effects reduce the overall metabolic rate of the cell, in preparation for transition to stationary phase.

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If nutrient levels do not rise or stress conditions do not dissipate, then the transition initiated by (p)ppGpp signaling is extended to the establishment of the stationary phase state (reviewed in Kolter et al., 1993; Hengge-Aronis, 1993; Kjelleberg, 1993). It has been reported that a homoserine lactone or a derivative of this molecule might also be involved in signaling the need to transition to stationary phase; this signal, like (p)ppGpp, is also likely to act intracellularly, unlike the extracellular homoserine lactone signal of Vibrio (Huisman and Kolter, 1994). During this transition, the E. coli cell changes morphologically from rod shaped to a much smaller, almost spherical cell by reductive division to segregate all available nucleoids to individual cells. The nucleoid condenses and the membrane and cell wall compositions change. Perhaps most significantly, synthesis of a new family of proteins is induced (reviewed in Matin el al., 1989). These proteins are induced at different times during entry into stationary phase; some are induced at the transition and others are induced later. A number of these genes are transcribed by a stationary phase-specific sigma factor, called us(reviewed in Loewen and Hengge-Aronis, 1994). The cellular concentration of this rpoS-encoded sigma factor is regulated at the level of transcription through several promoters, as well as at the levels of translation and protein stability (Lange and Hengge-Aronis, 1994). The functions of genes controlled by us include management of DNA damage, alteration of cell morphology, and establishment of a stress-resistant state, capable of enduring heat shock, changes in osmolarity, and exposure to toxins such as H,O, (Loewen and Hengge-Aronis, 1994). Expression of the rpuS gene is induced by the presence of (p)ppGpp, linking this early signal to stationary phase induction (Gentry et al., 1993; Lange et al., 1995). Use of a stationary phase-specific sigma factor appears to be relatively widespread among gram-negative bacteria, as rpoS homologs have been identified in Salmonella typhimurium (Kowarz et al., 1994) and Pseudomonas aeruginosa (Fujita et al., 1994), and E. coli rpoS-dependent promoters have been shown to be induced properly in stationary phase in several other gram-negative bacteria (Miksch and Dobrowolski, 1995). In addition to direct regulation of transcription initiation, us also functions indirectly to control transcriptional cascades through the activation of other transcription factors, including BolA, AppY, and Dps, each controlling different subsets of genes and responding to additional stress-related signals necessary for their induction (Loewen and Hengge-Aronis, 1994). This broadens the adaptability of the stationary-phase response to modulate gene expression depending on the nature of the stress the cell is facing. Additionally, it is clear that (p)ppGpp and rpoS are not the sole players involved in stationary-phase induction. Genes transcribed by other sigma factors, including the heat shock sigma factor and u54,are induced at the transition to stationary phase (Jenkins el al., 1991; Weiner and Model, 1994), and the genes encoding the subunits of the integration host factor (IHF) protein in E. coli are regulated differentially by rpoS and (p)ppGpp (Aviv et al., 1994). Thus, induction of the stationary-phase re-

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sponse appears to be a complex process that appears flexible enough to allow the inducing stimulus to set an appropriate pattern of gene expression. After the initial establishment of the stationary state, cellular metabolism is dramatically reduced, with protein synthesis continuing at only 0.05% the rate of logarithmically growing cells (Kolter et al., 1993). Survival depends on the conditions prevalent on entry into stationary phase; for example, cells grown in rich medium experience a reduction of one to two orders of magnitude in the number of viable cells, while cells in minimal medium may show little or no loss in viability (Siegele et al., 1993). As conditions change again to favor growth, E. coli rapidly reactivates its metabolic pathways. In another gram-negative bacterium, Vibrio sp. S14, a family of approximately 20 genes is induced on reentry into log phase (Albertson et al., 1990). In E. coli,however, only one gene, surB, has been identified thus far as required for this transition; the SurB protein shares homology with membrane transport proteins although its function is not yet known (Siegele and Kolter, 1993). An appreciation of the differentiation of bacteria on the transition between log and stationary phase is relatively new, yet a number of interesting players have been identified; these include (p)ppGpp, rpoS, and the genes regulated either directly or indirectly by them. Future efforts will certainly extend our understanding of the interplay of these components with each other as well as with other cellular components yet to be identified.

111. Control of Cellular Differentiation during the Caulobacter crescentus Cell Cycle A. Temporal and Spatial Control of Chromosome Replication An essential aspect of cell cycle regulation is the proper control of chromosomal replication and segregation. For eukaryotes, while the mechanisms involved in the initiation and control of DNA replication are not clearly defined, the mitotic and meiotic segregation of the replicated chromosomes is understood in much greater detail than for prokaryotes (reviewed in Wadsworth, 1993). In contrast, at least in E. coli, we have a good understanding of the biochemical mechanisms of the initiation and elongation of DNA replication. It appears that initiation is temporally regulated, occurring at a distinct time in the cell cycle (Nordstrom and Austin, 1993), and that the regulation of reinitiation tightly controls the number of chromosome copies (Lu e l al., 1994). A detailed model of the enzymology of the elongation process on both leading and lagging strands has been proposed (Kornberg and Baker, 1992; Stukenberg et al., 1994). However, the mechanisms responsible for segregation of the replicated nucleoids are still largely undefined. Spatially, no distinction appears to be made as to which of the two chromosomes is segregated to each daughter cell. Work in E. coli has identified

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some candidate genes that may be involved in the process of chromosome partitioning (reviewed in Hiraga, 1993), but the precise role of such genes is not clear. Even the timing of segregation has not been resolved, with some suggestion that partitioning occurs rapidly, near the time of cell division (Hiraga et a l . , 1990; Begg and Donachie, 1991), and other evidence favoring a slower separation of the nucleoids associated with cellular growth (van Helvoort and Woldringh, 1994). More is known about chromosome distribution during the asymmetric division of B . subtilis near the beginning of endospore production, where the SpoIIIE protein is involved in actively segregating a nucleoid into the forespore compartment (Wu and Emngton, 1994; L. J. Wu et al., 1995). Evidence indicates that SpoIIIE plays a role in chromosome segregation in vegetative cells as well (Sharpe and Errington, 1995). Still, the physical and biochemical mechanisms that orchestrate the partitioning of nucleoids remain elusive. Caulobacter crescentus provides an amenable system for addressing questions about the time and place of DNA replication. Unlike E . coli, the chromosome of C . crescentus initiates replication only once each cell cycle, eliminating the complexity of multiple chromosome copies within the cell. More interesting is the observation that the nucleoids from the two cell types have quite distinct sedimentation coefficients, with swanner cell nucleoids sedimenting faster (>6000S) than the stalked cell nucleoids (-3000s) (Evinger and Agabian, 1979; Swoboda et a l . , 1982). This implies distinct differences in higher order structure; and since no transition forms with intermediate sedimentation coefficients have been detected, it implies a specific reordering of chromosome structure during the cell cycle. It is also known that only the stalked cell chromosome is competent for initiation of DNA replication; initiation occurs at the time of the swarmer-to-stalked cell transition and in the stalked cell following cell division (Degnen and Newton, 1972; Dingwall and Shapiro, 1989). It is not known if the observed differences in higher order nucleoid structure are related to control of replication initiation. What factors are responsible for controlling the timing of C . crescentus DNA replication initiation during the cell cycle? Two major possibilities exist a priori; first, some or all of the machinery required to initiate replication may be specifically localized to stalk cells. Alternatively, initiation could be directly regulated, either by factors that specifically repress initiation in the swarmer cells or that specifically activate it in the stalked cells. It is unlikely that replication factors are absent in swanner cells, because it has been shown that RK2-derived plasmids, which rely heavily on the host replication machinery, are replicated in swarmer cells, albeit with lesser efficiency than in stalked cells (Marczynski et al., 1990). To address the possibility of direct regulation of initiation, the C . crescentus chromosomal origin of replication was identified and cloned as an autonomously replicating plasmid (Marczynski and Shapiro, 1992). The cloned origin displayed proper timing of replication initiation (Marczynski and Shapiro, 1992), indicating that all of the cis-acting elements required for replication control were

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present. Further analysis has revealed similarities to the E. coli chromosomal replication origin, including the presence of an A/T-rich sequence and essential DnaA boxes, but distinct differences were also found. These included two novel repeated motifs (an AAGCCCGG 8-mer and an interrupted GTTAA-n7-TTAA 9-mer) and dependence on transcription of the adjacent housekeeping hemE gene (Marczynski et al., 1995). The hemE gene is expressed from two promoters labeled Pweakand Pstrong,both located within the minimal original of replication. The Pweakpromoter is constitutively transcribed and is likely to be responsible for the synthesis of the HemE enzyme (a homolog of uroporphyrinogen decarboxylase). In contrast, transcription of Pstrongis temporally controlled; it is highest during the stalked cell phase of the cell cycle, and is specifically repressed both in the swanner compartment of the predivisional cell and also in the progeny swarmer cell (Marczynski et al., 1995). The mRNA initiating from Pstrongis not translated. On the basis of these findings, a model for the control of DNA replication initiation in C. crescentus has been proposed. This model centers on the hypothesis that the Pstrongtranscript remains associated with the template DNA, to produce an RNA::DNA hybrid important either in the priming of replication or in transcriptional activation of replisome formation (Marczynski et al., 1995). Inherent in this model is the suggestion that a transacting factor functions to control differentially the transcription of Pstrong. It was demonstrated that repression of Pstrongspecifically in the swarmer compartment of the predivisional cell and in the progeny swarmer cells coincides with the repression of DNA replication initiation in these cell types. Analysis of the Pstrongpromoter region showed that two of the conserved 9-mer interrupted repeats overlapped the Pstrong-10 and -35 promoter sequences. An essential response regulator protein, CtrA, initially identified as linking flagellar synthesis to cell cycle events (described below), has been shown to bind to these interrupted 9-mer repeats. Furthermore, in a strain bearing a temperature-sensitive ctrA allele, the activity of Pstrongis elevated severalfold (Quon et al., 1996). These observations strongly suggest that CtrA is the factor responsible for inhibition of DNA replication initiation in the swanner cell (G. Marczynski and L. Shapiro, unpublished observation; Quon et al., 1996). Thus, the regulation of replication initiation is associated with a regulatory protein known to play a central role in transmitting cell cycle cues to a variety of cellular processes. Further work remains to identify other factors that contribute to CtrA-mediated regulation of cell type-specific replication initiation during the C. crescentus cell cycle. As is the case for E. coli, no mechanism responsible for the segregation of the replicated C . crescentus chromosomes within the predivisional cell has been elucidated. The possibility has been raised that perhaps the replicated daughter chromosomes are specifically targeted to one pole or the other, based on a molecular marker that distinguishes the parent strands. Such distinctions could

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be made on the basis of properties of DNA methylation. A DNA methyltransferase, CcrM, that modifies the adenine residues in the sequence GAnTC has been identified (Zweiger et al., 1994). On replication, remethylation of the newly synthesized strand does not occur until just prior to cell division, meaning that the DNA remains hemimethylated, allowing efficient distinction between old and new strands (Zweiger et al., 1994; Stephens et al., 1995a). However, chromosomal segregation has been shown to be random, demonstrating that there is no predetermined fate for the differentially methylated chromosomal strands (Osley and Newton, 1974; Marczynski et al., 1990). Identification and characterization of the chromosomal origin of replication have fostered a deeper understanding of the mechanisms that regulate the temporal and spatial control of chromosome replication in C. crescentus. Further insight into the mode of action of the CtrA response regulator should broaden our understanding of this control. Transcriptional regulation of DNA replication genes may play an additional role in controlling the timing of replication initiation (see discussion below). Given that there is a link between DNA replication and expression of the hierarchy responsible for flagellar biogenesis (Osley and Newton, 1978, 1980; Dingwall et a l . , 1992a; Stephens and Shapiro, 1993), an understanding of the control of DNA replication will shed light on other questions of temporal expression and subcellular localization during the C. crescentus cell cycle. Caulobacter crescentus is also a likely organism in which to identify factors responsible for prokaryotic chromosome partitioning, because of the ease of cell synchrony and the differential replication potential of swarmer and stalked cell chromosomes.

B. Control of the Timing and Subcellular Location of Transcription Because synchronous swarmer populations of C . crescentus cells may be obtained easily, the temporal and spatial expression patterns of a variety of genes have been examined; these include genes involved in the well-studied process of flagellar biogenesis, and more recently those involved in DNA replication. These patterns of expression represent normal events occurring as part of each cell cycle, in contrast to the temporally regulated cascades of gene expression that occur during sporulation in Bacillus, Myxococcus, or Streptomyces or nodulation of legumes by Rhizabium. Temporal expression patterns of genes within the cell cycles of other bacteria have been demonstrated, such as the induction of gid and inhibition of mioC transcription concomitant with initiation of DNA replication from the flanking E . coli oriC replication origin (Ogawa and Okazaki, 1994). It is likely that temporal control of gene expression is significant in a number of contexts in prokaryotic cells, but is most accessible to analysis during the C . crescentus cell cycle.

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1. Genes Involved in Flagellar Biogenesis Within the life cycle of C . crescentus, motility of the swarmer cell plays a key role in cell dispersal. This bacterium synthesizes a single polar flagellum at a specific time in the cell cycle, and estimates indicate that synthesis of the flagellum requires at least 50 genes (Ely and Ely, 1989), many of which have now been identified, cloned, and sequenced (reviewed in Brun et al., 1994; Gober and Marques, 1995; Stephens et al., 1995b; Jenal et al., 1995). Analysis of the epistatic relationships and temporal patterns of expression of these genes allowed them to be grouped into three classes: class I1 genes are the earliest expressed, and include genes involved in export of flagellar components, synthesis of the earliest structural components of the basal body, and genes involved in activating transcription of the class I11 genes (Fig. 7). Mutations in class I1 genes not only block flagellar biogenesis, but also inhibit cell division, demonstrating that proper expression of these flagellar genes is linked to the execution of other cell cycle-related events. The expression of the class 111 genes, which encode the structural components of the remainder of the basal body and the hook, are in turn required for the synthesis of the class IV gene products, which are assembled into the flagellar filament. Members of each of these classes also negatively autoregulate their own expression in a largely uncharacterized manner. Perhaps one regulatory mechanism employed in this process is similar to the anti-sigma factor-mediated regulation discovered in Salmonella and E . coli flagellar biogenesis, whereby activation of transcription is dependent on export of an anti-sigma factor by a completed flagellar basal body (Hughes et al., 1993; Kutsukake et a l . , 1994). Until recently, no genes had been assigned to class I, which has been reserved for regulators of class I1 genes that link their expression to the cell cycle. However, the product of the ctrA gene, a response regulator shown to footprint class I1 flagellar promoters and to modulate the level of class I1 flagellar gene expression, is a likely class I gene candidate (Quon et al., 1996).

a. Regulation of Class I1 Flagellar Genes. The order of transcription of the flagellar genes reflects the order of assembly of their protein products into the nascent flagellum. Transcription of the class I1 genes is coordinately induced in the late stalked cell phase, while the chromosome is in the act of replication. Comparison of the class I1 promoter sequences revealed conserved motifs that differed from known prokaryotic promoters, including consensus C.crescentus housekeeping promoters (Malakooti et al., 1995). In vitro mutagenesis of the promoters of several of the class I1 operons, including t h e j i F operon (Van Way et al., 1993), the JiLM operon (Stephens and Shapiro, 1993), and the j i Q R operon (Zhuang and Shapiro, 1995), revealed that the -10 and -35 regions were essential for promoter activity. In addition, a highly conserved region between the - 10 and -35 sequences played an important role in transcriptional control. It is not clear whether these promoters are recognized by RNA polymerase with the

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(f7]\. &K,

Class IV

+ Hook

(f7gE)

L-nng

(flgH)

P-nng Distal rod

(j¶gG)

E-nng

Prommal rod (&I3 MSring

(pin

+

Class III

I+

Class I1

I+ Fig. 7 Diagram of the C . crescentus flagellum and the flagellar transcriptional cascade. The name of each structure or function is accompanied by the gene designation shown in parentheses. The structure of the C-ring complex is adapted from that proposed for the S. typhirnuriurn basal body (Francis et al., 1994). The known genes, whose products are thought to function in the flagellarspecific export pathway in C . crescentus, are grouped below the MS ring. The genes encoding structural and regulatory proteins are grouped in classes that are expressed in a temporal order that reflects the order of assembly of their protein products. Arrows indicate positive regulation (+), in which the expression of genes within a class requires the expression of the gene products of the preceding class. Boxed are the transcription factors known to function in the regulatory cascade. The regulatory cascade is initiated by as yet unidentified cell cycle cues that result in the synthesis of a class I transcriptional regulator CtrA (Quon et af., 1996). CtrA, an essential signal transduction protein, interacts directly with early flagellar gene (class 11) promoters, thus linking flagellar gene expression to cell cycle events. FlbD, another signal transduction protein, works in concert with u54 to activate transcription of the late class 111 and class IV promoters.

housekeeping u70 subunit (Malakooti and EIy, 1995) or with an alternate sigma factor. It has been shown that mutations in the ctrA response regulator DNAbinding protein altered transcription of the class I1 promoters, and further that the CtrA protein specifically bound to conserved sequences in these promoter regions (Quon et al., 1996). This class I ctrA gene encodes a member of the response regulator family, and presumably is itself regulated by a specific, unidentified sensor kinase linking it to cell cycle cues. In addition to CtrA, however, other elements also regulate expression of the class I1 gene family. It has been

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shown that DNA replication is required for flagellar biogenesis (Osley and Newton, 1978, 1980; Sheffery and Newton, 1981), and more recent studies have shown that a block in DNA synthesis disrupts expression of class I1 promoters (Dingwall et al., 1992a; Stephens and Shapiro, 1993). Whether DNA replication is linked to class I1 promoter expression via the CtrA response regulator or by another mechanism is unclear. In addition, transcriptional repression of at least the class I1 jliF operon appears also to be mediated by the product of the flbD gene, the last gene in theJliF operon (Benson et al., 1994; Mullin et al., 1994; Wingrove and Gober, 1994). This DNA-binding protein, key in regulation of the class 111 genes (see below), binds to thefliF promoter sequence and most likely restricts access of RNA polymerase. Since class I1 genes are maximally transcribed early in the predivisional cell, there is no spatial distinction made in expression between the nascent prestalk and preswarmer compartments, most likely because no physical barrier exists at this time between these compartments. Later in the cell cycle, when DNA replication and segregation are complete and the two compartments appear to be physically separated, the waning class I1 gene transcription from JliLM, jliQR, and more recently fromjliP promoters was found to show no polar bias (Yu and Shapiro, 1992; Zhuang and Shapiro, 1995; Gober et al., 1995). Therefore, except for the fliF promoter (which is specifically repressed by phosphorylated FlbD in the swarmer cell pole; Wingrove and Gober, 1994), spatial regulation is not apparent in class I1 gene transcription. In contrast, at the protein level, products of class I1 genes are generally targeted to the pole opposite the stalk, where the new flagellum is assembled (Gober et al., 1991b, 1995). The mechanism responsible for this protein targeting is currently an area of active investigation. b. Temporal and Spatial Regulation of Class 111 and Class IV Flagellar Gene Expression. Class 111 gene transcription is induced after the class I1 genes, and is dependent on expression of all of the class I1 gene products. If expression of one of the class I1 operons (the$iF operon) is delayed, then class 111 transcription is concomitantly delayed (Ohta et al., 1991). Analysis of class 111 and class IV promoter sequences revealed that they each have a consensus us4 promoter (Mullin et al., 1987; Mullin and Newton, 1989; Dingwall et al., 1990, 1992b; Khambaty and Ely, 1992; Mullin and Newton, 1993), and that class I11 genes can be expressed in vitro using a54-based transcriptional systems (Ninfa et al., 1989; Benson et al., 1994; J . Wu et al., 1995). The C. crescentus rpoN gene encoding 0 5 4 appears to be a member of the class I1 gene group (Brun and Shapiro, 1992). Bacterial genes transcribed from ~5~ promoters require a transcriptional activator, binding either upstream or downstream of the start site, to permit open complex formation and initiation of transcription (reviewed in Merrick, 1993). In C . crescentus flagellar biogenesis, three specific 0 5 4 activatorbinding sequences have been shown by mutagenesis to function in transcriptional

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activation of the crS4-dependent promoters among both class I11 and IV genes, varying in location from -350 bases upstream to -1 10 bases downstream of the site of transcriptional initiation. These include the ftr sequence (Mullin and Newton, 1989; Gober et al., 1991a; Gober and Shapiro, 1992; Mullin and Newton, 1993), the RE-1 and RE-2 sequences (Dingwall et al., 1992b; Khambaty and Ely, 1992; Marques and Gober, 1995), and the Rf-2 sequence (Dingwall et al., 1990). A class I1 gene product, FlbD, has been shown to be the activator that binds to the ftr site (Wingrove et al., 1993; Mullin et al., 1994; Benson et al., 1994). As is the case for other aS4-mediated transcriptional events (North et al., 1993), IHF (integration host factor) has been shown to bind to a site located between the promoter and transcriptional activator sequences (Gober and Shapiro, 1990, 1992; Marques and Gober, 1995). IHF is a DNA-bending protein (reviewed by Goosen and van de Putte, 1995) that is postulated to enhance transcriptional activation by bringing the activator into contact with the us4containing polymerase. The levels of the IHF subunits are modulated during the C. crescentus cell cycle; they are highest in predivisional cells, at the time of maximal transcription of the late flagellar genes expressed from as4-dependent promoters (Gober and Shapiro, 1992). How are the class 111 genes temporally regulated? Although the synthesis of us4,FlbD, and IHF proteins is maximal at the time of transcription of the class I11 genes, this does not appear to be the whole story. FlbD, like CtrA, is a member of the family of response regulator proteins whose activity is controlled by phosphorylation (Ramakrishnan and Newton, 1990). Experiments have shown that while the steady state level of this protein does not change appreciably during the cell cycle, FlbD phosphorylation is under cell cycle control, with maximal phosphorylation coinciding with the time of class 111 gene transcription (Wingrove e f al., 1993). These results suggest that control by phosphorylation of FlbD, in combination with us4 and IHF availability, plays a significant role in setting the timing of class I11 gene expression. The link between class I1 and class 111 gene expression may involve additional factors, identified by mutations in genes that bypass the class I1 requirement for the expression of class I11 and class IV genes (Mangan et al., 1995). In this same study, it was reported that while class I V j j K andjjL flagellin gene transcription was surprisingly not affected in class I1 and class Ill flagellar mutants, flagellin protein was not synthesized. This identifies temporal regulation at the level of translation for the class IV genes, in contrast to the transcriptional regulation seen for class I1 and class 111 genes (Mangan et al., 1995). A major challenge faced by the C . crescentus cell is to ensure that the products of the flagellar genes are correctly targeted to the incipient swarmer pole of the predivisional cell. As noted above, maximal transcription of class I1 genes occurs early in the cell cycle, and thus their mRNAs are not polarly localized. Maximal transcription of several of the class 111 and class IV genes occurs after completion

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of chromosomal replication, and evidence has been presented that these late flagellar genes are transcribed solely from the chromosome in the swarmer compartment of the predivisional cell (Milhausen and Agabian, 1983; Gober et al., 1991b; Gober and Shapiro, 1992). Spatially restricted expression of these genes requires the response regulator FlbD for transcriptional activation. Although the FlbD protein shows no spatial bias, phosphorylation of FlbD appears to occur primarily in the swarmer compartment, because transcriptional activation of promoters occurred in both compartments of a strain containing a constitutively active FlbD allele, locked in the "phosphorylated" state (Wingrove et al., 1993). The characterization of FlbD phosphorylation as significant in polar localization of gene expression indicates that a signal transduction cascade is likely to be involved in spatial control of transcription in this bacterium. The temporal and spatial regulation of gene expression evident in the biosynthesis of the C . crescentus polar flagellum has provided a valuable system with which to address the fundamental mechanisms of bacterial differentiation. The goal is now to determine what is initiating the regulatory signals that turn on flagellar biogenesis at a defined time in the cell cycle. Does chromosomal replication act as a cellular clock to set the timing of other cell cycle events? Is there a phosphorelay, akin to that identified for endospore formation in B . subtilis (Hoch, 1993), that coordinates cell cycle events? Is there a factor laid down at the site of division that marks the fate of the new pole? These and other hypotheses that have been put forward to explain temporal and spatial gene regulation are currently being tested.

c. %o-Component Systems and Cell Cycle Regulation. The CtrA protein is a global cell cycle regulator that has been linked to at least five distinct cell cycle events, including the activation and repression of early flagellar gene expression, DNA replication, cell division, and DNA methylation (Quon et al., 1996). Both CtrA, and FlbD, the transcriptional activator of class 111 flagellar genes, are members of the response regulator group of the bacterial signal transduction superfamily (for reviews see Stock er al., 1989; Parkinson and Kofoid, 1992). These types of proteins typically have a cognate histidine kinase. In the two-component signaling paradigm, the histidine kinase, in response to an input signal, autophosphorylates at a conserved histidine residue and then transfers its phosphate to a conserved aspartate residue on its cognate response regulator. In many cases, the phosphorylated response regulator functions to activate or repress transcription by binding to specific sites in the promoter region of its target genes. There are also response regulators that lack a DNA-binding domain, such as CheY (Stock et al., 1985), that modulate specific physiological responses of the cell. It has been estimated that in a single bacterium at least 50 of these systems operate, controlling a variety of adaptive responses ranging from monitoring changes in nutrient concentration, pH, osmolarity, and temperature, to the transmission of signals required for sporulation and fruiting body formation (Stock er al., 1989).

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The cognate histidine kinases for CtrA and FlbD have not yet been identified. However, the genes for several histidine kinases have been isolated, and strains with mutant alleles of one of these, pleC, display pleiotropic @le) defects in polar morphogenesis, including the assembly of paralyzed flagella and a failure to form stalks (Ely et a l . , 1984; Sommer and Newton, 1989). The PleC protein is capable of autophosphorylation in vitro (Wang et a l . , 1993), suggesting that PleC regulates the expression of genes involved in polar organelle development through the phosphorylation of key regulatory proteins. Pseudoreversion analysis of temperature-sensitive mutations in pleC identified three new cell division genes: d i d , divK, and divL (Sommer and Newton, 1991). One of these genes, divJ, like pleC, encodes a protein homologous to histidine protein kinases (Ohta et a l . , 1992). Another gene identified in this screen, divK, encodes an essential single-domain response regulator, which lacks a DNA-binding domain (Hecht et al., 1995). It has been proposed that the DivJ, DivK, and PleC proteins act in a signal transduction pathway in which the DivJ and PleC histidine kinases phosphorylate DivK to control cell division, stalk formation, and flagellar motility. However, neither the signals nor the target genes for this pathway have yet been identified. These findings suggest that an extensive phosphorelay network may be involved in coordinating the timing of events in the C. crescentus cell cycle. New components of this network, such as the pleD gene, encoding a protein homologous to members of the response regulator family (Hecht and Newton, 1995), continue to be added to the growing list. In all cases, the challenge is to identify the target genes, to determine the role of phosphorylation in these interactions, and to determine how the timing of these events is linked to cell cycle progression. In Bacillus, multiple environmental, metabolic, and cell cycle cues control the phosphorylation cascade that initiates sporulation (Hoch, 1993). A seminal question that these studies in C. crescentus will help to address pertains to the intrinsic signals responsible for activating the phosphorelay network during the progress of the normal cell cycle.

2. Genes Involved in DNA Metabolism Several C . crescentus genes directly involved in DNA replication have been identified to date, including genes encoding DnaA, involved in replication initiation (Zweiger and Shapiro, 1994), DnaN, encoding the homolog of the sliding clamp protein that imparts processivity to the DNA polymerase I11 holoenzyme (Roberts and Shapiro, 1996), and DnaX, a component of the complex that loads the sliding clamp onto the DNA (Winzeler and Shapiro, 1996). Both DnaN and DnaX are involved in the elongation phase of replication. GyrB is a subunit of DNA topoisomerase 11, likely also to be involved in the elongation process (Rizzo et al., 1993). Another gene, labeled dnaC but with no known relation to the E . coli gene of the same name, has also been isolated but has yet to be sequenced (Ohta et a l . , 1990). Other genes with auxiliary roles in replication

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have also been found, most notable including the chaperones encoded by the dnaKJ operon (Gomes et al., 1990; Avedissian et a l . , 1995), grpE (Roberts et al., 1995), and the groESL operon (Avedissian and Gomes, 1995). Transcriptional analysis of the grpE and groESL operons demonstrated that at physiological growth temperatures they are expressed constitutively (Roberts et al., 1995; Avedissian and Gomes, 1995). The striking observation is that most of the remainder of these genes, including dnaN, dnaX, gyrB, dnaC, and dnaKJ, are induced to maximal transcription around the time of transition from swarmer to stalked cell, with the elevated levels of transcription generally extending through the stalked cell phase (Ohta et al., 1990; Rizzo et al., 1993; Avedissian et al., 1995; Roberts and Shapiro, 1996; Winzeler and Shapiro, 1996). The dnuA gene shows a slightly different pattern of expression, peaking just before stalked cell phase (Zweiger et al., 1994). This is not unexpected, given the role of this gene in initiation while the rest of the genes function in elongation (where a function is known). Each of the genes also exhibits a lower level of transcription during the predivisional and swarmer phases of the cycle (between 10 and 50% of the maximal level), consistent with the observation that all of the factors necessary for plasmid replication do exist in the swarmer cell, in spite of the absence of chromosomal DNA replication (Marczynski et al., 1990). Unlike many other C . crescentus genes, transcription of this family of DNA replication genes does not appear to be modulated by the response regulator protein CtrA (R. Roberts and L. Shapiro, unpublished observation). For several of these temporally regulated genes (including d n d , d n d , dnaX, gyrB, and dnaK), the transcriptional start sites have been mapped, allowing comparison of their promoter sequences. In general, these regions did not show strong conservation to the deduced consensus a70 promoter sequence from C . crescentus (Malakooti et al., 1995). Comparison of these promoters demonstrated little conservation in the - 10 region (only a single A was invariant), but sequences at approximately -20 were extensively conserved, as were sequences in the -35 region. Closer analysis revealed another significant element: an 8-mer with well-conserved sequence at six of the eight nucleotides. However, the position of this 8-mer was not conserved, located at variable sites between -50 and -35 for d n d , dnaN, dnaX, and gyrB, but at + 15 or dnaKJ (Roberts and Shapiro, 1996; Winzeler and Shapiro, 1996). Current efforts are underway to address the roles of these conserved sequences in both the transcriptional activity of these promoters and in their induction at the time of swarmer-to-stalked cell transition. It is tempting to speculate that these conserved motifs might function as recognition sites for a regulatory protein(s), including perhaps yet another member of the response regulator family. As for spatial localization of the gene products involved in replication, the product of the dnaK gene has been shown to segregate preferentially to the stalked cell (Reuter and Shapiro, 1987), and transcription of the gyrB gene occurs predominantly in the stalked cell pole of the predivisional cell (Rizzo et

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al., 1993). These results are in agreement with the model that replication factors should preferentially accumulate in the stalked cell compartment, since DNA replication reinitiates immediately following division only in the stalked cell. However, neither functional significance of DnaK or GyrB locational nor spatial localization of other replication factors has yet been addressed. In addition to components of the DNA replication machinery, another important family of DNA metabolism proteins consists of the DNA methylases. One level proposed for the regulation of chromosomal replication in E. coli is by recognition of methylated sites in the origin (at the sequence GATC) and preferential sequestration by the SeqA protein, to block premature reinitiation (Lu et al., 1994; Brendler et al., 1995). When the adenines in GATC sites become hemimethylated on replication, they are normally remethylated very quickly by the Dam methylase, which plays a role not only in the regulation of replication initiation but also in repair of mutations (reviewed in Palmer and Marinus, 1994). However, like many bacteria, C. crescentus lacks a homolog of the dam gene, and GATC sites are not methylated (Barbeyron et al., 1984). However, another gene responsible for the methylation of the C. crescentus chromosome has been found (Zweiger et al., 1994). This gene, ccrM, encodes a DNA methyltransferase that recognizes the sequence GAnTC, methylating the adenine residue. No cognate restriction enzyme with GAnTC as its recognition sequence has been identified, and in addition, this methylase was shown to be essential, a first among bacterial DNA methylases (Stephens et al., 1996). Interestingly, CcrM appears to be a widely conserved methylase among many members of the a subdivision of the purple bacteria, where C. crescentus is phylogenetically placed, suggesting perhaps that its function is of general significance (Stephens et al., 1996). Analysis of the methylation state of the C. crescentus chromosome during the cell cycle demonstrated that on replication, the strands become hemimethylated, but unlike the rapid remethylation of E. coli GATC sites by Dam, the C . crescentus hemimethylated state persists through the stalked cell phase until very near to the time of completion of replication. The chromosomes are then both fully remethylated close to the time of cell division (Zweiger et al., 1994), because expression of the ccrM gene is limited to the predivisional cell. The promoter sequence of this gene resembles that of the flagellar class I1 gene consensus (Zweiger el al., 1994; Stephens et al., 1995a). It has been shown that, also like class I1 flagellar genes, the timing of transcription of the ccrM gene is regulated by the cell cycle-responsive transcriptional regulator CtrA (Quon et al., 1996). Following remethylation of the newly replicated chromosomes in the predivisional cell, the CcrM protein is rapidly degraded, most likely by the C. crescentus Lon protease homolog, thus excluding methylation activity from the daughter cells (Wright et al., 1996). Thus, the genes involved in DNA metabolism show two patterns of temporally regulated transcription: components of the DNA replication machinery are transcribed maximally during chromosomal replication, and the CcrM meth-

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yltransferase is transcribed later in the cycle; patterns that fit well with the times at which these gene functions are required. Of crucial importance is the identification of factors that set the timing of transcription of these genes, with the ultimate objective being to characterize how these events are tied to progression through the cell cycle. The factor regulating expression of the methylase, like the flagellar cascade, appears to be CtrA. However, transcription of DNA replication factors appears to be independent of CtrA, and therefore may represent the targets of another cell cycle signaling pathway.

C. Targeting of Proteins to Specific Locations within the Cell

The selective transcription of late flagellar genes in the C. crescentus swanner compartment of the predivisional cell is one mechanism accounting for the asymmetric positioning of their gene products in the predivisional cell. This alone, however, is inadequate to explain how the flagellum, which requires approximately 50 gene products for its assembly and function, is specifically placed at the pole opposite the stalk. Clearly, other mechanisms for the asymmetric distribution of proteins must be at work. One such mechanism, protein localization, functions in the generation of asymmetry in the dividing C . crescenzus cell and may play a general role in bacterial physiology. Much of our understanding of protein localization during the C . crescentus cell cycle has come from expression and immunolocalization studies of the chemoreceptor protein McpA. In many bacteria, chemotactic stimuli are detected with chemoreceptors, known as methylaccepting chemotaxis proteins (MCPs). MCPs are integral membrane proteins with two membrane-spanning domains (Parkinson, 1993). When the periplasmic domain of the MCP interacts with a ligand (an attractant or repellant), a conformational change is induced that mediates the transmission of a chemotactic signal to the flagellar motor via a phosphorelay system. The signal to the flagella facilitates net movement of the bacterium toward or away from high concentrations of the ligand. In C . crescentus, the chemoreceptor protein McpA is synthesized in the predivisional cell but appears only in the progeny swanner cell on cell division (Alley et al.,1991, 1992). lmmunoelectron microscopy revealed that wild-type McpA is present only at the flagellated pole of the predivisional cell (Alley e f al., 1992). On division, McpA remains at that pole in the progeny swarmer cell, then disappears later in the cell cycle when the swarmer cell differentiates into a stalked cell. The polar location of the chemoreceptor is not unique to C. crescentus, or polarly flagellated bacteria. MCP polar localization also occurs in E . coli cells, which have flagella distributed over their entire surface (Maddock and Shapiro, 1993). Polar localization of the MCPs to the E . coli cell pole requires the two cytoplasmic components of the signal transduction pathway, CheA and Chew, which have been shown to form a ternary complex with the chemorecep-

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tor in vitro (Gegner et al., 1992). Using immunogold electron microscopy and indirect fluorescence light microscopy, clusters of all three components have been observed primarily at the E. coli cell pole (Maddock and Shapiro, 1993), suggesting that subcellular localization of the MCPs occurs in a ternary complex with cytoplasmic CheA and Chew proteins. When an E. coli chemoreceptor gene is expressed in C. crescentus, the E . coli protein is also targeted to the swanner cell pole (Alley et al., 1992), suggesting that the C . crescentus CheA and Chew proteins are able to interact with the E . coli MCP and direct the protein complex to the cell poles. The E . coli chemoreceptor interacts with CheA and Chew via a 45-amino acid, highly conserved domain in its cytoplasmic portion (Liu and Parkinson, 1991). When this domain is deleted, the chemoreceptor is no longer polarly localized (Alley et al., 1992). Since the C. crescentus McpA protein also has the highly conserved domain, ternary complex formation with CheA and Chew may also contribute to its targeted localization. As yet unanswered questions relate to the mechanism by which the bacterial chemoreceptor is targeted to and maintained at the cell pole. Are there specific polar transport systems? Do large macromolecular assemblies of chemotaxis proteins hinder free membrane diffusion away from the pole? Further elucidation of discrete domains within the McpA protein that are required for both protein interactions and localization, as well as the identification of other proteins targeted to the cell pole, will certainly shed light on the mechanisms of transport and retention. Protein localization is also required for the initiation of flagellar biogenesis at the swanner pole of the C. crescentus predivisional cell. The assembly of the C. crescentus polar flagellum occurs in a cell-proximal to cell-distal order, beginning with insertion into the membrane of the early flagellar structural proteins. A key event in determining the polar site of flagellar assembly may be structural markers left at the site of the previous division plane, although no such markers have been identified. This structural “memory” might dictate the positioning of the earliest protein components of the flagellum at the cell pole. Once in place, these flagellar components could function as a nucleation site for the targeting and progressive assembly of proteins comprising the more distal substructures. The FliF protein is the structural subunit of the flagellar MS-ring, which is one of the earliest substructures of the flagellum to be assembled (Fig. 7). It has been shown that in C. crescentus, FliF is synthesized in the early predivisional cell, but specifically targeted to the swanner cell compartment (Jenal and Shapiro, 1996). This is most likely a consequence of protein localization and not localized transcription of theJEiF gene, because the synthesis of FliF takes place before the swarmer and stalked cell compartments are formed. It is likely that the FliF protein itself is sequestered to the swanner pole of the predivisional cell, where it is assembled into the MS-ring of the flagellar motor. Another flagellar protein that is synthesized early in the predivisional cell but is observed only in the progeny swarmer cell is the 29-kDa flagellin protein (Loewy et al., 1987).

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Compartmentalization of this filament subunit occurs even when flagellar morphogenesis is blocked, suggesting that the 29-kDa protein can be sorted to the swarmer compartment independently of other flagellar components. In addition to the single polar flagellum, pili are also specifically assembled at the C . crescentus swarmer cell pole (Schmidt, 1966; Sommer and Newton, 1988). This assembly occurs in the swarmer cell shortly before cell division. The pilin subunits are drawn from an intracellular pool of unassembled cytoplasmic pilin that is distributed equally between the swarmer and stalked cell compartments (Smit, 1987). This indicates that the localized assembly of pili at the swarmer cell pole requires the polar targeting of the assembly machinery, rather than compartmentalization of the pilin subunits. In M . xanthus pili are also localized at the cell pole (MacRae et al., 1977; Kaiser, 1979). Mutations that result in a complete loss of piliation were mapped to a M . xanthus region encoding at least three genes sharing homology with components of the type IV pilus biogenesis pathway (Wu and Kaiser, 1995). These pili-minus mutants are also defective in social gliding motility, the type of motility associated with cells in high density and close contact. It has been suggested that the M . xanthus pili may be involved in the intimate cell-cell interactions required for social motility (Wu and Kaiser, 1995) and it is possible that their polar localization could mediate the end-to-end interactions of aggregating cells. While the chemoreceptors, as well as proteins involved in flagellar and pili biogenesis, are specifically targeted to the swarmer cell, there is also evidence for the localization of proteins to the C . crescentus stalked cell. Two heat shock proteins, DnaK and Lon, are synthesized in the C . crescentus predivisional cell but show a bias to the stalked cell progeny (Reuter and Shapiro, 1987). Why these proteins are specifically targeted to the stalked cell is not known. It is possible that they play a role in the initiation of DNA replication, which the stalked cell, but not the swarmer cell, immediately undergoes following cell division. The targeting of Lon, an ATP-dependent protease, to the stalked cell may have other important ramifications, since this protein appears to play a role in the proteolysis of an essential DNA methyltransferase (see below). Protein localization also plays an important role in the establishment of cell type-specific gene expression during sporulation in B . subtilis. During sporulation, a polar septum is formed that partitions the developing cell into two unequal cellular compartments: the forespore and the mother cell (Fig. 3). As described above, cell type-specific gene expression in these compartments is accomplished through the differential activation of distinct genes by a series of developmental stage-specific RNA polymerase sigma factors. The activation of one of these factors, uF,in the forespore requires dephosphorylation of the SpoIIAA inhibitor by the serine phosphatase SpoIIE (Duncan et al., 1995). SpoIIE is an integral membrane protein that becomes localized to the polar septum during sporulation (Arigoni et al., 1995). What is the effect of localizing SpoIIE to the polar septum? One intriguing hypothesis is that localization displays SpoIIE equally on

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both the forespore and mother cell sides of the polar septum. However, since the forespore has a smaller volume than the mother cell (approximately one-fifth the size), this localization event would increase the concentration of SpoIIE in the forespore. The higher effective concentration of phosphatase would result in more UF activity in the forespore than in the mother cell. In this model, the developing cell has an elegant yet simple mechanism to link, via protein localization, asymmetric cell division to the pathway controlling cell-specific gene transcription. The process by which these asymmetrically localized bacterial cellular components are sequestered to specific regions of the bacterial cell during development is not understood. One hypothesis explaining the polar targeting of the flagellar and chemotaxis proteins in C. crescentus is that proteins are inserted directly into a site at the cell pole via a polar secretory signal. Complex formation with other proteins may then prevent the lateral diffusion of the proteins away from the site. Another possibility is that proteins are randomly inserted into the membrane and selectively retained at one of the poles. Both models require an identification mechanism by which the cell pole is somehow marked. The nature of such a marker, or targeting patch, is unknown. However, since the bacterial cell poles originate as the site of cell division, as mentioned above, a polar marker could be laid down at that time which could serve as a targeting or organizational center at the new poles. During the earliest stages of sporulation in B . subtilis, the SpoIIE protein is localized near both poles of the developing sporangia, and only later in development becomes sequestered at the position of the polar septum (Arigoni ef al., 1995). Since SpoIIE becomes localized only to one pole at the asymmetrically positioned polar septum, as opposed to the symmetrically placed septum during vegetative growth, it has been suggested that polar markers may exist that are either masked or degraded at specific times during development. The asymmetric distribution of proteins is an important component of the cell differentiation events in C. crescentus and most likely for bacteria in general. Early observations of bacterial polar structures implied the existence of protein localization, but in spite of this little is known about the mechanisms of localization. What has emerged, however, is the finding that the bacteria1 cell is in fact highly organized, with discrete functional domains located at specific sites in the cell. This discovery has pointed the way to the importance of an understanding of how these domains are created and the mechanisms involved in targeting proteins to these specific subcellular sites.

D. Specific Degradation of Proteins at Discrete Times during the Cell Cycle

Immunoelectron microscopic examination of proteins during the C. crescentus cell cycle or during sporulation in B . subtilis has shown that localized proteins

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appear and disappear at specific times during the cell cycle. As discussed earlier, the temporal control of gene expression and localization of the protein products are two mechanisms that help explain these temporal and spatial appearances of specific proteins. Because developmental progression requires not only the localization, but also the degradation, of various proteins, it is perhaps not surprising that proteolysis is emerging as another important regulatory mechanism in the C . crescentus cell cycle as well as the developmental programs of other bacteria. As described above, the C . crescentus McpA chemoreceptor is synthesized in the predivisional cell and targeted to the swarmer cell pole. When the swarmer cell differentiates into a stalked cell McpA is proteolytically turned over (Alley et al., 1993). The site that targets McpA for proteolysis appears to be a discrete domain located in its carboxy terminus. When 14 amino acids are removed from the C terminus of McpA, the mutant protein is not degraded when swarmer cells differentiate into stalked cells (Alley et al., 1993). Consequently, the chemoreceptors are now found at both poles of the predivisional cell; the presence of McpA at the swarmer pole is due to de novo synthesis and targeting of McpA, and the stalked pole carries the old McpA that was not degraded when the flagellum was released and a new stalk was assembled in its place. A similar localization pattern is seen when the wild-type McpA is overexpressed and not fully degraded in C . crescentus, indicating that the proteolytic machinery can be saturated. While proteolysis contributes to the asymmetric distribution of the McpA, it does not appear to be involved in polar targeting, since McpA is still properly targeted to the cell pole when the proteolytic signal domain is deleted. Proteolysis during the swarmer-to-stalked cell transition extends to other C . crescentus proteins. The cell cycle synthesis and targeting pattern of FliF, encoding the flagellar MS-ring, is similar to the McpA protein, in that both proteins are synthesized in the predivisional cell and specifically targeted to the swarmer cell progeny. The FliF protein is also subject to proteolytic degradation during the swarmer-to-stalked cell transition (Jenal and Shapiro, 1996). The turnover determinant, like that of the McpA, appears to be within the C terminus of the FliF protein; FliF derivatives lacking 26 amino acids at their C terminus are stable throughout the cell cycle (Jenal and Shapiro, 1996). One issue that arose from these studies was whether the FliF localized to the swarmer cell compartment of the predivisional cell was a result of protein targeting to the swarmer cell or due to proteolytic removal at other sites in the cell. This was addressed experimentally by analyzing the distribution of newly synthesized FliF protein in the predivisional cell. It was found by pulse-chase labeling that both wild-type FliF and a stable FliF C terminal deletion derivative synthesized in the predivisional cell segregated preferentially to the swarmer compartment (Jenal and Shapiro, 1996). Stalked pole-specific proteolysis, therefore, does not contribute to asymmetric distribution of the FliF protein in the predivisional cell. It was concluded that two independent mechanisms do contribute to the asymmetric distribution of FliF in the predivisional cell: degradation of the old FliF during the swarmer-to-

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stalked cell transition and targeting of newly synthesized FliF to the swarmer pole of the predivisional cell (Jenal and Shapiro, 1996). Not all C . crescentus flagellar components are degraded during the swarmerto-stalked cell transition. For example, FlgH, the protein subunit of the flagellar basal body L-ring, is stable throughout the C. crescentus cell cycle, as is FliL, a protein required for flagellar rotation (Jenal et al., 1994; Mohr et al., 1996). This raises a number of interesting questions. What targets certain proteins for their destruction during the C . crescentus cell cycle? What proteases are responsible and how are they regulated? The identification of discrete domains required for proteolysis of the McpA and FliF proteins has provided a clue to the mechanism of turnover for these proteins. The C terminus of both McpA and FliF may be recognized by a modification enzyme, as in the case of ubiquination in the eukaryotic cell (Hochstrasser, 1995), which marks the protein for subsequent proteolytic degradation. Alternatively, the site in the C terminus might itself be the recognition site for a specific protease. Is the protease or the modifying enzyme present only in a certain cell type? At this time, the protein(s) responsible for the degradation of the McpA and FliF proteins during the swarmer-tostalked cell transition have not been identified. There is evidence, however, that at least one protease, Lon, is targeted to the C . crescentus stalked cell, and that this protein plays a role in proteolysis of the DNA methyltransferase, CcrM. As mentioned earlier, the expression of the essential site-specific DNA methyltransferase, CcrM, is tightly restricted to the C . crescentus predivisional cell (Zweiger et al., 1994; Stephens et al., 1996). The CcrM protein is present only in predivisional cells, coincident with remethylation of the newly replicated chromosomes, and is rapidly degraded late in the cell cycle, just prior to cell division. There is also evidence that a cell division protein, FtsZ, is likewise degraded late in the cell cycle, perhaps in a cell type-specific manner (Quardokus et al., 1996). The protease responsible for CcrM degradation is likely to be a homolog of the ATP-dependent protease Lon (Wright et al., 1996). A null mutation in the C . crescentus lon gene results in the presence of CcrM in all cell types and the phenotype of lon mutant cells is similar to that of cells in which the wild-type ccrM gene is constitutively expressed (Zweiger et al., 1994; Wright et al., 1996). Moreover, it has been demonstrated that the chromosomal DNA in the Lon mutant is persistently methylated on both strands throughout the cell cycle. It should be pointed out, however, that it has not been demonstrated in vitro that the methyltransferase is the direct substrate for Lon and therefore its proteolytic role may be indirect. The mechanism responsible for defining the narrow window of CcrM activity during the cell cycle appears to be a finely choreographed balancing act between CcrM synthesis and degradation. The Lon protein, in contrast to CcrM, is expressed throughout the cell cycle at a relatively constant level (Wright et al., 1996). It is likely, therefore, that CcrM turnover is initiated as soon as CcrM is synthesized. Since the rate of CcrM synthesis peaks in early predivisional cells,

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its synthesis rate at this point would be expected to be greater than the rate of its degradation, and therefore the CcrM protein accumulates. As transcription of ccrM decreases in late predivisional cells, the proteolytic rate becomes predominant and CcrM levels diminish such that only negligible levels of CcrM can be detected in the two progeny cells. Why might the rapid Lon-dependent degradation of CcrM just prior to cell division be important? Since the Lon protease is preferentially targeted to the stalked cell compartment of the predivisional cell (Reuter and Shapiro, 1987), degradation of CcrM in this compartment could prevent the premature remethylation of the origin region in the progeny stalked cell to control replication; persistent methylation has been seen to result in inappropriate reinitiation of chromosomal replication (Zweiger et al., 1994). Proteolysis during the C. crescentus cell cycle cannot be attributed solely to Lon. The cell cycle degradation patterns of the FliF and McpA proteins are not affected in the lon mutant strain (M. R. K. Alley, U. Jenal, and L. Shapiro, unpublished results), indicating that other proteases must be at work. The identification of the genes encoding another ATP-dependent protease, Clp, in C . crescentus (U. Jenal and L. Shapiro, unpublished results) should enhance our understanding of proteolysis in this bacterium, and its role in the regulation of the cell cycle. Proteolysis by Lon may also play a role in the programmed development of other bacteria. Two lon genes have been identified in M . xanthus. The lonV gene is essential for vegetative growth, while the other, bsgA (lonD), is required for development (Tojo et al., 1993a, b; Gill et al., 1993). The developmental phenotypes associated with the bsgA mutation include abnormal developmental gene expression, as well as a failure to form fruiting bodies and myxospores (Gill et al., 1993; Tojo et al., 1993b). Substrates for the lonV and bsgA gene products have not yet been identified, and therefore their specific roles in M. xanthus growth and development have not been established. Furthermore, a gene, lon-I, encoding a Lon protease homolog in B . subtilis, has been identified (Riethdorf et al., 1994; Schmidt et al., 1994). A mutation in Lon-1 results in the inappropriate expression of genes under the control of uG,a cell type-specific sigma factor controlling the transcription of genes in the forespore compartment of the developing sporangium. It has been suggested that the Lon-1 gene product plays a role in preventing the accumulation of active uG,perhaps through direct proteolysis of the UG protein (Schmidt et al., 1994). We are just beginning to appreciate the role of proteolysis as a regulatory mechanism in bacterial development. In C. crescentus, future studies aimed at determining whether the CcrM methyltransferase is a direct substrate for Lon, understanding the role of proteolysis in the temporal and spatial control of the cell division protein FtsZ, and identifying other proteins targeted for degradation will certainly shed light on the role of selective proteolysis in the cell cycle. That the cell cycle-controlled proteolysis of the FliF and McpA proteins is not depen-

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dent on Lon points to the existence of additional, as yet undiscovered, proteases in this bacterium that play significant roles in cell cycle events. Advances in the understanding of the critical role of the timed destruction of cyclins in regu lating the eukaryotic cell cycle (reviewed in Gottesman and Maurizi, 1992; Hochstrasser, 1995) point to the importance of proteolysis as an essential regulatory component during development. A clearer view of the role of targeted protein destruction in the developmental pathways of prokaryotes should reveal commonalities, as well as a better understanding of the control of development in many organisms.

IV. Conclusions and Future Perspectives One of the current frontiers in biology is the study of development: how does a multipotent stem cell give rise to a spectrum of specialized cell types? Two fundamental processes in such cellular restructuring are the ability to localize components of the cell in a spatially directed manner, and the capacity to regulate gene expression temporally as a function of the cell cycle. These fundamental processes play a role in differentiation not only in higher eukaryotes, but also in a variety of bacteria. Development takes on many forms among the bacteria, from a natural part of the cell cycle as seen in C . crescentus to the sporulation pathways initiated by stress in a variety of other organisms. Developmental programs are evident even in the prototypical bacterium of molecular biology, E . coli, as it responds to stress and enters or exits the stationary phase. Dissection of the factors driving these programs in several different bacteria is well underway, revealing in each case a complex network of signals temporally and spatially controlling gene expression. As the components regulating development in bacteria are understood at a molecular level, this fundamental knowledge should illuminate basic principles that extend beyond the prokaryotic kingdom to aid in the understanding of developmental processes in a variety of other organisms.

Acknowledgments We thank Janine Maddock for providing the electron micrograph of C . crescentus shown in Fig. 1, and Yves Brun and Bert Ely for communicating unpublished data. We also thank Keith Chater, Alan Grossman, Dale Kaiser, Roberto Kolter, Sharon Long, and Dan Wall for critical reading of the manuscript, and members of the Shapiro laboratory for helpful discussions. R.C.R. was supported by American Cancer Society postdoctoral fellowship PF-3831, and C.D.M. was supported by American Cancer Society postdoctoral fellowship PF-3941. Work in the Shapiro laboratory was supported by Grants GM32506 and GM51426 from the National Institutes of Health.

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Shapiro, L., Agabian-Keshishian, N., and Bendis, 1. (1971). Bacterial differentiation. Science 173, 884-892. Sharpe, M. E., and Enington, 1. (1995). Postseptational chromosome partition in bacteria. Proc. Natl. Acud. Sci. U.S.A. 92, 8630-8634. Sheffery, M., and Newton, A. (1981). Regulation of periodic protein synthesis in the cell cycle: Control of initiation and termination of flagellar gene expression. Cell 24, 49-57. Shimkets, L. J. (1990). Social and developmental biology of the Myxobacteria. Microbiot. Rev. 54, 473-501. Shimkets, L. I., and Rafiee, H. (1990). CsgA, an extracellular protein essential for Myxococcus xanthus development. J . Bacteriol. 172, 5299-5306. Siegele, D. A,, and Kolter, R. (1993). Isolation and characterization of an Escherichia coli mutant defective in resuming growth after starvation. Genes Dev. 7, 2629-2640. Siegele, D. A., Almirh, M., and Koltet, R. (1993). Approaches to the study of survival and death in stationary phase Escherichiu coli. In “Starvation in Bacteria” (S. Kjelleberg, ed.), pp. 151-169. Plenum, New York. Singer, M., and Kaiser, D. (1995). Ectopic production of guanosine penta- and tetraphosphate can initiate early developmental gene expression in Myxococcus xanthus. Genes Dev. 9, 16331644. Smit, J. (1987). Localizing the subunit pool for the temporally regulated polar pili of Caulobacter crescentus. J. Cell Biof. 105, 1821-1828. Solomon, J. M., Magnuson, R., Srivastava, A., and Grossman, A. D. (1995). Convergent sensing pathways mediate response to two extracellular competence factors in Bacillus subtilis. Genes Dev. 9, 547-558. Sommer, I. M., and Newton, A. (1988). Sequential regulation of developmental events during polar morphogenesis in Caulobacter crescentus: Assembly of pili on swarmer cells requires cell separation. J. Bacteriol. 170, 409-415. Sommer, J. M., and Newton, A. (1989). Turning off flagellum rotation requires the pleiotropic gene pleD :pleA, pleC, and pleD define two morphogenic pathways in Caulobacter crescentus. J. Bacferiof. 171, 392-401. Sommer, J. M., and Newton, A. (1991). Pseudoreversion analysis indicates a direct role of cell division genes in polar morphogenesis and differentiation in Cuulobacter crescentus. Genetics 129, 623-630. Spira, B . , Silberstein, N., and Yagil, E. (1995). Guanosine 3’,5’-bispyrophosphate (ppGpp) synthesis in cells of Escherichia coli starved for Pi. J. Bacteriol. 177, 4053-4058. Stephens, C. M., and Shapiro, L. (1993). An unusual promoter controls cell-cycle regulation and dependence on DNA replication of the CaulobacterfliLM early flagellar operon. Mol. Microbiol. 9, 1169-1179. Stephens, C. M., Zweiger, G., and Shapiro, L. (1995a). Coordinate cell cycle control of a Caulobucter DNA methyltransferase and the flagellar genetic hierarchy. J. Bacteriol. 177, 16621669. Stephens, C., Jenal, U., and Shapiro, L. (1995b). Expression of cell polarity during Caulobacter differentiation. Semin. Dev. Biol. 6, 3- 1 1. Stephens, C. M., Reisenauer, A., Wright, R., and Shapiro, L. (1996). A cell cycle-regulated bacterial DNA methyltransferase is essential for viability. Proc. Nail. Acud. Sci. U.S.A. 93, 12 10- 12 14. Stock, A , , Koshland, D. E. J., and Stock, J. (1985). Homologies between the Saimonefla typhimurium CheY protein and proteins involved in the regulation of chemotaxis, membrane protein synthesis and sporulation. Proc. Natl. Acud. Sci. U.S.A. 82, 7989-7993. Stock, J. B., Ninfa, A. J., and Stock, A. M. (1989). Protein phosphorylation and regulation of adaptive responses in bacteria. Microbiol. Rev. 53, 450-490. Strauch, M. A., and Hoch, J. A. (1993). Transition-state regulators: Sentinels of Bacillus subrilis post-exponential gene expression. M o l . Microbiol. 7, 337-342.

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Stukenberg, P. T., ’hmer, J., and O’Donnell, M. (1994). An explanation for lagging strand replication: Polymerase hopping among DNA sliding clamps. Cell 78, 877-887. Swoboda, U. K., Dow, C. S., and Vitkovic, L. (1982). Nucleoids of Cuulobucfer crescenfus CB15. J. Gen. Microbiol. 128, 279-289. Tan, H., and Chater, K. F. (1993). Two developmentally-controlled promoters of Szreptomyces coelicolor A3(2) that resemble the major class of motility-related promoters in other bacteria. J. Bacteriol. 175, 933-940. Toal, D. R., Clifton, S. W., Roe, B. A , and Downard, J. (1995). The esg locus of Myxococcus xanthus encodes the Ela and E I P subunits of a branched-chain keto acid dehydrogenase. Mol. Microbiol. 16, 177-189. Tojo, N., Inouye, S., and Komano, T. (1993a). Cloning and nucleotide sequence of the Myxococcus xunfhus lon gene: Indispensability of Ion for vegetative growth. J . Bucferiol. 175, 22712277. Tojo, N., Inouye, S., and Komano, T. (1993b). The lonD gene is homologous to the lon gene encoding an ATP-dependent protease and is essential for the development of Myxococcus xunthus. J. Bucteriol. 175, 4545-4549. Udo, H., Munoz-Dorado, J., Inouye, M., and Inouye, S. (1995). Myxococcus xanzhus, a gramnegative bacterium, contains a transmembrane protein serinelthreonine kinase that blocks the secretion of beta-lactamase by phosphorylation. Genes Dev. 9, 972-983. Ueda, K., Miyake, K., Horinouchi, S., and Beppu, T.(1993). A gene cluster involved in aerial mycelium formation in Srrepfomyces griseus encodes proteins similar to the response regulators of two-component regulatory systems and membrane translocators. J . Bucferiol. 175, 2006-20 16. van Helvoort, J. M., and Woldringh, C. L. (1994). Nucleoid partitioning in Escherichiu coli during steady-state growth and upon recovery from chloramphenicol treatment. Mol. Microbiol. 13, 577-583. Van Rhijn, P., and Vanderleyden, J. (1995). The Rhizobium-plant symbiosis. Microbiol. Rev. 59, 124- 142. van Sinderen, D., Luttinger, A,, Kong, L., Dubnau, D., Venema, G., and Hamoen, L. (1995). comK encodes the competence transcription factor, the key regulatory protein for competence development in Bacillus subtilis. Mol. Microbiol. 15, 455-462. Van Way, S. M., Newton, A , , Mullin, A. H . , and Mullin, D. A. (1993). Identification of the promoter and a negative regulatory element, ffr4, that is needed for cell cycle timing of f7iF operon expression in Cuulobucfer crescentus. J. Bucteriol. 175, 367-376. Wadsworth, P. (1993). Mitosis: Spindle assembly and chromosome motion. Curr. Opin. Cell Biol. 5, 123-128. Wang, S. P., Sharma, P. L., Schoenlein, P. V., and Ely, B. (1993). A histidine protein kinase is involved in polar organelle development in Cuulobucfer crescentus. Proc. Nufl. Acud. Sci. U.S.A. 90, 630-634. Weiner, L., and Model, P. (1994). Role of an Escherichiu coli stress-response operon in stationary-phase survival. Proc. Nufl.Acud. Sci. U.S.A. 91, 2191-2195. Weinstein, M., Lois, A. F., Monson, E. K . , Ditta, G. S., and Helinski, D. R. (1992). Isolation of phosphorylation-deficient mutants of the Rhizobium meliloti two component regulatory protein, FixJ. Mol. Microbiol. 6, 2041-2049. Wemer, D. (1992). Physiology of nitrogen fixing legume nodules: Compartments and functions. In “Biological Nitrogen Fixation” (G. Stacey, R. H. Bums, and H. J. Evans, eds.), pp. 39943 1. Chapman and Hall, New York. Willey, J., Santamaria, R., Guijarro, J., Geistlich, M.. and Losick, R. (1991). Extracellular complementation of a developmental mutation implicates a small sporulation protein in aerial mycelium formation by S. coelicolor. Cell 65, 641-650. Willey, J., Schwedock, J., and Losick, R. (1993). Multiple extracellular signals govern the pro-

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Gametes and Fertilization in Flowering Plants Darlene Southworth Department of Biology Southern Oregon State College Ashland, Oregon 97520

I. Introduction 11. Male Gametes A. Meiosis to Gametogenesis B . Sperm Structure C. Pollen Tube Growth 111. Female Gametes A. Meiosis to Embryo Sac Formation B . Egg Structure C. Central Cell Structure IV. Double Fertilization A. Fertilization in Vivo B . Fertilization in V i m V. Summary References

1. Introduction The field of flowering plant gametes and fertilization is moving from descriptive ultrastructure to precise descriptions of development, experimental investigations, and molecular approaches, although the molecular and cellular activities that regulate gametogenesis and fertilization are as yet unknown. The purpose of this article is to provide an overview of plant gametes and fertilization, including current research and reviews on structure and function, and to compare plant gametes and fertilization events with those in animals. The process of fertilization in flowering plants is not well known, in part because the gametes are not free in the environment, but are embedded in other cells or tissues. Gametes in flowering plants are relatively sparse compared to gamete production by marine invertebrates or by brown algae. Neither sperm nor eggs are readily released as in vertebrates or ferns. Flowers enclose male and female organs in which meiosis occurs: anthers (“male gonads”) and ovaries (“female gonads”). In anthers, meiosis produces haploid cells that develop into pollen. In contrast to animal meiosis, in which sperm are produced directly, male haploid cells in flowering plants divide, evenCurrent Topics in Developmenral Bioloxy, Val 34 Copyright 0 1996 by Academic Press. Inc. All rights of reproduction in any form reserved.

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tually producing two sperm cells per pollen grain. Ovaries enclose ovules or immature seeds. In each ovule is one meiotic cell surrounded by protective tissues. In the most common pattern, three of the four meiotic products die, and the one surviving haploid cell divides three times. One of the resulting haploid cells becomes the egg. Pollen lands on the stigma, an extension of the ovary, and grows a long pollen tube into an ovule. Sperm are moved through the pollen tube and deposited near the egg. Fertilization, a fusion of gametes, takes place, resulting in a zygote that grows into an embryo. In addition, a second fertilization between a sperm and the central cell also occurs, giving rise to endosperm, a nutritive tissue.

II. Male Gametes Sperm cells in flowering plants are structurally unique among plant cells. Specialized features of sperm cells include the absence of cell walls (a departure from the textbook definition of plant cells as having cell walls), a small cytoplasmic volume, condensed chromatin, microtubule bundles, and a spindle shape with long extensions. Reviews have emphasized particular aspects of flowering plant sperm structure including the cytoskeleton of sperm and generative cells (Palevitz and Tiezzi, 1992), association of two sperm cells with the vegetative nucleus in the male germ unit (Mogensen, 1992), evolution of double fertilization (Knox et al., 1993), and isolation of sperm cells (Russell, 1991; Theunis et a l . , 1991; Chaboud and Perez, 1992).

A. Meiosis to Gametogenesis

1. Pollen Formation Meiosis takes place in anthers of very young flower buds. Following meiosis, haploid cells differentiate into pollen (Bedinger, 1992) (Fig. 1). They develop a complex cell wall, take up nutrients, and differentiate. Each haploid cell divides asymetrically into a larger vegetative cell and a smaller generative cell, both enclosed within the pollen grain wall. The vegetative cell will not divide again, but will develop a long extension that is the pollen tube. The vegetative cell acts as a nurse or Sertoli cell surrounding the generative cell and later the sperm. Contact between the vegetative cell and the generative cell is close, forming a type of cell-cell junction in which membranes closely parallel each other (Southworth, 1992). No membrane bridges link vegetative and generative cells. Although the separation distance between the cell membranes of the vegetative and generative cells is uneven in aldehyde-fixed thin sections (Fig. l), in freezefractured or freeze-substituted pollen, the separation distance is uniform, and

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!

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I

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Fig. 1 Sperm development pathways in flowering plants. Pollen mother cells in the anther (A) divide by meiosis, producing a tetrad of haploid cells (B) that grow and divide (C) to form the generative cell ( G ) and the vegetative cell (v). In bicellular pollen (D), division of the generative cell occurs in the pollen tube (E). In tricellular pollen (F) the generative cell divides before discharge of the pollen; the pollen tube develops with sperm cells already present (G). Both bicellular and tricellular pollen produce a pollen tube containing two sperm cells (H).

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plasma membranes are parallel (Cresti et al., 1987; Southworth et al., 1989a). In freeze-fractured pollen, the surface of the generative cell is indented. Distinctive fracture patterns of parallel ridges occur on the inner vegetative cell membrane at the indentations (Fig. 2). The ridged pattern suggests a junction, either for maintaining cell contact or for passage of materials, but does not precisely resemble tight or gap junctions. Undoubtedly, the vegetative cell is responsible for nourishment of the generative cell and for controlling its cell cycle; however, there is no direct evidence for transfer of molecules between them. The vegetative cell stores food during its development within the anther and later takes up nutrients from the style during pollen tube growth. The generative cell divides to form two sperm (Fig. 1). When pollen grains are released from the anther, they are either bicellular with one vegetative cell and one generative cell or tricellular with one vegetative cell and two sperm cells, depending on the timing of generative cell division. An old terminology inaccurately referred to pollen as “binucleate” or “trinucleate,” and failed to recognize that these nuclei were in separate cells.

2. Development of the Generative Cell The generative cell differentiates and develops structural properties later found also in sperm. After the first pollen mitosis, the wall around the generative cell decreases and in some species disappears completely. The generative cell rounds up and lies in a cavity or pocket within the vegetative cell. A generative cell is surrounded by its own cell membrane appressed against an inner cell membrane of the vegetative cell pocket. Generative cells are closely associated with the vegetative nucleus, often with a cellular extension entwined through the vegetative nucleus but always separated from it by the two cell membra,ies (Mogensen, 1992; Yu and Russell, 1993). Chromatin condenses to make a densely packed nucleus (Fig. 3). In lily, generative cells and sperm form male-specific histones that are variants of histones H2B and H3 (Ueda and Tanaka, 1995a,b). These altered histones are not found in vegetative cell nuclei. No specific function has been demonstrated yet, although they correlate with condensation of chromatin in generative cells and in sperm and suggest a diminished capacity for transcription. The cytoskeleton of generative cells is distinct from that of somatic cells. Immature generative cells are spheroidal with a meshwork of microtubules (Zhou and Yang, 1991). As microtubules elongate, the generative cell becomes ellipsoidal and finally spindle shaped with cytoplasmic extensions at one or both ends of the cell (Fig. 3). Parallel microtubule bundles form a slightly spiraled cage or basket of microtubule bundles in the thin cytoplasmic layer around the nucleus (Del Casino et a l . , 1992; Bohdanowicz et al., 1995). The extensions vary in precise shape and location, but consistently include ends of microtubule bundles.

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Fig. 2 Electron micrographs of the generative cell of Amaryllis belladonna. (A) Thin section across the sperm cell, through the nucleus and microtubule bundles in the cytoplasm. The interface between the generative cell and vegetative cell is convoluted (at right). GN, Generative cell nucleus; VC, vegetative cell. Bar: 0.2 pm. (B) Freeze-fracture face of the vegetative cell membrane appressed to the generative cell. Bar: 0.5 pm. (C) Ridges on the fracture face of the vegetative cell at indentations into generative cell. Bar: 0.2 pm. [Adapted from Southworth er al. (1994), Fig. 1. p. 539; Figs. 5 and 6 , p. 541.1

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Fig. 3 Light micrographs of generative cells. (A) Generative cell of lily; phase contrast. Bar: 10 pm. (B) Fluorescence of the generative cell nucleus of lily as in (A), stained with DAPI. (C) Microtubule bundles in the generative cell in a pollen tube of Nicoriana tubacum stained with FITClabeled antibodies to a-tubulin (Del Casino et al., 1993). confocal scanning laser microscopy (Y.-Q. Li, unpublished). Bar: 5 pm.

Microtubule bundles are not regularly bridged to the plasma membrane or to the nuclear envelope (Cresti et al., 1990) (Fig. 2). Evidence for actin microfilaments in generative cells is conflicting (Taylor er al., 1989; Palevitz and Liu, 1992; Knox et al., 1993) although microfilaments have been observed in vegetative cells (Lancelle et al., 1987; Palevitz and Liu, 1992). In Brassica, actin filaments encircled the generative cell, but it was not clear whether the actin was in the generative cell or in the vegetative cell surrounding it (Hause er d . ,1992). Knox et al. (1993) showed short actin microfilaments in generative cells of lily in isolated vegetative cell protoplasts in culture. In contrast to the rapid movement of organelles in the vegetative cell, cytoplasmic streaming is minimal in generative cells (Pierson et al., 1990). This is consistent with the absence of actin microfilaments that would be involved in cytoplasmic streaming (Palevitz and Liu, 1992). Generative cell division occurs either within the pollen grain before shedding

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of pollen from anthers or in the pollen tube after pollination and pollen tube germination (Fig. 1). Timing of cell division is hereditary, but regulative mechanisms are unknown. Generative cell division within the pollen grain is a typical mitosis with formation of a cell plate. Division within the pollen tube is unusual in that the cell plate is oblique or longitudinal to the long axis of the pollen tube (Terasaka and Niitsu, 1989). A skewed spindle is formed with chromosomes moving nearly parallel to the metaphase plate. The cell plate is reduced. The cytoskeleton of the generative cell gives rise to the cytoskeleton of the sperm cell. In tobacco, generative cell microtubule bundles disassemble, then reorganize as a spindle apparatus, followed by formation of new microtubule bundles in sperm (Palevitz, 1993; Yu and Russell, 1993). In Trudescuntiu, generative cell microtubule bundles participate directly in spindle formation by kinetochore fiber capture (Palevitz and Cresti, 1989; Palevitz, 1990; Liu and Palevitz, 1991, 1992; Palevitz and Tiezzi, 1992). Interphase microtubules do not disassem ble. Microtubule bundles are present during prophase and become part of the spindle apparatus. After cytokinesis, the original microtubule bundles of generative cells form the microtubule bundles of sperm. In sperm, microtubule bundles generally consist of fewer microtubules than in generative cells but not precisely half as many (Cresti er al., 1990; Yu and Russell, 1993).

B. Sperm Structure 1. Morphology

Sperm in pollen grains or pollen tubes are surrounded by two membranes: the sperm cell membrane plus the inner cell membrane of the vegetative cell (Fig. 4). Their shape ranges from spheroidal to highly elongate with one or more thin extensions 30 ym or longer. They are unusually small, as little as 3 ym in diameter, with few organelles, little cytoplasm, and condensed chromatin. The cytoskeleton is similar to that in generative cells, a basket or cage of microtubule bundles arranged around the nucleus. The small size of sperm cells derives from the small size of the generative cell, from their lack of growth, and from cytoplasmic diminution observed in serial reconstructions (Yu and Russell, 1992). In tobacco, vesicles or cytoplasmic bodies containing membranous organelles and rarely microtubules pinch off from sperm cells (Mogensen and Rusche, 1985; Yu et ul., 1992). Vesicles remain in pockets in the vegetative cell. Further loss of cytoplasm occurs at fertilization when a cytoplasmic body is Ieft outside the egg at fertilization (Mogensen, 1982, 1988). Sperm nuclei are oval to elongate, with most of the nuclear volume occupied by condensed chromatin in a tightly packed mass that fluoresces brightly with

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Fig. 4 Sperm of Gerberujumesonii. (A) Freeze-fracture of sperm in pollen. N, Nucleus. Bar: 0.5 pm. (B) Thin section of sperm; nucleus with condensed chromatin. Bar: 0.5 pm. (C) Isolated sperm with extension (arrowhead); phase contrast. Bar: 5 pm. (D) Isolated sperm as in (C); DAPI staining, fluorescence microscopy. [(A) and (B) from Southworth (1990),Fig. 3, p. 99; Fig. 8, p. 101; (C) and (D) from Southworth and Knox (1989), Fig. l c and d, p. 275.1

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DNA-fluorochromes (Fig. 4). In thin sections, condensed chromatin is unevenly stained (Fig. 4). Nucleoli are difficult to identify. Occasional nuclear “vacuoles,” spheroidal zones free of chromatin, are observed. Mitochondria have been observed in nuclei of sperm cells, apparently trapped there during reconstitution of the nuclear envelope under close condition!: in the pollen tube (Yu and Russell, 1994~).The nuclear envelope lies in contact with condensed chromatin, and nuclear pores are sparse to absent (Southworth et al., 1989b; Southworth, 1990) (Fig. 4). Sperm organelles include mitochondria, plastids, ribosomes, endoplasmic reticulum, dictyosomes, and vesicles. No structure comparable to an acrosome has been detected. As in generative cells, bundles of microtubules, with a predominantly axial orientation, branch and rejoin to form a slightly twisted basket or cage of microtubule bundles surrounding the nucleus (Cresti et al., 1992; Palevitz and Tiezzi, 1992; Pierson and Cresti, 1992; Knox et a/., 1993). Microtubule bundles show no particular association with the plasma membrane or nuclear envelope, although the small volume of cytoplasm leads to close proximity of the bundles to both nucleus and plasma membrane (Fig. 5). Similar patterns of microtubule bundles are found in sperm cells located in tricellular pollen at anthesis and in pollen tubes derived from bicellular pollen after mitosis. Microtubule bundles terminate in the cellular extensions. Because microtubule bundles join and diverge, the number of bundles and the microtubules per bundle change along the length of the sperm so that a single cross-section provides little quantitative information. Microtubules in flowering plant sperm are particularly labile and depolymerize when sperm are released from the pollen tube by osmotic shock. Isolated sperm change shape from spindle shaped to spheroidal within a few minutes (Russell, 1991; Theunis er al., 1991; Palevitz and Tiezzi, 1992; Zee, 1992). This correlates with loss of microtubule bundles and of polymerized microtubules. Only in high osmotica was the spindle shape maintained (Southworth and Knox, 1989). The presence of actin in sperm, as in generative cells, is uncertain. Taylor et al. (1989) reported actin in sperm in pollen tubes of Rhododendron. However, Palevitz and Liu (1992) were unable to replicate these results. Actin was not detected in sperm cells of Brussica (Hause et al., 1992). Sperm of some species are dimorphic with respect to size and number of organelles (Russell, 1991; Zhu et al., 1992; Yu and Russell, 1994b). Other species show slight differences in size and shape, but no distinct dimorphism (Mogensen and Rusche, 1985). In Plumbago, with the most dimorphic sperm of any described, sperm closest to the vegetative nucleus had no plastids and more than 200 mitochondria whereas the other sperm contained fewer mitochondria and more than 20 plastids (Russell, 1984). Reports of sperm dimorphism are of particular interest because of their possible relationship to double fertilization (see Section IV).

Fig. 5 Sperm of Brussicu sp. (A) Microtubule bundles in cross-section of sperm in the pollen tube of B. oleruceu. A gap, probably induced by aldehyde fixation, separates the sperm cell from the surrounding vegetative cell (M. Cresti, unpublished). Bar:0.1 krn. (B) Sperm and vegetative nucleus in pollen tube of B. campestris. This constitutes the male germ unit. DAPI staining; fluorescence microscopy. Bar: 10 km. (D) Isolated sperm cells as in (C); DAPI staining, fluorescence microscopy.

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2. Male Germ Unit Sperm in pollen tubes remain in close association with each other and with the vegetative nucleus. In many species, one sperm is attached, via a long extension, to the vegetative cell membrane appressed to the vegetative nucleus. The second sperm is attached to the opposite end of the first either by membrane contact or by extracellular matrix. This tripartite structure, two sperm plus vegetative nucleus, is called the male germ unit (Dumas et al., 1984; Mogensen, 1992; Yu et al., 1992) (Fig. 5). Contacts between vegetative cell and generative cell are transient, separating during division of the generative cell and reforming as a contact between one sperm and the vegetative cell (Yu and Russell, 1994a). The two sperm cells and vegetative nucleus remain together during pollen tube growth and passage of sperm through the pollen tube, but separate quickly when the pollen tube is ruptured. This association serves to deliver two sperm cells simultaneously to the embryo sac for double fertilization.

3. Sperm in Vitro Sperm cells have been isolated from pollen grains and pollen tubes (Southworth and Knox, 1989; Theunis et al., 1991; Russell, 1991) (Fig. 5). Methods include osmotic shock, gentle grinding, and enzymatic digestion of the pollen tube cell wall. Sperm and generative celis apparently contain a lower osmoticum than the vegetative cell, so a solution that is hypotonic to the vegetative cell may be isotonic or hypertonic to sperm (Southworth and Morningstar, 1992). Isolated sperm retain membrane integrity as measured by osmotic response, exclusion of charged dyes, and the fluorochromatic reaction (Southworth and Knox, 1989). Isolated sperm can function in in vitro fertilization experiments (see Section IV, B) . C. Pollen Tube Growth

Pollen is carried by animals, primarily insects, or by wind to the stigma, the receptive portion of female flower parts. This process is pollination, but not fertilization. Pollen and stigma recognize each other, and the stigmatic cells signal the pollen to germinate. The vegetative cell in the pollen grain hydrates and elongates, forming a pollen tube (Mascarenhas, 1993). Sperm cells are in the pollen grain and move into the pollen tube (Fig. 5). Sperm reach the egg by passing through the pollen tube as it grows toward the egg.

1. Mechanisms of Pollen Tube Growth Pollen germinates on the surface of the stigma, penetrates between stigmatic cells, and grows within the cell walls of stigmatic cells and between cells of the

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style (Heslop-Hamson and Heslop-Harrison, 1994). Often stylar tissue is loosely compacted, and pollen tube growth proceeds through a gellike extracellular matrix that nourishes the pollen tube and determines the route of growth. Pollen tubes grow out the base of the style into the ovary cavity, where they grow along tissue surfaces and into the micropyles of ovules. Pollen tubes burst in cells adjacent to the egg, freeing the sperm at a distance of a few microns from the egg and central cells (Fig. 6). Pollen tubes grow at their tips, away from the pollen grain (Pierson and Cresti, 1992). They develop osmotic pressure, and with the constraint of cell wall polysaccharides, retain plasticity only at the tip where new wall material is added from dictyosome vesicles. Pollen tubes have an extensive cytoskeleton including microtubules, actin microfilaments, myosin on organelles and on the generative cell surface, and a number of motor proteins including dynein and kinesin (Lan1987; h t r o m et d., 1995; Miller et d., 1995; Tirlapur et d., 1995). celle et d., The presence of the microtubule motor protein kinesin could be detected in the pollen tube, but not in the generative cell (Cai et al., 1993). Sanders and Lord (1989, 1992) observed that polystyrene latex beads placed on whole or cut stigmas were translocated in and through the style, suggesting that the extracellular matrix of stylar tissues interacts with cell surfaces to pro-

Hg. 6 Organization of the embryo sac in an ovule. The tip of the pollen tube has entered one of the two synergids and ruptured, releasing two sperm (S). One sperm will fuse with the egg and the other with the central cell.

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mote pollen tube extension. A molecule recognized by antibodies to vitronectin, a surface adhesion molecule, was found in styles (Sanders et al., 1991). Sanders and Lord (1992) argue that growth of the pollen tube by cellular extension of pollen tube resembles the directed cell movements of animal cells, particularly the cells of the neural crest. 2. Movement of Sperm Cells

Sperm and generative cells travel through the pollen tube; however, spontaneous movements, of sperm or generative cells have not been reported (Pierson et al., 1990; Pierson and Cresti, 1992). Flowering plant sperm are not flagellated. They lack autonomous motility, exhibiting no ameboid motion or twisting. It is likely that sperm are moved by the actions of the vegetative cell surrounding it. The cytoskeleton of sperm cells is not organized to promote movement. Microtubules are infrequently linked to the plasma membrane (Cresti et al., 1990; Del Casino et aE., 1992). Because actin is usually a component of ameboid motion, the lack of actin microfilaments in sperm cells is consistent with lack of self-generated motility. By contrast, the pollen tube cytoplasm of the surrounding vegetative cell is well suited for intracellular movements (Palevitz and Tiezzi, 1992; Pierson and Cresti, 1992). Cytoplasmic streaming is a major feature of the pollen tube cytoplasm. A myosin-like protein has been observed at the interface of vegetative and generative cells (Pierson and Cresti, 1992; Bohdanowicz et al., 1995; Tirlapur et al., 1995), and capitate projections on the inner plasma membrane of vegetative cells are sometimes associated with microtubules or microfilaments in vegetative cell cytoplasm (VanWent and Gori, 1989; Cresti et al., 1991; Southworth et al., 1994; Bohdanowicz et al., 1995). Microtubules in the pollen tube form a network around the generative cell and vegetative nucleus. Movement is slowed after oryzalin treatment, which disrupts microtubules (Astrom et al., 1995). This evidence suggests that microfilaments in vegetative cell cytoplasm have a key role in sperm movement within the pollen tube, with the direction of movement determined by polarity of microfilaments or microtubules (Pierson and Cresti, 1992). Sperm movement depends on a connection between the sperm plasma membrane and the inner vegetative cell plasma membrane lining the pocket in which the sperm is located. In turn, the inner vegetative cell membrane may be attached to the pollen tube cytoskeleton that provides the motive force (Lancelle et al., 1987; Southworth et al., 1989a,b; Southworth, 1990, 1992).

111. Female Gametes Meiosis leading to egg formation occurs within the ovary of a flower bud. In each ovary are one to many ovules. Each ovule is an immature seed consisting of one

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meiocyte surrounded by several tissue layers and attached by a vascular connection to the placenta or inner lining of the ovary (Fig. 6).

A. Meiosis to Embryo Sac Formation

Events beginning with meiosis and ending in differentiation of an egg cell vary among flowering plants. In the most common pattern (called the Polygonum type), three of the four meiotic products die, leaving one functional haploid cell per meiocyte, a situation roughly comparable to the loss of polar bodies in animal oogenesis (Gifford and Foster, 1989; Huang and Russell, 1992). After meiosis, the functional haploid nucleus divides three times to produce eight nuclei. The cytoplasm then cleaves to form the cells of an embryo sac (Fig. 6). One of these differentiates into an egg. The factors that determine which haploid cell will become the egg appear to be related to the position of the cell within the embryo sac. Two cells adjacent to the egg become synergids that function as the destination of the pollen tube. Adjacent to the egg in the center of the embryo sac, two nuclei fuse before cytokinesis to form a diploid central cell. An embryo sac thus consists of seven cells: one egg, two synergids, one diploid central cell, and three other haploid cells that do not function in fertilization. Following meiosis and several cycles of mitosis in an ovule, one haploid cell becomes the egg, and one cell becomes the central cell. Both are fertilized by sperm, usually from the same pollen tube. This is the process of double fertilization that is characteristic of flowering plants.

B. Egg Structure

Distinguishing features of flowering plant eggs include the location in the embryo sac, polarity of cell shape and organelle distribution, and incompleteness of the cell wall (Russell et a l . , 1990; Huang and Russell, 1992). The egg cell differentiates at the end of the embryo sac nearest the opening (micropyle) through which the pollen tube will enter (Fig. 6). The egg itself is polarized, with the nucleus either central or at one end of the egg and with a large vacuole either surrounding the nucleus or at the opposite end of the cell (Zhu el al., 1993). A partial cell wall encloses the egg, leaving a portion of the plasma membrane adjacent to the plasma membrane of the synergids with no wall between them. The organelles of egg cytoplasm suggest a cell that is quiescent but potentially active. The nucleus is relatively large with a large nucleolus. Ribosomes are abundant in the cytoplasm, but not as polysomes. Dictyosomes and endoplasmic reticulum are present but with few vesicles. Plastids contain few internal membranes and infrequently starch. The location of plastids within the egg varies

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from an even distribution to a polarized distribution away from the micropylar end of the egg (Zhu et al., 1993). Mitochondria are often clustered around the nucleus. Embryo sacs and eggs have been isolated from ovules in 27 genera by a combination of enzymatic digestion and squashing or micromanipulation (Huang and Russell, 1992; Huang et al., 1992). Egg cells can be distinguished from other cells of the embryo sac by their location in partially intact isolated embryo sacs and by their relative size among the cells freed from embryo sacs. These eggs are alive as judged by the fluorochromatic reaction and by their ability to fuse with isolated sperm in vitro (see Section IV,B).

C. Central Cell Structure

The central cell is distinguished by its location in the embryo sac, the incompleteness of the cell wall, the number of nuclei and their state of fusion, and the large size of the vacuole (Huang and Russell, 1992; Huang et al., 1992). Cytoplasmic strands traverse the vacuole, and organelles pass along these strands. Central cells are commonly uninucleate (diploid) or binucleate (haploid). Usually nuclei fuse before fertilization.

IV. Double Fertilization Double fertilization is a characterizing feature of flowering plants (Gifford and Foster, 1989; Russell, 1992; Knox et al., 1993). The primary event of fertilization is the fusion of egg and sperm, producing a zygote that develops into an embryo. A second fertilization or fusion event occurs between the second sperm and the central cell adjacent to the egg. This second fertilization forms endosperm, a nutritive tissue that supports growth of the immature embryo. Both fertilizations are essential for reproductive success. Ultrastructural details of fertilization are available for few species. The events of fertilization in flowering plants are not easy to follow because the egg is embedded in tissues of the ovule.

A. Fertilization in Vivo A pollen tube with two sperm enters the ovule through an opening called the micropyle (Fig. 6). The pollen tube generally ruptures in one synergid, but in

plants lacking synergids, it bursts between cells of the embryo sac. Sperm are released from the pollen tube near the wall-less portions of the egg and central cell.

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Fertilization occurs with fusion of egg and sperm membranes. Membranes make contact and fuse at numerous sites along the contact zone. Vesicles form as fusion occurs. Russell (1983) estimated that the time for fusion was less than 1 min in Plumbago because partially fused gametes were rare in ovules sampled at 5-min intervals in a precisely timed system. Fusion of sperm and central cell membranes proceeds similarly. The sperm nucleus enters the egg or central cell. Entry of organelles is more variable. In barley (Mogensen, 1982, 1988) and in Plumbago, a cytoplasmic body derived from the sperm cell is eliminated or prevented from entering the egg, and only a portion of the sperm plasma membrane incorporates into the zygote membrane. Sperm mitochondria and plastids may be excluded from the egg during the fusion process. This explains the maternal inheritance pattern of mitochondria and plastids in most species. Because there are two events of fertilization and two sperm per pollen tube, the question arises whether there is preferential fertilization; that is, whether the two sperm cells differ in such a way that each sperm has a predetermined destination. The sperm of Plumbago have the most extreme dimorphic distribution of any sperm yet observed (Russell, 1984). In Plumbago, the sperm that is not attached to the vegetative nucleus and that contains more plastids and fewer mitochondria fuses with the egg in 94% of cases (Russell, 1985). This observation suggests some mechanism for preferential fertilization in this species; however, preferential fertilization may not occur in plants that do not have dimorphic sperm.

B. Fertilization in Vitro Because sperm cells are encased in pollen tubes, and egg cells are embedded in ovular tissue, in vitro fertilization cannot be achieved readily. With the development of sperm isolation techniques and advances in the separation of eggs from ovules and embryo sacs in the 1980s, experimental approaches to in vitro fertilization of flowering plants have been successful. Kranz et al. (1991) pioneered fusion of isolated gametes by electrofusion. Sperm isolated by hypoosmotic shock were brought into surface contact with eggs isolated by enzymatic digestion and micromanipulation. They fused rapidly (in 1 sec) by electrofusion. These in vitro zygotes divided and gave rise to masses of cells (Kranz et ul., 1992). While these results were exciting, they most likely did not represent the method of fusion of plant gametes in vivo. Faure et al. (1994) demonstrated fusion of plant gametes in mannitol with 5 mM calcium. Gametes, isolated as by Kranz et al. (1992), were brought into contact by manipulation with needles in a fusion medium under mineral oil. After 4 min of adhesion, gametes fused within 10 sec. There was no fusion of a

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second sperm in 20 trials. In controls, sperm fused with other sperm cells in 2% of trials and with leaf mesophyll protoplasts in 17% of trials. This method may more closely imitate conditions in vivo. These results support the hypothesis that there is a block to polyspermy in flowering plant eggs (Faure et al., 1994).

V. Summary Flowering plant sperm resemble animal sperm with condensed chromatin, reduced cell volume, and specialized cytoskeleton. The structure of plant eggs is unlike that of animal eggs; however, egg activation by fertilization is a similar phenomenon requiring sperm-egg fusion, migration of gamete nuclei in the egg, and nuclear fusion followed by cell division. Sperm can be isolated readily from pollen grains or pollen tubes by osmotic shock. Eggs are more difficult to isolate, requiring both enzymatic digestion and micromanipulation. Research on fertilization, including blocks to polyspermy and egg activation, is at its inception. The tools for experimentation on fertilization are complex and not widely available. Simpler methods with which to test in vitro fertilization would be valuable. The events of gametogenesis, fertilization, and activation in animals provide useful hypotheses for work on plant gametes. For example, it should be possible to characterize gamete cell surfaces, especially of sperm, and to raise antibodies to surface molecules that function in recognition and fertilization. Second, it ought to be possible to identify membrane dimorphism between the two sperm in one pollen grain or tube to determine the underlying basis for double fertilization. The microtubular cytoskeleton is a highly distinctive feature of sperm. Quantitative comparisons of the cytoskeleton of generative cells and sperm in the same species, in bi- and tricellular pollen, e.g., counts of microtubule bundles and microtubules per bundle and quantitative comparison of cytoskeletons of generative cells and sperm in pollen grains and pollen tubes, would determine the stability of the cytoskeleton. A more complete description of the cytoskeleton, including location of y-tubulins (Palevitz et al., 1994), of microtubule-associated proteins (e.g., dynein and kinesin), and of components of centrioles and basal bodies, could identify dispersed microtubule-organizing centers and help us to understand the evolution of the distinctive sperm cytoskeleton. There is a growing base of information on the molecular biology of anther development (Mascarenhas, 1992; Twell, 1994). Currently, gene expression can be identified at stages of anther development and in tissues of the anther, including pollen grains. Because sperm and generative cells represent such a small fraction of the protein and RNA of anthers and pollen, most research on molecular biology of pollen is concerned with the vegetative cell. Further research resolving gene expression in developing sperm would be useful.

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Acknowledgments This work was supported by NSF Grants IBN-9305453 and IBN-9418178 through Research at Undergraduate Institutions.

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Index

A

Acrosome reaction, ion channel signaling, I 3 1 - 144 mammals, 137- I44 sea urchin, 131-136 starfish, 136-137 Axial structures, early activation cues. 189190

B Burillus subtilis. starvation-induced endospore formation, 21 1-215 Bacteria, development, 207-245 cellular differentiation control, 226-245 chromosome replication, 226-229 protein degradation timing, 241-245 protein targeting, 238-241 transcription location, 229-238 concepts, 207-208 future research directions, 245 normal differentiation events, 209-210 starvation-induced events endospore formation, 21 1-215 fruiting body development, 215-218 sporulation, 218-221 stationary phase adaptation, 224-226 symbiotic relationships, 22 1-224 C

Caenorhabditis elegans, myogenic helix-loophelix transcription factor myogenesis characteristics, 180-181 mutational analysis, 182- 183 Caulobacter crescentus cellular differentiation control, 226-245 chromosome replication, 226-229 protein degradation timing, 241-245 protein targeting, 238-241 transcription location, 229-238 normal differentiation events, 209-21 0 Chromatin, decondensation, sperm nuclei to male pronuclei transformation, 4 1-52

in virro conditions, 47-52 in vivo conditions, 41-46 Chromosome replication, bacteria differentiation control, 226-229 Cytoplasmic incompatibility, Drosophilu fertilization, 103-107

D Deadhead effect, Drosophila embryo development, 101-102 Development models bacteria, 207-245 cellular differentiation control, 226-245 chromosome replication, 226-229 protein degradation timing, 241 -245 protein targeting, 238-241 transcription location, 229-238 concepts, 207-208 future research directions, 245 normal differentiation events, 209-210 starvation-induced events endospore formation, 21 1-215 fruiting body development, 215-218 sporulation, 218-221 stationary phase adaptation, 224-226 symbiotic relationships, 221-224 Drosophila embryos, deadhead effect, 101102 myogenic helix-loop-helix transcription factor myogenesis developmental expression, 175- 179 gene expression patterns, 177- 178 somite subdomains, 178-179 somitogenesis, 175-176 myocyte enhancer factor 2 family, 196197 Differentiation, bacteria development, 226-245 chromosome replication, 226-229 normal events, 209-210 protein degradation timing, 241 -245 protein targeting, 238-241 transcription location, 229-238 281

282 Double fertilization, flowering plant fertilization, 273-275 Drosophila myogenic helix-loop-helix transcription factor myogenesis characteristics, 179- 181 mutational analysis, 182- 183 sperm-egg interactions, 89- 112 cytoplasmic incompatibility, 103- I07 genetics deadhead effect, 101-102 maternal-effect mutations, 100-102 paternal-effect mutations, 102- 103 young arrest, 100-101 karyogamy, 97 overview, 89-92 pronuclear maturation, 95-97 sperm characteristics penetration, 95 production, 92-94 storage, 94 transfer, 94 utilization, 94 sperm-derived structure analysis, 98- 100 sperm function models, 107-1 11 diffusion-gradient production, 109-1 I0 nutritive protein import, 107-109 specific protein import, 109 structural role, 110-1 11 syngamy, 95 sperm nuclei to male pronuclei transformation cell-free preparation comparison, 29-3 1 chromatin decondensation in vitro conditions, 49-50 in vivo conditions, 42-43 male pronuclear activities development comparison, 27-29 replication, 75-76 transcription reinitiation, 77 maternal histone exchange, 36 nuclear envelope alterations formation, 60-61 lamin role, 67-68 removal, 56 nucleosome formation, 54 sperm protein modifications, 36 DSS locus, sex determination role, 16-17

E Eggs Drosophila sperm-egg interactions, 89-1 12

Index cytoplasmic incompatibility, 103-107 genetics deadhead effect, 101-102 maternal-effect mutations, 100-102 paternal-effect mutations, 102- 103 young arrest, 100-101 karyogamy, 97 overview, 89-92 pronuclear maturation, 95-97 sperm characteristics penetration, 95 production, 92-94 storage, 94 transfer, 94 utilization, 94 sperm-derived structure analysis, 98-100 sperm function models, 107-1 11 diffusion-gradient production, 109- 110 nutritive protein import, 107-109 specific protein import, 109 structural role, 110-11 1 syngamy, 95 flowering plant fertilization, 27 1-273 ion channel signaling, 117-149 egg activation, 144-148 egg characteristics, 1 19-121 importance, 117- 118 ionic environment influence fish sperm, 121-122 mammalian sperm, 123 sea urchin sperm, 121-122 long-range gametic communication, 124131 mammals, 130-131 sea urchins, 124-130 short-range gametic communication, 131144 mammals, 137-144 sea urchin, 131-136 starfish, 136-137 Embryo development Drosophila deadhead effect, 101-102 maternal-effect mutations, 100- 102 paternal-effect mutations, 102- 103 young arrest, 100-101 myogenic helix-loop-helix transcription factor myogenesis, 169-199 developmental expression, 175- 179 gene expression patterns, 177- 178 somite subdomains, 178- 179 somitogenesis, 175- 176

Index

283

early activation, 188- 191 axial structure cues, 189-190 regulatory element analysis, 191-194 invertebrate models, 179- 181 mutational analysis, 183- 188 invertebrate genes, 182-183 mouse mutations, 183-188 myocyte enhancer factor 2 family, 194198 characteristics, 194- 196 MyoD family, 171- 175 cloning, 171-172 properties, 172- 174 regulation, 174-175 nonmammalian vertebrate models, 18 1182 overview, 169- 171 Endospores, starvation-induced formation, 21 1-215 Escherichia coli, stationary phase adaptation, 224-226

male gametes, 260-27 1 generative cell development, 262-265 pollen formation, 260-262 pollen tube growth, 269-271 sperm structure, 265-269 overview, 259-260 Fish, gamete signaling, 121-122 Flowering plants, fertilization mechanisms, 259-275 double fertilization, 273-275 female gametes, 271-273 male gametes, 260-271 generative cell development, 262-265 pollen formation, 260-262 pollen tube growth, 269-271 sperm structure, 265-269 overview, 259-260 Fruit flies, see Drosophila Fruiting bodies, starvation-induced development, 215-218

F

Gametes, see Eggs; Fertilization; Sperm Gonadogenesis SRY gene role, 2-3 testis-determining factor, 1-2

G Female pronuclei, maturation, 95-97 Fertilization Drosophila sperm-egg interactions, 89- 1 12 cytoplasmic incompatibility, 103- 107 genetics deadhead effect, 101-102 maternal-effect mutations, 100- 102 paternal-effect mutations, 102- 103 young arrest, 100-101 karyogamy, 97 overview, 89-92 pronuclear maturation, 95-97 sperm characteristics penetration, 95 production, 92-94 storage, 94 transfer, 94 utilization, 94 sperm-derived structure analysis, 98- I00 sperm function models, 107-1 I 1 diffusion-gradient production, 109-1 10 nutritive protein import, 107-109 specific protein import, 109 structural role, 110- 1 1 I syngamy, 95 flowering plants, 259-275 double fertilization, 273-275 female gametes, 271-273

H High mobility group box, sex determination role, 7-8 Histone, sperm nuclei to male pronuclei transformation, 33-40

I Incompatibility, cytoplasm, Drosophila fertilization, 103-107 Ion channels, gamete signaling, 117-149 egg activation, 144- 148 gamete characteristics egg, 119-121 sperm, 118-119 importance, 117-118 ionic environment influence fish sperm, 121-122 mammalian sperm, 123 sea urchin sperm, 121-122 long-range gametic communication, 124131 mammals, 130-131 sea urchins, 124-130

284 Ion Channels, gamete signaling (conrinued) short-range gametic communication, 131I44 mammals, 137-144 sea urchin, 131-136 starfish. 136-137

K Karyogamy, Drosophila sperm-egg interactions, 97

L Lamin, sperm nuclei to male pronuclei transformation, 64-69 Legumes, Rhizobriurn symbiotic relationship, 221-224

M Male pronuclei, sperm nuclei transformation, 26-78 chromatin decondensation, 41-52 in v i m conditions, 47-52 in vivo conditions, 41-46 male pronuclear activities, 75-78 replication, 75-76 transcription reinitiation, 76-77 maternal histone exchange, 33-40 nuclear envelope alterations, 55-74 formation, 59-64 lamin role, 64-69 nuclear pores, 69-71 removal, 55-59 nuclear protein changes, 32-40 nucleosome formation, 52-55 overview, 26-32 cell-free preparation comparison, 29-32 male pronuclear development comparison, in vivo. 27-29 Mammals ion channel signaling ionic environment influence, 123 long-range gametic communication, 130131 short-range gametic communication, 137144 myogenic helix-loop-helix transcription factor myogenesis, mutational analysis, 183-1 88

Index sex determination, 1- 18 DSS locus, 16-17 Miillerian inhibitory substance, 13-14 overview, 1-2 SOX9 gene, 15-16 SRY gene, 2-7 gonadogenesis, 2-3 transcription, 3-5 transcript structure, 5-7 SRY protein, 7-13 DNA-binding properties, 8-9 high mobility group box, 7-8 transcript activation, 9-12 steroidogenic factor 1, 14-15 Tas locus, 16-17 testis-determining factor, 1-2 Wilms' tumor-associated gene, 15 Maternal-effect mutations, Drosophila embryo development, 100- 102 Maternal histone, sperm nuclei to male pronuclei transformation, 33-40 Mouse, myogenic helix-loop-helix transcription factor myogenesis, mutational analysis, 183-188 Miillerian inhibitory substance, sex determination role, 13-14 Mutational analysis Drosophila sperm-egg interactions maternal-effect, 100- 102 paternal-effect, 102- 103 myogenic helix-loop-helix transcription factor myogenesis, 183-188 Myoblasts, myogenic helix-loop-helix transcription factor early activation, 190- 191 Myogenesis, myogenic helix-loop-helix transcription factors. 169-199 developmental expression, 175- 179 gene expression patterns, 177- 178 somite subdomains, 178-179 somitogenesis, 175- 176 early activation, 188- 194 axial structure cues, 189-190 migratory versus myotomal myoblasts, 190- 191 regulatory element analysis, 191-194 invertebrate models, 179- 181 mutational analysis, 183- 188 invertebrate genes, 182-183 mouse mutations, 183-188 myocyte enhancer factor 2 family, 194-198 characteristics, 194- 196

Index developmental expression, 196- 197 myogenesis regulation, 197- 198 MyoD family, 171-175 cloning, 171- 172 properties, 172-174 regulation, 174-175 nonmammalian vertebrate models, 181 - 182 overview, 169- 171 Mysococcus sunthus, starvation-induced fruiting body development, 215-218

N Nuclear envelope, sperm nuclei to male pronuclei transformation, 55-74 formation, 59-64 lamin role, 64-69 nuclear pores, 69-71 removal, 55-59 Nuclear pores, sperm nuclei to male pronuclei transformation, 69-71 Nucleosomes, sperm nuclei to male pronuclei transformation, 52-55

P Paternal-effect mutations, Drosophilu embryo development, 102- 103 Plants, see Flowering plants Pollen formation, 260-262 tube growth, 269-271 Pronuclei, see Female pronuclei; Male pronuclei

R Replication, chromosomes, bacteria differentiation control, 226-229 Rhizobrium, legume symbiotic relationship, 221-224

S Sea urchin ion channel signaling ionic environment influence, 12 I- 122 long-range gametic communication, 124 130 short-range gametic communication, 13 136

285 sperm nuclei to male pronuclei transformation cell-free preparation comparison, 3 1-32 chromatin decondensation in vitro conditions, 49-50 in vivo conditions, 44-46 male pronuclear activities development comparison, 27-29 replication, 76 transcription reinitiation, 77 maternal histone exchange, 37-40 nuclear envelope alterations formation, 63-64 lamin role, 68-69 removal, 57-59 nucleosome formation, 54-55 sperm protein modifications, 37-40 Sex determination, 1-18 DSS IOCUS, 16-17 Miillerian inhibitory substance, 13- 14 overview, 1-2 SOX9 gene, 15-16 SRY gene, 2-7 gonadogenesis, 2-3 transcription, tissue distribution, 3-5 transcript structure, 5-7 SRY protein, 7- 13 DNA-binding properties, 8-9 high mobility group box, 7-8 transcript activation, 9-12 steroidogenic factor I , 14- 15 TUSIOCUS,16-17 testis-determining factor, 1-2 Wilms’ tumor-associated gene, 15 Somites, myogenic helix-loop-helix transcription factor myogenesis, developmental expression somite subdomains, 178- 179 somitogenesis, 175-176 SOX9 gene, sex determination role, 15-16 Sperm cytoplasmic incompatibility, 103- 107 fertilization, 89- 1 12 characteristics penetration, 95 production, 92-94 storage, 94 transfer, 94 utilization, 94 flowering plants, 260-27 1 generative cell development, 262-265

286 Sperm, flowering plants (continued) pollen formation, 260-262 pollen tube growth, 269-271 sperm structure, 265-269 genetics deadhead effect, 101-102 maternal-effect mutations, 100- 102 paternal-effect mutations, 102- 103 young arrest, 100-101 karyogamy, 97 overview, 89-92 pronuclear maturation, 95-97 sperm-derived structure analysis, 98-100 sperm function models, 107- I 1 1 diffusion-gradient production, 109- 110 nutritive protein import, 107- 109 specific protein import, 109 structural role, 110-1 11 syngamy, 95 ion channel signaling, 117-149 egg activation, 144-148 importance, 117-1 18 ionic environment influence fish sperm, 121-122 mammalian sperm, 123 sea urchin sperm, 121-122 long-range gametic communication, 124131 mammals, 130- 131 sea urchins, 124-130 short-range gametic communication, 131144 mammals, 137-144 sea urchin, 131-136 starfish, 136- 137 sperm characteristics, 118- I19 Sperm nuclei, male pronuclei formation, 2678 chromatin decondensation, 41-52 in vitro conditions, 47-52 in vivo conditions, 41-46 male pronuclear activities, 75-78 replication, 75-76 transcription reinitiation, 76-77 maternal histone exchange, 33-40 nuclear envelope alterations, 55-74 formation, 59-64 lamin role, 64-69 nuclear pores, 69-71 removal, 55-59

Index nuclear protein changes, 32-40 nucleosome formation, 52-55 overview, 26-32 cell-free preparation comparison, 29-32 male pronuclear development comparison, in vivo, 27-29 Spisula solidissima. sperm nuclei to male pronuclei transformation cell-free preparation comparison, 3 1 chromatin decondensation in virro conditions, 50 in vivo conditions, 43-44 male pronuclei development comparison, 27-29 maternal histone exchange, 37 nuclear envelope alterations formation, 61-63 lamin role, 68 removal, 56-57 sperm protein modifications, 37 SRY gene, sex determination role, 2-7 gonadogenesis, 2-3 transcription, tissue distribution, 3-5 transcript structure, 5-7 SRY protein, sex determination role, 7-13 DNA-binding properties, 8-9 high mobility group box, 7-8 transcript activation, 9-12 Starfish, acrosome reaction, 136-137 Starvation-induced events, bacteria development endospore formation, 21 1-215 fruiting body development, 215-218 sporulation, 218-221 Steroidogenic factor 1, sex determination role, 14-15 Strepfomyces, starvation-induced sporulation, 218-221 Surf Clams, see Spisula solidissima Symbiosis, Rhizobrium and legume relationship, 221-224 Syngamy, Drosophila sperm-egg interactions, 95

T Tas locus, sex determination role, 16-17 Testis-determining factor, sex determination role, 1-2 Transcription bacteria models

Index location timing, 229-238 protein location targeting, 238-241 male pronuclei, 76-77 SRY gene activation, 9-12 tissue distribution, 3-5 transcript structure, 5-7 Transcription factors, myogenic helix-loophelix transcription factor myogenesis, 169-199 developmental expression, 175- 179 patterns, 177-178 somite subdomains, 178-179 somitogenesis, 175-176 early activation, 188-194 axial structure cues, 189- 190 myoblasts, 190-191 regulatory element analysis, 191-194 invertebrate models, 179-1 81 mutational analysis, 183- 188 invertebrate genes, 182- 183 mouse mutations, 183- 188 myocyte enhancer factor 2 family, 194-1 98 MyoD family, I7 1- 175 cloning, 171-172 properties, 172- 174 regulation, 174-175 nonmammalian vertebrate models, 181- 182 overview, 169-171 Tumor-associated genes, sex determination role, 15

287 W Wilms’ tumor-associated gene, sex determination role. 15

X Xenopus laevis myogenic helix-loop-helix transcription factor myogenesis, 181-182 sperm nuclei to male pronuclei transformation cell-free preparation comparison, 29-3 I chromatin decondensation in vitro conditions, 47-49 in vivo conditions, 41-42 male pronuclear activities development comparison, 27-29 replication, 75 transcription reinitiation, 76-77 maternal histone exchange, 33-36 nuclear envelope alterations formation, 60 lamin role, 66-67 nucleosome formation, 53-54 sperm protein modifications, 33-36

Y Young arrest, Drosophila embryo develop ment. 100-101

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