Current Topics in Developmental Biology, Volume 52

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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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

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 <1 <1 <1 <1 <1 <1 ND

59 5 10 9 5 4 1 ND <1 2 1 1 1 1 <1 <1 1 <1 <1 ND ND ND ND

6 11 33 8 ND 2 11 ND ND 5 3 3 2 1 1 1 1 2 1 ND ND ND ND

2 11 47 5 6 1 4 4 2 4 4 1 2 1 2 1 1 <1 2 1 <1 <1 ND

2 12 44 17 3 2 2 ND 2 2 2 1 2 1 1 <1 1 1 1 <1 4 1 <1

1 18 37 5 2 1 1 2 3 2 4 6 5 2 2 1 3 3 2 <1 <1 <1 ND

1 7 65 1 2 1 <1 8 1 1 2 3 2 1 1 <1 1 1 1 <1 <1 <1 ND

13 9 47 <1 2 <1 1 10 <1 3 2 4 2 1 1 <1 1 3 1 1 <1 <1 ND

a

Values are given as a percentage of total amino acids, and were obtained from published work. When amino acid contents were given as absolute concentrations in the original reference, they have been converted here to percentage of total amino acids, rounded to the nearest whole number. Amino acids present at 0.5% or less are shown as <1. b Gu´erin et al. (1995). c Dumoulin et al. (1992a). d Miller and Schultz (1987). e Percentages were calculated assuming that glutamine, hypotaurine, and proline (not measured) constitute 12% of the total as in Gu´erin et al. (1995). f Elhassan et al. (1999b). g β-alanine was measured only in cow, where it composed 0.4% of the total.

Whether betaine, an excellent osmolyte in embryos, is present in oviductal fluid or within PI embryos in vivo has not been investigated. It will be interesting to determine whether betaine is present in oviductal fluid, and whether it can be transported into embryos and accumulated to high levels. As mentioned above, the kidney synthesizes and uses betaine as a major organic osmolyte. Because the oviduct and kidney share similar embryological origins (Byskov and

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Høyer, 1988), it is possible that the cells of the oviduct also have this capacity, and can provide this osmolyte to the PI embryo.

C. Oviductal Fluid Osmolarity Organic osmolytes would be required by embryos in vivo only if the osmolarity of oviductal fluid is high enough. For many years, the only indication of mouse oviductal fluid osmolarity was indirect, obtained by adding up the concentrations of Na+, Cl−, K+, Ca2+, and Mg2+ as measured by electron probe X-ray microanalysis (Borland et al., 1977). Van Winkle et al. (1990a) estimated oviductal fluid osmolarity from total ion concentrations by assuming that these would exist largely as osmotically active ions, yielding a value of 360 mOsM (Van Winkle et al., 1990a). However, salts at physiological concentrations, such as in oviductal fluid, are incompletely dissociated, and correcting for this yields an actual osmolarity closer to 345 mOsM. Several direct measurements of the osmolarity of oviductal fluid in species other than mouse have been reported. Olds and VanDemark (1957) measured the osmolarity of fluid derived by passing bovine oviducts through a clothes ringer, and found an osmolarity of about 350 mOsM, while more recent measurements of the osmolarity of small samples of neat bovine oviductal fluid yielded values of about 330 mOsM (luteal) or 295 mOsM (nonluteal; Killian et al., 1989) and 370 mOsM (Elhassan et al., 1999a). The osmolarity of equine oviductal fluid was found to be about 280 mOsM during estrus (Engle and Foley, 1975). Waring (1976) measured the osmolarity in small samples of rat oviductal fluid obtained by cannulation, and found an osmolarity of about 290 mOsM. It is not clear why the measured osmolarities vary so widely, even within species (e.g., bovine). However, the measured osmolarity of oviductal fluid during the time embryos are present would appear to be 290 mOsM or greater for most species. Another approach is to determine the tonicity of oviductal fluid with respect to the embryos themselves, rather than measuring osmolarity. This has been attempted by monitoring the change in size of one-cell and two-cell embryos immediately after their removal from the oviduct into media of various osmolarities, in effect using the embryos themselves as osmometers (Collins and Baltz, 1999). By this method, the tonicity of mouse oviductal fluid for both one-celland two-cell-stage embryos was estimated to be about 300 mOsM. In addition, measurement of the diameters of one-cell embryos in situ in undiluted oviductal fluid indicated that their sizes corresponded to the size of embryos in 300 mOsM medium (Collins and Baltz, 1999). A tonicity of 300 mOsM is high enough to require the use of organic osmolytes for optimal development of PI embryos in vivo, and thus accounts for high intracellular concentrations of osmolytes such as glycine, taurine, alanine, and glutamine in embryos freshly removed from the oviduct.

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VI. Discussion and Summary Various organic osmolytes have been shown conclusively to be beneficial to early embryo development in vitro, and most importantly to protect their development against the deleterious effects of increased osmolarity. At least one—glycine—has been shown to be accumulated to a higher level in early PI embryos when osmolarity is increased, and to support cell volume maintenance at increased tonicities. It therefore seems safe to conclude that the ability to use organic osmolytes is a feature of the physiology of early mammalian embryos. What role do inorganic ions play? Although active inorganic ion transport may play a part in acute volume regulation in embryos as in other cells, this has not yet been shown. Instead, the major role intracellular ions are currently known to play is a deleterious one, in which a level of intracellular ions sufficient to balance even a moderate external tonicity of about 300 mOsM severely decreases development of one-cell embryos in the absence of organic osmolytes. Several lines of evidence indicate that intracellular ionic strength is maintained at a low level in early PI embryos, with organic osmolytes providing the needed additional osmotic support, to avoid the deleterious effects of high ionic strength. Thus, it is proposed here that early mammalian embryos, especially during the earliest cleavage stages, ideally maintain their total intracellular ionic strength at a level approximately equivalent to that needed to balance an external tonicity of about 250–270 mOsM, which is approximately the optimal tonicity determined in vitro in the absence of organic osmolytes. Under conditions like those experienced by the embryo in vivo, where tonicity appears to be 300 mOsM or greater, the balance of intracellular osmolarity would normally be provided by organic osmolytes. Furthermore, judging from the presence of appropriate osmolytes in the oviduct, and the large intracellular concentration of these same osmolytes in eggs and embryos when they are freshly removed from the oviduct, the use of organic osmolytes would appear to be a normal property of embryos rather than an artifact of in vitro culture or a response to stress. Volume and osmotic regulation in early PI embryos thus seems to consist primarily of adjustments in the intracellular content of organic osmolytes, and three proposals can be advanced on the basis of the work described above. First, glycine, alanine, glutamine, possibly proline, and β-amino acids such as taurine and possibly hypotaurine and β-alanine are the major osmolytes normally used by one-cell- or two-cell-stage embryos (although nonamino acid osmolytes such as betaine, if present, and myo-inositol in the rabbit, may also be found to play a role). Second, these osmolytes are taken up from the external environment (i.e., the oviduct) via transporters of the Na+- and Cl−-dependent neurotransmitter transporter family— GLY and system β. Third, osmolytes are released from early PI embryos via a swelling-activated anion/organic osmolyte channel that is indistinguishable from the VSOAC channel in other cells. A schematic model incorporating these proposals is shown in Fig. 17A.

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Figure 17 Model of osmotic and volume regulation in preimplantation embryos. (A) One-cell stage and early cleavage stage embryos. Early in PI development, embryos are proposed to regulate cell volume and intracellular osmolarity at least in part by accumulating organic osmolytes via GLY and system β, while osmolyte efflux is mediated by a VSOAC channel activated by increased cell volume. The osmoprotective GLY substrates available in the embryo environment include glycine and glutamine, both of which are accumulated by embryos. In addition, others such as proline and betaine may be accumulated via GLY as well (although a betaine-specific transporter has not been ruled out). System β substrates accumulated by embryos include taurine and alanine, and others such as β-alanine and hypotaurine may also be accumulated. (B) Changing activities of osmoregulatory mechanisms over PI development. The mechanisms of osmoregulation are proposed to change over the course of PI development. After the cleavage stages, GLY declines in activity, and the ability of the embryo to accumulate glycine in response to hypertonicity is lost. Preliminary data indicate that the VSOAC channel also declines after the early cleavage stages. In contrast, system β increases in activity after the cleavage stages. Thus, the only known osmoregulatory mechanism active after the cleavage stages is system β.

This is in part a novel scheme for organic osmolyte regulation. While glycine is a common osmolyte in mammalian cells, the usual route for its uptake into osmotically challenged cells is via system A. Surprisingly, the PI embryo appears to lack any system A activity prior to the blastocyst stage and the only known route for glycine accumulation in early cleavage-stage embryos is GLY, whose major role in embryos thus appears to be to provide glycine as an osmolyte. This is a previously undescribed role, and the widely expressed GLY system does not appear to have been proposed to have a role in osmotic regulation in other cells. If this role in embryos is confirmed, it would be tempting to speculate that GLY has a similar role in other cells as well. Thus, it could be that the osmoregulatory function of system A is to respond to large increases in osmolarity, after a delay to accommodate its synthesis, and permit a greatly increased rate of amino acid osmolyte accumulation in response to sustained osmotic perturbations, while GLY mediates the normal “housekeeping” role of accumulating a level of intracellular glycine in cells that allows continuous control over cell volume and intracellular osmolarity independent of intracellular ionic strength. However, there is still considerable work to be done

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before GLY can be firmly established as an osmolyte transporter even in embryos. At minimum, it remains to be demonstrated that glycine cannot be accumulated as an osmolyte in embryos lacking GLY activity (e.g., via “knockout” of the glytl gene), or where its activity is blocked by a specific inhibitor.6 How are the intracellular concentrations of organic osmolytes regulated in embryos? Thus far, there is evidence in early PI embryos for quick stimulation of system β by hypertonicity, but only minimal stimulation of GLY over the relevant range. Such stimulation would serve to increase the rate of osmolyte accumulation but would not on its own regulate the final intracellular concentration, which is the important parameter for governing cell volume and intracellular ionic strength. It is indeed not clear whether GLY and system β regulate the intracellular concentration of their substrates, or whether they simply mediate concentrative uptake of these substrates without being subject to feedback control. If these transport systems cannot themselves regulate intracellular osmolyte concentrations, then the mechanism whereby osmolyte levels in embryos are determined could instead depend on the swelling-activated VSOAC-like channel. In this scenario, continued accumulation of organic osmolytes would swell the cell, activating the channel whenever the volume exceeded a threshold. This release of osmolytes via the channel would thus regulate intracellular osmolyte levels and maintain volume near the threshold. In any case, further work is clearly needed to elucidate the regulation of osmolytes in embryos. Another area of active future investigation is likely to be the developmental regulation of osmoregulatory mechanisms in PI embryos. While eggs, one-cell embryos, and early cleavage-stage embryos clearly possess transport mechanisms that mediate the uptake of organic osmolytes and a swelling-activated channel that mediates osmolyte release, at least two of these mechanisms appear to be diminished or lost during PI development (Fig. 17B). GLY activity and intracellular glycine content decreases by the four- to eight-cell stage, and GLY activity is absent from blastocysts. This parallels the loss of the embryos’ ability to accumulate increased amounts of glycine in response to hypertonicity, implying that glycine can no longer be used by PI embryos as a major osmolyte after the cleavage stages. Similarly, preliminary data indicate that swelling-activated channel activity decreases significantly by the four-cell stage. In contrast, system β activity and taurine content in embryos increase with PI development, perhaps indicating an increased role. Thus, the ability to use and regulate glycine and other GLY substrates as osmolytes would appear to be a property of only the earliest stages of PI embryo development, while other mechanisms may come into play as PI development proceeds. The complete story of cell volume and osmotic regulation during the earliest phase of embryogenesis remains to be investigated. 6

No inhibitor of GLY transport activity has been available until recently, when Bergeron et al. (1998) reported the use of a sarcosine derivative which blocks glycine uptake via glyt1-encoded GLY in neurons. It is not widely available as yet, but may prove useful if it is found to block GLY activity in nonneuronal cells.

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Acknowledgments Work from the author’s laboratory that has been cited here was supported by the Canadian Institutes of Health Research (formerly the Medical Research Council; Operating Grant MT12040) and the Whitaker Foundation (Biomedical Engineering Grant). Jay M. Baltz is a recipient of a Premier’s Research Excellence Award from the Government of Ontario. Work on human spare embryos (released for research with patient consent) was supported by the Division of Reproductive Medicine, Department of Obstetrics and Gynecology, University of Ottawa. I would like to thank Drs. John Biggers, George Seidel, Patrick Quinn, Michael Kane, Keith Betteridge, Robert Foote, and Richard Gordon for providing helpful information and input. I would especially like to thank Dr. Lon Van Winkle for extensive discussions and help, and Dr. Cathy Morris for stimulating discussions on the biophysics of cells. A substantial amount of previously unpublished work by Diane G. S´eguin, Kerri M. Dawson, and Mary-Anne Hammer is presented here, and their contributions are gratefully acknowledged.

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3 Cell–Cell Interactions in Vascular Development Diane C. Darland 1 and Patricia A. D’Amore 1,2,∗

1

Schepens Eye Research Institute Department of Ophthalmology Harvard Medical School Boston, Massachusetts 02114 2

Department of Pathology Harvard Medical School Boston, Massachusetts 02114

I. Introduction II. The Cardiovascular System A. Cardiac Development B. Early Development of the Vasculature C. Vascular Tree Formation D. Vessel Maturation III. Differences in Vascular Beds A. Variations in Endothelial Cell Gene Expression B. Endothelial Cells of Barrier Vessels C. Arterial versus Venous Endothelial Cells IV. Parallels between Angiogenesis in Development and Pathology A. Tumor Angiogenesis B. Ocular Angiogenesis V. New Directions A. Endothelial Cells as Mural Cell Precursors B. Cell–Cell Communication and Gap Junctions C. Neuronal–Vascular Interactions D. Vasculogenesis in the Adult VI. Summary References

I. Introduction The literal meaning of “vascular development” derives from two separate definitions: “of or relating to a channel for the conveyance of a body fluid (blood) or to a system of such channels” and “the act, process, or result of making visible ∗ To whom correspondence should be addressed. E-mail: [email protected]. 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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or manifest” (Merriam-Webster, 1987). The combination of these two ideas leads to a whole that is greater than the sum of its parts. A vast array of factors and mechanisms must work in concert to form a system as necessary and complex as the vasculature. The circulatory system is the earliest functioning organ system of the embryo and is absolutely required for survival of the embryo. It has evolved to service the function of gas exchange, nutrient delivery, and waste product removal. The larger vessels function predominantly as conduits to facilitate fluid transport, whereas the smaller vessels function in diffusion (Nguyen and D’Amore, 2000). A review of targeted gene disruptions that lead to cardiovascular abnormalities in mice yields an ever-increasing list (reviewed in Bazzoni et al., 1999; Gale and Yancopoulos, 1999; Hynes, 1996), and examination of this list leads one to the following ideas. First, many factors are necessary, but not likely sufficient, for the formation of the vascular system. Second, an abnormal vascular system becomes a critical developmental limitation beyond which an embryo can rarely progress to full maturation. Establishment of a normal vascular system requires a myriad of cell–cell interactions that occur in a precise temporal and spatial sequence. An examination of the nature of these cell–cell interactions reveals the impact of the microenvironment on the course and extent of cellular differentiation. This review emphasizes specification as influenced by cellular context and environment, as opposed to differentiation directed by the inherent potential of a particular cell type. Cells utilize a variety of mechanisms to communicate. In this review we focus on those areas where there is a body of literature to support a function in vascular development in vivo. This list includes, but is not restricted to, soluble growth factors that act in an autocrine or paracrine fashion, cell adhesion molecules that act directly through cell contact, and extracellular matrix (ECM) interactions that function via cell adhesion and migration.

II. The Cardiovascular System A. Cardiac Development If the vascular system is the first functional unit, the heart by association is the first functional organ. The heart arises from bilateral clusters of precardiac cells derived from presumptive mesodermal cells that are specified at the time of gastrulation (DeHaan, 1959; Jacobson, 1961). A bilateral origin for the heart has been described for all animal species examined, regardless of whether the early embryo derives from a blastomere or blastodisc (Gilbert, 1997). There is mounting evidence to suggest that cell–cell interactions play an integral role in specification of the cardiac precursors and in their eventual differentiation into heart primordia. Considerable evidence comes from the study of avian cardiac development, which indicates that precardiac tissue is specified in presumptive anterior mesoderm (Garc´ıa-Martinez and Schoenwolf, 1993; Schultheiss et al.,

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1995). A strong candidate for the induction of precardiac tissue is the underlying endoderm. Explant and recombination experiments in Xenopus (Nascone and Mercola, 1995) and chick/quail chimera transplantation studies (Schultheiss et al., 1995; Sugi and Lough, 1994) have shown that anterior endoderm, but not posterior endoderm, can induce precardiac tissue in the anterior mesoderm. Interestingly, when posterior primitive streak tissue (presumptive posterior mesoderm) in the chick is in contact with anterior endoderm, it also becomes precardiac tissue (Schultheiss et al., 1995). The caudal tissue is never specified for the heart lineage in vivo, indicating that local environmental signals and cell–cell interactions have a significant impact on cell fate determination in heart development. The endoderm-derived factor(s) that specifies heart tissue is as yet unidentified. However, if precardiac tissue and anterior endoderm from chick are combined, only transplants with direct contact give rise to heart tissue (Sugi and Lough, 1994). These results indicate that the endoderm-derived inducing factor has a short diffusion distance or that it is dependent on intercellular contact. One hypothesis to explain the inductive potential of the anterior endoderm is that precardiac mesoderm migrates caudally until it contacts anterior endoderm (Linask and Lash, 1986). There is a corresponding caudal to rostral increase in the ECM protein, fibronectin, along this migratory path. If the precardiac cells and the associated endoderm tissue are rotated after establishment of the fibronectin gradient, anterior migration does not occur. It is unclear which cells deposit the fibronectin gradient, but the importance of the gradient in guiding precardiac tissue to the correct geographic location at the appropriate time is clear. After specification by the endoderm, the precardiac tissue differentiates to form a double-walled tube on either side of the embryo. The exterior cell layer, or epimyocardium, gives rise to the heart muscles and the interior cell layer or endocardium gives rise to the inner lining of the heart, again demonstrating how cell–cell interactions specify differentiation. As the lateral portions of the embryo migrate ventrally and fuse at the midline, the bilateral tubes fuse and give rise, on the right side, to the ventricle leading to the pulmonary trunk and, on the left side, to the aorta. If a barrier is placed ventrally to prevent fusion in a developing chick embryo, then two separate heart structures form on either side of the body, resulting in cardia bifida (DeHaan, 1959). Another aspect of heart formation that is likely to be mediated by cell–cell interactions is the formation of the valves and septa. A subset of the endothelial cells (ECs) that line the interior of the heart at the atrioventricular cushions undergo a transition to mesenchymal tissue, which subsequently gives rise to the valves and septa. It is likely that this transition is due, at least in part, to ECM molecules, or to growth factors bound to the matrix secreted by the underlying endoderm (Krug et al., 1987). The transforming growth factor β (TGF-β) family of proteins, which bind to ECM and are activated in the extracellular environment, are likely candidates to be involved in this process. In contrast to control-treated explants, atrioventricular cushion explants treated with antisense oligonucleotides to TGF-β3 do not undergo the epithelial-to-mesenchymal transition (Nakajima

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et al., 1998; Potts et al., 1991). In a chick atrioventricular explant assay, neutralizing antibodies to TGF-β2 and TGF-β3 were used to determine that TGF-β2 was required for the separation of ECs prior to the transition to mesenchyme, and that TGF-β3 likely mediated the mesenchymal transition itself (Boyer et al., 1999). Bone morphogenetic protein 2 (BMP2), another member of the TGF-β superfamily, may also promote the transition, possibly in conjunction with TGF-β3 (Yamagishi et al., 1999; Yamada et al., 1999). Collectively, these results indicate a central role for TGF-β superfamily members in the cell–cell interactions that modulate septation of the heart. Two additional factors, angiopoietin (ang) 1 and 2, have been implicated in heart development (Gale and Yancopoulos, 1999). By in situ hybridization, mRNA for ang 1 has been shown to be localized predominantly to the developing heart myocardium and surrounding endocardium, mural cells of most blood vessels, and hematopoietic clusters in the liver (Davis et al., 1996; Maisonpierre et al., 1997). Ang 2 is expressed by the mesenchyme of developing vessels, but is also found in the smooth muscle cells (SMCs) in the dorsal aorta and in the major aortic branches (Maisonpierre et al., 1997). These expression patterns shift in the adult, where ang 1 has a widespread expression pattern and ang 2 expression is restricted to the reproductive organs that undergo angiogenesis (Maisonpierre et al., 1997). The receptor for these ligands, the Tie 2 (Tek) tyrosine kinase, is expressed on ECs and on cells of the hematopoietic lineage (Dumont et al., 1992; Korhonen et al., 1994). Ang 1 and ang 2 can both bind to the Tie 2 receptor, yet only ang 1 can activate the receptor. An antagonistic relationship between ang 1 and ang 2 has been proposed so that the outcome of their actions would be dependent on their relative expression levels, with ang 1 stimulating receptor phosphorylation and ang 2 blocking phosphorylation (Davis and Yancopoulos, 1999). Transgenic mice overexpressing ang 2 and knockout mice lacking ang 1 and Tie 2, all of which are embryonic lethal, share many phenotypic abnormalities, including abnormal heart development. The ang 1-deficient mice display defective cardiac trabeculation (Suri et al., 1996). In addition, the ECs of the endocardium do not adhere well to the underlying matrix and appear loosely attached and rounded. Similar heart defects are observed in the ang 2-overexpressing mice (Maisonpierre et al., 1997) and the Tie 2 knockout mice (Dumont et al., 1992). A similar phenotype has been observed in mice with targeted disruption of the genes for neuregulin and neuregulin receptor (Kramer et al., 1996; Lee et al., 1995), which have been shown to be involved in growth and differentiation of a number of cell types including glial, epithelial, and striated muscle (Marchionni, 1995). An intriguing hypothesis has been proposed suggesting that ang 1, produced in the myocardium, sends differentiation signals to the endocardium via the Tie-2 receptor. Conversely, the endocardium produces neuregulin that signals through its receptor on the myocardium. In this way, cell–cell communication between the adjacent layers may lead to functional septation in the heart (Gale and Yancopoulos, 1999). It has been suggested that the transcription factor MEF2C (myocyte enhancer factor 2C) is involved in coordinating expression of some of the factors involved

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in normal cardiogenesis (Bi et al., 1999). The MEF2C null mouse displays a phenotype that is similar to the ang 2 null mouse, having enlarged cardiac and cranial vessels, although the dorsal aorta appears normal in the absence of MEF2C (Lin et al., 1997). The MEF2C–/– embryos had increased mRNA for ang 1, Tie 2, and vascular endothelial growth factor (VEGF), relative to wild-type embryos (Bi et al., 1999). Interestingly, analysis of cardiac tissue revealed reduced expression of mRNA [by reverse transcriptase-polymerase chain reaction (RT-PCR) analysis] for ang 1 and VEGF (Bi et al., 1999) as well as the dHAND (deciduum, heart, autonomic nervous system, and neural crest derivatives) transcription factor required for cardiac morphogenesis (Lin et al., 1997; Srivastava et al., 1995). Taken together, these results suggest that the vascular phenotype in the mice lacking MEF2C is due to abnormal regulation of the genetic program for angiogenesis, downstream effectors of which include ang 1 and VEGF. Cell–cell communication at several levels is required for normal heart formation, and a number of factors have been proposed to be involved at various steps in this process. The role of neural crest in cardiovascular patterning, cardiac looping, and chamber formation is discussed elsewhere (Kirby and Waldo, 1995). B. Early Development of the Vasculature 1. Endothelial Cell Specification The earliest identifiable ECs can be pinpointed to the yolk sac of the developing embryo. The yolk sac is part of the extraembryonic membranes and is surrounded by endoderm, the innermost of the three embryonic layers in a developing embryo. Early studies of “wandering mesenchymal” cells in teleost embryos showed that the mesenchymal cells migrate throughout the yolk sac and give rise to vascular endothelium and blood cells (Noden, 1989; Stockard, 1915). The earliest structures formed, blood islands, line the yolk sac and arise from groups of cells called angiogenic clusters. The angiogenic clusters of the yolk sac are unique in that they consist of hemangioblasts that are common precursors of ECs and hematopoietic cells (Risau, 1995). As differentiation of the clusters proceeds, the hemangioblasts differentiate to ECs on the outside of the cluster and hematopoietic precursors on the inside. The hemangioblasts are an early source of blood that is later derived from mesodermal cells located around the aorta. In postnatal life the aorta-derived blood stem cells take up residence in the bone marrow and are the principal source of blood precursors in the adult. A mutation in zebrafish called cloche negatively affects both endothelial and hematopoietic cells, resulting in an absence of endocardium, head and trunk ECs and most blood cells, indicating that the mutation affects a precursor population for both cell lineages (Fouquet et al., 1997; Liao et al., 1998; Stainier et al., 1995). Transplantation of wild-type cells into cloche mutants allowed for normal formation of endocardium, whereas transplantation of cloche mutant cells into wild-type hosts prevented normal cardiac development (Stainier et al., 1995).

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Analysis of hematopoiesis in similar transplant studies indicated that cloche is cell-autonomous for differentiation of red blood cells, but not for their survival or proliferation, which are affected by microenvironmental influences (Parker and Stainier, 1999). Differentiation of ECs and hematopoietic cells from a common hemangioblast precursor in zebrafish is an example of how cell–cell interactions can alter cell fate. Specifically, the positional orientation of the cells appears to determine lineage specification. Lineage may be an inherent property of prespecified cell types, or a combination of both processes may be involved (Mills et al., 1999). It is important to note that the differentiation of ECs in the extraembryonic membranes is different from intraembryonic ECs differentiation (Pardanaud et al., 1989). Where vasculogenesis occurs within the embryo proper, the ECs are specified from precursor cells called angioblasts. Within the embryo, angioblasts are not found in association with hematopoietic precursors. Studies using chick/quail chimeric transplants demonstrated two potential sources of ECs during embryo development (Pardanaud et al., 1996). ECs derived from the paraxial mesoderm colonize the cephalic (head) vessels and the somitic (trunk) vessels, including vessels of the kidney, limbs, body wall, and some portions of the dorsal aorta (Fig. 1). The ECs that are derived from the splanchnopleural mesoderm give rise to the vessels of the visceral organs, gut, and the floor of the dorsal aorta. These developmental derivations are thought to be roughly similar for ECs of other animal species, including humans (reviewed in Noden, 1989; Patterson et al., 1998; Risau, 1995). Analysis of vasculogenesis in mouse [embryonic day (E) 6.5–E9.5]

Figure 1 Embryonic derivation of angioblasts. The schematic demonstrates the three primary layers of developing embryo, the ectoderm, mesoderm, and endoderm, before neurulation. The yolk sac lies beneath the endoderm and is encircled by it. Avian and mammalian embryos are flattened discs on the yolk sac, whereas amphibian embryos completely encircle the yolk sac. In the chick, angioblasts arise from the paraxial mesoderm; the cephalic portion contributes to the vessels of the head and the somitic portion contributes to the trunk vessels and the dorsal aorta. The splanchnopleural mesoderm, located laterally in the embryo, contributes angioblasts for the vessels of the visceral organs and the ventral portion of the aorta. NC, Notochord.

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3. Cell–Cell Interactions in Vascular Development Table I Relative Expression of Endothelial Cell–Associated Proteins during Mouse Vasculogenesisa ECs

TAL-1 VEGFR-2 PECAM CAM34 VE-cadherin Tie 2 a

Angioblasts

Primitive vessels

Mature vessels

+ +

+ + + + +/− +

− + + + + +

− − −

Modified with permission (Drake and Fleming, 2000).

has revealed a temporal and spatial expression of genes associated with specific endothelial lineages (Drake and Fleming, 2000). The transcription factor TAL1/SCL (T cell acute leukemia 1/Sezary cell leukemia) is expressed in angioblasts prior to their expression of EC-specific proteins and is downregulated in ECs of mature vessels (Table I) (Drake and Fleming, 2000). TAL-1/SCL is also expressed in angioblasts of birds (Drake et al., 1997) and zebrafish (Liao et al., 1998), marking regions of early vasculogenesis (Table I). Basic fibroblast growth factor (FGF2) has been implicated in the specification of angioblasts (Risau, 1995). When quail blastodiscs are explanted and disrupted in culture, the mesoderm cells do not form angioblasts or express proteins associated with the EC lineage (Flamme and Risau, 1992) unless they are treated with FGF2, in which case blood island-like structures are formed (Flamme and Risau, 1992). FGF2 has been identified in the chick chorioallantoic membrane (CAM) during vasculogenesis and can promote vessel growth when exogenously added to the CAM during embryo development (Ribatti et al., 1995). Moreover, neutralizing antibodies to FGF2 inhibited vessel growth when applied locally, suggesting that FGF2 normally functions to promote vessel growth, possibly by inducing angioblast formation from mesoderm (Ribatti et al., 1995). Although not all of the factors responsible for specification of the blood islands and subsequent differentiation of ECs are known, there is evidence to suggest that VEGF is involved. VEGF functions by signaling through tyrosine kinase transmembrane receptors VEGFR-1, -2, and -3 (Klagsbrun and D’Amore, 1996; Neufeld et al., 1999). VEGFR-2 (Flk-1; KDR) is expressed in the yolk sac blood islands and is the earliest marker of EC specification identified to date (Dumont et al., 1995). Interestingly, the TAL-1/SCL-positive angioblasts also express VEGFR-2, whereas the hematopoietic precursors express only the TAL-1/SCL protein (Drake and Fleming, 2000). Mice that have been engineered to lack VEGFR-2 display abnormalities in yolk sac and blood island formation. In these mice, EC precursors do not form in the blood islands (Shalaby et al., 1995), indicating a requirement for signaling through the VEGFR-2 in EC specification.

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ECs that form in blood islands and within the embryo establish connections with one another and form primitive vessels. This type of de novo vessel formation is referred to as vasculogenesis. Vascularization via vasculogenesis is not uniform throughout the embryo, but does occur in the gut, lung, and aorta. ECs in the early vasculature of these organs and ECs that are specified later in development are recruited from the splanchnic or visceral mesoderm, a portion of the lateral plate mesoderm (Fig. 1). Although some embryonic vessels are formed by vasculogenesis, the majority of vessels in the developing embryo are formed through a process called angiogenesis, the formation of new vessels by sprouting from preexisting vessels. During angiogenesis, ECs are induced to proliferate and migrate to form new vessel branches. VEGF, as the name implies, was identified as a mitogen for ECs (Ferrara and Henzel, 1989; Leung et al., 1989; Plou¨et et al., 1989). The VEGF gene encodes at least three protein isoforms that arise from alternative splicing. Of these, the VEGF165 isoform (human) is reportedly the most potent mitogen for ECs (Keyt et al., 1996). VEGF is a likely candidate to mediate the early phase of EC proliferation that is common to both vasculogenesis and angiogenesis. Evidence to support this idea comes from observations of VEGFR-1-deficient mice that develop EC precursors, but have abnormal formation of the primitive vessels (Fong et al., 1995), likely because of excess production of EC precursors (Fong et al., 1999). Thus, the phenotypes of the VEGF receptor knockout mice, VEGFR-2 and -1, support the hypothesis that VEGF signaling is required for EC differentiation from precursors and for EC proliferation to form nascent vessel structures, respectively. 2. Mural Cell Specification Mural or “perivascular” cells were first identified as cells that are closely associated with, and external to, the ECs. Mural cell function and morphology vary depending on their location within the vascular tree (Rhodin, 1968). In large vessels, such as arteries, the mural cells are referred to as smooth muscle cells (SMCs) and form a multilayered sheath around the elastic artery wall. These large vessels are fed by a distinct vessel supply, the vaso vasorum, located within the medial layer of the vessel wall. In the arterioles and venules, the SMCs form a continuous sheet that can be single or multilayered. These vessels lack the thickened medial layer of the larger vessels, but have a basement membrane layer that lies between the ECs and the SMCs and is produced by contributions of the two cell types (reviewed in Ruoslahti and Engvall, 1997; Shepro and Morel, 1993; Steinberg, 1963). Acquisition of a basement membrane, including the protein laminin, is characteristic of maturing vessels (Risau and Lemmon, 1988). The mural cells that form a noncontinuous abluminal layer at the level of the capillary and postcapillary venule are referred to as pericytes (Doherty and Canfield, 1999; Sims, 1986). Pericytes send extensive processes along the length of and around the capillary tube. Although the pericytes

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and ECs are separated by a thin basement membrane, there are sites of direct contact between pericyte processes and the ECs (Fujiwara and Uehara, 1984; Zweifach, 1937) (Fig. 2; see color insert). Because most new vessel growth occurs as sprouts from the smaller vessels, we focus our attention on the pericytes. Microvascular mural cells were first identified by Rouget (1873) and were later referred to as Rouget cells. In 1923 Zimmerman coined the descriptive term “peri,” meaning near or around, and “cyte,” referring to cell; microvascular mural cells have subsequently been referred to as pericytes (Zimmerman, 1923). There has been much controversy over the years regarding the exact nature of the pericyte, particularly as it relates to the SMC. Pericytes are of mesenchymal origin and have been referred to as less differentiated SMCs (Rouget, 1873), although there is also variability in the degree of differentiation among SMCs identified in the large arteries (Gittenberger-de-Groot et al., 1999). Chimera studies with quail limb buds transplanted onto the CAM indicate that pericytes can be recruited from surrounding mesenchymal tissue and incorporated into new vessels (Stein et al., 1996). There is also evidence to suggest that pericytes can be derived from nearby vascular SMCs that “dedifferentiate,” proliferate, and then associate with new microvascular sprouts (Hellstr¨om et al., 1999; Lindahl et al., 1997; Nicosia and Villaschi, 1995; Skalli et al., 1989). It seems likely that both methods of pericyte derivation can occur during neovascularization. Whether tissue-specific differences determine the nature of the recruitment is unclear.

C. Vascular Tree Formation Because of the structural parallels observed between main trunk and limb branching in trees and the progressive reduction of vessel size from arteries through to arterioles and capillaries, the vascular system is often referred to as the “vascular tree.” This progression is reversed from capillaries to postcapillary venules and through the veins that then reconnect to complete the circuitry of the vasculature (reviewed in Rhodin, 1968; Sims, 1986). While vasculogenesis and angiogenesis initiate vessel formation via two distinct processes, the later steps of vessel assembly are shared by both processes (reviewed in Darland and D’Amore, 1999; Noden, 1989). Common building blocks in both vasculogenesis and angiogenesis are the EC and the mural cell. ECs are the foundation of the vasculature in that they form the vessel lumen and directly interface with the circulating cells and plasma-derived factors. The mural cell is located external to the EC and has been proposed to function in vessel stabilization and contraction, and normally does not have direct contact with the blood. These preliminary descriptions aside, it is important to note that the characteristics of these two cell types vary according to the structural and functional requirements of their location within the vascular system. In that

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regard, the combination of ECs and mural cells is definitely greater than the sum of the individual components. Other cell types that are involved in formation and maintenance of the vasculature include tissue parenchymal cells, platelets, and lymphatic cells. Furthermore, biomechanical forces resulting from fluid flow affect vascular development at both the cellular and system levels. Although we acknowledge the importance of these aspects of the vascular system, their consideration is beyond the scope of this discussion and the reader is referred to excellent reviews on these topics (Foldi, 1999; Gimbrone et al., 1997; Gimbrone, 1999; Kato, 2000; Wilting et al., 1999). 1. Primitive Network Formation One of the earliest steps in the formation of the vessel system is association of ECs to form cords that then elongate and connect with one another, generating a primitive network (reviewed in Drake et al., 1998). Initial network formation occurs predominantly via vasculogenesis. Expansion of the network is likely to involve angiogenesis from the earliest established vessels. Evidence from a number of studies indicates that VEGF is likely to be involved in vasculogenesis. In mice that have been engineered to lack the VEGFR-1 receptor there are abnormalities in the primitive vessel network (Fong et al., 1995). ECs are specified from angioblasts in these mice, indicating that the VEGFR-1 signaling is not required for EC lineage specification but is required for differentiation into a network. Further analysis of these mice has led to the suggestion that the absence of VEGFR-1 leads to an excess of EC precursors that would not arise in the presence of functional receptor signaling (Fong et al., 1999). Interestingly, a number of studies involving overexpression of VEGF have resulted in developmental abnormalities at early stages of vascular development. Overexpression of VEGF (VEGF122) by microinjection of plasmid DNA or mRNA in early-stage Xenopus embryos resulted in ectopic EC precursors in formation of the primitive network (Cleaver et al., 1997). When embryos were allowed to develop further, ectopic vessels formed (likely from the excess precursors) and often showed an enlarged, sinusoidal phenotype (Cleaver et al., 1997). Overexpression of VEGF (VEGF165) in mice resulted in an increased mass of disorganized vasculature (M¨uhlhauser et al., 1995). Morever, injection of VEGF into avian embryos also resulted in excess vasculature as well as sinusoid-like vessels that the authors suggest may be due to hyperfusion of primitive vessels (Drake and Little, 1995) (Fig. 2; see color plates). A similar disorganization in the vasculature has been observed in mice lacking vascular endothelial-cadherin (VE-cadherin), a protein that is localized to the adherens junctions of ECs (Lampugnani et al., 1992). The disruption in EC intercellular signaling resulted in disruption of EC organization in large vessels and in the yolk sac vasculature (Carmeliet et al., 1999; Gory-Faur´e et al., 1999), although the ECs still showed some adherence to each other (Gory-Faur´e et al., 1999). Another

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adhesion molecule that is a candidate for affecting EC contact during early plexus formation is platelet endothelial cell adhesion molecule (PECAM) (reviewed in Newman, 1999). Neutralizing antibodies to PECAM have been shown to inhibit angiogenesis capillary tube formation in vitro and cytokine-induced neovascularization in the rat cornea and in a mouse implant model (DeLisser et al., 1997). There is an absence of PECAM-1-expressing cells in the rostral portion of VEcadherin null mice, suggesting that there may be coordinate expression of these two adhesion molecules during development (Gory-Faur´e et al., 1999). Of note, mice generated that lack PECAM show no obvious abnormalities in either vasculogenesis or angiogenesis (Duncan et al., 1999), which the authors suggest is due to functional redundancy during vascular development. In fact, several adhesion molecules that have been identified as potential candidates for affecting vasculogenesis and angiogenesis have been eliminated in mice by genetic recombination strategies, with the resulting null mice having no vascular phenotype (Bazzoni et al., 1999; Hynes, 1996). The absence of overt vascular abnormalities does not imply that these molecules do not function in vascular development, only that there is the possibility of redundancy in the system. After formation of the initial network, the cord structures form lumens and become recognizable as primitive vessels. Interactions with the ECM are likely to be involved in lumen formation. Cells bind to ECM proteins in the environment via cell membrane-associated receptors called integrins. Integrins are heterodimers of α and β receptors that signal matrix-based information via intracellular second messengers (reviewed in Eliceiri et al., 1998; Huang and Ingber, 1999; Ruoslahti and Engvall, 1997). The ECM protein fibronectin has been shown to be present in the area of migrating ECs and immature capillaries (Risau et al., 1988). In addition, in an in vitro model of vessel assembly in three-dimensional collagen gels, lumen formation has been shown to be dependent on α 2β 1 (Davis and Camarillo, 1996). If antibodies to the β 1 integrin, a fibronectin receptor, are added to developing chick embryos during vasculogenesis, lumen formation is inhibited (Drake et al., 1992). A blocking antibody specific for a different integrin, α vβ 3, perturbed lumen formation as well and had the additional effect of inducing clusters of ECs throughout the primordial vascular network (Drake et al., 1992). Cluster formation reduced cell interactions with the ECM and prevented the formation of the normal array that results as the vessels coalesce (Drake et al., 1995). When CAMs were implanted with melanoma tumor fragments (human M21-L) and then treated with α vβ 3-blocking antibody or cyclic peptide antagonists to α vβ 3, the vessel investment normally observed in these tumors was prevented (Brooks et al., 1994b). The tumor vessels on the treated CAMs were disrupted and leaky (Brooks et al., 1994a,b). Interestingly, mice lacking all α v integrins have been generated and most vascular development proceeds normally in these mice, although there is extensive hemorrhage and only about 20% of the embryos survive to birth (Bader et al., 1998). The α vβ 3 integrin remains a good candidate for regulating at least some of the cell–ECM interactions that are involved in vascular development.

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2. Recruitment of Mural Cells Once a new vessel has formed, whether by vasculogenesis or angiogenesis, there is an important component of vessel maturation that involves recruitment of mural cells. Two basic mechanisms have been proposed for mural cell recruitment to sites of neovascularization (Hellstr¨om et al., 1999; Hirschi et al., 1998). The first involves recruitment of pericytes/SMCs from the mesenchymal tissue surrounding the new vessel. The second involves recruitment of SMCs from the vessel that is the source of the new sprout. The first mechanism is probably utilized in both angiogenesis and vasculogenesis, but the second is likely restricted to angiogenesis. The first phase of mural cell recruitment to a new vessel or sprout involves both migration of the responsive cell and, subsequently or concomitantly, induction of proliferation of the cell to produce sufficient progeny to outline the vessel. Evidence is accumulating to suggest a role for platelet-derived growth factor B (PDGF-B) in this early phase of recruitment. PDGF-B is produced by proliferating and/or activated ECs, and the mesenchymal cells express the cognate receptor (Hellstr¨om et al., 1999; Lindahl et al., 1997). PDGF-B promotes migration of mesenchymal cells (Hirschi et al., 1998) and induces their proliferation in vitro (Hirschi et al., 1999). Analysis of PDGF-B knockout mice has revealed that pericyte recruitment to new vessels is abnormal in these animals, particularly in the brain microvasculature (Hellstr¨om et al., 1999; Lindahl et al., 1997). Surprisingly, not all perivascular cells are affected by the absence of PDGF-B. This suggests that there are differences in the pericytes of different vascular beds that result in a differential sensitivity to the absence of PDGF-B. Alternatively, it could be that different molecular mechanisms mediate recruitment of pericytes in different tissues. In addition, SMCs in the limb arteries of the PDGF-B knockout mice display a reduced labeling index for bromodeoxyuridine (BrdU), relative to wild-type animals, indicating that fewer proliferating cells are recruited to the new vessel site and/or that there is reduced SMC proliferation (Hellstr¨om et al., 1999). As there is in vitro evidence to suggest that PDGF-B may be involved in both recruitment and proliferation of perivascular cells, it is possible that the knockout mice are affected at both levels. The absence of PDGF-B leads to embryonic lethality, with the animals dying of massive hematoma and cardiac abnormalities (Lev´een et al., 1994; Soriano, 1997), further emphasizing the critical requirement for PDGF-B in the formation of a functional vascular system. 3. Differentiation of Mural Cells Once the mural cell has become associated with the nascent vessel, it is induced to differentiate toward a pericyte or SMC lineage. There is considerable evidence to indicate that members of the TGF-β family of proteins are likely to be involved in this process (Hirschi et al., 1998; Pepper, 1997). When mesenchymal cells contact ECs in vitro, there is localized activation of latent TGF-β (Antonelli-Orlidge et al., 1989). Furthermore, when mesenchymal cells are cocultured with ECs or treated with TGF-β1, they express SMC markers, indicating differentiation toward an

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SMC lineage (Hirschi et al., 1998). The induction of mesenchymal cell differentiation to SMCs in coculture with ECs can be blocked with neutralizing antibodies against TGF-β (Hirschi et al., 1998). TGF-β1 has also been shown to direct neural crest cells toward a smooth muscle lineage (Shah et al., 1996), and TGF-β expression has been correlated with SMC differentiation in a rat model of arterial injury (Grainger et al., 1998). Mice lacking the gene for TGF-β1 exhibited abnormalities in vasculogenesis and haematopoiesis, although the majority of the defects were isolated to the extraembryonic membranes (Dickson et al., 1995). Specifically, EC progenitors were induced from yolk sac mesenchyme; however, they formed abnormal tube structures and the vessels were weak and leaky (Dickson et al., 1995). Similar developmental abnormalities were observed in the yolk sacs of mice lacking the TGF-β serine/threonine kinase receptor (type II) (Oshima et al., 1996). Although neither the ligand nor the receptor knockout mice were examined for presence or absence of mural cells, it is interesting to speculate whether the vascular phenotypes may have been due in part to abnormalities in or lack of differentiated mural cells. Further evidence to support this idea was obtained from mice engineered to lack endoglin, a protein that binds the TGF-β ligand receptor complex (Barbara et al., 1999). In the absence of endoglin, blood vessels are formed but are not invested with SMCs (Li et al., 1999). Thus, TGF-β appears to be required for several aspects of vessel formation. Pericytes are similar to SMCs in that they express an array of SMC-associated proteins, yet have characteristics that distinguish them from SMCs (reviewed in Hirschi and D’Amore, 1996; Owens, 1995; Sims, 1986). The primary distinguishing characteristics of the pericytes are their location within the vascular tree, the nature and extent of their cell processes and their density along the vessel (Fig. 3). At the level of the smallest vessels, where the greatest diffusion occurs, pericytes are sparse and there is controversy as to whether they are contractile in nature (Clark and Clark, 1925; Zweifach, 1934; and reviewed in Sims, 1986). There is some evidence to indicate that there are pericyte-like cells in the subendothelial layer of arterial vessel walls, which form an extensive network of process contact and may function similar to the pericytes in capillaries (Andreeva et al., 1998). The differences between pericytes and SMCs are likely due to slight variations in gene expression patterns regulated by the microenvironment (Miano et al., 1993; Nehls and Drenckhahn, 1991, 1993; Schlingemann et al., 1991; Tilton et al., 1979). Although there is no specific protein that distinguishes pericytes/SMCs from other cell types, there is a collection of proteins that are collectively associated with the SMC lineage (reviewed in Doherty and Canfield, 1999; Owens, 1995). These proteins include (but are not limited to) several cytoskeletal or cytoskeletonassociated proteins such as the α chain of SM-actin (Skalli et al., 1989), calponin (Duband et al., 1993), SM22-α (Duband et al., 1993), caldesmon (Frid et al., 1992; Matsumura and Yamashire, 1993), SM-myosin heavy chain (Miano et al., 1994), and smoothelin (van der Loop et al., 1997). Other proteins associated with the pericyte lineage more specifically are desmin (Nehls et al., 1992), aminopeptidase A

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Figure 3 Scanning electron micrograph of pericytes/SMCs associated with microvessels in the rat mammary gland. (A) The SMCs circumferentially surround the vessel in a transitional region between a terminal arteriole and a precapillary arteriole. (B) The SMCs wrap split processes around the vessel in a precapillary arteriole, with a nerve fiber running the ventral length of the vessel (arrow). (C and D) The pericytes wrap multiple processes around the vessel to form a network of contacts on an arterial capillary (C), and on a postcapillary venule (D). Original magnification: (A) ×5000; (B) ×5800; (C) ×4800; (D) ×3600. [Reprinted with permission (Fujiwara and Uehara, 1984).]

(Ramsauer et al., 1998; Schlingemann et al., 1990), and the high molecular weight melanoma-associated antigen, also known as NG2 proteoglycan (Grako and Stallcup, 1995; Schlingemann et al., 1991). Aminopeptidase A is expressed in pericytes throughout the microvasculature (Schlingemann et al., 1991) and the vasoconstrictor, angiotensin II, is a substrate of this enzyme (Healy and Wilk, 1993). Aminopeptidase A is strongly upregulated in activated pericytes such as those found in areas of active angiogenesis (Schlingemann et al., 1996). The NG2 proteoglycan has been suggested as a marker of activated pericytes and immature SMCs and may be involved in mediating cell–matrix interactions (Burg et al., 1996; Tillet et al., 1997) and growth factor responsiveness in pericytes (Goretzki et al., 1999; Grako et al., 1999).

D. Vessel Maturation 1. Remodeling Remodeling is defined as a process that involves an increase or a decrease in vessel diameter to compensate for changes in flow, alterations in perivascular cells as the vessel diameter changes, and expansion or regression of vessels to accommodate

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changing tissue needs. The developing vasculature responds dynamically to the growing needs of the embryo by remodeling vessels as required. The impact of remodeling on vessel bed characteristics was documented many years ago in the rabbit ear chamber model, where dramatic changes in the vessel bed over time could be observed (Clark and Clark, 1939). The authors noted anastomosis or fusion of vessel sprouts, degeneration of extraneous vessels, and thickening and reduction of other vessel structures with variation in flow and perfusion (Clark and Clark, 1939) (Fig. 4). These studies could be interpreted as a wound-healing model; however,

Figure 4 Vessel changes in a capillary bed in a rabbit ear chamber. Changes in microvessels were monitored from 1 h to 8 days after “round table” implant into the rabbit ear. The vessels and sprouts are numbered consistently so that the change in their structure can be seen. The capillary bed is most extensive at 24 h after implantation. Anastomoses of adjacent vessels and degeneration of other vessels result in a single primary vessel remaining in the bed area by day 8. The asterisks indicate vessels that undergo remodeling. [Modified with permission (Clark and Clark, 1939).]

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it is becoming clear that many of the processes of remodeling in healing vasculature are recapitulations of those that occur during development (Cliff, 1963). Remodeling in developing vasculature was observed in early descriptive studies examining flow effects in the developing amphibian (Clark and Clark, 1940). The authors reported dramatic changes in mural cells and suggested that “. . . the longitudinally arranged adventitial cells may increase in number and become transformed into circularly arranged smooth muscle cells following increased pressure in the vessel and that, conversely, muscle cells may revert to the longitudinal arrangement or degenerate following a return to a slower type of circulation with reduced blood pressure . . .” (Clark and Clark, 1940). EC and mural cell proliferation and cell death are likely to occur as vessels grow or regress, respectively, although this has yet to be clearly demonstrated in vivo. Angiogenesis in the corpus luteum of the cycling ovary, where there are rapid changes in the vasculature, is illustrative of vascular remodeling. In the corpus luteum of the pregnant rat there is a direct correlation between proliferation of EC and anastomosis of vessel sprouts in newly formed capillary beds (Meyer and McGeachie, 1988). Similar vessel bed remodeling was observed in the bovine ovary as vessel density increased prior to implantation (Augustin et al., 1995). In a bovine model of corpus luteum formation in the ovarian cycle, the authors closely monitored angiogenesis in the cycle with the criteria of “. . . vessel density, percentage of vessels with lumen and ratio of Bandeiraea simplicifolia-I to von Willebrand factor-positive” ECs. They observed formation of a dense, mature vasculature that slowly regressed (Augustin et al., 1995). Although apoptosis was not detected during the stage of vessel regression, other studies indicate that apoptosis would be the expected mechanism of reduction of vessel structures (Augustin et al., 1995; Saraste and Pulkki, 2000). During the phase of EC proliferation and formation of dense vasculature, there is increased expression of VEGF and its receptors, which does not decrease until the vessels regress (Goede et al., 1998). There is evidence to suggest that VEGF expression is correlated with steroid responsiveness in cells adjacent to regions of rapid angiogenesis in the rat reproductive cycle (Shweiki et al., 1993). Cumulatively, these data indicate a correlation between VEGF expression and the initial phase of EC proliferation and new sprout formation. As VEGF levels are reduced, no new angiogenesis occurs (Goede et al., 1998). It is unclear whether absence of VEGF is sufficient for vessel regression or whether additional signals are required. In addition to the changes in VEGF during angiogenesis in the ovarian cycle are changes in the levels of ang 2. Ang 2 mRNA is expressed at relatively constant levels throughout most of the cycle, but is dramatically increased during vessel regression (Maisonpierre et al., 1997). Similarly, the ratio of ang 2 to ang 1 has been shown to be higher during the period of regression relative to the period of angiogenesis (Goede et al., 1998). Although VEGF has been shown to increase ang 2 expression by microvascular ECs in culture, no direct induction has been demonstrated in vivo (Oh et al., 1999). Ang 1 is expressed throughout the cy-

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cle in the ovary (Maisonpierre et al., 1997). Of note, ang 1 has been reported to act as a survival factor for ECs in vitro (Kwak et al., 1999) and to promote network stabilization in an in vitro assay of EC cord formation (Papapetropoulos et al., 1999). One hypothesis suggests that VEGF promotes angiogenesis, whereas ang 1 promotes stabilization and ang 2 promotes changes in vascular beds (destabilization). This hypothesis was proposed for tumor angiogenesis (Holash et al., 1999a); however, it has also been suggested as a mechanism for developmental vessel remodeling (Gale and Yancopoulos, 1999; Goede et al., 1998). One dramatic example of vessel regression is evinced by the hyaloid vascular system that nourishes the lens, vitreous, and retina during development. When development has proceeded to the point where the hyaloid vessel system is no longer required (prior to birth in humans and early in postnatal life in other mammals), the vessels regress (Oyster, 1999). Ultrastructural studies of the hyaloid system indicate that vessel regression is via apoptosis, although there is no clear indication what initiates the onset of apoptosis (Ito and Yoshioka, 1999). In the hyaloid vessels undergoing regression, ECs and pericytes have a characteristic apoptotic appearance (Saraste and Pulkki, 2000) and there is a dramatic influx of macrophages (Ito and Yoshioka, 1999) and leukocytic cells (Zhu et al., 1999), indicating a requirement for these cells for progression of the apoptosis process in vivo (reviewed in Allen et al., 1998; Jacobson et al., 1997). In an explant model of vascular regression in the pupillary membrane, one component of the hyaloid vascular system, VEGF, was shown to protect vascular cells from apoptosis (Meeson et al., 1999). It may be that the angiopoietins are involved in hyaloid vessel regression as well. One prediction would be that ang 2 would be upregulated, relative to VEGF and ang 1, during the period of active vessel regression. It may be that blocking the vessel-destabilizing signal of ang 2 would result in persistence of the vessel structure, although this has yet to be examined in the hyaloid vasculature. 2. Mural Cells in Remodeling Because mural cells have a physical and temporal association with ECs of new vessels, they are uniquely suited to influence vessel remodeling. Evidence is accumulating to suggest that new vessels lacking mural cells are less stable than those with associated mural cells (Ausprunk and Folkman, 1977; Orlidge and D’Amore, 1987). This hypothesis derives from two concepts. The first is that an unstable vessel is in a phase of development that allows for further growth or regression. The second is that stable vessels are less resistant to change because mural cells suppress EC proliferation and prohibit new vessel growth. The possible role of cell–cell interactions was tested in adult skeletal and cardiac tissue by determining whether pericyte association correlated with new capillary growth (Egginton et al., 1996). Capillaries of skeletal and cardiac muscle were electrically stimulated and then assessed for angiogenesis. The authors noted that capillary numbers increased in areas where pericyte processes had withdrawn (Egginton et al., 1996). Although

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extended times of dissociation between pericytes and ECs were not tested in this study, one prediction would be that a lengthy period without mural cell–EC contact might result in vessel instability and subsequent regression. The authors did not note whether the new capillary structures induced after indirect electrical stimulation became fully invested with pericytes. More recently, the idea that mural cells influence EC stability has been assessed in the retinal vasculature, which undergoes dramatic vascular remodeling during the formation of the mature vasculature. The retina was examined when there were mixed stages of vascular development including a capillary plexus lacking pericytes, vessels with newly associated pericytes (α-SMA positive), and mature vessels with stably invested pericytes (Benjamin et al., 1998). The authors observed that intraocular injection of PDGF-BB resulted in pericyte withdrawal from the vessels and that vessels lacking pericytes underwent regression (Benjamin et al., 1998) (Fig. 5; see color insert). In another series of experiments, application of VEGF increased pericyte investment of vessels and promoted vessel stabilization (Benjamin et al., 1998). There are a number of levels at which pericytes may mediate vessel stabilization. These include, but are not limited to, provision of growth factors to promote survival of ECs and production of basement membrane proteins to enhance adhesion and survival of ECs (reviewed in Doherty and Canfield, 1999). Further evidence of the importance of mural cells in normal vessels can be gleaned from examination of the ang 1 knockout mice, which displayed abnormalities in mural cell investment (Suri et al., 1996). The vasculature appeared to be less complex with reduced vessel branching and fewer large vessels relative to wild type. For example, the ECs of head fold vessels in the ang 1-deficient mice were only loosely associated with underlying matrix. The collagen fiber network below the ECs was less organized in the ang 1 null mice and few mural cells were associated with the vessels (Suri et al., 1996). Similar results were observed with the Tie 2 knockout mice (Dumont et al., 1994). One role of pericytes is to contribute to the production of the basement membrane (Shepro and Morel, 1993). It is possible that in the mice lacking ang 1 and Tie 2, the absence of perivascular cells, which normally secrete angiopoietins, prevents organization of the basement membrane, thereby causing EC instability. It is unclear whether the abnormalities observed are a direct result of the absence of ang 1, are the result of the absence of perivascular cells, or are the outcome of a combination of both conditions.

III. Differences in Vascular Beds A. Variations in Endothelial Cell Gene Expression Although ECs throughout the vasculature originate from similar sources, not all ECs are alike. Aside from the obvious differences in ECs from large versus small

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vessels (Kumar et al., 1987), there are also differences in the repertoire of genes expressed by ECs of different vessel beds. The idea of tissue-specific EC genes was indirectly demonstrated in a study using phage libraries to target specific tissues within an organism (Pasqualini and Ruoslahti, 1996). The surprising outcome of this and subsequent studies was that distinct peptides could mediate phage homing to different populations of ECs throughout the vasculature, including the lung, kidney, and brain (Pasqualini and Ruoslahti, 1996; Rajotte et al., 1998). Each peptide binds to a distinct EC surface protein, a few of which have been identified to date (Burg et al., 1999; Koivunen et al., 1999). In addition to the distinct surface antigens identified above, there are subtle differences in the expression of genes normally associated with the differentiated EC phenoytpe. An example is the von Willebrand factor (VWF) gene, which is expressed by all ECs and is a cofactor for platelet adhesion (reviewed in Ruggeri and Ware, 1993). Message levels for VWF vary among different EC populations (Rand et al., 1987). Reporter studies using a portion of the flanking sequence and the first exon of the VWF gene coupled to lacZ have been able to direct expression to some microvascular ECs in the brain (Aird et al., 1995). Further studies using transgenic mice expressing larger segments of the VWF promoter coupled to lacZ revealed VWF gene expression in microvessels in the brain, heart, and skeletal muscle (Aird et al., 1997). Plasminogen activator inhibitor gene expression also varies among different vascular beds. Plasminogen activator inhibitor is high in aorta, heart, and adipose tissue, but low in liver, adrenal gland, and kidney vasculature (Sawdey et al., 1989). There are also differences in the antigenicity of various EC populations (Page et al., 1992) as well as variations in gene expression after injury, depending on the tissue bed affected (Gerritsen and Bloor, 1993). Adenoviral vectors have been used to express VEGF164 (mouse isoform) in a variety of normal tissues in adult mice and rats (Pettersson et al., 2000). All overexpressing tissues produced enlarged and dilated vessels (mother vessels) after 3 days; but there were differences in the temporal and qualitative response in various vascular beds, with fat showing sustained angiogenesis relative to striated muscle, for example (Pettersson et al., 2000).

B. Endothelial Cells of Barrier Vessels One of the clearest examples of how the microenvironment can influence gene expression in ECs is found in the blood–brain barrier (BBB) (Bar, 1980) and the blood–retina barrier (BRB) (Sagaties et al., 1987). These vascular beds differ from others both structurally and functionally. The BBB and BRB form a semipermeable barrier that limits and controls diffusion of substances across the vessel wall into the neural tissue (Reese and Karnovsky, 1967) to a much greater degree than that observed in other vascular beds. The ECs of the BBB and the BRB form tight junctions, central structures in the establishment of the semipermeable

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barrier (reviewed in Yap et al., 1998; Spring, 1998). The ECs of the barrier vessels are characterized by elevated expression of metabolic enzymes such as glucose transporter (Virgintino et al., 1998) and γ -glutamyltransferase (Lawrenson et al., 1999). In addition, the ECs of the barrier vessels express high levels of several of the proteins commonly associated with tight junctions (Tsukita et al., 1999), including ZO-1 (Zonula occludens 1; Stevenson et al., 1986), ZO-2 (Jesaitis and Goodenough, 1994), occludin (Furuse et al., 1993; Sakakibara et al., 1997), and claudin (Furuse et al., 1998a,b). Cell–cell contact between ECs and astrocytes is thought to be involved in establishing the unique properties of the BBB and the BRB (Meyer et al., 1991). Astrocytes have fine processes that have been shown to come into close proximity with the ECs (Young, 1991). In an in vivo model of barrier formation, astrocytes were injected into the anterior eye chamber of rats, where the cells formed aggregates and became vascularized (Janzer and Raff, 1987). The new vessels with associated astrocytes displayed barrier properties not observed in the new vessels without associated astrocytes (Janzer and Raff, 1987). In developing cortical vessels, ECs migrate into the avascular zones in the vicinity of astrocytes (Bar, 1980). In the retinal vasculature, EC precursors arise in the avascular zone and are thought to induce a wave of astrocyte cell migration from the optic nerve, closely followed by a wave of migrating ECs (Chan-Ling and Stone, 1991; Ling and Stone, 1988; McLeod et al., 1987). In both vascular beds, EC association with astrocytes precedes expression of proteins associated with the differentiated ECs (Jiang et al., 1995). Once EC migration has occurred, the astrocytes associate with newly forming vessels (Chan-Ling and Stone, 1991; Gariano et al., 1996). In a study examining development of the human fetal retinal vasculature, astrocyte migration was shown to be temporally correlated with the transition of ECs from migrating, elongated precursors to rounded ECs in cordlike formations (Hughes et al., 2000). Some factors have been proposed as candidates to mediate the astrocytedependent differentiation of the BBB and BRB. One such candidate is fibronectin, which has been shown to be expressed at the time of astrocyte migration and may be a component of the guidance pathway that directs EC migration (Jiang et al., 1994). In the retina there is evidence to suggest that PDGF-A is involved in astrocyte migration. PDGF-A is expressed locally by the retinal ganglion cells, and the PDGF-A receptor PDGFRα is expressed by the astrocytes (Mudhar et al., 1993). When neonatal rats were injected with Cos cells expressing soluble PDGFRα receptor (dominant negative), astrocyte migration into the retina was reduced compared with control-treated animals and there was an accompanying reduction in vessel growth, most likely due to the abnormal astrocyte migration (Fruttiger et al., 1996). Similar results were obtained in mice given subcutaneous injections of a PDGFRα-neutralizing monoclonal antibody; vessel growth was observed only where astrocytes could be detected (Fruttiger et al., 1996). In the same study, the authors overexpressed PDGF-A in retinal ganglion cells and observed an increase

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in the number of astrocytes and a coincident increase in vascularization (Fruttiger et al., 1996). There is also evidence that factors such as VEGF, which is involved in neovascularization in other vessel beds, are also involved in the BBB and BRB vessel beds (Nag et al., 1997; Provis et al., 1997; Yi et al., 1998). A potential role for the microvascular pericyte in the formation of the barrier has been proposed, but the specifics remain unclear (Balabanov and Dore-Duffy, 1998; Dehouck et al., 1997; Ramsauer et al., 1998). However, it is clear that the neural microenvironment contributes to the establishment of the barrier properties and that the environment in the retina and the central nervous system is distinct from that of other tissues.

C. Arterial versus Venous Endothelial Cells Another example of the variability in microvascular beds is the distinctive pattern of ephrin expression in the ECs of arteries versus veins. The ephrins are a growing family of membrane-associated proteins that interact with and signal through receptor tyrosine kinases called Ephs (Br¨uckner and Klein, 1998). Studies revealed that arteries and veins express different combinations of these proteins. Most distinctive is the pattern of the Eph B4 receptor, which was confined to the venous side of the vasculature, whereas the specific ligand for this receptor, ephrin B2, was expressed in arterial ECs (Wang et al., 1998). Undoubtedly, the designation of arterial versus venous ECs is not due entirely to expression of these two proteins. In fact, analysis of other ephrin and Eph receptor isoforms indicated that the arterial/venous division may be further complicated by overlapping expression of ephrins and their receptors (Adams et al., 1999). In situ hybridization and ligand-binding studies revealed that ephrins B1 and B2 are expressed in arteries, ephrin B1 and Eph B4 are expressed in veins, and Eph B3 is expressed in veins and in some arteries (Adams et al., 1999). These results suggest that combinations of ephrins and Ephs are likely to play a role in establishment of the arterial and venous vessels. In ephrin B2-deficient mice the venous system was developed, but it did not have the normal branching pattern observed in a mature vasculature (Wang et al., 1998). Therefore, in the absence of artery-derived ephrin B2, remodeling in the adjacent venous system was perturbed. Analysis of mice deficient in both Eph B2 and Eph B3 exhibit a phenotype similar to that of the Eph B2-deficient mice in that the cardiac, cranial, and intersomitic vessels are generally smaller and less organized than their wild-type counterparts (Adams et al., 1999). The role that ephrins play in vascular development is unclear; however, evidence suggests that ephrin B1 can activate cell attachment via integrins, thus establishing a connection to the ECM (Huynh-Do et al., 1999). Moreover, ephrin B2 and the B2/B3 null mice have abnormal cardiac trabeculation (Adams et al., 1999), with a phenotype quite similar to that of mice with targeted disruptions of ang 1 (Suri et al., 1996), neuregulin (Meyer and Birchmeier, 1995), and neuregulin receptor (Lee et al., 1995). Although

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no distinct pattern of expression, such as arterial versus venous, has been described for the angiopoietins or the neuregulins, further examination may reveal whether their expression patterns are cell-autonomous or microenvironmentally controlled. Another example of differences in gene expression between arterial and venous EC is seen with the gridlock gene product in zebrafish (Weinstein et al., 1995). gridlock is clearly expressed in the presumptive dorsal aorta, but not in the axial or bilateral cardinal veins (Zhong et al., 2000). A mutation in this gene results in abnormal aortic development and a lack of blood flow to the caudal region of the embryo (Zhong et al., 2000). The mechanism of how gridlock functions has yet to be elucidated. More systematic analysis of arterial and venous ECs is likely to reveal additional proteins that are differentially expressed. It will be interesting to determine whether these proteins are responsible for or are a result of arterial versus venous patterning.

IV. Parallels between Angiogenesis in Development and Pathology The focus of this review has been largely developmental; however, it is increasingly apparent that there are similarities in the mechanisms that regulate angiogenesis during development and in the postnatal organism (Carmeliet and Collen, 1997). Although there are many examples of postnatal neovascularization, such as in wound healing (Carmeliet and Collen, 1997; Cliff, 1963) or atherosclerosis (Schwartz et al., 1990), we focus our attention on two representative examples, tumor angiogenesis and ocular neovascularization.

A. Tumor Angiogenesis Although there is tremendous variability among the vessels that invest tumors, from sinus-like large vessels to unstable microvessels (Dvorak et al., 1999), they appear to share a degree of instability not observed in “normal” vasculature (Folkman, 1996). One explanation for instability of tumor vasculature is that it represents a developmentally “suspended” vascular bed that contains regions of relatively “stable” vessels, but does not progress to the mature vessel state seen in the normal vasculature. ECs during development and in tumors display similar properties as they invade surrounding tissue, migrate, and form new basement membrane (Ausprunk and Folkman, 1977; B¨ohle and Kalthoff, 1999; Dvorak et al., 1999). One example of angiogenesis is seen in hemangiomas, which commonly occur in premature babies or in early postnatal life (Folkman and D’Amore, 1996; Takahashi et al., 1994). These benign tumors regress over time, but their early phase is

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characterized by rapid capillary vessel growth and they have high numbers of activated perivascular cells and EC (Mulliken and Glowacki, 1982; Mulliken et al., 1982). There are contradicting reports regarding the expression of the adherens junction protein, VE-cadherin, in hemangiomas, with some reported as having poor expression of VE-cadherin (Dejana et al., 1995) and others having a high expression of VE-cadherin (Mart`ın-Padura et al., 1995). The differences may be explained by the fact that there are different subtypes of hemangiosarcomas and angiosarcomas. Regardless, it is apparent that abnormal regulation of EC intercellular adhesion may be one of the factors that characterizes pathologic versus developmental angiogenesis (Dejana, 1996). A number of the proteins that have been demonstrated to regulate developmental angiogenesis have been shown to play a role in tumor angiogenesis as well. Most prominent among these are VEGF and the angiopoietins (Folkman and D’Amore, 1996). VEGF has been shown to be upregulated in hemangiomas (Takahashi et al., 1994) and glioblastoma in vivo (Plate et al., 1992, 1994). In a glioblastoma tumor model, stable lines of U87MG cells engineered to overexpress the three isoforms of VEGF (121, 165, and 189) (Cheng et al., 1997) were injected intracerebrally into mice. The resulting tumors were all vascularized, but the VEGF121- and VEGF165expressing cells had leaky and ruptured vessels, whereas the VEGF188-expressing mice did not (Cheng et al., 1997). In work done to test the instability of tumor vessels, xenografted gliomas induced to express VEGF were shown to have a high number of vessels without associated pericytes/SMCs (Benjamin et al., 1999). When VEGF expression was turned off in these tumors, the ECs in vessels lacking pericytes detached and underwent apoptosis (Benjamin et al., 1999). Thus, although tumor vessels appear to be abnormal in that they are chronically immature, the vessels have a VEGF-dependent survival phase that is similar to that observed newly formed normal vessels. The angiopoietins, which have been shown to be critical in developmental vascularization, have also been examined for their possible role in tumor vascularization. Expression of ang 1, ang 2 mRNA, and Tie 2 protein has been described in glioma vasculature (Stratmann et al., 1998). Ang 1 and Tie 2 mRNA levels were higher in glioma and astrocytoma than in normal brain samples and ang 2 mRNA was detected only in the tumor samples (Stratmann et al., 1998). It is interesting to note that high VEGF expression has been localized to areas of active angiogenesis in glioblastomas (Plate et al., 1992; Shweiki et al., 1992) and that VEGF can upregulate ang 2 in microvascular ECs in vitro (Oh et al., 1999). When C6 glioma cells or RBA mammary adenocarcinoma cells were implanted into rat brains there was induction of ang 2 in the resulting tumors, with regression of “coopted” vessels in regions lacking VEGF and angiogenesis in regions expressing VEGF (Holash et al., 1999a). Ang 1 has also been proposed to antagonize the permeabilityinducing effects of VEGF (Thurston et al., 1999, 2000). Thus, one possible explanation for the instability of tumor vasculature is that localized expression

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of the angiopoietins and VEGF mediates variations in vessel permeability, remodeling, and angiogenesis throughout the tumor (Holash et al., 1999b). Attention has been given to the role of homeobox genes in the development of the cardiovascular system and in tumorigenesis (Patterson et al., 1998). The Hox D3 gene, specifically, has been proposed to act in the transition from resting to invasive ECs (Boudreau et al., 1997). Hox D3 is coexpressed with the α vβ 3 integrin in endotheliomas in vivo and expression of a Hox D3 retrovirus in chick CAM resulted in endothelioma-like dilated vessels (Boudreau et al., 1997). The α vβ 3 integrin has been proposed to function during angiogenesis by mediating EC binding to the temporary matrix that is established during EC invasion and migration into the surrounding tissue (Brooks et al., 1994a, 1996). Although Hox D3 has not been shown to directly upregulate α vβ 3, application of Hox D3 antisense oligonucleotides to human umbilical vein endothelial cells (HUVECs) can block the activation-induced upregulation of α vβ 3 (Brooks et al., 1996). Thus, Hox D3 is likely to play a role in the gene expression switch required for developmental as well as tumorigenic angiogenesis. An interesting correlation between developmental and pathologic angiogenesis has been made with regard to expression of the Id proteins (Id1–Id4) (Carmeliet, 1999). Id proteins are transcription factors that have been implicated in developmental regulation of neurogenesis (Duncan et al., 1992; Ishibashi et al., 1995) and lymphangiogenesis (Pan et al., 1999; Yokota et al., 1999). Expression of Id proteins has been most closely associated with the transition from a proliferative precursor to a differentiated cell (Norton et al., 1998); during differentiation of myoblasts to striated muscle, Id protein levels are reduced and overexpression of Id proteins in C2C12 muscle cells reduced expression of several striated muscle markers and inhibited differentiation (Jen et al., 1992). In addition, differentiation of a mammary epithelial cell line (SCp2) a loss of Id1 protein (Desprez et al., 1995) and is associated with a corresponding decrease in matrix metalloproteinase (MMP) expression that is correlated with reduced invasiveness (Desprez et al., 1998). An interesting pattern of Id mRNA expression occurs during mouse development; there is reduced expression of Id genes during the differentiation of a number of tissues including the vasculature (Ellmeier and Weith, 1995; Jen et al., 1997). Id1, Id2, and Id3 are expressed in developing EC, but only Id1 and Id3 are expressed in the central nervous system (CNS) vasculature (Jen et al., 1997; Lyden et al., 1999). In mice lacking both Id1 and Id3, CNS vessels were dilated and there was an absence of branching and remodeling with reduced production of the basement membrane protein laminin (Lyden et al., 1999). Mice with reduced Id levels (Id1+/–; Id3–/–) also showed reduced levels of MMP2 and a corresponding reduction in their ability to support growth of several implanted tumors (Lyden et al., 1999). Id1 and Id3 expression levels were low in quiescent vessels and high in vessels actively undergoing angiogenesis in development or during tumor growth (Lyden et al., 1999). Thus, Id proteins are likely to be involved in the switch from undifferentiated to differentiated ECs and from quiescent to activated ECs.

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B. Ocular Angiogenesis An example of pathologic angiogenesis that is uniquely developmental in origin is retinopathy of prematurity (ROP), a condition of premature infants that is thought to be initiated by a transition from hyperoxia to hypoxia (James, 1976). Normal vessel growth in the mammalian retina occurs by a combination of vasculogenesis and angiogenesis (Chan-Ling, 1994; Hughes et al., 2000; McLeod et al., 1987; Provis et al., 1997) and the neovascularization characteristic of ROP may result from a combination of these processes as well (Chan-Ling and Stone, 1993). Briefly, retinal vascularization is thought to be induced by a “physiologic hypoxia” that results when the differentiation of neural retina leads to greater oxygen consumption and a thicker retina. This hypoxia causes elevated VEGF expression by astrocytes, which in turn directs the vascularization of the retina. Exposure of premature infants to hyperoxia, required because of pulmonary distress, suppresses hypoxiainduced VEGF expression, leading to regression of existing immature vessels and reduced vascularization. On return to room air, the now undervascularized retina senses hypoxia, upregulates VEGF, and a pathologic process of neovascularization occurs. The vessels observed in ROP are disorganized and leaky, resulting in poor oxygenation of the retinal tissue and subsequent loss of retinal ganglion cells and photoreceptors (Chan-Ling and Stone, 1993). The source of the neovascularization signal was identified as the retina, itself (Glaser et al., 1980), and the activity was increased in extracts from hypoxic retinas (Garner and Kissun, 1987). VEGF is thought to mediate developmental vascularization in the retina (Provis et al., 1997; Yi et al., 1998) as well as neovascularization in the progression of ROP (Vinores et al., 1997). Experiments utilizing slow release of VEGF from pellets implanted in the vitreous of primates and rabbits resulted in dilation of retinal vessels with some accompanying remodeling and iris neovascularization (Ozaki et al., 1997). Moreover, a soluble form of the VEGF receptor that inhibits VEGF signaling was used in a mouse model of ROP to inhibit neovascularization (Aiello et al., 1995), further confirming the integral role that VEGF plays in ocular neovascularization. Many of the characteristics of vessel bed remodeling that occur during development are mirrored in diabetic retinopathy, a pathology that is characterized by vascular dysfunction and a progressive loss of retinal vessels (Garner, 1993). The retinal vasculature appears to be particularly sensitive to the fluctuations in glucose levels experienced in the diabetic condition (reviewed in Archer, 1999). The early stages of diabetic retinopathy are characterized by loss of pericytes from the vasculature and a subsequent onset of vessel instability and acellularity (Cogan et al., 1961; Kuwabara and Cogan, 1963). In hypertensive diabetic rats, there were dramatic changes in retinal capillaries including pericyte rounding, thickening of the basement membrane between pericytes and microvascular ECs, and increasing vessel leakiness (Dosso et al., 1999). The vascular degeneration in diabetes appears to be due to apoptotic cell death of the pericytes prior to and concomitant with the

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loss of ECs (Mizutani et al., 1996). Genes that regulate apoptosis (e.g., bax) have been shown to be upregulated in bovine retinal pericytes in vitro (Li et al., 1998) and in human retinal pericytes from diabetics in vivo (Podesta et al., 2000). This is apparently the reverse of the situation observed during development, during which vessel beds are remodeled and stabilized after investment with pericytes. The integrin α vβ 3, which has been implicated extensively in vascular development, has been shown to be expressed in eye tissue from patients undergoing ocular neovascularization (Friedlander et al., 1996). Expression of α vβ 3 was detected in choroidal neovascularization tissue from patients with the wet form of age-related macular degeneration and from ocular histoplasmosis tissue as well as in retinas of patients with proliferative diabetic retinopathy (Friedlander et al., 1996). Since α vβ 3 is predominantly expressed on angiogenic ECs (Brooks et al., 1994a), it is interesting to note that this integrin is likely to be involved in EC adhesion to matrix in pathologic angiogenesis. A considerable body of literature exists linking VEGF expression in ischemic conditions and ocular neovascularization, both in development (Stone et al., 1995) and pathology (Vinores et al., 1997), where the common denominator appears to be a period of relative hypoxia that stimulates VEGF expression (reviewed in Adamis et al., 1999; Dor and Keshet, 1997; Miller, 1997). Neutralizing antibodies to VEGF have been used to inhibit the iris neovascularization that accompanies retinal ischemia, marking the dramatic involvement of VEGF in pathologic neovascularization (Adamis et al., 1996a,b) (Fig. 6; see color insert). Under hypoxic/ischemic conditions, both developmental and pathological, VEGF expression is likely controlled, at least in part, by members of the hypoxiainducible factor (HIF) family (reviewed in Flamme et al., 1998; Guillemin and Krasnow, 1997; Ozaki et al., 1999; Semenza, 1998). HIF-1, a dimer of α and β chains, is a transcription factor that upregulates a number of genes in response to hypoxic and stress conditions (Semenza, 1998; Wenger and Gassmann, 1997). HIF-1-based regulation of VEGF expression is probably the most extensively studied means for induction of VEGF. VEGF and HIF-1α were upregulated in the retinas of mice exposed to hypoxic conditions for 2 h, while HIF-1β levels remained high, but constant (Ozaki et al., 1999). Mice lacking HIF-1α are embryonic lethal at E10.5 and have several vascular abnormalities, including an absence of cephalic vasculature (by E8.5) and disorganization and a lack of branching in the yolk sac vessels (Iyer et al., 1998; Ryan et al., 1998). Embryonic stem (ES) cells from the null mice form smaller tumors with reduced vasculature and VEGF expression, relative to wild-type mice, when induced to form teratocarcinomas in immune-compromised mice (Ryan et al., 1998). Moreover, HIF-1α –/– ES cells are resistant to apoptosis induced by hypoxic or hypoglycemic conditions, but not to apoptosis induced by cytokine withdrawal (Carmeliet et al., 1998). In contrast, other studies have shown a clear induction of VEGF in HIF-1α knockout mice in response to glucose deprivation, but not hypoxia, suggesting a clear distinction between the two stress pathways (Iyer et al., 1998; Kotch et al., 1999).

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V. New Directions The forefront of vascular research is advancing rapidly with the widespread use of molecular techniques. We have identified a few topics that are of particular interest. These fall within the broad area of cell–cell interactions and effects of microenvironment on cell plasticity and phenotype. These topics are ripe for revealing new insights into the mechanisms of cell–cell interactions that underlie the development of the vascular system. A. Endothelial Cells as Mural Cell Precursors Pericytes are thought to arise from mesenchymal cells or from SMCs locally available at sites of new vessel growth, but an additional possible source of perivascular cells has been suggested. Within larger vessels one source of SMCs may be the EC themselves (reviewed in Majesky and Schwartz, 1997). ECs are hypothesized to have the potential to transdifferentiate into pericytes/SMCs, although normally the majority of ECs do not. Preliminary evidence to support this hypothesis comes from studies to determine where circulating ECs incorporate in the developing avian embryo (DeRuiter et al., 1997). Quail ECs (E2.5) were labeled with wheat germ agglutinin/colloidal gold and allowed to develop. After 19 h the embryos were examined for endocardial colocalization of QH1 (an EC marker) and lectin/gold or α-smooth muscle actin (α-SMA), an SMC marker. Gold-labeled, α-SMA-positive cells were detected in the subendothelial layer (DeRuiter et al., 1997). These results are intriguing, but controversial because of the choice of markers, neither of which is absolutely cell type specific. Another study used an in vivo model of large vessel transplants to examine mural cell recruitment via transdifferentiation of ECs (reviewed in Williams, 1995). In preparing ECs for autologous graft implants, SMCs were consistently observed in the subendothelial space, separate from the multilayered SMCs. The authors suggest that transdifferentiation of ECs to SMCs and subsequent seeding of the EC layer could explain their observation. If future evidence supports this idea of EC “transdifferentiation” to mesenchymal cell within larger arteries, it would parallel the endothelial-to-mesenchymal transdifferentiation observed in septation of the heart (Eisenberg and Markwald, 1995). It is interesting to speculate whether any of the factors identified in the atrioventricular transition, such as TGF-β family members (Boyer et al., 1999; Brown et al., 1999; Potts et al., 1991), angiopoietins (Suri et al., 1996), ephrins (Adams et al., 1999), or neuregulins (Lee et al., 1995; Meyer and Birchmeier, 1995), may play a role in the EC transdifferentiation to SMCs. EC plasticity, potential differentiation pathways, and stabilization are particularly important with regard to studies aimed at generating arteries in vitro for transplantation (Niklason et al., 1999). Perhaps ECs are sufficiently plastic in their phenotype to switch from a modified epithelial cell

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to a mesenchymal cell on the basis of their location within the intimal wall and in response to microenvironmental influences.

B. Cell–Cell Communication and Gap Junctions One possible mode for cell–cell communication in the vasculature is gap junctions. Gap junctions are hexameric channels in the cell membrane that are direct points of connection between cells and through which small molecules may move by passive diffusion (reviewed in Kumar and Gilula, 1996). Hydrophobic molecules of ≤1.5 nm in diameter can readily pass through gap junctions, thus allowing rapid intercellular exchange of ions and small molecules, such as those utilized in second-messenger signaling pathways (reviewed in Loewenstein, 1981; Spray, 1994). Connexin genes have interesting expression patterns in the cardiovascular system (Delorme et al., 1997; Gourdie et al., 1993; Ko et al., 1999; Yeh et al., 1997). The α 5 connexin/connexin 40 (Cx40) (Bastide et al., 1993) and α 4 connexin/connexin 37 (Cx37) (Reed et al., 1993) proteins are expressed in ECs, and α 1 connexin/connexin 43 (Cx43) (Lo et al., 1997; Waldo et al., 1999) is expressed in selective regions of the heart. Blocking gap junctional communication during development in Xenopus (Levin and Mercola, 1998), overexpressing a dominant negative form of Cx43 in mice (Sullivan et al., 1998), or knocking out the gene for Cx43 (Huang et al., 1998; Reaume et al., 1995) results in left/right asymmetry and abnormal cardiac development, including septation defects. Both the dominant negative Cx43 transgenic mice and the Cx43 null mice are embryonic lethal (Reaume et al., 1995; Sullivan et al., 1998). Overexpression of the Cx43 gene in mice also resulted in heart malformations. When the Cx43-overexpressing mice were bred with the Cx43 null mice, the resulting offspring had increased viability (relative to the Cx43 knockout), indicating that there was a partial rescue with overexpression of Cx43 in the phenotype of the null mice (Ewart et al., 1997). Mice lacking Cx40 are viable, but develop arrhythmia, likely due to the absence of Cx40 in the atrioventricular node (Hagendorff et al., 1999; Kirchhoff et al., 1998; Simon et al., 1998). These results lend further support to the importance of connexin function in development and maintenance of the cardiovascular system. Increasing evidence indicates that expression of the connexin genes varies throughout the vasculature. For example, Cx37, Cx40, and Cx43 show different expression patterns in aortic versus coronary vessels among bovine, pig, and rat aorta (van Kempen and Johngsma, 1999). In this study both ECs and SMCs (except rat aortic SMCs) were found to express Cx40, whereas Cx43 was found predominantly in SMCs with only an occasional positive EC (van Kempen and Johngsma, 1999). Cx37 was lowest overall and was restricted to ECs (van Kempen and Johngsma, 1999). Using triple immunolabeling to visualize connexins in rat aorta and pulmonary arteries, Cx40, Cx37, and Cx43 were detected within the same junction in ECs (Yeh et al., 1998). Dye tracer analysis of junctional connections

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indicates that functional communication exists between ECs and between ECs and SMCs, although the latter appears to be directionally biased, favoring the EC-to-SMC flow of tracer dyes (Little et al., 1995). Gap junction genes are regulated both temporally and spatially, particularly in response to environmental cues. In vitro evidence has shown that Cx43 expression is rapidly upregulated in bovine aorta-derived ECs in response to nonlaminar flow (DePaola et al., 1999) and in SMCs by stretch (Cowan et al., 1998). Cx43 has been shown to be expressed in vivo at vessel branch points and regions of nonlaminar flow, whereas Cx37 and Cx40 are primarily localized to regions of laminar flow in rat aortic endothelium (Gabriels and Paul, 1998). Of note, there was an upregulation of Cx43 in rat aortic ECs in response to shear stress (induced by aortic coarctation), but no apparent change in the level of Cx37 or Cx40 (Gabriels and Paul, 1998). Thus, gap junctional communication between cells may play a critical role not only in differentiation of the vasculature, but also in vessel function in response to changes in flow and pressure and in maintenance of vascular tone (Christ et al., 1996).

C. Neuronal–Vascular Interactions One area of importance that has yet to be fully examined is the effect of innervation on vascular development. Perivascular nerves that innervate large vessels form a plexus that has no direct contact with the ECs, but the nerves do form “en passant” synapses with SMCs (Burnstock, 1986). Electron microscopic examination of the rat mammary gland vasculature revealed dense innervation of the terminal arterioles, less innervation in the capillary arterioles, an occasional process associated with the arterial capillaries, and no innervation in the venous microvasculature (Fujiwara and Uehara, 1984) (e.g., Fig. 2B; see color plates). Similarly, in the rabbit ear arteriovenous anastomoses, peptidergic innervation is detected only on the arterial and not on the venous portion of the vessel (Golfert et al., 1998). Peptidergic neurons associated with the vasculature have been localized to small blood vessels as early as 16–17 weeks of gestation in humans (Terenghi et al., 1993). Innervation, then, is an interesting distinction between arterial and venous vessels that may prove to be a fruitful area of research, particularly with regard to the potential influence of the neural contact on the differentiation/specification of the vascular cells. Innervation may influence vessel remodeling during development by altering vascular tone in response to alterations in flow and tissue need (reviewed in Burnstock, 1985; Burnstock and Ralevic, 1994). There is considerable evidence to indicate that vascular tone is regulated by two components: vasoeffectors released by the ECs (Vanhoutte and Rimele, 1983) and neurotransmitters/neuropeptides released by autonomic and sensory nerves; both affect mural cell contractility (Dhital and Burnstock, 1989). Some of the neurohumoral substances may have additional effects, such as altering vascular tone. The neuropeptide substance P

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induces proliferation of microvascular ECs and HUVECs in vitro and stimulates neovascularization in a corneal pocket assay (Ziche et al., 1990). Neuropeptide Y, which is expressed in cardiac sympathetic nerves, acts as a mitogen for SMCs in culture (Zukowska-Grojec et al., 1993a,b) and has been shown to induce tube formation and capillary sprouting in vitro (Zukowska-Grojec et al., 1998). In addition, rat superior cervical ganglion neurons stimulate proliferation of rat arterial SMCs, and TGF-β and endothelin signaling are likely mediators of the proliferation observed in the cocultures (Damon, 2000). Although these data are obtained from tissue culture studies, the results suggest that ECs and SMCs can respond to factors normally considered to be neuronal inflammatory factors (substance P) or autocrine modulators of neurotransmitter release (neuropeptide Y), respectively. Neuropeptide Y, in particular, has been suggested to exacerbate heart disease and hypertension (Zukowska-Grojec and Wahlestedt, 1993a,b). Another circumstance in which innervation may affect vascular development is via interactions between the neuron and its target cell, the SMC. There is considerable evidence in neural development that innervation of a target initiates a cascade of events that induce the target to provide trophic support (in the form of survival factors) for the neurons (reviewed in Pettmann and Henderson, 1998). Survival of the neurons and successful innervation of the target may result in differentiation of the target; thus, both the neuron and its target cell are altered by the interaction (Landis, 1994). One trophic factor, nerve growth factor (Levi-Montalcini, 1987), can induce nerve growth in the cerebral artery (sympathetic) and the tail vein (nonsympathetic) in rats that have reduced innervation due to aging (Andrews and Cowen, 1994). Moreover, the age of the target vasculature (Cowen et al., 1982; Gavazzi et al., 1992) and the presence of Alzheimer’s disease (Tong and Hamel, 1999) have been shown to determine the innervation pattern, indicating the importance of neuron–target interactions in the development and stabilization of the vasculature, particularly with regard to the influence of local microenvironmental conditions. The interaction between neurons and the vasculature takes on even more relevance with the growing list of proteins that play critical roles in both vascular and neuronal development. For example, the ephrins and the Eph receptors were shown to be involved in establishing repulsive cues in cell–cell contact mediated neuronal migration during development (reviewed in O’Leary and Wilkinson, 1999). These proteins have been identified as components of arterial and venous specification, vascular remodeling, and cardiac development (Gale and Yancopoulos, 1999). Another neuronal migration factor that does double duty is neuropilin. Neuropilin, which is involved in cell–cell contact-mediated axon guidance (Tessier-Lavigne and Goodman, 1996), has been identified as a receptor for VEGF (Soker et al., 1996, 1998); neuropilin is also a receptor for collapsin 1, which signals the repulsive cues in the nervous system (He and Tessier-Lavigne, 1997). Collapsin and VEGF165 compete for binding to neuropilin and have opposing effects on EC motility, inhibition, and promotion (Miao et al., 1999). As functional overlap dovetails

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between these and doubtless other factors involved in neuronal and vascular development, guiding principles that govern cell–cell interactions in development of both systems will become clear.

D. Vasculogenesis in the Adult Until recently, it was thought that vasculogenesis occurred only in the early stages of embryogenesis. However, evidence is accumulating to indicate the existence of EC precursors, as a source of stem cells for vasculogenesis, in the bone marrow of the adult (Hatzopoulos et al., 1998; Isner, 1998; Lin et al., 2000; Shi et al., 1998). Stem cells can be defined as “. . . cells that are capable of extensive proliferation and that are able to give rise to additional cells similar to themselves (“self-renewal”), as well as to differentiated progeny” (Siminovitch et al., 1963). While this functional description of a stem cell was originally applied to hematopoietic stem cells, it is valid with regard to EC progenitor cells as well (Morrison et al., 1997; Till, 1981). EC progenitor cells were isolated from human peripheral blood on the basis of cell surface expression of CD34/PECAM and Flk-1/VEGFR-2 (Asahara et al., 1997). The progenitor cells differentiated into ECs in vitro and became incorporated into new vessels in an in vivo model of hind-limb ischemia (Asahara et al., 1997). Further confirmation of the existence of EC progenitors came from a series of experiments utilizing transgenic mice expressing the lacZ gene under the control of the Tie 2 or VEGFR-2 promoter (Asahara et al., 1999a). Putative progenitor cells isolated from the blood of these lacZ-expressing donor mice were transplanted into host mice that had been irradiated to destroy the bone marrow. Histologic examination of the mice showed β-galactosidase (β-Gal)-positive ECs in new vessels that had formed in several pathologic models including tumor growth, wound healing, hind-limb ischemia, and cardiac ischemia, as well as in physiologic models such as corpus luteum formation and endometrial growth (Asahara et al., 1999a). Further evidence suggested that administration of VEGF led to increased numbers of circulating EC progenitor cells and their subsequent incorporation into new vessels in a corneal pocket neovascularization assay (Asahara et al., 1999b). It is unclear whether VEGF directly increases the numbers of EC progenitor cells, increases the incorporation of EC progenitors in new vessels, or acts via a combination of both effects. The existence of EC progenitor cells provides new possibilities for promoting vascular growth in the adult. In cases of hypertensive vascular damage, ischemia– reperfusion, or vessel loss during peripheral vascular disease, circulating EC progenitors may be tapped as a source for promoting neovascularization. This approach would not require immune system compromise, since the source of the progenitors would be the patients themselves. Indeed, in a development for treatment of peripheral vascular disease (Rivard et al., 1999), patients were treated

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with intramuscular injection of plasmid DNA-encoding VEGF (Baumgartner et al., 1998) and had an overall increase in neovascularization and reperfusion in injured/ischemic areas. One possible mechanism of VEGF action in these patients is that the VEGF may have stimulated vasculogenesis by increasing the numbers of circulating EC precursors. Alternatively, and more likely, local production of VEGF may have stimulated angiogenesis to promote revascularization and reperfusion. The latter explanation is supported by studies in which myoblasts, engineered to express VEGF165, were injected into adult mouse muscle (Springer et al., 1998). Examination of the injection site showed increased numbers of ECs, increased vascular channels, and the development of hemangiomas adjacent to the injection site, indicating that uncontrolled angiogenesis had occurred (Springer et al., 1998). Although these studies may be interpreted to suggest that normal, rather than compromised, tissues were responding to local signals of VEGF, it may be that the injection itself elicits a local wounding response that induces angiogenesis. In addition, it is difficult in these studies to distinguish between neovascularization by vasculogenesis versus angiogenesis (Rafii, 2000), but future efforts combining basic research and clinical applications in this area are likely to be fruitful.

VI. Summary Research into areas as divergent as hemangiopoiesis and cardiogenesis as well as investigations of diseases such as cancer and diabetic retinopathy have converged to form the face of research in vascular development today. This convergence of disparate topics has resulted in rapid advances in many areas of vascular research. The focus of this review has been the role of cell–cell interactions in the development of the vascular system, but we have included discussions of pathology where the mechanism of disease progression may have parallels with developmental processes. A number of intriguing questions remain unanswered. For example, what triggers abnormal angiogenesis in the disease state? Are the mechanisms similar to those that control developmental neovascularization? Perhaps the difference in development in angiogenesis versus in disease is context driven, that is, an adult versus an embryonic organism. If this is the case, can the controls that curtail developmental vessel formation be applied in pathologies? Can cell–cell interactions be targeted as a control point for new vessel formation? For instance, can perivascular cells be stimulated or eliminated to result in increased vessel stability or instability, respectively? If the hypothesis that mural cell association is required for vessel stabilization is accurate, are there mechanisms to promote or inhibit mural cell recruitment and differentiation as needed? These and other questions lie in wait for the next generation of approaches to discern the mechanisms and the nature of the cell–cell interactions and the influence of the microenvironment on vascular development.

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Acknowledgments The authors gratefully acknowledge Dr. Laura Benjamin, Dr. Charles Little, Dr. Chris Drake, and Dr. Anthony Adamis for contributing their data (published and unpublished); and Dr. Takashi Fujiwara, Yasuo Uehara, Dr. Eliot Clark, and Dr. Eleanor Clark for generating data used with permission to illustrate the ideas put forth in this review. Because it is not possible to refer to all primary references in the field of vascular development, we have referred to representative references and topic reviews where relevant. D. Darland is supported by the NHLBI and the Susan Komen Breast Cancer Foundation. P. D’Amore is a Jules and Doris Stein Research to Prevent Blindness Professor.

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4 Genetic Regulation of Preimplantation Embryo Survival Carol M. Warner1 and Carol A. Brenner 2 1

Department of Biology Northeastern University Boston, Massachusetts 02115 2

Gamete and Embryo Laboratory Institute for Reproductive Medicine and Science of Saint Barnabas Medical Center West Orange, New Jersey 07052

I. Introduction II. Preimplantation Development III. Environmental Effects on Preimplantation Embryo Survival A. Culture Media B. Exposure of Embryos to Toxic Substances C. Protection of Embryos by the Zona Pellucida IV. Genetic Effects on Preimplantation Embryo Survival A. Embryonic Genes B. Lethal Mutations C. Mitochondrial Genes D. Genomic Imprinting E. Nuclear Transfer and Reprogramming F. Telomeres and Telomerase V. Genes That Regulate Preimplantation Growth A. Growth Factors B. The Ped Gene C. Genes That Regulate Cell Cycle Processes VI. Genes That Regulate Preimplantation Death VII. Conclusions References

I. Introduction All mammals undergo a period of development between fertilization and implantation that is called the preimplantation period. Embryo survival during this preimplantation period is dependent on both environmental and genetic factors and is crucial for a successful pregnancy. The elucidation of the genes that control 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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preimplantation embryo survival is complex. Revolutionary new methods in biology, including whole genome DNA sequence analysis, detection of single nucleotide polymorphisms (SNPs) and mRNA expression with DNA chips and microarrays, cloning of animals by nuclear transfer, and creation of gene knockout mice and embryonic stem cells, should all aid in furthering the understanding of the genetic control of preimplantation embryo survival. The appearance of these new methodologies makes this a particularly timely moment at which to evaluate the status of what is known about the regulation of preimplantation embryo survival and to speculate about what the future might hold. In this review we focus on two species, the mouse and the human, with only occasional references to other species. The reasons are that the mouse is the most thoroughly studied mammalian system with respect to preimplantation development, and that the advent of assisted reproductive technologies (ART), including in vitro fertilization (IVF), has allowed access to spare human embryos for genetic studies. Because of the vast magnitude of this field and the limited space for this review, we apologize to those researchers whose work is not cited. The emphasis of this review is on particular genes that influence the growth and death of preimplantation mouse and human embryos.

II. Preimplantation Development Preimplantation mammalian development is defined as the period between fertilization of the oocyte (egg) and implantation of the embryo into the uterus (Menezo and Renard, 1993; Hogan et al., 1994; Schultz, 1999). During the preimplantation period mammalian embryos undergo a series of cleavage divisions leading to the formation of a blastocyst. Images depicting preimplantation mouse embryo development and preimplantation human embryo development from the fertilized egg to the blastocyst stage are shown in Figs. 1 and 2, respectively. The embryos are surrounded by a porous extracellular coat, the zona pellucida (Wassarman et al., 1999). Cell division in mammalian embryos is somewhat asynchronous after the two-cell stage, but the cleavage stages are referred to as “four cell,” “eight cell,” etc., even though these numbers might not reflect the exact number of cells in a particular embryo. At the 8-cell stage in mice and at approximately the 16-cell stage in humans, the embryos undergo a morphological event called “compaction,” in which the distinction between the blastomeres is lost. Next, a solid ball of cells called the morula is formed, and on further cleavage division, the morula gives rise to the blastocyst. The blastocyst is a fluid-filled, ball-shaped object consisting of two types of cells, an outer layer termed the trophectoderm (TE), and a clump of inner cells called the inner cell mass (ICM) (see Figs. 1E and 2E). Depending on the location of the TE cells with respect to the ICM, the TE cells are defined as polar TE (abutting the ICM) or mural TE (distal to the ICM). The cells of the ICM that

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Figure 1 Mouse preimplantation embryos.

154 Figure 2 Human preimplantation embryos.

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4. Genetic Regulation of Preimplantation Embryo Survival Table I Experimentally Determined Protein, RNA, and DNA Content of Preimplantation Mouse Embryosa

Stage One-cell Two-cell Eight-cell Morula Blastocyst

Hours post-hCG

Protein/embryo (ng)

RNA/embryo (ng)

DNA/embryo (pg)

18 43 71 80 90

28 26 24 21 24

0.55 0.40 0.46 Not reported 1.37

29 41 155 Not reported 439

hCG, Human chorionic gonadotropin. a Protein data from Brinster (1967); RNA and DNA data from Olds et al. (1973).

contact the blastocoelic cavity form the primitive endoderm of the embryos; the other ICM cells form the primitive ectoderm from which the primitive ectodermal and germ cells of the embryos arise. A few primitive ectoderm cells from the ICM also differentiate into some extraembryonic tissues. The rest of the extraembryonic tissues, including the placenta, arise from the TE. There is a polarity to zygotes that sets the stage for development of the ICM and the TE in the blastocyst (Gardner, 1997). The size of mouse embryos, about 100 μm in diameter, and of human embryos, about 155 μm in diameter, does not change during the preimplantation period, which is why the preimplantation cell divisions are called “cleavage divisions.” The cellular portion of mouse embryos is about 70 μm in diameter and the thickness of the zona pellucida is about 15 μm, to give a total diameter of about 100 μm; the cellular portion of human embryos is about 115 μm and the thickness of the zona pellucida is about 20 μm, to give a total diameter of about 155 μm. The protein, RNA, and DNA content of different stages of preimplantation mouse embryos are shown in Table I. Similar experimental data are not available for human

Table II Calculated Estimates of Protein, RNA, and DNA Content of Preimplantation Human Embryos

Stage One-cell Two-cell Eight-cell Morula Blastocyst

Hours post-hCG

Protein/embryo (ng)

RNA/embryo (ng)

DNA/embryo (pg)

18 42 72 96 120

124 115 106 93 106

2.4 1.8 2.0 — 6.1

128 182 691 — 1945

hCG, Human chorionic gonadotropin.

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Carol M. Warner and Carol A. Brenner Table III Characteristics of Preimplantation Embryos from a Variety of Mammalian Species

Species

Major ZGAa (cell stage)

Blastocyst formationb (days postfertilization)

Implantationb (days postfertilization)

Mouse Human Rabbit Cow Sheep Pig

2-cell 4-cell 8- to 16-cell 8-cell 8-cell 4-cell

4–5 5–6 3–4 7–8 6–7 5–6

5–6 6–7 6 22 15 13

ZGA, Zygotic genomic activation. a Data from Exley and Warner (1999b). b Data on mouse and human from Hardy (1993); data on rabbit from Davies and Hesseldahl (1971); data on cow, sheep, and pig from Bazer et al. (1993).

embryos. However, on the basis of the relative size of human embryos compared with mouse embryos one can calculate the probable amounts of protein, RNA, and DNA in human embryos. These estimated numbers are shown in Table II. Because of the small quantities of material available from preimplantation mammalian embryos, highly specialized microtechniques are often used in experimental embryo manipulation. The time between fertilization and blastocyst formation varies among species and this is summarized for a variety of species in Table III. After blastocyst formation occurs, a period of blastocyst expansion follows that ends with the “hatching” of the blastocyst from the zona pellucida. For some species, including the mouse and human, implantation occurs directly after hatching, while for other species, such as the pig, there is a period of embryo elongation after hatching and before implantation. Differences in the timing of implantation for a variety of species are also shown in Table III. The final parameter shown in Table III is the time of major zygotic genomic activation (ZGA). The unfertilized oocyte has stores of maternal mRNA that are largely degraded after fertilization. ZGA refers to the time after fertilization at which transcription of new mRNAs from the embryonic genome occurs (Exley and Warner, 1999b).

III. Environmental Effects on Preimplantation Embryo Survival A. Culture Media All phenotypes, including embryo survival, are dependent on both environmental and genetic factors. However, it is often difficult to separate environmental from

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genetic factors in ascertaining parameters involved in embryo survival because the embryos are in the milieu of the maternal oviductal or uterine fluid, depending on their stage of development. A major breakthrough in this area was the development of chemically defined culture media in which mouse embryos could be grown in vitro (Whitten and Biggers, 1968). Despite extensive efforts to improve culture media over the years, rates of in vitro development still do not match up with rates of in vivo development (Biggers, 1998; Loutradis et al., 2000). Moreover, there are changing metabolic requirements for preimplantation embryos during the course of development, so it is probably not possible to design a single ideal culture medium for the entire preimplantation period (Gardner, 1998). It has been shown that certain culture media interfere with normal gene expression. For instance, gene imprinting (preferential expression of paternally inherited vs. maternally inherited genes) is perturbed for the imprinted H19 gene when embryos are grown in certain culture media (Doherty et al., 2000). Another example of the importance of the environment to embryo health and survival is highlighted by cloning experiments in sheep and cattle that are often plagued with a phenomenon called “large offspring syndrome” (Young et al., 1998; Pennisi and Vogel, 2000). The consensus is that the poking and prodding of the embryos during the cloning procedures may be one culprit that leads to large (and often unhealthy) offspring, but the chemical composition of the culture media is also most probably another culprit. In addition, it has been hypothesized that a compromised environment of the embryo during the preimplantation period can lead to health problems later in life, such as cardiovascular disease and high blood pressure (Barker, 1995; Kwong et al., 2000). One solution to the problem of inappropriate culture conditions for preimplantation embryos would be to conduct a complete chemical analysis of the components of the oviductal and uterine fluids throughout the period of preimplantation development. This may be possible with future advances in analytical chemistry that use highly sensitive mass spectrometric. methods

B. Exposure of Embryos to Toxic Substances Embryo survival is directly affected by exposure to toxic substances (Kimmel et al., 1993). For instance, in the mouse, there is a decreased rate of preimplantation development and dose-dependent toxicity on exposure to alcohol (Cebral et al., 1999, 2000), azidothymidine (AZT) (Sieh et al., 1992; Toltzis et al., 1993; C. M. Warner, unpublished data, 2001), ethylene oxide (Polifca et al., 1996), mitomycin C (Nagao et al., 2000), and excess glucose (Moley, 1999). Exposure to excess glucose causes an increase in apoptosis in preimplantation mouse embryos; this is discussed in more detail below. For acetaminophen, different effects on preimplantation development were observed when the embryos were exposed to this compound in vitro compared with in vivo. The embryos exposed in vitro showed depressed embryo development from the morula to the blastocyst stage, but embryos exposed

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in vivo had no deleterious effects of acetaminophen exposure (Laub et al., 2000). The most likely reason is that the maternal liver is active in eliminating substances, such as acetaminophen, that could potentially harm the developing embryo. Another example of a reported deleterious effect of an exogenous compound on preimplantation mouse embryo development is the finding that exposure of two-cell mouse embryos to active cannabinoids inhibits blastocoel formation, trophoblast proliferation, and hatching from the zona pellucida (Wang et al., 1999). Also, brief exposure of embryos to N-methyl-N-nitrosourea during the preimplantation period can lead to malformations and poor fetal outcome later in the pregnancy (Bossert et al., 1990). Thus, many of the parameters described above, with respect to morphological changes in the embryo during the preimplantation period, are subject to modification by toxic substances in the environment, and the effects of these toxic substances may occur long after the period of preimplantation exposure is over. Interestingly, there is some indication that substances toxic to mouse embryos are not necessarily the same as substances toxic to human embryos. For instance, one study has shown that endotoxins are not deleterious to mouse embryos but cause embryo fragmentation and low pregnancy rates in humans undergoing IVF (Dumoulin et al., 1991). To increase the viability of human embryos cultured in vitro, coculture with fetal bovine uterine fibroblasts or bovine oviductal epithelial cells is often used. The coculture cells may metabolize toxins, thus reducing their levels, and may also provide beneficial growth factors for the embryos (Wiemer et al., 1998). Coculture results in an increased rate of cleavage, a significantly reduced rate of embryo fragmentation, and higher implantation and pregnancy rates. In addition, it is thought that the origins, effects, and control of air pollution in laboratories used for human embryo culture must be routinely assayed for aldehydes (Hall et al., 1998).

C. Protection of Embryos by the Zona Pellucida A unique feature of preimplantation embryos is the presence of the zona pellucida. The zona pellucida is responsible for species specificity of fertilization and is a secondary site to the block of polyspermy (Wassarman and Mortillo, 1991; Yanagimachi, 1994). One might initially think that the zona pellucida would provide some protection from toxic chemicals, but this is apparently not the case. The zona pellucida is composed of about 80% protein (three unique proteins called ZP1, ZP2, and ZP3) and 20% carbohydrate. It is a porous structure and is permeable to most small molecules (Turner and Horobin, 1997), to large proteins such as immunoglobulins (Sellens and Jenkinson, 1975), and even to small viruses (Wassarman, 1988). It is particularly intriguing that antibodies can cross the zona pellucida because this raises the question of how embryos are protected from cytolysis by

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autoantibodies that could potentially lyse the embryos in the presence of complement. We have shown that preimplantation mouse embryos express mRNA for FcRn (Warner and Paschetto, 2000). It is unknown whether FcRn protein is also expressed by the embryos. If FcRn protein is found to be expressed by preimplantation embryos, it is possible that any antibodies in the oviductal or uterine fluid could be bound to the embryos by the Fc region of the antibody, making the initiation of the complement cascade impossible. The whole area of protection of preimplantation embryos from complementmediated lysis is an interesting one that deserves further investigation. One study has shown that knockout of a complement regulatory gene in mice, Crry, results in fetal death midgestation (Xu et al., 2000), but no studies have yet been reported on the expression of the Crry gene in preimplantation embryos. In humans, it has been reported that complement-binding proteins are expressed on blastocysts (Taylor and Johnson, 1996), but it remains to be determined how human embryos escape complement-mediated lysis. One area that has received little attention is the possible role of the zona pellucida in protecting the embryo from attack by cells of the maternal immune system. A diagram depicting three possible types of maternal killer cells, cytotoxic T lymphocytes (CTLs), natural killer (NK) cells, and macrophages (M), and their potential interaction with embryonic target molecules, is shown in Fig. 3 (see color insert). It has been demonstrated that killing of preimplantation mouse embryos by CTLs directed to major histocompatibility complex (MHC) class Ia proteins is possible only after removal of the zona pellucida (Ewoldsen et al., 1987). The role of the zona pellucida in NK cell- and macrophage-mediated killing of preimplantation embryos is unknown. Future research aimed at defining a topological map of the proteins on the cell surface of preimplantation embryos should lead to an eventual understanding of how preimplantation embryos escape killing by the maternal immune system.

IV. Genetic Effects on Preimplantation Embryo Survival Almost any gene that is expressed in preimplantation embryos could play a role in regulating their survival. For the mouse the expression of hundreds of genes has been assayed, but because of limited availability of human embryos, the number of genes whose expression has been assayed is still quite low. The presence of genes, including particular alleles, mutations, and SNPs, can be detected by using polymerase chain reaction (PCR), or by using DNA chips or microarrays. Likewise, the presence of mRNAs for particular genes can be assayed by using reverse transcriptase (RT)-PCR, or by using DNA chips or microarrays. It should be noted that the presence of mRNA for a particular protein does not guarantee that the protein will be expressed, although the absence of mRNA for a particular protein does indicate that new protein cannot possibly be synthesized. Moreover, several

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articles have shown that there is little correlation of mRNA levels with protein levels in various types of cells (Anderson and Seilhamer, 1997; Gygi et al., 1999). Therefore, as a general principle, it is crucial to measure protein levels as well as mRNA levels to understand protein expression and function in preimplantation embryos. In this section we review the expression, in preimplantation mouse and human embryos, of a select set of genes that seem particularly relevant to development and/or have particular significance with respect to embryo survival in the ART clinic. This section includes those nuclear and mitochondrial genes that affect many aspects of embryo development and survival, but excludes those genes that regulate growth and death. These latter two types of genes will be discussed in subsequent sections.

A. Embryonic Genes Genetic effects on preimplantation embryo survival may be of maternal or embryonic origin. Of particular interest to this review are genes that are expressed by the embryos themselves independent of the maternal environment. To this end, DNA chip and microarray technology should allow the complete description of all genes that are expressed during each stage of development. The first study of this type has been reported by Ko et al. (2000). They found that preimplantation mouse embryos expressed mRNAs for more than 9000 genes and that the pattern of gene expression changed throughout development. These results confirmed the idea that there is a genetically programmed sequence of events in which different genes are turned on and off during the preimplantation period. When the entire mouse genome has been sequenced (expected by the time this article is published), it will be possible to screen all of the genes (estimated at 30–100,000) to determine which ones are transcribed throughout preimplantation development. Since the complete human DNA genomic sequence has been reported, this is already possible with human embryos, and preliminary studies along these lines have been initiated in several laboratories (Brenner and Cohen, 2000). The analysis of the gene expression data to be generated is quite complex and will need sophisticated approaches from the field of bioinformatics to yield information that is biologically useful. A major challenge will be to move from genomics to proteomics to identify functions for each of the proteins that arises from mRNA transcribed during the preimplantation period of development. Simultaneous with the global approach of the genomics/proteomics revolution, studies of individual genes that affect preimplantation embryo survival are being conducted by a number of laboratories. As described previously, there are several morphological events during the preimplantation period of development that are candidates for genes that mediate preimplantation survival. These can be broken down into genes that regulate ZGA, genes that regulate compaction, genes that regulate blastocyst formation, and genes that regulate blastocyst elongation. The

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last category of genes is not discussed in detail since neither mouse nor human preimplantation embryos undergo an elongation phase before implantation. However, it is noteworthy that a study of gene expression during elongation of the preimplantation pig embryo has been published (Wilson et al., 2000). Examples of genes that influence ZGA, compaction, and blastocyst formation are given in the next sections. 1. Genes That Influence Zygotic Genomic Activation As shown in Table III and mentioned previously, the time of ZGA varies among species. In the mouse, a limited amount of transcription occurs in zygotes, but the major onset of ZGA is at the two-cell stage. Several genes that are involved in mouse ZGA have been described, including eIF-1A and transcription-requiring complex (TRC) (Schultz et al., 1999), as well as GAGA box-binding factor and Sp1 (Bavilacqua et al., 2000). ZGA is related to nuclear reprogramming, which is particularly relevant to cloning from adult cells, a subject discussed later in this article. In human preimplantation embryos a dramatic reprogramming of gene expression occurs from the four- to the eight-cell stage (Braude et al., 1988). More precise estimates of the timing of ZGA depend on the detection of de novo transcripts from the zygotic genome, which varies according to the sensitivity of the techniques used and the specific genes analyzed. Braude et al. (1988) have hypothesized that reprogramming of gene expression in the human preimplantation embryo mostly likely involves DNA replication and chromatin remodeling. Although the gene expression of eIF-1A and TRC has not been assayed in human embryos, paternal transcripts for the Y-linked genes ZFY and SRY, as well as the myotonic dystrophy-associated protein kinase gene DK, have been detected as early as the late pronuclear, one-cell stage (Pergament and Fiddler, 1998). Furthermore, since the high incidence of human embryonic arrest coincides with the transition from maternal to embryonic regulation of development, it has been proposed that the failure of ZGA is responsible. 2. Genes That Influence Compaction In the mouse, several genes have been described that influence compaction. Compaction is basically a cell adhesion event that is regulated by the expression of a series of cell surface proteins including E-cadherin (also known as uvomorulin) (Kemler et al., 1977; Vestweber et al., 1985). Other proteins involved in the adhesion process are the catenins and the actin cytoskeleton and the interaction of these proteins is now well documented (Nieset et al., 1997). Compaction is known to be controlled at the posttranslational level since exposure to protein synthesis inhibitors does not impede compaction (Kidder and McLachlin, 1985; Levy et al., 1986). It is still not clear exactly what the posttranslational modifications are that control compaction, although it is thought that phosphorylation events mediated by

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phosphatases and/or kinases are largely responsible (Goval and Alexandre, 2000). One of these, protein kinase C (PKC), is definitely involved in the regulation of compaction. It has been shown that seven isotypes of PKC are expressed by preimplantation mouse embryos, and that a key redistribution of the PKC isotypes occurs at the time of compaction (Pauken and Capco, 2000). Thus, quite a bit is known about the genetic regulation of compaction in preimplantation mouse embryos, but complete elucidation of all the genes involved is yet to come. Compaction of cleaving human preimplantation embryos changes them from a collection of individual cells into a solid mass with indistinguishable cell membranes. As in the mouse, compaction is due to the formation of tight junctions causing the blastomeres to become closely apposed. The positioning increases the extent of contact between the blastomeres that is essential for subsequent development. In humans this usually occurs at about the 8- to 16-cell stage (Nikas et al., 1996). Although compaction has been observed at earlier stages, it is not known whether this represents a normal occurrence. When compaction takes place, the cells lose their totipotency as the result of cell–cell interactions and it is believed that the onset of compaction marks the beginning of major ZGA in human embryos. During compaction a number of cellular changes occur within the embryo that ensure the subsequent establishment of the trophectoderm. These changes include the formation of tight junctions, gap junctions, and cytoskeletal connections and reorganization between blastomeres. Prior to implantation, the human embryo expresses connexin-containing gap junctions (Hardy et al., 1996). Desmosomes appear between outer cells prior to cavitation and are retained in the trophectoderm of the late blastocyst. Other relevant cell adhesion molecules active during compaction in human preimplantation embryos still remain to be elucidated. 3. Genes That Influence Blastocyst Formation After compaction, the next major morphological landmark in preimplantation development is blastocyst formation. It is generally agreed that the major control of blastocyst formation is by the plasma membrane sodium pump, Na+/K+-ATPase (Watson, 1992). Studies of bovine embryos have shown that disruption of Na+/K+ATPase gene expression by antisense oligonucleotides abolishes blastocyst formation (Watson et al., 1999). In the mouse there are several isozymes of Na+/K+ATPase, and similar to the observation reported above on the role of different isozymes of PKC in compaction, there are probably differential roles for as many as six isozymes of Na+/K+-ATPase in blastocyst formation (MacPhee et al., 2000). There are, undoubtedly, multiple genes other than Na+/K+-ATPase that are involved in blastocyst formation. For instance, it has been reported that there is an association between insulin-like growth factor (IGF-I) expression and blastocyst formation in mouse embryos (Kowalik et al., 1999). In humans, IGF-I not only enhances growth of preimplantation embryos, but also reduces apoptotic cell death (Spanos et al., 2000). Another report has suggested that the splicing factor SRp20

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is involved in mouse blastocyst formation (Jumaa et al., 1999). In this study the SRp20 gene was knocked out by using the Cre–loxP system and it was found that development in the resulting mice was blocked at the morula stage. Tight junction assembly during blastocyst formation is also under genetic control in the mouse (Sheth et al., 1997, 2000). In humans, approximately 24 h after the morula has formed and compaction has taken place, the intercellular spaces begin to enlarge to create the central fluid-filled cavity, the blastocoel. As in the mouse, the human blastocyst consists of ICM and TE cells (Figs. 1 and 2). Expanded human blastocysts have a total cell number of 60 cells on day 5, which increases to 80 and 125 on days 6 and 7, respectively (Gardner and Schoolcraft, 1999). (This compares with about 32 cells in day 4 mouse blastocysts.) The proportion of ICM cells in human blastocysts ranges from 33 to 50% between days 5 and 7. It has also been observed that although the human blastocyst expands readily in vitro, about 20% of such blastocysts have hatching problems. Rescue of these blastocysts can be accomplished by assistedhatching (AHA), using acidic Tyrode’s solution to create a gap in the zona pellucida before intrauterine transfer (Cohen et al., 1991). The AHA technique is also used to save blastocysts trapped in overtly thick zonae pellucidae. Selective embryo hatching protocols in the IVF clinic have promoted both higher implantation rates and pregnancy outcomes for infertile women. There are multiple genes that affect human blastocyst development, including genes that encode proteins that can be characterized as growth factors and their receptors, gene regulators, and transcription factors. Some examples are the transcription regulators OCT4 and OCT6 (Abdel-Rahman et al., 1995; Ben-Shushan et al., 1998; Hansis et al., 2000), a cell surface glycoprotein CD44, insulin growth factors and their receptors, heparin-binding epidermal growth factor (HB-EGF) and its associated EGF receptor, leukemia inhibitory factor (LIF) and its receptor, as well as matrix metalloproteinases (MMPs), which are crucial proteases for the implantation process (Brenner et al., 1989; Pergament and Fiddler, 1998). Implantation rates, although increasing in the IVF clinic, range from 5 to 30% and repeated implantation failures occur commonly in clinical practices. One suggestion to improve pregnancy rates has been to transfer blastocysts on day 5, rather than eight-cell embryos on day 3, to recipient mothers. In this way any embryos with aberrant gene expression leading to embryonic arrest would be eliminated and the resulting blastocysts would presumably have a better chance of leading to a successful pregnancy. Thus, a full understanding of the genes that regulate blastocyst formation in human embryos has potential clinical importance.

B. Lethal Mutations Mutations that arrest preimplantation development are scarce compared with mutations that affect postimplantation development (Magnuson et al., 1993). The

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reasons for this are not clear, but it is possible that stores of maternal mRNAs and proteins in the oocyte can provide a boost that allows embryos to develop to the blastocyst stage in spite of potentially lethal mutations. Another possibility is that major genes regulating preimplantation development have other genes with redundant functions capable of substituting for them, so that a point mutation in one particular gene would not necessarily be lethal during the preimplantation period. A few preimplantation embryonic lethal mutations that have been described in the mouse are t12 (located in the T/t complex on chromosome 17), which is lethal at the morula stage (Cheng et al., 1983); pin (located in the albino complex on chromosome 7), which arrests cell division some time between the two- and six-cell stages and causes embryo death 1–2 days later (Magnuson et al., 1993); and mpg or dist1 (located near the α-globin complex on chromosome 11), which causes abnormal development of both the ICM and TE in blastocysts, leading to periimplantation death (Hendrey et al., 1995). A gene encoding a zinc finger transcription factor, Wt1, has been found to have an indirect effect on preimplantation embryo survival by controlling the oviductal environment in conjunction with a modifier oviductal protein that has not yet been identified (Kreidberg et al., 1999). Another gene, Traube (Trb), halts mouse preimplantation development at the compacted morula stage, presumably by interfering with the synthesis of ribosomes (Thomas et al., 2000). So far, in human preimplantation embryos no lethal mutations have been described.

C. Mitochondrial Genes Cellular genes are found not only in the nucleus, but also in cytoplasmic mitochondria. It has been ascertained, using highly sensitive quantitative PCR techniques, that the mouse oocyte contains a mean copy number of about 150,000 mitochondria compared with about 300,000 in the human oocyte (Steuerwald et al., 2000a). Thus, there is a significant amount of mitochondrial DNA (mtDNA) in oocytes and embryos that may contribute to genetic regulation of preimplantation embryo survival. 1. Mitochondrial Mutations There are now more than 150 known mtDNA rearrangements, including deletions, insertions, and duplications (Wallace, 1993). Such mutations in mtDNA are responsible for a number of catastrophic neuromuscular diseases, such as Kearns– Sayre syndrome (KSS), chronic progressive external ophthalmoplegia (CPEO), and Pearson’s syndrome. Mitochondrial DNA rearrangements have been shown to accumulate with age, and prevalently appear in postmitotic, nondividing tissues (Cortopassi and Arnheim, 1990). When aging tissues accumulate mtDNA rearrangements, and it reaches a significant threshold level, a reduction in the

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efficiency of oxidative phosphorylation occurs. Since the oocyte is a nondividing tissue, which may be subject to meiotic arrest in the ovary for up to 50 years, there must be sufficient copies of the mitochondrial genome to allow the mammalian preimplantation embryo to develop and implant. It is postulated that mitochondrial replication occurs initially during oocyte maturation and then not again until after the egg cylinder stage. Therefore a high frequency of abnormal mitochondria in the oocyte could reduce the number of functional mitochondria leading to embryonic arrest, failed implantation, or mitochondrial disease. In the future, mice carrying mtDNA with pathogenic mutations may provide a system with which to study how the mutant mtDNAs are transmitted and distributed in tissues, resulting in expression of mitochondrial diseases. Mouse cells without mitochondria have been isolated and utilized to trap mtDNAs with somatic mutations (cybrids). Mice can now be generated to model mtDNA disease by electrofusion of fertilized mouse eggs with enucleated cybrids (Inoue et al., 2000). These mice should be useful for studying the precise mechanisms of formation, transmission, and pathogenic expression of duplicated mtDNA and the variation of mtDNA in specific tissues and thus could be models of mtDNA-based diseases. Multiple laboratories have detected a particular mtDNA mutation called the “common deletion,” mtDNA4977, in human oocytes (Chen et al., 1995; Keefe et al., 1995; Brenner et al., 1998; Barritt et al., 1999). The mtDNA4977 mutation can be detected at a frequency of 30 to 50% in human oocytes. Although there is no age-related accumulation of this mutation, there is a significant reduction of mtDNA4977 detected in embryos compared with oocytes (Brenner et al., 1998; Barritt et al., 1999). These findings suggest that a selection mechanism of some type may be working on the oocyte and early preimplantation embryo to reduce or eliminate the inheritance of mitochondrial mutations. In addition, 23 novel mtDNA rearrangements have been identified in human oocytes and embryos (Barritt et al., 1999). Using a nested PCR strategy, 51% of human oocytes and 32% of embryos exhibited mtDNA rearrangements. Multiple rearrangements were detected in 31% of oocytes and 14% of embryos. Thus, a significant reduction in both mtDNA rearrangements and multiple mtDNA rearrangements is found in embryos compared with oocytes. A mtDNA point mutation was discovered that is predominantly present in oocytes from women of advanced age (Barritt et al., 2000a,b). Interestingly, this mtDNA mutation affects the mitochondrial control region responsible for transcription and replication regulation. This mutation represents a single base pair transversion of a thymine (T) to guanine (G) at base pair 414 (T414G) in the mitochondrial genome. In women <37 years of age the frequency of this oocyte mutation was 4% compared with 40% in women older than 37 years of age. The link between mitochondrial mutations, functional impairment of the aging oocyte, and the regulation of mitochondrial replication still remains an unsolved mystery. An interesting proposition is that it should be possible to cure mitochondrial disease by nuclear transfer methods similar to cloning methods discussed below.

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2. Cytoplasmic Transplantation Reproductive biologists in IVF clinics have utilized cloning manipulations and techniques to solve many different problems. Transplantation of ooplasm from healthy donor oocytes into oocytes from patients with recurrent implantation failure after ART has led to the birth of healthy babies (Cohen et al., 1998, 1999). The volume of ooplasm transferred, 5–15% of the total ooplasm volume, almost certainly involves the transfer of mitochondria as well as mRNAs, proteins, and other factors. It is not known precisely how ooplasmic transfer affects the physiology of the early human embryo, but it appears that the introduction of a small amount of ooplasm from a donor oocyte may correct certain as yet unspecified ooplasm deficiencies. The clinical experience gained so far supports the idea that the procedure may be useful when conventional ART has repeatedly and consistently failed and when cytoplasmic deficiency of the oocytes is the suspected cause of poor embryo viability. 3. Mitochondrial DNA Heteroplasmy The infusion of mitochondria from a different source may lead to heteroplasmy, that is, the presence of more than one population of mtDNA in a single cell, and the heteroplasmy may persist if the transferred mitochondria survive and replicate throughout embryonic and fetal development. By examining mtDNA from the donor and recipient it is possible to distinguish differences in the mtDNA hypervariable region or the mtDNA fingerprint. Ooplasmic transfer can result in sustained mtDNA heteroplasmy representing both donor and recipient. This was demonstrated by mtDNA fingerprinting of embryos, amniocytes, fetal placenta, and cord blood. These results show that the donor-derived mitochondrial population persists after ooplasmic transfer and may be replicated during fetal development (Brenner et al., 2000). In addition, two mtDNA populations have been found in blood samples from healthy 1-year-old children resulting from ooplasmic transplantation (Barritt et al., 2001). These mtDNA fingerprints demonstrate that the transferred mitochondria can be replicated and maintained in the offspring, without potentially altering mitochondrial function. However, the mechanisms of regulation of the donor’s mitochondrial population in the presence of the recipient’s mitochondrial and nuclear genomes are unknown. These heteroplasmic children will be monitored for the foreseeable future to determine whether the heteroplasmy persists throughout life.

D. Genomic Imprinting Genomic imprinting is the epigenetic mechanism that distinguishes whether genes that are inherited from the maternal versus the paternal genome result in parentspecific gene expression. Genomic imprinting is important in the development and

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survival of mammalian embryos. Genomic imprinting is established according to specific markers that are imposed on the genome during gametogenesis; the allelespecific gene expression is then maintained throughout embryogenesis and is an absolute prerequisite for normal development. The pioneering work of McGrath and Solter (1984) and Surani et al. (1984) enabled researchers to reject lack of a cytoplasmic component from the fertilizing male gamete or homozygosity for recessive lethal alleles as explanations for the abnormal development of uniparental haploid and diploid murine embryos, in favor of parental nonequivalence (i.e., genomic imprinting). In these experiments, nuclear transplantation was used to create embryos with differing parental contributions from inbred mice. In common with haploid parthenotes activated by fertilization with a spermatozoon, uniparental diploid gynogenotes (two maternal nuclei) and androgenotes (two paternal nuclei) also developed to the blastocyst stage but no further. These and other experiments confirmed that the parental genetic contributions were somehow imprinted, distinguishing paternal and maternal components. Imprinting of a parental allele is established during gametogenesis and is intimately associated with the methylation of cytidines located within dinucleotide CG repeating sequences, commonly referred to as CpG islands, which lie outside the coding regions of the gene. We now understand that the paternal genome is more important in the development of extraembryonic tissues, whereas the maternal genome is more closely associated with postimplantation embryonic development. It is thought that there are more than 30 genes that are marked during sperm and egg formation that are selectively switched off in the embryo. Understanding the molecular basis for the parental specific expression of Igf 2 is of long-standing interest in the field of imprinting. Igf 2 is part of a cluster of imprinted genes whose organization is well conserved in mice and humans. The paternal-specific Igf 2 gene and its neighbor, the maternal-specific H19 gene, are coregulated since they share enhancers. Dissecting monoallelic expression pathways will therefore contribute toward an understanding of normal gene regulation and of the molecular basis for diseases associated with disregulation of the imprinted loci.

E. Nuclear Transfer and Reprogramming It has become possible to make exact genetic copies (clones) of sheep, cows, goats, monkeys, and mice. It is also possible to utilize techniques such as cytoplasmic and nuclear transplantation in human reproduction. With the development of these new technologies, patterns of gene expression may be examined as well as manipulated. The pattern of gene expression in adult cells is different from that in embryonic cells. Some genes are expressed in adult cells but not in embryonic cells and vice versa. When embryos are analyzed a few hours after the transfer of an adult cell nucleus to an enucleated metaphase II egg, the pattern of gene

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expression cannot be distinguished from that in embryos grown from normally fertilized eggs. This means that the egg cytoplasm causes a dramatic switch in gene expression in the transferred nucleus in only a few hours. This switch is called nuclear reprogramming. 1. Nuclear Transfer and Cloning In mammals, normal fertilization begins with the union of the egg and the sperm. The unfertilized oocyte is stopped at metaphase II until the sperm can provide an activation signal that triggers the resumption and the completion of cell division. Nuclear transfer subverts fertilization by replacing the female genetic material from the unfertilized egg with the nucleus from a different somatic or embryonic cell. This was first done successfully on frogs. Nuclear transplantation in mammals has proved more difficult than in frogs. Although it is easier to clone mammals from embryonic or fetal cells, it is now possible to use adult nuclei for the cloning procedure. The cloning of the first mammal from an adult cell, Dolly the sheep, has been followed by successful cloning from an adult cell of cows, mice, and pigs (reviewed in Gurdon and Colman, 1999; Pennisi and Vogel, 2000). 2. Gene Reprogramming Reprogramming normal developmental genes after nuclear transplantation requires approximately 30 genes, which are actively expressed by both the sperm and the oocyte during meiotic maturation. These genes are switched off in the embryo until after ZGA. Imprinted genes are unlikely to be reprogrammed by nuclear transfer because if they were, the embryos would not survive. Another focus of reprogramming is the inactivation of one X chromosome in female mammals (Clerc and Avner, 2000; Eggan et al., 2000). During early development of female mammals, one of the two X chromosomes is randomly inactivated in those tissues contributing to the fetus. Gene reprogramming in mammals seems to occur on the same genes active before nuclear transfer, and does not require the formation of new DNA (Surani, 1998). Key molecules found in the egg, such as telomerase and embryo-specific histones, may be involved in the regulation of reprogramming. 3. Nuclear Transfer at the Germinal Vesicle Stage Nuclear transfer may be used for other purposes than cloning. It may be possible to use transfer of nuclei to germinal vesicle (GV) stage oocytes to reduce the incidence of aneuploidy. The proper segregation of chromosomes is a fundamental prerequisite for the orderly completion of cell division. Inappropriate chromosome separation can result in the production of aneuploid cells and abnormal gametes. While aneuploidy in somatic cells is likely responsible for the development of various cancers, the incidence of aneuploidy in gametes contributes to birth defects

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and pregnancy loss. It is now well established that nondisjunction of bivalent chromosomes during oogenesis, and therefore chromosomal abnormalities, increases significantly with advancing maternal age. The risk of conceiving a chromosomally abnormal fetus during IVF increases from 6.8% for women 35–39 years old to ∼50% in women 45 years of age or older (Liu, et al., 1999; Takeuchi et al., 1999; Zhang et al., 1999). However, it is not known what molecular mechanisms orchestrate chromosome distribution during human oocyte maturation (Steuerwald et al., 2001). Studies have explored whether nuclear transplantation at the GV stage can reverse the effects of aneuploidy (Liu et al., 1999). A GV is removed with a small amount of cytoplasm from an older woman and transferred into an enucleated oocyte from a young woman. Such reconstituted oocytes are placed in culture and allowed to mature so that the extruded first polar bodies can be processed for chromosomal analysis. Many laboratories believe that a complete haploid set of chromosomes can be obtained from the first polar body. Although more extensive research is necessary before GV transfer is used as a therapeutic technique for rescuing genomes from the maternal age-related increase in chromosomal nondisjunction during the first meiotic division in human oocytes, this technique seems promising. However, if the rescue is to be applied to reconstructed mammalian oocytes, they must be capable of fertilization and subsequent embryonic growth. The reprogramming of development genes in the mouse and human oocyte and embryo must be examined to ensure normal human development before this technique can be safely and ethically implemented.

F. Telomeres and Telomerase One of the more intriguing questions in the area of biology is whether mechanisms are already programmed in our genes to determine our life span. Is such a genetic program already determined at birth? Or even prior to birth in the oocyte itself? It is now believed that telomere shortening is a major event in biological aging. Telomeres are repeats of DNA protecting the stability of chromosomes and are essential for chromosome end maintenance. The ribonucleoprotein enzyme that adds these DNA repeats to the ends or tails of the chromosomes is called telomerase. The telomerase catalytic subunit (TCS) is structurally related to reverse transcriptase and thus represents the first member of this family with essential cellular function (Nakamura et al., 1997). Interestingly, telomerase activity is high in germ, embryonic, and cancer cells. All of these are considered telomerase positive. The enzyme is missing in telomerase-negative cells, such as most somatic cells. Immortal cancer cells divide uncontrollably and are telomerase positive. The mechanisms that convert somatic cells to cancer cells are unknown, but may well involve telomerase reactivation. Reactivation of telomerase results in the unscheduled additions of TTAGGG repeats that normally cap human chromosome ends. Cells with elongated telomeres show a spectacular alteration in their growth potential.

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It is now evident that human oocytes and embryos have high levels of telomerase. The oocyte is the most likely place for the setting of the telomeric clock, since telomeres should be added to the chromosomes prior to fertilization and before the onset of ZGA and blastocyst formation. Logically, mammalian spermatozoa and oocytes should have the longest telomeres to ensure the transmission of full-length chromosomes to the progeny (Kozik et al., 1998). Human oocytes and embryos indeed express the human telomerase catalytic subunit (hTCS). Surprisingly, multiple transcripts of hTCS were also found at different meiotic stages and in embryos (Brenner et al., 1999). Potentially normal donated oocytes did not reveal any alternately spliced variants compared with developmentally abnormal human oocytes and embryos. In early human development the expression of the enzyme may have at least two regulatory mechanisms controlling the activity of telomerase: transcriptional control of the hTCS gene and alternate splicing of its transcripts. Clearly differing telomerase activity in individual mammalian oocytes and embryos may serve as a marker for embryonic health and may even predict life span. The correlation between telomerase activity, telomere length, and cellular replicative capacity mechanisms in oocytes and embryos remains to be fully explored and understood. When researchers announced in 1997 that they had cloned the sheep, Dolly, many scientists asked the question: Are her cells older than she is? Dolly was cloned from an adult cell and everyone wondered whether her own cells would show some of the hallmarks of an older animal. It transpired that Dolly’s telomeres were shorter than normal (Shiels et al., 1999), and because telomeres normally shrink with age, this was a disturbing sign that her cellular clock had not been reset. Mice have also been cloned by nuclear transfer into enucleated oocytes and bred for six generations. Successive generations of these mice showed no signs of premature aging. There was also no evidence of shortening of the telomeres, and in fact there apparently was a slight increase in telomere length (Wakayama et al., 2000). In cattle cloning, transferred nuclei from nearly 100 cultured cells into enucleated eggs eventually produced six cattle. When the blood cells from the young cattle were analyzed, it was found that the calves’ telomeres were at least as long as the telomeres of normal cattle the same age and in some cases even longer than the telomeres of the normal cattle (Lanza et al., 2000). Nuclear transfer from adult somatic cells has produced cloned pigs; however, at this time their telomere length has not been reported (Polejaeva et al., 2000). Thus, telomere length after cloning seems to vary among species and may have an impact on life span.

V. Genes That Regulate Preimplantation Growth Growth during the preimplantation period is one of the most important parameters involved in the regulation of embryo survival. An increased number of cells in the ICM of blastocysts is associated with a greater chance of subsequent fetal survival (Tam, 1988; Brison and Schultz, 1996; Lane and Gardner, 1997; Van Soom

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et al., 1997; Chan et al., 2000), and growth during the preimplantation period is important for synchronization of the embryo with the maternal reproductive tract so that successful implantation can occur (Rogers and Leeton, 2000). Moreover, as pointed out previously, slow growth during the preimplantation period has been shown to be associated with impaired cardiovascular function and high blood pressure later in life (Barker, 1995; Kwong et al., 2000). A. Growth Factors Growth factor receptors are present on preimplantation embryos (Kaye, 1997). The sources of the growth factors that could potentially interact with these growth factor receptors are from the environment, the mother, or the embryos themselves. The latter two sources of growth factors are referred to as paracrine and autocrine, respectively. A diagram of the possible interactions of growth factors with preimplantation embryos is shown in Fig. 4 (see color insert). Effects of various exogenous substances, such as vitamins, oxygen, and lactate, on preimplantation growth have been reported (McKiernan and Bavister, 2000; Trimarchi et al., 2000a,b; Lane and Gardner, 2000). It should be noted that there is no requirement for exogenous growth factors in media used to culture preimplantation embryos in vitro in order to obtain live offspring. However, the embryos themselves produce autocrine growth factors that are required for survival (Lane and Gardner, 1992). Quite a number of studies have appeared defining growth factors produced by preimplantation embryos. These autocrine growth factors include interleukin 1 (IL-1), interleukin 6 (IL-6), interleukin 10 (IL-10), colony-stimulating factor (CSF), leukemia inhibitory factor (LIF), transforming growth factor (TGF), epidermal growth factor (EGF), interferon γ (IFN-γ ), gonadotrophin releasing hormone (GnRH), vascular endothelial growth factor (VEGF) (reviewed in Krussel et al., 2000), as well as platelet-activating factor (PAF) (O’Neill, 1998; Emerson et al., 2000), early pregnancy factor (EPF) (Athanasas-Platsis et al., 2000), and acrogranin (Diaz-Cueto et al., 2000). This list will undoubtedly be expanded as the genomics/proteomics revolution proceeds. B. The Ped Gene 1. The Mouse Ped Gene One particularly interesting gene that regulates preimplantation embryonic growth is the Ped gene discovered in our laboratory (Verbanac and Warner, 1981). The properties of the Ped gene have been reviewed by ourselves (Warner et al., 1998a–c) and by others (Fernandez et al., 1999; Gill, 1999). We discovered the Ped gene when we observed that the rate of preimplantation development varied among inbred strains of mice. Although a few other workers had reported apparent genetic differences in the rate of development of preimplantation mouse embryos (Whitten and Dagg, 1962; McLaren and Bowman, 1973; Titenko, 1977), our key

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observation was that this difference was correlated with the major histocompatibility complex (MHC) haplotype. This led us to propose that there is a gene, Ped (Preimplantation embryo development), in the MHC that influences the rate of cleavage division of early embryos. We defined two functional alleles of the Ped gene, fast and slow, on the basis of the number of cells per embryo at 89 h after human chorionic gonadotropin (hCG) injection (Verbanac and Warner, 1981; Goldbard et al., 1982b), and showed that the fast Ped allele is dominant ( Goldbard et al., 1982a). We also showed that fast and slow development of mouse embryos is maintained in vitro independent of the maternal uterine environment (Brownell and Warner, 1988). Thus, fast and slow development of preimplantation mouse embryos is an intrinsic genetic property of the embryos themselves. In our next set of studies we located the Ped gene to the Q region of the mouse MHC (Warner et al., 1987, 1988, 1991). We were then able to show that Qa-2 protein, the product of two almost identical (one nucleotide difference) Q region genes, Q7 and Q9, was responsible for the Ped gene phenotype (Xu et al., 1994; Wu et al., 1999). Those embryos that express Qa-2 protein cleave at a fast rate whereas those embryos that are missing Qa-2 protein cleave at a slow rate. Embryos missing Qa-2 protein have a deletion of both the Q7 and Q9 genes. Figure 5 (see color insert) shows images of embryos with the Ped fast and Ped slow alleles and also shows that Qa-2 is present on the cell surface of Ped fast embryos, but missing from the surface of Ped slow embryos. It is noteworthy that Qa-2 present on the embryos is below the level of detectability by immunofluorescence, but can be detected by the highly sensitive method of Immuno-PCR (McElhinny and Warner, 1997; Ke and Warner, 2000). The mystery is to determine how a protein on the cell surface can transmit a signal to the embryos to cleave at a faster rate than those embryos that are missing the protein. There are additional functions of the Ped gene beyond the preimplantation period of development. This observation complements many of the studies cited above, in which it was pointed out that rate of growth during the preimplantation period affects health later in life. We have shown that survival to birth, birth weight, and weaning weight are all influenced by the Ped gene (Warner et al., 1991, 1993; Exley and Warner, 1999a). Those embryos with the Ped fast allele have a higher chance of their resulting pups surviving to term, and the pups have a higher birth weight and weaning weight compared with those pups arising from embryos with the Ped slow allele. Interestingly, both Ped fast and Ped slow embryos have an equal chance of surviving to midgestation; fetal loss occurs between midgestation and birth (Exley and Warner, 1999a). Thus, the Ped gene does not seem to influence whether implantation occurs, but rather the effect is on fetal survival later in pregnancy. Since embryos with the Ped fast allele have a higher number of cells both in their trophectoderm (TE) and inner cell mass (ICM) (McElhinny et al., 1998), this may lead to better vascularized implantation sites, better placentation, and enhanced fetal development even though implantation rates are similar for Ped fast and Ped slow embryos. There are many possibilities to explain

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fetal loss of Qa-2-negative fetuses between midgestation and birth. These include a possible protective role of Qa-2 in warding off attack by maternal NK cells, CTLs, or macrophages, similar to the mechanisms illustrated previously in Fig. 3 (see color plates). It is also possible that the Ped gene influences growth and health throughout adult life; life span has been suggested to be affected by the Ped gene (Tarin, 1997), but this suggestion has not been tested experimentally. The Ped gene is a good example of how growth during the preimplantation period influences subsequent embryo survival and health later in life. There are other examples of this same phenomenon. For instance, knocking out the gene for alkaline phosphatase (EAP) results in slower development during the preimplantation period, a longer gestation time, and smaller litter size, similar to the effects described above for the Ped gene (Dehghani et al., 2000). Thus, certain genes can be deleted from the mouse genome (e.g., Q7, Q9, and EAP) with the result that reproductive capacity of the embryos is diminished, but not completely abolished. 2. A Human Homolog of the Mouse Ped Gene Since the mouse Ped gene has such an important influence on overall reproductive success, it would be most interesting to identify a similar gene in humans. The Ped gene phenotype, a range of developmental stages among embryos fertilized at the same time coupled to preferential survival of the faster developing embryos, exists in the human population, as well as in other animal species including mice, rats, hamsters, pigs, cows, and monkeys (Wilmut et al., 1986; Bazer et al., 1988; McKiernan and Bavister, 1994; Gonzales et al., 1995; Warner et al., 1998a–c; Cohen et al., 1999; Edwards and Beard, 1999). Thousands of slow-developing human embryos have been carefully checked for dozens of morphological criteria and chromosomal abnormalities, with the conclusion that a large proportion of slow-developing embryos are perfectly healthy and can give rise to healthy offspring, although there is a greater chance of obtaining a successful pregnancy when the faster developing embryos are used for embryo transfer (Buster et al., 1985; Cummins et al., 1986; Claman et al., 1987; Puissant et al., 1987; Clark, 1988; Bolton et al., 1989; Levy et al., 1991; Trounson and Osborn, 1993; Warner et al., 1998a; Trounson and Gardner, 2000). Figure 6 shows a picture of two healthy human embryos fertilized by IVF at the same time, but showing different rates of development 3 and 4 days later. Thus, the two major hallmarks of the mouse Ped gene phenotype, a range of developmental stages among preimplantation embryos fertilized at the same time and preferential survival of the faster developing embryos, is present in the human population. The challenge is to identify the genes that regulate rate of development and fetal survival in humans. Data suggest that human HLA-G is the functional homolog of mouse Qa-2, the Ped gene product (Allcock et al., 2000). This conclusion is based on the analysis of the complete DNA sequence of the human MHC, the HLA complex (Beck et al., 1999; Beck and Trowsdale, 2000), along with new findings about the expression

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Figure 6 Fast- and slow-developing human embryos.

and function of HLA-G (reviewed in Le Bouteiller and Blaschitz, 1999). Although Qa-2 and HLA-G are not strict genetic orthologs, they are believed to be functional homologs as the result of convergent evolution (Allcock et al., 2000). Table IV shows a comparison of many of the features of Qa-2 and HLA-G (based on the following articles and references cited within: Cai et al., 1996; Stroynowski and Tabaczewski, 1996; Manilay and Sykes, 1998; O’Callaghan and Bell, 1998; Warner et al., 1998a–c; Braud et al., 1999; Cao et al., 1999; Fernandez et al., 1999; Hiby et al., 1999; Le Bouteiller and Blaschitz, 1999; Munz et al., 1999; Ke and Warner, 2000; McElhinny and Warner, 2000; Morales et al., 2000). Table IV shows that the vast majority of properties of Qa-2 and HLA-G are similar, with only one major difference. This difference is that Qa-2 is attached to the cell membrane by a glycosylphosphatidylinositol (GPI) linkage whereas HLA-G is simply inserted into the cell membrane by a short (six-amino acid) tail. We originally thought that this might have important functional significance (Cao

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Table IV Comparison of HLA-G and Qa-2, the Ped Gene Product Feature MHC class Ib molecule DNA sequence Membrane-bound and soluble forms Attached to membrane by a short tail GPI linkage of short tail to membrane Limited tissue expression Expression by preimplantation embryos Expression by placenta Nonapeptides bound Amino acid frequency Peptide sequences Same peptides in membrane and soluble forms Cell surface expression dependent on TAP Increased expression with IFN-γ Increased cell proliferation with cross-linking Deletion polymorphism compatible with embryo survival Interacts with T cell receptor (TCR) Interacts with CD8 (on T cells) Interacts with NK cell receptors Interacts with other accessory proteins Acts as a signal transduction molecule Effect on Ped gene phenotype Increases preimplantation growth rate Enhances fetal survival Increases birth weight Increases weaning weight

Qa-2

HLA-G

Yes Similar to HLA-G Yes Yes Yes Yes Yes Yes

Yes Similar to Qa-2 Yes Yes No Yes Yes Yes

Similar to HLA-G Unknown Unknown Yes Yes Yes Yes

Similar to Qa-2 Self-peptides Yes Yes Yes Unknown Yes

Yes Yes Unknown Probable Probable

Yes Yes Yes Unknown Unknown

Yes Yes Yes Yes

Yes Yes Unknown Unknown

a

Based on Cai et al. (1996), Stroynowski and Tabaczewski (1996), Manilay and Sykes (1998), Warner et al. (1998a–c), O’Callaghan and Bell (1998), Braud et al. (1999), Cao et al. (1999), Fernandez et al. (1999), Hiby et al. (1999), Le Bouteiller and Blaschitz (1999), Munz et al. (1999), Ke and Warner (2000), McElhinny and Warner (2000), and Morales et al. (2000).

et al., 1999), but in further researching this finding we discovered that a short tail (six amino acids, as is found in HLA-G) confers many of the same properties to proteins as a GPI linkage: increased lateral mobility in the membrane, higher cell surface half-life, and a requirement for accessory proteins to transmit a signal across the membrane (Medof et al., 1996; Davis et al., 1997). We therefore now believe that the GPI linkage of Qa-2 is not likely to be a defining feature of its function. However, the GPI linkage of Qa-2 does provide an unusual experimental opportunity: GPI-linked proteins can be spontaneously incorporated into plasma membranes in a procedure called “protein painting,” to cause a transient change in the phenotype of the painted cell (McElhinny et al., 2000).

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For HLA-G to be a functional homolog of Qa-2 in conferring the Ped phenotype, it must be expressed in human embryos. Four sets of human embryos have been analyzed for mRNA for HLA-G. Three of these studies analyzed almost 300 human embryos and found that 40–90% of the embryos were positive for HLA-G mRNA depending on the primers that were used (Jurisicova et al., 1996a,b, 1999; Cao et al., 1999). The fourth study did not find any HLA-G mRNA in the human embryos that they analyzed, but their sample size was only 11 embryos (Hiby et al., 1999). The reasons for these negative results are unknown, but include the sensitivity of the assay used, the particular HLA-G primers used, the developmental stage of the embryos tested, and the ethnic makeup of the donors of the embryos. One must conclude from these studies that a significant proportion of human preimplantation embryos express mRNA for HLA-G. However, as pointed out previously, mRNA levels are not necessarily indicative of protein levels. One of the above-cited studies did measure protein expression for HLA-G in human embryos (Jurisicova et al., 1996a), and this study did, indeed, find a correlation of rate of embryonic development with HLA-G protein expression. Thus, these results corroborate the idea that HLA-G is a functional homolog of Qa-2, the Ped gene product. It should be pointed out that studies of expression of HLA-G in human embryos are complicated by the outbred nature of the human population. There are presently 14 known alleles of HLA-G (Morales et al., 2000), and the allelic frequencies differ among different ethnic groups (e.g., Alizadeh et al., 1993; Karhukorpi et al., 1996; Ober et al., 1998; van der Ven et al., 1998a,b; Penzes et al., 1999; Yamashita et al., 1999). Therefore, further studies are needed to fully understand the significance of the expression of different alleles in reproductive success. Interestingly, we have found that the presence of particular alleles of HLA-G in women attending an IVF clinic makes pregnancy success after IVF more likely to occur (Warner et al., 2001). It has been suggested that HLA-G may help protect fetuses from destruction by the maternal immune system by inhibiting NK cells, T cells, and/or myelomonocytic cells, thus enhancing the probability of fetal survival (Pazmany et al., 1996; King et al., 1997; Pende et al., 1997; Rouas-Freiss et al., 1997a,b; Rolstad and Seaman, 1998; Allan et al., 1999; Le Gal et al., 1999; Munz et al., 1999). A complete understanding of the role of HLA-G in the regulation of preimplantation embryo survival remains to be elucidated.

C. Genes That Regulate Cell Cycle Processes Cell cycle signals have been observed in both mouse and human preimplantation oocytes and embryos. These signals occur in gametes during the period preceding fertilization and are induced in the oocyte by the fertilizing spermatozoon on gamete fusion. Possible mechanisms of abnormal cell cycle signaling can impair oocyte maturation and embryo development. These mechanisms may include

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failure of first and second meiotic divisions leading to aneuploidy; incomplete failure of the second meiotic division, leading to de novo chromosomal numerical abnormalities; abnormal pronuclear development and function; and abnormalities of the blastomere cell cycle, possibly leading to embryo cleavage arrest or to mosaicisms and problems with blastomere allocation to embryonic cell lineages. Studies have led to significant improvements in the understanding of molecular mechanisms controlling cell division and growth. A key regulator of these processes has been shown to be maturation promoting factor (MPF). Molecular characterization of MPF has shown that active MPF is a protein dimer composed of a catalytic serine/threonine kinase subunit (p34cdc2 kinase) and a regulatory cyclin B subunit. Nuclear laminins and histone H1 are some of the substrates that are phosphorylated by p34cdc2 kinase; thus MPF has been proposed to be involved in several features of cell division, such as disassembly of the nucleus and chromosome condensation. Periodic activity is a characteristic of MPF since it is needed for each cell division. A protein kinase that has been implicated in upregulation of MPF activity both at the reinitiation of meiosis and during metaphase arrest, is the cellular component of the viral oncogene mos. c-Mos kinase has been proposed to enhance MPF activity via several mechanisms. In mouse oocytes, c-Mos kinase has been proposed to inhibit proteolytic degradation of cyclin B, which in turn leads to accumulation of cyclin-β 1 between meiosis I and II and thus maintains the high MPF activity seen during metaphase arrest. The mRNA expression patterns of the cell cycle genes, c-mos and cyclin-β 1, have been characterized in both mouse and human oocytes and embryos by both qualitative and semiquantitative RT-PCR. The protooncogene c-mos is expressed as a maternal message in an oocyte-specific manner (O’Keefe et al., 1991; Heikinheimo and Gibbons, 1998). The expression of c-mos is transient and little c-mos mRNA can be detected in the human embryo, suggesting meiosis-stage functions for c-mos in the human oocyte. As judged by the disappearance of c-Mos, the maternal pool seems to be degraded by the six- to eight-cell stage before ZGA. Lack of c-Mos might thus allow the two stages of meiosis to continue uninterrupted, resulting in parthenogenic activation. By using specific mRNA degradation by double-stranded RNA (dsRNA), which is termed RNA interference, the targeted reduction of c-mos mRNA in mouse oocytes resulted in parthenogenic activation (Svoboda et al., 2000). On the other hand, abundant expression of cyclin-β 1 seen in both mouse and human oocytes and in embryos from the six-cell stage onward indicates that active transcription and activation of the embryonic genome is necessary for mitotic cell division. The current model of eukaryotic cell cycle regulation suggests that there is an oscillating biochemical clock, which is regulated by surveillance systems, called cell cycle checkpoints. The spindle assembly checkpoint modulates the timing of anaphase initiation in response to the improper alignment of chromosomes at the metaphase plate. If defects are detected, a signal is transduced to halt further progression of the cell cycle until correct bipolar attachment to the spindle is achieved. MAD2 and BUB1 genes encode conserved kinetochore-associated

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proteins believed to be components of the checkpoint regulatory pathway. A failure in this surveillance system could lead to genomic instability that may underlie the increased incidence of aneuploidy in the gametes of older women. To explore this possibility, the copy number of these transcripts has been determined in human oocytes at various stages of maturation, using a real-time rapid cycle fluorescence RT-PCR method (Steuerwald et al., 1999, 2000b, 2001). The results obtained following quantitative analysis suggest that these messages degrade as oocytes age. Potentially, this may impair checkpoint function in older oocytes and may be a contributing factor to age-related aneuploidy (Steuerwald et al., 2001). Work now confirms the presence of the MAD2 protein at the kinetochores of normal, seconddivision mouse oocytes at metaphase (A. Blaszczyk et al., unpublished, 2001). It has also been postulated that postzygotic errors in cell cycle checkpoint gene expression in human embryos may lead to genomic instabilities causing chromosomal mosaicisms (D. Wells et al., unpublished results, 2001).

VI. Genes That Regulate Preimplantation Death There is a current fascination with the topic of cell death, with more than 20,000 publications appearing on this topic in the past 5 years (Golstein, 1998). The reason is that cell death affects crucial normal and pathological biological processes such as development, aging, and cancer, in a wide range of organisms from the nematode Caenorhabditis elegans to humans. Cell death can occur by necrosis or apoptosis. Most of the fascination with cell death has centered on apoptosis because the molecular pathways leading to apoptosis have yielded to biochemical characterization. The pioneering work on genes that mediate apoptosis in C. elegans (Ellis and Horvitz, 1986) has led to the definition of three mammalian genes (or gene families) homologous to three genes in C. elegans that mediate apoptosis: ced-3, which encodes a protein product similar to mammalian caspases; ced-4, which encodes a product similar to mammalian Apaf-1; and ced-9, which encodes a product similar to the mammalian Bcl-2 family of proteins. The first suggestion that apoptosis occurs in embryos was made in 1974 (ElShershaby and Hinchcliffe, 1974). This study and subsequent studies have suggested that a few cells in normal blastocysts die by apoptosis in what appears to be part of a normal developmental program (El-Shershaby and Hinchliffe, 1974; Mohr and Trounson, 1982; Handyside and Hunter, 1986; Hardy et al., 1989; Pierce et al., 1989; Parchment, 1993; Hardy, 1997, 1999; Warner et al., 1998b; Exley et al., 1999a). Abnormal preimplantation embryos often have extensive cellular fragmentation suggesting that they are undergoing death by apoptosis (Jurisicova et al., 1996c, 1998a,b; Levy et al., 1997; Warner et al., 1998c; Yang et al., 1998). DNA fragmentation has been observed with the TUNEL [terminal deoxynucleotidyl transferase [TdT]-mediated dUTP nick-end labeling]

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assay (Ben-Sasson et al., 1995) in fragmented blastomeres, confirming that many fragmented embryonic cells die by apoptosis (Jacobsen et al., 1996; Jurisicova et al., 1996c; Weil et al., 1996; Brison and Schultz, 1997, 1998; Levy et al., 1998; Warner et al., 1998c; Exley et al., 1999). In these studies the embryos were either induced to undergo fragmentation by withdrawal of growth factors or introduction of drugs, or were found in a fragmented state after development in vivo or in vitro. Studies of the genes that regulate apoptosis in preimplantation embryos are still in their infancy. A few studies have appeared showing by RT-PCR that members of the Bcl-2 and caspase families of genes are transcribed in preimplantation mouse and human embryos (Moley et al., 1998; Warner et al., 1998a–c; Jurisicova et al., 1998a; Exley et al., 1999). In addition, caspase 3 activity, a hallmark of apoptosis, has been detected in fragmented preimplantation embryos (Exley et al., 1999). Bcl-2 and Bax protein have been shown to be present in both normal and fragmented preimplantation mouse embryos (Moley et al., 1998; Exley et al., 1999), and there is some indication that the ratio of Bcl-2 to Bax may influence whether an embryo lives or dies during the preimplantation period of development (Moley et al., 1998; Exley et al., 1999). An intriguing set of studies has shown that hyperglycemia can induce apoptosis in preimplantation mouse embryos (Moley et al., 1998; Moley, 1999). Hyperglycemia in diabetic mice and humans during the preimplantation period not only leads to a decreased rate of preimplantation development and some preimplantation embryonic death, but can also lead to fetal loss during gestation and to congenital malformations. It is hypothesized that the mechanism of the effect of excess glucose is indirect. Two possibilities are that excess glucose induces uterine secretion of tumor necrosis factor α (TNF-α), a known inducer of apoptosis, and that excess glucose shuts down glucose transport into the embryo, which also induces apoptosis. Both of these mechanisms would result in the experimentally observed decrease in the number of cells in the ICM. And it is known that a critical number of cells in the ICM is required for normal development and pregnancy outcome (Tam, 1988; Brison and Schultz, 1996; Lane and Gardner, 1997; Van Soom et al., 1997; Chan et al., 2000). Thus, the effect of maternal hyperglycemia on preimplantation embryos is another example, along with those cited previously, that growth during the preimplantation period of development has a marked effect on later development and health.

VII. Conclusions Genes regulate mammalian preimplantation embryo survival. In this review we have considered the expression of genes of both nuclear and mitochondrial origin and their effect on preimplantation embryonic development, growth, and death. We have emphasized the mouse as a model system, and have included data on

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human embryos wherever possible. One major theme is that what happens to an embryo during the preimplantation period of development, whether of genetic or environmental origin, can influence health later in life. Another theme that we have emphasized is that the rate of preimplantation development and the number of cells in the ICM both have a marked effect on subsequent embryo health and survival. In a larger overview, the evaluation of mammalian preimplantation embryonic health and the assessment of the chance that a particular embryo will implant in the uterus and survive to term are provided by two complementary technologies, morphological assessment and genetic analysis. Advances in preimplantation embryo imaging using state-of-the-art technologies such as two-photon laser scanning fluorescence microscopy, polarization microscopy, and quadrature tomographic microscopy (Stott et al., 2001) are underway in a number of laboratories, including our own. Embryo imaging is an exciting new field and the data garnered by these new imaging modalities may in due course significantly enhance and complement the genetic data discussed in this review. We are at the cusp of a revolution in biology with the acquisition of the complete DNA sequence of the human genome and the imminent availability of the sequence of the mouse genome. Thus, large amounts of data from imaging and genetic studies present a challenge for analysis by new methods being developed by researchers working in bioinformatics and data management. The start of the new millennium is an exciting time for biological research. One of the most fascinating fields in biology is the study of preimplantation embryonic development. Preimplantation embryos are being used for a variety of emerging reproductive technologies including IVF, intracytoplasmic sperm injection (ICSI), preimplantation genetic diagnosis, nuclear transfer, cytoplasmic transfer, and the creation of embryonic stem (ES) cells. With the perfection of cloning technologies in animal models, it will be possible in the near future to clone humans. In addition to cloning, ES cells derived from human preimplantation embryos may soon be used for transplantation after their differentiation into particular tissues and organs is perfected. Extensive discussions of scientific benefits and risks of these new technologies, as well as the ethical and moral aspects of the new techniques, are currently underway. These are exciting times for preimplantation embryo research. The future is here!

Acknowledgments We are grateful for financial support from the NIH (HD31505, HD39215, HD40309), the NSF Engineering Research Center for Subsurface Sensing and Imaging Systems (CenSSIS) (EEC-9986821), and the Institute for Reproductive Medicine and Science of Saint Barnabas Medical Center. We also express appreciation to Judy Newmark and Mina Alikani for providing the images of mouse and human embryos, and to Judy Newmark and Martina Comiskey for critical reading of this manuscript.

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Index

A Acetaminophen preimplantation embryo survival, 157–158 AEs, 22–24 Alanine, 66, 75, 91, 92 PI development, 65 preimplantation embryo protection, 70 –71 Alcohol preimplantation embryo survival, 157–158 Aldose reductase, 64 Amino acids efflux two-cell mouse embryos, 85 eggs, 91 embryos, 91 neutral transport, see System A oviductal fluid, 92 PI development, 66 zwitterionic transport, 64 Ang, 110, 122, 129 Angioblasts embryonic derivation, 112 Angiogenesis, 114, 128–132 corpus luteum, 122 ocular, 131–132 Angiopoietin (ang), 110, 129 Anterior mesoderm presumptive, 108–109 Antibodies zona pellucida, 158–159 Apoptosis, 178–179 Arabidopsis thaliana, 8, 9, 12, 23, 27 Arginine, 66, 91, 92 Arterial endothelial cells vs. venous, 127–128 Asparagine, 66, 91, 92 Aspartate, 66, 83 Aspartic acid, 91, 92 Astrocytes, 126

Axial elements (AEs), 22–24 Azidothymidine (AZT) preimplantation embryo survival, 157–158 AZT preimplantation embryo survival, 157–158

B B-alanine, 91, 92 preimplantation embryo protection, 69–70 B-amino acids oviductal fluid, 90 B-amino acid transport system, see System β Barrier vessels EC, 125–127 BBB, 125–127 Betaine, 92 accumulation, 63 preimplantation embryo protection, 68–69 transporter, 63, 64, 71 BGT1 transporter, 63, 64 Blastocyst, 152 expansion, 155 formation genes, 162–163 hatching, 155 Blood-brain barrier (BBB), 125–127 Blood stem cells, 111 Bovine oviductal fluid osmolarity, 93

C Caenorhabditis elegans, 6, 8, 13, 24, 25, 27, 30, 178–179 CAM, 113 Cannabinoiods preimplantation embryo survival, 158 CAP (catabolite gene activator protein), 10

193

194 Cardia bifida, 109 Cardiac development, 108–111 Cardiovascular system, 108–124 Ced-3, 178–179 Ced-4, 178–179 Ced-9, 178–179 Cell-cell communication gap junctions, 134 –135 interactions vascular development, 107–139 Cell cycle checkpoints, 177–178 control, 36–37 genes, 176 –178 Cell death, 178–179 Cell division mammalian embryos, 152 Cell volume increases preimplantation mammalian embryos, 80–88 inorganic ion transport, 61– 62 mammalian organic osmolytes, 62– 63 organic osmolytes, 62– 65 osmolarity, 60– 61 regulation model, 95 preimplantation embryo, 80–97 Chick chorioallantoic membrane (CAM), 113 Chromosome pairing homologous, 22, 24–25, 27, 36 Chromosome segregation premature meiosis I, 26–27 Chromosome structure higher order alterations, 22–24 Chronic progressive external ophthalmoplegia (CPEO), 164 Cl− channel blockers, 87 2Cl− cotransporter, 61– 62 Cleavage divisions, 155 Cloche mutant cells, 111–112 Cloning nuclear transfer, 168 CoHR (common homology region), 32 Collapsin, 136 Colony-stimulating factor (CSF), 171 Common homology region, 32

Index Compaction genes, 161–162 Complement-mediated lysis preimplantation embryo, 159 Connexin genes expression, 134–135 Coprinus cinereus, 6, 8, 12, 13, 18, 23, 24, 25 Corpus luteum angiogenesis, 122 CPEO, 164 CSF, 171 CTLs, 159 Culture media preimplantation embryo survival, 156–157 Cysteine, 66, 91, 92 Cytoplasmic transplantation, 166 Cytotoxic T lymphocytes (CTLs), 159 D Death preimplantation embryo genes, 178–179 Diabetic retinopathy, 131–132 mtDNA4977, 164 DNA cleavage mechanism, 11–12 physical analysis, 3 preimplantation embryo human, 156 mouse, 155 replication connection, 35–36 DNA heteroplasmy mitochondria, 166 Double-Holliday junction, 5 Double-strand break (DSB), 2 catalyst Spo11, 7–8 genes, 27–28 in meiotic chromosome structure development, 22–27 initiate meiotic recombination in Saccharomyces cerevisiae, 3–5 intergenic interactions, 28–29 in organisms, 6 pathway budding yeast, 3 in S. cerevisiae, 4 sites

195

0. Index competition, 34 nonrandomly dispersed, 34 promoter regions, 32 Drosophila melanogaster, 8, 24, 25, 30 Drosophila melanogaster Mei-W68, 12 DSB, see Double-strand break E Early pregnancy factor (EPF), 171 ECs, see Endothelial cells EGF, 171 Eggs amino acids, 91 Embryos amino acids, 91 genes, 159–163 Endocardium, 109 Endoderm, 108–109 Endoderm-derived inducing factor, 109 Endothelial cells (ECs), 115–116 arterial vs. venous, 127–128 barrier vessels, 125–127 development, 111–112 differentiation, 112 gene expression, 124 –125 mural cell precursors, 133–134 specification, 111–112 Endotoxins preimplantation embryo survival, 158 Environment preimplantation embryo survival, 156–159 EPF, 171 Eph receptors, 136 Ephrins, 127–128, 136 Epidermal growth factor (EGF), 171 Epimyocaridum, 109 Equine oviductal fluid osmolarity, 93 Escherichia coli, 6, 10–11, 13, 38 Ethylene oxide preimplantation embryo survival, 157–158

G GABA transporter, 63 Gap junctions cell-cell communication, 134 –135 Gene activator protein, 10 Genes blastocyst formation, 162–163 cell cycle, 176–178 compaction, 161–162 preimplantation embryo death, 178–179 growth, 170–178 reprogramming, 168 ZGA, 161 Genetic effects preimplantation embryo survival, 159–170 Genomic imprinting, 166–167 Germinal vesicle state nuclear transfer, 168–169 Glioblastoma, 129 Glutamate, 66 Glutamic acid, 91, 92 Glutamine, 76, 91, 92 PI development, 65 preimplantation embryo protection, 67 GLY, 55–56, 74–75, 95 Glycine, 66, 90, 91, 92, 95 accumulation, 77–80, 85 oviductal fluid, 90 PI development, 65 preimplantation embryo protection, 67–68, 70–71 Glycine transport system (GLY), 55–56, 74 –75, 95 Glytl gene, 96 GnRH, 171 Gonadotrophin releasing hormone (GnRH), 171 GPC accumulation, 63 Gridlock gene product zebrafish, 128 Growth factors, 171

H F FGF, 113 Fibroblast growth factor (FGF), 113 Fibronectin, 117, 126–127

HB-EGF, 163 Heart development, 108–111 Hemangioblasts, 111

196 Hemangiomas, 129 Heparin-binding epidermal growth factor (HB-EGF), 163 Higher order development chromosome structure, 34–35 Higher order chromosome structure alterations, 22–24 development, 34–35 Histidine, 66, 91, 92 Homeobox genes, 130 Homologous chromosome pairing, 22–27, 36 Homologous recombination, 2 HOPI, 27–28 Human HLA-G, 173–176 Hyaloid vessel system, 123 Hypertonicity mammalian embryos cell volume, 60 –80 Hypotaurine, 91, 92 PI development, 65 preimplantation embryo protection, 69–70 Hypotonic swelling permeability one-cell mouse embryo, 86 I ICM, 152–156 Id proteins, 130 IL-1, 171 IL-6, 171 IL-10, 171 Inner cell mass (ICM), 152–156 Inorganic ion transport cell volume, 61–62 Integrins, 117 Interferon, 171 Interleukin 1 (IL-1), 171 Interleukin 6 (IL-6), 171 Interleukin 10 (IL-10), 171 Isoleucine, 66, 91, 92 K KAR3, 28 K+ channels RVD, 82 K+ cotransporter, 61–62 Kearns-Sayre syndrome (KSS), 164 KSS, 164

Index L LacZ gene, 137–138 Lateral elements (LEs), 22–24 LEs, 22–24 Lethal mutations, 163–164 Leucine, 66, 91, 92 Leukemia inhibitory factor (LIF), 163, 171 LIF, 163, 171 Lymphatic cells, 116 Lysine, 66, 91, 92

M Macrophages, 159 Madin-Darby canine kidney (MDCK) cells, 67 Major histocompatibility complex (MHC), 159 Mammalian embryos preimplantation, see Preimplantation mammalian embryos Mammalian organic osmolytes cell volume, 62–63 transport systems, 63 Mannitol osmolarity, 57 Maturation promoting factor (MPF), 177 MDCK cells, 67 MeAIB, 2 MEF2C, 110 Meiosis-specific, 20, 25 Meiosis-specific alteration chromatin nuclease hypersensitivity, 33 Meiotic prophase progression, 22–27 Meiotic recombination, 19 evolutionary conservation, 18 initiation cast of players, 6–31 chromatin structure, 31–33 double-strand breaks, 3–6 factors, 31 mechanism and control, 1–38 molecular model, 37–38 promoters, 31–33 sequence specificity, 31–33 site selectivity, 31–33 Mei-P22, 24 Mei-W68, 24 MEK1, 27–28

197

0. Index ME14 (meiosis-specific), 20, 25 MER2, 25 MER1 (meiotic recombination), 19 Methanococcus jannaschi, 10–11 Methionine, 66, 92 MHC, 159 Mice ped gene, 171–173 human homolog, 173–176 preimplantation embryos, 153 RVD, 81 – 83 in vitro, 58, 59 vasculogenesis EC-associated protein expression, 113 zygotes swelling-activated current, 84 Micrococcal nuclease (MNase) hypersensitivity, 33 Mitochondria DNA heteroplasmy, 166 genes, 164 –166 mutations, 164 –165 Mitomycin C preimplantation embryo survival, 157–158 MNase hypersensitivity, 33 MPF, 177 MRE2, 19 MRE11, 13–18 nonull alleles, 16–17 Mural cell precursors EC, 133–134 Mural cells, 115–116 differentiation, 118–120 identification, 114 recruitment, 118 remodeling, 123–124 specifications, 114–120 Mural trophectoderm, 152 Myocyte enhancer factor 2C, 110 Myocyte enhancer factor 2C (MEF2C), 110 Myo-Inositol preimplantation embryo protection, 70–71 Myo-inositol transporter, 63, 64, 71 N Na+ cotransporter, 61– 62 NAM8, 19

Natural killer cells, 159 NCT/TAUT transporter, 63 Neovascularization ocular, 132 Neuronal-vascular interactions, 135–137 Neuropilin, 136 Neurospora crassa, 9, 13 Neutral amino acids transport, see System A Nitrosourea preimplantation embryo survival, 158 Nuclear transfer, 167–169 cloning, 168 germinal vesicle state, 168 –169 Nucleotide-resolution mapping Spo11, 33 O Ocular angiogenesis, 131–132 Ocular neovascularization, 132 Oocytes telomerase, 170 Organic osmolytes, 55 accumulation, 63–65 tonicity, 64–65 cell volume, 62–65 efflux, 81 mammalian cell volume, 62–63 transport systems, 63 osmolarity in vivo, 88–93 oviductal fluid, 90 –93 preimplantation embryo, 88–90 mammalian, 65–80 improved development, 65–67 osmotically regulated accumulation, 77–80 permeability, 83–88 protection, 67–71 transport, 71–72 Osmolarity in vitro, 57–60 mammalian embryos, 57–60 organic osmolytes in vivo, 88–93 oviductal fluid, 93 proportional dilution, 57

198 Osmolytes organic, see Organic osmolytes Osmotic regulation preimplantation embryo, 55–80 model, 95 Oviductal fluid amino acids, 92 bovine, 93 equine, 93 organic osmolytes, 90 –93 osmolarity, 93 P PAF, 171 Parenchymal cells, 116 Patch-clamp measurement, 84 PDGF-B, 118 Pearson’s syndrome, 164 PECAM, 117 Ped gene mice, 171–173 human homolog, 173–176 Peptidergic neurons, 135 Pericytes, 114–115, 123–124, 133–134 scanning electron micrograph, 120 Perivascular cells identification, 114 Permeability hypotonic swelling one-cell mouse embryo, 86 Phenylalanine, 66, 91, 92 PKC, 162 Platelet-activating factor (PAF), 171 Platelet-derived growth factor B (PDGF-B), 118 Platelet endothelial cell adhesion molecule (PECAM), 117 Platelets, 116 Polarity gradient, 5 Polar trophectoderm, 152 Posterior mesoderm presumptive, 109 Posterior primitive streak tissue, 109 Precardiac tissue differentiation, 109 induction, 108–109 Preimplantation embryos cell volume regulation, 80–97 model, 95 complement-mediated lysis, 159 culture media, 156–157

Index development, 152–156 genes death, 178–179 growth, 170–178 human, 154 mammalian cell division, 152 characteristics, 156 mice, 153 RVD, 81–83 in vitro, 58, 59 one-cell vs. two-cell osmolarity, 60 organic osmolytes osmotically regulated accumulation, 77–80 permeability, 83–88 protection, 67–71 transport, 71–72 osmotic regulation, 55–80 model, 95 sodium chloride, 59 survival environmental effects, 156–159 genetic effects, 159–170 genetic regulation, 151–180 toxic substances, 157–158 in vivo organic osmolytes, 88–90 Preimplantation mammalian embryos cell division, 152 cell volume increases, 80–88 regulation, 81–88 characteristics, 156 hypertonicity cell volume, 60 –80 organic osmolytes, 65–80 improved development, 65–67 osmolarity, 57–60 volume regulation, 56 Premature chromosome segregation meiosis I, 26–27 Premeiotic S phase duration, 25–26 Presumptive anterior mesoderm, 108–109 Presumptive posterior mesoderm, 109 Primitive network formation, 116–118 Primitive vessels, 117 Prokaryotic type II enzymes, see Topoisomerase VI Proline, 66, 76, 91, 92 preimplantation embryo protection, 70–71

199

0. Index Proportional dilution osmolarity, 57 Protein preimplantation embryo human, 156 mouse, 155 Protein kinase C (PKC), 162 Pyrococcus horikoshii, 9 Q Qa-2, 175 R Rad50, 13–18 biochemical properties, 13–15 null mutants, 23 point mutations, 15 –16 Rad32 function, 18 Radiolabeled compounds permeability swelling, 87 Raffinose osmolarity, 57, 59 Rec102, 20–21, 25 Rec103, 21 Rec104, 20–21 Rec107, 19 Rec114, 20–21 Recombination homologous, 2 Recombination events chromatin structure, 31–33 promoters, 31–33 sequence specificity, 31–33 site selectivity, 31–33 Recombination hot spots, 33 RED1, 27–28 Regulatory volume decrease (RVD), 80, 87 K+ channels, 82 mouse embryos, 81–83 swelling-activated permeability, 89 Regulatory volume increase (RVI), 61–62 Remodeling mural cells, 123–124 Retinopathy of prematurity (ROP), 131 RNA preimplantation embryo human, 156 mouse, 155

ROP, 131 RVD, 80, 87 K+ channels, 82 mouse embryos, 81–83 swelling-activated permeability, 89 RVI, 61–62 S Saccharomyces cerevisiae, 2, 9, 12, 13, 15, 18, 21, 22, 23, 25, 26, 30, 32, 36 chromosome III, 34 enzyme, 37 Spo11 identification and early characteristics, 6 –7 Saccharomyces paradoxus, 20–21 Saccharomyces pastorianus, 20–21 SAE2/COM1, 13–18 evolutionary conservation, 13–15 meiotic recombination roles, 15–18 Sarcosine, 75–76 SC, 22–24 Schizosaccharomyces pombe, 6, 9, 12, 13, 21, 26, 29, 30, 32, 33 SCL TAL-1, 113 Septa formation, 109 Serine, 66, 91, 92 Sezary cell leukemia (SCL) TAL-1, 113 SK18 (SuperKiller), 21 SMCs, 114, 118–119, 133–134 scanning electron micrograph, 120 Smooth muscle cells (SMCs), 114, 118–119, 133–134 scanning electron micrograph, 120 Sodium chloride osmolarity, 57 preimplantation embryo, 59 Sorbitol, 64 accumulation, 63 Sordaria macrospora, 9 Splanchnopleural mesoderm, 112 Spo11, 6 –13 double-strand break catalyst, 7–8 evolutionary conservation, 8 functions, 30 –31 mechanism, 11–12 molecular model, 37–38

200 Spo11, (continued ) nonmeiotic cells, 12–13 Saccharomyces cerevisiae identification and early characteristics, 6–7 Top6A family structure, 10 –11 SRp20, 162–163 Sucrose osmolarity, 57 Sulfolobus shibatae enzyme, 8 SuperKiller, 21 Swelling-activated Cl− current, 80 –81, 83 Swelling-activated current mouse zygotes, 84 Swelling-activated permeability RVD, 89 screening, 87– 88 Synaptonemal complex (SC), 22–24 System A, 63, 64, 71–72 System β, 55–56, 63, 64, 71, 72–73 T TAL-1 SCL, 113 Taurine, 66, 75, 90, 91, 92 accumulation, 63 PI development, 65 preimplantation embryo protection, 69–70 swelling-activated Cl− current, 83 transport system, 63 T cell acute leukemia 1(TAL-1) SCL, 113 TCS, 169–170 TE, 152–156 Telomerase, 169–170 Telomerase catalytic subunit (TCS), 169–170 Telomeres, 169–170 TGF, 171 TGF-β, 109, 118–119 Threonine, 66, 91, 92 TonE, 64 Tonicity response element (TonE), 64 Top6A family Spo11 structure, 10 –11 Topoisomerase VI, 8 Toprim domain, 10, 12 Toprim metal-binding pocket, 37 Toxic substances preimplantation embryo survival, 157–158

Index Transforming growth factor-β (TGF-β), 109, 118–119 Transforming growth factor (TGF), 171 Transport systems mammalian organic osmolytes, 63 Trophectoderm (TE), 152–156 Tryptophan, 66, 91, 92 Tumor angiogenesis, 128–130 Tyrosine, 66, 91, 92 V Valine, 66, 91, 92 Valves formation, 109 Vascular beds differences, 124 –128 Vascular development cell-cell interactions, 107–139 defined, 107 Vascular endothelial-cadherin (VE-cadherin), 116–117 Vascular endothelial growth factor (VEGF), 113, 116, 122, 129, 131, 137–138, 171 Vascular tree formation, 115–116 Vasculature, see also Vessels disorganization, 116 –117 early development, 111–114 Vasculogenesis, 114, 116 adult, 137–138 mice EC-associated protein expression, 113 VE-cadherin, 116 –117 VEGF, see Vascular endothelial growth factor Venous endothelial cells vs. arterial, 127–128 Vessels maturation, 120 –124 remodeling, 120 –123 Volume regulation mammalian embers, 56 VSOAC channel, 81, 86–87, 96 W Wandering mesenchymal cells, 111–112 WD motif, 21 Whole-cell patch-clamp electrophysiological recordings, 83

201

0. Index X Xenopus, 108–109, 116 XRS2, 13–18 Z Zebrafish gridlock gene product, 128 ZGA genes, 161

Zip2, 23 Zip23, 23 Zona pellucida, 152, 155, 158–159 Zwitterionic amino acids transport, 64 Zygotes mice swelling-activated current, 84 Zygotic genomic activation (ZGA) genes, 161

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Contents of Previous Volumes

Volume 47 1 Early Events of Somitogenesis in Higher Vertebrates: Allocation of Precursor Cells during Gastrulation and the Organization of a Moristic Pattern in the Paraxial Mesoderm Patrick P. L. Tam, Devorah Goldman, Anne Camus, and Gary C. Shoenwolf

2 Retrospective Tracing of the Developmental Lineage of the Mouse Myotome Sophie Eloy-Trinquet, Luc Mathis, and Jean-Franc¸ois Nicolas

3 Segmentation of the Paraxial Mesoderm and Vertebrate Somitogenesis Olivier Pourqul´e

4 Segmentation: A View from the Border Claudio D. Stern and Daniel Vasiliauskas

5 Genetic Regulation of Somite Formation Alan Rawls, Jeanne Wilson-Rawls, and Eric N. Olsen

6 Hox Genes and the Global Patterning of the Somitic Mesoderm Ann Campbell Burke

7 The Origin and Morphogenesis of Amphibian Somites Ray Keller

8 Somitogenesis in Zebrafish ¨ Scott A. Halley and Christiana Nusslain-Volhard

9 Rostrocaudal Differences within the Somites Confer Segmental Pattern to Trunk Neural Crest Migration Marianne Bronner-Fraser

203

204

Contents of Previous Volumes

Volume 48 1 Evolution and Development of Distinct Cell Lineages Derived from Somites Beate Brand-Saberi and Bodo Christ

2 Duality of Molecular Signaling Involved in Vertebral Chondrogenesis Anne-H´el`ene Monsoro-Burq and Nicole Le Douarin

3 Sclerotome Induction and Differentiation Jennifer L. Docker

4 Genetics of Muscle Determination and Development Hans-Henning Arnold and Thomas Braun

5 Multiple Tissue Interactions and Signal Transduction Pathways Control Somite Myogenesis Anne-Ga¨elle Borycki and Charles P. Emerson, Jr.

6 The Birth of Muscle Progenitor Cells in the Mouse: Spatiotemporal Considerations Shahragim Tajbakhsh and Margaret Buckingham

7 Mouse –Chick Chimera: An Experimental System for Study of Somite Development Josiane Fontaine-P´erus

8 Transcriptional Regulation during Somitogenesis Dennis Summerbell and Peter W. J. Rigby

9 Determination and Morphogenesis in Myogenic Progenitor Cells: An Experimental Embryological Approach Charles P. Ordahl, Brian A. Williams, and Wilfred Denetclaw

Volume 49 1 The Centrosome and Parthenogenesis ¨ Thomas Kuntziger and Michel Bornens

2 ␥-Tubulin Berl R. Oakley

3 ␥-Tubulin Complexes and Their Role in Microtubule Nucleation Ruwanthi N. Gunawardane, Sofia B. Lizarraga, Christiane Wiese, Andrew Wilde, and Yixian Zheng

Contents of Previous Volumes

205

4 ␥-Tubulin of Budding Yeast Jackie Vogel and Michael Snyder

5 The Spindle Pole Body of Saccharomyces cerevisiae: Architecture and Assembly of the Core Components Susan E. Francis and Trisha N. Davis

6 The Microtubule Organizing Centers of Schizosaccharomyces pombe Iain M. Hagan and Janni Petersen

7 Comparative Structural, Molecular, and Functional Aspects of the Dictyostelium discoideum Centrosome Ralph Gr¨af, Nicole Brusis, Christine Daunderer, Ursula Euteneuer, Andrea Hestermann, Manfred Schliwa, and Masahiro Ueda

8 Are There Nucleic Acids in the Centrosome? Wallace F. Marshall and Joel L. Rosenbaum

9 Basal Bodies and Centrioles: Their Function and Structure Andrea M. Preble, Thomas M. Giddings, Jr., and Susan K. Dutcher

10 Centriole Duplication and Maturation in Animal Cells B. M. H. Lange, A. J. Faragher, P. March, and K. Gull

11 Centrosome Replication in Somatic Cells: The Significance of the G1 Phase Ron Balczon

12 The Coordination of Centrosome Reproduction with Nuclear Events during the Cell Cycle Greenfield Sluder and Edward H. Hinchcliffe

13 Regulating Centrosomes by Protein Phosphorylation Andrew M. Fry, Thibault Mayor, and Erich A. Nigg

14 The Role of the Centrosome in the Development of Malignant Tumors Wilma L. Lingle and Jeffrey L. Salisbury

15 The Centrosome-Associated Aurora/lpl-like Kinase Family T. M. Goepfert and B. R. Brinkley

16 Centrosome Reduction during Mammalian Spermiogenesis G. Manandhar, C. Simerly, and G. Schatten

17 The Centrosome of the Early C. elegans Embryo: Inheritance, Assembly, Replication, and Developmental Roles Kevin F. O ′Connell

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Contents of Previous Volumes

18 The Centrosome in Drosophila Oocyte Development Timothy L. Megraw and Thomas C. Kaufman

19 The Centrosome in Early Drosophila Embryogenesis W. F. Rothwell and W. Sullivan

20 Centrosome Maturation Robert E. Palazzo, Jacalyn M. Vogel, Bradley J. Schnackenberg, Dawn R. Hull, and Xingyong Wu

Volume 50 1 Patterning the Early Sea Urchin Embryo Charles A. Ettensohn and Hyla C. Sweet

2 Turning Mesoderm into Blood: The Formation of Hematopoietic Stem Cells during Embryogenesis Alan J. Davidson and Leonard I. Zon

3 Mechanisms of Plant Embryo Development Shunong Bai, Lingjing Chen, Mary Alice Yund, and Zinmay Rence Sung

4 Sperm-Mediated Gene Transfer Anthony W. S. Chan, C. Marc Luetjens, and Gerald P. Schatten

5 Gonocyte–Sertoli Cell Interactions during Development of the Neonatal Rodent Testis Joanne M. Orth, William F. Jester, Ling-Hong Li, and Andrew L. Laslett

6 Attributes and Dynamics of the Endoplasmic Reticulum in Mammalian Eggs Douglas Kline

7 Germ Plasm and Molecular Determinants of Germ Cell Fate Douglas W. Houston and Mary Lou King

Volume 51 1 Patterning and Lineage Specification in the Amphibian Embryo Agnes P. Chan and Laurence D. Etkin

2 Transcriptional Programs Regulating Vascular Smooth Muscle Cell Development and Differentiation Michael S. Parmacek

Contents of Previous Volumes

207

3 Myofibroblasts: Molecular Crossdressers Gennyne A. Walker, Ivan A. Guerrero, and Leslie A. Leinwand

4 Checkpoint and DNA-Repair Proteins Are Associated with the Cores of Mammalian Meiotic Chromosomes Madalena Tarsounas and Peter B. Moens

5 Cytoskeletal and Ca2+ Regulation of Hyphal Tip Growth and Initiation Sara Torralba and I. Brent Heath

6 Pattern Formation during C. elegans Vulval Induction Minqin Wang and Paul W. Sternberg

7 A Molecular Clock Involved in Somite Segmentation Miguel Maroto and Olivier Pourqui´e

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