Interestingly, these mutant mouse models also exhibit defects in melanogenesis and hematopoiesis because these genes are also required for the migration of melanocytes and mast cells.¹⁴ PGCs also express integrin subunits, which may potentially interact with laminin and fibronectin produced by cells in the genital ridges during the formation of the gonads. For example, integrin β₁ is required for PGC colonization of the gonads, as the migration of PGCs is arrested at the gut endoderm in integrin β₁ null mutants.¹⁵

OOGONIA AND THEIR MEIOTIC ENTRY TO BECOME OOCYTES

Colonization of the genital ridges by PGCs occurs concomitantly with the formation of the ovaries, at 12 to 13 dpc. Postmigratory PGCs, together with somatic epithelial cells that have derived from the coelomic epithelium and mesenchyme, form the ovarian cortical sex cords. Germ cell survival beyond 13 dpc requires the C2H2-type zinc-finger transcription factor Zpf148, which may activate tumor suppressor protein p53 in germ cells at this stage of development.¹⁶ Around the same time, PGCs differentiate into oogonia, which are sexually differentiated germ cells that would undergo the last rounds of mitotic divisions to establish the germ cell population at 11 to 13 dpc.¹⁷

Our understanding of mammalian oogonium function has been limited, perhaps due to their transient existence. Incomplete cytokinesis during synchronous, mitotic divisions of the oogonia results in the formation of clusters, or cysts, that are connected by intercellular bridges 0.5 to 1.0 μm in diameter.¹⁷ Oogonia become oocytes by entering meiosis I at 14 dpc; by 17 dpc, most oogonia have transitioned to oocytes. In the perinatal period, these germ cell cysts undergo breakdown, after which only a third of the oocytes survive and become enclosed in a single layer of granulosa cells to form primordial follicles.¹⁷ Thus, perinatal germ cell apoptosis appears to be a developmental process that is differentially regulated compared to the loss of oocytes during follicular atresia that occurs in the sexually mature animal. Pepling and Stradling suggested that cyst formation and subsequent breakdown may serve to ensure the incorporation of mitochondria from dying oocytes into those that survive.¹⁷

Oocyte Development

The female germ cell becomes an oocyte when it enters prophase of meiosis I. The signals that instruct the germ cell to enter meiosis are not known. Further, we do not know whether this decision is cell-autonomous or whether it is dependent on signaling from adjacent somatic cells. Nevertheless, we do know that all oogonia have entered prophase to become oocytes by 17 dpc. DNA replication, chromosome pairing, and recombination, which are hallmarks of sexual reproduction ensuring that all eggs contain a unique haploid complement of DNA, occur in prophase.

Prophase of Meiosis I

The four stages of prophase I are leptotene, zygotene, pachytene, and diplotene. DNA replication is finalized in preleptotene, and in leptotene, sister chromatids search for their homologous counterparts. Formation of recombination nodules also commences at this time to facilitate interaction between homologous chromosomes.

In zygotene, homologous chromosomes pair and begin to synapse. Their synapses are maintained by the synaptonemal complex, which is formed by many protein subunits, including synaptonemal complex protein 3 (Scp3) in the axial elements, Scp1 in the central element, and Scp2.¹⁸ Synapsis is completed in pachytene and continues to be maintained until diplotene, when homologous chromosomes are held together mainly at sites of chiasmata. The crossing-over and recombination of chromosomes occur over 4 days in the pachytene stage, prior to the formation of ovarian follicles.

Many proteins, including those with functions in DNA mismatch repair, recombination, and DNA damage checkpoint, have essential roles during prophase I. Gene-targeted mouse models deficient in these proteins exhibit female meiosis phenotypes ranging from premature ovarian failure due to perinatal oocyte loss, to defects that become apparent only later during oocyte maturation. These proteins are also required in spermatogenesis, which demonstrates mechanisms of prophase I that are common to both sexes, although the meiosis phenotypes tend to be more severe and occur in slightly earlier stages of development in males. Our discussion focuses on the requirement of these proteins in oogenesis.

MutL and MutS Families of Proteins

DNA mismatch recognition and repair by the MutL and MutS families of proteins are well conserved from yeast to humans.¹⁹ MutL and MutS heterodimeric protein complexes are thought to interact to activate DNA mismatch repair. Although the MutL and MutS vertebrate homologs (Mlh and Msh, respectively) are generally known for their roles in maintaining genomic stability against tumor formation, gene targeting in mice revealed that Mlh1, Mlh3, Msh4, and Msh5 have essential functions in meiosis in both males and females.²⁰^–²⁴

Mlh1 and Mlh3 homozygous mutant females have similar infertility phenotypes, in which the newborn ovaries appear normal with a normal number of follicles at various stages of development. However, there is a significant decrease in the number of oocytes that can complete meiosis I, meiosis II, and undergo subsequent development to two-cell embryos.^21,²³ Both Mlh1 and Mlh3 colocalize to chromosomes in pachytene and are thought to have critical functions in meiotic recombination, as supported by the decreased number of chiasmata formed in the oocytes of these mutant animals.^21,^23,²⁵

Although oocytes seem to enter and arrest at diplotene, chromosomes are unpaired and cannot stably attach to the bipolar spindle, which leads to abnormal meiotic I spindle formation, abnormal or incomplete meiosis I, and fertilization failure.²⁵ Further, in both human and mouse pachytene oocytes, MLH1 and MLH3 serve as molecular markers of recombination nodules, which are necessary for the crossover structures that are present in the diplotene stage.²⁶

Similarly, Msh4 and Msh5 have essential functions in pachytene of prophase I, although their phenotypes were more severe.^20,^22,²⁴ Female null mutants of Msh4 and Msh5 have significant loss of oocytes by postnatal days 2 to 4, before meiotic arrest at the diplotene stage.^20,^22,²⁴ Their oocytes enter leptotene to form synaptonemal complexes but fail to undergo complete pairing at zygotene and fail to enter diplotene. These oocytes undergo apoptosis, which results in atretic follicles, loss of follicular architecture, and premature ovarian failure. Both Msh4 and Msh5 localize to chromosomes, can form heterodimers in vitro, and are thought to function at the same time point during chromosomal synapsis.²² The relevance of these findings from the mouse model to human reproduction is further supported by the expression of MSH4 and MSH5 in the human testes and ovaries.²⁷

We do not yet know the exact mechanisms by which these Mlh and Msh proteins act or how they bind to chromosomes. The oocyte apoptosis phenotype has been attributed to failure of chromosome pairing or formation of synapses. However, recent genetic analyses based on the Spo11/Msh or Spo11/Mlh double null mutants indicate that oocyte apoptosis is a consequence of failure to repair double-strand breaks.²⁸ Spo11 initiates double-strand breaks, which are required for recombination. In the absence of Spo11, the phenotype of Mlh1 and Msh5 was less severe and resembled that of Spo11. Similarly, Dmc1, another protein involved in recombination, and ataxia-telengiectasia mutated (Atm), a DNA damage checkpoint protein, are both required for the repair of double-strand breaks; in their absence, persistent double-strand breaks lead to oocyte death via apoptosis.²⁸

The functional mechanism of a protein is often elucidated by identifying proteins with which it associates and the nature of their interactions. The colocalization of Mlh1 proteins with Scp3 and cyclin-dependent kinase 2 (Cdk2) may suggest interaction or related functions, especially since Scp3 and Cdk2 have also been shown to be required in meiosis.^29,³⁰ Scp3 is a key protein subunit in the synaptonemal complex, which holds homologous chromosome pairs to facilitate synapsis. Female mice lacking Scp3 are subfertile, because many of their oocytes contain univalents (unpaired chromosomes), exhibit abnormal chromosomal segregation, and produce a decreased number of viable embryos. Interestingly, their fertility worsens with age, so that this model is thought to be potentially powerful in studying the age-related subfertility and aneuploidy that are major problems in human reproduction.³⁰

Cdk Family of Proteins

Cyclin-dependent kinase (Cdk2) belongs to the family of Cdk proteins, which are considered master regulators of the cell cycle.³¹ However, Cdk2-null mutants are unexpectedly viable; thus, Cdk2 is not required for mitosis during development in vivo to be required for prophase I of meiosis in the oocytes instead. Oocyte loss and premature ovarian failure occur by the first few postnatal days. Although the precise molecular defects differ, a similar phenotype is seen in the null mutants of Dmc1, Msh4, Msh5, and Atm, presumably because these oocytes fail to enter or complete pachytene, and enter the common apoptotic pathway by default. In contrast, by postnatal day 5, all oocytes in wild type mice have progressed to the diplotene stage, where they remain arrested until peri-ovulation in the sexually mature adult, when they resume and complete meiosis.

OOCYTE DEVELOPMENT AND EARLY STAGES OF FOLLICULOGENESIS

The transition from pachytene to diplotene marks the beginning of folliculogenesis, which is closely associated with subsequent oocyte development. Whereas no pachytene oocytes are found within follicles, approximately 80% of diplotene oocytes are enclosed in primordial or more developmentally advanced follicles.³² Primordial follicles are formed by the encapsulation of a diplotene-arrested oocyte by a single flat layer of granulosa cells. Subsequently, the granulosa cells, still in a single layer, assume a cuboidal shape to form primary follicles.

Transcription Factors

Our understanding of primordial and primary follicular development has recently been enlightened by the discovery of the requirement of two transcription factors at these stages of development (Fig. 3-2). Figα, a β-loop-helix-loop transcription factor, is required for oocyte survival into the primordial follicle stage.³³ Based on its known role in the transcription of oocyte-specific genes zona pellucida 1 (zp1), zp2, and zp3, its role in primordial follicle formation is presumed to be the transcription of other oocyte-specific transcription factors.³³

Figure 3-2 Schematic representation of oogenesis and folliculogenesis during development in the mouse model. This figure shows and correlates the developmental stages of the oocyte and follicle, and highlights examples of essential regulators at each stage. The “Types 1-8” nomenclature, as described by Pedersen and Peters ¹⁹¹, is also shown to facilitate interpretation of the literature. PGCs, primordial germ cells; dpc, number of days postcoitum; dpp, number of days postpartum.

Another transcription factor, Nobox, which is an oocyte-specific homeobox protein, has a critical role in the transition of primordial to primary follicles.³⁴ In the absence of Nobox, mouse ovarian follicles do not develop beyond the primordial stage, which results in massive oocyte loss in the early postnatal period.³⁴ Identification of novel downstream target genes of Figα and Nobox in the future should provide insight into these early stages of folliculogenesis.

Zona Pellucida

The growing oocyte synthesizes zona pellucida glycoproteins, which comprise approximately 17% of the total cellular protein content. The zona pellucida proteins zp1, zp2, and zp3 are encoded by distinct genes and are coordinately expressed under the control of Figα during oocyte growth.³³ These zona proteins form the zona pellucida or zona matrix that surrounds the growing oocyte, but each one has a critical role in proper oocyte and embryo development. zp1, the only zp protein that forms intermolecular disulfide bonds, contributes to 10% to 15% of the zona mass and is thought to provide structural integrity to the zona pellucida. zp3 serves as the receptor for sperm binding and the subsequent acrosome reaction; zp2 functions as a secondary receptor. Together, zp2 and zp3 mediate sperm binding and species specificity during fertilization.

These three glycoproteins are essential for fertilization and viability of growing oocytes and early embryos. Mice lacking zp3 cannot form the zona matrix and are sterile,³⁵ and zp1 knockout mice have decreased fecundity and precocious hatching of early embryos from the structurally compromised zona matrix.³⁶ Further, ectopic granulosa cells are seen between the oolemma and zona pellucida, and may increase the perivitelline space prior to ovulation.³⁶ It is important to note that these may be four zona pellucida genes that are expressed in humans (ZP1, ZP2, ZP3, and zPB). Interestingly, abnormalities and lack of zona pellucida in human oocytes have been observed during IVF; however, the cause and implication of this defect in human reproduction is not well understood.

Regulation of Folliculogenesis by Oocyte Morphogens

Although bidirectional communication between the oocyte and follicular cells is required for the development of the meiotically competent oocyte, there is increasing support to view the oocyte as autonomous in determining its own fate by regulating follicular development via the secretion of various growth factors.^37,³⁸ The establishment of a morphogen gradient by the oocyte regulates follicular development via differential gene expression and function in granulosa cells. In the absence of these morphogens, follicle-stimulating hormone (FSH) induces all cumulus cells, which are directly adjacent to the oocyte and have distinct functions, to differentiate into membrane granulosa cells. Currently, several members of the TGFβ superfamily are the most well-understood candidate oocyte morphogens.

Transforming Growth Factor β

The mammalian oocyte expresses at least three TGFβ superfamily members: growth differentiation factor 9 (Gdf9), Bmp15, and Bmp6.³⁸ Although Gdf9 is expressed throughout the reproductive tract and bone marrow,³⁹ its oocyte expression is restricted to oocytes in follicles at the primary (type 3a), preantral, or more advanced stages of development.^39,⁴⁰ After fertilization, the level of Gdf9 transcripts decreases to low or undetectable in preimplantation embryos.⁴¹

Gdf9 is a critical regulator of follicular development. In Gdf9-deficient female mice, primordial and primary follicles appear normal, but development beyond type 3a follicles is blocked and the animals are sterile.⁴² Further, in the absence of Gdf9, theca cell precursors are absent around the follicles, and the expression of kit ligand and inhibin-α is upregulated in the granulosa cells of the primary follicles.⁴³ The increased levels of kit ligand are thought to mediate excessive oocyte growth via interaction with the c-kit tyrosine kinase receptor on the oocyte, leading to abnormally large oocytes and their cell death.⁴³ Interestingly, Gdf9 deficiency also causes failure of granulosa cells to proliferate or undergo cell death, which presumably leads to their abnormal differentiation into clusters of steroidogenic cells reminiscent of corpora lutea.⁴³ Therefore, the oocyte can be viewed as the master regulator of early folliculogenesis via its secretion of Gdf9.

The oocyte expression pattern of Gdf9 is shared by its family member Bmp15, which is encoded by an X-linked gene.^44,⁴⁵ Bmp15 homozygous null mutant females are subfertile due to decreased rates of ovulation and early embryo survival. Further, an effect of gene dosage is observed in Gdf9^+/− Bmp15^−/− mice, which have a more severe phenotype than Bmp15^−/− mice.⁴⁶ In addition, cumulus–oocyte reconstitution and RNA interference (RNAi) experiments demonstrated that both proteins regulate cumulus expansion in vitro. Therefore, Gdf9 and Bmp15 are thought to have synergistic roles in the regulation of folliculogenesis.

Interestingly, naturally occurring mutations in GDF9 and BMP15 are common in certain strains of domestic sheep, perhaps as a result of breeding practices. Although sheep carrying mutations in both alleles of GDF9 or BMP15 are sterile, those carrying a single mutant allele have increased ovulation rates and are “super fertile.” Superovulation in the presence of heterozygosity of either of these genes is thought to be caused by decreased inhibin production by granulosa cells, which in turn leads to increased pituitary FSH secretion, development of more than one dominant follicle, and ultimately superovulation. Thus, the paradoxical increase in fertility when one allele is mutated exemplifies the impact of gene dosage on ovarian follicular function.⁴⁵

RECRUITMENT OF OVARIAN FOLLICLES

Nongrowing mouse oocytes within primordial follicles have a diameter of approximately 12 μm and comprise the resting pool. Cohorts from the resting pool are continuously induced by ovarian paracrine signals to undergo coordinated oocyte and follicular growth during a process called initial recruitment (Fig. 3-3 ⁴⁷). The recruited, growing follicles are called primary follicles, but this growth phase is very protracted so that primordial and primary follicles may not be easily distinguishable. These growing follicles subsequently develop into secondary and antral follicles. (The timeline of different stages of follicular development in humans and rodents are shown in Fig. 3-4 and 3-5.⁴⁷)

Figure 3-3 Life history of ovarian follicles: endowment, maintenance, initial recruitment, maturation, atresia or cyclic recruitment, ovulation, and exhaustion. A fixed number of primordial follicles are endowed during early life, and most of them are maintained in a resting state. Growth of some of these dormant follicles is initiated before and throughout reproductive life (initial recruitment). Follicles develop through primordial, primary, and secondary stages before acquiring an antral cavity. At the antral stage most follicles undergo atresia; however, under the optimal gonadotropin stimulation that occurs after puberty, a few of them are rescued (cyclic recruitment) to reach the preovulatory stage. Eventually, depletion of the pool of resting follicles leads to ovarian follicle exhaustion and senescence.

Rights were not granted to include this figure in electronic media. Please refer to the printed book.

Figure 3-4 Duration of follicle recruitment and selection in human and rat ovaries. Primordial follicles undergo initial recruitment to enter the growing pool of primary follicles. Due to its protracted nature, the duration required for this step is unknown. In the human ovary, more than 120 days are required for the primary follicles to reach the secondary follicle stage, whereas 71 days are needed to grow from the secondary to the early antral stage. During cyclic recruitment, increases in circulating follicle-stimulating hormone (FSH) allow a cohort of antral follicles (2 to 5 mm in diameter) to escape apoptotic demise. Among this cohort, a leading follicle emerges as dominant by secreting high levels of estrogens and inhibins to suppress pituitary FSH release. The result is a negative selection of the remaining cohort, leading to its ultimate demise. Concomitantly, increases in local growth factors and vasculature allow a positive selection of the dominant follicle, thus ensuring its final growth and eventual ovulation. After cyclic recruitment, it takes only 2 weeks for an antral follicle to become a dominant graafian follicle. In the rat, the duration of follicle development is much shorter than that needed for human follicles. The time required between the initial recruitment of a primordial follicle and its growth to the secondary stage is more than 30 days, whereas the time for a secondary follicle to reach the early antral stage is about 28 days. Once reaching the early antral stage (0.2–0.4 μm in diameter), the follicles are subjected to cyclic recruitment, and only 2 to 3 days are needed for them to grow into preovulatory follicles.

Rights were not granted to include this figure in electronic media. Please refer to the printed book.

Figure 3-5 Landmarks of follicular development during fetal and neonatal life in humans and rodents. In the human ovary, primordial follicles are present by 20 weeks of fetal life, whereas primary follicles are found by 24 weeks. By 26 weeks, some follicles have progressed to the secondary stage. Antral follicles develop in the third trimester and are also seen postnatally when follicle-stimulating hormone (FSH) levels are elevated. After puberty, cyclic increases in serum gonadotropins stimulate the antral follicles to become preovulatory follicles during each menstrual cycle. In the rat ovary, primordial follicles are formed by 3 days after birth when the first wave of follicles begins growth. These follicles progress to the early antral stage during the third week of life when serum gonadotropin levels are elevated. After early antral follicles are formed, ovarian cell apoptosis increases. FSH receptors are found by day 7 of age, when secondary follicles are present, followed closely by the formation of luteinizing hormone (LH) receptors in the thecal cells. Cyclic ovarian function begins around day 35 of age. The neonatal rodent model allows analysis of early follicle development in a synchronized population of growing follicles. Pmd, primordial.

Rights were not granted to include this figure in electronic media. Please refer to the printed book.

The oocyte grows from approximately 12 μm to approximately 80 μm in the fully grown state, which is achieved prior to formation of the antral follicle in sexually mature mice. In general, antral follicles tend to undergo follicular atresia, which is initiated by apoptosis in the granulosa cells, unless they are rescued by increased levels of FSH in the estrus or menstrual cycles in mice and humans, respectively. This cyclic recruitment of antral follicles results in the rescue and continued development of a few follicles, but usually only one dominant follicle prevails and develops to the periovulatory stage in the human menstrual cycle.⁴⁷ Although cyclic recruitment is operative after puberty in humans, most antral follicles ultimately undergo atresia, which results in oocyte death.

Primordial follicles that have not been recruited are thought to remain dormant and be available for recruitment at a later time. Therefore, maintenance of the pool of nongrowing oocytes is necessary to sustain a normal reproductive life span. Müllerian inhibiting substance or antimüllerian hormone, a TGFβ superfamily member well-known for its regulatory role in the sexual differentiation of the reproductive tract during development, is expressed in granulosa cells of growing follicles starting in the perinatal period.⁴⁸ During folliculogenesis, it inhibits the recruitment of primordial follicles to the pool of growing follicles by decreasing the responsiveness of granulosa cells to FSH.⁴⁹ Antimüllerian hormone is recognized as an important clinical marker of the ovarian reserve of viable follicles.

Regulation of Folliculogenesis by Gonadotropins

Preantral follicle development, which involves oocyte growth with limited granulosa cell proliferation, is regulated predominantly by paracrine and autocrine signals. Although gonadotropins are not required for early follicle development in vivo, these follicles can develop in response to FSH in vitro.⁴⁷ In contrast, gonadotropins are required for antrum formation and the rapid granulosa cell proliferation that results in the graafian or antral follicle.

Follicular growth from the primary and secondary follicles to antral stages takes several weeks in the mouse and several months in many larger mammals, including humans. The Graafian follicle can exceed 600 μm and contains more than 50,000 granulosa cells.

Granulosa cells fall into two distinct subtypes—cumulus and mural granulosa cells—that exhibit differential morphologic features and functions. Mural granulosa cells, located in the periantral and outer membrane regions, express FSH receptor (FSHR) and are responsive to FSH.

FSHR comprises a large extracellular domain that binds the FSH hormone with high affinity, and a seven membrane-spanning domain that activates downstream cyclic adenylate cyclase signaling via the heterotrimeric G proteins.⁵⁰ Mice deficient in the FSH β subunit, which normally forms heterodimers with the α subunits, are infertile due to arrested follicular development prior to formation of the antrum.⁵¹

Consistent with the critical role of FSH signaling via its receptor, follitropin receptor knockout (FORKO) mice are also infertile.⁵² At postnatal day 2, there are fewer nongrowing but more growing follicles, but by postnatal day 24, the numbers of resting and growing follicles are both decreased. Therefore, postnatal recruitment of resting follicles into the growing phase appears to be aberrantly accelerated, followed by arrest in further follicular recruitment later on.

Further, no antral follicles are observed. Although these mouse models confirm that FSH signaling is required for follicular development beyond the preantral stage, the phenotype of the FORKO mice also suggests that FSHR signaling is essential in the regulation of the timing and rate of follicular recruitment.⁵² In contrast, luteinizing hormone receptor knockout mice (LuRKO) females are infertile, presumably due to defects in the later stages of folliculogenesis because follicles up to the early antral stage are present, but preovulatory follicles and corpora lutea fail to form.⁵³

FSH and FSHR-mediated downstream intracellular signaling results in gene and protein expression that exemplifies granulosa cell function. For example, antimüllerian hormone, which is implicated in primordial follicle recruitment, exhibits an aberrant expression pattern in FORKO mice.⁵² Although recruitment of theca cells and their expression of the LH receptor and P450 aromatase genes are independent of FSH signaling, expression of these genes in granulosa cells is dependent on FSH signaling.⁵⁴ Similarly, the genes encoding for inhibin and activin subunits, whose homodimers and heterodimers are thought to be important paracrine and autocrine mediators of follicular growth, have decreased expression levels in granulosa cells in the FSHβ knockout model.⁵⁴ Other genes that are dysregulated as a result of FSHβ deficiency include those encoding for androgen receptor (AR), estrogen receptor β (ERβ), and cyclin D2, all of which have been shown to be essential in the final stages of follicular development.⁵⁴

Estrogen Receptors

Two estrogen receptors, ERα and ERβ, are expressed in granulosa cells in the antral follicle. Different knockout models have been generated for these two receptors based on different strategies of gene targeting, but they produced similar phenotypes.⁵⁵ ERαKO females are infertile; ERβKO females are subfertile. In both ERαKO and ERβKO mice, folliculogenesis progresses normally until the large antral stage, when ERβ is critical for ER-mediated granulosa cell proliferation and subsequent ovulation.

In contrast, ERα is not critical for follicular growth but is required for ovulation. The ERαβKO females clarify the question of functional redundancy between the two ERs by demonstrating that in the absence of ERs, antral follicles only have single layers of granulosa and theca cells, multiple small pockets of antral fluid, and oocytes that are dissociated from cumulus cells.⁵⁵

D-type Cyclin

Granulosa cell proliferation in late folliculogenesis also requires cyclin D2, which belongs to the D-type cyclin family of cell cycle regulators that promote the transition from G₁ to S phases via binding to Cdk4 or Cdk6.⁵⁶ Cyclin D2–deficient females have decreased FSH-mediated granulosa cell proliferation, which results in antral follicles that have significantly fewer layers of granulosa cells. This defect in granulosa cell proliferation also results in ovulation failure on stimulation by LH. However, the number of follicles is not decreased, luteinization of theca cells proceeds, and follicles differentiate to become corpora lutea.

We learn from the cyclin D2–deficient mutants that although these final stages of folliculogenesis and ovulation require cyclin D2 via FSHR-mediated activation, specifically of protein kinase A (PKA), oogenesis per se does not require these last rounds of granulosa cell proliferation. In fact, the oocyte not only appears normal, but it is also competent to undergo meiotic resumption and fertilization to produce viable embryos.⁵⁶ Therefore, although oogenesis and folliculogenesis are closely intertwined, oocyte development is a cell-autonomous process to a certain extent, especially in the advanced stages and at periovulation.

THE ROLE OF CUMULUS CELLS IN OOGENESIS

Cumulus cells, also called corona radiata, are specialized granulosa cells that directly line the oocyte. In addition to their supportive role in cytoplasmic maturation of the oocyte, they have important functions in oocyte development, including the maintainenance of meiotic arrest and induction of ovulation. Ovulation is a broad term that encompasses the processes of follicular luteinization, follicle rupture, and meiotic resumption in the oocyte.

A critical step during LH-induced ovulation is the cumulus cell-mediated cumulus expansion. Mural granulosa cells express LH receptor, which allows them to respond to LH by secreting proteins of the epidermal growth factor (EGF) family—amphiregulin, epiregulin, and betacellulin—which are thought to act as paracrine signals that lead to cumulus expansion.⁵⁷ During this process, cumulus cells disperse in the extracellular matrix, which comprises hyaluronic acid, tumor necrosis factor-stimulated gene 6 (TSG6), and serum-derived inter-α-inhibitor, all of which are essential for follicular rupture.⁵⁸

The cumulus expression of Tsg6 and several other proteins implicated in cumulus expansion is regulated by prostaglandin E₂ (PGE₂) via its receptor EP₂.⁵⁸ Consistent with the critical role of PGE₂ in cumulus expansion and ovulation, mice deficient in either EP₂ or cycloxygenase-2 (Cox-2), which is the rate-limiting enzyme for PGE₂, are infertile due to their inability to ovulate.⁵⁸^–⁶⁰

Connexin and Gap Junctions

Cumulus cells mediate many of their important functions by communicating through gap junctions among themselves as well as with the oolemma. These gap junctions are intercellular channels formed by proteins of the connexin family for the diffusion of sugars, amino acids, lipid precursors, nucleotides, metabolites, and signaling molecules. Connexin family members share the same protein domains, including four membrane-spanning domains, two extracellular loops, a cytoplasmic loop, and cytoplasmic N- and C-terminals.⁶¹ In mice, there are at least 17 connexin proteins, whose distinct sequence or length in the cytoplasmic loops and C-terminal tails, as well as heterodimeric and homodimeric coupling, allow functional diversity.

In mice, connexin (Cx) 32, 37, 43, 45, and 57 are expressed in the cumulus–oocyte complex, where they are located between cumulus cells, on cumulus transzonal projections that anchor cumulus cells into the zona pellucida, or on the microvilli or plasma membrane of the oocyte.⁶¹

The critical role of gap junctions in oogenesis is exemplified by the sterility phenotype found in Cx37-deficient mice. Cx37 is made by both the oocyte and granulosa cells, but it may be the only connexin member in cumulus–oocyte gap junctions that is contributed by the oocyte.⁶¹ In the absence of Cx37, follicular development fails at the preantral-to-antral transition, so that most follicles are arrested at the primary follicle stage, and only a few small antral follicles are present. Further, ovulation does not occur despite the formation of numerous corpora lutea.⁶²

Similarly, in vitro experiments show that Cx43, which is also expressed in granulosa cells, is required for folliculogenesis beyond the primary follicle stage.⁶³ Most importantly, oocytes from these two mutant mouse models are meiotically incompetent, which may reflect the requirement of cumulus cell communication in the acquisition of meiotic competence.

These connexin-deficient experimental models demonstrate the critical role of gap junctions in folliculogenesis, but they do not explain the role of cumulus cells in the regulation of meiotic arrest and resumption in the oocyte. Numerous models and hypotheses attempt to explain how and to what extent cumulus cells mediate the high oocyte intracellular cAMP levels that are required to maintain meiotic arrest even after the oocyte has acquired meiotic competence.⁶⁴ Although the complete story remains to be told, several key players have been identified.

MAINTENANCE OF MEIOTIC ARREST

The oocyte is maintained in meiotic arrest via developmental stage-specific mechanisms. Before reaching the fully grown stage, the oocyte is intrinsically incompetent, but once the fully grown oocyte acquires meiotic competence, cumulus cells are thought to contribute at least partially to the maintenance of meiotic arrest.

It is well known that fully grown oocytes isolated from surrounding cumulus cells spontaneously undergo meiotic resumption independent of LH signaling. Although the signal that confers cumulus regulation of arrest has not been identified, the signaling cascade within the oocyte compartment has become much better understood.

GTP-binding Protein

The stimulatory heterotrimeric GTP-binding protein (G_s)-linked receptor 3 (GPR3) is expressed in the oocyte plasma membrane and is thought to mediate cumulus control of meiotic arrest. However, GPR3 is capable of activity independent of LH or cumulus cells also. The role of GPR3 is exemplified by ovarian sections from prepubertal GPR3^−/− females in which meiotic stages beyond prophase I are seen in oocytes in intact early antral follicles in the absence of LH signaling or cumulus expansion.⁶⁵

Similarly, inactivation of G_s by microinjection of an antibody specifically targeting G_s into the oocyte within an intact follicle allows the oocyte to resume meiosis even if cumulus cells remain adherent to the oocyte.⁶⁵ GPR3 maintains activity of G_s in stimulating adenylyl cyclase to produce high cAMP levels, which in turn maintain PKA activity.⁶⁵ Specifically, the isoform adenylyl cyclase 3 expressed by the oocyte is critical in generating sufficient cAMP.⁶⁶ Acting downstream of the GPR3→G_s→cAMP→PKA signaling cascade, PKA-dependent phosphorylation of substrate proteins presumably inhibits meiotic resumption through as-yet unknown mechanisms that include suppression of M-phase promoting factor (MPF) activity; this represents an area of intense investigation.⁶⁴

MEIOTIC RESUMPTION

Whereas LH triggers ovulation by the cumulus cell compartment and release of the oocyte from meiotic arrest, the resumption and completion of meiosis may be regulated intrinsically to a large extent by the fully grown oocyte. Resumption of meiosis requires both decreased levels of intracellular cAMP and activation of MPF.⁶⁴ Decreased production and increased degradation of cAMP in the oocyte appear to contribute to meiotic resumption.

Decreased cAMP production is mediated by the inhibitory G protein, which in turn is activated by an oocyte-expressed G protein-coupled receptor called leucine-rich repeat-containing G protein-coupled receptor 8 (LGR8).⁶⁷ Interestingly, the role of LH in meiotic resumption is demonstrated by this LGR8-mediated decrease in cAMP levels, because LGR8 is stimulated by insulin-like 3, which is expressed and secreted by ovarian theca cells in response to LH.⁶⁷

Phosphodiesterase (PDE)

On the other hand, the degradation and inactivation of cAMP in the oocyte are mediated by the oocyte-expressed cyclic nucleotide phosphodiesterase family member, PDE3A. GV oocytes from PDE3A^−/− mice fail to resume meiosis in vivo and remain in the GV stage even after ovulation. In addition, meiosis does not resume even when fully grown GV oocytes from these mice are isolated from cumulus cells and cultured in vitro.⁶⁸ Thus, the GPR3 and PDE3A knockout mouse models demonstrate their critical and opposing roles in the regulation of meiotic arrest and resumption, respectively. More importantly, these models also prove that ovulation and meiotic resumption, though usually occurring synchronously, are independent processes that can be differentially regulated.

Meiotic Competence and Oocyte Maturation

Meiotic progression requires activation of MPF, which is composed of cyclin B and its protein kinase Cdk1 (also called p34^cdc2).⁶⁹ The requirement of MPF for the G→M transition in mitosis and meiosis is highly conserved across species. Activation and inactivation of many cell-cycle regulators are mediated via phosphorylation, a form of post-translational protein modification of specific residues. The dual phosphatase, Cdc25b, is required for the dephosphorylation of residues 14 and 15 of p34^cdc2, which results in activating p34^cdc2. Oocytes from Cdc25b^−/− mice are unable to resume meiosis and remain arrested in prophase unless rescued by microinjection of Cdc25b mRNA.⁷⁰ Active p34^cdc2 accumulates rapidly through an autoamplification loop. Therefore, the rate-limiting step of MPF activation is thought to be the synthesis and accumulation of cyclin B. During oocyte growth, the level of active MPF increases and eventually reaches the threshold above which the oocyte becomes meiotically competent, which refers to its ability to undergo meiotic resumption.

In general, meiotic competence is related to oocyte size, presumably because cytoplasmic volume reflects the accumulation of synthesized proteins, including cyclin B, p34^cdc2, and Cdc25b, which are essential for meiosis.⁷¹ Oocytes that are smaller than 70 to 80 μm have not reached their full growth potential and have low rates of meiotic competence, whereas those approximately 100 μm are considered fully grown and tend to be able to undergo meiosis. Another predictor of meiotic competence is the presence of a surround nucleolar (SN) chromosomal localization pattern that can be visualized in the live oocyte with culture media containing Hoescht stain.^72,⁷³

Germinal Vesicle Breakdown (GVBD)

The first observable morphologic change of meiotic resumption is GVBD, which is characterized by the sequential disappearance of GV and nucleolus that is frequently concurrent with a shift from central to ectopic localization of the GV. After GVBD, bivalents start to become organized at prometaphase and then align at the metaphase plate in metaphase I.

At the anaphase–metaphase transition, chromosomes begin to segregate, and a cytoplasmic protrusion signifying late anaphase may be visible. MPF activity increases as the oocyte progresses from GV, GVBD, to metaphase I, but then declines transiently between metaphase I and metaphase II.^74,⁷⁵ However, progress beyond metaphase I may not occur in some oocytes if molecular prerequisites are not met.⁷⁶

Metaphase–Anaphase Transition

The metaphase–anaphase transition is regulated by decreased MPF activity, suppression of protein kinase C (PKC) activity, and the activity of many other proteins.⁷⁷^–⁷⁹ Decreased MPF activity is achieved by proteasome degradation of cyclin B, which is required for polar body extrusion (metaphase I–metaphase II) and at metaphase II exit on fertilization.⁸⁰^–⁸³ Like many mitotic and meiotic cell cycle regulators, the proteolysis of cyclin B is mediated by ubiquitin and the anaphase-promoting complex (APC), which are well conserved and required for proteolysis of many proteins across species.

The anaphase-promoting complex is an E3 ubiquitin ligase that transfers ubiquitin “tags” from E2 ubiquitin conjugating enzymes to substrates, such as cyclin B, thereby “tagging” them for recognition and proteolysis by the 26S proteasome.⁸² The essential role of the proteasome in meiosis is further supported by the metaphase I arrest phenotype that is seen in mouse and rat oocytes cultured in media containing chemical inhibitors of the proteasome.^84,⁸⁵

Meiotic Spindle

Achievement of metaphase I depends on the formation of an intact meiotic spindle, which requires the cell cycle regulator, cell division cycle-6 homologue (Cdc6). Insight into this process is provided by oocytes in which Cdc6 expression is abrogated by the RNA interference technique.⁸⁶ These oocytes undergo GVBD, but the meiotic spindle is not formed, and they fail to reach metaphase I. In the absence of the meiotic spindle, condensed chromosomes fail to form bivalents and do not align to form the metaphase plate. Instead, the chromosomes form a halolike structure and meiosis I is arrested with no cell division or extrusion of polar body.⁸⁶

In the presence of a normal meiotic spindle, the oocyte will reach metaphase I and then undergo the metaphase–anaphase transition, which is an important event because defective chromosome segregation at this stage can lead to aneuploidy in the resulting egg and embryo.

In mitosis, the centromeres of sister chromatids are held together by cohesin until the onset of anaphase, when cohesin cleavage by separase allows separation of sister chromatids.⁸⁷^–⁸⁹ Inhibition of separase activity by securin prior to anaphase onset, and its subsequent activation at anaphase upon securin degradation via the APC, are critical in ensuring correct chromosomal segregation.^90,⁹¹ The APC→securin→separase cascade is also required to ensure correct spindle function and homologous chromosome disjunction during the metaphase–anaphase transition in meiosis in the oocyte.^92,⁹³

Spindle Assembly Checkpoint (SAC)

Another mechanism that the oocyte has in common with mitotic cells to ensure the correct segregation of chromosomes is the spindle assembly checkpoint (SAC). SAC proteins detect whether chromosomes are properly aligned and whether kinetochores are attached to the spindle. SAC proteins control the onset of anaphase I through interactions with one another, the kinetochores of unaligned chromosomes, and Cdc20, which is a protein that mediates APC/C proteolysis. (Kinetochores are the sites of attachment of centromeres of the chromosomes to the microtubules of the spindle. Homologous chromosomes migrate to opposite poles through tension exerted by the spindle on the kinetochores.) If chromosome alignment is not complete, the SAC protein would inhibit APC/C to prevent precocious sister chromatid separation, which can cause aneuploidy in the resulting egg and embryo.

Mitotic SAC proteins that also have essential spindle checkpoint functions during maturation of the mouse oocyte include Mad2, BubR1, and Bub1. Interference of the function of these proteins results in accelerated completion of meiosis I, presumably because anaphase occurs prematurely.⁹⁴ Further, the abrogation of Mad2 expression has specifically been shown to increase the rate of aneuploidy, whereas its overexpression inhibits disjunction of homologous chromosomes.⁹⁵ Therefore, the expression and function of these cell-cycle regulators are candidates to be investigated for their potential involvement in age-related aneuploidy in human oocytes.

At the end of anaphase I, the first polar body is extruded, which is followed by reaccumulation of cyclin B levels. MPF reactivation is required for the synthesis of c-mos, which leads to activation of MAP kinase kinase and MAP kinase.⁹⁶^–¹⁰⁰ This c-mos-mediated MAP kinase activity is critical for the maintenance of metaphase II arrest, as shown by the c-mos^−/− mouse model. Unfertilized eggs from c-mos^−/− mice fail to arrest at metaphase II, and undergo parthenogenetic activation, during which the second polar body is extruded in the absence of fertilization.^101,¹⁰²

NUCLEAR AND CYTOPLASMIC CHANGES DURING OOCYTE GROWTH

The growing oocyte undergoes nuclear and cytoplasmic changes that pertain to nuclear organization, epigenetic regulation, transcription, translation, ultrastructural morphology, and organelle function. These changes are sometimes referred to as nuclear or cytoplasmic maturation, broadly defined based on their association with oocyte maturation, subsequent fertilization, or embryo development. Many of the morphologic changes of the oocyte have been elegantly described at the cellular and ultrastructural levels.¹⁰³ Further, expression, translational modification, and function of cell-cycle regulators, also considered indices of cytoplasmic maturation, are discussed in this chapter under “Meiotic Competence and Oocyte Maturation.” In this section, we highlight some biochemical features characteristic of the oocyte around the time of maturation.

Transcriptional Regulation

The growing oocyte is characterized by a high level of transcriptional activity that increases its total RNA content from approximately 0.2 ng in a small oocyte to approximately 0.6 ng in a fully grown oocyte.¹⁰⁴ In growing oocytes, gonadotropin stimulation of follicles increases the proportion of transcriptionally inactive oocytes; thus granulosa cells are thought to have an important role in the regulation of transcriptional activity in the growing oocyte.¹⁰⁵ The fully grown oocyte undergoes global transcriptional silencing, which is a prerequisite for meiotic resumption, oocyte maturation, and subsequent embryo development.¹⁰⁶ Therefore, messenger RNAs (mRNAs) that are synthesized during oocyte growth are not only utilized for the translation of proteins during the growth phase, but are also stored for protein synthesis in the fully grown stage, during oocyte maturation, or later in the early embryo. Thus, precise control of translation is vital to the function and ultimate developmental potential of the oocyte and embryo.

Regulation of Translation

Translation requires modification of the mRNA transcript such as polyadenylation at the 3′ end (addition of a poly-A tail) and cap binding at the 5′ end. Such modifications are controlled by cis and trans elements. For example, the translation and accumulation of cyclin B1 is regulated via polyadenylation and cis-regulatory mechanisms in the 3′ untranslated region (3′UTR) of the cyclin B mRNA transcript.¹⁰⁷

Cis elements in the 3′UTR consist of a highly conserved U-rich sequence (CPE) in conjunction with a downstream hexanucleotide cytoplasmic polydenylation element, AAUAAA. Cis elements mediate polyadenylation via binding to the phosphorylated form of the CPE binding protein (CPEB), which is a trans-acting factor specific to the oocyte. The critical function of CPEB in the oocyte was demonstrated in CPEB knockout mice.¹⁰⁸ In the absence of CPEB oogenesis arrested at the pachytene stage on 16.5 dpc, presumably because translation of SCPs 1 and 3 did not occur, which led to death and resorption of the oocytes.^108,¹⁰⁹ However, the requirement of CPEB at this early stage also precluded the function of this protein during oocyte maturation to be ascertained.

Other regulatory mechanisms of translation involve inhibition of translation if the presence of certain proteins would interfere with oocyte maturation. For example, some mRNAs can be stored in the form of ribonuclear particles (RNPs), which do not support translation and prevent the misexpression of certain proteins.¹¹⁰

The importance of this mechanism is shown by the relative abundance of the Y-box protein family member, MSY2, which has an essential role in mediating translational silencing by RNPs and comprises 2% of the total protein content in the fully grown oocyte.^111,¹¹² In addition, poly-A-specific ribonuclease (PARN) mediates widespread deadenylation and subsequent degradation of mRNAs encoding for a variety of proteins, including zona pellucida proteins, actin, alpha tubulin, and Cx43, at the time of meiotic resumption.¹¹⁰ In contrast, the polyadenylation of certain cell-cycle proteins required during oocyte maturation (i.e., cyclin B1) is enhanced at meiotic resumption.¹¹³

Epigenetic Regulation

In addition to regulation at the levels of transcription and translation, epigenetic regulatory mechanisms in the oocyte, sperm, and early embryo affect the expression of specific genes through the process of genomic imprinting, as well as the entire molecular program through global changes in methylation status and structural chromatin remodeling. Structural remodeling of chromatin is associated with significant changes in the nuclear architecture during oocyte growth and maturation.

Chromosomes in the live mouse oocyte can be viewed using culture media containing Hoescht stain, a chemical that chelates with the minor groove of DNA and emits blue fluorescent light upon UV light absorption. Centromeres and pericentromeric heterochromatin are initially located at the periphery of the nucleus in oocytes in primordial and primary follicles.¹⁰⁶ Then during oocyte growth, they are spread within the nucleus, followed by localization to the periphery of the nucleolus. This perinucleolar heterochromatin rim, or karyosphere, appears as a bright rim around the nucleolus and is also known as the surround nucleoli (SN) pattern.^73,¹¹⁴

Although the SN pattern is associated with global transcriptional repression and high potential for meiotic competence and embryo development, we now know that chromatin remodeling and transcriptional repression are distinct processes controlled by different mechanisms.¹⁰⁶ As in somatic cells, histone deacetylases were also found to be critical for the large-scale chromatin remodeling in the fully grown oocyte because their inhibition resulted in a breakdown of the SN architecture and aberrant configuration of the meiotic chromosomes and spindle.¹⁰⁶

Another important epigenetic mechanism that has emerged is control of transcriptional activation and repression via demethylation and methylation, respectively. Whereas the genome of somatic cell precursors undergoes remethylation before gastrulation at 6.5 dpc, the genome of PGC precursors remains demethylated at 12.5 dpc. Then partial methylation occurs at 15.5 dpc and methylation is completed by 18.5 dpc.³² The embryonic genome is inactivated until the two-cell stage in mice (eight-cell stage in humans), when it becomes activated by global demethylation.

These global changes in methylation status are distinct from X-inactivation and genomic imprinting. Briefly, X-inactivation is a complex and random process in which one of the X chromosomes in XX mammalian females is inactivated to ensure an equivalent dosage of X-linked genes in XX females and XY males.¹¹⁵ Initiation of this process is controlled by a locus called X-inactivation center (Xic) at Xq13. This locus contains X-inactive specific transcript (Xist), which encodes a non-coding mRNA that coats the X-chromosome in cis to trigger inactivation¹¹⁵ (see Chapter 5).

Genomic Imprinting

Finally, a more gene-specific mechanism of epigenetic regulation is genomic imprinting, which mediates predesignated, differential silencing of the maternal and paternal alleles.¹¹⁶ During gametogenesis, certain genes on autosomes become hypermethylated, or silenced, based on their parental origin. Once established, these imprints are thought to be protected from genome-wide demethylation.

Imprinting possibly occurs at different time points in oogenesis, spermatogenesis, and embryogenesis for different genes. Congenital syndromes or cancer can result from a failure of the allele from the designated parental origin to be silenced or an effective null mutation based on a heterozygous mutation in the nonimprinted allele. These disease mechanisms and syndromes are further discussed in Chapter 5.

The mechanism of imprinting is an area of research that is particularly relevant to reproductive endocrinology and infertility treatment because of reports associating assisted reproductive technologies (ART) with possible increased risk of imprinting defects.¹¹⁷ In addition, it has been suggested that in the mouse model, oocyte maturation and embryo culture in vitro may interfere with imprinting.^118,¹¹⁹ However, because defects in imprinting are rare in the general population and in children conceived through ART, it is difficult to determine the statistical significance in these studies. Consequently, whether there is a true association or a causal link remains an open question.

If ART is indeed associated with imprinting defects, one would have to examine all possible causes, including controlled ovarian hyperstimulation (COH), clinical embryology, and the cause of infertility itself, because imprinting defects may be the cause of infertility that becomes unmasked by infertility treament. Therefore, this area warrants intense investigation to determine whether and how mechanisms of imprinting impact on future directions in ART.

THE OOCYTE–EMBRYO TRANSITION

In addition to the maternal genome, the oocyte contributes specific gene products that are required for early embryo development prior to zygote gene activation. In the mouse model, zygote genome activation occurs at the two-cell stage, while in the human it occurs at the four- to eight-cell stage.

Maternal Effect Genes

Because the oocyte does not synthesize new mRNA transcripts after meiotic resumption and the embryonic genome is not activated until the two-cell stage, the oocyte must store transcripts and proteins for all of its requirements during the oocyte–embryo transition. Genes encoding oocyte proteins that persist and have critical functions in the early stages of embryogenesis are called maternal effect genes.

It is not surprising, therefore, that several proteins encoded by maternal effect genes—oocyte-specific linker histone H1 (H1F00), nucleoplasmin 2 (Npm2), DNA methyltransferase1 oocyte isoform (Dnmt1o)—are involved in mechanisms of epigenetic gene regulation; these processes are initiated in the oocyte, but continue during early embryo development.¹²⁰^–¹²⁴ All of these maternal effect genes, and potentially ones that have yet to be discovered, serve as critical links to demonstrate how the unfertilized oocyte can affect subsequent embryo development, besides contributing to the embryonic genome.

H1FOO is an oocyte-specific H1 linker histone that associates with chromatin during the GV and maturation stages; this association continues until the late two-cell to four-cell embryo stages when they are gradually replaced by somatic H1. Because somatic H1 linker histones are important in the regulation of chromatin function, the oocyte- and early embryo-specific expression of H1FOO suggests that it may be critical in chromatin remodeling during the oocyte–embryo transition.^120,¹²¹

In contrast, Npm2, a protein that is present in the nuclei of oocytes and peri-implantation embryos, is proven to have an essential role in chromatin structure. Embryos of Npm2^−/−mutant females lack Npm2, and die before implantation because Npm2 is required for the maintenance of normal nucleolar structures. Specifically, heterochromatin and deacetylated histone H3, which are usually localized to the rim of the nucleolus in oocytes and early embryos, are absent.¹²² Therefore, this mouse model also demonstrates the importance of chromatin regulation and nuclear architecture in early embryo development.

Another maternal effect gene that is important in epigenetic regulation is Dnmt1. The oocyte variant, Dnmt1o, is an isoform of the Dnmt1 protein that is expressed only in the growing oocyte and early embryo, under an oocyte-specific promoter. Embryos derived from Dnmt1o-deficient oocytes are also deficient in Dnmt1o and develop into fetuses that die during gestation.¹²³ These embryos have defective imprinting, as shown by inappropriate methylation at certain gene loci and loss of allele-specific gene expression. The critical function of Dnmt1o is thought to occur at the eight-cell stage, when the protein transiently translocates from its usual location in the cytoplasm to the nucleus.¹²³

Zar1-null mutant females appear to have normal oogenesis and fertilization appears to be initiated, but the two pronuclei remain separated and most embryos arrest at the one-cell stage.¹²⁵ Further, of the small percentage of embryos lacking Zar1 that survive to the two-cell stage, there is decreased expression of proteins in the transcription-requiring complex, which indicates a defect in embryonic genome activation. Whereas Zar1 is critical for progression to the two-cell stage, Mater, another protein encoded by a maternal effect gene, is expressed in oocytes and preimplantation embryo and is critical for embryonic development beyond the two-cell stage.¹²⁶ Investigation into the functions of these proteins will contribute to our understanding of mechanisms that control the merging of paternal and maternal genomes, embryonic genome activation, and early embryo development.

Polarity and Cytoplasmic Reorganization

Unlike other model organisms such as the fruit fly, the determination and establishment of cell polarity, or asymmetric distribution of cytoplasmic contents, in the mammalian oocyte and embryo is not well understood. Part of the difficulty is that although polarity in the form of an eccentric GV is observed in oocytes of some mammalian species, such as human, Rhesus monkey, bovine, and porcine, GV oocytes isolated from mouse and rat ovarian follicles do not demonstrate polarity, perhaps due to artificial effects of the in vitro culture system.¹²⁷

The growing oocyte has one dominant microtubule organizing center (MTOC). The MTOC comprises centrosome, which organizes microtubules in an astral configuration. In mitotic cells, centrosomes are made up of a pair of centrioles, which are cylindrical structures containing nine triplets of microtubules that form the astral fibers required for cell division. However, centrosomes in the oocyte do not have centrioles and are thus called MTOCs. The fully grown, meiotic competent oocyte has multiple MTOCs that mediate migration of the GV from the center to an eccentric, cortical position during oocyte maturation. The eccentric position of the GV is thought to be related to or to determine the location of the meiotic spindle, which in turn determines the site of polar body extrusion.¹²⁷

The eccentric position of the meiotic spindle defines the animal–vegetal axis, in which the animal pole contains the spindle-associated cortex, and the vegetal pole is characterized by clusters of endoplasmic reticulum. This asymmetric organization of the cytoplasm is thought to be critical in ensuring that both cell divisions at meiosis I and meiosis II will produce polar bodies, rather than daughter cells of equal size. This asymmetry in the generation of the egg is well conserved across species and is thought to be nature’s way of maximally conserving resources for the resulting embryo.

It is controversial whether the meiosis II cell division further determines the point of fusion between sperm and egg, the position of the first mitotic spindle, and even the spatial organization of the first mitotic division.¹²⁸ The mos/MAP kinase pathway, which regulates the metaphase II arrest, is also required for transitioning the meiotic spindle from a cortical location back to the central location of a mitotic spindle.¹²⁹ Therefore, determinants of oocyte polarity likely have as-yet unrecognized impact on the development and survival of the early embryo. Indeed, the cytoplasm undergoes drastic reorganization from the asymmetric oocyte and egg, to become an embryo that is capable of symmetric cell divisions.

APOPTOSIS: THE ULTIMATE CELL FATE FOR MOST OOCYTES

The oocyte is perhaps the most limiting factor in human reproduction because only about 300 to 400 eggs are ovulated during a woman’s entire reproductive lifespan. In other words, more than 99.9% of all oocytes undergo programmed cell death, or apoptosis, without ever having the opportunity to become fully grown, ovulate, or fertilize.

The first wave of apoptosis in the mouse model is thought to occur when germline cysts containing oogonia break down, and a similar process likely occurs in the human female fetus. The number of oocytes reaches a peak of approximately 7 million by midgestation in humans.^32,¹³⁰^–¹³³ Subsequently, oocytes that do not become encapsulated by granulosa cells undergo apoptosis. Further, growing follicles undergo atresia as they reach the antral stage, so that there are approximately 300,000 to 400,000 follicles at birth and about 200,000 follicles remaining by puberty.^132,^134,¹³⁵

Although apoptosis is the most common cell fate, the initiating factor and the mechanism may differ depending on when it occurs during development. For example, the apoptosis that affects primary oocytes before their incorporation into primordial follicles is thought to be oocyte-autonomous, whereas oocytes in growing preantral and antral follicles die secondary to follicular atresia that is caused by apoptosis of the granulosa cells.

Many hypotheses have been raised in an attempt to explain the high attrition rate, including that it is advantageous for the species to eliminate germ cells containing chromosomal or other errors. However, no hypothesis has been rigorously tested and the question remains open.

Environmental Factors

In addition to apoptosis that is part of “normal” development, environmental factors and genetic susceptibility can exacerbate depletion of the oocyte pool, which is often manifested clinically as premature ovarian failure or early menopause. Environmental factors that have demonstrated oocyte toxicity include organochlorine chemicals from pesticides and industrial processes,¹³⁶ polycyclic aromatic hydrocarbons from tobacco smoke or fuel combustion,^137,¹³⁸ and chemotherapeutic agents (i.e., doxorubicin) that activate the apoptotic pathway via ceramide, a lipid that is generated by acid sphingomyelinase.¹³⁹ The harmful effect of ceramide is further supported by the ability of its metabolite and antagonist, sphingosine-1-phosphate, to confer protection in preventing oocytes from undergoing apoptosis when exposed to chemotherapy.^140,¹⁴¹

Genetic Factors

Genetic mouse models of oocyte apoptosis may exhibit phenotypes varying from subfertility (i.e., small litter size or infrequent litters), premature ovarian failure, or infertility due to congenital ovarian atresia. Broadly, genetic defects may primarily be due to structural or aneuploid chromosomal abnormalities; defective X chromosome inactivation; mutation in single genes that are essential in oogenesis, folliculogenesis, or gonadogenesis; or single-gene defects affecting apoptotic pathways in the ovary. Examples of the latter are highlighted here; the other genetic abnormalities are discussed elsewhere in this book. Most importantly, apoptosis should be recognized as a common final pathway that will follow defects interfering with oogenesis.

As in somatic cells, appropriate expression of pro-apoptotic proteins and those protecting against apoptosis is required to maintain the “normal” rate of attrition. Bcl-2 and Bcl-x proteins are expressed in the oocyte and granulosa cells and “protect” the oocyte from apoptosis. Although Bcl-2-deficient mice are fertile with normal litter size, there is a decrease in the number of primordial follicles as well as an increase in the number of abnormal primordial follicles that contain no oocyte or a dying oocyte.¹⁴² Similarly, Bcl-x hypomorphic mutant mice have significantly decreased numbers of primordial and primary follicles, but their phenotype is more severe and fertility is impaired.¹⁴³ Therefore, these two genes are required for the maintenance of both the oocyte pool and follicular architecture. In contrast, targeted overexpression of Bcl-2 in granulosa cells results in enhanced folliculogenesis, larger litter size, and an increased incidence of germ cell tumors with age.¹⁴⁴

Interestingly, female mice that are deficient in Bax are fertile and have significantly more primordial follicles than wild type mice even with advancing age. Further, they are able to superovulate in response to exogenous gonadotropins even at an advanced age, and these oocytes develop into viable embryos.¹⁴⁵ Similar results were seen in the mouse model in which the pro-apoptotic gene caspase-2 is deleted. In addition, in the absence of caspase-2, oocytes do not undergo drug-induced apoptosis on exposure to the chemotherapeutic agent doxorubicin.¹⁴⁶

These data support the relevance of this research area in understanding and potentially extending the female reproductive lifespan. Most importantly, understanding the regulation of apoptosis in the mammalian oocyte is vital to the development of oocyte- or follicle-sparing chemotherapeutic and other pharmaceutical agents.

CLINICAL RELEVANCE AND FUTURE DIRECTIONS

Many key challenges in clinical reproductive medicine today relate to the availability and function of human oocytes and the follicles that enclose them. However, we are very far from understanding what proportion of “unexplained infertility” is due to an oocyte-specific factor and even further from having any oocyte-specific treatment.^147,¹⁴⁸ Significant advances in our management of oocyte-related infertility will require a better understanding of the mechanisms through which maternal age, environmental effects, and patient-specific genetic susceptibility impact on oocyte function.

Human Gene Mutations and Oocyte Phenotypes

Human and mouse genetic models of oocyte apoptosis may exhibit phenotypes varying from subfertility, premature ovarian failure, or infertility due to congenital ovarian atresia. Although these conditions are usually considered distinct clinical entities that are managed differently, their genetic basis may be closely related. Broadly, genetic defects may primarily be due to structural or aneuploid chromosomal abnormalities involving the X chromosome (i.e., Turner’s syndrome), defective X chromosome inactivation, and mutation or epigenetic defects involving single genes that are essential for oogenesis, folliculogenesis, or gonadogenesis. In particular, familial or de novo single-gene mutations affecting any stage of oogenesis can theoretically result in a spectrum of conditions varying from congenital ovarian dysgenesis, to premature ovarian failure, to “poor oocyte quality” identified during COH or IVF treatment.

Although gene targeting in experimental mouse models has contributed significantly to our understanding of oogenesis, it is critical that we increase our effort to relate human phenotypes encountered in clinical reproductive endocrinology and infertility treatment, such as “poor ovarian response” and developmental abnormalities of oocytes and embryos, to our understanding of the mechanisms regulating oogenesis.

Premature Ovarian Failure

Of all oocyte- or ovarian follicle-related conditions, identification of human gene mutations has thus far largely focused on familial premature ovarian failure. Using cytogenetic and fluorescent in situ hybridization techniques, familial cases of premature ovarian failure associated with balanced translocation could be studied to identify candidate genes that are disrupted by the translocation breakpoint. For example, many autosomal dominant mutations in FOXL2, a transcription factor that is expressed only in granulosa and eyelid cells, have been identified as the cause of a syndrome of premature ovarian failure and eyelid malformation called blepharophimosis ptosis epicanthus inversus syndrome.¹⁴⁹^–¹⁵³

Another genetic mutation that has been linked to familial hypergonadotropic gonadal failure in humans is localized to the X-linked gene BMP15.^45,¹⁵⁴ Further, the “critical region” on the long arm of the X chromosome (Xq13-Xq26) is proposed to contain several genes that are required for oogenesis or ovary development. A familial case of premature ovarian failure has been linked to a gene located in this critical region, DIA, which is the human homolog of the Drosophila diaphanous gene that is critical for oogenesis in Drosophila.¹⁵⁵ Because mouse homologs of FOXL2 and BMP15, as well as the D. diaphanous gene, have critical functions in oogenesis or folliculogenesis, these examples underscore the value of connecting what we learn from animal models and scenarios encountered in the clinic.

Familial hypergonadotropic ovarian dysgenesis has also been identified in patients with autosomal recessive, inactivating point mutations in the FSHR gene,¹⁵⁶^–¹⁵⁸ which presumably abrogated follicular response to FSH, resulting in atresia and premature ovarian failure. Conversely, there are also reports of various FSHR mutations that increase the sensitivity of the FSH receptor to FSH, human chorionic gonadotropin (hCG), and thyroid-stimulating hormone (TSH), which led to spontaneous ovarian hyperstimulation syndrome and spontaneous superovulation.¹⁵⁹^–¹⁶⁵

These human gene mutations serve as spontaneous gene targeting models that demonstrate the role of these genes in human reproduction in vivo. Further, biochemical investigations of proteins encoded by these genes may provide insight into the pathogenesis of, and potential therapy for poor ovarian response to COH and the prevention of premature ovarian failure and ovarian hyperstimulation syndrome.

In the distant (or not so distant) future, it is conceivable that as more human genetic mutations causing ovarian dysfunction are identified, genetic screening panels combined with proper counseling may help predict risks of premature “oocyte aging” to help young women with their family planning. Similarly, poor responders and patients at risk for ovarian hyperstimulation syndrome may be identified before their COH/intrauterine insemination or IVF treatment. Establishment of reliable predictors will not only help optimize cycle outcomes, but will also be valuable in counseling couples regarding their treatment options and risks of multiple gestation and ovarian hyperstimulation syndrome.

Aneuploidy

Abnormal chromosomal segregation in meiosis results in aneuploidy, which increases in incidence with advancing maternal age in humans. Aneuploidy, considered “the most important problem in human reproduction,” affects an estimated 25% to 50% of human eggs, accounts for at least 35% of spontaneous abortions and approximately 4% of stillbirths, and is the leading genetic cause of congenital mental retardation and developmental disabilities.¹⁶⁶^–¹⁷²

The key risk factors for maternally derived aneuploidy in humans are maternal age and susceptible patterns of meiotic recombination, such as recombination sites that are close to the centromere or the telomere (reviewed in Lamb and colleagues ¹⁷³). Compared to the mouse model, there is an extreme variability in the location of recombination nodules in human oocytes,²⁶ which may explain the high rates of aneuploidy in our eggs and embryos. However, although susceptible locations of recombination nodules are thought to be the main risk factor in young women, they do not explain the high aneuploidy rates seen with increasing maternal age.¹⁷³ In fact, the association between these recombination patterns and aneuploidy decreases with increasing age. Therefore, as-yet undefined recombination-independent factors are proposed to be risk factors for older women.¹⁷³

IN VITRO FERTILIZATION

IVF allows us to see the entire developmental process from GV, GVBD, to metaphase II arrest, formation of the one-cell embryo with two pronuclei, all the way to the blastocyst stage. Although we expect to see some abnormal oocytes or embryos during IVF treatment, when an unusual oocyte or embryo phenotype is consistently observed in the same couple, especially if they have “unexplained infertility,” we are alerted to the possibility that those abnormalities may be related to the underlying cause of the couple’s infertility.

Repeated IVF treatment failure in these patients would further identify them as the patient population that may benefit from having a molecular diagnosis and nonempirical treatment that specifically targets the molecular diagnosis. For example, recurrent or familial cases of zona pellucida abnormalities,¹⁷⁴ maturation arrest,¹⁷⁵^–¹⁷⁹ parthenogenetic activation,¹⁸⁰ triploidy,¹⁸¹ abnormal fertilization,¹⁸² abnormal cell division in meiosis II,¹⁸³ and empty follicle syndrome^184,¹⁸⁵ have been reported. Indeed, the development of molecular diagnosis and treatment for this subset of patients is a major challenge in reproductive medicine.

FUTURE DIRECTIONS

Outcome-based, systematic surveys of human oocyte and embryo phenotypes encountered in IVF treatment are critical in guiding clinicians and scientists to focus on conditions that are prevalent and relevant in patients seeking infertility treatment. Further, although mammalian oocyte biology research has traditionally been based on the candidate gene approach, complementary experimental strategies by more unbiased, forward genetic methods such as random chemical mutagenesis of the mouse genome followed by screening for infertility phenotype has begun to uncover novel genes that have critical functions in oogenesis.¹⁸⁶ Newly identified genes with potentially critical functions in oogenesis can then be further studied by various gene targeting techniques.

Another forward genetic approach is chemical genetics in the form of high throughput small molecule (SM) library screening of processes such as oocyte maturation and follicular development. SM library screening followed by functional experiments have led to the identification of lead compounds in drug discovery and the identification of novel proteins in other areas of research.¹⁸⁷^–¹⁸⁹ Although the technical challenges involved in the high throughput experimentation of the oocyte or follicle may seem quite overwhelming, they may not be truly insurmountable. We can look beyond medicine and biology, and engage bioengineers to bring microfluidics and other microsystems-based technologies to help us accelerate discovery in the field of oocyte biology and infertility.¹⁹⁰ Such interdisciplinary effort will be instrumental in giving us the ability to treat or prevent oocyte-related reproductive problems. It will also facilitate the development of oocyte- or fertility-sparing pharmaceuticals and the identification of environmental aneugens to direct preventive strategies to preserve reproductive health.

New findings on functions and human mutations of ovarian genes can be searched online on several databases, including the Ovarian Kaleiodoscope (http://ovary.stanford.edu/) and Online Mendelian Inheritance in Man (OMIM) (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=OMIM),¹⁹¹ OMIM and Online Mendelian Inheritance in Man are trademarks of the Johns Hopkins University.

PEARLS

• During prophase of meiosis I, oocytes undergo DNA replication, homologous pairing, synapse, and recombination, but then remain arrested at the diplotene stage of prophase I until sexual maturity.

• Primordial germ cells are derived extragonadally from the yolk sac endoderm. They migrate from the base of the allantois to the genital ridges.

• Members of the TGF superfamily of growth factors are important signaling molecules for the migration and survival of primordial germ cells.

• Primordial germ cells differentiate into oogonia, which are sexually differentiated germ cells.

• Oogonia become oocytes by entering into prophase of meiosis I.

• Primordial follicles are diplotene-arrested oocytes surrounded by a single layer of flattened granulosa cells.

• Primary follicles are characterized by a single layer of cuboidal granulosa cells.

• Prophase I has four stages: leptotene, zygotene, pachytene, and diplotene.

• The transition from pachytene to diplotene marks the beginning of folliculogenesis.

• Oocytes arrest at the diplotene stage, when homologous chromosomes are held together at chiasmata.