Current Topics in Developmental Biology, Volume 52 (Current Topics in Developmental Biology)

Current Topics in Developmental Biology

Volume 52

Series Editor Gerald P. Schatten Departments of Obstetrics–Gynecology and Cell and Developmental Biology Oregon Regional Primate Research Center Oregon Health Sciences University Beaverton, Oregon 97006-3499

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

Philip Ingham University of Sheffield, United Kingdom

Mary Lou King University of Miami, Florida

Story C. Landis National Institutes of Health/ National 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, Palo Alto, California

Founding Editors A. A. Moscona Alberto Monroy

Current Topics in Developmental Biology Volume 52 Edited by

Gerald P. Schatten Departments of Obstetrics–Gynecology and Cell and Developmental Biology Oregon Regional Primate Research Center Oregon Health Sciences University Beaverton, Oregon

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Cover photo credit: GUS activity in shoot apex of a two-day-old seedling. From Figure 6 in Chapter 3 “Mechanisms of Plant Embryo Development” by Shunong Bai, Lingjing Chen, Mary Alice Yund, and Zinmay Renee Sung. See text and color plate for further details.

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Academic Press Harcourt Place, 32 Jamestown Road, London NW1 7BY, UK http://www.academicpress.com International Standard Book Number: 0-12-153150-3 PRINTED IN THE UNITED STATES OF AMERICA 00 01 02 03 04 05 EB 9 8 7 6

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Contents

Contributors

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1 Mechanism and Control of Meiotic Recombination Initiation Scott Keeney I. II. III. IV.

Introduction 2 Double-Strand Breaks as Initiators of Meiotic Recombination The Cast of Players, from Fungi to Mammals 6 Additional Factors That Influence or Are Influenced by Recombination Initiation 31 V. A Molecular Model for Spo11 Action 37 References 39

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2 Osmoregulation and Cell Volume Regulation in the Preimplantation Embryo Jay M. Baltz I. Introduction 56 II. Sensitivity of Mammalian Embryos to Osmolarity 57 III. Response to Hypertonicity and Regulation against Cell Volume Decreases by Mammalian Embryos 60 IV. Regulation against Volume Increases by Mammalian Embryos 80 V. Organic Osmolytes and Osmolarity in Vivo 88 VI. Discussion and Summary 94 References 97

3 Cell–Cell Interactions in Vascular Development Diane C. Darland and Patricia A. D’Amore I. II. III. IV. V. VI.

Introduction 107 The Cardiovascular System 108 Differences in Vascular Beds 124 Parallels between Angiogenesis in Development and Pathology New Directions 133 Summary 138 References 139

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4 Genetic Regulation of Preimplantation Embryo Survival Carol M. Warner and Carol A. Brenner I. II. III. IV. V. VI. VII.

Introduction 151 Preimplantation Development 152 Environmental Effects on Preimplantation Embryo Survival Genetic Effects on Preimplantation Embryo Survival 159 Genes That Regulate Preimplantation Growth 170 Genes That Regulate Preimplantation Death 178 Conclusions 179 References 181

Index 193 Contents of Previous Volumes

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Contributors

Numbers in parentheses indicate the pages on which authors’ contributions begin.

Jay M. Baltz (55), Hormones, Growth, and Development Unit, Ottawa Health Research Institute, and Departments of Obstetrics and Gynecology (Division of Reproductive Medicine) and Cellular and Molecular Medicine, University of Ottawa, Ottawa, Ontario, Canada K1Y 4E9 Carol A. Brenner (151), Gamete and Embryo Laboratory, Institute for Reproductive Medicine and Science of Saint Barnabas Medical Center, West Orange, New Jersey 07052 Patricia A. D’Amore (107), Schepens Eye Research Institute, Department of Ophthalmology, and Department of Pathology, Harvard Medical School, Boston, Massachusetts 02114 Diane C. Darland (107), Schepens Eye Research Institute, Department of Ophthalmology, Harvard Medical School, Boston, Massachusetts 02114 Scott Keeney (1), Molecular Biology Program, Memorial Sloan-Kettering Cancer Center, and Weill Graduate School of Medical Sciences of Cornell University, New York, New York 10021 Carol M. Warner (151), Department of Biology, Northeastern University, Boston, Massachusetts 02115

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1 Mechanism and Control of Meiotic Recombination Initiation Scott Keeney Molecular Biology Program Memorial Sloan-Kettering Cancer Center, and Weill Graduate School of Medical Sciences of Cornell University New York, New York 10021

I. Introduction II. Double-Strand Breaks as Initiators of Meiotic Recombination A. Overview of the Double-Strand Break Pathway in Budding Yeast B. Evidence That Double-Strand Breaks Initiate Meiotic Recombination in Saccharomyces cerevisiae C. Probable Initiation of Meiotic Recombination in Other Organisms by Double-Strand Breaks III. The Cast of Players, from Fungi to Mammals A. SPO11 B. RAD50, MRE11, XRS2, and SAE2/COM1 C. Other Genes That Are Absolutely Required for Double-Strand Break Formation in Budding Yeast D. Roles of Double-Strand Break Genes in Development of Meiotic Chromosome Structure, Homologous Chromosome Pairing, and Progression through Meiotic Prophase E. Genes That Are Involved in, but Not Absolutely Required for, Double-Strand Break Formation F. Intergenic Interactions Important for Double-Strand Break Formation G. Other Potential Double-Strand Break Genes H. Possible Functions for the Friends of Spo11 IV. Additional Factors That Influence or Are Influenced by Recombination Initiation A. Site Selectivity: Chromatin Structure, Promoters, and Sequence Specificity B. Meiosis-Specific Alteration of Nuclease Hypersensitivity in the Chromatin at Recombination Hot Spots C. Interplay with the Development of Higher Order Chromosome Structure D. The DNA Replication Connection E. Homologous Chromosome Pairing F. Cell Cycle Control V. A Molecular Model for Spo11 Action References

Current Topics in Developmental Biology, Vol. 52 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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Scott Keeney Homologous recombination is essential during meiosis in most sexually reproducing organisms. In budding yeast, and most likely in other organisms as well, meiotic recombination proceeds via the formation and repair of DNA double-strand breaks (DSBs). These breaks appear to be formed by the Spo11 protein, with assistance from a large number of other gene products, by a topoisomerase-like transesterase mechanism. Recent studies in fission yeast, multicellular fungi, flies, worms, plants, and mammals indicate that the role of Spo11 in meiotic recombination initiation is highly conserved. This chapter reviews the properties of Spo11 and the other gene products required for meiotic DSB formation in a number of organisms and discusses ways in which recombination initiation is coordinated with other events occurring in the meiotic cell.  2001 Academic Press. C

I. Introduction In most sexually reproducing organisms, meiotic crossover recombination forms physical connections between homologous chromosomes that allow them to orient properly on the spindle and to segregate accurately at the first division. If recombination fails, chromosome disjunction also frequently fails, with disastrous consequences for gamete formation. Studies of a number of organisms are illuminating the molecular mechanism of meiotic recombination and are revealing how recombination is temporally and spatially coordinated with other cellular events. One of the most striking features of meiotic recombination is its frequency. During vegetative growth in Saccharomyces cerevisiae, spontaneous recombination typically occurs at a rate of 10−6–10−7 per locus per generation (e.g., Steele et al., 1991). During meiosis, in contrast, the frequency can reach 1–10%, a jump of 10,000-fold or more. This increase results from the induction of a highly regulated pathway in which meiosis-specific gene products redirect the activities of proteins that also function in recombinational DNA repair in vegetative cells. In budding yeast—and probably in other organisms as well—this pathway has at its heart the formation and subsequent repair, via homologous recombination, of programmed DNA double-strand breaks (DSBs) catalyzed by the Spo11 protein. This chapter reviews the mechanisms by which meiotic recombination initiates and is controlled. The review is divided into four main parts: an overview of the role of DSBs in meiotic recombination; a review of the proteins involved in DSB formation; a summary of how recombination initiation is coordinated with other events, such as the development of higher order chromosome structures; and discussion of a molecular model for DSB formation. Where appropriate, the similarities and often surprising differences between organisms are highlighted. Such comparisons provide useful insights, especially into the remarkable degree to which meiotic recombination mechanisms are evolutionarily conserved.

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II. Double-Strand Breaks as Initiators of Meiotic Recombination A. Overview of the Double-Strand Break Pathway in Budding Yeast Physical analysis of DNA isolated from meiotic cultures has defined the major steps along the recombination pathway in S. cerevisiae (Fig. 1). As had been suggested earlier (Resnick, 1976; Szostak et al., 1983), the initiating lesion is a DSB (Game et al., 1989; Sun et al., 1989; Cao et al., 1990; Zenvirth et al., 1992), most likely with a short (2-nucleotide) 5 overhang (de Massy et al., 1995; Liu et al., 1995; Xu and Kleckner, 1995; Xu and Petes, 1996) (see Section III,A,5 for more detailed discussion). Each DSB end is covalently attached to the Spo11 protein, presumably through a phosphodiester linkage between the 5 terminus and a tyrosine side chain on the protein (de Massy et al., 1995; Keeney and Kleckner, 1995; Liu et al., 1995; Bergerat et al., 1997; Keeney et al., 1997). In wild-type cells, Spo11 is removed and the 5 -strand termini are nucleolytically resected to yield molecules with variable-length, 3 single-stranded tails (Sun et al., 1991; Bishop et al., 1992). DNA strand exchange proteins (including Dmc1 and Rad51) catalyze invasion of these tails into intact homologous duplexes, giving rise to double-Holliday junction intermediates (Collins and Newlon, 1994; Schwacha and Kleckner, 1994, 1995, 1997) and, ultimately, mature recombinant products. The molecular details of the steps after DSB resection have been extensively reviewed (e.g., Kupiec et al., 1997; Roeder, 1997; Smith and Nicolas, 1998; Paques and Haber, 1999). A large number of genes are absolutely required for meiotic recombination initiation. These are listed in Fig. 1 and described in detail in Section III. A null mutation in any of these genes eliminates DSB formation and meiotic recombination, resulting in meiotic lethality from chromosome nondisjunction at the first division. In strains carrying certain rad50 or mre11 point mutations (e.g., a “rad50S” or “mre11S ” mutation), DSBs are formed but are not resected (Alani et al., 1990; Cao et al., 1990; Nairz and Klein, 1997). Because of this, strand invasion and double-Holliday junction formation do not occur (Schwacha and Kleckner, 1994). Deletion of the SAE2/COM1 gene also causes this phenotype (McKee and Kleckner, 1997; Prinz et al., 1997). B. Evidence That Double-Strand Breaks Initiate Meiotic Recombination in Saccharomyces cerevisiae Largely on the basis of studies of recombination between chromosomes and linearized plasmids, Szostak and colleagues (1983) proposed a model describing the repair of double-stranded gaps by homologous recombination and further suggested that this model could explain observed patterns of meiotic recombination. Although initiation mechanisms other than double-stranded cleavage cannot be

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Figure 1 Overview of the DSB pathway in S. cerevisiae. An intact DNA duplex is cleaved by Spo11 protein to yield a covalent Spo11–DNA complex (a and b). Spo11 is then released and the 5 -terminal strands are degraded to yield 3 single-stranded tails (c). These tails undergo strand invasion into an intact, homologous duplex (d), giving rise to double-Holliday junction intermediates (e) and, ultimately, mature recombinant products [an example with a crossover configuration is shown in (f)]. Genes required for formation and resection of DSBs are indicated. Those listed on the right are meiosis specific (except for NAM8/MRE2); those on the left have functions in vegetative cells as well. Mutations affecting the boxed genes reduce but do not eliminate DSB formation. MER1 and NAM8/MRE2 are required for DSB formation solely because they facilitate proper splicing of the MER2 transcript.

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entirely ruled out, there is no evidence to date to support their existence. For example, other models suggest that recombination could be initiated by the formation of single-stranded nicks (Holliday, 1964; Meselson and Radding, 1975), but sensitive assays have failed to detect such nicks at several recombination hot spots (Liu et al., 1995; Xu and Kleckner, 1995). There are important differences between current versions of the DSB repair model and its original incarnation (Gilbertson and Stahl, 1996; Stahl, 1996; Paques and Haber, 1999), but several lines of evidence strongly indicate that DSBs initiate meiotic recombination. First, DSBs occur transiently in the DNA of meiotic cells (Game et al., 1989; Sun et al., 1989; Cao et al., 1990; Zenvirth et al., 1992). The appearance and disappearance of these breaks match the timing of meiotic recombination, and the location and relative strength of DSB sites correlate well with recombination frequencies measured by genetic assays (Rocco et al., 1992; de Massy et al., 1994; Wu and Lichten, 1994, 1995; Fan et al., 1995; Borde et al., 1999). Moreover, physical assays have demonstrated other intermediates predicted by the DSB repair model, such as heteroduplex DNA (Lichten et al., 1990; Goyon and Lichten, 1993; Nag and Petes, 1993) and double-Holliday junctions (Bell and Byers, 1983; Collins and Newlon, 1994; Schwacha and Kleckner, 1994, 1995). Second, DSBs occur at sites that are inferred from genetic data to be recombination initiation sites. At certain loci, recombination frequencies vary with physical position along the DNA (Nicolas and Petes, 1994). Such a “polarity gradient” is thought to reflect the presence of a preferred site for recombination initiation at the high end of the gradient. The high ends of polarity gradients at several loci coincide with prominent DSB sites (Nicolas and Petes, 1994). Third, the DSB model predicts that the chromosome that initiates recombination will be the recipient of genetic information because it copies information from its intact homologous partner. For allele combinations that show disparity in the direction of information transfer, this prediction is met (e.g., Nicolas et al., 1989). Fourth, cis-acting mutations that raise or lower the recombination frequency at recombination hot spots also raise or lower the DSB frequency (de Massy and Nicolas, 1993; Fan et al., 1995; Wu and Lichten, 1995; Xu and Kleckner, 1995; Bullard et al., 1996). Likewise, mutations in trans-acting factors coordinately affect DSBs and recombination, both for mutations that affect only a particular hot spot (Fan et al., 1995) and for mutations that affect recombination globally (such as spo11). Importantly, mutations that allow DSBs to form but that block subsequent processing also eliminate recombination, as expected if DSBs are recombination precursors (e.g., rad50S) (Alani et al., 1990; Schwacha and Kleckner, 1994). Fifth, DSBs provided from an exogenous source (such as ionizing radiation) partially rescue the recombination and spore viability defects of a spo11 mutant (Thorne and Byers, 1993). Also, DSBs made by the HO (Malkova et al., 1996) or VDE (Gimble and Thorner, 1992) endonucleases during meiosis give rise to recombinant products. Moreover, the HO-mediated recombination events have been shown to be largely indistinguishable from normal meiotic events (Malkova et al., 1996, 2000).

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C. Probable Initiation of Meiotic Recombination in Other Organisms by Double-Strand Breaks The mechanism of meiotic recombination initiation is almost certainly conserved. Transient, meiosis-specific DSBs have been demonstrated in Schizosaccharomyces pombe, dependent on genes known to be required for meiotic recombination, including the SPO11 homolog rec12+ (Cervantes et al., 2000; Zenvirth and Simchen, 2000). These DSBs accumulate in mutant strains defective for the recA homolog rhp51+, as expected if the breaks are precursors of meiotic recombination. These are exciting results, but some puzzles remain. No DSBs have been detected at the strong meiotic recombination hot spot ade6-M26 (Bahler et al., 1991; Cervantes et al., 2000). Moreover, there is no clear correlation between DSB location and crossover distribution at several loci examined. To reconcile these anomalies, it has been proposed that meiotic DSBs serve as entry points for a recombination machine, which translocates along the chromosome until it generates a crossover at some distance from the DSB site (Cervantes et al., 2000). Such a model is similar in some regards to the mechanism by which RecBCD initiates recombination from DSBs in Escherichia coli (Eggleston and West, 1997). Alternatively, a double-Holliday junction formed at a DSB site could branch migrate along the chromosome and be resolved a considerable distance away (J. Kohli and G. Smith, personal communication, 2000). No reports of meiotic DSBs in other organisms have been published to date. However, DSBs are potent inducers of homologous recombination in many organisms (e.g., Lankenau, 1995; Liang et al., 1998). Moreover, Spo11 is widely conserved (Section III,A,3), and meiotic defects in spo11 mutants of Coprinus cinereus and Caenorhabditis elegans can be partially rescued by ionizing radiation (Dernburg et al., 1998; Celerin et al., 2000). Other proteins required for DSB formation are also conserved (Section III), as are many of the downstream DSB-processing activities, such as the meiosis-specific recA homolog DMC1 (Bishop et al., 1992; Pittman et al., 1998; Yoshida et al., 1998). Finally, γ -H2AX (a phosphorylated form of histone H2AX that is thought to be a marker for the location of DSBs; Rogakou et al., 1998) appears on meiotic chromosomes from leptonema through zygonema in mouse in a Spo11-dependent fashion (Mahadevaiah et al., 2001). Taken together, these observations make a strong argument that DSB formation is a universal feature of meiotic recombination.

III. The Cast of Players, from Fungi to Mammals A. SPO11 1. Identification and Early Characterization Saccharomyces cerevisiae SPO11 was identified in a screen for temperaturesensitive mutations that eliminated ascospore formation (Esposito and Esposito,

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1969, 1974; Esposito et al., 1972). SPO11 is absolutely required for intragenic and intergenic meiotic recombination, and spo11 mutants make no DSBs (Klapholz et al., 1985; Wagstaff et al., 1985; Cao et al., 1990). In many commonly used laboratory strains, spo11 mutants often have only a minor defect in spore formation. However, without crossovers to ensure accurate chromosome segregation, spo11 mutants show wholesale chromosome nondisjunction at the first division and thus generate aneuploid, inviable spores. SPO11 encodes a 45-kDa protein and its message is transcribed only during meiosis (Atcheson et al., 1987; Giroux et al., 1989). The cis- and trans-acting factors responsible for repression in vegetative cells and for activation in early meiosis have been characterized in detail (Mitchell, 1994; Kupiec et al., 1997; Vershon and Pierce, 2000). SPO11 and other meiotic transcripts have a much shorter half-life in vegetative cells than during meiosis, indicating that posttranscriptional mechanisms also control SPO11 expression (Surosky and Esposito, 1992). The mechanisms responsible for destabilizing these transcripts in mitotic cells, or stabilizing them in meiotic cells, are not understood. The spore inviability of spo11 mutants can be rescued by a spo13 mutation, which causes cells to undergo a single-division meiosis and package two diploid spores (Klapholz and Esposito, 1980; Klapholz et al., 1985). In a spo11 spo13 double mutant, the single meiotic division is predominantly equational (i.e., sister chromatids segregate from one another, as in mitosis) and crossovers are eliminated, resulting in complete linkage of genes along a chromosome (Klapholz et al., 1985). This property was used as an early tool for mapping mutations in other genes (Klapholz and Esposito, 1982) and, more importantly, has been exploited to study mutants (such as spo11 and rad50) that would otherwise give inviable meiotic products (Malone and Esposito, 1981; Klapholz et al., 1985). For example, recombination mutants can be classified as “early” or “late” on the basis of the ability of a spo13 mutation to rescue spore viability (Malone, 1983; Malone et al., 1991). Early mutants can be rescued by spo13 and include spo11 and rad50. In contrast, late mutants (e.g., rad52) are not rescued by spo13. This is because late gene functions are required to repair DSBs once they are formed, and the unrepaired DSBs that persist in a double mutant with spo13 are lethal. Early mutants are epistatic to late mutants in a spo13 background (i.e., a spo11 rad52 spo13 triple behaves similarly to a spo11 spo13 double mutant) because DSB repair functions become dispensable if no DSBs are made. This property has been used in genetic screens for mutations that abolish recombination initiation (Malone et al., 1991) as well as for mutants defective for meiotic DSB repair (McKee and Kleckner, 1997; Prinz et al., 1997). 2. Spo11 as the Double-Strand Break Catalyst Protein is covalently bound to the 5 -strand termini of the unresected DSBs that accumulate in rad50S and sae2 mutants, but not to the resected DSBs that accumulate transiently in wild-type strains (de Massy et al., 1995; Keeney and Kleckner,

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1995; Liu et al., 1995). Spo11 was shown to be the DSB-associated protein by direct purification and microsequencing of the covalent protein–DNA complexes (Keeney et al., 1997) and Spo11 was found to be homologous to a subunit of an archaeal topoisomerase (Bergerat et al., 1997). These observations strongly suggest that Spo11 is the catalytic subunit of the meiotic DNA cleaving activity and that it cuts DNA by a topoisomerase-like transesterification reaction to generate a covalent protein–DNA intermediate. It should be noted, however, that Spo11 protein has not been directly demonstrated to cleave DNA. 3. Evolutionary Conservation A large number of Spo11 homologs have been identified in archaebacteria and eukaryotes (Fig. 2; see color insert) but not in eubacteria. The archaeal homologs are all likely to function as type II topoisomerases, but topoisomerase activity has been directly demonstrated only for the Sulfolobus shibatae enzyme. Protein purified from S. shibatae cultures, or overexpressed and purified from E. coli, has ATP-dependent double-stranded DNA decatenating and relaxing activities (Bergerat et al., 1994; Buhler et al., 1998). Drugs that inhibit eukaryotic topoisomerase II by stabilizing a covalent protein–DNA complex also inhibit the purified archaeal enzyme (Bergerat et al., 1994). However, the mechanism of inhibition by these compounds appears to be different for the archaeal protein because they do not stabilize a covalent protein–DNA complex with this enzyme. The amino acid sequence of this new topoisomerase is unlike the previously known eukaryotic and prokaryotic type II enzymes and was named topoisomerase VI to distinguish it from these proteins (Bergerat et al., 1997). Topoisomerase VI is an A2B2 heterotetramer, the smaller subunit of which (called Top6A) shares similarity with Spo11. The Top6A subunit can bind DNA nonspecifically (Nichols et al., 1999), but does not catalyze detectable levels of DNA cleavage by itself (Buhler et al., 1998). The Top6B subunit contains an ATP-binding motif characteristic of type II topoisomerases, heat shock proteins, and mismatch repair proteins (Bergerat et al., 1997). The eukaryotic members of the Spo11/Top6A family all appear to be essential for meiotic recombination. Mutants in S. pombe (rec12 mutants; Lin and Smith, 1994), Drosophila melanogaster (mei-W68 mutants; McKim et al., 1998; McKim and Hayashi-Hagihara, 1998), and C. elegans (spo-11 mutants; Dernburg et al., 1998, 2000) exhibit no meiotic recombination and have increased levels of meiosis I chromosome nondisjunction. Interestingly, Arabidopsis thaliana has at least three SPO11 homologs (Hartung and Puchta, 2000; Grelon et al., 2001). Mutation of one of these, AtSPO11-1, diminishes meiotic crossing over by roughly 10-fold, but does not completely abolish it, suggesting that the different copies can partially substitute for one another (Grelon et al., 2001). Whether the three copies play additional, nonoverlapping roles as well is not currently known. spo11 mutants in the mushroom C. cinereus (Celerin et al., 2000) and in the mouse (Baudat et al., 2000; Romanienko and Camerini-Otero, 2000) arrest during meiotic prophase I

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and undergo programmed cell death, so it has not been possible to test directly for a defect in meiotic recombination. However, other lines of evidence strongly suggest that Spo11 is required for recombination in these organisms as well: (1) mutants in both organisms show chromosome structure defects similar to those in S. cerevisiae (Section III,A,6); (2) the C. cinereus mutant is partially rescued by X-irradiation, as are the S. cerevisiae and C. elegans mutants (Thorne and Byers, 1993; Dernburg et al., 1998); and (3) the mouse mutant fails to assemble chromosome-associated Dmc1/Rad51 complexes and the phosphorylated histone γ -H2AX (Baudat et al., 2000; Romanienko and Camerini-Otero, 2000; Mahadevaiah et al., 2001), similar to the S. cerevisiae mutant (Bishop, 1994). Potential SPO11 orthologs have also been identified in humans (Romanienko and Camerini-Otero, 1999; Shannon et al., 1999), and expressed sequence tags (ESTs) with sequence similarities to SPO11 have been isolated from rice, maize, and trypanosomes. A homologous sequence has also been identified in Neurospora crassa, but the predicted sequence currently available in sequence databases lacks several key residues thought to be critical for Spo11 catalytic activity (see below and Fig. 2). This discrepancy is probably due to misassignment of introns and exons, judging from analysis of the closely related sequence from Sordaria macrospora (A. Storlazzi and D. Zickler, personal communication, 2000). The role of these genes in meiosis has not yet been established. Alignment of available Spo11/Top6A family members (Fig. 2) reveals conservation over most of their lengths, with sequence identity of approximately 20–30% for most pairwise combinations. Several blocks of greater similarity correspond to portions of two structural domains discussed in more detail in the next section. The N-terminal sequences show little conservation, except in comparisons among the archaeal species or between mouse and human. The sizes of most of the family members fall in a fairly narrow range. Exceptions include Top6A from Pyrococcus horikoshii, which lacks the poorly conserved N-terminal domain, and the C. elegans Spo-11 protein, which has a long, C-terminal acidic tail. Attempts to cross-complement spo11 mutants in S. cerevisiae and S. pombe with genes cloned from other organisms have been unsuccessful (G. Smith, personal communication, 2000; P. Romanienko, personal communication, 2000; B. de Massy, personal communication, 2000). Perhaps this species specificity arises because of substantial divergence of protein interacting surfaces. Despite conservation of the Spo11/Top6A family, there is no obvious homolog of the Top6B protein in most eukaryotic genomes (A. thaliana appears to be the only exception to date). It is possible that an equivalent subunit exists but has diverged substantially from the archaeal proteins. Alternatively, if eukaryotic Spo11 proteins cleave DNA but do not act as topoisomerases, they may not require an associated ATP-binding subunit. ATP binding and hydrolysis by classic type II topoisomerases appear to drive conformational changes important for capturing a second DNA duplex and for opening and closing the DNA gate during strand transfer (Lindsley and Wang, 1993; Baird et al., 1999).

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4. Structure of Members of the Spo11/Top6A Family The crystal structure of a fragment of the Top6A subunit of Methanococcus ˚ resolution (Nichols et al., 1999) (Fig. 3; see color jannaschi was solved at 2.0-A insert). The structural model encompasses the portion that is most conserved in the Spo11/Top6A family (see Fig. 2), without the poorly conserved N-terminal domain. The model reveals similarities between secondary structure elements in Top6A and other topoisomerases that are otherwise unrelated at the sequence level, as well as critical differences in the tertiary arrangement of these elements. The structure also provides insight into how this family of proteins binds and cleaves DNA and points to potential protein-binding surfaces. Earlier studies identified two units of secondary structure in the type II topoisomerases of bacteria and eukaryotes (Berger et al., 1998); the enzymes from these organisms share substantial sequence similarity (Caron and Wang, 1994). These units are also found in two topoisomerases with dissimilar sequences: topoisomerase I from E. coli and archaeal Top6A. These motifs appear to be diagnostic of enzymes that generate 5 -phosphodiester linkages because they are not found in the type I topoisomerases of vaccinia virus or humans, which cleave DNA to form a 3 -phosphodiester linkage (Lima et al., 1994; Cheng et al., 1998). The first of these structural domains is an α-helical fold similar to the E. coli CAP (catabolite gene activator protein) DNA-binding domain (Schultz et al., 1991) (indicated in yellow in Figs. 2 and 3). In the eukaryotic and bacterial type II topoisomerases and in bacterial type I topoisomerases, this domain contains the tyrosine that attacks the DNA backbone (Berger et al., 1998). In the Spo11/Top6A family, this domain contains the only conserved tyrosine residue (highlighted in red in Fig. 2); mutation of this residue in Spo11 results in a null phenotype for meiotic recombination (Bergerat et al., 1997). Because the domain is common to all topoisomerases that generate a 5 -tyrosyl phosphodiester, it has been termed the “5Y-CAP” motif (Nichols et al., 1999). The structures of the 5Y-CAP domains from different topoisomerase families are highly superimposable, even though there is no detectable sequence homology between them (Berger et al., 1998; Nichols et al., 1999). The second domain consists of a four-stranded parallel β sheet sandwiched between two pairs of α helices (indicated in green in Figs. 2 and 3). This domain shows modest sequence similarity among different families of topoisomerases and corresponds to a sequence motif identified by an iterative database search seeded with the sequence of E. coli primase (Aravind et al., 1998). This motif has been termed the “Toprim” domain (for topoisomerases and primases). Only three residues—a glutamate and two aspartates—are conserved in nearly all Toprim-motif-containing proteins (highlighted in red in Fig. 2). They occur with a characteristic spacing and have been proposed to function in general acid/base chemistry or to bind a divalent metal ion known to be important for the activities of Toprim-containing enzymes.

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Consistent with the latter idea, a Mg2+ ion is bound by these three residues in the Top6A crystal (Nichols et al., 1999). Methanococcus jannaschi Top6A forms a U-shaped dimer with a channel about ˚ wide and 50 A ˚ long thought to be where DNA binds to the protein (Fig. 3). 18 A The conserved tyrosine in the 5Y-CAP domain and the metal-binding pocket of the Toprim domain are presumably part of the active site. Mutagenesis studies support this conclusion for all of the families of 5 -cleaving topoisomerases (Chen and Wang, 1998; Liu and Wang, 1998; Diaz et al., 2001; P. C. Varoutas and A. Nicolas, personal communication, 2000). In the Top6A structure, the catalytic tyrosine of each monomer lies closer to the Toprim metal-binding pocket of its dimer partner than it does to its own Toprim domain. This arrangement suggests that the dimer forms two hybrid active sites, each responsible for cleaving a single strand of the DNA duplex. (One of the active sites is circled in Fig. 3B.) Similar hybrid active sites are thought to form in eukaryotic and prokaryotic type II topoisomerases (Berger et al., 1998). Consistent with this idea, yeast topoisomerase II heterodimers formed between mutant and wild-type protomers or between two different mutant protomers (e.g., one carrying a mutation in the 5Y-CAP domain and the other a mutation in the Toprim domain) assemble one mutant active site and one fully wild-type active site and are capable of nicking DNA (Liu and Wang, 1998, 1999). This model also predicts that active site mutants should confer a semidominantnegative phenotype when coexpressed with wild type. This prediction is met for SPO11 under at least some conditions (Diaz et al., 2001; P. C. Varoutas and A. Nicolas, personal communication, 2000). The Top6A structure provides clues about potential surfaces on Spo11 that might be involved in protein–protein interactions. Berger and colleagues proposed that each of a pair of Top6B subunits binds to an α-helical region on the upper surface of each Top6A subunit (arrows in Fig. 3A) (Nichols et al., 1999). Although there is no clear homolog of Top6B in most eukaryotes, the equivalent portion of Spo11 is an attractive candidate for a protein interaction surface. The archaeal proteins show significant sequence conservation across this region, but the eukaryotic Spo11 sequences are more highly diverged (Fig. 2). Perhaps the divergence in this region contributes to the observed inability of Spo11 homologs to support interspecific cross-complementation (Section III,A,3). 5. The Mechanism of DNA Cleavage by Spo11 Early reports conflicted as to whether Spo11-generated DSBs have 5 overhangs (Liu et al., 1995; Xu and Kleckner, 1995) or blunt ends (de Massy et al., 1995; Xu and Petes, 1996). More recent analysis (B. de Massy, personal communication, 2000) suggests that the primary cleavage species has a 2-nucleotide 5 overhang that is sometimes filled in by DNA polymerase to yield blunt ends. Whether this fill-in

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event occurs in vivo or is an artifact of the DNA isolation procedure is not currently known. In the Top6A structural model, the catalytic tyrosine is located about ˚ from the Toprim metal-binding pocket and is far from the sugar–phosphate 9A backbone when a DNA duplex is modeled into the putative DNA-binding channel. If a 2-nucleotide 5 overhang is assumed for the cleavage product, the scissile phosphodiester bonds lie close to the metal-binding pockets (Fig. 3B). To allow the catalytic tyrosines to attack these bonds, the 5Y-CAP domains may flex inward relative to the Toprim domains (Nichols et al., 1999). DNA cleavage and religation by topoisomerases are isoenergetic. The religation reaction is normally faster than the cleavage reaction, so equilibrium favors the intact DNA duplex, but certain circumstances can perturb this balance. Several topoisomerase inhibitors stabilize the cleaved DNA complex, perhaps by slowing the rate of religation or by sterically blocking it (Wang, 1994; Fortune and Osheroff, 2000). Substituting Ca2+ for Mg2+ can also have this effect for some enzymes. In E. coli topoisomerase IV, a mutation in the ParE subunit that destabilizes the ParE–ParC interaction generates an enzyme with hyper-DNA cleavage activity. It is thought that the enzyme falls apart after DNA cleavage, leaving the ParC subunit covalently attached to the DNA and thus preventing religation (Mossessova et al., 2000; Nurse et al., 2000). These characteristics of other topoisomerases raise questions about Spo11 activity. For example, is the Spo11 DNA cleavage reaction reversible? If so, what drives the reaction forward in wild-type cells? What is the molecular basis for the stabilization of the Spo11–DNA complex in rad50S mutants? 6. Does Spo11 Play a Role in Nonmeiotic Cells? SPO11 orthologs in S. cerevisiae, S. pombe, and C. cinereus are expressed only on entry into meiosis and no mitotic phenotypes have been described in these organisms (Atcheson et al., 1987; Lin and Smith, 1994; Celerin et al., 2000). Similarly, nematodes that lack germ cells do not express Spo-11 (Dernburg et al., 1998). Expression is not restricted to germ line or meiotic cells in all organisms, however. Mouse and human Spo11 mRNAs were detected in several adult somatic tissues, including brain and thymus, although at significantly lower levels than in testis or fetal ovary (Keeney et al., 1999; Romanienko and Camerini-Otero, 1999). It is not known whether functional Spo11 protein is made in these nonmeiotic tissues. Similarly, at least one SPO11 homolog in A. thaliana (AtSPO11-1) is expressed in both meiotic and somatic tissues (Hartung and Puchta, 2000). In D. melanogaster, Mei-W68 transcripts are readily detectable in somatic tissues at several larval stages, but expression in adult flies is limited to the ovaries (McKim and Hayashi-Hagihara, 1998). In mei-W68 mutants, the frequency of recombinant somatic clones in the abdomen and of mitotic crossing over in the male germ line are increased (Lutken and Baker, 1979; McKim and HayashiHagihara, 1998). Thus, paradoxically, a mei-W68 mutation confers a hypo

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recombination phenotype in female meiosis as well as a hyper recombination phenotype in some mitotically dividing cells.

B. RAD50, MRE11, XRS2, and SAE2/COM1 RAD50, MRE11, and XRS2 are unique among the many genes required for meiotic DSB formation in S. cerevisiae because they are also required for DNA metabolic events in vegetative cells. They are involved in telomere maintenance, cell cycle responses to DNA damage, and DSB repair by both homologous recombination and nonhomologous end joining (reviewed in Haber, 1998; Paques and Haber, 1999; Petrini, 1999). Null mutants grow slowly, are hypersensitive to DNA-damaging agents, and show an increase in spontaneous recombination. In meiotic cells, these genes are required for DSB formation and resection. Whether they can also function as dessert toppings and floor waxes, as recently suggested (Haber, 1998), has yet to be definitively addressed. This section focuses primarily on properties of these genes critical for meiotic recombination. 1. Biochemical Properties and Evolutionary Conservation Yeast Rad50, Mre11, and Xrs2 proteins make a physical complex (Johzuka and Ogawa, 1995; Bressan et al., 1998; Usui et al., 1998), as do mammalian Rad50, Mre11, and Nbs1 proteins (Dolganov et al., 1996; Carney et al., 1998; Paull and Gellert, 1998; Trujillo et al., 1998). Nbs1 and Xrs2 are similar in size but share little if any sequence similarity, so it is not known whether they are functionally equivalent to one another. Rad50 and Mre11 are essential for cell viability in vertebrates (Xiao and Weaver, 1997; Luo et al., 1999; Yamaguchi-Iwai et al., 1999). Hypomorphic alleles of human NBS1 cause an inherited chromosome instability disorder, Nijmegen breakage syndrome, and mutations of MRE11 cause an ataxia telangiectasia-like disorder (Carney et al., 1998; Varon et al., 1998; Stewart et al., 1999). Both syndromes exhibit cellular hypersensitivity to X rays. MRE11 orthologs are also required for radiation resistance in S. pombe (rad32+; Tavassoli et al., 1995), C. cinereus (Gerecke and Zolan, 2000), N. crassa (mus-23; Watanabe et al., 1997), and C. elegans (A. Villeneuve, personal communication, 2000). Thus it appears that many of the somatic roles of these proteins in DNA repair are conserved. Rad50 and Mre11 are homologous to the E. coli SbcC and SbcD proteins, respectively, and to the bacteriophage T4 proteins gp46 and gp57, respectively (Leach et al., 1992). SbcC and SbcD proteins form a complex with properties similar to those of the Rad50-containing complex (see below). However, prokaryotes and phage T4 have no obvious equivalent of Xrs2 or Nbs1. The S. cerevisiae RAD50 gene product is a 152.4-kDa protein with a consensus Walker-type nucleotide-binding motif and a long central region of heptad repeats

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(Alani et al., 1989). This general arrangement places Rad50 in the structural maintenance of chromosomes (SMC) family of proteins, which play roles in chromosome condensation and segregation in eukaryotes and prokaryotes (Hirano, 1999). From crystallographic and electron microscopy (EM) studies of purified proteins (Melby et al., 1998; Hopfner et al., 2000), SMC family members appear to form head-to-tail dimers in which the N-terminal globular domain of one protomer (containing the consensus Walker A box) associates with the C-terminal globular domain of the other protomer (containing the consensus Walker B box). The central heptad repeat regions of the two protomers are thought to associate with one another to form an α-helical coiled coil containing a flexible hinge. Consistent with this structural organization, purified yeast Rad50 protein forms dimers and exhibits ATP-dependent DNA-binding activity (Raymond and Kleckner, 1993). The 77.6-kDa Mre11 protein and its homologs contain sequences conserved in a diverse group of phosphodiesterases (Sharples and Leach, 1995). The presence of these sequence elements, in combination with earlier studies suggesting that T4 gp46 and gp47 had associated nuclease activities (Kutter and Wiberg, 1968; Prashad and Hosoda, 1972), led to the demonstration that the E. coli SbcCD complex has ATP-dependent 3 → 5 double-strand exonuclease and ATP-independent single-strand endonuclease activities and that it can cleave stem–loop structures at the 5 end of the loop. All nuclease activities require Mn2+ as a cofactor (Connelly and Leach, 1996; Connelly et al., 1997, 1998, 1999). Similarly, human and budding yeast Mre11, by themselves or in complex with Rad50, have Mn2+-dependent single-strand endonuclease, 3 → 5 double-strand exonuclease, and stem–loop opening activities (Furuse et al., 1998; Paull and Gellert, 1998; Trujillo et al., 1998; Usui et al., 1998; Moreau et al., 1999). Human Mre11 exonuclease activity is stimulated by Rad50, but ATP is neither required nor stimulatory, in contrast to SbcCD. Adding Nbs1 to the complex reveals several new activities, including partial unwinding of duplex DNA and cleavage of fully paired hairpins (as opposed to stem–loop structures) (Paull and Gellert, 1999). Both of these activities are stimulated by ATP. All of the Mre11 nucleolytic activities in vitro require nonphysiological Mn2+ concentrations. This might indicate that some unidentified factor(s) is required to allow Mre11 to use another metal cofactor or to form a stable manganesecontaining metalloenzyme. The observed 3 → 5 polarity of the Mre11 exonuclease is also surprising because genetic and physical analyses of meiotic recombination and mitotic DSB repair had long suggested that Mre11, Rad50, and Xrs2 are involved in 5 → 3 exonucleolytic resection (Paques and Haber, 1999). One possible explanation for this paradox is that the Mre11 complex is not the main resection exonuclease, but instead recruits or controls other exonuclease activities (Paques and Haber, 1999; Paull and Gellert, 1999). One candidate for such a meiotic DSB resection activity is the 5 → 3 exonuclease product of the EXO1 gene. Exo1 protein functions in vegetative cells in mismatch repair and mitotic recombination (Szankasi and Smith, 1995; Fiorentini et al., 1997; Symington et al.,

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2000) and is required for normal DSB processing and recombination in meiotic cells (Khazanehdari and Borts, 2000; Tsubouchi and Ogawa, 2000). Exo1 cannot be the only such activity, however, because DSB resection does still occur in an exo1 mutant. 2. Roles in Meiotic Recombination The prominent meiotic phenotypes of S. cerevisiae rad50, mre11, and xrs2 null mutants are largely the same as those described above for spo11 mutants: no DSBs, no meiotic recombination, and extremely low spore viability (Malone and Esposito, 1981; Alani et al., 1990; Cao et al., 1990; Ajimura et al., 1992; Ivanov et al., 1992). RAD50 and MRE11 are also required for the resection of DSBs, on the basis of characterization of nonnull alleles. It is not known whether XRS2 is also required for DSB resection. a. Point Mutations of RAD50. Random mutagenesis of the budding yeast RAD50 coding sequence generated 10 rad50S (for “Separation-of-function”) alleles that were meiosis defective but conferred nearly wild-type resistance to DNA damage (Alani et al., 1990). The meiotic defect in rad50S is distinct from rad50: DSBs are made, but they accumulate in an unresected form and do not give rise to recombinant DNA products (Alani et al., 1990; Cao et al., 1990). The mutated residues lie near the N-terminal Walker A box or the C-terminal Walker B box in the primary sequence. Mapping several of these residues onto the crystal structure of an archaeal Rad50 homolog suggests that they reside in a surface patch that may be involved in protein–protein interactions (Hopfner et al., 2000). One known temperature-sensitive allele (rad50-ts1) is hypersensitive to methyl methane sulfonate (MMS) at the restrictive temperature (similar to a rad50 mutant) but produces unresected DSBs in meiosis (like rad50S) (Alani et al., 1990). This allele encodes two amino acid changes in the second heptad repeat region. Two site-directed mutants within the Walker A box consensus have phenotypes indistinguishable from rad50, suggesting that ATP binding is critical for Rad50 function (Alani et al., 1990). The rad50S mutations have proved useful for mapping and quantitating DSBs because the unresected DSBs do not turn over, because they give sharper bands on Southern blots, and because DSB-proximal DNA sequences can be purified by virtue of their covalent association with Spo11 protein (e.g., Baudat and Nicolas, 1997; Gerton et al., 2000). This use begs the question of whether the amount and distribution of DSBs are the same in the mutants as in wild type. By some criteria, the DSBs in rad50S strains appear to be faithful reporters of wild-type break patterns. The relative strengths of many DSB sites are similar in RAD50 and rad50S (e.g., Cao et al., 1990; de Massy et al., 1995; Liu et al., 1995). Also, the locations of DSB 3 -strand termini are the same in both genotypes, suggesting that cleavage specificity is unaltered (de Massy et al., 1995; Liu et al., 1995).

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However, other observations indicate that there may be critical differences. For example, a rad50S mutant gave different break patterns across a large section of Chromosome III when compared with another mutant that accumulates DSBs, dmc1 (Dresser et al., 1997). Moreover, DSBs in late-replicating regions of the genome appear to be specifically reduced in rad50S (and sae2) mutants (Borde et al., 2000). This apparent lack of neutrality may account in part for observed discrepancies between the genetic map and the rad50S DSB map in certain regions of Chromosome III (Baudat and Nicolas, 1997; see also Borde et al., 1999). In any case, it is important to be aware of this caveat when interpreting rad50S-based DSB measurements. In rad50S cells, Rad50, Mre11, and Xrs2 proteins colocalize with one another in cytologically observable complexes (called foci) on spread meiotic chromosomes (Usui et al., 1998). However, significantly fewer foci were observed than expected for the number of meiotic DSBs (30–35 foci versus about 200 DSBs per cell), the intensity of immunostaining varied greatly from focus to focus in the same nucleus, and foci could not be detected on spread meiotic chromosomes from wild-type cells. Thus, the molecular nature of these foci is not clear. They might represent coalescence of several sites at which DSB formation has occurred. Alternatively, the foci might reflect a response to unrepairable DSBs unlike any events that occur during normal meiotic recombination. b. Nonnull Alleles of MRE11. The spectrum of phenotypes obtained with nonnull alleles of MRE11 is more complex, but the following generalizations appear to hold: (1) Mre11 must be able to form complexes with Rad50 and Xrs2 for efficient DNA repair in vegetative cells, but not for formation of meiotic DSBs; (2) Mre11 nuclease activity is not required for most roles in vegetative cells, or for DSB formation in meiosis, but is required for meiotic DSB resection; (3) the C-terminal region of Mre11 is dispensable for mitotic DNA repair and meiotic DSB resection, but is essential for meiotic DSB formation. The “mre11S” allele was isolated in a screen for mutations that caused SPO11dependent lethality during meiosis (Nairz and Klein, 1997). mre11S and rad50S mutants have similar phenotypes: vegetative resistance to DNA-damaging agents is nearly (but not quite) normal and meiotic DSBs accumulate in an unresected form. The mre11S mutation provided the first evidence that MRE11, like RAD50, is involved in DSB resection. This conclusion was confirmed by characterization of a previously identified radiation-sensitive mre11 allele originally designated rad58 (Chepurnaya et al., 1995; Tsubouchi and Ogawa, 1998). This “mre11-58” mutant blocks DSB resection, but is unlike mre11S in that it is hypersensitive to DNA damage during vegetative growth. Both mre11S and mre11-58 contain substitutions within the conserved phosphodiesterase motifs, and the mre11-58 protein is nuclease defective in vitro, suggesting that absence of nuclease activity might underlie the mutant effects (Nairz and Klein, 1997; Tsubouchi and Ogawa, 1998). Several groups targeted the conserved phosphodiesterase motifs for mutagenesis (Bressan et al., 1998; Furuse et al., 1998; Moreau et al., 1999). In general, these

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mutants fall into two classes: those similar to mre11S (DNA damage-resistant, unresected meiotic DSBs) and those similar to mre11-58 (DNA damagehypersensitive, unresected meiotic DSBs). The products of two of the mre11S-like alleles, mre11-D56N and mre11-H125N, are nuclease defective in vitro (Moreau et al., 1999). Thus, the nuclease function of Mre11 is essential for meiotic DSB resection, but is dispensable for most of the functions of the protein in mitotic DNA repair. However, the weak MMS and ionizing radiation sensitivity of these mutants may indicate that Mre11 nuclease is required to process a subset of damaged DNA ends (Moreau et al., 1999; Paques and Haber, 1999). Where tested, all of the phosphodiesterase mutants were nuclease defective, so why are there differences in the DNA repair phenotypes? Mutant alleles such as mre11-58 that confer DNA damage sensitivity also show defects in binding to Rad50 and/or Xrs2 (Bressan et al., 1998; Usui et al., 1998; Chamankhah and Xiao, 1999). Perhaps Mre11–Rad50–Xrs2 complex formation is critical for mitotic DNA repair, but is dispensable for formation (if not resection) of meiotic DSBs. Whether Rad50 and Xrs2 can associate with the products of the meiosis-defective, repair-proficient alleles (such as mre11S, mre11-D56N, and mre11-H125N ) has not yet been reported. Several other alleles specifically affect meiotic DSB formation. C-terminal truncation mutants mre11-5 (Usui et al., 1998) and mre11C49 (Furuse et al., 1998) show nearly wild-type resistance to DNA damage, but cannot support formation of meiotic DSBs. The C-terminal region of Mre11 interacts with at least three proteins present in extracts from meiotic cells but not from vegetative cells (Usui et al., 1998). These interactions may be important for DSB formation but the meiosis-specific polypeptides have not yet been identified. A separate C-terminal truncation allele (mre11-T4) was generated by transposon mutagenesis (Nairz and Klein, 1997). Although the DSB phenotype of this allele has not been reported, the sporulation phenotype suggests that it is DSB defective. (mre11-T4, like rad50 and mre11 null mutants, sporulates well, although spore viability is low. In contrast, rad50S and mre11S mutants sporulate poorly.) Unlike the mre11-5 and mre11C49 truncation alleles, however, mre11- T4 mutants are hypersensitive to DNA damage. Under some conditions, mre11S and mre11-T4 show interallelic complementation for DNA damage resistance and spore viability, presumably reflecting the ability of Mre11 to selfassociate (Nairz and Klein, 1997). This result suggests that mutant proteins that cannot support DSB formation (because of a C-terminal truncation), can support DSB resection. c. SAE2/COM1. Two independent screens for mutations that conferred SPO11-dependent sporulation defects identified the SAE2/COM1 gene (for Sporulation in the Absence of Spo Eleven, or Completion of meiotic recombination) (McKee and Kleckner, 1997; Prinz et al., 1997). The phenotypes of sae2/com1 null mutants are identical to rad50S mutants in nearly every respect. No motifs suggestive of biochemical function are apparent in the 40.0-kDa Sae2 protein

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sequence, and there are no homologs in available databases. The similar phenotypes of sae2/com1, rad50S, and mre11S mutants suggest that Sae2 functions as part of, or regulates the activity of, the Mre11–Rad50–Xrs2 complex, but this idea has not been directly tested. d. Evolutionary Conservation of Roles in Meiotic Recombination. The function of Rad50 and Mre11 in meiosis appears to be evolutionarily conserved. In C. elegans, mre-11 mutations confer a meiotic phenotype largely similar to that of spo-11 mutants, except that the meiotic defects in mre-11 mutants are not rescued by γ irradiation. In fact, irradiation reduces the production of viable progeny in the mre-11 mutants, consistent with an additional role subsequent to DSB formation (perhaps similar to that in S. cerevisiae) (A. Villeneuve, personal communication, 2000). In rodent spermatocytes, Rad50 and Mre11 localization are consistent with a role for these proteins in meiotic recombination (Goedecke et al., 1999; Eijpe et al., 2000), but the cell-lethal phenotype of targeted disruptions of these genes has hindered a direct genetic test thus far. The mre11-1 mutation in C. cinereus confers meiotic defects in homologous pairing, chromatin condensation, and synaptonemal complex (SC) formation (Gerecke and Zolan, 2000). As for spo11 mutants in this organism, mre11 mutants do not progress past metaphase I, so it is not known whether recombination is affected. However, the similarities of mre11-1 phenotypes to spo11 mutants in C. cinereus and to mre11 null mutants in S. cerevisiae make this likely. The N. crassa MRE11 ortholog mus-23 is also required for meiosis and sporulation (Watanabe et al., 1997). In S. pombe, deletion of the N-terminal two-thirds of the Mre11 ortholog Rad32 results in a 10- to 15-fold decrease in meiotic recombination and a reduction in spore viability to 0.5% (Tavassoli et al., 1995). Fox and Smith (1998) have pointed out that this spore viability defect is more severe than expected for a recombination initiation mutant—random segregation of the three chromosomes of S. pombe in the absence of meiotic recombination (e.g., in a rec12 or rec6 mutant) is sufficient to support a relatively high spore viability of about 20%. These findings raise the possibility that rad32+ may be required for DSB processing but not for DSB formation in fission yeast. Consistent with this interpretation, combining a rad32 mutation with a mutation in rec6 resulted in an increase in spore viability to values typical for the rec6 mutation alone (J. Bedoyan, unpublished results cited in Fox and Smith, 1998). A mutation equivalent to the S. cerevisiae mre11S mutation had no effect on rad32 function, but other mutations in the conserved phosphodiesterase motifs of rad32 resulted in recombination and spore viability phenotypes indistinguishable from the partial deletion mutation (Wilson et al., 1998). These results support the idea that Rad32-associated nuclease activity is critical for its function in meiosis, but biochemical analysis has not yet been reported for wild-type or mutant proteins.

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C. Other Genes That Are Absolutely Required for Double-Strand Break Formation in Budding Yeast 1. MER1, NAM8/MRE2, and MER2/REC107 MER1 (for MEiotic Recombination) was originally identified in a screen for mutants that produce inviable spores (Rockmill and Roeder, 1988; Engebrecht and Roeder, 1989). Null mutants are less recombination defective than spo11 mutants, and spo11 and rad50 are epistatic to mer1 for meiotic intragenic recombination (Engebrecht and Roeder, 1989). The MER1 transcript is induced in meiosis and encodes a 31-kDa protein with no known homologs. Mutations in MRE2 (which also stands for Meiotic REcombination) were isolated in a screen for recombination-defective mutants in a haploid strain disomic for Chromosome III (Ajimura et al., 1992). Null mutants have phenotypes similar to those of mer1 mutants (Ajimura et al., 1992; Ogawa et al., 1995; Nakagawa and Ogawa, 1997). MRE2 is identical to NAM8, isolated as a multicopy suppressor of a mitochondrial splicing defect (Ekwall et al., 1992). MER2 was isolated as a multicopy suppressor of the intragenic recombination defect in mer1 (Engebrecht et al., 1990) and mre2 mutants (Nakagawa and Ogawa, 1997). It was also identified independently (as REC107) in a screen for mutations that rescued the meiotic lethality in rad52 spo13 haploid strains (Malone et al., 1991). Null mer2 mutants are severely recombination defective and produce inviable spores because they cannot make DSBs (Engebrecht and Roeder, 1990; Malone et al., 1991; Cool and Malone, 1992; Rockmill et al., 1995). The 35.5-kDa Mer2 protein has no obvious homologs in other organisms but it shares sequence similarity with myosin-related proteins within a stretch of heptad repeat sequence predicted to form α-helical coiled coil (Rockmill et al., 1995). The MER2 transcript contains an 80-nucleotide intron that is inefficiently spliced in vegetative cells, at least in part because of a noncanonical 5 splice site sequence (Engebrecht et al., 1991; Nandabalan et al., 1993). Mer1 and Nam8 proteins stimulate efficient removal of this intron during meiosis (Engebrecht et al., 1991; Nandabalan and Roeder, 1995; Ogawa et al., 1995; Nakagawa and Ogawa, 1997). Mer1 appears to act by binding to a splicing enhancer in the MER2 transcript and physically interacting with U1 snRNP (Spingola and Ares, 2000). Nam8 is required for Mer1 to activate splicing and is itself a nonessential U1 snRNPassociated protein (Spingola and Ares, 2000). Overexpression of wild-type MER2 or expression of an intronless version completely suppresses the DSB defects of mer1 and nam8/mre2 mutants (Engebrecht et al., 1991; Nakagawa and Ogawa, 1997). Thus, the roles of MER1 and NAM8/MRE2 in DSB formation are limited to promoting the splicing of MER2. However, an intronless copy of MER2 does not fully suppress the crossover or spore viability defects of mer1 and nam8/mre2 mutants (Engebrecht et al., 1990; Storlazzi et al., 1995; Nakagawa and Ogawa, 1997), presumably because Mer1 and Nam8 are required for meiosis-specific

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splicing of other mRNAs, including the MER3 and SPO70 transcripts (Chu et al., 1998; Nakagawa and Ogawa, 1999; Davis et al., 2000; Spingola and Ares, 2000). No vegetative cell defects have been described for mer2 mutants, so it is a puzzle why cells use this unique splicing mechanism to control the expression of MER2. 2. MEI4 MEI4 (for MEIosis-specific) was first identified in a screen for UV-induced mutants defective for intragenic meiotic recombination (Menees and Roeder, 1989). Null mutants are resistant to MMS but are defective for meiotic recombination and heteroduplex DNA formation and are epistatic to rad52 mutations (Menees and Roeder, 1989; Menees et al., 1992; Nag et al., 1995). Sporulation occurs efficiently, but spores are inviable and the inviability can be rescued by a spo13 mutation. As expected for mutants with this array of phenotypes, mei4 strains make no detectable meiotic DSBs (Keeney et al., 1997; Jiao et al., 1999). MEI4 is not required for transcriptional induction of other DSB genes (e.g., SPO11). Its transcript is expressed only in meiosis and contains a single intron whose splicing is not dependent on MER1 (Menees et al., 1992). The 48.1-kDa Mei4 protein has no obvious homologs or sequence motifs suggestive of a biochemical function. 3. REC102, REC104, and REC114 Mutations in REC102, REC104, and REC114 were identified in a screen for suppressors of the meiotic lethality in rad52 spo13 haploids (Malone et al., 1991). REC102 was also isolated in a screen for sporulation-proficient, meiotic lethal mutants (Bhargava et al., 1992), and REC114 was independently isolated in a screen for meiotic recombination defects in haploids disomic for Chromosome III (Ajimura et al., 1992). Null mutants are recombination defective and produce inviable spores and spore inviability is rescued by spo13 (Malone et al., 1991; Bhargava et al., 1992; Cool and Malone, 1992; Galbraith and Malone, 1992; Pittman et al., 1993; Mao-Draayer et al., 1996). All three are required for meiotic DSB formation (Bullard et al., 1996). Interestingly, overexpression of REC114 inhibits DSB formation, suggesting that the dosage of Rec114 protein is critical for proper function of the DSB machinery (Bishop et al., 1999). Transcription of all three genes is meiosis specific (Cool and Malone, 1992; Galbraith and Malone, 1992; Pittman et al., 1993). REC102 encodes a 23.2-kDa nuclear-localized protein with no known homologs. Rec102 interacts physically and genetically with Spo11 (see Section III,F), and associates with meiotic chromosomes during early meiotic prophase in a Spo11-dependent manner (Kee and Keeney, 2001; and our unpublished results, 2000). REC104 encodes a 20.6-kDa protein, the only known homologs of which were isolated from other Saccharomyces species (S. paradoxus and S. pastorianus).

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Both homologs complement an S. cerevisiae rec104 mutant (Nau et al., 1997). REC114 encodes a 49.5-kDa protein. Homologs from S. paradoxus and S. pastorianus complement an S. cerevisiae rec114 mutant (Malone et al., 1997). The REC114 transcript contains an intron that is unusual in two regards: (1) it is located near the 3 end of the coding region, whereas most introns in budding yeast are located near the 5 end; and (2) splicing uses a noncanonical 3 splice site (AAG instead of CAG) (Pittman et al., 1993; Malone et al., 1997). Splicing of this intron does not require MER1. A portion of Rec114 protein shares modest similarity with the meiotically induced S. pombe Rec7 protein (Malone et al., 1997; Fox and Smith, 1998). rec7 mutants are severely defective for meiotic recombination and have reduced spore viability, but show no detectable mitotic defects (Ponticelli and Smith, 1989; Lin et al., 1992; Fox and Smith, 1998). It is not known whether this sequence similarity reflects functional conservation. None of the proteins has sequence motifs that suggest a biochemical function. 4. SKI8/REC103 The same screen that identified rec102, rec104, and rec114 mutants also identified a recombination-defective mutant that was named rec103 (Malone et al., 1991). Where tested, the meiotic recombination phenotypes of rec103 mutants were indistinguishable from those of the other rec mutants (Gardiner et al., 1997). REC103 is presumably required for DSB formation, but direct analysis has not been reported. REC103 is identical to SKI8 (for Super-Killer). ski8 mutants derepress the replication of double-stranded RNA viruses and sensitize cells to the toxic effects of the increased viral load (Wickner, 1996). SKI8/REC103 is required to inhibit translation of nonpolyadenylated RNA (including that of L-A and M viruses), probably because it is required for 3 → 5 exonucleolytic RNA degradation (Masison et al., 1995; Jacobs et al., 1998). It is not clear how this RNA metabolism role relates to a role in meiotic DSB formation. It is possible that the effects of ski8/rec103 mutations on meiotic recombination are indirect, via effects on expression of other DSB genes. Alternatively, Ski8 might play a direct role in DSB formation. A two-hybrid interaction between Ski8 and Spo11 has been reported (Uetz et al., 2000). Ski8 is similar to the S. pombe Rec14 protein, which is also required for meiotic recombination (Evans et al., 1997; Fox and Smith, 1998). rec14+ is expressed in vegetative cells, and mutants have a slow-growth phenotype. It is possible that SKI8/REC103 and rec14+ perform similar roles during meiosis and in mitotically dividing cells. It will be interesting to see whether S. pombe Rec14 and Rec12 proteins physically interact. Ski8 (44.0 kDa) and Rec14 (32.9 kDa) each contain multiple repeats of the “WD” motif originally identified in β-transducin (Matsumoto et al., 1993; Evans et al., 1997). WD motifs are found in a functionally diverse range of proteins and are thought to mediate protein–protein interactions (see Smith et al., 1999, for a review).

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D. Roles of Double-Strand Break Genes in Development of Meiotic Chromosome Structure, Homologous Chromosome Pairing, and Progression through Meiotic Prophase Mutants defective for SPO11 or other of the genes required for DSB formation show several phenotypes in addition to a complete absence of homologous recombination and the resulting defects in chromosome segregation. Many of these phenotypes are observed in only a subset of organisms for which the relevant mutation(s) has been described. Some phenotypes appear to reflect secondary consequences of a lack of DSBs. Others may represent roles for the affected gene product independent of its role in DSB formation.

1. Alterations in Higher Order Chromosome Structures Meiotic cells develop a series of specialized higher order chromosome structures (Roeder, 1997; Moens et al., 1998; Zickler and Kleckner, 1998, 1999). Early in meiotic prophase, pairs of sister chromatids develop short stretches of proteinaceous axial structure termed axial elements (AEs) that can be observed cytologically by immunofluorescence or electron microscopy. As prophase continues, the AE segments of a given chromosome elongate and coalesce as they become juxtaposed (“synapsed”) with the AE of its homologous partner, forming a tripartite structure known as the synaptonemal complex (SC), which eventually extends the length of each chromosome pair. The SC consists of a pair of coaligned AEs (now termed lateral elements, or LEs) held together by a proteinaceous central element. Spherical or ovoid structures (ranging from 30 to 200 nm in diameter) called recombination nodules are associated with AEs and SCs. They contain recombination factors such as the strand exchange proteins Dmc1 and Rad51 and are presumed to be the sites where meiotic recombination occurs (reviewed in Zickler and Kleckner, 1999). In S. cerevisiae, SPO11 is required for normal SC formation (Giroux et al., 1989; Loidl et al., 1994). [Note, however, that the extent of the synapsis defect in spo11 mutants varies among published reports (cf. Klapholz et al., 1985). Reports also vary as to whether SPO11 is required for AE formation (Giroux et al., 1989; Loidl et al., 1994). The reason for these differences is not known.] The presence or absence of Spo11 also influences the outcome of recombinational repair of a DSB made by the HO endonuclease ectopically expressed during meiosis, perhaps reflecting a role for Spo11 in promoting formation of a meiotic chromosome structure that directs the outcome of DSB repair (Malkova et al., 2000). Other DSB genes are also required for SC formation: mer2, mei4, and rec102 mutants each make long AEs that fail to synapse (Bhargava et al., 1992; Menees et al., 1992; Rockmill et al., 1995). Whether REC104, REC114, or SKI8/REC103 is required has not been determined. mer1 mutants are defective for SC formation, but this

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defect is suppressed by MER2 overexpression, suggesting that the role of MER1 in SC formation is limited to its role in facilitating splicing of the MER2 transcript (Engebrecht et al., 1990). Null mutants for RAD50 confer a severe chromosome structure defect. By EM analysis, only short stretches of AE were formed in rad50 strains, with no tripartite SC detected (Alani et al., 1990; Loidl et al., 1994). Effects of mre11 or xrs2 mutations have not been described for S. cerevisiae, but the mre11-T4 mutation confers a similar phenotype to rad50, except that rare mre11-T4 nuclei contained tripartite SC with frequent partner switches indicative of nonhomologous synapsis (Nairz and Klein, 1997). As discussed in Section III,B,2,b, this mutation is likely to cause a defect in DSB formation. AE formation is more extensive in rad50S strains than in rad50, with many nuclei showing fairly long stretches of AE (Alani et al., 1990; Loidl et al., 1994). Tripartite SC formation is defective in rad50S, although the extent of synapsis varied between different studies (Alani et al., 1990; Loidl et al., 1994). Fairly extensive AE formation was also observed in mre11S and sae2/com1 mutants, with a fraction of nuclei showing significant levels of tripartite SC, at least some of which was between nonhomologous chomosomes (Nairz and Klein, 1997; Prinz et al., 1997). Based in part on comparisons between the rad50 and rad50S phenotypes, it has been argued that meiotic recombination and meiotic chromosome synapsis are intimately coupled to one another in budding yeast, either because they share common molecular steps (such as a search for homology) or because of regulatory events that coordinate them (Alani et al., 1990). The different effects of the earlier recombination block (rad50) compared with the later recombination block (rad50S) may indicate that recombination and chromosome structure “pathways” are interdigitated at more than one point. Studies suggest that the Zip2 and Zip3 proteins may play a direct role in coupling recombination to the initiation of synapsis (Chua and Roeder, 1998; Agarwal and Roeder, 2000). These proteins, which are required for SC formation, colocalize with Mre11 on spread meiotic chromosomes. Moreover, Zip3 interacts physically with Zip2 as well as with Mre11 and other meiotic recombination proteins (Agarwal and Roeder, 2000). SC formation requires meiotic recombination functions in a number of other organisms as well. Spo11 is required for normal SC formation in C. cinereus and mouse (Baudat et al., 2000; Celerin et al., 2000; Romanienko and Camerini-Otero, 2000). Likewise, a C. cinereus mre11 mutant shows less extensive AE formation than wild type, with incomplete, nonhomologous synapsis (Gerecke and Zolan, 2000). Moreover, several other mouse genes presumed to be required for meiotic recombination are also required for normal homologous synapsis, similar to Spo11: these include Dmc1, Msh4, and Msh5 (Pittman et al., 1998; Yoshida et al., 1998; de Vries et al., 1999; Edelmann et al., 1999; Kneitz et al., 2000). Mutation of AtSPO11-1 in A. thaliana also appears to disrupt SC formation (Grelon et al., 2001). Thus, it appears that formation and correct processing of homologous recombination intermediates are necessary to promote proper homologous synapsis

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in these organisms. However, it should be noted that significant amounts of SC do form in many of these instances, indicating that recombination is not essential for synapsis per se. Moreover, the requirement for Spo11 and Rad50 in SC formation in C. cinereus is bypassed by a spo22 mutation, which abolishes premeiotic DNA replication (Merino et al., 2000). The molecular basis of this bypass is not yet clear, but it could indicate that the presence of a sister chromatid and/or a replication-dependent chromosome structure imposes constraints on synaptonemal complex formation and that recombination initiation is required to overcome these constraints. The genetic dependence of SC formation on recombination functions is not universal. spo-11 mutants in C. elegans and mei-W68 and mei-P22 mutants in D. melanogaster females make normal amounts of SC that is indistinguishable from wild type by ultrastructural analysis (Dernburg et al., 1998; McKim et al., 1998). Likewise, mre-11 mutations in C. elegans have no discernible effect on pachytene chromosome morphology (A. Villeneuve, personal communication, 2000). Why are Spo11 and other recombination initiation proteins required for normal SC formation in some organisms but not in others? It is a formal possibility that there is a fundamental difference in the timing and relative dependency of these events (McKim et al., 1998; Walker and Hawley, 2000). However, it has not yet been tested whether recombination initiation requires synapsis in C. elegans or D. melanogaster, nor has it been possible to determine the relative timing of these events in normal meiosis. We have argued elsewhere for an alternative explanation that variations between organisms reflect species-specific differences in the balance between factors that contribute to proper synapsis (Baudat et al., 2000). Such factors could include facilitated versus nonfacilitated nucleation of SC formation, competition between self-aggregation and assembly of SC components along chromosome axes, and the choice of a synaptic partner. For example, D. melanogaster has robust pathways for ensuring homologous chromosome pairing (Walker and Hawley, 2000). These pathways are independent of homologous recombination and are manifested in various somatic tissues and in both the male germ line (where recombination is absent) and the female germ line (in which the fourth chromosome does not recombine, yet still develops SC). Perhaps these pairing mechanisms obviate a requirement for recombination initiation to nucleate SC formation. Similar arguments have been put forth by others (McKim and Hayashi-Hagihara, 1998). 2. Homologous Chromosome Pairing Pairing of homologous chromosomes is a prominent feature of nuclear architecture in many organisms, in both meiotic and mitotically dividing cells (e.g., Burgess et al., 1999). For the purposes of this review, homologous pairing is defined as the coalignment of homologous chromosomes, irrespective of whether the

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chromosome axes are held together by the SC. In diploid S. cerevisiae strains, homologous chromosomes are paired during G1 and G2 phases (Weiner and Kleckner, 1994; Burgess and Kleckner, 1999; Burgess et al., 1999). As cells enter meiotic S phase, premeiotic pairing is disrupted and then reestablished on a region-byregion basis. Restoration of pairing during meiosis requires SPO11 (Loidl et al., 1994; Weiner and Kleckner, 1994), but a spo11 point mutation that changes the catalytic tyrosine to phenylalanine (spo11-Y135F) supports normal levels of meiotic pairing even though it results in a complete defect in DSBs and recombination (Cha et al., 2000). Thus, SPO11 plays a role in chromosome pairing independent of its role in catalyzing DSB formation. The molecular nature of this role is not clear. One possibility is that Spo11 protein, or a Spo11-dependent multiprotein complex, is necessary for formation of a chromatin or higher order chromosome structure that is permissive for homologous pairing. MER2, MEI4, and REC102 are also required for normal homologous pairing (Nag et al., 1995; Rockmill et al., 1995). Moreover, restoration of homologous pairing during meiosis is drastically reduced in S. cerevisiae rad50 mutants, nearly as much as in spo11 mutants (Loidl et al., 1994; Weiner and Kleckner, 1994). However, it should be noted that premeiotic chromosome pairing also appears to be affected in rad50 null mutants (Weiner and Kleckner, 1994). Pairing is decreased in rad50S, mre11S, and sae2/com1 mutants relative to wild type, but not as severely as in a rad50 null (Loidl et al., 1994; Weiner and Kleckner, 1994; Nairz and Klein, 1997; Prinz et al., 1997). The pairing phenotypes of mre11 and xrs2 mutants in budding yeast have not been reported. As for defects in SC formation, effects of recombination initiation mutants on homologous pairing vary from organism to organism. For example, homologous pairing is defective in an mre11 mutant of C. cinereus (Gerecke and Zolan, 2000), but apparently not in C. elegans (A. Villeneuve, personal communication, 2000). Similarly, Spo11 is not required for meiotic homologous pairing in D. melanogaster or C. elegans (Dernburg et al., 1998; McKim et al., 1998), but in C. cinereus, a spo11 mutant shows a partial defect for meiotic homologous pairing (Celerin et al., 2000). There may be a similar defect during meiosis in Spo11-deficient mice, because much, if not all, of the residual synapsis in the mutants appears to be nonhomologous, as judged by the presence of frequent synaptic partner switches (Baudat et al., 2000; Romanienko and Camerini-Otero, 2000). 3. Duration of Premeiotic S Phase In many organisms, premeiotic S phase is significantly longer than the S phase of the preceding mitotic divisions (see Cha et al., 2000). In one study, bulk premeiotic DNA synthesis in S. cerevisiae took ∼80 min to complete, as opposed to ∼20 min in vegetative cells (Cha et al., 2000). spo11 mutants showed a 20 –30% decrease in the length of S phase, indicating that SPO11 is required for at least some element of this prolongation. The molecular role of SPO11 in this process is not clear, but

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the spo11-Y135F allele supports normal replication timing, indicating that this function is independent of DSB formation. REC102 is not required for normal kinetics of premeiotic S phase (Cha et al., 2000); none of the other DSB genes has been examined for this property. Studies suggest that rec12+ is not required for normal kinetics of S phase progression in S. pombe (Forsburg and Hodson, 2000). Whether S phase length is affected in spo11 mutants in other organisms has not been reported. 4. Premature Chromosome Segregation at Meiosis I In S. cerevisiae, spo11 mutants appear to carry out the first meiotic division earlier than wild type, on the basis of the kinetics of formation of binucleate cells (Klapholz et al., 1985; Giroux et al., 1993). This phenotype is closely tied to Spo11 catalytic activity because spo11 active site point mutants (e.g., spo11-Y135F ) are indistinguishable from deletion mutants in this respect (Cha et al., 2000; R. Diaz and S. Keeney, unpublished data, 2000). Several other DSB-defective mutants also show this property: rec102 deletion mutants generate binucleate cells earlier than normal (although this effect may be strain-dependent in some cases) (Bhargava et al., 1992; R. Cha, personal communication, 2000). Null rad50 mutants also form binucleates early, whereas rec104 and rec114 mutants divide earlier than wild type, but not as early as rec102 or rad50 (Galbraith et al., 1997; Jiao et al., 1999). In contrast, a mei4 mutation has no effect on division timing (Menees et al., 1992; Galbraith et al., 1997). The earlier division mutants rec102 and rad50 are epistatic to the rec104 mutant (i.e., a rec102 rec104 double mutant divides as early as a rec102 single mutant). Similarly, rad50, rec102, rec104, and rec114 are epistatic to the normally timed mei4 mutant. The molecular basis of these effects is not known. One interpretation is that the early divisions represent early onset of anaphase, meaning that the length of prophase I is shortened in the mutants. Malone and colleagues proposed that the “early division” genes are required to provide an inhibitory signal that delays meiotic progression, perhaps involving an altered chromatin or higher order chromosome structure (Jiao et al., 1999). Neither DSBs nor later recombination intermediates can be the source of such a signal, because mei4 mutants show normal division kinetics and because introduction of a DSB by HO endonuclease in an early-dividing mutant (rec104) does not affect division kinetics (Jiao et al., 1999). An alternative interpretation is suggested by the observation that spo11 mutants show a spindle checkpoint-dependent delay in degradation of the Pds1 protein, which normally happens at the onset of anaphase I (Shonn et al., 2000). Thus, spindle elongation and separation of chromatin masses occur in the mutant prior to progression of the cell cycle machinery to the anaphase stage. It was proposed that early division is caused by a failure of achiasmate chromosomes to resist the tension imposed by the prometaphase spindle (Shonn et al., 2000). However, this idea does not explain why the achiasmate mei4 mutant has normal division kinetics.

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One possibility is that mei4 cells have a compensatory delay in spindle assembly that causes the mutant to mimic normal division timing. Another possibility is that mei4 mutants are not completely DSB defective and that one or a few residual crossovers are sufficient to prevent premature spindle elongation. The epistasis relationship between early division mutants and mei4 favors the latter idea.

E. Genes That Are Involved in, but Not Absolutely Required for, Double-Strand Break Formation Several additional genes are important for formation of normal levels of DSBs. These include HOP1, RED1, MEK1, and KAR3. Many other genes (e.g., DMC1, RAD51, and RAD52) are required for normal repair of meiotic DSBs, but are not known to have effects on DSB formation per se. The latter genes have been extensively reviewed elsewhere (Roeder, 1997; Smith and Nicolas, 1998; Paques and Haber, 1999). 1. Chromosome Structure Elements: HOP1, RED1, and MEK1 Red1 (95.5 kDa), Hop1 (68.7 kDa), and Mek1 (56.9 kDa; also known as Mre4) are abundant, meiosis-specific proteins that localize to meiotic chromosomes (Thompson and Roeder, 1989; Hollingsworth et al., 1990; Leem and Ogawa, 1992; Smith and Roeder, 1997; Bailis and Roeder, 1998). Red1 and Hop1 are thought to be structural components of meiotic chromosomes and Mek1 is a kinase thought to regulate their activities. Mutations in each cause chromosome structure defects: red1 mutants do not make any AEs or SC (Rockmill and Roeder, 1990); hop1 mutants make AEs, but not SC (Hollingsworth and Byers, 1989; Loidl et al., 1994); and mek1 mutants make AE and discontinuous stretches of SC (Rockmill and Roeder, 1991). The three proteins interact physically and genetically with one another in a variety of assays (e.g., Hollingsworth and Johnson, 1993; Hollingsworth and Ponte, 1997; Bailis and Roeder, 1998; de los Santos and Hollingsworth, 1999). Mek1 can phosphorylate Red1 and itself in vitro (Bailis and Roeder, 1998; de los Santos and Hollingsworth, 1999; Woltering et al., 2000), and reversal of Mek1-dependent phosphorylation of Red1 has been proposed to control exit from the pachytene stage (Bailis and Roeder, 2000). Potential Hop1 orthologs were identified in C. elegans and A. thaliana (Zetka et al., 1999; Caryl et al., 2000). Mutations in the yeast genes cause defects in meiotic recombination and DSB formation. Recombination is reduced to ∼10–20% of normal in red1 and mek1 mutants (Rockmill and Roeder, 1990; Rockmill and Roeder, 1991), and to even lower levels in hop1 mutants (Hollingsworth and Byers, 1989; Rockmill and Roeder, 1990). These defects are mirrored by severe defects in DSB formation, although the extent of the defect may be strain and/or locus dependent (Mao-Draayer et al., 1996; Xu et al., 1997; Woltering et al., 2000). These results imply that activity of

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the DSB machinery is tightly coordinated with the development of meiotic chromosome structures. Red1 and Mek1 have been proposed to act at the point of DSB formation, perhaps engaging the DSB machinery along a pathway biased toward interhomolog recombination (Schwacha and Kleckner, 1997; Xu et al., 1997). Because of its more severe mutant phenotypes, Hop1 has been proposed to be even more intimately connected to the DSB machinery (Zickler and Kleckner, 1999). 2. KAR3 Somewhat surprisingly, the KAR3 gene is also required for normal levels of meiotic DSBs. kar3 mutants arrest before the first meiotic division and are severely defective for SC formation (Meluh, 1992; Bascom-Slack and Dawson, 1997). KAR3 encodes a kinesin-like motor protein essential for karyogamy and for normal mitotic spindle function (Meluh and Rose, 1990; Roof et al., 1992). Its role in meiotic recombination is not yet understood. It is possible that the recombination defects in kar3 mutants are indirect effects of problems in the mitotic divisions prior to meiosis. In keeping with this idea, IME1 transcription may be deregulated in kar3 mutants: expression in vegetative cells was derepressed and no further induction was seen on shift to sporulation conditions (Meluh, 1992). Expression of IME1 is a critical regulatory event for initiation of meiosis in budding yeast (Mitchell, 1994; Kupiec et al., 1997). Alternatively, Kar3 might play a more direct role by promoting chromosome movements that are important for pairing of homologous chromosomes and for recombination initiation (Bascom-Slack and Dawson, 1997). This proposal fits with the reorganization of the nucleus observed during meiosis in S. cerevisiae (e.g., Hayashi et al., 1998) and is in accord with observations in fission yeast (Chikashige et al., 1994; Kohli and Bahler, 1994; Svoboda et al., 1995). Early meiotic prophase in S. pombe is characterized by microtubule-driven, telomereled nuclear movements back and forth along the cell axis during and after karyogamy. These movements are thought to help reorganize the chromosomes, perhaps to facilitate homologous pairing and recombination. Indeed, several mutations that disrupt this nuclear movement cause a severe reduction in meiotic recombination: kms1 (Shimanuki et al., 1997; Niwa et al., 2000), taz1 (Cooper et al., 1998; Nimmo et al., 1998), and a mutation in the cytoplasmic dynein heavy chain (Yamamoto et al., 1999). It will be interesting to see whether these S. pombe mutations affect recombination at the initiation stage, as for budding yeast kar3.

F. Intergenic Interactions Important for Double-Strand Break Formation Many of the genes and gene products involved in DSB formation interact with one another. Several of these interactions are discussed above: Rad50, Mre11,

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and Xrs2/Nbs1 proteins form stable complexes, and several unidentified meiosisspecific proteins bind to the C-terminal portion of Mre11, which is critical for DSB formation (Section III,B). Spo11 and Ski8 interact in a yeast two-hybrid assay (Uetz et al., 2000). Hop1, Red1, and Mek1 interact physically and genetically with one another (Section III,E,1). Overexpression of REC104 partially suppresses the spore viability defect conferred by the temperature-sensitive hop1-628 allele (Hollingsworth and Johnson, 1993). This suppression is also observed with a hop1 null allele, suggesting that rescue is due to bypass of the need for HOP1 (Friedman et al., 1994). REC102 overexpression suppresses the recombination defect conferred by several temperaturesensitive rec104 alleles. A rec104 null mutation is not suppressed, however, suggesting that this interaction is not due to bypass of the mutant defect (Salem et al., 1999). A synthetic cold-sensitive phenotype was uncovered when certain alleles of SPO11 and REC102 were combined, and Rec102 and Spo11 proteins are present in a common multiprotein complex in meiotic cells (Kee and Keeney, 2001). Overexpression of REC102 or REC104 partially suppresses a temperature-sensitive spo11 mutant (Rieger, 1999). The molecular basis of these suppression and synthetic phenotypes is not known, but these observations point to an intriguing daisy chain of genetic interactions that connects the chromosome structure (i.e., Hop1 and Red1) to the business end of the DSB machinery (Spo11) via Rec102 and Rec104.

G. Other Potential Double-Strand Break Genes It is possible that additional factors required for DSB formation remain to be identified in budding yeast. Previous screens for recombination-defective mutants may not have achieved saturation, and redundant functions or genes required for vegetative growth would not have been uncovered. In other organisms, several DSB gene candidates have been identified from mutational analysis and homologybased searches. Except in S. pombe, direct demonstration of a requirement in DSB formation will be elusive because physical assays for DSBs are not yet available. However, by applying the criteria of a profound recombination defect and, if applicable, similarity to the phenotype of a spo11 null in the same organism, mutants can be classified as being likely to affect DSB formation. Whether a mutant can form cytologically observable deposits of γ -H2AX or complexes of strand exchange proteins (such as Dmc1 and Rad51) on meiotic chromosomes also appears to be a useful criterion (Gasior et al., 1998; Baudat et al., 2000; Romanienko and Camerini-Otero, 2000; Mahadevaiah et al., 2001). Several S. pombe rec genes were identified in a screen for meiotic recombination defects at the ade6 locus (Ponticelli and Smith, 1989). Mutations in rad32, isolated in a screen for radiation-sensitive mutants, also decrease meiotic recombination (Tavassoli et al., 1995). Of these genes, several are candidates to be involved in recombination initiation. rad32+ (an MRE11 ortholog), rec7 + (similar to REC114),

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+

rec12 (a SPO11 ortholog), and rec14 (similar to SKI8) are discussed above. Mutations in two others, rec6+ and rec15+, cause severe meiotic recombination defects and reduced spore viability, but show no mitotic phenotypes (Lin and Smith, 1994, 1995). rec6+, like rec12+, is required for meiosis-specific DSBs in S. pombe (Cervantes et al., 2000). Transcription of rec6+ and rec15+ is meiosis specific. Neither shows obvious sequence similarity to known proteins. Rec15 has a region of heptad repeat sequence predicted to form α-helical coiled coil, reminiscent of Mer2 from S. cerevisiae, but the two proteins share no sequence similarity otherwise. Several other genes are also important for meiotic recombination in S. pombe, but are considered more likely to be involved in later steps (Fox and Smith, 1998). Several D. melanogaster mutations drastically reduce meiotic recombination. These include mei-W68 (McKim et al., 1998) and mei-P22 (McKim et al., 1998; Sekelsky et al., 1999), which eliminate meiotic gene conversion, crossing over, and recombination nodule formation without giving an increase in sister chromatid exchanges (McKim et al., 1998). On the basis of these characteristics, the mutations were proposed to abolish initiation of recombination. This idea was confirmed by the subsequent demonstration that Mei-W68 is homologous to SPO11 (McKim and Hayashi-Hagihara, 1998). The mei-P22 mutation was isolated in a large-scale screen for P-element insertions that cause an increase in homolog nondisjunction (Sekelsky et al., 1999). The affected gene encodes a small, basic protein whose molecular function is not yet known (K. McKim, personal communication, 2000). Neither mei-W68 nor mei-P22 mutations have any discernible effect on SC formation, similar to spo-11 mutants in C. elegans (Section III,A,6). Another D. melanogaster mutation, c(3)G, also reduces meiotic recombination (Hall, 1972; Hawley et al., 1993), but c(3)G mutants are defective for SC formation (Smith and King, 1968). It is not certain whether c(3)G specifically affects initiation or a later step in recombination. Other recombination-defective mutants (e.g., mei-9 and mei-41) do not appear to meet the criteria discussed above for mei-W68 and mei-P22, so they are not likely to affect the initiation step (Hawley et al., 1993; Sekelsky et al., 1999). H. Possible Functions for the Friends of Spo11 The molecular roles for most of the proteins involved in DSB formation are not known, but several possibilities can be envisioned. Some might play an indirect role by controlling expression of other gene products. Mer1 and Nam8 are clear examples of this sort because they promote splicing of the MER2 transcript. Other DSB genes could encode transcription, splicing, or translation factors necessary for proper expression of Spo11 or other proteins. For most, this possibility has not been systematically addressed. How Spo11 activity is preferentially targeted to specific sites in the genome is poorly understood (Section IV,A). Some of the gene products required for DSB

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formation might play a role in determining this site specificity, either through their own DNA-binding properties or through interactions with other sequence-specific DNA-binding proteins. DSB formation is coordinated with the development of higher order chromosome structures (Section IV,C). Some of the proteins required for DSBs might participate in transducing input to or from chromosome structural elements, perhaps by interacting simultaneously with Spo11-containing complexes and with chromosome structure proteins. The genetic and/or physical interactions between HOP1, REC104, REC102, and SPO11 (Section III,F) point to Rec102 and Rec104 as possible candidates. Also, the Zip3 protein has been proposed to coordinate chromosome structure development with recombination because it is required for SC formation and interacts with Mre11 and other recombination proteins (Agarwal and Roeder, 2000). Finally, at least some of the proteins required for recombination initiation are likely to participate directly in the catalytic mechanism of DSB formation and resection. Factors of this type could stabilize the covalent Spo11–DNA complex, catalyze the release of Spo11 from its covalent attachment to the DNA, or function as an analog of the B subunit of archaeal topoisomerase VI (although none of the known players has obvious sequence similarity to Top6B proteins).

IV. Additional Factors That Influence or Are Influenced by Recombination Initiation A. Site Selectivity: Chromatin Structure, Promoters, and Sequence Specificity Recombination events are distributed nonrandomly along chromosomes (Lichten and Goldman, 1995; Baudat and Nicolas, 1997; Nicolas, 1998; Wahls, 1998; Gerton et al., 2000). In budding yeast, at least one component of this distribution arises from the preferential formation of DSBs at some sites (termed hot spots) but not at others (cold spots). The factors that determine whether a given sequence will be hot or cold are not completely understood, but some general rules have emerged. One important determinant is the chromatin structure. Essentially all known DSB sites are nuclease-hypersensitive sites in both mitotic and meiotic chromatin (Ohta et al., 1994; Wu and Lichten, 1994; Fan and Petes, 1996; Keeney and Kleckner, 1996). Tracts of simple repeats that exclude nucleosomes can activate meiotic recombination initiation (Kirkpatrick et al., 1999b). Moreover, opening of the chromatin structure at the PHO5 promoter when it is shifted from a repressed to an activated state correlates with an increase in DSB frequency (Wu and Lichten, 1994). These results suggest that an open chromatin configuration is necessary for DSB formation. But chromatin structure cannot be the sole arbiter of DSB site selectivity, because not all nuclease-hypersensitive sites are DSB sites,

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there is no correlation between the degree of nuclease hypersensitivity of a site and the frequency of DSB formation, and the fine pattern of nuclease hypersensitivity often does not precisely match the distribution of DSBs (Ohta et al., 1994; Wu and Lichten, 1994, 1995; Fan and Petes, 1996; Keeney and Kleckner, 1996; Borde et al., 1999). Moreover, under certain conditions a nuclease-hypersensitive region can show large variations (10- to 15-fold) in DSB frequency with no detectable changes in the accessibility to DNase I (Wu and Lichten, 1995; Borde et al., 1999). Most naturally occurring DSB sites appear to lie in promoter regions (Baudat and Nicolas, 1997; Nicolas, 1998). Nevertheless, several lines of evidence indicate that transcription per se is not required for DSB formation. First, deletion of the TATA box at HIS4 drastically diminishes transcription without affecting the recombination frequency (White et al., 1992). Second, some DSB sites are in intergenic regions without nearby promoters, and some are within coding sequences (Bullard et al., 1996; Baudat and Nicolas, 1997). Third, artificial hot spots have been created by insertion of heterologous sequences into the genome (Cao et al., 1990; Wu and Lichten, 1995) or by incorporation of DNA structures such as large palindromes or trinucleotide repeat tracts (Nag and Kurst, 1997; Jankowski et al., 2000; Nasar et al., 2000). Most of these potent DSB sites appear not to be transcriptionally active. The tendency of DSB sites to occur in promoter regions may reflect the fact that promoter regions tend to have an open chromatin configuration. Alternatively, sequence-specific binding proteins (such as transcription factors) may play a role in targeting the DSB machinery to particular sites, either by inducing a local chromatin structure that is permissive for DSB formation or by recruiting the recombination initiation machinery by direct protein–protein interactions (White et al., 1992, 1993; Fan et al., 1995; Kirkpatrick et al., 1999a). Petes and colleagues have argued that there are two classes of recombination hot spot: those that require transcription factors for targeting recombination complexes, and those that are transcription factor independent because they are constitutively in a permissive state for DSB formation (Kirkpatrick et al., 1999a). A transcription factor complex is required for activity of the M26 hot spot in S. pombe as well (Wahls and Smith, 1994; Kon et al., 1997). Until recently, no clear sequence consensus had been established for DSB sites in S. cerevisiae. To address this issue, Simchen and colleagues compared sequences flanking six DSB hot spots and derived a conserved sequence motif that they termed CoHR (for Common Homology Region) (Blumenthal-Perry et al., 2000). The 50-bp profile contains a central poly(A) tract and can tolerate fairly large (up to 250-bp) gaps. Good matches to this sequence profile are associated with the majority of strong DSB sites mapped on Chromosomes I, III, and VI. Moreover, two CoHR profile matches are found near the ARG4 DSB hot spot, and genomic deletions in this region that remove these sequences also abrogate DSB formation. One possibility is that the CoHR sequence provides a preferred site for assembly of

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Spo11 and/or associated proteins. Alternatively, this sequence might be associated with a chromatin structure that is especially permissive for changes that are required for DSB formation (Blumenthal-Perry et al., 2000). However, it is important to note that this profile is neither necessary nor sufficient for DSB formation, because not all mapped DSB sites have an associated CoHR motif and not all CoHR motifs are associated with DSB sites (especially near telomeric regions). It will be interesting to see whether more subtle targeted mutations within the profile affect DSB frequencies at CoHR-associated hotspots. Nucleotide-resolution mapping reveals that Spo11-mediated cleavage at several prominent DSB loci occurs at any of a number of positions distributed across fairly large stretches of the DNA (∼70–250 bp) (de Massy et al., 1995; Liu et al., 1995; Xu and Kleckner, 1995; Xu and Petes, 1996). Within these regions, break positions are distributed nonrandomly, but no rules for the nucleotide-resolution determinants of strand cleavage have been discerned. This may indicate that positioning of Spo11 within an acceptable DSB site is relatively sequence nonspecific.

B. Meiosis-Specific Alteration of Nuclease Hypersensitivity in the Chromatin at Recombination Hot Spots There is a meiosis-specific increase in the micrococcal nuclease (MNase) hypersensitivity of the chromatin at DSB sites at or before DSB formation (Ohta et al., 1994). Nuclease-hypersensitive regions that are not prominent DSB sites show little or no change as cells enter meiosis. Moreover, certain circumstances that eliminate hot spot activity without affecting the premeiotic chromatin structure also eliminate the meiosis-specific increase in MNase hypersensitivity (Ohta et al., 1999). Interestingly, a change in nuclease sensitivity is not seen when DNase I is used instead of MNase (Wu and Lichten, 1994). The reason for the nuclease-specificity of the phenomenon is not known. A meiosis-specific increase in MNase hypersensitivity has also been observed at the ade6-M26 hot spot in S. pombe, perhaps reflecting the operation of similar processes (Mizuno et al., 1997). MRE11, NAM8/MRE2, REC102, and REC104 are required for this meiotic increase in MNase hypersensitivity (Furuse et al., 1998; Ohta et al., 1998; K. Ohta, personal communication, 2000). In contrast, rad50 and xrs2 null mutants increase meiotic MNase hypersensitivity to higher levels than normal. mre11 mutants are epistatic to rad50 in this respect (Ohta et al., 1998). The requirement for NAM8/MRE2 suggests that Mer2 protein is essential for this process, but this has not been directly tested. For Mre11, the C-terminal portion that is required for DSB formation (Section III,B,2,b) is also required for induction of MNase hypersensitivity, suggesting that these phenomena are intimately connected. The molecular basis of this process is not understood, but the genetic requirements make it likely that the changes in MNase hypersensitivity are caused by binding of components of the DSB machinery.

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C. Interplay with the Development of Higher Order Chromosome Structure As discussed above, mutations that block DSB formation or resection also block the development of meiotic chromosome structures, and normal DSB formation requires the chromosome structure proteins Red1 and Hop1. Thus, there is substantial interplay between the recombination initiation machinery and the development of higher order structure of meiotic chromosomes. Additional observations may also reflect this interplay. First, DSB sites are nonrandomly dispersed among large chromosomal domains (Zenvirth et al., 1992; Baudat and Nicolas, 1997; Gerton et al., 2000). On S. cerevisiae Chromosome III, for example, large regions near the telomeres show few or no prominent DSBs. There are also few DSBs near the centromere, whereas interstitial regions in each of the chromosome arms show a high frequency of DSBs. In principle, the positions of these domains could be determined by the nonrandom distribution of potential DSB sites or by features of higher order chromosome structure that inhibit or potentiate DSB formation within specific regions. In studies to distinguish between these possibilities, Lichten and colleagues found that a recombination reporter construct placed at various sites along Chromosome III took on the properties of its position. Insertions into cold regions gave low meiotic DSB levels and low recombination frequencies, whereas insertions into hot regions gave higher DSB levels and higher recombination frequencies (Wu and Lichten, 1995; Borde et al., 1999). There was no detectable difference in DNase I hypersensitivity within the constructs, suggesting that the effects were not due to changes in the chromatin structure. This striking chromosomal position effect supports a model in which higher order structures and/or chromosome dynamics control the location of recombination initiation events. Because the same reporter construct was used in all cases, this domainal control must be largely independent of the local sequence at the DSB site. The molecular basis of these position effects is not currently known. Second, DSB sites compete with other sites nearby. Each recombination reporter construct used in the studies described above consists of an arg4 mutant allele, a selectable marker for integration (URA3), and vector sequences derived from pBR322 (Wu and Lichten, 1995). DSBs occur at high frequency within the pBR322-derived sequences, but a site in the arg4 promoter that gives rise to DSBs when in its normal context on the yeast chromosome (Sun et al., 1989; de Massy and Nicolas, 1993) does not give rise to DSBs in this construct (Wu and Lichten, 1995). Removing the pBR322 sequences from the construct restores DSBs to the arg4 promoter region. Thus, the strong DSB sites in the pBR322 sequences inhibit DSB formation at a nearby site. DNase I hypersensitivity at the arg4 promoter in the constructs is unaffected by the presence or absence of adjacent pBR322 sequences. Thus, the effects on DSB formation are not indirect consequences of changes in chromatin structure. Similar competition phenomena have been seen

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at other DSB hot spots (Xu and Kleckner, 1995; Fan et al., 1997). Moreover, it appears that competition can operate over relatively large distances because insertion of a strong DSB hot spot at the HIS4 locus inhibits recombination as much as 40 kb away on the same chromosome (Wu and Lichten, 1995; M. Lichten, personal communication, 2000).

D. The DNA Replication Connection Initiation of meiotic recombination occurs after premeiotic DNA replication (Padmore et al., 1991; Borde et al., 2000). It now appears that this temporal coordination is achieved through a mechanistic connection(s) between replication and DSB formation (reviewed in Baudat and Keeney, 2001). A number of studies have shown that mutations such as cdc8, cdc21, or pol1 that block premeiotic DNA replication also prevent meiotic recombination (Schild and Byers, 1978; Budd et al., 1989). Similarly, cells that are defective for CLB5 and CLB6 are incapable of carrying out either replication or recombination in meiosis (Stuart and Wittenberg, 1998; Smith et al., 2001). These genes encode B-type cyclins that promote the G1-to-S transition in vegetative cells (Stuart and Wittenberg, 1998). Defects in premeiotic DNA replication and recombination are also caused by mum2 mutations, but the molecular role of the Mum2 protein is not yet known (Davis et al., 2001). At least for mum2 and clb5 clb6 mutants, the recombination block occurs at or prior to the initiation step and is not due to a failure to induce the transcription of meiotic recombination genes (Davis et al., 2001; Smith et al., 2001). In contrast, treating meiotic cells with the ribonucleotide reductase inhibitor hydroxyurea similarly blocks both DNA synthesis and DSB formation (Simchen et al., 1976; Borde et al., 2000), but hydroxyurea treatment also prevents transcriptional induction of a number of meiotic genes, including SPO11 and HOP1, confounding the interpretation of this finding (Davis et al., 2001; S. Keeney, unpublished data, 2000; V. Borde and M. Lichten, personal communication, 2000). In principle, the block to recombination in the above-described mutants could be a consequence of inducing a regulatory arrest via a replication checkpoint. However, this appears not to be the case because the recombination defect in mum2 mutants (or hydroxyurea-treated cells) is not suppressed by a mec1 mutation, which eliminates the replication checkpoint (Borde et al., 2000; Davis et al., 2001). Moreover, the replication defect in clb5 clb6 double mutants does not trigger the MEC1-dependent checkpoint at all, so the recombination defect in this mutant cannot be a consequence of checkpoint induction (Stuart and Wittenberg, 1998; Smith et al., 2001). In a more direct analysis of the temporal relationship between replication and recombination, Borde and colleagues (2000) showed that DSB formation follows DNA replication by roughly 1.5–2 h. Delaying replication timing on one arm of Chromosome III (by deleting replication origins on that arm or by juxtaposing

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recombination initiation sites to a telomere) delayed DSB formation by a corresponding amount, without affecting DSB timing on the unaltered chromosome arm. These results strongly indicate that the timing of DSB formation is controlled on a chromosomal region-by-region basis rather than globally. In contrast, these studies revealed that the timing of disappearance of DSBs (as they are processed into further recombination intermediates) is controlled on a nucleus-wide basis. This temporal and regional correlation between replication and recombination initiation, combined with a lack of a clear regulatory (checkpoint) connection, supports the idea that the two processes are mechanistically coupled (Borde et al., 2000; Cha et al., 2000). One way in which they could be coupled would be if passage of the replication fork is required to establish a higher order chromosome structure that is permissive for DSB formation. Or, factors involved in DSB formation (including Spo11) might assemble on the chromosomes only during DNA replication. The latter idea fits with the observed effects of a spo11 mutation on the length of premeiotic S phase (Cha et al., 2000; see Section III,D,3) and by the fact that clb5 clb6 mutants do not show a meiotic increase in MNase hypersensitivity at DSB hot spots (Smith et al., 2001), perhaps reflecting a failure to assemble a pre-DSB complex (see Section IV,B).

E. Homologous Chromosome Pairing Homologous chromosome pairing is a universal feature of meiosis. In S. cerevisiae, there is interplay between the DSB machinery and the pathways that establish meiotic levels of chromosome pairing. As discussed in Section III,D,2, SPO11 and several other DSB genes are required for normal levels of DSB-independent homologous pairing. In addition, DSB frequencies on one chromosome can be influenced in trans by sequences at an allelic position on its homologous partner (Xu and Kleckner, 1995; Bullard et al., 1996; Keeney and Kleckner, 1996; Rocco and Nicolas, 1996). These observations may indicate that homology-dependent physical interactions between chromosomes influence chromatin structure and DSB formation. However, such interactions appear not to be obligatory for DSB formation. DSBs are formed with essentially normal kinetics and frequency in the absence of a homolog (i.e., in haploid cells undergoing meiosis) (de Massy et al., 1994; Gilbertson and Stahl, 1994; Fan et al., 1995) and at sites that lack local homology with the allelic positions on their homologous partners (e.g., Wu and Lichten, 1995).

F. Cell Cycle Control DSB formation is clearly under cell cycle control. First, formation of DSBs is normally limited to a fairly narrow window of time in meiosis (e.g., Padmore et al., 1991), even though Spo11 protein persists in the cell after DSBs are no longer

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made (S. Keeney, unpublished observations, 2000). Second, when meiotic cells are transferred to vegetative growth medium during prophase I, DSBs disappear rapidly, although they would have continued to form if the cells had been left in meiosis (Schwacha and Kleckner, 1997; Zenvirth et al., 1997; Arbel et al., 1999). This result indicates that existing DSBs are rapidly repaired on shift from meiosis to vegetative growth, but also implies that de novo formation of DSBs ceases. This in turn suggests that the DSB machinery is responsive to the physiological state of the cell. Third, regulatory arrest at the pachytene stage caused by ndt80, cdc28, cdc36, or cdc39 mutations is accompanied by an increased frequency of total meiotic recombination (both noncrossover and crossover associated) (Shuster and Byers, 1989; Xu et al., 1995). One interpretation of this observation is that a cell-cycleregulated transition is required to end the period during which recombination can initiate. There are many ways in which DSB formation could be tied to cell cycle progression. Cell-cycle-regulated posttranslational modifications could control the activity, stability, or subcellular localization of one or more critical factors (such as Spo11). Or, DSB formation might be controlled by cell-cycle-driven transitions in chromosome structure (see below).

V. A Molecular Model for Spo11 Action A working model for the mechanism of DSB formation is shown in Fig. 4 (see color insert). This model combines previously proposed features (de Massy et al., 1995; Keeney and Kleckner, 1995; Liu et al., 1995; Keeney et al., 1997) with details suggested by the structure of Top6A (Nichols et al., 1999). The first step is the assembly of pre-DSB “potential” (Xu et al., 1997): a Spo11 dimer binds a target site, presumably as part of a multiprotein complex (Fig. 4A and B; only Spo11 is diagrammed). A specific side chain on each Spo11 protomer (Tyr-135 in the S. cerevisiae enzyme) then attacks the DNA backbone, generating a reversible covalent complex with tyrosyl phosphodiester linkages between the proteins and the 5 -terminal strands. By analogy with type II topoisomerases, the author proposes that the noncovalent and covalent forms of the Spo11–DNA complex are in equilibrium at this point, with the balance favoring the noncovalent complex. In the second step, the pre-DSB potential is activated such that the Spo11 complex becomes committed to generating a DSB (Fig. 4B and C). This step is proposed to result in the formation of the irreversible covalent intermediate detected in rad50S and mre11 nuclease-defective mutants. One way this could be achieved would be for the Spo11 dimer interface to be disrupted and the protomers to be physically separated, similar to an earlier proposal by de Massy and colleagues (1995) (Fig. 4C). This movement would disrupt the active sites of the enzyme by separating each catalytic tyrosine from the Toprim metal-binding pocket on the other protomer. This intermediate could be directly equivalent to the strand passage intermediate proposed for the topoisomerase VI catalytic cycle

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(Nichols et al., 1999). Or, it could be analogous to the irreversible DNA cleavage complexes formed by gyrase or topoisomerase IV when a helicase or replication fork encounters a ternary topoisomerase–drug–DNA complex (Hiasa et al., 1996; Shea and Hiasa, 1999, 2000). In the third step, the covalent complex is converted to a protein-free DSB. Two general mechanisms for the release of Spo11 have been proposed (Keeney et al., 1997). In one, the 5 -tyrosyl phosphodiester bond is hydrolyzed, releasing intact Spo11 protomers (Fig. 4D). This reaction could be catalyzed by Spo11 itself; an E. coli topoisomerase IV mutant thought to be capable of such a reaction has been described (Nurse et al., 2000). Alternatively, another protein might catalyze the hydrolysis. An enzyme specific for 3 -tyrosyl linkages has been described (Pouliot et al., 1999), so a 5 -phosphodiesterase might also exist. In the second proposed release mechanism, single-strand endonucleolytic cleavage at some distance from the covalent protein–DNA linkage releases oligonucleotide-bound Spo11 and a partially or fully resected 5 strand (Fig. 4E). The DSB ends may or may not be processed further by one or more 5 → 3 exonucleases, perhaps including Exo1 (see Section III,B,1). Within the framework of this model, the proteins involved in DSB formation (Section III) are most likely to be involved in establishing DSB potential or in activating it subsequently. Sae2, the “S” function of Rad50, and the nuclease activity of Mre11 all appear to be specifically required for the third step. Thus, the Rad50–Mre11–Xrs2 complex (perhaps in conjunction with Sae2) is an attractive candidate to carry out the endonucleolytic release reaction (Fig. 4E). The helicase activity of this complex (Paull and Gellert, 1999) might partially unwind the DNA duplex at some distance from the covalently bound Spo11, preparing a singlestranded bubble as a substrate for Mre11 endonuclease activity. What is the signal that activates a potential DSB complex and triggers formation of an irreversible Spo11–DNA complex? One possibility arises from a model of Kleckner and colleagues, in which cyclical changes in the tensional state of chromosomes result in successive stages of stress and stress relief that drive DNA metabolic events and chromosome morphogenesis (Kleckner, 1997; Zickler and Kleckner, 1999; N. Kleckner, personal communication, 2000). Stress along the chromosome (perhaps induced by cell-cycle-driven changes in chromatin expansion) could provide a physical force favoring Spo11 dimer disruption. If a critical level of stress is required to trigger irreversible cleavage by a Spo11 complex, and if cleavage is accompanied by relief of stress that is propagated for some distance along the chromosome, then formation of a DSB would prevent nearby Spo11 complexes from giving rise to breaks. Thus, such a scenario could account for observed patterns of interference between DSB sites on the same chromosome (Section IV,C). Of course, other possibilities can also be envisioned. Undoubtedly, further analysis of Spo11 and its partners in a variety of organisms will continue to unravel the molecular details of recombination initiation and the means by which it is controlled.

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Acknowledgments I thank the many colleagues who generously provided reprints, preprints, and unpublished results. I am especially indebted to Fr´ed´eric Baudat, Sean Burgess, Neil Hunter, Michael Lichten, and members of my laboratory for critical reading and insightful comments on the manuscript. Preparation of this review and work from my laboratory were supported in part by grants from the NIH and the New York City Council Speaker’s Fund for Biomedical Research.

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2 Osmoregulation and Cell Volume Regulation in the Preimplantation Embryo Jay M. Baltz Hormones, Growth, and Development Unit, Ottawa Health Research Institute, and Departments of Obstetrics and Gynecology (Division of Reproductive Medicine) and Cellular and Molecular Medicine University of Ottawa Ottawa, Ontario, Canada K1Y 4E9

I. Introduction II. Sensitivity of Mammalian Embryos to Osmolarity A. Osmolarity in Vitro B. Are One-Cell Embryos More Sensitive Than Two-Cell Embryos? III. Response to Hypertonicity and Regulation against Cell Volume Decreases by Mammalian Embryos A. Dependence of Cell Volume on Osmolarity B. Short-Term Volume Regulation by Inorganic Ion Transport C. Organic Osmolytes D. Organic Osmolytes and Embryos IV. Regulation against Volume Increases by Mammalian Embryos A. Regulation against Volume Increases B. Regulation against a Volume Increase in Embryos V. Organic Osmolytes and Osmolarity in Vivo A. Organic Osmolytes within in Vivo-Derived Preimplantation Embryos B. Organic Osmolytes in Oviductal Fluid C. Oviductal Fluid Osmolarity VI. Discussion and Summary References

The early preimplantation mammalian embryo possesses mechanisms that regulate intracellular osmolarity and cell volume. While transport of osmotically active inorganic ions might play a role in this process in embryos, the major mechanisms that have been identified and studied are those that employ organic osmolytes. Organic osmolytes provide a substantial portion of intracellular osmotic support in embryos and are required for their development under in vivo conditions. The main osmolytes that have been identified in cleavage stage embryos are accumulated via two transport systems of the neurotransmitter transporter family active in early preimplantation embryos—the glycine transport system (GLY) and the β-amino acid transport system (system β). While system β has been established to have a similar role in Current Topics in Developmental Biology, Vol. 52 C 2001 by Academic Press. All rights of reproduction in any form reserved. Copyright  0070-2153/01 $35.00

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Jay M. Baltz many other cells, this is a novel function for the GLY transport system. The intracellular concentration of organic osmolytes such as glycine in early preimplantation embryos is regulated by tonicity, allowing the embryo to regulate its volume against shrinkage and to control its internal osmolarity. In addition, the cells of the embryo can regulate against an increase in volume via controlled release of osmolytes from the cytoplasm. This is mediated by a swelling-activated anion channel that is also highly permeable to a range of organic osmolytes, and which closely resembles similar channels found in many other cell types (VSOAC channels). Together, these mechanisms appear to regulate cell volume in the egg through the early cleavage stages of embryogenesis, after which there are indications that the mechanisms of osmoregulation change.  2001 Academic Press. C

I. Introduction Animal cells regulate their size precisely. This is true not only for somatic cells, but also for the cells of eggs and early embryos. Oocytes grow to a precise diameter characteristic of each species and the egg maintains that diameter for an extended period, implying that mechanisms must exist by which an egg can determine and control its size. Subsequently, as the embryo cleaves into successively smaller cells, each embryonic stage possesses blastomeres that are maintained at characteristic dimensions. The mechanisms determining blastomere size appear to be exceedingly ancient, because the same precise embryo cleavage patterns and equal-sized blastomeres can be clearly seen in the oldest known multicellular fossils (Xiao et al., 1998). Most work on elucidating volume regulation in mammalian embryos is recent, and was spurred by the realization that early preimplantation (PI) embryos, especially cleavage-stage embryos, will not develop in vitro when osmolarity is increased to approximately the levels thought to exist in vivo in the oviduct, but will develop at lower osmolarity. Because animal cells control their volumes osmotically, this observation implicated cell volume regulation as an important determinant of embryo viability. Further work has led to the realization that early PI embryos preferentially use organic osmolytes rather than inorganic ions for intracellular osmotic support, and that embryo viability at elevated osmolarities is rescued by the presence of such osmolytes. Organic osmolytes such as glycine and β-amino acids are almost certainly accumulated by early PI embryos in vivo. Evidence suggests that at least some of the transport systems that mediate intracellular accumulation of organic osmolytes in embryos are not the same as those that perform this function in other cells. Thus, the study of cell volume regulation in embryos not only sheds light on early embryo physiology, but may also reveal novel volume-regulatory mechanisms that might operate in other cells as well. This review details what is currently known about mechanisms of cell volume

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control in mammalian PI embryos. Although there has been an increase in information on this area, there is still much to be learned, especially in the area of how volume-regulatory mechanisms change during development and how they are regulated.

II. Sensitivity of Mammalian Embryos to Osmolarity A. Osmolarity in Vitro There have been a number of attempts to determine the optimal osmolarity1 for PI embryo development in vitro. Brinster (1965) demonstrated that there was a permissive range of osmolarities for two-cell embryo development to blastocysts, with an optimum at approximately 275 mOsM (Fig. 1). Subsequent studies in which mouse embryos have been cultured from the one-cell or two-cell stages have generally reported similar osmosensitivity (Fig. 1), and this has also been reported for rabbit (Naglee et al., 1969; Li and Foote, 1996), bovine (Seidel, 1977; Liu and Foote, 1996), rat (Miyoshi et al., 1994,1995), and porcine (Beckmann and Day, 1993) embryos. Osmolarity has generally been altered either by varying NaCl concentration or by adding a presumably inert component such as mannitol, raffinose, or sucrose.2 Thus, an effect on embryo development which had been ascribed to altered osmolarity might potentially be due to altered composition of the medium. Thus, it was necessary to demonstrate that the effects on embryo development were similar regardless of the means used to vary osmolarity. To this end, it has been shown that one-cell mouse embryo development was identically affected by hypertonicity regardless of whether osmolarity is varied with NaCl or the inert trisaccharide raffinose, indicating that embryo development is sensitive to increased osmolarity 1 Most measured values are actually osmolality (osmoles per kilogram) rather than osmolarity (osmoles per liter, or OsM). However, in the relatively dilute solutions that are physiologically relevant, these two measures differ negligibly. Thus, the more convenient osmolarity (usually expressed as mOSM) is used here throughout. Both osmolarity and osmolality are intrinsic properties of a solution, and should be distinguished from tonicity, which describes the osmotic effect of a solution on a particular entity such as a cell and is thus context specific. 2 Another method used to alter embryo culture medium osmolarity is proportional dilution or concentration (e.g., Hay-Schmidt, 1993; Li and Foote, 1995, 1996; Liu and Foote, 1996), which has the advantage of maintaining the proportions between the various components. However, in most cases the absolute concentration of each component of the medium is important, and the interactions between components are complex rather than proportional (Lawitts and Biggers, 1991a,b,1992,1993; Biggers, 1998). Thus, when diluting or concentrating media proportionally, the most important effects may be due to changes in the concentration of an essential nutrient or an inhibitory component, or a perturbation of the interactions between a number of components, rather than to osmolarity. Because of these difficulties, it may be inadvisable to alter media by this method if the aim of the experiment is primarily to assess the effects of osmolarity.

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Figure 1 Development of mouse embryos in vitro as a function of medium osmolarity: Frequency of development to blastocysts from the one-cell stage (1c, top) or two-cell stage (2c, bottom) as reported in the literature. The results of all studies found by the author in which in vitro mouse embryo development was compared in media covering a significant range of osmolarities are shown here. Only studies or portions of studies in which no organic osmolytes were included in the media are presented here, as the utilization of organic osmolytes by embryos is discussed separately. In each case, embryos were cultured continuously to the blastocyst stage in media with osmolarity adjusted to the value indicated. In those cases in which the data were given only in graphic form, values (osmolarity and/or percent development) have been estimated by the author. The sources of the data are indicated by letters labeling each curve, as follows: (A) Whitten (1971) (note: data were presented in a transformed form, but the transform used was not indicated in the original reference; thus these data are presented as “mean angular response” as reported, rather than as percent development); (B) Davidson et al. (1988a); (C) Dawson and Baltz (1997); (D) Lawitts and Biggers (1992); and Biggers et al. (1993); (E) HaySchmidt (1993); (F) Van Winkle et al. (1990a) (note: the original data are in the form of total ion concentrations; osmolarities of media formulated as indicated were measured by the author and used here); (G) Brinster (1965). Details can be found in the original articles.

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Figure 2 Development of mouse one-cell embryos in vitro as a function of osmolarity varied with NaCl or raffinose. Mouse one-cell embryos were cultured in media whose osmolarity was adjusted either by varying the NaCl concentration (circles) or by adding raffinose (triangles). Similar sets of experiments were done either in the absence (solid symbols) or presence (open symbols) of 1 mM glutamine. Embryo development, measured as percent blastocysts, was essentially identical irrespective of whether osmolarity had been adjusted with NaCl or raffinose (compare circles with triangles). The presence of glutamine, however, protected embryo development at increased osmolarities (compare open with closed symbols). The protection was identical regardless of whether osmolarity was increased with NaCl or raffinose. [Adapted from Dawson and Baltz (1997) with permission of the Society for the Study of Reproduction.]

per se3 (Fig. 2; Dawson and Baltz, 1997). Nevertheless, it can be seen that the range of osmolarities found to support PI mouse embryo development differed between the various studies (Fig. 1), indicating that variables such as composition of medium, method of altering osmolarity, or mouse strain affect the response of embryos to osmolarity so that there is not a single “permissive range” of osmolarities for PI embryo development under all conditions. This can be clearly seen in the difference in osmolarity ranges supporting embryo development in the presence 3 NaCl may also exert a concentration-dependent effect on PI embryo development independent of osmolarity. Too low a concentration of either Na+ or Cl− can have a detrimental effect on blastocyst development (Manejwala et al., 1989; Zhao et al., 1997). Increased NaCl may be detrimental as well, because Li and Foote (1995,1996) found that varying NaCl even at constant osmolarity affected development of rabbit zygotes or two-cell embryos, although the interpretation of these experiments is not quite straightforward as osmolarity was kept constant by proportionally adjusting all other constituents of the medium.Van Winkle et al. (1990a) reported that increasing osmolarity by adding NaCl had a greater detrimental effect on two-cell embryo development than increasing osmolarity by adding mannitol, and Liu and Foote (1996) reported a similar finding for bovine embryo development and sorbitol, although this did not seem to be the case with rabbit embryos (Li and Foote, 1996). These findings may indicate a toxic effect of NaCl, but alternatively, it could be that compounds such as sorbitol and mannitol, which can serve as organic osmolytes, protect against increased osmolarity by entering the cells, as suggested for mannitol in two-cell mouse embryos (Van Winkle et al., 1990a). In support of this, raffinose, which does not enter mammalian cells or serve as an organic osmolyte, was found to be equivalent in its effect to NaCl (Dawson and Baltz, 1997).

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or absence of glutamine under otherwise identical conditions (Fig. 2). As is seen below, interactions between osmolarity and components of the medium determine embryo viability and reveal physiologically important processes in early embryos. B. Are One-Cell Embryos More Sensitive Than Two-Cell Embryos? It is generally accepted that one-cell embryos are more susceptible to adverse culture conditions than are later stage PI embryos, but a comparison of the data in the literature does not clearly indicate a much narrower permissive range of osmolarities for one-cell culture versus two-cell culture (Fig. 1). However, it is difficult to draw any firm conclusions, as each study was done with different media, different strains of mice, and different means of altering osmolarity. Only one direct comparison of the effects of osmolarity on one-cell versus two-cell embryo development has apparently been carried out. Davidson et al. (1988b) cultured mouse embryos from the one-cell stage to blastocysts and exposed them to altered osmolarity only during 24-h periods coinciding either with the one-cell or two-cell stage. They thus found that two-cell embryos would tolerate a larger perturbation of osmolarity than one-cell embryos, indicating a greater sensitivity of one-cell embryos to osmolarity. Mammalian embryos cultured from the one-cell stage exhibit a tendency to block at specific points in development. In the mouse, this is manifested as a block at the two-cell stage in inbred and outbred strains (see Biggers, 1998). Improved culture media have led to the alleviation of the mouse two-cell block (Chatot et al., 1989; Lawitts and Biggers, 1991a,b, 1993; Erbach et al., 1994), as well as similar blocks in other species (Schini and Bavister, 1988; McKiernan et al., 1995; Miyoshi et al., 1994, 1995). A common feature of many of these media is that their osmolarity is markedly lower than those of media in which the two-cell block occurs: CZB medium has an osmolarity of about 275 mOsM (Devreker and Hardy, 1997) and KSOM is about 250 mOsM (Devreker and Hardy, 1997; J. M. Baltz, unpublished data, 2000). The need to reduce osmolarity in order to culture one-cell embryos to blastocysts, whereas two-cell embryos will culture to blastocysts at a high rate in higher osmolarity media (e.g., M16, whose osmolarity is about 290 mOsM), provides additional indication that one-cell embryos are more sensitive to increased osmolarity than two-cell or later stage embryos.

III. Response to Hypertonicity and Regulation against Cell Volume Decreases by Mammalian Embryos A. Dependence of Cell Volume on Osmolarity Even a small difference between the intracellular and extracellular concentrations of osmolytes results in an osmotic pressure differential that is too large

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for animal cells to withstand. Most cells therefore immediately respond to osmotic perturbations by behaving as nearly ideal osmometers (Lang et al., 1998a), with their volumes changing by exactly the amount needed to equalize osmotic pressure across the membrane. Thus, the short-term mechanism for alleviating osmotic pressure differences is a change of cell volume. Mouse eggs and embryos have accordingly been found to behave as ideal osmometers in response to rapid changes in osmolarity, reaching a new steady-state volume within minutes (Oda et al., 1992; Gao et al., 1996; Collins and Baltz, 1999). In the longer term, however, cells require the ability to actively regulate their volumes and thus have evolved mechanisms that allow the regulated recovery from swelling or shrinking (Hallows and Knauf, 1994; Lang et al., 1998a; Chamberlin and Strange, 1989). It is a mistaken, but all too common, notion that active regulation of cell volume is required only in extreme circumstances, such as when large osmotic perturbations of the external environment occur. In practice, cells, including oocytes and PI embryos, are required to regulate their volumes constantly in the face of osmotic changes continually arising from metabolic generation of solutes, solute transport, and normal variations in extracellular fluid composition (Lang et al., 1998a).

B. Short-Term Volume Regulation by Inorganic Ion Transport Many cells are capable of regaining normal volume after a volume loss, an ability termed a “regulatory volume increase” (RVI). Short-term RVI is mediated by transport of inorganic ions into the cell, which increases osmotic pressure and thus increases cell volume. One major mechanisms responsible for RVI in animals is activation of Na+,K+,2Cl− cotransporters of the NKCC family (Sarkadi and Parker, 1991; Levinson, 1992; Haas and Forbush, 1998; O’Neill, 1999). Whether PI mammalian embryos express the Na+, K+,2Cl− cotransporter is unknown. The other major mechanism of RVI in mammalian cells is the coupled activation of Na+/H+ antiporters and HCO3−/Cl− exchangers mediating the net influx of Na+ and Cl− (Parker, 1988; Hallows and Knauf, 1994; Kapus et al., 1994; Humphreys et al., 1995; Orlowski and Grinstein, 1997). A functional HCO3−/Cl− exchanger has been shown to be present in PI embryos of the mouse (Baltz et al., 1991; Zhao et al., 1995; Zhao and Baltz, 1996), hamster (Lane et al., 1999), human (Phillips et al., 2000), and possibly cow (Lane and Bavister, 1999). An Na+/H+ antiporter has been demonstrated in PI embryos of hamster (Lane et al., 1998), mouse (Gibb et al., 1997; Barr et al., 1998; C. L. Steeves et al., 2001), cow (Lane and Bavister, 1999), and human (Phillips et al., 2000). Together, these two transporters could potentially mediate RVI in PI embryos, but such a function has not been demonstrated. There is some indirect evidence that mouse embryos accumulate inorganic ions when osmolarity is increased, implying that RVI might be occurring. Biggers et al. (1993) measured the intracellular ion content of two-cell embryos cultured

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in media of about 230 or 300 mOsM (produced by increasing NaCl from 85 to 125 mM), using the electron probe X-ray microanalysis technique. Calculations with their data indicate that the sum of the total content of major intracellular ions (Na+ + Cl− + K+) was slightly more than 20% higher when external osmolarity was higher, consistent with net ion import during RVI. There have been, however, no reported direct demonstrations of RVI in mammalian eggs or embryos. Unfertilized eggs and zygotes behave as ideal osmometers when challenged with an abrupt increase in osmolarity (Leibo, 1980; Oda et al., 1992; Dawson et al., 1998), but most other cells are similarly incapable of recovering from direct hypertonic shrinkage. Instead, more elaborate manipulations designed to produce volume loss “isosmotically” are required to reveal RVI (cf. O’Neill, 1999). Such studies have not been reported in embryos yet, and thus it remains to be determined whether mammalian PI embryos are capable of RVI.

C. Organic Osmolytes Despite the availability of inorganic ions, many cells preferentially accumulate organic compounds as major intracellular osmolytes. Across osmotolerant species from diverse phyla within every kingdom, the same small set of organic osmolytes has been found to be used to provide osmotic support, namely small neutral α- and β-amino acids, methylamines such as betaine, and small polyols such as sorbitol and myo-inositol (Yancey, 1994). The common property that distinguishes these organic osmolytes is their compatibility with biochemical function even at high concentrations (Brown and Simpson, 1972; Brown, 1976; Yancey et al., 1982); a large body of work convincingly demonstrates that high ionic strength severely disrupts macromolecular functions such as enzyme activity and macromolecular assembly, while in contrast, high concentrations of organic osmolytes—even molar concentrations—are nearly without effect (Yancey et al., 1982; Somero, 1986; Yancey, 1994). Thus, intracellular ionic strength can be kept constant and at an optimal level while the cell independently controls intracellular osmolarity. 1. Organic Osmolytes in Mammals The use of organic osmolytes by mammals was initially shown in the kidney, which is subjected to high and variable external osmolarity (Garcia-Perez and Burg, 1991), and in the brain, which is extremely susceptible to damage on swelling (McManus and Churchwell, 1994). The major organic osmolytes of the mammalian kidney are betaine, sorbitol, myo-inositol, taurine, and glycerophosphorylcholine (GPC) (Uchida et al., 1991; Garcia-Perez and Ferraris, 1994; Burg, 1995; Beck et al., 1998). Organic osmolytes used by cells in the mammalian brain include taurine, myo-inositol, and amino acids (Strange et al., 1991; S´anchez-Olea et al., 1992, Lang et al., 1998b). More recently, a number of other cell types in

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mammals have been shown to utilize organic osmolytes (especially amino acids), and an exhaustive survey of the literature documents widespread use of organic osmolytes throughout the body (Lang et al., 1998b). Thus, it may be that most mammalian cells require that a portion of their intracellular osmolarity be provided by organic osmolytes even if they are normally exposed to nearly constant external osmolarity in their environments. 2. Mechanisms Mediating Organic Osmolyte Accumulation Sorbitol and GPC are synthesized within the cells in which they are accumulated as organic osmolytes (Nakanishi and Burg, 1989; Burg, 1995). However, most organic osmolytes in mammalian cells are instead accumulated via specific transport systems situated in the plasma membranes. Four such osmolyte transport systems have been identified in mammals: the betaine transporter, system β or the taurine (β-amino acid) transport system, the Na+/myo-inositol transporter, and system A, which transports amino acids (Fig. 3). The betaine transport system is highly concentrative and requires cotransport of both Na+ and Cl− (Kwon and Handler, 1995; Matskevitch et al., 1999). The transporter protein, designated BGT1 [for betaine/γ -aminobutyric acid (GABA) transporter; Yamauchi et al., 1992], is closely related to the GABA transporters and is thus a member of the neurotransmitter transporter family (Schloss et al., 1994). It is highly expressed in renal tissues, brain, and liver (Liu et al., 1993a; Zhang et al., 1996). Taurine, like betaine, is accumulated as an osmolyte via an Na+- and Cl−dependent transporter that is also a member of the neurotransmitter transporter family (Uchida et al., 1991, 1992; S´anchez-Olea et al., 1992; Kwon and Handler, 1995; Liu et al., 1992) and that has been designated either NCT (for Na+-coupled taurine transporter) or TAUT. NCT/TAUT is widely expressed (Uchida et al., 1992; Kwon and Handler, 1995; Warskulat et al., 1997). In addition to taurine, the NCT/TAUT protein transports other β-amino acids such as hypotaurine and β-alanine with high affinity (Liu et al., 1992; Uchida et al., 1992; Warskulat et al.,

Figure 3 Mammalian organic osmolyte transport systems. The four known mammalian organic osmolyte transport systems and the ions each cotransports with its substrates are shown schematically. Details are given in text.

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1997), and thus is responsible for the well-described β-amino acid transport system or “system β” (the designation that will be used here for β-amino acid transport activity) found in many cells. The myo-inositol transporter is widely expressed and is especially abundant in renal and brain tissues. It is responsible for the accumulation of myo-inositol as an osmolyte (Kwon et al., 1992; Kwon and Handler, 1995; Porcellati et al., 1999). myo-inositol transport is Na+ dependent, but independent of Cl−. The transporter protein is designated SMIT (for sodium myo-inositol transporter), and is a member of the Na+-coupled glucose transporter family (Kwon et al., 1992). Neutral (zwitterionic) amino acids are transported by a nearly ubiquitous Na+dependent mechanism that has been designated system A (Christensen, 1984; Pastor-Anglada et al., 1996; Castagna et al., 1997), and that was recently cloned (Reimer et al., 2000). It functions to load cells with small, neutral amino acid osmolytes such as glycine, proline, and alanine (Yamauchi et al., 1994; Øyaas et al., 1995; Pastor-Anglada et al., 1996). A defining characteristic of system A is acceptance of a range of N-methyl amino acids such as N-methylamino-α-isobutyric acid (MeAIB) and methylated glycine derivatives such as sarcosine and betaine (Christensen et al., 1965; Petronini et al., 1994; Øyaas et al., 1995). 3. Regulation of Osmolyte Accumulation by Tonicity The known osmolyte transport systems—the betaine transporter (BGT1), myoinositol transporter (SMIT), system β (NCT/TAUT), and system A (Fig. 3)—all respond to increased tonicity by increasing their rate of substrate transport (Burg, 1995, 1997; Kwon and Handler, 1995). In each case the increased rate of transport is due to an increase in maximal transport velocity (Vmax) (Nakanishi et al., 1989, 1990; Uchida et al., 1991; S´anchez-Olea et al., 1992; Yamauchi et al., 1994). The rate of osmolyte transport increases slowly in response to hypertonicity, reaching a maximal level only hours to days after external osmolarity is increased (Garcia-Perez and Burg, 1991; Pastor-Anglada et al., 1996). Similarly, the rate of intracellular sorbitol synthesis via aldose reductase (AR) increases over the course of hours or days after an increase in osmolarity (Garcia-Perez and Ferraris, 1994; Beck et al., 1998). The slow time course of upregulation reflects a requirement for mRNA and protein synthesis, which in the case of the cloned osmolyte transporters has been shown to involve synthesis of the transporters themselves and for sorbitol involves synthesis of AR (Kwon et al., 1992; Uchida et al.,1992,1993; Yamauchi et al., 1992, 1993, 1994; Garcia-Perez and Ferraris, 1994; Warskulat et al., 1997). A cis-acting element required for the hypertonic stimulation of transcription, designated TonE (for tonicity response element), has been identified in several of the osmolyte transporter and AR genes (Takenaka et al., 1994; Miyakawa et al., 1999a; Rim et al., 1998; Ferraris et al., 1999), and a tonicity-induced transcription factor, termed TonE-binding protein, or TonEBP, has been identified (Miyakawa et al.,1999b).

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Thus, the generally accepted pattern of organic osmolyte accumulation is one of slowly increasing accumulation, which is mediated by increased transcriptionrequiring synthesis of the proteins that mediate accumulation of the osmolytes in the cytoplasm. This pattern is consistent with a model in which inorganic ion accumulation occurs quickly in response to an increase in osmolarity as the first defense against hypertonic insult, while the accumulation of organic osmolytes is a secondary process that acts to replace inorganic ions with benign osmolytes, thus keeping intracellular ionic strength at an optimal low level.

D. Organic Osmolytes and Embryos Van Winkle et al. (1990a) first proposed that mammalian embryos utilize organic osmolytes, and that their presence protects embryos against increased osmolarity. Lawitts and Biggers (1991b, 1992) similarly proposed that organic components of newly developed embryo culture media exerted their beneficial effects by functioning as compatible organic osmolytes. Evidence has since accumulated that early mammalian embryos can use a range of organic compounds for osmoprotection. 1. Improved Embryo Development in Media Containing Osmolytes Only one study has been done in which the effect of a large number of amino acids, which include many of the potential osmolytes, on PI embryo development in vitro was assessed systematically. McKiernan et al.(1995) examined the effect of each of the common α-amino acids and taurine, separately and in combination, on the development of hamster one-cell embryos to blastocysts. Three amino acids— glutamine, glycine, and taurine—were found to greatly stimulate development, and several others were found to stimulate development in the presence of glutamine (Table I). Unfortunately, no comparable in-depth studies comparing a large number of amino acids under the same conditions have been done with embryos of other species, although several individual amino acids have been shown to stimulate PI embryo development of a number of species. Glutamine has been found to be especially beneficial for the culture of PI embryos of mouse, hamster, cow, and human (Chatot et al., 1989; Lawitts and Biggers, 1991b,1993; Bavister and McKiernan, 1993; Biggers, 1998; Devreker et al., 1998; Steeves and Gardner, 1999), and is now a standard component of embryo culture media. Glycine and alanine, alone or in combination, were shown to improve development of bovine zygotes to blastocysts (Lee and Fukui, 1996). Taurine has also been reported to improve PI embryo development of these same species (Dumoulin et al., 1992a,b; Bavister and McKiernan, 1993; Spindle, 1995; Liu et al., 1995; Devreker et al., 1999), and hypotaurine, the direct precursor to taurine in its synthesis, improves hamster embryo development (Barnett and Bavister, 1992). An extensive body of

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Table I Comparison of Amino Acids That Stimulate Embryo Development and Organic Osmolytes Amino acids that area: Amino acid Alanine Arginine Asparagine Aspartate Cysteine Glutamate Glutamine Glycine Histidine Isoleucine Leucine Lysine Methionine Phenylalanine Proline Serine Taurine Threonine Tryptophan Tyrosine Valine

MDCK osmolytesb √

Stimulatory in hamsterc

ND √

— — √

— — — √ √ √f

— —e — √ √ √

— — — √

— — √

— √ √ √g √ ND — —

NESS + Gln + Taud √ — √ √ — √ √ √

— — √ √ √

— — — — — — √ √ √

— — — —

— — — —

√ A checkmark ( ) indicates amino acid is included in category; a dash (—) indicates it is not. ND, Not determined. b Defined as showing a 3-fold or higher increase in accumulation after 8 h in hypertonic versus isotonic medium (data from Horio et al., 1997). MDCK, Madin–Darby canine kidney cells. c McKiernan et al., (1995). d These are amino acids (Eagle’s nonessential amino acids plus glutamine and taurine) that as a group have been found to stimulate development of cleavage-stage embryos (Gardner and Lane, 2000). e Cysteine is inhibitory, but is stimulatory at a low concentration (McKiernan et al., 1995). f Histidine showed only a 2.6-fold hypertonic stimulation of accumulation, but then increased to a 7-fold increase after 6 days (Horio et al., 1997). Thus, it is included here. g Hypertonic stimulation of taurine accumulation was only about 2-fold (Horio et al., 1997), but has been well characterized as an osmolyte in MDCK cells (Uchida et al., 1991) as well as many other cell types (see text). a

work by Gardner, Lane, and co-workers has shown that, when added as a group, Eagle’s nonessential amino acids plus glutamine and taurine (NESS + Gln + Tau; Table I) are stimulatory to cleavage-stage mouse, bovine, and human embryo development (reviewed in Gardner and Lane, 2000; Steeves and Gardner, 1999). This group overlaps considerably with the stimulatory amino acids identified by McKiernan et al. (1995) in the hamster (Table I).

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Many of these amino acids that have been found to be beneficial to embryo development, especially glutamine, taurine, glycine, and alanine, are established organic osmolytes in other cell types (Yancey, 1994). In Madin–Darby canine kidney (MDCK) cells, in which the effect of osmolarity on intracellular amino acid content has been studied most systematically, a subset of the amino acids has been shown to be accumulated in response to hypertonicity and to act as organic osmolytes (Table I; Uchida et al., 1991; Horio et al., 1997). Comparing these with those amino acids found to stimulate PI embryo development reveals that the two groups are almost identical (Table I and above), raising the possibility that one of the major functions of beneficial amino acids in embryos is to act as organic osmolytes. 2. Organic Osmolytes Protect Preimplantation Embryos against Increased Osmolarity A compound that functions as an organic osmolyte must at least be able to protect the cell against the deleterious effects of increased osmolarity. Several compounds have been explicitly shown to confer such protection on PI embryos. a. Glutamine. Glutamine protects one-cell mouse embryos against the deleterious effects of increased osmolarity (Figs. 2 and 4). Lawitts and Biggers (1992) first showed that glutamine protected one-cell embryo development against increased NaCl concentration, which led them to propose that glutamine acts as an organic osmolyte in embryos (Lawitts and Biggers, 1992). Glutamine has the same protective effect against raised osmolarity regardless of whether osmolarity is raised with NaCl or with raffinose, indicating protection against osmolarity per se rather than NaCl (Fig. 2; Dawson and Baltz, 1997). Indeed, even when all NaCl in culture medium was replaced with raffinose, glutamine was still found to protect development against increased osmolarity (Dawson and Baltz, 1997). b. Glycine. Glycine also confers protection on one-cell mouse embryos against increased osmolarity (Fig. 4). It also is a good candidate for an endogenous organic osmolyte used by early PI embryos, because it is found at high concentrations in eggs and cleavage-stage embryos and is available in the oviduct (see below). Van Winkle et al. (1990a) found that glycine partially rescued the development of two-cell embryos to blastocysts from the deleterious effect of increased osmolarity in either a modified Spindle’s or “oviductal” medium (345–370 mOsM4). However, glycine was without effect at the normal osmolarity of Spindle’s medium 4 Higher osmolarities are cited in the original article (Van Winkle et al., 1990a) and were calculated as the sum of the concentrations of all components of the media, assuming all salts were completely ionized. This inadvertently overestimated the osmolarity, as the salts were not completely ionized at physiological concentrations. The corrected osmolarities cited here were measured in the author’s laboratory.

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Figure 4 Osmoprotective effects of several compounds in mouse one-cell embryos. In parallel experiments involving a number of compounds, glutamine, betaine, glycine, and proline (excluding β-amino acids, which are discussed separately) were found to be most effective at protecting mouse one-cell development to blastocysts in 310 mOsM medium (adjusted by adding raffinose; similar results were obtained when adjusting osmolarity with NaCl; Dawson and Baltz, 1997).∗∗∗ p < 0.001. [Adapted from Dawson and Baltz (1997) with permission of the Society for the Study of Reproduction.]

(265 mOsM4). Sarcosine, the N-methyl derivative of glycine, similarly protected two-cell embryos against increased NaCl, indicating that the protective effect was not due to glycine metabolism. Subsequently, glycine was found to be one of the most effective compounds at protecting mouse one-cell embryos against increased osmolarity when tested in parallel against a number of other compounds (Fig. 4 and Dawson and Baltz, 1997). Glycine improved embryo development in hypertonic medium in a dose-dependent manner, with maximal effectiveness reached by 0.5–1.0 mM (Fig. 5; Van Winkle et al., 1990a; Dawson and Baltz, 1997). It may also function as an osmolyte in rabbit embryos (Li and Foote, 1995). Glycine appears to be protective against increased osmolarity per se, rather than against increased NaCl concentration, because nearly identical results were obtained when osmolarity was increased with raffinose or NaCl (Dawson and Baltz, 1997). c. Betaine. Betaine is highly effective at protecting one-cell mouse embryos against hypertonicity (Fig. 4; Biggers et al., 1993). Development of mouse one-cell embryos derived from random-bred females was found to be virtually nonexistent in medium with an osmolarity increased to about 300 mOsM, while the addition of betaine allowed a significant proportion to develop to blastocysts at the higher osmolarity (Biggers et al., 1993). Betaine was also able to rescue PI mouse embryos from the depression of protein synthesis caused by hypertonicity (Anbari and

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Figure 5 Glycine and β-alanine act as an organic osmolyte in mouse embryos. Mouse one-cell embryos cultured in medium with osmolarity raised to 310 mOsM (with raffinose) will not develop to blastocysts in the absence of an organic osmolyte. However, either glycine or β-alanine could rescue development in a dose-dependent manner. Glycine was found to be effective at much lower concentrations than β-alanine, but each supports similar levels of development when present at maximally effective concentrations. [Adapted from Dawson and Baltz (1997) with permission of the Society for the Study of Reproduction.]

Schultz, 1993). Betaine protects one-cell mouse embryo development regardless of whether osmolarity has been raised to 310 mOsM by increasing NaCl, or by the addition of raffinose, indicating that betaine, like glycine and glutamine, protects against increased osmolarity rather than NaCl concentration (Dawson and Baltz, 1997). It was interchangeable with glutamine in providing osmoprotection, and the effects of suboptimal concentrations of glutamine and betaine were additive, indicating a common mechanism (Biggers et al., 1993). Thus, mouse embryos appear to be able to use betaine as an organic osmolyte. It is not known whether embryos other than mouse are protected from high osmolarity by the presence of betaine; at least in one other case, bovine embryos, betaine was not found to protect against high osmolarity under the conditions used (Liu and Foote, 1996). d. Taurine, Hypotaurine, and β-alanine. The β-amino acids taurine, hypotaurine, and β-alanine each protect mouse embryos against increased osmolarity to varying degrees. Taurine, hypotaurine, and β-alanine were equally effective at supporting development of mouse one-cell embryos to the four-cell or greater stages in hypertonic medium, approximately doubling development over that observed in the absence of added organic compounds (Dawson and Baltz, 1997). However, the efficacy of the three β-amino acids differed in supporting subsequent development to the blastocyst stage, with β-alanine or hypotaurine supporting maximal development, while taurine exhibited an insignificant effect (Fig. 6). This is similar to the results of Van Winkle et al.(1990a), where taurine was found to be ineffective at protecting mouse two-cell embryo development against increased osmolarity. Taurine is also reported to protect porcine germinal vesicle stage oocytes against

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Figure 6 Osmoprotective effect of substrates of the system β transport mechanism. System β substrates (5 mM each, chosen because β-alanine was shown to be maximally protective at this concentration) were tested for the ability to protect mouse one-cell embryo development to blastocysts against the detrimental effect of 310 mOsM medium. Columns with different letters are significantly different (p< 0.05). [Adapted from Dawson and Baltz (1997) with permission of the Society for the Study of Reproduction.]

the detrimental effect of increased NaCl (Funahashi et al., 1996). A dose response for β-alanine showed that maximal protection of mouse one-cell embryo development was reached at concentrations well above 1 mM, with a half-maximally effective concentration (EC50) of about 1 mM; this contrasts with the much lower EC50 of glycine, which is about 50 μM (Fig. 5; Dawson and Baltz, 1997). e. Other Osmolytes. Proline is highly effective at protecting one-cell mouse embryo development to blastocysts at increased osmolarity (Fig. 4; Dawson and Baltz, 1997). Alanine has been shown to protect mouse two-cell embryos against increased osmolarity, although maximal development to blastocysts was only about half that obtained with glycine under the same conditions (Van Winkle et al., 1990a). myo-Inositol, a major osmolyte in kidney, does not protect mouse or bovine embryo development against increased osmolarity (Dawson and Baltz, 1997; Liu and Foote, 1996). In addition, myo-inositol did not rescue the maturation of porcine germinal vesicle-stage oocytes from a deleterious effect of increased NaCl (Funahashi et al., 1996). In contrast, rabbit one-cell or two-cell embryos were protected against increased NaCl (at constant osmolarity) by myo-inositol in the medium, which was interpreted as evidence of its serving as an osmolyte in these embryos (Li and Foote, 1995).

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Nothing is known about whether PI embryos can synthesize either of the organic osmolytes that are produced intracellularly by other cells: sorbitol and GPC. It has been shown, however, that external sorbitol can protect porcine germinal vesicle-stage oocytes from damage by high NaCl (Funahashi et al., 1996), and that raising osmolarity with sorbitol was found to be less detrimental to rabbit and bovine embryo development than raising it with NaCl (Li and Foote, 1996; Liu and Foote, 1996). This might be consistent with a possible role for external sorbitol as an osmoprotectant in these species, which would have to enter the cell by an unknown mechanism rather than being synthesized intracellularly as in other cells. Alternatively, however, some other property of sorbitol might benefit embryos stressed by increased NaCl. 3. Osmolyte Transport by Embryos Because the osmoprotective effect of including organic osmolytes in embryo culture media is evident within the first two cell cycles (Lawitts and Biggers, 1992; Biggers et al., 1993; Dawson and Baltz, 1997), any osmolyte transport systems must be present and active at least in zygotes and early cleavage-stage embryos. Thus, it is especially important to determine which osmolyte transport systems exist in zygotes and early cleavage-stage embryos. There are four organic osmolyte transport systems described in a variety of cell types in mammals, as detailed previously: the betaine transporter, the myo-inositol transporter, system A for amino acids, and system β for β-amino acids (Fig. 3). There is no information available about whether a betaine-specific transport mechanism exists in cleavage-stage PI embryos, or whether the betaine transporter BGT1 is expressed. Betaine might be transported at low affinity via other mechanisms in embryos (see below), which may account for its ability to confer protection against hypertonicity. However, it is also possible that embryos express a high-affinity betaine transporter such as BGT1. myo-inositol is specifically transported by PI mouse, bovine, and rabbit embryos. At least a portion of myo-inositol transport is Na+ dependent (Kane et al., 1992; Hynes et al., 2000), indicating the presence of a mechanism resembling SMIT. Because rabbit embryos may be able to utilize myo-inositol as an osmolyte, SMIT may function in protection against hypertonicity in this species. However, in species such as mouse and cow, imported myo-inositol is likely used in the synthesis of phosphoinositides rather than having a role in osmotic protection. The widespread neutral amino acid transport mechanism, system A, can transport all of the α-amino acids shown to protect PI embryos from hypertonicity, such as glutamine, glycine, proline, and alanine (Yamauchi et al., 1994). In addition, it can transport betaine (Christensen et al., 1965; Petronini et al., 1994). Thus, system A could potentially be a mechanism responsible for transporting many organic osmolytes used by PI embryos. Surprisingly, however, cleavagestage PI embryos are devoid of activity attributable to the nearly ubiquitous

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system A. Through at least the two-cell stage, all detectable glycine transport occurs via the GLY transport system, with no component that could be attributed to system A (see below; Hobbs and Kaye, 1985, 1986, 1990; Van Winkle et al., 1988). In addition, transport of methionine, as well as other amino acids that are system A substrates, is entirely Na+ independent before the morula stage, indicating that there is no system A activity before this point (Borland and Tasca, 1974; Kaye et al., 1982; Van Winkle, 1988). Also, the model system A substrate, MeAIB, is not transported by embryos before the blastocyst stage, nor does it compete effectively for uptake of other amino acids (Kaye et al., 1982; Van Winkle et al., 1988). Indeed, system A is not detectable at any stage before the blastocyst, where it is expressed in the inner cell mass but not trophectoderm (Jamshidi and Kaye, 1995). Broad-scope Na+-dependent amino acid transport does appear by the eight-cell stage; however, this is attributable to the appearance of an Na+ and Cl−dependent transport system designated B0,+, which predominates in the blastocyst but is not found in early cleavage-stage embryos (Van Winkle et al., 1985). It is possible, however, that system A is normally quiescent and activated only by hypertonicity, and therefore would not have been detected in the studies cited above. However, culture of mouse zygotes in hypertonic medium for 24 h does not increase the rate of glycine uptake (Dawson et al., 1998), nor does hypertonicity cause an acute increase in the rate of glycine transport (Van Winkle et al., 1988; Dawson et al., 1998), making it unlikely that system A appears in embryos as a response to hypertonicity, although this possibility remains to be directly tested. a. System β. The only one of the four mammalian osmolyte transport mechanisms shown to be present and active in mouse PI embryos is system β, the β-amino acid transport mechanism. Taurine transport in embryos is entirely Na+ and Cl− dependent, taurine and β-alanine serve as competitive inhibitors of each other’s transport consistent with system β characteristics, and mRNA encoding NCT/TAUT can be detected at every stage from oocytes through blastocysts (see Van Winkle et al., 1994; Van Winkle and Campione, 1996). In addition to transporting β-amino acids, the β-amino acid transport mechanism in embryos displays the expected high affinity for GABA, and probably also transports alanine and arginine with low affinity (Van Winkle et al., 1994). Transporter activity stays approximately constant from the one-cell through four- to eight-cell stages, and then increases about 3-fold at the blastocyst stage, with a concomitant increase in the taurine content of embryos (Van Winkle and Campione, 1996). Whether the synthesis of β-amino acid transporter (NCT/TAUT) mRNA in PI embryos can be regulated by tonicity as in other cells is not known. In other cells, there is a lag time of hours or even days between an increase in tonicity and increased transport rates, caused by the time needed for transcription and synthesis of new transporter proteins. However, even the longest persisting PI embryo stage—the one-cell stage—lasts less than 24 h, and thus the most osmotically sensitive stages would be largely past before upregulation could occur.

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Furthermore, the osmolarity of the oviductal environment probably does not vary significantly. Therefore, if synthesis of a β-amino acid transporter protein in embryos proves to respond to changes in osmolarity, it would seem probable that the function of such a feedback would be to ensure a continuous production of transporters to maintain a constant optimal transport rate, rather than to respond to osmotic perturbations. An osmotically induced increase in system β activity in early mouse embryos may instead be regulated by a quickly responding mechanism not common or obvious in other cells. One-cell-stage mouse embryos respond to increased osmolarity by quickly increasing the rate of taurine transport via system β (Fig. 7; Van Winkle et al., 1994), a phenomenon that is possibly similar to the fast stimulation of taurine transport by hypertonicity reported to occur in fish erythrocytes (Fincham et al., 1987). In contrast, system β activity in blastocysts was not affected by osmolarity over the same range (Van Winkle et al., 1994). Thus, the early PI embryo may possess a mechanism that permits a rapid increase in the accumulation of taurine and other β-amino acids in response to hypertonicity, and that is lost prior to the blastocyst stage.

Figure 7 Osmotic response of GLY and system β activities in one-cell mouse embryos. Taurine transport via system β in one-cell embryos was stimulated by increased osmolarity over the range from 200 and 300 mOsM. In contrast, glycine transport by GLY was relatively constant over that range, decreasing only when osmolarity fell well below 200 mOsM. Although the precise range over which system β activity in embryos is osmosensitive is not known, it is possible that its responsive range covers physiologically relevant changes in embryo volume. However, GLY activity may not be regulated by changes in cell volume normally encountered by the embryo, because it is affected by osmolarity only at hypotonic values. The curves are shown only for clarity, and were fit by nonlinear regression to a sigmoid. The data for taurine transport were taken from Van Winkle et al. (1994) with the rate at 440 mOsM (not shown) normalized to 100, and those for glycine transport were taken from Dawson et al. (1998) with the rate at 300 mOsM normalized to 100.

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b. GLY Transport System. Although it is clear from the foregoing discussion that any use of β-amino acids as organic osmolytes by PI embryos can be accounted for by the activity of system β, there are a number of other compounds that have excellent osmoprotective properties in embryos but are certainly not system β substrates, including glycine, betaine, glutamine, and proline. There are several transport systems active in early PI embryos which can accept at least some of these as substrates. However, the Na+- and Cl−-independent amino acid transport mechanisms in embryos, which are predominantly the L system in cleavage-stage embryos, and the b0,+ system in blastocysts (Van Winkle et al., 1990b), cannot mediate the highly concentrative intracellular accumulation required for osmolytes. Indeed, the one-cell and early cleavage-stage embryo has only one other major ion-dependent amino acid transport mechanism, the GLY transport system, in addition to system β. As a consequence, most amino acid transport except for that of glycine and β-amino acids is Na+ independent in PI embryos, not becoming Na+ dependent until compaction and the blastocyst stage (Borland and Tasca, 1974; Kaye et al., 1982; Van Winkle, 1988). Given that osmolyte accumulation must be mediated by concentrative, ion-linked transport, it has therefore been proposed that GLY and system β account for the ability of early embryos to accumulate organic osmolytes (Van Winkle et al., 1990a; Dawson and Baltz, 1997). The GLY transporters are members of the neurotransmitter transporter family (Schloss et al., 1994), and are therefore related to both the system β transporter (NCT/TAUT) and the betaine transporter (BGT1). Like other members of this family, GLY transport is dependent on both Na+ and Cl− (Aragon et al., 1987; Roux and Supplisson, 2000). The widely expressed GLY transport activity is encoded by the Glyt1 gene (Guasatella et al., 1992; Borowsky et al., 1993), from which arise several alternate transcripts; the Glyt1a isoform appears to be responsible for general GLY activity (Smith et al., 1992; Borowsky et al., 1993; Kim et al., 1994; Borowsky and Hoffman, 1998). Another gene, Glyt2, encodes a neuronal GLY-like transporter that differs from the common GLY activity in rejecting sarcosine as a substrate (Liu et al., 1993b). In neurons, GLY-like transport mediates reuptake of glycine from glycinergic and N-methyl-D-aspartate (NMDA)-ergic synapses. However, why a highly concentrative transporter specific for glycine, such as GLY, is required in nonneuronal tissue is unknown, but implies that it may have a function other than simply supplying glycine for metabolism, because other widespread transport mechanisms could adequately supply glycine for metabolic requirements (Christensen, 1984). Glycine is transported exclusively via GLY in early cleavage-stage mouse embryos (Hobbs and Kaye, 1985, 1986, 1990; Van Winkle et al., 1988). GLY activity is highest at the one-cell to two-cell stages, decreasing before compaction, and completely disappearing by the blastocyst stage (Hobbs and Kaye, 1985; Van Winkle et al., 1988; Van Winkle and Campione, 1996; Hammer et al., 2000). Glycine continues to be transported robustly after compaction, but occurs predominantly via the B0,+ transport system rather than GLY in blastocysts (Van Winkle et al.,

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1985,1988). In mouse embryos, GLY activity appears to be encoded by the Glyt1 gene, because Glyt1 mRNA is detectable in unfertilized eggs and in embryos before compaction, but not in morulae or blastocysts, consistent with the expression of GLY transport activity (Van Winkle and Campione, 1996). Glycine transport in mouse eggs and cleavage-stage embryos is completely inhibited by excess sarcosine (Van Winkle et al., 1988; Dawson et al., 1998; Hammer et al., 2000), and saturable glycine transport by cleavage-stage human embryos is similarly sarcosine inhibitable (Hammer et al., 2000), consistent with transport of glycine and sarcosine in embryos by GLY. It is unclear whether GLY can accept betaine as a substrate, but given the tolerance of the transporter for N-methyl groups, it would appear possible that betaine (N,N,N-trimethylglycine) could be a low-affinity substrate, analogous to the tolerance of system A for sarcosine, N,N-dimethylglycine, and betaine (Christensen et al., 1965). In addition to glycine and sarcosine, GLY accepts a limited array of other substrates. Proline competitively inhibits GLY activity in mouse PI embryos, indicating that it can be transported by GLY [tested as proline (Dawson et al., 1998) or as its analog pipecolate (Van Winkle et al., 1988)] and in other cells (Ellory et al., 1981; Kim et al., 1994). About one-third of glutamine transport in two-cell mouse embryos is Na+ dependent, and glutamine inhibits the uptake of glycine by two-cell embryos when present at millimolar levels, characteristics proposed to indicate low-affinity transport of glutamine in embryos via GLY (Lewis and Kaye, 1992). The GLY transport system in mouse one-cell embryos does not appear to be stimulated by increased osmolarity over the same range of osmolarities as system β (Fig. 7). Instead, the rate of transport was essentially constant from 200 to 350 mOsM, with an osmotic effect on transport seen only at about 150 mOsM. Indeed, the rate of glycine transport increased by only 30% when osmolarity was increased from 250 to 350 mOsM with raffinose (Dawson et al., 1998). Van Winkle et al. (1990a) also reported a similar lack of significant acute stimulation of glycine uptake in one- and two-cell mouse embryos immediately after osmolarity was increased with mannitol. Furthermore, total GLY activity is also not increased after prolonged culture at increased osmolarity. When mouse zygotes were cultured overnight at several osmolarities, the activity of GLY in the resulting two-cell embryos was identical regardless of whether they had been cultured at 250, 310, or 340 mOsM (Dawson et al., 1998). c. Do GLY and System β Account for All Osmolyte Transport in Embryos? So far, there is convincing evidence that early cleavage-stage mouse embryos are effectively protected against increased tonicity by glycine, glutamine, betaine, proline, β-alanine, and hypotaurine, at concentrations between 0.5 and 5.0 mM (see above; Van Winkle et al., 1990a; Biggers et al., 1993; Lawitts and Biggers, 1992; Dawson and Baltz, 1997). To a lesser extent, sarcosine, alanine, and possibly taurine are also osmoprotective (Van Winkle et al., 1990a; Dawson and Baltz, 1997). It was proposed (Dawson and Baltz, 1997) that these putative organic osmolytes

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in embryos could be divided into two groups consisting of proved or possible substrates of either GLY or system β. Glycine and sarcosine are classic highaffinity substrates of GLY, while glutamine and proline are probably low-affinity substrates. Betaine, as an N-methyl derivative of glycine, might be transported by GLY with low affinity as proposed above. Therefore, the osmoprotective effects of glycine, glutamine, proline, sarcosine, and possibly betaine could be explained by their transport via the GLY system in embryos. The remaining proved osmoprotective compounds are β-alanine, hypotaurine, alanine, and to a lesser extent taurine (Van Winkle et al., 1990a; Dawson and Baltz, 1997). The three β-amino acids are clearly system β substrates, while alanine has been shown to be a low-affinity system β substrate. Thus, all those osmoprotective compounds that are not proved or potential GLY substrates are system β substrates. Substrates of either transport mechanism apparently exert their protective effect via a common mechanism, because the effect of submaximal concentrations of two model osmoprotective substrates of each system, glycine and β-alanine, were additive (Fig. 8). For such structurally and functionally different compounds, the most likely mechanism for a common action is as intracellularly accumulated organic osmolytes. It seems likely, therefore, that both GLY and system β function as organic osmolyte transporters in one-cell and early cleavage-stage mouse embryos, and this proposal is reiterated here. It should be noted, however, that while

Figure 8 Additive effect of submaximal concentrations of glycine and β-alanine. Glycine or β-alanine was added to 310 mOsM culture medium near their half-maximally effective concentrations (0.05 and 1.0 mM, respectively). Their ability to rescue development of one-cell mouse embryos to blastocysts was assessed either singly or in combination, and it was found that they exerted an additive effect, indicating a shared mechanism. Columns with different letters are significantly different ( p < 0.05). [Adapted from Dawson and Baltz (1997) with permission of the Society for the Study of Reproduction.]

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system β is a well-established osmolyte transport mechanism in an array of cell types, this would be a novel function for GLY, and thus extensive work must be done to firmly establish GLY as an osmolyte transport system in embryos. The question of whether GLY and system β are the only osmolyte transporters in embryos has not been resolved. Although a number of amino acids and potential osmolytes have been tested for osmoprotective effects, most of these are either GLY or system β substrates. Only a few other amino acids have been assessed, such as leucine (the model substrate of the L amino acid transport system), and lysine (a substrate of the b0,+ and B0,+ systems), and were not found to protect embryos against increased osmolarity (Dawson and Baltz, 1997). In addition, myo-inositol was not found to act as an osmolyte in mouse embryos, as described above, although it may function as such in rabbit embryos. Therefore, no organic compounds identified as osmoprotective in mouse embryos can be clearly excluded as substrates of either GLY or system β, and so there is no compelling evidence yet for additional osmolyte transport systems in embryos. Perhaps the most likely candidate at present for a third osmolyte transporter in embryos would be one that mediates high-affinity betaine transport. Betaine is an excellent osmoprotective compound in early mouse embryos (see above; Biggers et al., 1993; Anbari and Schultz, 1993; Dawson and Baltz, 1997), and it is tempting to speculate that, in addition to GLY and system β, early embryos might possess a betaine transporter such as BGT1. 4. Osmotically Regulated Accumulation of Osmolytes in Embryos Strong evidence that a compound is utilized as an organic osmolyte would be the demonstration that its intracellular accumulation depends on external tonicity, with accumulation increasing as tonicity increases. There is indirect evidence of increased accumulation of two putative osmolytes—betaine and taurine—and direct evidence of osmotically stimulated accumulation of glycine in embryos. The total Na+ and Cl− contents of two-cell mouse embryos were found to be lower when betaine was present in hypertonic medium than when it was absent, while in contrast the presence of betaine did not affect the intracellular content of Na+ or Cl− in medium of lower tonicity (Biggers et al., 1993). This would imply that betaine is accumulated intracellularly to displace Na+ and Cl− only under hypertonic conditions and thus that its accumulation might be stimulated by increased tonicity. For taurine, the hypertonic stimulation of system β activity that has been demonstrated in mouse embryos (see above; Van Winkle et al., 1994) would likely result in the increased accumulation of intracellular taurine or other β-amino acids as in other cells. Glycine is the only putative organic osmolyte whose total intracellular concentration in embryos has been shown directly to be a function of osmolarity. Dawson et al. (1998) measured total intracellular glycine accumulation by mouse zygotes cultured for 24 h at different osmolarities (adjusted with raffinose) in the presence

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Figure 9 Glycine accumulation by mouse one-cell embryos is regulated by osmolarity. The amount of glycine accumulated after 24 h at 250, 310, or 340 mOsM is shown at left as a function of external glycine concentration (numbers labeling curves indicate osmolarity). At right, the result for 1.0 mM glycine is shown, demonstrating that a significantly higher amount of glycine is accumulated by embryos after 24 h at 310 or 340 mOsM than at 250 mOsM. This result with 1.0 mM glycine has subsequently been repeated numerous times in the author’s laboratory and is highly reproducible (Hammer et al., 2000; and M. A. Hammer, C. L. Steeves, and J. M. Baltz, unpublished data, 2000). [Adapted from Dawson et al. (1998) with permission of the Society for the Study of Reproduction.]

of glycine. The amount of glycine accumulated when osmolarity was 310 or 340 mOsM was found to be significantly increased over the amount accumulated when the osmolarity was 250 mOsM, consistent with a role for glycine as an osmolyte (Fig. 9). Maximal intracellular accumulation was achieved at external glycine concentrations above about 0.5–1.0 mM (Dawson et al., 1998), approximately the same glycine concentration that was found to maximally protect one- and twocell embryo development against increased osmolarity (Fig. 5; Van Winkle et al., 1990a; and see above). How increased tonicity results in an increase in intracellular glycine concentration has not been determined, because GLY transport activity seems to be only slightly stimulated by increased tonicity (above). Although the maximum concentrations of glycine accumulated in embryos cultured in 310 and 340 mOsM media were not different, it is interesting to note that the maximal accumulation of glycine was reached at a lower external glycine concentration, at 340 rather than 310 mOsM (Fig. 9). Although this phenomenon has not yet been investigated, it may indicate that the transport of glycine is regulated either at the level of its affinity, or by a mechanism sensitive to the glycine gradient, such as a leak pathway mediating the efflux of glycine from the embryo (e.g., the swelling-activated channel discussed below). Another indirect indication that glycine is accumulated intracellularly by PI embryos in response to increased tonicity is its effect on cell volume. Mouse zygotes transferred to 310 mOsM medium maintained larger cell volumes if glycine was present in, rather than absent from, the medium (Fig. 10; Dawson et al., 1998). Zygotes in 310 mOsM medium that contained glycine maintained approximately the same sizes in 310 mOsM medium as zygotes in 250 mOsM medium, while in contrast zygotes at 310 mOsM in the absence of glycine were found to

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Figure 10 Volume of one-cell embryos in medium of different osmolarities in the presence or absence of glycine. (A–C) Volumes of one-cell embryos as a function of time after transfer to media of osmolarities of 250, 310, and 340 mOsM, respectively, as indicated. At 310 mOsM, the presence of glycine allowed the embryos to maintain a significantly higher volume than in the absence of glycine (∗∗∗ p < 0.001 or ∗∗ p < 0.01; dashed line added to facilitate comparison). [From Dawson et al. (1998), used with permission of the Society for the Study of Reproduction.]

shrink significantly (Fig. 10). A similar support of cell volume in hypertonic media has been shown for glycine accumulated via system A in Ehrlich ascites tumor cells (Hacking and Eddy, 1981; Hudson and Schultz, 1988). Thus, glycine appears to be established as an organic osmolyte used by early PI embryos, because its intracellular concentration is sensitive to external osmolarity, it allows cell volume to be maintained when tonicity is raised, and the embryo possesses a specific concentrative transporter for glycine.

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The ability of embryos to use glycine as an osmolyte is largely restricted to the one-cell and early cleavage-stages. One-cell mouse embryos cultured to the twocell stage show a marked osmosensitivity of glycine accumulation as described above (Fig. 9). However, glycine accumulation in two-cell embryos cultured to the four-cell stage was barely dependent on osmolarity, while those cultured to greater than four cells, and morulae cultured to blastocysts, did not respond at all to increased tonicity (Hammer et al., 2000). It is interesting that the loss of the ability of embryos to accumulate glycine as an osmolyte in response to increased tonicity essentially parallels the decrease in GLY activity after the two-cell stage of PI embryo development and its complete disappearance by the blastocyst stage (Van Winkle and Campione, 1996), providing further indication that transport via GLY is required for the use of glycine as an osmolyte in embryos.

IV. Regulation against Volume Increases by Mammalian Embryos A. Regulation against Volume Increases Cells are able to recover from volume increases by exporting osmotically active substances from the cytoplasm. Such an ability to recover from cell swelling is termed a regulatory volume decrease, or RVD. A few types of mammalian cells such as erythrocytes mediate RVD by exporting K+ and Cl− via cotransporters of the KCC family (Sarkadi and Parker, 1991; Warnock and Eveloff, 1989; Gillen et al., 1996; Lang et al., 1998b; Mount et al., 1999). However, the vast majority of cell types release osmolytes on swelling primarily via separate but functionally coupled K+ and anion channels (Lang et al., 1998b). Thus, in most cells RVD is found to be inhibited by an array of chemically unrelated Cl− channel and K+ channel blockers (Knoblauch et al., 1989; Sarkadi and Parker, 1991; MacLeod et al., 1992; Lippmann et al., 1995; S´anchez-Olea et al., 1996; Pasantes-Morales et al., 1997). Consistent with the participation of a swelling-activated Cl− channel in RVD, almost all vertebrate cell types examined have been found to exhibit a pronounced swelling-activated Cl− current (Strange et al., 1996; Okada, 1997; Lang et al., 1998b). Although there may be several types of swelling-activated Cl− channel, the most widespread swelling-activated Cl− current has a unique set of characteristics. This current is outwardly rectified in symmetric Cl− and has a higher permeability to I− than Cl− (Strange et al., 1996; Okada, 1997). It is inhibited not only by typical Cl− channel blockers (Strange et al., 1996; Okada, 1997), but also by external ATP at millimolar levels (Jackson and Strange, 1995; Tsumura et al., 1996). Perhaps the most important feature of the swelling-activated anion channel is its substantial permeability to organic osmolytes in addition to inorganic anions. Swelling-activated currents mediated by anionic amino acids such as aspartate and glutamate (Jackson et al., 1994; Roy, 1995), the ionized forms of glycine and taurine (Jackson and

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Strange, 1993; Roy, 1995; Manolopoulos et. al., 1997), or by other organic anions such as pyruvate, butyrate, and acetate (Jackson et al., 1994), can be directly demonstrated when these organic anions are substituted for inorganic anions. Furthermore, competition experiments have shown that the same swelling-activated channel carries organic compounds such as inositol, aspartate, and glutamate as carries Cl− (Jackson and Strange, 1993; Levitan and Garber, 1998). A swelling-activated efflux of organic osmolytes with the same time course as RVD has been demonstrated in a large number of cell types (Kirk et al., 1992; Strange and Jackson, 1995; Strange et al., 1996). This organic osmolyte efflux has the same pharmacology as the swelling-activated current (Jackson and Strange, 1993; Strange et al., 1996; Okada, 1997), and therefore it was postulated that the osmolyte efflux was mediated by the same channels that give rise to the swellingactivated Cl− and organic anion currents (Roy and Malo, 1992; Kirk et al., 1992; Jackson and Strange, 1993). It is now generally accepted that, at least in the majority of cases, swelling-activated organic osmolyte efflux and swelling-activated Cl− and organic anion currents are both manifestations of the same underlying anion channel that is activated by cell swelling. Thus, because it is activated by cell swelling, participates in RVD, and is permeable to organic osmolytes, the swelling-activated channel involved in all these processes has been termed the volume-sensitive organic osmolyte/anion channel (VSOAC; Jackson et al., 1996). Several proteins have been proposed to underlie the VSOAC current, including the small protein pICln (Paulmichl et al., 1992), the multidrug resistance P glycoprotein (Valverde et al., 1992), and two members of the ClC Cl− channel family, ClC-2 (reviewed in Strange et al., 1996) and ClC-3 (Duan et al., 1997, 1999). It now seems unlikely that the first two function as swelling-activated channels, although they may influence channel activity (Pasantes-Morales et al., 1997; Mor´an et al., 1997; Strange, 1998; Emma et al., 1998), and the properties of ClC-2 differ from those of the widespread VSOAC current (Strange et al., 1996). The best candidate at present for a VSOAC-like channel is ClC-3, because its electrophysiological and pharmacological properties largely match those of VSOAC (Duan et al., 1997, 1999). However, whether ClC-3 is the channel responsible for VSOAC currents in most cells, or whether there are other VSOAC-like channels, is not yet certain (Strange, 1998; Wang et al., 2000).

B. Regulation against a Volume Increase in Embryos 1. Regulatory Volume Decrease in Embryos Mouse one-cell embryos are capable of recovering from an increase in volume. After hypotonic swelling, mouse one-cell embryos recovered with a mean halftime of about 10 min after peak swelling, and were fully recovered and restored

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Figure 11 Recovery from hypotonic swelling (RVD) by one-cell mouse embryos. One-cell mouse embryos recover from swelling in hypotonic medium (150 mOsM hypotonic exposure indicated by solid bars at bottoms of each graph) within about 10 min (control). RVD is blocked by either K+ channel blockers (Ba2+, quinine) or Cl− channel blockers (DIDS, NPPB). These are typical examples of 10–51 replicates, as described in S´eguin and Baltz (1997). [Figure adapted from S´eguin and Baltz (1997) with permission of the American Physiological Society.]

to their initial volumes within about 25 min (Fig. 11; S´eguin and Baltz, 1997). RVD in mouse embryos appears to be mediated by separate Cl− and K+ channels, because either Cl− channel blockers or K+ channel blockers separately prevented RVD in mouse one-cell embryos (Fig. 11; S´eguin and Baltz, 1997). Consistent with there being separate Cl− and K+ channels functioning in parallel, addition of the cation-selective ionophore gramicidin partially restored RVD in one-cell embryos when recovery had been blocked by the K+ channel blocker quinine, but not when it was blocked by the Cl− channel blocker diisothiocyanatostilbene 2,2 -disulfonic acid (DIDS) (S´eguin and Baltz, 1997).Van Winkle et al. (1994) also reported that mouse one-cell embryos could recover from swelling, although the RVD they observed was slower, with full recovery occurring within about 60 min after peak swelling. The K+ channels mediating RVD in mouse embryos have not been identified. Oocytes and early PI embryos have a number of different types of K+ channels (Day et al., 1991), including Ca2+-activated K+ channels (Yoshida et al., 1990). The K+ channels required for RVD in the mouse embryo would not seem to be Ca2+ dependent, however, because chelating intracellular Ca2+ did not slow RVD, and toxins that block several subtypes of Ca2+-dependent K+ channels were without effect (S´eguin and Baltz, 1997). It is also not known whether the K+ channels required for embryo RVD are activated by cell swelling; no evidence of a hypotonically induced increase in K+ current has been observed in mouse onecell embryos (M. Kolajova and J. M. Baltz, unpublished data, 2000). In contrast, swelling-activated Cl− channels are now well characterized in one-cell mouse embryos (Kolajova and Baltz, 1999).

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2. Swelling-Activated Cl− Channels in Embryos The presence of Cl− channels that are activated by swelling has been directly demonstrated in mouse one-cell embryos, using whole-cell patch-clamp electrophysiological recordings (Fig. 12A and B; Kolajova and Baltz, 1999). This swelling- activated current is outwardly rectified in symmetric Cl− (Fig. 12C) and inhibited by Cl− channel blockers [e.g., DIDS and 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB); Fig. 12D and E] and by millimolar amounts of external ATP (Fig. 12F), each of which had also been shown to block RVD in embryos (S´eguin and Baltz, 1997). The swelling-activated current has an anion selectivity of I− > Cl−, and both aspartate (Fig. 12G) and taurine were capable of carrying this current in embryos (Kolajova and Baltz, 1999). These characteristics are consistent with the presence in embryos of a VSOAC-type swelling-activated Cl− and organic osmolyte channel. A similar swelling-activated current has also been demonstrated in human failed-fertilized eggs (Hammer et al., 2000). Thus, it is proposed that the one-cell embryo possesses swelling-activated channels that are identical to the VSOAC channels in other cells, and that these are part of the mechanism by which embryos regulate their volumes. The unfertilized mouse egg also exhibits this swelling-activated Cl− current, although to a somewhat reduced degree (Kolajova and Baltz, 1999). In preliminary work, the current appeared to be less sensitive to DIDS than in fertilized eggs (Kolajova and Baltz, 1999), but subsequent work has shown that the majority of the swelling-activated current in unfertilized eggs is inhibitable by DIDS and has the same general characteristics as that in one-cell embryos, indicating that the same type of channel is likely present (M. Kolajova and J. M. Baltz, unpublished data, 2000). It has not been reported whether the swelling-activated current is present in later stage embryos. Our initial investigations indicate that the swellingactivated channel is present through the two-cell stage, but that activity declines sharply by the four-cell stage and remains low during the remainder of the cleavage stages (M. Kolajova and J. M. Baltz, unpublished data, 2000). 3. Permeability to Osmolytes Is Increased by Swelling in Embryos If early PI embryos mediate RVD via the swelling-induced opening of a VSOAClike channel, then embryos should be able to regulate against a volume increase by releasing intracellular organic osmolytes through this channel, as shown in other cells (above). Consistent with this, two-cell mouse embryos that had been preloaded with either 3H-labeled taurine or glycine were found to retain most of the labeled amino acid for at least several hours under isotonic conditions, but to release almost all the labeled amino acid when hypotonically swelled (Fig. 13; Dumoulin et al., 1997; Dawson et al., 1998). An identical result was also reported for human failed-fertilized eggs and spare embryos (Dumoulin et al., 1997). Hypotonically stimulated osmolyte release was complete within 30 min, a period

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Figure 12 Swelling-activated currents in mouse zygotes. (A) Current-versus voltage plot showing the swelling-induced activation of current in a mouse one-cell embryo obtained from patch-clamp measurement in whole-cell configuration as described in Kolajova and Baltz (1999). Minimal current was evident when osmolarity was 250 mOsM (250), but increased substantially in hypotonic 180 mOsM medium (180). The current increase was reversed when the embryo volume was again decreased in 330 mOsM medium (330). (B) Mean current and conductance (slope) at +60 mV derived from current– voltage plots as shown in (A) obtained for a number of one-cell embryos, as a function of their measured diameter (n = 58 at 250 and 180, n = 7 at 330 mOsM). Both current and conductance increase with cell swelling. (C) Outward rectification of swelling-activated current evident in symmetric Cl− (80 mM Cl− inside and out). (D and E) Inhibition of swelling-activated current by the Cl− channel blockers DIDS (100 μM) and NPPB (100 μM) in one-cell embryos swelled in 180 mOsM medium. (F) Inhibition of swelling-activated current in 180 mOsM medium by external ATP (5 mM). (G) Swelling-activated current in 180 mOsM medium was observed when all Cl− was replaced with aspartate (Asp), and was inhibitable by DIDS (Asp + DIDS). [Adapted from Kolajova and Baltz (1999) with permission of the Society for the Study of Reproduction.]

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Figure 13 Increased efflux of accumulated amino acids from two-cell mouse embryos induced by swelling. Two-cell mouse embryos were allowed to accumulate either [3H]taurine (circles) or [3H] glycine (triangles) and then transferred to amino acid-free medium. The osmolarity of the amino acid-free medium differed from the osmolarity during accumulation by the amount indicated on the horizontal axis. The embryos were incubated in this medium for 4 h and then the amount of 3 H-labeled amino acid remaining was determined. Efflux of either taurine and glycine was substantially higher when the efflux medium was hypotonic than when it was isotonic or hypertonic, indicating that cell swelling increased the efflux of taurine and glycine from embryos. Data for taurine were taken from Dumoulin et al. (1997), and those for glycine were taken from Dawson et al. (1998). The osmolarity during amino acid accumulation was 280 mOsM for taurine, and either 250 mOsM (open triangles) or 310 mOsM (closed triangles) for glycine. Glycine efflux reached a plateau within 30 min (Dawson et al., 1998) but the 4-h time point was used to facilitate direct comparison between the two studies. The striking similarity of the responses for the two different amino acids in the two separate studies implies that a common mechanism mediates the swelling-induced release of both.

that corresponded to the RVD observed in these embryos under the same conditions (Dawson et al., 1998). Interestingly, not only was significantly more glycine accumulated when the embryos had been cultured at 310 versus 250 mOsM, as described above, but a larger proportion of the increased amount of glycine accumulated at 310 mOsM was released on hypotonic swelling, indicating that an increased proportion was present in an osmotically active form (Dawson et al., 1998). Another method of demonstrating a swelling-activated increase in permeability to organic osmolytes is to determine whether there is a swelling-induced increased in the rate of influx of labeled osmolytes present in the external medium. This is done in Na+-depleted medium to eliminate interference by Na+-dependent transport systems (Van Winkle et al., 1994). Although the direction of transport in this case is “backward” from that which would mediate RVD, this method quickly reveals any increase in permeability induced by swelling without the need for preloading. In this way, the permeability of one-cell mouse embryos to external taurine had been shown to increase by more than 10-fold by hypotonic swelling (Fig. 14; Van Winkle et al., 1994), and the increased permeability was reversed

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Figure 14 Increase in permeability to a subset of compounds induced by hypotonic swelling in onecell mouse embryos. One-cell mouse embryos were swelled by exposure to hypotonic medium, and their permeability to 3H-labeled amino acids in the external medium was determined. Permeability was assessed in swelled and unswelled embryos by measuring the amount of amino acid that entered the embryo under each condition within a fixed period of time (see text). The increase in permeability in swelled embryos (expressed relative to the permeability of unswelled embryos) is shown. Thus, the value of 1 for alanine indicates no increase in permeability on swelling, whereas for example, glycine permeability increased by about 4-fold in swelled embryos. These data are from two sources, indicated by the letters after the amino acids at bottom: (a) unpublished data from the author’s laboratory, obtained by K. M. Dawson and M. A. Hammer. Embryos were swelled in 190 mOsM medium, and compared with controls in 290 mOsM medium. A 1–2 μM concentration of each 3H-labeled compound (10– 130 Ci/mmol; Amersham or New England Nuclear) was used, and uptake by groups of 10 embryos was assayed after a 15-min exposure as described in Dawson et al., (1998). N = 4–12 groups of 10 embryos for each compound tested; (b) data estimated from Fig. 6 inVan Winkle et al. (1994), where details can be found. The symbols above the columns indicate significant difference (∗∗ p < 0.01 and ∗∗∗ p < 0.001, by t test versus unswelled; # significantly different as described in Van Winkle et al., 1994; NS, p > 0.05). The second column (Gly + DIDS) shows the relative permeability to glycine in swelled embryos in the presence of 100 μM DIDS. DIDS largely blocked the increase in permeability (comparison indicated by the line, ∗∗∗ p < 0.001), but did not affect the extent of swelling (not shown).

by Cl− channel blockers, which inhibit the VSOAC channel.5 The permeability of 1-cell embryos to external glycine was also increased by swelling, and this increased permeability was inhibited by the Cl− channel blocker DIDS (Fig. 14; Van Winkle et al., 1994). In addition, β-alanine (Fig. 14; Van Winkle et al., 1994) 5 The effectiveness of several inhibitors (niflumate and furosemide) differed from what had been reported for other cells at the time some of this work was done (Van Winkle et al., 1994), and thus it was postulated that the swelling-activated pathway in embryos differed from that in other cells. However, it has since become clear that the various inhibitors vary among cell types in their effectiveness in inhibiting VSOAC currents (Strange et al., 1996; Okada, 1997). Thus, coupled with direct electrophysiological evidence of VSOAC currents in mouse embryos (Kolajova and Baltz, 1999), it would seem probable that the same mechanism underlies the swelling-activated increase in taurine permeability.

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and to a lesser extent proline and aspartate (Fig. 14) exhibit increased permeabilities in hypotonically swelled one-cell embryos, while, in contrast, the influx of alanine and lysine were not significantly affected by hypotonic swelling (Fig. 14; Van Winkle et al., 1994). The inhibition by Cl− channel blockers and the observed amino acid selectivity are again consistent with a swelling-activated VSOAC-like channel in early PI embryos. As discussed above, a high level of swelling-activated channel activity may be restricted to the earliest stages of PI embryo. While blastocysts exhibited an increased influx of taurine in hypotonic media, it was smaller in magnitude than that seen in one-cell embryos, not observed until osmolarity was decreased to levels significantly below those eliciting a response in one-cell embryos, and not inhibitable by a Cl− channel blocker effective in one-cell embryos (DIDS), all of which might indicate a mechanism distinct from that of one-cell embryos (Van Winkle et al., 1994). Radiolabeled compounds are clearly useful in directly demonstrating an increased permeability on swelling, but this method has drawbacks, principally the high expense of the radiolabeled compounds. In addition, while much of the amino acid transport activity in embryos can be eliminated by performing experiments in Na+-depleted media, there are several Na+-independent transporters in embryos (Van Winkle, 1988) that will interfere with measurements involving amino acids that are their substrates. Another technique for assessing the permeability of compounds through the swelling-activated channel (Fig. 15), which avoids these problems, has been developed by Pasantes-Morales et al. (1994). Here, a large external concentration of a putative osmolyte is present during RVD. Because permeation through the swelling-activated pathway is passive, the flux of any permeable molecule will be a function of its concentration gradient and permeability. Therefore, if there is a high enough concentration of a permeable compound outside the cell, a large influx of this osmotically active solute will occur, and RVD will be opposed by the influx of osmolyte. In this way, compounds with a large permeability in swelled cells can be identified by the inhibitory effect of high external concentrations of such compounds on RVD. This technique allows a large number of compounds to be screened for a swelling-activated permeability (Pasantes-Morales et al., 1994). It should be noted, however, that while any endogenous osmolyte used by the cell should be found by this technique to have a high permeability in swelled cells, the converse, that all compounds exhibiting a high permeability are endogenous osmolytes, is not true. Thus, permeable compounds identified in this way are potential osmolytes, but they must also be shown to be normally present in the cell to be identified as an osmolyte used by that cell type. A subset of compounds tested in this way inhibited experimentally induced RVD in mouse one-cell embryos, indicating significant swelling-activated permeability (Fig. 16). Compounds that had been directly shown by other methods to be highly permeable through a swelling-activated pathway in embryos, such as glycine,

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Figure 15 Method for demonstrating swelling-activated permeability increase. A cell (left) is hypotonically swelled. Normally, swelling induces a greatly increased permeability to intracellular osmolytes already present in the cell (O), which exit the cell and allow volume to recover (RVD). If a large external concentration of a substance (S) that is also permeable only to swelled cells is present, however, it would enter the cell, on swelling, down its concentration gradient (top left). The introduction of a large amount of the externally present substance (S) opposes RVD, because its influx increases intracellular osmolarity even as the efflux of endogenous osmolytes attempts to allow RVD (top right). In contrast, if the substance S does not have a significantly increased permeability in the swelled cell, it will not enter and will not affect RVD, which will then occur normally (bottom left and right).

aspartate, β-alanine, and taurine (Van Winkle et al., 1994; Dumoulin et al., 1997; Dawson et al., 1998), were also identified by this method as highly permeable (i.e., blocking RVD). Conversely, those known not to permeate, such as lysine and raffinose, and positively charged amino acids, which should not permeate the VSOAC anion channel, had no effect on RVD. Interestingly, many of the amino acids that are beneficial to early PI embryo culture (see Table I and above) were also found by this method to be permeable through the pathway opened by hypotonic swelling (Fig. 16). It is tempting to speculate that inclusion of these amino acids in the medium is beneficial in part because they can be lost through the swelling-activated pathway, and thus are more easily depleted than amino acids that are not permeable. It should also be noted that pyruvate, a major metabolic substrate of cleavage-stage embryos, can apparently be lost via this pathway (Fig. 16).

V. Organic Osmolytes and Osmolarity in Vivo A. Organic Osmolytes within in Vivo-Derived Preimplantation Embryos The amino acids reported to be present at highest concentration within mouse eggs or embryos freshly removed from the oviduct are glycine and taurine, with significant amounts of alanine, glutamine, and glutamic and aspartic acids present as well.

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Figure 16 Swelling-activated permeability to a number of compounds assessed by their effect on recovery from hypotonic swelling (RVD) in mouse one-cell embryos. The inset shows RVDs in two individual mouse zygotes. The trace marked “raffinose” is a normal RVD that occurred in 150 mOsM medium with 50 mM raffinose replacing a portion of NaCl. This RVD is indistinguishable from that which occurs in normal medium lacking raffinose (see Fig. 11) (S´eguin and Baltz, 1997). The trace marked “glycine” shows the complete inhibition of RVD when 50 mM glycine is instead present in the medium, which is taken as an indication of a high swelling-activated permeability to glycine (see text and Fig. 15). The main part of the figure shows mean ( ± SEM) rates of RVD in the presence of a large number of different compounds whose identities are indicated at bottom. Each compound was added to a final concentration of 50 mM, replacing a portion of NaCl (except for aspartic and glutamic acids, which were added as 25 mM of the K+ or Na+ salts to maintain iso-osmolarity; no difference in effect was seen between salts). The rate of RVD was calculated from the slope of the recovery after peak swelling was achieved (determined by linear regression), expressed in arbitrary diameter units per minute, with the larger negative rates indicating robust RVD, and a rate of zero indicating complete inhibition. The results have been arbitrarily arranged in order of effectiveness of the compound, so that those judged most permeable to swelled one-cell embryos appear at the right, while those that did not affect RVD are at the left. When significance was tested by ANOVA followed by Dunnett’s multiple comparisons test, all compounds from valine to glycine, inclusive (to the right of the vertical dashed line), were significantly different from the negative control, raffinose ( p < 0.01), indicating some swelling-activated permeability to each of these compounds in one-cell mouse embryos. This work was done in the author’s laboratory (D. G. S´eguin and J. M. Baltz, unpublished data, 1998) .

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The glycine content of mouse ovulated eggs and two-cell embryos corresponds to intracellular concentrations of about 20 and 50 mM, respectively (Table II). Taurine is present in ovulated mouse eggs at about 30 mM, apparently decreasing to about 5 mM by the two-cell stage (Table II). Glutamine is not present at high intracellular concentrations in mouse eggs or two-cell embryos, but reaches about 10 mM at the four- to eight-cell stage and thus may become osmotically significant at that point (Table II; Van Winkle and Campione, 1996). Alanine is also present at significant levels, appearing highest at the two-cell stage (Table II). While aspartic and especially glutamic acids are also present at fairly high levels, these are charged molecules and thus probably do not have a role as organic osmolytes. Thus, the amino acids that mouse embryos accumulate in vivo and that could function as organic osmolytes appear to be mainly glycine, taurine, glutamine, and alanine, which is consistent with the set of amino acids identified as osmoprotective in vitro. The sum of the intracellular concentrations of these four amino acids in eggs and embryos immediately after removal from the oviduct is roughly 55–65 mM in unfertilized eggs and two-cell embryos (with the majority being taurine in eggs and glycine in two-cell embryos), and about 35 mM at the four- to eight-cell stages (Table II). These concentrations, if they represent free cytoplasmic amino acids, are certainly sufficient to explain the osmoprotective effect of these osmolytes. In addition, the concentrations of several amino acids that act as osmolytes in embryos, including proline, β-alanine, and hypotaurine, have not been determined in embryos and may be present at osmotically significant levels intracellularly. Rabbit eggs, in contrast, contain a total concentration of only about 12 mM of these amino acids (Table II; Miller and Schultz, 1987). This observation makes it tempting to speculate that the osmoprotective effect of myo-inositol in rabbit but not mouse embryos (above) is due to the latter utilizing high levels of amino acids as osmolytes, while rabbit embryos also accumulate myo-inositol.

B. Organic Osmolytes in Oviductal Fluid The same amino acids present at high concentrations in eggs and embryos are also major components of oviductal fluid (Table III). Glycine is the major α-amino acid present in oviductal fluid of all species examined (Table III), with absolute concentrations in the millimolar range (Gu´erin et al., 1995). This would be sufficient for osmoprotection, because 0.5–1.0 mM external glycine was found to be maximally effective in mouse embryos (Figs. 5 and 9; Van Winkle et al., 1990a; Dawson and Baltz, 1997; Dawson et al., 1998). Several other α-amino acids with osmoprotective properties in embryos are also present, with alanine comprising the second most significant fraction after glycine in all species examined, and glutamine and proline each present in significant amounts (Table III). β-Amino acids are also present in mouse oviductal fluid, with taurine composing the major

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2. Osmoregulation in Embryos Table II Amino Acid Content of Eggs and Embryos

Mouse egga

Mouse two-cellb

Mouse fourto eight-cellb

Rabbit eggc

Taurine Alanine Glycine Glutamic acid Glutamine Aspartic acid Threonine Hypotaurine Proline Serine Valine Lysine Leucine Phenylalanine Tyrosine Methionine Isoleucine Arginine Histidine Asparagine Cystine Tryptophan β-Alanine

33d 3.0 20 4.5 0.5 2.3 0.6 NDe ND 0.9 0.4 0.4 0.3 0.1 0 0.1 0.1 0 0.1 ND ND ND ND

5.2 7.5 53 14 1.0 3.5 ND ND ND 1.2 ND ND ND ND ND ND ND ND ND ND ND ND ND

7.4 3.8 16 8.5 9.5 1.3 ND ND ND 1.1 ND ND ND ND ND ND ND ND ND ND ND ND ND

7.6 2.1 2.6 2.7 ND 3.0 0.2 ND ND 0.4 0.3 0.1 0.4 0.1 0.1 0.1 0.2 0.2 0.2 ND ND ND ND

Gly + Tau + Ala + Gln

57

67

37

12

Amino acid

a

Schultz et al. (1981). Van Winkle and Dickinson (1995). c Miller and Schultz (1987). d Concentrations are in millimolar units. For mouse, concentrations were calculated by assuming the egg has a volume of 200 pl and the embryo a volume of 190 pl (the latter is the volume of a one-cell embryo in the oviduct; Collins and Baltz, 1999). This is similar to the calculation done by Van Winkle and Campione (1996). e ND, Not determined. b

fraction of total amino acids (both α and β combined) and present in millimolar amounts in mouse (Gu´erin et al., 1995). However, it is a relatively minor component of oviductal amino acids in other species, with the possible exception of pig (Table III;Gu´erin et al., 1995). In addition, the β-amino acids hypotaurine and β-alanine are present in oviductal fluids (Gu´erin et al., 1995; Gu´erin and Menezo, 1995). Thus, amino acids that are proved to be osmoprotective in embryos and to be accumulated by embryos to high levels in vivo, such as glycine, taurine, glutamine, alanine, and proline, are also present in significant amounts in oviductal fluid and available to the egg and embryo in vivo.

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Table III Amino Acid Content of Oviductal Fluid of Several Species Mouse

Amino acid Taurine Alanine Glycine Glutamic acid Glutamine Aspartic acid Threonine Hypotaurine Proline Serine Valine Lysine Leucine Phenylalanine Tyrosine Methionine Isoleucine Arginine Histidine Asparagine Cyst(e)ine Tryptophan β-Alanineg

Rabbit

a

Cow

Sheep Pig (Gu´erin (Dumoulin (Miller and (Gu´erin (Elhassan (Gu´erin (Gu´erin (Gu´erin et al.b ) et al.e ) Schultzd,e ) et al.b ) et al.f ) et al.b ) et al.b ) et al.b ) 39 10 11 10 7 4 4 3 3 3 2 2 1 1 1 1