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Chapter 3 Oogenesis


Oogenesis is an area that has long been of interest in medicine, as well as biology, economics, sociology, and public policy. Almost four centuries ago, the English physician William Harvey (1578–1657) wrote ex ovo omnia—“all that is alive comes from the egg.” Oogenesis exemplifies many fundamental biologic processes, such as mitosis, meiosis, and intracellular and intercellular signaling. The process of oogenesis begins with migratory primordial germ cells (PGCs) and results in an ovulated egg containing genetic material, proteins, mRNA transcripts, and organelles that are essential to the early embryo.

Although our understanding of these basic mechanisms has increased significantly over the past few decades, we are still limited in our ability to treat female infertility due to “poor oocyte quality” or to identify women at risk for this condition to allow more patient-specific family planning. Currently, poor oocyte quality refers to a broad spectrum of clinical conditions, including poor response to exogenous gonadotropins, recurrent pregnancy loss or congenital anomalies due to aneuploidy, and abnormalities of oocyte maturation, fertilization, and early embryo development that are observed during in vitro fertilization (IVF) treatment. Because these conditions are more prevalent in older women, they also carry serious social implications for women and their partners in family planning.

Many aspects of oogenesis and meiosis are well conserved across sexually reproducing species. Experimental work on the nematode Caenorhabditis elegans, the fruit fly Drosophila melanogaster, the frog Xenopus laevis, and the fission and budding yeasts Saccharomyces cerevisiae and S. pombe, respectively, has provided us with much insight into these processes. However, there are key differences in oogenesis between mammalian and nonmammalian species. As with many processes in development, oogenesis is extremely similar between mice and humans. Thus, in vivo gene-targeted and in vitro oocyte maturation models in the mouse have been most informative and relevant to our understanding of mammalian oogenesis.

The generation of healthy eggs with the correct genetic complement and the ability to develop into viable embryos requires that oocyte development and maturation be tightly regulated. In an effort to describe in detail our current understanding of the basic biologic mechanisms behind this process, this chapter discusses mammalian oogenesis as it is understood in the mouse model (Fig. 3-1) and highlights clinical conditions that are related to defects in oogenesis.

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Figure 3-1 Stages of oocyte maturation. Oocytes and their corresponding chromosomes at various stages of maturation, as visualized by differential interference contrast (DIC) [A, C, E, G, I, K] and 4′6-diamidino-2-phenylindole (DAPI) stain [B, D, F, H, J, L], respectively. DAPI stain binds to natural double-stranded DNA in the chromosomes and allows chromosomes to be visualized in live cells. A, An oocyte at the germinal vesicle (GV) stage (center). GV, germinal vesicle; N, nucleolus; ZP, zona pellucida. B, The same oocyte under milliseconds of exposure to UV light, upon which DAPI-stained chromosomes are visualized as fluorescent structures. The chromosomes (arrow) are localized to the periphery of the nucleolus and assume the “surrounding nucleoli” configuration indicative of meiotic competence (see text for detailed explanation). C, D, An oocyte that has undergone germinal vesicle breakdown and is in pro-metaphase I, as indicated by complete disappearance of both GV and nucleoli on DIC and the presence of condensed bivalents (arrow). E, F, An oocyte in metaphase I. The chromosomes (arrow) are aligned at the metaphase plate and individual bivalents cannot be distinguished. G, H, An oocyte that has been cultured in media containing the proteasome inhibitor MG132 (2.5 μM) for 24 hours. Proteasome inhibition causes this oocyte to be arrested at metaphase I, and congression of chromosomes (arrow) remains at the metaphase plate.

I, J, An oocyte (top) in the metaphase–anaphase transition, as indicated by chromosomes that are in the process of segregating. The bottom oocyte has extruded the first polar body, which is indicated by the condensed chromosomes (single spot). (The chromosomes in the oocyte proper are not seen on this plane.) K, L, An oocyte (center) that is in metaphase arrest, which is indicated by extrusion of the first polar body. Note a very faint fluorescent spot, which represents very condensed chromosomes that are tightly localized in the first polar body. Chromosomes in this metaphase II-arrested oocyte are aligned at the metaphase plate. These chromosomes appear fainter and not in optimal focus because their focal planes are slightly different. (Due to the thickness of the oocyte [≈80–100 μm], optimal visualization of both sets of chromosomes may not be possible if they are in different focal planes.)


Oogenesis begins in the early embryo with PGCs, which migrate from the yolk sac to the urogenital ridge to establish the germline. Upon sexual differentiation, PGCs give rise to oogonia, which undergo mitosis to expand the population in the developing female gonad. Oogonia become oocytes as they enter meiosis I.

During prophase of meiosis I, oocytes undergo DNA replication, homologous chromosome pairing, synopsis, and recombination, but then remain arrested at the diplotene stage of prophase I until sexual maturity. While the oocyte remains in meiotic arrest, it continues to increase in size coordinately with follicular growth. Before meiotic resumption, the fully grown oocyte is characterized by the presence of the germinal vesicle (GV), which is the meiotic counterpart of the nuclear envelope in somatic cells.

Gonadotropin-dependent follicular growth is followed by oocyte maturation, which refers to the resumption and completion of meiosis I in the oocyte. This process occurs at each estrus cycle in mice and at each menstrual cycle after the onset of puberty in humans. On mating in an estrus cycle in the mouse, luteinizing hormone (LH) and cumulus–oocyte signaling result in decreased cyclic adenosine monophosphate (cAMP) in a cohort of oocytes. These oocytes resume meiosis and undergo GV breakdown (GVBD). Condensation of chromosomes is followed by congression of homologous chromosomes at metaphase I and segregation at anaphase I. Extrusion of the first polar body signifies exit from meiosis I and concomitant entry into meiosis II, whereupon the eggs are arrested at metaphase II until fertilization with sperm.

After fertilization, they exit meiosis II and transition into one-cell embryos. In the human menstrual cycle, a cohort of follicles is similarly recruited to resume meiosis, but typically only one dominant follicle develops, so that only one oocyte completes meiosis I and is released during the endocrine- and paracrine-mediated process of ovulation.


Primordial germ cells are the founder cells of the gametes. In many species, including Drosophila, Danio rerio (zebrafish), and C. elegans, germ cells are derived from germ plasm, which is maternally derived cytoplasm that contains germ cell determinants. In contrast, mammalian PGC fate is thought to be induced by cell–cell interaction requiring the adhesion molecule E-cadherin and signaling from surrounding tissues.1,2

The migration of PGCs from the base of allantois to the genital ridges has been mapped out by alkaline phosphatase staining; more recently, their migration in vivo has been visualized by green fluorescent protein (GFP) expressed under the control of a truncated Oct-4 promoter (GFP:Oct-4) acting specifically in PGCs.3,4 Primordial germ cells are derived extragonadally from yolk sac endoderm and part of the allantois that arises from the posterior part of the primitive streak.

Primordial germ cells can be distinctly identified at 7.5 days post-coitum (dpc, a measure of embryonic age) during gastrulation by their high expression of alkaline phosphatase or GFP:Oct-4. At 7.5 dpc, PGCs are seen at the root of the allantois, from which they migrate along the endoderm via passive transfer mechanisms, to arrive at the epithelial layer of the hindgut at 8 dpc. Then, with the acquisition of motility via long pseudopodia, they traverse through the wall of the hindgut at 9 dpc.3

At 9.5 to 11.5 dpc, they migrate along the dorsal mesentery toward the genital ridges in the roof of the coelomic cavity. PGCs remain sexually undifferentiated until 12.5 dpc, when they coalesce with the mesodermally derived somatic components of the gonads to form ovaries or testes. During the process of migration, PGCs undergo proliferation by mitosis at approximately every 18 hours, so that their population increases from fewer than 100 founder cells to approximately 3,000 cells by 11.5 dpc and approximately 25,000 cells by 13.5 dpc.

Specific Genes Expressed for Migration of Primordial Germ Cells

Several genes are known to be required for PGC cell fate determination, migration, and survival in the mouse embryo. Bone morphogenetic protein 4 (Bmp4) and Bmp8b are maximally expressed in the extraembryonic ectoderm adjacent to the proximal epiblast, where precursors of PGCs are localized at 6.5 dpc.5,6 These two proteins are thought to have essential roles in the differentiation of PGC precursors into PGCs because PGCs are absent in both Bmp4 and Bmp8b homozygous null mutant mouse models.6,7

Bone morphogenetic proteins are members of the transforming growth factor β (TGFβ) superfamily of growth factors, which also includes the TGFβ family and activins. These signaling molecules have diverse and often critical functions during embryonic development and in adult tissue homeostasis. They signal via formation of heterodimers or homodimers of the highly conserved carboxyl-terminal domain, which is characterized by six or seven cysteine residues that are critical to the structure, stability, and function of these proteins. TGFβ superfamily members signal by binding to serine/threonine kinase cell membrane receptors that leads to phosphorylation of proteins of the Smad family (named after the C. elegans gene sma and Drosophila gene Mad, reviewed in Shi and Massague8).

Smad complexes can bind DNA and mediate transcriptional activation or repression. Smad1 and Smad5 are known to act downstream of the Bmp signaling pathway and may have important functions in PGC development because loss of these proteins results in decreased size of the PGC population.911

Migration of PGCs through the hindgut and along the mesentery is mediated by the binding of c-Kit, a tyrosine kinase receptor expressed in PGCs, to its ligand, stem cell factor (Scf). Scf is expressed by somatic cells and its gradient along the migratory path is thought to direct the migration of PGCs. Mutations in the dominant white spotting (W) and Steel loci, which encode c-Kit and Scf, respectively, result in defects in gametogenesis. Although PGCs form, they do not proliferate, migrate to ectopic sites, and aggregate prematurely, so that the genital ridges are poorly populated by germ cells and the animals are sterile.12,13

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


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.2024

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.


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

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


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. 3-347). 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 3-4 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.

(From McGee EA, Hsueh AJW: Initial and cyclic recruitment of ovarian follicles. Endocrine Reviews 21:200–214, 2000; with permission. Copyright 2000, The Endocrine Society.47)


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.

(From McGee EA, Hsueh AJW: Initial and cyclic recruitment of ovarian follicles. Endocrine Reviews 21:200–214, 2000; with permission. Copyright 2000, The Endocrine Society.47)

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

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.