Human Reproduction Update Advance Access originally published online on January 22, 2007
Human Reproduction Update 2007 13(3):289-312; doi:10.1093/humupd/dml062
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Molecular mechanisms of ovulation: co-ordination through the cumulus complex
Research Centre for Reproductive Health, School of Paediatrics and Reproductive Health, The University of Adelaide, Adelaide, South Australia 5005, Australia
1 Correspondence address. School of Paediatrics and Reproductive Health, The University of Adelaide, Adelaide, South Australia 5005, Australia Tel: +61 8 8303 4096; Fax: +61 8 8303 4099; E-mail: darryl.russell{at}adelaide.edu.au
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Successful ovulation requires that developmentally competent oocytes are released with appropriate timing from the ovarian follicle. Somatic cells of the follicle sense the ovulatory stimulus and guide resumption of meiosis and release of the oocyte, as well as structural remodelling and luteinization of the follicle. Complex intercellular communication co-ordinates critical stages of oocyte maturation and links this process with release from the follicle. To achieve these outcomes, ovulation is controlled through multiple inputs, including endocrine hormones, immune and metabolic signals, as well as intrafollicular paracrine factors from the theca, mural and cumulus granulosa cells and the oocyte itself. This review focuses on the recent advances in understanding of molecular mechanisms that commence after the gonadotrophin surge and culminate with release of the oocyte. These mechanisms include intracellular signalling, gene regulation and remodelling of tissue structure in each of the distinct ovarian compartments. Most critical ovulatory mediators exert effects through the cumulus cell complex that surrounds and connects with the oocyte. The convergence of ovulatory signals through the cumulus complex co-ordinates the key mechanistic processes that mediate and control oocyte maturation and ovulation.
Key words: ovulation / fertility / ovarian follicle / oocyte / ovary
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Successful ovulation is a complex process whereby ovarian follicles reactivate oocyte meiosis, create a rupture pore in the apical follicle wall and initiate tissue restructuring and differentiation to form the corpus luteum. These processes are fundamental to successful establishment of pregnancy, but importantly also impact on the developmental potential of resultant embryos. A cascade of events drive ovulation, initiated upon receipt by the follicle of a single trigger, the surge of gonadotrophins: luteinizing hormone (LH) and follicle-stimulating hormone (FSH) from the pituitary (Figure 1). In response, detailed changes in gene expression and follicular structure occur with overlapping control and interdependent consequences in the theca, granulosa, cumulus and oocyte compartments of the ovarian follicle. Systemic and local inputs co-ordinate with signals from the oocyte; thus ovulation is under multipartite control facilitating synchronization of oocyte maturation with release and permitting the selection of oocytes with full developmental competence for succession to the reproductive pool.
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This review focuses on the dynamic tissue remodelling, cell–cell communication, intracellular signalling and transcriptional changes that occur in each compartment of the ovulating follicle. Most of the recent advances in understanding of the molecular mechanisms of ovulation have come from rodent models, with the most complete knowledge of genes and molecular pathways in ovulation derived from studies with mice and these are a focus of this review. However, a considerable contribution from studies in domestic animal and primate species is also important. Which model species is most appropriate for understanding critical genes and processes in human ovulation is currently unclear, making translational research in humans very important, and available information from such research is highlighted here.
Mural granulosa cells: targets and transducers of ovulatory signals
Pre-ovulatory follicles contain two distinct sublineages of granulosa cells that arise during folliculogenesis as the cell populations segregate upon formation of the fluid-filled follicular antrum. The mural granulosa cells line the follicle wall and reside very close to the basement membrane; and the cumulus cells are those directly adjacent to the oocyte (Figure 1). These two cell populations exhibit highly divergent responses during ovulation. Indeed, the direct response to the ovulatory LH surge is predominant in mural granulosa cells due to greater receptor levels than in cumulus cells. Mural granulosa layers express LH-receptors (LH-R), at levels typically an order of magnitude higher than those in cumulus cells. Binding of human chorionic gonadotrophin (hCG) within intact follicles is consistently reported to be at least 9-fold higher in mural granulosa cells than in cumulus of rat (Amsterdam et al., 1975
; Bortolussi et al., 1979; Lawrence et al., 1980), pig (Channing et al., 1981), hamster (Oxberry and Greenwald, 1982) and mouse (Wang and Greenwald, 1993b). The earliest studies utilized in vivo and in vitro binding of radiolabelled hCG to detect sites of action within the ovary and found that cumulus cells bound little or no hCG in contrast to mural granulosa cells within the same proestrus follicles (Amsterdam et al., 1975; Bortolussi et al., 1979; Channing et al., 1981; Oxberry and Greenwald, 1982; Wang and Greenwald, 1993a). Isolated cumulus–oocyte complex (COC) were similarly reported to have a 10-fold less hCG-binding capacity than isolated mural granulosa cells (Lawrence et al., 1980; Channing et al., 1981). Subsequent studies using in-situ hybridization showed LH-R mRNA expression is stratified within the rat pre-ovulatory follicle, with mural granulosa cells expressing the highest, levels and cumulus cells and oocytes having low to undetectable levels (Peng et al., 1991). An immunohistochemical study in rats supported mRNA analyses, demonstrating far greater LH/hCG receptor protein in theca, stroma and luteal tissue than mural or cumulus, but low levels of cell surface receptor were detected on cumulus cells after ovulation (Bukovsky et al., 1993). Confirming these in vivo observations of follicular expression, LH-R mRNA was low or undetectable in freshly isolated murine, bovine and equine cumulus cells but highly expressed in mural cells (Eppig et al., 1997; Goudet et al., 1999; Robert et al., 2003).
Postovulatory human cumulus cells undergo structural changes consistent with luteinization (Motta et al., 1995; Familiari et al., 2006), and this process includes the induction of LH-R. In a recent study, the relative levels of LH-R mRNA 36 h after hCG treatment in human granulosa versus cumulus cells were reported to be similar (Foong et al., 2006). However, LH-R mRNA levels prior to LH treatment or the responsiveness of human COC to FSH versus LH was not reported. This is essential information for understanding whether human, similar to animal models, have higher LH-R in the mural granulosa of pre-ovulatory follicles.
Numerous studies indicate that pre-ovulatory cumulus cells respond poorly, if at all, to direct LH exposure. Highly purified LH alone could not cause cumulus expansion in isolated mouse COCs (Eppig, 1979a, 1979b) in contrast to highly purified FSH. Furthermore, although cumulus expansion in vivo is induced by pure hCG treatment, this effect required the presence of other follicular compartments indicating the response in cumulus is indirect (Eppig, 1980). In rats, Hillensjo et al. (1981) reported that COC mucification is an FSH-specific effect and that FSH was 10-fold more potent than LH in stimulating progesterone (P) secretion by cumulus cells (Hillensjo et al., 1981). Subsequent experiments in mouse showed that FSH and LH can sequentially stimulate cumulus matrix synthesis, with FSH required prior to LH responsiveness, and that FSH or FSH plus estradiol induced LH-binding activity on cumulus cells, peaking after 8 h when cumulus expansion is well advanced in the mouse (Chen et al., 1994). Indeed most studies demonstrating LH action on COCs or maturation of oocytes in vitro include prior treatment or co-culture in the presence of FSH (Shimada et al., 2003; Foong et al., 2006). The preponderance of current evidence indicates that the ovulatory LH signal is received and responded to in the mural granulosa and theca cell layers and these may transmit a secondary signal to cumulus cells inducing myriad changes including the up-regulation of LH-R expression.
LH-induced intracellular signals
The LH surge triggers the activity of multiple intracellular signalling pathways in granulosa cells that culminate in altered transcriptional complexes mediating expression of ovulatory genes. The LH-R, a classical G
s G-protein-coupled receptor (GPCR) activates adenylate cyclase, resulting in a large intracellular cAMP increase that activates the cAMP-dependent serine kinase protein kinase A (PKA), a reaction which has been intensively studied (Marsh, 1976; Richards, 1994). Downstream of PKA, the cAMP regulatory element-binding protein (CREB) is phosphorylated at serine 133 (Mukherjee et al., 1996; Gonzalez-Robayna et al., 1999; Salvador et al., 2002; Russell et al., 2003b) and then recruits the CBP/p300 transcriptional coactivator (Arias et al., 1994). Phosphodiesterases are required to maintain tonic cAMP levels in responsive cells and PDE4D, in particular, regulates cAMP levels in granulosa cells (Tsafriri et al., 1996).
The LH surge is thought to stimulate additional signalling pathways, since LH treatment increased inositol triphosphate production in rat granulosa cells (Davis et al., 1986) as well as increased phospholipase C activity and intracellular calcium in non-ovarian cells (Gudermann et al., 1992; Zhu et al., 1994). Furthermore, cAMP and PKC activators, such as phorbol ester, are both required to emulate LH responses in cultured granulosa cells (Morris and Richards, 1995; Sriraman et al., 2003). Paradoxically studies have shown that LH-R activation (by hCG) does not activate PKC nor is a PKC inhibitor able to block hCG-mediated effects on rat granulosa cells (Salvador et al., 2002). Thus, although LH signalling likely involves activation of multiple parallel signal transduction pathways, PKC may not be one.
It is agreed that LH action on granulosa cells stimulates the extracellular regulated kinase (Erk1/2 or MAPK) pathway (Das et al., 1996; Maizels et al., 2001; Seger et al., 2001; Salvador et al., 2002; Choi et al., 2005), with activation of Erk being very rapid (Sela-Abramovich et al., 2005) and dependent on PKA (Salvador et al., 2002; Russell et al., 2003b). It is not yet entirely clear whether Erks are activated via direct phosphorylation by PKA or through some intermediate step such as inactivation of Erk phosphatases (Cottom et al., 2003), nor is the downstream function of Erks in granulosa cells, particularly in relation to ovulation, well understood; reports to date have primarily focused on their role in progesterone synthesis as cells luteinize (Seger et al., 2001; Dewi et al., 2002; Tajima et al., 2003). Recently, the transcription factor Rhox5 was identified as a transcriptional target of LH via Erks in ovulating follicles (MacLean et al., 2005). Ovulatory hCG also activates AP1 transcription factors of the Jun, Fos and Fra family (Sharma and Richards, 2000), consistent with general observations that Erk1/2 typically phosphorylate Fos, Myc, Elk-1 and Stat3 (Roux and Blenis, 2004). The effectors of Erk-mediated transcriptional regulation of granulosa cell gene expression during ovulation remain to be determined.
Follicular cGMP levels are increased during the course of ovulation, and at least one cGMP effector, the cGMP-dependent kinase cGKII, is also up-regulated at the time of ovulation by cAMP and Erk1/2 via progesterone receptor (PR) and epidermal growth factor (Egf) mediated pathways (Sriraman et al., 2005). Guanylyl cyclase receptors which generate cGMP, have been localized within the ovary (LaPolt et al., 2003). Expression of guanylyl cyclase-A receptor, and its ligand atrial natriuretic peptide (ANP), is up-regulated in mouse granulosa cells in response to ovulatory hCG (Sriraman et al., 2005); whereas guanylyl cyclase isoforms GC-B, GC-C and sGC are expressed at low levels and not hormonally regulated. Elevated production and activity of cGMP during the course of ovulation may alternatively occur via guanylyl cyclase activation by nitric oxide (NO), since nitric oxide synthase (NOS) in the theca produces NO during ovulation (Zackrisson et al., 1996). Clear effects of cGMP on granulosa cell gene expression, steroidogenesis and oocyte maturation are reported, but how this cascade impacts ovulation in particular is not known (LaPolt et al., 2003).
Studies using LH- or hCG-stimulated granulosa-lutein cells obtained from IVF patients confirm that PKA and PKC pathways are active in human granulosa cells (Freimann et al., 2004, 2005). More work is needed, however, to understand, as in rodents, the complex array of intracellular signalling pathways activated in response to the simple LH surge.
Periovulatory regulation of gene transcription
Signalling pathways induced by LH rapidly modify the transcriptional machinery of mural granulosa cells, reprogramming cellular function towards ovulation concurrent with luteinization. Extensive reprogramming of gene expression after the LH surge is achieved through modulation of a suite of transcriptional regulators, which in turn mediate effector gene transcription. A cohort of periovulatory genes with key roles in ovulation exhibits a common pattern of transcriptional induction involving binding of Sp1/Sp3 transcription factors to GC-box promoter elements. Sp1 and Sp3 are constitutively present in granulosa cells but exhibit sequence-specific, LH-regulated binding to gene promoters (Russell et al., 2003b). The signal transduction mechanism that modifies Sp1 binding activity in granulosa cells is uncertain, but in most tissues involves post-translational phosphorylation or glycosylation (Li et al., 2004).
Activity of Sp1/Sp3 complexes induces expression of additional transcription factors that further contribute to a cascade of ovulatory gene expression (Figure 2). Induction of PR in mouse and rat mural granulosa cells occurs in response to the LH surge through a cAMP/PKA-dependent mechanism, whereas phorbol ester (PKC activation) has a synergistic effect (Clemens et al., 1998; Sriraman et al., 2003; Sriraman and Richards, 2004). The cAMP-mediated induction of PR requires Sp1/Sp3 binding to multiple GC-box elements in the PR promoter (Sriraman et al., 2003) and consequent recruitment of additional transcription factors. In the rhesus monkey, PR is similarly induced in the periovulatory period (Stouffer, 2002). PR is present in human pre-ovulatory granulosa cells (Suzuki et al., 1994; Revelli et al., 1996; Chang et al., 2005). Two isoforms of PR are produced from distinct promoters on the same gene; PR-A is predominant in rat granulosa cells (Natraj and Richards, 1993) and is the most critical for ovulation, since mice lacking only PR-A have severely reduced ovulation (Mulac-Jericevic et al., 2000). Human ovaries express PR-A and PR-B isoforms, with PR-A most consistently found to predominate (Stouffer, 2003), however, the regulation of each isoform in ovulating follicles is not yet determined.
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Early growth response-1 (Egr-1) is a transcription factor induced through cAMP/PKA and Erk signalling. Transcriptional induction of Egr-1 requires Sp1/Sp3 and phosphorylated-CREB binding to the proximal promoter. This is thought to lead to recruitment of a combinatorial transcription initiator complex involving CBP and other cofactors (Russell et al., 2003b
). Another transcription regulator Rhox5 is also regulated by the combination of Ets/GABP, Sp1 and Sp3 (MacLean et al., 2005).
These LH-induced transcription factors in turn co-ordinately activate transcription of a further suite of ovulatory effector genes. For instance, the protease cathepsin L is induced through Sp1/Sp3, PR, Egr-1 and CREB-mediated transactivation (Jaffe et al., 1989; Sriraman and Richards, 2004), and another ovulatory protease ADAMTS-1 is induced through Sp1/Sp3 and PR in combination with C/EBPß, and NF-1-like factor (Doyle et al., 2004). The Egf-like factor epiregulin is also regulated by Sp1/Sp3 binding its promoter in rat granulosa cells (Sekiguchi et al., 2002).
Thus, the cascade of diverse periovulatory gene expression in mural granulosa cells involves the induction and recruitment of a cohort of transcription factors that subsequently induce effector gene products, including ADAMTS-1 and cathepsin L proteases, Egf-like ligands (Egf-L) and others. These ovulatory genes exhibit a signature pattern of mRNA and protein expression: a rapid, transient increase after the LH surge followed by down-regulation. Recurring modifications of the transcriptional machinery interacting with these gene promoters mediate this characteristic expression. The common denominator is Sp1/Sp3 transcription factor binding, suggesting that these are universal mediators of periovulatory gene expression. The Sp factors recruit chromatin-remodelling complexes that can switch in response to changing cell context to modify transcriptional complex assembly, or alter promoter occupancy at neighbouring enhancer sites (Li et al., 2004) (Figure 2). This process has been well described for LH-R transactivation by Sp and nuclear receptor transcription factors in granulosa cells (Zhang and Dufau, 2003). Histone deacetylases recruited to Sp1/Sp3-bound promoters silence transcription, but can be rapidly exchanged for histone-acetylating complexes that open local chromatin structure to RNA polymerase access where Sp1/Sp3 are present. More detail is needed to better understand how LH-induced intracellular signals impact specific transcriptional complexes that distinctly regulate ovulation versus luteinization events and how this model relates to human periovulatory gene regulation.
Essential effectors of ovulation in mural granulosa cells
A number of mural granulosa cell genes play essential roles in ovulation. Mouse models, in particular, have yielded a great deal of information about the role of key genes in specific aspects of the ovulatory process. Of those identified to date, a few are expressed prior to the LH surge, such as the transcription co-regulator, RIP140, and the phosphodiesterase, PDE4D; however, the majority are transiently expressed during the discrete ovulatory process. Many are transcriptional regulators, including PR, Egr-1 and CAAT enhancer binding protein (C/EBPß). These co-ordinately mediate expression of ovulatory effector genes.
Transcriptional regulators: PR, RIP140, Egr-1 and C/EBPß
Progesterone and PR are very specific key regulators of follicular rupture. Antagonists of PR, including RU486 (Tsafriri et al., 1987; Loutradis et al., 1991), or inhibitors of P synthesis each reduces or completely blocks ovulation in several species including rats (Espey, 1998), sheep (Murdoch et al., 1986) and humans (Baird et al., 2003). Pre-ovulatory granulosa cells begin expressing progesterone in the days prior to the LH surge (Gore-Langton and Armstrong, 1994; Richards, 1994), whereas PR is acutely induced after the LH surge and is immediately activated by the high local concentration of ligand, before its expression is down-regulated. Mice with a targeted deletion of the PR gene (PRKO) exhibit normal follicle growth and luteinization, but a complete and specific block in ovulation (Lydon et al., 1996; Robker et al., 2000). Key PR-dependent genes expressed in the periovulatory period, but absent in PRKO mice and most likely contributing to their anovulation, include proteases ADAMTS-1 and cathepsin L (Robker et al., 2000).
The nuclear receptor co-regulator RIP140 is most highly expressed in pre-ovulatory granulosa cells and is rapidly down-regulated after the LH surge in the periovulatory period (White et al., 2000). RIP140 plays a broad role in the regulation of genes that are involved in follicle rupture, suggesting that this transcription cofactor is part of a cell-specific transcriptional complex formed in the granulosa compartment of ovulating follicles. Mice null for RIP140 exhibit apparently normal follicular growth and luteinization, but a block in ovulation due to the failed induction of numerous critical mural and cumulus cell genes, such as ADAMTS-1, epiregulin, amphiregulin, betacellulin, hyaluronan synthase-2(HAS-2), cyclooxygenase-2(COX-2), pentraxin-3(PTX-3) and tumour necrosis factor stimulator gene-6(TSG-6) (Tullet et al., 2005). Thus, RIP140 may regulate differentiation of follicles to pre-ovulatory maturity or mediate a single master event in the periovulatory response to LH. The close similarity of the PRKO phenotype to that of RIP140 null mice suggests that these two transcriptional regulators are in the same essential pathway.
The LH surge also initiates rapid and transient induction of transcription factors Egr-1 (Russell et al., 2003b) and C/EBPß (Sirois and Richards, 1993) in mural granulosa cells. Egr-1 null mice are infertile, primarily due to loss of LH secretion due to its role in LHß gene synthesis in the pituitary (Lee et al., 1996; Topilko et al., 1998). Thus, fertility may be controlled at multiple levels by Egr-1, including both pituitary and ovarian roles in the ovulatory process. It has been reported that ovarian LH-R expression is activated by Egr-1 and is disrupted in Egr-1 null mouse ovaries (Topilko et al., 1998; Yoshino et al., 2002). However, closer analysis of these mice indicates that after exogenous gonadotrophin replacement, ovulation remains impaired but that ovarian LH-R is normally expressed (Russell 2004). Null mutants for C/EBPß show oocytes entrapped in CL; however, evidence suggests these are among the few models with affected luteinization in addition to ovulation (Sterneck et al., 1997). Consistent with this it was shown that, C/EBPß may regulate steroidogenesis through the StAR gene (Christenson et al., 1999).
cAMP metabolism
The phosphodiesterase PDE4D, an important regulator of LH-mediated cAMP accumulation in granulosa cells (Tsafriri et al., 1996), is required for normal ovulation. PDE4D null mice exhibit dramatically reduced ovulation rates and litter size due to altered responsiveness of granulosa cells to the LH surge and reduced expression of ovulatory genes, including PR, cathepsin L and COX-2 (Jin et al., 1999; Park et al., 2003). Many follicles fail to reach pre-ovulatory stage due to premature luteinization. However, by in-situ hybridization, it was shown that the morphologically normal pre-ovulatory follicles, although reduced in number, still express ovulatory genes such as the Egf-L (Park et al., 2004). Thus, rather than rendering the granulosa cells unable to respond to ovulatory LH, the lack of PDE4D primarily permits accumulation of cAMP such that follicles exhibit premature luteinization and entrapment of oocytes in immature luteinized follicles.
Egf-L: amphiregulin, betacellulin and epiregulin
The Egf-L, epiregulin, amphiregulin and betacellulin, are very rapidly induced specifically in mural granulosa cells within 1–3 h after the LH surge and act as major secondary signals that transmit the ovulatory signal to the cumulus complex (Park et al., 2004) (Figure 1). Epiregulin and amphiregulin are also LH inducible in human granulosa cells (Freimann et al., 2004). Owing presumably to functional redundancy between epiregulin, amphiregulin and betacellulin, null mutant mice for each of these genes are fertile (Luetteke et al., 1999; Lee et al., 2004). Each of the Egf-L factors are synthesized as integral membrane proteins, and cleavage of the precursor forms is required for their interaction with cumulus-expressed receptors since protease inhibitors can block transmission of the LH signal to cumulus cells, whereas cotreatment with epiregulin reverses this block (Ashkenazi et al., 2005). Identification of the processing enzyme will reveal a key regulatory event in intrafollicular co-ordination of the ovulatory processes.
Extracellular matrix protease ADAMTS-1
The extracellular protease ADAMTS-1 was identified as LH- and PR-induced gene in mural granulosa cells in the rat (Espey et al., 2000) and mouse (Robker et al., 2000). This gene has been identified in bovine (Madan et al., 2003), equine (Boerboom et al., 2003), primate (Young et al., 2004) and human ovaries (Freimann et al., 2005) and may be associated with the aetiology or symptoms of anovulatory human polycystic ovarian syndrome (Jansen et al., 2004), but has yet to be studied in human normal ovulation. The latent pro-form of ADAMTS-1 is synthesized by the mural granulosa cells, but the secreted, mature form is selectively localized to the extracelullar matrix (ECM) and cell surfaces of the expanded cumulus complex (Russell et al., 2003a), suggesting this is where it mediates its principal effects. Two lines of mice with null mutation of the ADAMTS-1 gene have 70–90% reduced ovulation rates and 75% reduction in litter size (Shindo et al., 2000; Mittaz et al., 2004). From these studies, it is clear that ADAMTS-1 is an important mediator of LH and progesterone effects in ovulation across species. Substrates for ADAMTS-1 include versican, which is also produced by rodent mural granulosa cells and stimulated by the LH surge (Russell et al., 2003a) and has been identified in human ovaries (Eriksen et al., 1999; Relucenti et al., 2005). As with ADAMTS-1, versican is produced in mural granulosa cells, but the majority of protein in the periovulatory period is incorporated into the cumulus cell matrix.
Clearly, many mural granulosa cell gene products are required for successful and optimal ovulation. Many of these genes encode proteins that sense and respond to systemic inputs, including the LH surge. A cohort of essential ovulatory mediators are transcription factors that are up- and down-regulated by the LH surge and function to translate the ovulatory trigger into global reprogramming of gene transcription and ultimately cell function. Interestingly, a new group of effector gene products is emerging, including the Egf-L, versican and ADAMTS-1, which are produced by mural granulosa cells, but specifically translocate to and act within the COC. This combined endocrine and paracrine intrafollicular communication enables the co-ordination of maternal receptivity with oocyte maturation and ovulation.
Cumulus cells: conduit of ovulatory signals
The pre-ovulatory oocyte is surrounded by several cell layers, known as the cumulus granulosa cells, which are tightly connected to each other and the oocyte through intercellular membrane processes and gap junctions that facilitate exchange of glucose metabolites, small signalling molecules and ions. Oocytes depend on the cumulus cells for metabolism of glucose and supply of pyruvate for energy production (Biggers et al., 1967; Gardner et al., 1996; Preis et al., 2005). The cumulus cells exhibit hormone responsiveness and gene expression profiles distinct from those of the mural granulosa layers. In preovulatory follicles, cumulus and mural cells present abundant cell surface FSH receptors, but the cumulus mass and granulosa cells very close to the antrum show the greatest proliferative response to FSH during follicle growth (Robker and Richards, 1998). The cumulus are the cells most directly exposed to mitogenic factors secreted by the oocyte (Erickson and Shimasaki, 2000). Oocyte-secreted factors (OSF) belonging to the Tgfß superfamily establish a morphogen gradient that promotes the cumulus cell characteristics including enhanced cell proliferation, and repression of LH-R and PR gene transcription, and progesterone biosynthesis (Eppig et al., 1998; Elvin et al., 1999a; Gilchrist et al., 2004). The rate of proliferation of these cells has been suggested to correlate with implantation potential in human assisted reproduction (Gregory, 1998). Following the ovulatory LH surge, cumulus cells respond with a unique pattern of gene induction leading to production and stabilization of a local ECM that envelopes the COC. This process, known as cumulus expansion or mucification, is dependent on a specific cascade of intracellular signals and ECM gene expression within the COC and plays a key role in ovulation (Chen et al., 1993; Hess et al., 1999; Fulop et al., 2003; Salustri et al., 2004) and subsequent fertility success (Tanghe et al., 2003; Somfai et al., 2004; Yang et al., 2005).
Intracellular signalling in periovulatory cumulus cells
Cumulus cells respond rapidly and change dramatically after the gonadotrophin surge. The program of gene induction in the cumulus cells responding to the gonadotrophin surge is potentially mediated, at least in part, by high FSH that accompanies the LH surge. In women, as in other mammalian species, FSH levels surge with similar timing kinetics to LH at the midcycle (Hoff et al., 1983). Local paracrine effectors including Egf-L from granulosa cells, interleukin-1 from the thecal compartment and oocyte-derived signals also contribute to cumulus cell responses (Figure 1).
Cumulus response to endocrine and paracrine stimuli
Follicle stimulating hormone
In isolated COCs in vitro, FSH induces periovulatory changes in cumulus cells and oocytes (Su et al., 2002) and in vivo very high concentrations of pure FSH have even been found to initiate oocyte maturation and ovulation in rodents (Tsafriri et al., 1976b; Galway et al., 1990; Wang and Greenwald, 1993a) and primates (Zelinski-Wooten et al., 1998). However, the levels of FSH used to activate COC expansion in vitro are supraphysiological, and in animal models or human IVF cycles, levels of FSH, which reflect physiological circulating levels during the ovulatory surge, do not enhance molecular events associated with ovulation or oocyte development (Tsafriri et al., 1976b; Vermeiden et al., 1997). Although periovulatory gene expression is induced by FSH in the cumulus, it has been widely shown that ovulation and oocyte maturation are efficiently induced in vivo by pure exogenous LH preparations that mimic the levels of an endogenous LH surge. These observations indicate that FSH may play a minor role, but is not essential, whereas intrafollicular responses to the LH surge mediate ovulation.
Since FSH is effective in stimulating ovulatory cumulus gene expression, it is widely used to study molecular aspects of this process. The FSH receptor is a GPCR closely related to LH-R that activates adenylate cyclase mediating cAMP synthesis and resulting in PKA-activated intracellular signalling (Figure 3). In addition, FSH independently, activates the PI3K/Akt pathway (Gonzalez-Robayna et al., 2000; Zeleznik et al., 2003) and recruits the 14-3-3 adaptor protein (Cohen et al., 2004) to mediate cell survival and proliferation in granulosa cells. Through the Akt pathway, FSH also increases transcription of the hypoxia inducible factor-1 (Alam et al., 2004) leading to the induction of expression from genes such as vascular endothelial growth factor (VEGF), a critical mediator of angiogenesis during ovulation/luteinization (see below). Most importantly, FSH activates ERK signalling in granulosa (Das et al., 1996) and in cumulus cells (Su et al., 2002) and Erk activity is requisite in cumulus cells but not in oocytes for COC expansion and oocyte meiotic maturation (Su et al., 2002). Inhibitors of Erk, p38 kinase or Akt block COC expansion induced by FSH or cAMP treatment (Ochsner et al., 2003a), indicating that each signalling kinase pathway converges to mediate important periovulatory cumulus gene expression.
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Prostaglandins
Cumulus cells in ovulating COC produce prostaglandins via induction of COX-2 and other prostaglandin synthetic enzymes. Induction of COX-2 in COC can be stimulated by Egf in cows (Nuttinck et al., 2002
) and mice (Park et al., 2004) or by FSH through cAMP/PKA activation (Joyce et al., 2001). Pharmacological activators of cAMP synthesis induce COX-2, but this is sensitive to inhibition of p38-MAPK or Erk1/2 (Ochsner et al., 2003a), indicating that Erk-mediated transcriptional induction, distinct from or in conjunction with the classical cAMP pathway, is required for COX-2 expression. The COX-2 gene promoter has been studied in detail and shown in mouse granulosa cells to be dependent on oocyte-derived Gdf-9 (Elvin et al., 2000) and induced through an E-box element that interacts with the upstream stimulatory factor (USF) transcription factor in a manner regulated by PKA (Morris and Richards, 1996; Sayasith et al., 2004; Sayasith et al., 2005). The predominant prostaglandin produced is prostaglandin (PGE2) which interacts with the PGE2-receptor (EP-2) to induce changes in cumulus gene expression (Segi et al., 2003) critical for COC expansion and ovulation (Hizaki et al., 1999). Prostaglandins made by the cumulus cells potentially mediate important signalling effects through other receptors also, since disruption to fertility in EP-2 null mice is far less severe than in COX-2 nulls that produce no prostaglandins (Table I). Very recently, the PGE2 receptor, EP-4, was also shown to be induced in periovulatory cumulus cells and to participate with EP-2 in oocyte maturation, cumulus cell survival and matrix production (Takahashi et al., 2006). Both PGE2 receptors are GPCR that stimulate cAMP production; thus in cumulus cells, prostaglandin synthesis establishes an autocrine signalling loop synergizing with FSH and reinforcing cAMP/PKA and Erk-dependent gene induction in cumulus cells (Figure 3). This loop of cAMP-mediated PKA activation in cumulus cells may be controlled by the regulator of G-protein signalling (RGS-2), which has a temporal and spatial pattern of expression very similar to COX-2 and EP-2 in periovulatory COCs (Ujioka et al., 2000). Adenylate cyclase activity can be either augmented or inhibited by RGS-2 (Kehrl and Sinnarajah, 2002), and its role in COC has yet to be more fully investigated. Human granulosa-lutein cells express EP-2 (Harris et al., 2001)and secrete PGE2 (Chandras et al., 2004), indicating that a similar signalling loop likely exists in human ovulation.
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Epidermal growth factor-like ligands
Egf-L (amphiregulin, betacellulin and epiregulin) from LH-stimulated mural granulosa cells also activate periovulatory COC gene expression (Espey and Richards, 2002
; Park et al., 2004; Rimon et al., 2004). These ligands activate the Egf-receptor tyrosine kinase and subsequently the Erk activity (Kansra et al., 2004) (Figure 3). Cumulus cell expression of ECM genes, COC expansion and ovulation are blocked by pharmacological inhibition of the Egf-receptor tyrosine kinase or downstream Erk function (Lorenzo et al., 2001; Su et al., 2002; Park et al., 2004; Tsafriri et al., 2005). Ovulation induction by hCG is also blocked by inhibition of Egf-receptor activity; oocytes are entrapped in follicles surrounded by compact cumulus cells and cumulus matrix gene induction in cultured follicles is inhibited (Ashkenazi et al., 2005). Thus, both FSH and Egf-L signals promote cumulus periovulatory gene expression through activation of Erk signalling.
Androgen
Cumulus cells of several species, including rat (Pelletier et al., 2000; Szoltys et al., 2003) and human (Hickey et al., 2005), express the androgen receptor and thus are thought to respond directly to androgens produced by theca cells. There is limited evidence for the consequences of androgen action in ovulation; however, in female mice lacking androgen receptor a minor, albeit significant reduction in ovulation rate was reported (Hu et al., 2004). Furthermore, ovulatory genes, including PR, HAS2 and TSG-6, were not induced, and the cumulus complexes ovulated were reported to have a disorganized structure, suggesting a role for androgen receptor in COC expansion and the ovulatory process (Hu et al., 2004).
Interleukin-1ß
Other cells within the theca interna, possibly leukocytes surrounding periovulatory follicles, may also contribute to cumulus expansion in vivo (Figure 1). These cells produce IL-1ß (Jasper and Norman, 1995; Wang et al., 1997; Minge et al., 2005) in response to LH (Kol et al., 1999). The IL-1ß receptor activates a receptor-associated kinase, again resulting in Erk activation (Eder, 1997). Thus, in vivo, paracrine signals from the theca also transmit signals to the cumulus and co-ordinate the cumulus response to the LH surge. Hyaluronan (HA) synthesis and cumulus expansion are induced by IL-1ß (Kokia et al., 1993; Singh and Armstrong, 1997), and cumulus expansion and ovulation can be blocked by IL-1 antagonism in rodents (Simon et al., 1994). In humans, IL-1ß and IL-1 have been found to be induced by gonadotrophins and serum levels correlate with successful fertilization (Mendoza et al., 1999) or implantation in assisted reproductive cycles (Karagouni et al., 1998). However, subsequent studies evidenced either no correlation (Barrionuevo et al., 2000) or reverse correlation (Mendoza et al., 2002).
Extracellular matrix
The COC matrix itself may have important effects in mediating or modulating cumulus intracellular signalling. The HA receptor, CD44 is expressed on cumulus cells, induced after the LH surge in many species including humans (Ohta et al., 1999; Yokoo et al., 2002; Schoenfelder and Einspanier, 2003) and can transduce multiple intracellular signals (Bourguignon, 2001) via cytoskeletal changes, which transactivate the Egf-R family member ErbB2 and activate Erk. Proteoglycan components of the COC matrix, such as CD44 and versican, likely bind growth factors and modulate signalling. However, CD44 null mice have normal fertility (Schmits et al., 1997; Protin et al., 1999), suggesting the involvement of redundant HA receptors such as RHAMM, which is present in bovine COC (Turley et al., 2002). Actions of the proteoglycan inter-alpha trypsin inhibitor (II) found in the COC matrix include modulating kinases Erk and Akt to mediate gene expression changes (Kobayashi et al., 2001). Microarray analysis identified 29 genes up-regulated and 5 genes down-regulated 12 h after hCG in COC of mice that lack II relative to wild-type controls (Suzuki et al., 2004), suggesting that it regulates gene expression changes in addition to being a key matrix component.
To date, transcription regulators modulating periovulatory cumulus cell gene expression have not been studied in detail due to lack of an appropriate in vitro model for such studies. Repeatedly it has been reported that signals that activate periovulatory cumulus gene expression act through the Erk kinase family. The transcription factors most commonly phosphorylated by Erk kinases include AP-1 factors, signal transducer and activator of transcription (STAT) and serum-responsive transcription factors. However, in addition, cumulus-cell-specific gene expression requires oocyte-derived signals.
Cumulus cell response to signals from the oocyte
Mature oocytes, although transcriptionally inactive, secrete signals that play a key permissive role in periovulatory cumulus gene expression in mice (Eppig et al., 2002). This oocyte-derived activity, originally referred to as cumulus-expansion-enabling factor (CEEF), is secreted by mouse, pig and cow oocytes (Eppig et al., 1993; Singh et al., 1993; Vanderhyden et al., 1993; Gilchrist et al., 2004). This ability is linked to the oocytes' developmental competence (Eppig et al., 1993). A range of experiments seeking to identify the CEEF in mice suggests that Gdf-9 as well as other Tgfß superfamily members can support cumulus ECM gene expression (Vanderhyden et al., 2003; Dragovic et al., 2005). However, the identity of the true oocyte-secreted factor(s) facilitating COC expansion remains controversial, since antibody neutralization of Gdf-9 secreted by oocytes only partially blocks COC expansion (Dragovic et al., 2005), whereas suppression of Gdf9 mRNA in mouse oocytes was more highly effective (Gui and Joyce, 2005).
The specific roles for Gdf-9, BMP-15 and perhaps other Tgfß family members such as BMP-6, in ovulation have also been difficult to fully interpret because of species variations and their critical role early in folliculogenesis. In pigs and cows, the oocyte does not need to be present at the time of cumulus expansion, but in cows, BMP-15 is a necessary cumulus cell survival factor during oocyte maturation and COC expansion (Hussein et al., 2005). The impact of growth factor signals from the human oocyte has not been directly tested due to obvious constraints on available COCs for analysis. Thus, the model system that best represents the human is unknown.
In all species, the initiator of ovulatory cumulus gene expression is clearly not Gdf-9 and/or BMP-15, since these factors are constitutively synthesized by oocytes and affect follicular somatic cell development throughout folliculogenesis (Elvin et al., 1999b; Juengel et al., 2002), including in humans (Di Pasquale et al., 2004), without activating matrix production. It is thought that in all species, oocyte signals specify the cumulus cell lineage which orchestrates the cumulus-specific responses after the LH surge (Erickson and Shimasaki, 2000; McNatty et al., 2005). Therefore, by producing Gdf-9 and BMP-15, the oocyte controls the differentiation of its immediate environment and hence its ovulation.
Signalling by Gdf-9 in cumulus cells is achieved by binding to the BMP type-II receptor (BMPRII), followed by recruitment and activation of the type-I receptor, Alk5 (Mazerbourg et al., 2004) (Figure 3). Thus Gdf-9 uses similar intracellular signalling to Tgfß. Activated Alk5 in human granulosa-lutein cells phosphorylates transcription factors SMAD2 and SMAD3, which dimerize with SMAD4, translocate to the nucleus and bind to SMAD-response elements in gene promoters containing the core nucleotide sequence CAGA, such as the inhibin-B promoter (Kaivo-Oja et al., 2005). Not surprisingly, factors that share the capacity to activate SMAD2/3 in cumulus cells are those previously shown to share the CEEF activity.
The mRNA and protein for SMAD2 and 4 are induced in whole mouse ovary by FSH (Gueripel et al., 2004), indicating that the pre-ovulatory follicle is primed for a robust response to Gdf-9 and other Tgfß-related ligands in the periovulatory phase. Knockout of the SMAD3 gene results in anovulatory infertility primarily due to defects in follicle growth (Tomic et al., 2004), a similar but not identical phenotype to Gdf-9–/–. This may indicate that the Gdf-9 functions partially but not exclusively through SMAD3. SMAD2 and SMAD4 knockout mice die in embryonic development. Whether SMADs bind directly to promoters of each cumulus-expressed periovulatory gene or regulate these genes through indirect means is not known. It has been suggested that Gdf-9 also activates Erk signalling in cumulus (Su et al., 2003), but since other Erk activators such as Egf cannot substitute for the signal from oocytes, SMAD 2/3 activation is more likely to be the requisite mechanism of Gdf-9-mediated cumulus gene expression.
Essential cumulus gene expression and regulation
The set of genes expressed in periovulatory cumulus cells are critical for normal rates of ovulation and fertility (Table I). Among the earliest induced genes is COX-2, the rate-limiting enzyme in prostaglandin production (Sirois and Richards, 1992; Wong and Richards, 1992). COX-2 production is rapidly induced within 2 h of an hCG stimulus in rodent mural and cumulus cells (Sirois et al., 1992), but expression is higher and persists longer in the COC (Elvin et al., 1999a; Joyce et al., 2001; Segi et al., 2003). In larger species with longer periovulatory periods, this induction happens later and COX-2 has been proposed to be part of a molecular clock that sets the species-specific timing of ovulation (Sirois et al., 2004). This suggestion is supported by studies in the macaque monkey, but in this model, the timing of prostaglandin secretion is dependent on induction of COX-2, microsomal prostaglandin-E synthase and most particularly on expression and post-translational activation of phospholipase A2 in granulosa cells (Duffy et al., 2005). This mechanism places more stringent control on the process required for prostaglandin synthesis and the timing of ovulation. It is not known whether a similar complex regulatory mechanism operates in human ovulation.
Null mutation of the mouse COX-2 gene results in defective cumulus expansion, reduced ovulation rate and infertility (Dinchuk et al., 1995; Lim et al., 1997; Davis et al., 1999; Ochsner et al., 2003b). This is explained by the reduced expression of PGE2/EP-2-dependent genes such as TSG-6 in COX-2–/– as well as EP-2–/– mice (Ochsner et al., 2003a). Likewise, treatment of mice and rats with COX-2 inhibitors indomethacin or NS398 also blocks ovulation and TSG-6 expression (Mikuni et al., 1998; Yoshioka et al., 2000; Espey and Richards, 2002). Indomethacin treatment also caused delayed ovulation by 2–12 days in women, (Athanasiou et al., 1996) and another COX-2 inhibitor, rofecoxib, had a similar effect (Pall et al., 2001). These studies collectively demonstrate that prostaglandin production via COX-2 enzyme is a key event in the cumulus gene expression cascade and vital to ovulation across mammalian species.
Genes encoding COC matrix components are also induced in cumulus cells by FSH or Egf in conjunction with oocyte signals including HAS-2, the enzyme that synthesizes the HA backbone of the cumulus matrix, (Eppig, 1979a, 1981; Tirone et al., 1997; Elvin et al., 1999a; Dragovic et al., 2005) and HA binding proteins TSG-6 (Fulop et al., 1997; Yoshioka et al., 2000) and pentraxin-3 (PTX-3) (Varani et al., 2002; Salustri et al., 2004). The inhibitor of BMP signalling Gremlin is expressed in pre-ovulatory mural and cumulus cells, but maintained only in cumulus cells after ovulation induction through Gdf-9 action (Pangas et al., 2004). The need for oocyte factors as well as FSH or Egf for induction of these cumulus specific genes has been characterized, but the full regulatory mechanism in cumulus cells has not been studied in detail. Interestingly, induction of TSG-6 in cumulus cells is dependent on autocrine/paracrine prostaglandin action, yet HAS-2 expression is not (Ochsner et al., 2003b). However, PGE2 treatment of isolated COC can induce HAS-2 mRNA (Eppig, 1981). These observations indicate that HAS-2 induction requires cAMP-PKA and/or Erk signalling in addition to Gdf-9/BMP-15 and that diverse regulatory mechanisms control different periovulatory genes in cumulus cells in vivo.
Cumulus matrix function
Appropriate formation of the expanded cumulus matrix is critical for ovulation. In fact, the success of follicular rupture as well as fertilization is exquisitely sensitive to perturbations in the composition and hence functional capacity of the cumulus matrix. This has been clearly established in mice by gene ablation of COC matrix proteins (Sato et al., 2001; Zhuo et al., 2001; Varani et al., 2002; Fulop et al., 2003; Salustri et al., 2004) (Table I). Other approaches that inhibit the expansion, organization or function of cumulus matrix proteins support the observations from knockout studies (Chen et al., 1993; Hess et al., 1999; Ochsner et al., 2003a).
Composition of expanded cumulus matrix
The backbone of the expanded cumulus matrix is HA, a large disaccharide chain common to many extracellular matrices including cartilage where it provides strong osmotic force and viscoelastic properties. Synthesis of HA requires glucose as a substrate for the hexosamine pathway, which generates the disaccharide subunits of HA. Glucose uptake and glycolytic activity in cumulus cells is markedly stimulated by the LH surge in rodents, cows and humans (Tsafriri et al., 1976a; Zuelke and Brackett, 1992; Roy and Terada, 1999; Sutton et al., 2003). Glucose flux through the hexosamine pathway in bovine COC increases during oocyte maturation (Sutton-McDowall et al., 2004). Strands of HA matrix are stabilized and organized into a specific functional structure by cross-linking proteins that bind HA through canonical link-protein-like modules. Protein cross-linkers play a fundamental role of stabilization of the HA matrix, as demonstrated by its dissolution by treatment with protease (Cherr et al., 1990). Organizers of the COC matrix, which bind HA directly, include TSG-6 (Carrette et al., 2001; Mukhopadhyay et al., 2001; Ochsner et al., 2003a), synthesized by cumulus cells and versican synthesized by the mural granulosa cells (Russell et al., 2003c) and the heavy chains (HCs) of II, a serum protein secreted by hepatocytes (Steinbauch and Loeb 1961). Circulating II is a trimeric protein complex comprising two HC subunits joined through chondroitin sulphate bridges to bikunin (Bost et al., 1998). It circulates in high concentrations and enters the ovulating ovary in blood plasma as a result of increased follicular vascular permeability during ovulation (Castillo and Templeton, 1993). The HC binds HA in the expanding COC ECM, whereas the bikunin subunit remains in solution and takes no further part in the COC matrix formation. Cartilage link protein may also be a product of FSH or IGF-1-stimulated mural granulosa cells (Sun et al., 2003) and is found in pre-ovulatory follicles of rodents and humans (Kobayashi et al., 1999). Exogenous link protein enhances volumetric COC expansion in vitro, but is not an essential matrix component (Sun et al., 2002). PTX-3 is an immune protective and anti-inflammatory protein that does not possess an HA-binding motif, but localizes intensely to the cumulus ECM in rodents and human. In addition, fibronectin, tenascin-C and laminin have been reported in the human COC matrix (Familiari et al., 1996).
Function of HA cross-linking proteins in cumulus matrix
Evidence suggests that the function of the cumulus matrix is dependent on a specific structure produced through the binding of HC and HC-TSG-6 complexes to HA (Carrette et al., 2001; Mukhopadhyay et al., 2001; Fulop et al., 2003; Ochsner et al., 2003a) (Figure 4). Formation of the appropriate covalent complex of II-HC to HA requires an intermediate interaction with TSG-6 (Chen et al., 1996; Carrette et al., 2001; Jessen and Odum, 2003). Loss of either TSG-6 or II synthesis results in almost identical phenotypes: HA is produced by cumulus cells but not stabilized, resulting neither in matrix aggregation nor morphological COC expansion in vitro or in vivo. As a result, fewer (<50%) oocytes are ovulated (Sato et al., 2001; Zhuo et al., 2001; Fulop et al., 2003). Of the few oocytes that are ovulated in TSG-6–/– mice, no fertilization or implantation has been observed in natural matings, but fertilization under IVF conditions has not been assessed (Fulop et al., 2003). A TSG-6 neutralising antibody can prevent cumulus matrix assembly in vitro (Ochsner et al., 2003a). Treatment of mice with inhibitors of the HA synthetic pathway (Chen et al., 1993) or short HA oligosaccharides that adsorb TSG-6 (Hess et al., 1999) also block COC expansion and ovulation. These data combined with TSG-6–/– studies indicate that TSG-6 as well as II interactions with HA are required for the formation of the COC matrix.
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The role of PTX-3 is different; mice with PTX-3 null mutation achieve COC matrix expansion in vivo, but the cumulus cells appear abnormally distributed. Oocytes are ovulated but fail to fertilize because of the rapid degradation and loss of the COC matrix during or after release from the follicle (Varani et al., 2002
; Salustri et al., 2004). Thus, PTX-3 appears to be required for matrix stability and retention. In vivo fertilization was <2% of the normal rate in one PTX-3–/– mouse line (Varani et al., 2002) and completely deficient in another (Salustri et al., 2004). Under IVF conditions, fertilization of PTX-3–/– oocytes occurred at the normal rate. This is expected since it is known that the need for the COC matrix can be circumvented in vitro by the use of high sperm concentrations; it also indicates that there is no profound oocyte deficiency in the absence of PTX-3 (Salustri et al., 2004). PTX-3 protein forms homo-decameric complexes that may interact with HA-bound TSG-6 molecules to form anchoring complexes that cross-link multiple independent HA molecules (Salustri et al., 2004). The above observations together demonstrate that distinct functional aspects of the COC matrix composition and structure are involved in release of the oocyte and its protection during passage through the oviduct.
Versican, a chondroitin-sulphate-substituted large aggregating proteoglycan of the lectican family, is closely related to aggrecan that cross-links HA providing strength and elasticity to cartilage. Versican is produced primarily by mural granulosa cells and rapidly incorporated into the cumulus matrix as it forms, suggesting that it binds HA through its link module and is another organizer of COC matrix structure (Russell et al., 2003c). Whether versican is obligatory for COC matrix expansion or ovulation has not been directly confirmed. Mutation of the versican gene in mice is uninformative since these mice die before embryonic day 7 because of defective heart development (Mjaatvedt et al., 1998). However, an important role for versican protein synthesis and proteolytic processing may be implied from other studies. The protease ADAMTS-1 selectively accumulates in the expanding COC matrix possibly through direct interaction with versican and specifically cleaves versican within the COC matrix (Russell et al., 2003a). Ovaries of PRKO mice have 10% of normal ADAMTS-1 protein abundance and 50% lower versican processing, but morphologically normal COC expansion occurs. PRKO mice are anovulatory due in part to reduced cleavage of versican resulting in altered COC matrix function (Russell et al., 2003a). ADAMTS-1 null mutation also results in reduced ovulation rate and subfertility in two independent null lines (Shindo et al., 2000; Mittaz et al., 2004). These observations suggest that cross-linking of COC HA by versican occurs prior to ovulation, but rapid cleavage of versican is requisite for normal matrix function and ovulation of COC.
Oocyte release and COC matrix expansion seem inseparable, co-dependent processes, yet how the cumulus matrix affects or participates in ovulation remains unknown. The composition and structure of the expanded cumulus matrix is of clear importance for fertility in rodents. A matrix of similar composition forms around human oocytes (Relucenti et al., 2005), and the degree of cumulus expansion and expression of mRNA for the HAS-2 enzyme have been correlated with human oocyte quality (McKenzie et al., 2004), but composition and function have been less well studied.
Additional key properties of the cumulus matrix
In all the disrupted cumulus matrix models thus far tested, in vivo fertilization rate is the most dramatically reduced fertility parameter (Table I). Disrupted cumulus matrix may alter oviductal transport of the COC. After ovulation, oocyte capture requires selective adhesion between the COC matrix and ciliated infundibular epithelium of the oviduct (Lam et al., 2000). Matrix enclosed COC is efficiently transported through the oviductal lumen (Lam et al., 2000; Talbot et al., 2003), whereas cumulus-denuded oocytes are not (Odor and Blandau, 1973; Mahi-Brown and Yanagimachi, 1983). Impaired oocyte capture or transport in women can cause infertility and/or ectopic pregnancy (Harper, 1994). However, animal models of cumulus matrix disruption and reduced ovulation frequently show entrapment of oocytes in luteinized follicles (Davis et al., 1999; Robker et al., 2000; Shindo et al., 2000; Varani et al., 2002; Mittaz et al., 2004), consistent with specific effects on oocyte release rather than capture or transport of the oocyte.
Cumulus matrix also plays a role in capacitation of sperm (Van Soom et al., 2002) and may present a selective barrier that screens for sperm with high fertilization potential (Hong et al., 2004). A proteoglycan component of the COC consistent in size and composition to versican accelerates human sperm acrosome reaction (Tesarik et al., 1988; Drahorad et al., 1991). PTX-3 binds sperm and may either anchor or attract sperm to the COC in the oviduct in vivo (Salustri et al., 2004). Furthermore, a form of glycodelin in the COC matrix appears to have the effect of promoting sperm binding to the oocyte (Yeung et al., 2006). Thus components of the COC matrix may be an active sperm attractant and capacitator.
The presence of cumulus has also been found to reduce oxidative stress on the oocyte (Fatehi et al., 2005); this may be an important role for cumulus and one explanation for the reported beneficial effect of cumulus inclusion in bovine IVF (Luciano et al., 2005). Pericellular versican is known to be a potent inhibitor of oxidative damage in cells (Wu et al., 2005), and hence versican in the COC matrix may protect the oocyte during the events of tissue hyperaemia and inflammation, which increase oxidative free radicals during ovulation.
The synthesis of COC matrix and cumulus gene expression has been linked to the determination of oocyte quality and developmental capacity. Degree of cumulus expansion has been used as a criterion for selection of oocytes for IVF for many years (Ball et al., 1983; Laufer et al., 1984; Foote, 1987). More recently, PTX-3 expression was correlated with oocyte quality in one study (Zhang et al., 2005); however, in another study, PTX-3 expression was not significantly correlated with blastocyst grade, whereas COX-2 and HAS-2 were (McKenzie et al., 2004). In each case, these parameters most likely serve as surrogate measures of the level of endocrine, paracrine and OSF action on cumulus cells. The resulting cumulus matrix gene expression and structure is thus indicative of follicular maturity and oocyte quality, which are predictive of developmental competence.
The oocyte: arbiter of its own release
Throughout follicle growth, oocytes are arrested at meiotic prophase-I, but progress through a series of maturational stages, both cytoplasmic and nuclear, which determine its eventual developmental competence. The timing whereby the oocyte reaches its fully grown state is determined by bi-directional feedback signalling between the oocyte and its companion somatic cells and is critical to its post-fertilization developmental success (De La Fuente and Eppig, 2001). By the pre-ovulatory stage, mature oocytes are transcriptionally silent until activation of the embryonic genome occurs after fertilization. The mechanism of transcription silencing involves sequestering of transcription factors and RNA polymerases into inactive complexes as well as condensation of chromatin structure (De La Fuente et al., 2004) and histone deacetylation (Borsuk and Milik, 2005). This specific chromatin organization is a key aspect of oocyte developmental competence and meiotic progression after ovulatory initiation. Thus, the follicle-oocyte dialogue synchronizes the activation of oocyte growth and developmental capacity with its release, and this process is co-ordinated by signals received and transmitted through ovarian somatic cells, in particular the cumulus cells.
Follicular control of oocyte developmental competence
Cyclic AMP transport
Meiotic arrest is maintained prior to ovulation through communication of a suppressive effect from the surrounding somatic cells; purines in follicular fluid (Eppig et al., 1985) and high levels of cAMP in the oocyte are particularly important (Schultz et al., 1983; Bornslaeger et al., 1986). During folliculogenesis, cumulus cells synthesize cAMP in response to FSH and deliver cAMP through gap junctions shared with oocytes (Webb et al., 2002). Intra-oocyte cAMP levels are balanced by a range of factors, including the rate of synthesis and metabolism in each compartment and the maintenance of gap junctional transport from cumulus to oocyte. The gonadotrophin surge elicits a strong increase in cAMP in cumulus cells; yet in the oocyte, cAMP levels fall, promoting the resumption of meiosis. This partitioning of responses is in part achieved by cessation of oocyte and somatic cell gap junctional communication (Dekel et al., 1981) via phosphorylation of gap junction proteins and down-regulation of connexins that assemble gap junctions (Granot and Dekel, 1994; Kalma et al., 2004; Sela-Abramovich et al., 2005). Physical dissociation of cumulus cells from the oocyte also occurs as a result of COC expansion (Larsen, 1996). Partitioning of cAMP metabolizing phosphodiesterases also accounts for the disparity in cAMP levels in oocytes and cumulus. Oocytes metabolize cAMP via a specific PDE3 isoform (Wiersma et al., 1998; Richard et al., 2001; Shitsukawa et al., 2001), and selective pharmacological inhibition of PDE3 blocks meiotic resumption without affecting ovulation in rats (Wiersma et al., 1998). Likewise, PDE3 inhibition delays spontaneous or gonadotrophin-induced meiotic resumption in the rhesus monkey (Jensen et al., 2002; Jensen et al., 2005) and spontaneous maturation of denuded human oocytes (Nogueira et al., 2006).
Oocytes enter a critical stage of development in response to diminishing cAMP after the LH surge, resuming meiosis and progressing through precisely synchronized nuclear and cytoplasmic maturation, to achieve full developmental competence. Nuclear maturation involves breakdown of the oocyte nuclear membrane, formation of the meiotic spindle and extrusion of half the oocyte chromosomes into the first polar body to complete metaphase-I.
Meiosis-activating sterol
Progression of oocytes through metaphase-I involves accumulation of a meiosis-activating sterol (MAS; 4,4-dimethyl-5a-cholest-8,14,24,triene-3ß-ol) after the LH-surge (Byskov et al., 1995; Marin Bivens et al., 2004a; Marin Bivens et al., 2004b), which may contribute to synchronization of oocyte developmental competence with follicular development and ovulation. However, whether MAS is a critical mediator of meiotic progression remains controversial (Tsafriri et al., 2005), and control of meiotic resumption is a complex process certainly modulated by numerous input signals.
Paracrine control of oocyte maturation by follicular compartments
Signals produced by cumulus cells appear to influence oocyte maturation even after loss of gap junctions, since bovine-denuded oocytes matured in culture have minimal developmental potential unless intact COCs are co-cultured (Luciano et al., 2005). Cytoplasmic maturation of human oocytes is also improved by maintaining cumulus cells in the IVF culture (Goud et al., 1998). Maturation of bovine oocytes in co-culture with isolated cumulus cell monolayers or conditioned media induced no similar effect (Ikeda et al., 2000; Tanghe et al., 2003; Luciano et al., 2005), suggesting that the appropriate organization and communication between the cumulus cells, oocyte and/or the associated matrix is important for promoting oocyte developmental competence. Calcium mobilization by Egf has also been suggested to depend upon the organization of cumulus cells in intact mouse COC (O'Donnell et al., 2004). Human immature oocytes matured in vitro without cumulus cells also exhibit a lack of co-ordination of maturation processes (Combelles et al., 2002) and co-culture with cumulus cells in a 3D matrix is coming under investigation as a means to more successfully generate mature, MII-arrested oocytes in vitro (Combelles et al., 2005). Co-culture of human embryos with cumulus cells has been also suggested to increase fertilization rate (Tanghe et al., 2003), embryo grade (Saito et al., 1994) and clinical pregnancy rates (Carrell et al., 1999). These results suggest that cumulus cells support both oocyte and embryo development.
Three recently described paracrine signals from follicular somatic compartments may enhance oocyte maturation. Thecal cells produce insulin-like factor-3 (Insl-3), a member of the relaxin family of insulin-related peptides, in response to the LH- surge (Bathgate et al., 1999; Kawamura et al., 2004) (Figure 1). The Insl-3 receptor (LGR-8) present on oocytes is a Gi-coupled GPCR. Oocytes (and male germ cells) undergo a reduction in intracellular cAMP production via LGR8/Gi-mediated inhibition of adenylate cyclase activity. The reduction in cAMP initiates meiotic resumption and germinal vesicle breakdown (Kawamura et al., 2004). Insl-3 null mice exhibit female subfertility and disrupted estrous cyclicity (Nef and Parada, 1999), but oocyte competence has not been analysed in detail. Leydig cells of the human testis produce Insl-3, but its expression has not been investigated in women.
Secondly, brain-derived neurotropic factor (BDNF) is expressed by mural and cumulus granulosa cells after hCG treatment and its receptor TrkB is exclusively expressed in the oocyte. Recombinant BDNF enhanced the progression of in vitro fertilized mouse embryos to blastocysts, whereas treatment of mice with a Trk receptor antagonist in vivo during the periovulatory period reduced the progression of fertilized embryos to blastocyst (Kawamura et al., 2005).
Finally, meiotic resumption and oocyte developmental competence are also increased by Egf-L signals from granulosa cells. Egf itself has long been known to be an effective inducer of oocyte meiotic resumption in mice (Downs et al., 1998; Smitz et al., 1998; Su et al., 2002), cows (Lorenzo et al., 2001), pigs (Grupen et al., 1997; Procházka, 2000; Prochazka et al., 2003) and rats (Dekel and Sherizly, 1985). The Egf-L amphiregulin, epiregulin and betacellulin are also efficient inducers of meiosis in follicle enclosed oocytes from mice (Park et al., 2004) or rats (Ashkenazi et al., 2005; Tsafriri et al., 2005). Signalling through the Egf pathway is superior to FSH in promoting oocyte polarity and developmental competence (De La Fuente et al., 1999; Sanfins et al., 2003; Sanfins et al., 2004; Rossi et al., 2006). This effect is specifically mediated through cumulus cells (O'Donnell et al., 2004; Jamnongjit et al., 2005). Furthermore, Egf-L signals appear to require synthesis of Egf-L followed by proteolytic cleavage and changes in cumulus cell gene expression, which potently mediate the LH effect on meiotic resumption in vivo (Ashkenazi et al., 2005). Protease activators of Egf-L, though as yet unidentified, may place a further level of control on oocyte maturation and ovulation via somatic follicular cells.
The multiple overlapping mechanisms of follicular control of oocyte maturation testify to the complexity and fine balance of this process. Fertilization and especially the development of fertilized oocytes to healthy blastocyst is limited by the quality of the oocyte, and cumulus cells play a critical role in determining oocyte developmental potential both before and after ovulation.
Oocyte control of ovulation rate
Genetic studies have confirmed that oocytes produce signals that can regulate the ovulation quota of humans, sheep and mice. The source of this effect is difficult to confirm because of differing regulation and activity of Gdf-9 and BMP-15 in species with different ovulation rates. Mice lacking Gdf-9 exhibit a block in folliculogenesis at the early primary (type-IV) stage (Dong et al., 1996; Elvin et al., 1999a; Elvin et al., 1999b), and thus no pre-ovulatory follicles occur. Mutations in the related BMP-15 gene in mice produce subtle alterations of follicle growth and ovulation, but BMP-15 appears to play a more important role in combination with Gdf-9 (Yan et al., 2001). It is thought that a reduced role for BMP-15 has evolved in mice because of alteration of the pro-domain sequence that perturbs secretion of mouse BMP-15, leading to reduced function (Hashimoto et al., 2005). In sheep, BMP-15 is a critical, but complex mediator of folliculogenesis since ewes with a single copy mutant gene have an increased ovulation rate, whereas homozygosity for null BMP-15 mutations results in complete ovulatory failure (Galloway et al., 2000). The same relationship between gene dosage and ovulation rate has been observed for one mutation of Gdf-9 in sheep (Hanrahan et al., 2004). Short-term immunoneutralization of Gdf-9 or BMP-15 can also raise ovulation rates in sheep without overtly harming the developmental trajectories of the resulting embryos and fetuses (Juengel et al., 2004), but importantly, longer term neutralization perturbs follicular and oocyte function (Juengel et al., 2002). Humans carrying a heterozygous mutation in BMP-15 suffer ovarian failure and infertility (Di Pasquale et al., 2004), whereas ovulation rate may be enhanced by a mutation of Gdf-9 (Montgomery et al., 2004). Oocyte health, follicle growth and ovulation are dependent on some level of Gdf-9 and BMP-15 signalling, and mildly reducing their activity elicits an intraovarian response increasing ovulatory follicle development. Thus, in most species including humans Gdf-9 and BMP-15 are oocyte-specific genes that influence ovulation via responses elicited in the follicular somatic cells. This oocyte/somatic cell regulatory loop guides follicle growth and sets the ovulation quota limit (Moore et al., 2004; McNatty et al., 2005).
Appropriate promotion of periovulatory cumulus gene expression can be achieved only by matured oocytes (Joyce et al., 2001; Latham et al., 2004; Eppig et al., 2005), but this effect also requires an interaction with unknown products of granulosa cells from mature follicles (Joyce et al., 2001). Thus, stage-specific feedback communication between oocytes and somatic cells co-ordinates control of meiotic resumption and ovulation, whereas immature or incompetent oocytes fail to interact appropriately in this communication loop and are not released (Albertini, 1992; Albertini et al., 2003; Plancha et al., 2005). Follicular and oocyte signals converge through the cumulus cells that transmit maturation-promoting effects to the oocyte. It can be supposed that a good quality follicular/oocyte unit generates a signature gene expression profile in cumulus cells. Indeed, correlative relationships between follicular gene expression levels and oocyte developmental trajectory have been reported in humans (McKenzie et al., 2004; Hasegawa et al., 2005; Zhang et al., 2005). The dependence on this fine internal balance is demonstrated when deviations are introduced in the normal regulation of follicle and oocyte maturation such as through exogenous ovarian hyperstimulation which overrides the normal control permitting ovulation of oocytes with inferior developmental competence (Sato and Marrs, 1986; Yun et al., 1987; Greve and Callesen, 2001).
Structural remodelling and rupture of the follicle
Prior to ovulation, the ovarian and follicular walls present a multilaminar barrier that must be breached for release of the COC into the oviduct. High tensile strength of these tissues is derived from the granulosa layers, the follicular basement membrane, the theca interna and externa layers with associated collagenous ECM, the type-I collagen-rich tunica albuginea and the ovarian surface epithelium with its underlying basal lamina. After the LH surge, cellular and ECM layers at the follicular apex become thinned and the circumferential basement membrane is degraded through proteolytic actions. At the same time, the entire follicular structure is remodelled through rapid angiogenesis and infiltration of the neovasculature, thecal cells and immune cells into the antral space, accompanying differentiation of granulosa cells to luteal cells. Thus, remodelling of the follicle wall structure is an important process for oocyte release as well as CL formation, requiring basal lamina degradation and driven by rapid angiogenic morphogenesis of the thecal layers.
Thecal and vascular remodelling
Several proteases have been shown to increase in expression and activity in the periovulatory period. These include matrix metalloproteinases, MMP-2 (Russell et al., 1995; Curry et al., 2001; Jo et al., 2004; Smith et al., 2005), MMP-9 (Hagglund et al., 1999; Robker et al., 2000), MT1-MMP (MMP-14) and MMP-19 (Hagglund et al., 1999; Jo and Curry, 2004; Jo et al., 2004), ADAMTS-1 (Espey et al., 2000), cathepsin L (Robker et al., 2000) and kallikreins (Holland et al., 2001). Type-1 collagenase, MMP-1, is induced in primate granulosa cells by LH (Stouffer and Duffy, 2003). The role of MMP, their inhibitors (TIMP) and other proteases in ovarian function has been reviewed elsewhere (Curry and Osteen 2003; Ohnishi et al., 2005) and few have emerged as obligatory for ovulatory follicle remodelling. This is, at least in part, due to redundant actions of multiple proteases with overlapping substrate specificity being co-ordinately involved in matrix degradation of the various follicular compartments. Thus, although protease activities are certainly required for the remodelling processes, the identification of the key individual enzymes is ongoing.
Pharmacological inhibition, which blocks activity of specific enzyme families, is likely to be a better experimental approach than targeted genetic deletions because it can account for redundant functionality as well as compensatory enzyme expression. Reduced ovulation has been reported using broad spectrum MMP inhibitors administered into the ovarian bursa (Reich et al., 1985) or in vitro perfused rat ovaries (Brannstrom et al., 1988; Butler et al., 1991). Furthermore, specific intrafollicular inhibition of MMP-2 in ewes blocked ovulation and the normal thecal vascular projections associated with CL morphogenesis (Gottsch et al., 2002). This result was very similar to the effect seen in earlier studies actively immunising ewes against the N-terminal peptide of inhibin-, which reduced MMP-2 activation, thecal/vascular invagination and ovulation (Russell et al., 1995).
Simultaneous with follicular basement membrane remodelling, very rapid angiogenesis is initiated in the follicular vasculature (Hazzard and Stouffer, 2000). This process is required for neovascularization of the forming corpus luteum; however, several models of ovulatory insufficiency show a link between disruption of angiogenesis and ovulatory failure. Angiogenesis in ovulating follicles appears to be primarily regulated through induction of VEGF expression (Neulen et al., 1995; Gomez et al., 2003). Several approaches that block the action of VEGF and hence angiogenesis result in a severe failure of follicle wall remodelling and ovulation, resulting in entrapment of oocytes in corpora lutea and infertility. These studies include immuno-inhibition of VEGF-A (Fraser et al., 2000; Zimmermann et al., 2001) or VEGF receptor-2 in primates (Zimmermann et al., 2002; Zimmermann et al., 2003), antagonism with soluble VEGFR decoy in rats (Ferrara et al., 1998) and primates (Wulff et al., 2001; Hazzard et al., 2002; Wulff et al., 2002) or inhibition of angiogenesis through angiopoietin antagonism in monkeys (Xu et al., 2005). Thus, VEGF mediated angiogenic remodelling of the follicle wall is necessary for efficient follicle rupture.
Contribution of leukocytes
Considerable evidence indicates that immune cells contribute to the ECM remodelling of follicular rupture (Wu et al., 2004). In particular macrophages and neutrophils accumulate around periovulatory follicles, potentially through binding to the ICAM-1 leukocyte receptor induced in ovarian vascular endothelium of ovulating follicles (Bonello et al., 2004). The important role for immune cells in ovulation is demonstrated through enhanced ovulation rate after their addition to in vitro ovarian perfusion medium (Hellberg et al., 1991).
Macrophages gather in the follicular thecal layers during follicle growth of many species (Suzuki et al., 1998; Tamura et al., 1998) and may secrete factors involved in follicular growth and development (Matsuura et al., 2002; Wang et al., 2005). Macrophages are also specifically recruited in greater numbers after the LH surge and mediate aspects of ovulation since depletion of macrophages in perfused mouse ovaries results in reduced ovulation rate and delayed estrous cycle progression (Van der Hoek et al., 2000).
Neutrophil invasion of the perifollicular thecal tissues after the LH surge has been demonstrated in humans (Brannstrom et al., 1994), rats (Brannstrom et al., 1993), sheep (Cavender and Murdoch, 1988) and pigs (Standaert et al., 1991), particularly near the follicular apex (Brannstrom et al., 1993). Neutrophils are important mediators of tissue remodelling (Vaday et al., 2001) and may secrete neutrophil elastase (MMP-8) and other matrix-degrading proteases in ovulating ovaries. Acute neutrophil depletion using a cytotoxic antibody reduced ovulation rate in rats demonstrating their role in facilitating ovulation (Brannstrom et al., 1995).
Leukocytes release cytokines and free radicals, such as NO which are likely to mediate the inflammatory-like aspects of ovulation and contribute to processes of tissue destruction (Brannstrom and Enskog 2002; Minge et al., 2005). Studies inhibiting NOS in vivo or ex vivo in rodents demonstrate that NO is required for ovulation (Shukovski and Tsafriri 1994; Bonello et al., 1996). Both inducible NOS and endothelial NOS isoforms have been identified in the ovary, predominantly in the thecal compartment, and are hormonally regulated (Jablonka-Shariff and Olson, 1997), but their specific roles are still unclear. It is likely that NO acts at several levels in the follicle to optimally control blood vessel dilation and permeability, and leukocyte recruitment (Bonello et al., 1996), in addition to the stimulation of nitrosylation-sensitive signalling cascades in neighbouring ovarian compartments.
Platelet activating factor (PAF), a mediator of inflammatory processes, may also mediate aspects of ovulation. Inhibition of PAF blocked ovulation in rats (Abisogun et al., 1989; Kikukawa et al., 1991), and PAF secretion was found to increase in ovulating sheep ovaries (Alexander et al., 1990); however, in rat ovaries, PAF mRNA decreased after ovulatory hCG (Espey et al., 1989) and the PAF receptor was found to be the lowest in proestrous mouse ovaries (Lash and Legge, 2001). Thus, PAF may play a role in the inflammatory aspects of ovulation; however, this role appears complex or may vary among species.
Basement membrane degradation
The basement membrane, which surrounds the granulosa layers of every follicle, separates these epithelial-like cells from the mesenchyme-derived thecal cells. Laminin, collagen type-IV, fibronectin and proteoglycans are the major ECM components of this basement membrane found in most species (Bortolussi et al., 1989). Versican is also abundant either within or adjacent to the basement membrane of preovulatory follicles (Rodgers et al., 2003; Russell et al., 2003c). An intriguing, but unusual study that induced precocious ovulation in 13-day-old mice by blocking the interaction between laminin and its integrin receptor (6ß1) (Nakamura et al., 1997) suggests that disruption of cellular interactions with basement membrane components can promote follicular rupture. After the LH surge, the basement membrane ECM undergoes rapid dissolution early in the periovulatory period (Palotie et al., 1984). This is essential both for apical perforation and passage of the COC, as well as for the basal infiltration of theca and vascular components during the formation of the corpus luteum structure (Rodgers et al., 2003).
Structural components of basement membranes, such as collagen type-IV and laminin, are targets for proteolysis by specific enzymes including the gelatinases (MMP-2 and -9), plasmin and the plasminogen activators (tPA and uPA). Gelatinases and uPA degrade and remodel the basal lamina of the ovarian surface epithelium (Yang et al., 2004), and thus it is likely they target the follicular basal lamina as well. Interestingly, a cohort of infertile women undergoing IVF showed significantly lower follicular fluid MMP-2 and -9 as well as increased tissue inhibitor of metalloproteinases (TIMP-1) suggestive of impaired basement membrane remodelling in these infertile women (D'Ascenzo et al., 2004). The gelatinases, MMP-2 and -9, are found in growing follicles of most species and MMP-9, in particular, is induced during ovulation (Robker et al., 2000; Curry et al., 2001; D'Ascenzo et al., 2004). However, null mutation of each of these genes has not yielded any evidence that they are essential for basement membrane degradation or follicular rupture (Itoh et al., 1997; Vu et al., 1998). Similarly, inhibitor studies and null mutation in mice indicate that the PAs and plasmin are not individually or jointly essential for ovulation (Ny et al., 1999). Furthermore, among the many anovulatory experimental models, including those with altered thecal/vascular remodelling (Russell et al., 1995; Gottsch et al., 2002), there are no experimental reports with demonstrated persistence of the basement membrane structure after induction of the ovulatory cascade. Thus, a critical protease or enzyme system has not yet been elucidated in this process. This indicates that basement membrane degrading proteases are highly redundant, but also suggests that the basal lamina does not present a key barrier to COC release in ovulatory failure.
Rupture at the follicular apex
At the follicular apex, the cellular and ECM layers are first thinned and then breached by a combination of proteolytic action as well as cell death and migration of thecal fibroblasts away from the apical site (Espey, 1994, 1998). The surface epithelial cells die through apoptosis and are sloughed from the follicle apex region (Murdoch et al., 1999). Fibrillar collagenase activity is presumably required to degrade the thick collagen fibres found in the theca and tunica albuginea (Espey and Rondell, 1968; Espey, 1970; Tsafriri, 1995). In the rabbit, pro-MMP-1 (interstitial type-I collagenase) was identified in increased concentration at the follicle apex around the time of ovulation (Tadakuma et al., 1993). However, as with basement membrane degradation, enzymes directly involved have not been clearly identified and there is little strong evidence to implicate MMP-1 or other specific proteases in follicle rupture. Gene deletions of MMPs with interstitial collagenase activity in mice have yielded no follicular or ovulatory defects. These include MMP-3 (Alexander et al., 2001), MMP-7 (Wilson et al., 1997) and MMP-13 (Stickens et al., 2004). The Medaka Teleost (bony fish) provides a more primitive and perhaps simplified model of rupture of the follicle apex in which combined actions of membrane-bound matrix metalloproteinases MT1-MMP and MT2-MMP along with MMP-2 were shown to be necessary and sufficient (Ogiwara et al., 2005). However, the degree to which this species model reflects the mammalian system is not known. Inactivation of the MT1-MMP (MMP-14) gene in mice results in quite severe growth and connective tissue defects, yet ovulation was not apparently influenced (Holmbeck et al., 1999). Thus, it can be concluded that ovulation is assured through redundant and overlapping activities of proteases that rupture the follicular apex.
A few studies have reported treatments that encourage follicle rupture at regions other than the apex. Inhibition of COX-2 using indomethacin disrupted the guidance of follicle rupture, causing random rupture into the interstitium or vascular spaces at the basal side of some follicles (Osman and Dullaart, 1976; Gaytan et al., 2003). But surprisingly, this treatment did not appear to block cumulus expansion, which is an expected consequence of blocking COX-2 action (Eppig, 1981; Downs and Longo, 1983; Davis et al., 1999; Ochsner et al., 2003b). Basal rupture of follicles was also reported after inhibition of the plasminogen activator system by antibodies or antiplasmin administered intrabursally (Tsafriri et al., 1989). This may result from surrounding the ovary in a high dose of antiprotease, since in other studies, particularly in plasmin or plasminogen-activator-deficient mice, misguided follicle rupture location has not been reported.
To date, the only individual protease that has been demonstrated to be essential for follicle rupture is ADAMTS-1. Aspects of follicle matrix dissolution appear to be dependent on ADAMTS-1, since ADAMTS-1 null mice have minimal ovulation rates and exhibit oocytes retained within luteinized follicles (Mittaz et al., 2004). The incidence of pregnancy and the litter size of females that do become pregnant are each reduced >50% (Table I) (Shindo et al., 2000; Mittaz et al., 2004). Functionally related family members are also expressed in the bovine ovary (Madan et al., 2003), and in mice, ADAMTS-4 is induced by ovulatory LH in mural and cumulus cells and ADAMTS-5 is expressed in granulosa cells of all healthy follicles (Richards et al., 2005). Although these related ADAMTS proteases likely rescue aspects of ADAMTS-1 action in ADAMTS-1–/– ovaries, reduced fertility is still observed. Mice deficient in ADAMTS-4 or ADAMTS-5 show no fertility defects (Stanton et al., 2005), indicating that ADAMTS-1 uniquely plays a critical and non-redundant role in ovulation.
Reduced expression of ADAMTS-1 was also identified in PRKO mice (Robker et al., 2000) and RIP140 null mice (White et al., 2000), both models with complete, and specific, defects in follicle rupture. The anovulation of PRKO and RIP140 null mice is more severe than that of ADAMTS-1 null mice and likely due to the loss of co-operative proteases. ADAMTS-4 and -5 are not dysregulated in PRKO follicles (Richards et al., 2005), but other ADAMTS family members have not been specifically assessed. Members of the MMP family showed no difference in expression or activation, whereas cathepsin L, a cysteine protease active in both lysosomes and the extracellular space, is deficiently expressed in PRKO mice (Robker et al., 2000). This and perhaps other unidentified proteins may act in concert with ADAMTS-1 in tissue destruction and follicular rupture. It can be concluded that PR is a key mediator of ovulatory structural remodelling involving at least two protease classes. In addition, ADAMTS-1 localization and cleavage of versican in the COC and its essential role in ovulation again implicate the COC matrix structure as key to ovulation. Overall, tissue remodelling appears co-ordinated across the follicular compartments by a series of tightly regulated proteases, many with overlapping roles.
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TOPAbstractIntroductionConclusionReferences |
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The molecular processes set into motion when a pre-ovulatory follicle receives an ovulatory stimulus are a complex, interconnected cascade of events that occur in the theca, mural granulosa, cumulus and oocyte compartments. Remarkable reprogramming of cell functions occurs in all tissue compartments and restructuring of the follicle ensues. The success of these events is of premium importance to successful establishment of pregnancy and to the continuing developmental potential of the resultant embryo. Many questions remain to be definitively answered however, and, in particular, the relevance of species differences and the aspects most relevant to human ovulatory control must be defined.
This review of the state of current research leads to the conclusion that the oocyte's companion somatic cells, the cumulus cells, play a central role in ovulation. In growing follicles, the cumulus cells help maintain oocyte meiotic arrest as well as providing nutrients that support the metabolic needs and development of the oocyte. Success of ovulation is intimately coupled to cumulus matrix expansion, which responds to endocrine, mural granulosa, theca and oocyte inputs. The resultant formation of cumulus matrix is thus hypothesized to facilitate detachment and expulsion of the COC mass from the follicle. Ovulatory changes in the cumulus mass involve a complex series of tissue remodelling and transcriptional changes in cumulus cells that contribute to meiotic resumption. Evidence is accumulating to demonstrate that the capacity of oocytes to support cumulus gene expression is linked to the oocyte's developmental competence, a mechanism that elegantly co-ordinates oocyte release with developmental potential.
The ovulation process may therefore be considered a checkpoint event whereby healthy and meiotically competent oocytes engineer their own release. Conversely, poor endocrine, follicular or oocyte signals indicate a suboptimal cycle that would result in production of inferior oocytes. Multifactorial regulation of ovulation may ensure the prevention of incompetent oocytes from entering the reproductive pool through anovulation and/or prevention of fertilization. Thus, the co-ordination and synchronization of endocrine, paracrine, immune and metabolic signals acting mainly through the cumulus compartment exert control on oocyte maturation, developmental potential and ultimately ovulation.
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Abisogun AO, Braquet P, Tsafriri A. (1989) The involvement of platelet activating factor in ovulation. Science 20:381–383.
Alam H, Maizels ET, Park Y, et al. (2004) Follicle-stimulating hormone activation of hypoxia-inducible factor-1 by the phosphatidylinositol 3-kinase/AKT/ras homolog enriched in brain (Rheb)/mammalian target of rapamycin (mTOR) pathway is necessary for induction of select protein markers of follicular differentiation. J Biol Chem 279:19431–40.
Albertini DF. (1992) Cytoplasmic microtubular dynamics and chromatin organization during mammalian oogenesis and oocyte maturation. Mutat Res/Genet Toxicol 296:57–68.
Albertini DF, Sanfins A, Combelles CM. (2003) Origins and manifestations of oocyte maturation competencies. Reprod Biomed Online 6:410–15.[Medline]
Alexander BM, Van Kirk EA, Murdoch WJ. (1990) Secretion of platelet-activating factor by periovulatory ovine follicles. Life Sci 47:865–8.[CrossRef][Web of Science][Medline]
Alexander CM, Selvarajan S, Mudgett J, et al. (2001) Stromelysin-1 Regulates Adipogenesis during Mammary Gland Involution. J Cell Biol 152:693–703.
Amsterdam A, Koch Y, Lieberman ME, et al. (1975) Distribution of binding sites for human chorionic gonadotropin in the preovulatory follicle of the rat. J Cell Biol 67:894–900.
Arias J, Alberts AS, Brindle P, et al. (1994) Activation of cAMP and mitogen responsive genes relies on a common nuclear factor. Nature 370:226–229.[CrossRef][Medline]
Ashkenazi H, Cao X, Motola S, et al. (2005) Epidermal growth factor family members: endogenous mediators of the ovulatory response. Endocrinology 146:77–84.
Athanasiou S, Bourne TH, Khalid A, et al. (1996) Effects of indomethacin on follicular structure, vascularity, and function over the periovulatory period in women. Fertil Steril 65:556–60.[Web of Science][Medline]
Baird DT, Brown A, Cheng L, et al. (2003) Mifepristone: a novel estrogen-free daily contraceptive pill. Steroids 68:1099–105.[CrossRef][Web of Science][Medline]
Ball G, Leibfried M, Lenz R, et al. (1983) Factors affecting successful in vitro fertilization of bovine follicular oocytes. Biol Reprod 28:717–25.[Abstract]
Barrionuevo MJ, Schwandt RA, Rao PS, et al. (2000) Nitric oxide (NO) and interleukin-1beta (IL-1beta) in follicular fluid and their correlation with fertilization and embryo cleavage. Am J Reprod Immunol 44:359–64.[Medline]
Bathgate R, Moniac N, Bartlick B, et al. (1999) Expression and regulation of relaxin-like factor gene transcripts in the bovine ovary: differentiation-dependent expression in theca cell cultures. Biol Reprod 61:1090–8.
Biggers JD, Whittingham DG, Donahue RP. (1967) The pattern of energy metabolism in the mouse oocyte and zygote. Proc Natl Acad Sci U S A 58:560–7.
Boerboom D, Russell DL, Richards JS, Sirois J. (2003) Regulation of transcripts encoding ADAMTS-1 (a disintegrin and metalloproteinase with thrombospondin-like motifs-1) and progesterone receptor by human chorionic gonadotropin in equine preovulatory follicles. Endocrinology Submitted December 2002.
Bonello N, Jasper MJ, Norman RJ. (2004) Periovulatory expression of intercellular adhesion molecule-1 in the rat ovary. Biol Reprod 71:1384–90.
Bornslaeger EA, Mattei P, Schultz RM. (1986) Involvement of cAMP-dependent protein kinase and protein phosphorylation in regulation of mouse oocyte maturation. Dev Biol 114:453–62.[CrossRef][Web of Science][Medline]
Bonello N, McKie K, Jasper M, et al. (1996) Inhibition of nitric oxide: effects on interleukin-1 beta-enhanced ovulation rate, steroid hormones, and ovarian leukocyte distribution at ovulation in the rat. Biol Reprod 54:436–45.[Abstract]
Borsuk E and Milik E. (2005) Fully grown mouse oocyte contains transcription inhibiting activity which acts through histone deacetylation. Mol Reprod Dev 71:509–15.[CrossRef][Web of Science][Medline]
Bortolussi M, Marini G, Reolon ML. (1979) A histochemical study of the binding of 125I-HCG to the rat ovary throughout the estrous cycle. Cell Tissue Res 197:213–26.[Web of Science][Medline]
Bortolussi M, Zanchetta R, Doliana R, et al. (1989) Changes in the organization of the extracellular matrix in ovarian follicles during the preovulatory phase and atresia. An immunofluorescence study. Basic Appl Histochem 33:31–8.[Web of Science][Medline]
Bost F, Diarra-Mehrpour M, Martin J-P. (1998) Inter- -trypsin inhibitor proteoglycan family. A group of proteins binding and stabilizing the extracellular matrix. Eur J Biochem 252:339–46.[Web of Science][Medline]
Bourguignon LYW. (2001) CD44-mediated oncogenic signaling and cytoskeleton activation during mammary tumor progression. J Mammary Gland Biol Neoplasia 6:287–97.[CrossRef][Web of Science][Medline]
Brannstrom M and Enskog A. (2002) Leukocyte networks and ovulation. J Reprod Immunol 57:47–60.[CrossRef][Web of Science][Medline]
Brannstrom M, Mayrhofer G, Robertson S. (1993) Localization of leukocyte subsets in the rat ovary during the periovulatory period. Biol Reprod 48:277–86.[Abstract]
Brannstrom M, Woessner JFJ, Koos RD, et al. (1988) Inhibitors of mammalian tissue collagenase and metalloproteinases suppress ovulation in the perfused rat ovary. Endocrinology 122:1715–21.
Brannstrom M, Pascoe V, Norman RJ, et al. (1994) Localization of leukocyte subsets in the follicle wall and in the corpus luteum throughout the human menstrual cycle. Fertil Steril 61:488–95.[Web of Science][Medline]
Brannstrom M, Bonello N, Norman RJ, et al. (1995) Reduction of ovulation rate in the rat by administration of a neutrophil-depleting monoclonal antibody. J Reprod Immunol 29:265–70.[CrossRef][Web of Science][Medline]
Bukovsky A, Chen T, Wimalasena J, et al. (1993) Cellular localization of luteinizing hormone receptor immunoreactivity in the ovaries of immature, gonadotropin-primed and normal cycling rats. Biol Reprod 48:1367–82.[Abstract]
Butler TA, Zhu C, Mueller RA, et al. (1991) Inhibition of ovulation in the perfused rat ovary by the synthetic collagenase inhibitor SC 44463. Biol Reprod 44:1183–8.[Abstract]
Byskov AG, Andersen CY, Nordholm L, et al. (1995) Chemical structure of sterols that activate oocyte meiosis. Nature 374:559–62.[CrossRef][Medline]
Carrell DT, Peterson CM, Jones KP, et al. (1999) A simplified coculture system using homologous, attached cumulus tissue results in improved human embryo morphology and pregnancy rates during in vitro fertilization. J Assist Reprod Genet 16:344–9.[CrossRef][Web of Science][Medline]
Carrette O, Nemade RV, Day AJ, et al. (2001) TSG-6 is concentrated in the extracellular matrix of mouse cumulus oocyte complexes through hyaluronan and inter-alpha-inhibitor binding. Biol Reprod 65:301–8.
Castillo GM and Templeton DM. (1993) Subunit structure of bovine ESF (extracellular-matrix stabilizing factor(s)): A chondroitin sulfate proteoglycan with homology to human I[alpha]i (inter-[alpha]-trypsin inhibitors). FEBS Lett 318:292–6.[CrossRef][Web of Science][Medline]
Cavender J and Murdoch W. (1988) Morphological studies of the microcirculatory system of periovulatory ovine follicles. Biol Reprod 39:989–97.[Abstract]
Chandras C, Ragoobir J, Barrett GE, et al. (2004) Roles for prostaglandins in the steroidogenic response of human granulosa cells to high-density lipoproteins. Mol Cell Endocrinol 222:1–8.[CrossRef][Web of Science][Medline]
Chang SY, Kang H-Y, Lan K-C, et al. (2005) Expression of steroid receptors, their cofactors, and aromatase in human luteinized granulosa cells after controlled ovarian hyperstimulation. Fertil Steril 83:1241–7.[CrossRef][Web of Science][Medline]
Channing CP, Bae IH, Stone SL, et al. (1981) Porcine granulosa and cumulus cell properties. LH/hCG receptors, ability to secrete progesterone and ability to respond to LH. Mol Cell Endocrinol 22:359–70.[CrossRef][Web of Science][Medline]
Chen L, Russell PT, Larsen WJ. (1993) Functional significance of cumulus expansion in the mouse: roles for the preovulatory synthesis of hyaluronic acid within the cumulus mass. Mol Reprod Dev 34:87–93.[CrossRef][Web of Science][Medline]
Chen L, Russell P, Larsen W. (1994) Sequential effects of follicle-stimulating hormone and luteinizing hormone on mouse cumulus expansion in vitro. Biol Reprod 51:290–5.[Abstract]
Chen L, Zhang H, Powers RW, et al. (1996) Covalent linkage between proteins of the inter-alpha -inhibitor family and hyaluronic acid is mediated by a factor produced by granulosa cells. J Biol Chem 271:19409–14.
Cherr GN, Yudin AI, Katz DF. (1990) Organization of the hamster cumulus extracellular matrix: a hyaluronate-glicoprotein gel, which modulates sperm access to the oocyte. Dev Growth Differ 32:353–65.[Medline]
Choi J-H, Choi K-C, Auersperg N, et al. (2005) Gonadotropins upregulate the epidermal growth factor receptor through activation of mitogen-activated protein kinases and phosphatidyl-inositol-3-kinase in human ovarian surface epithelial cells. Endocr Relat Cancer 12:407–21.
Christenson LK, Johnson PF, McAllister JM, et al. (1999) CCAAT/enhancer-binding proteins regulate expression of the human steroidogenic acute regulatory protein (StAR) gene. J Biol Chem 274:26591–8.
Clemens JW, Robker RL, Kraus WL, et al. (1998) Hormone induction of progesterone receptor (PR) messenger ribonucleic acid and activation of PR promoter regions in ovarian granulosa cells: evidence for a role of cyclic adenosine 3',5'-monophosphate but not estradiol. Mol Endocrinol 12:1201–14.
Cohen BD, Nechamen CA, Dias JA. (2004) Human follitropin receptor (FSHR) interacts with the adapter protein 14-3-3[tau]. Mol Cell Endocrinol 220:1–7.[CrossRef][Web of Science][Medline]
Combelles CMH, Cekleniak NA, Racowsky C, et al. (2002) Assessment of nuclear and cytoplasmic maturation in in-vitro matured human oocytes. Hum Reprod 17:1006–16.
Combelles CMH, Fissore RA, Albertini DF, et al. (2005) In vitro maturation of human oocytes and cumulus cells using a co-culture three-dimensional collagen gel system. Hum Reprod 20:1349–58.
Cottom J, Salvador LM, Maizels ET, et al. (2003) Follicle-stimulating hormone activates extracellular signal-regulated kinase but not extracellular signal-regulated kinase kinase through a 100-kda phosphotyrosine phosphatase. J Biol Chem 278:7167–79.
Curry TE Jr and Osteen KG. (2003) The matrix metalloproteinase system: changes, regulation, and impact throughout the ovarian and uterine reproductive cycle. Endocr Rev 24:428–65.
Curry TE Jr, Song L, Wheeler SE. (2001) Cellular localization of gelatinases and tissue inhibitors of metalloproteinases during follicular growth, ovulation, and early luteal formation in the rat. Biol Reprod 65:855–65.
Das S, Maizels E, DeManno D, et al. (1996) A stimulatory role of cyclic adenosine 3',5'-monophosphate in follicle- stimulating hormone-activated mitogen-activated protein kinase signaling pathway in rat ovarian granulosa cells. Endocrinology 137:967–74.[Abstract]
D'Ascenzo S, Giusti I, Millimaggi D, et al. (2004) Intrafollicular expression of matrix metalloproteinases and their inhibitors in normally ovulating women compared with patients undergoing in vitro fertilization treatment. Eur J Endocrinol 151:87–91.[Abstract]
Davis JS, Weakland LL, West LA, et al. (1986) Luteinizing hormone stimulates the formation of inositol trisphosphate and cyclic AMP in rat granulosa cells. Evidence for phospholipase C generated second messengers in the action of luteinizing hormone. Biochem J 238:597–604.[Web of Science][Medline]
Davis BJ, Lennard DE, Lee CA, et al. (1999) Anovulation in cyclooxygenase-2-deficient mice is restored by prostaglandin E2 and interleukin-1beta. Endocrinology 140:2685–95.
De La Fuente R and Eppig JJ. (2001) Transcriptional activity of the mouse oocyte genome: companion granulosa cells modulate transcription and chromatin remodeling. Dev Biol 229:224–36.[CrossRef][Web of Science][Medline]
De La Fuente R, O'Brien MJ, Eppig JJ. (1999) Epidermal growth factor enhances preimplantation developmental competence of maturing mouse oocytes. Hum Reprod 14:3060–8.
De La Fuente R, Viveiros MM, Burns KH, et al. (2004) Major chromatin remodeling in the germinal vesicle (GV) of mammalian oocytes is dispensable for global transcriptional silencing but required for centromeric heterochromatin function. Dev Biol 275:447–58.[CrossRef][Web of Science][Medline]
Dekel N and Sherizly I. (1985) Epidermal growth factor induces maturation of rat follicle-enclosed oocytes. Endocrinology 116:406–9.
Dekel N, Lawrence TS, Gilula NB, et al. (1981) Modulation of cell-to-cell communication in the cumulus-oocyte complex and the regulation of oocyte maturation by LH. Dev Biol 86:356–62.[CrossRef][Web of Science][Medline]
Dewi DA, Abayasekara DRE, Wheeler-Jones CPD. (2002) Requirement for ERK1/2 activation in the regulation of progesterone production in human granulosa-lutein cells is stimulus specific. Endocrinology 143:877–88.
Di Pasquale E, Beck-Peccoz P, Persani L. (2004) Hypergonadotropic ovarian failure associated with an inherited mutation of human bone morphogenetic protein-15 (BMP15) gene. Am J Hum Genet 75:106–11.[CrossRef][Web of Science][Medline]
Dinchuk JE, Car BD, Focht RJ, et al. (1995) Renal abnormalities and an altered inflammatory response in mice lacking cyclooxygenase II. Nature 378:406–9.[CrossRef][Medline]
Dong J, Albertini DF, Nishimori K, et al. (1996) Growth differentiation factor-9 is required during early ovarian folliculogenesis. Nature 383:531–5.[CrossRef][Medline]
Downs SM and Longo FJ. (1983) Prostaglandins and preovulatory follicular maturation in mice. J Exp Zoolog 228:99–108.[CrossRef]
Downs SM, Daniel SA, Eppig JJ. (1998) Induction of maturation in cumulus cell-enclosed mouse oocytes by follicle-stimulating hormone and epidermal growth factor: evidence for a positive stimulus of somatic cell origin. J Exp Zoolog 245:86–96.
Doyle KMH, Russell DL, Sriraman V, et al. (2004) Coordinate transcription of the adamts-1 gene by luteinizing hormone and progesterone receptor. Mol Endocrinol 18:2463–78.
Dragovic RA, Ritter LJ, Schulz SJ, et al. (2005) Role of oocyte-secreted growth differentiation factor 9 in the regulation of mouse cumulus expansion. Endocrinology 146:2798–806.[CrossRef][Web of Science][Medline]
Drahorad J, Tesarik J, Cechova D, et al. (1991) Proteins and glycosaminoglycans in the intercellular matrix of the human cumulus-oophorus and their effect on conversion of proacrosin to acrosin. J Reprod Fertil 93:253–62.
Duffy DM, Seachord CL, Dozier BL. (2005) An ovulatory gonadotropin stimulus increases cytosolic phospholipase a2 expression and activity in granulosa cells of primate periovulatory follicles. J Clin Endocrinol Metab 90:5858–65.
Eder J. (1997) Tumour necrosis factor [alpha] and interleukin 1 signalling: do MAPKK kinases connect it all? Trends Pharmacol Sci 18:319–22.[Medline]
Elvin JA, Yan C, Matzuk MM. (2000) Growth differentiation factor-9 stimulates progesterone synthesis in granulosa cells via a prostaglandin E2/EP2 receptor pathway. Proc Natl Acad Sci U S A 97:10288–93.
Elvin JA, Clark AT, Wang P, et al. (1999a) Paracrine actions of growth differentiation factor-9 in the mammalian ovary. Mol Endocrinol 13:1035–48.
Elvin JA, Yan C, Wang P, et al. (1999b) Molecular characterization of the follicle defects in the growth differentiation factor 9-deficient ovary. Mol Endocrinol 13:1018–34.
Eppig JJ. (1979a) FSH stimulates hyaluronic acid synthesis by oocyte-cumulus cell complexes from mouse preovulatory follicles. Nature 281:483–4.[CrossRef][Medline]
Eppig JJ. (1979b) Gonadotropin stimulation of the expansion of cumulus oophori isolated from mice: general conditions for expansion in vitro. J Exp Zool 208:111–20.[CrossRef][Web of Science][Medline]
Eppig JJ. (1980) Regulation of cumulus oophorus expansion by gonadotropins in vivo and in vitro. Biol Reprod 23:545–52.[Abstract]
Eppig JJ. (1981) Prostaglandin E2 stimulates cumulus expansion and hyaluronic acid synthesis by cumuli oophori isolated from mice. Biol Reprod 25:191–5.[Abstract]
Eppig J, Pendola F, Wigglesworth K. (1998) Mouse oocytes suppress cAMP-induced expression of LH receptor mRNA by granulosa cells in vitro. Mol Reprod Dev 49:327–32.[CrossRef][Web of Science][Medline]
Eppig JJ, Wigglesworth K, Chesnel F. (1993) Secretion of cumulus expansion enabling factor by mouse oocytes: relationship to oocyte growth and competence to resume meiosis. Dev Biol 158:400–9.[CrossRef][Web of Science][Medline]
Eppig JJ, Wigglesworth K, Pendola FL. (2002) The mammalian oocyte orchestrates the rate of ovarian follicular development. Proc Natl Acad Sci U S A 99:2890–4.
Eppig J, Ward-Bailey P, Coleman D. (1985) Hypoxanthine and adenosine in murine ovarian follicular fluid: concentrations and activity in maintaining oocyte meiotic arrest. Biol Reprod 33:1041–9.[Abstract]
Eppig J, Wigglesworth K, Pendola F, et al. (1997) Murine oocytes suppress expression of luteinizing hormone receptor messenger ribonucleic acid by granulosa cells. Biol Reprod 56:976–84.[Abstract]
Eppig JJ, Pendola FL, Wigglesworth K, et al. (2005) Mouse oocytes regulate metabolic cooperativity between granulosa cells and oocytes: amino acid transport. Biol Reprod 73:351–7.
Erickson GF and Shimasaki S. (2000) The role of the oocyte in folliculogenesis. Trends Endocrinol Metab 11:193–8.[CrossRef][Web of Science][Medline]
Eriksen GV, Carlstedt I, Morgelin M, et al. (1999) Isolation and characterization of proteoglycans from human follicular fluid. Biochem J 340:613–20.[CrossRef][Web of Science][Medline]
Espey LL. (1970) Effect of various substances on tensile strength of sow ovarian follicles. Am J Physiol 219:230–3.
Espey LL. (1994) Current status of the hypothesis that mammalian ovulation is comparable to an inflammatory reaction. Biol Reprod 50:233–8.[Abstract]
Espey LL. (1998) An Overview of 37 Years of Research on Ovulation(Springer, New York).
Espey LL and Richards JS. (2002) Temporal and spatial patterns of ovarian gene transcription following an ovulatory dose of gonadotropin in the rat. Biol Reprod 67:1662–70.
Espey LL and Rondell P. (1968) Collagenolytic activity in the rabbit and sow Graafian follicle during ovulation. Am J Physiol 214:326–9.
Espey L, Tanaka N, Woodard D, et al. (1989) Decrease in ovarian platelet-activating factor during ovulation in the gonadotropin-primed immature rat. Biol Reprod 41:104–10.[Abstract]
Espey LL, Yoshioka S, Russell DL, et al. (2000) Ovarian expression of a disintegrin and metalloproteinase with thrombospondin motifs during ovulation in the gonadotropin-primed immature rat. Biol Reprod 62:1090–5.
Familiari G, Verlengia C, Nottola SA, et al. (1996) Heterogeneous distribution of fibronectin, tenascin-C, and laminin immunoreactive material in the cumulus-corona cells surrounding mature human oocytes from IVF-ET protocols–evidence that they are composed of different subpopulations: an immunohistochemical study using scanning confocal laser and fluorescence microscopy. Mol Reprod Dev 43:392–402.[CrossRef][Web of Science][Medline]
Familiari G, Heyn R, Relucenti M, et al. (2006) Ultrastructural dynamics of human reproduction, from ovulation to fertilization and early embryo development. Int Rev Cytol 249:53–141.[Web of Science][Medline]
Fatehi AN, Roelen BA, Colenbrander B, et al. (2005) Presence of cumulus cells during in vitro fertilization protects the bovine oocyte against oxidative stress and improves first cleavage but does not affect further development. Zygote 13:177–85.[CrossRef][Web of Science][Medline]
Ferrara N, Chen H, Davis-Smyth T, et al. (1998) Vascular endothelial growth factor is essential for corpus luteum angiogenesis. Nat Med 4:336–40.[CrossRef][Web of Science][Medline]
Foong SC, Abbott DH, Zschunke MA, et al. (2006) Follicle Luteinization in Hyperandrogenic Follicles of Polycystic Ovary Syndrome (PCOS) Patients Undergoing Gonadotropin Therapy For In Vitro Fertilization (IVF). J Clin Endocrinol Metab jc.2005–2142.
Foote RH. (1987) In vitro fertilization and embryo transfer in domestic animals: applications in animals and implications for humans. J In Vitro Fertilization Embryo Transfer 4:73–88.[CrossRef]
Fraser HM, Dickson SE, Lunn SF, et al. (2000) Suppression of luteal angiogenesis in the primate after neutralization of vascular endothelial growth factor. Endocrinology 141:995–1000.
Freimann S, Ben-Ami I, Dantes A, et al. (2004) EGF-like factor epiregulin and amphiregulin expression is regulated by gonadotropins/cAMP in human ovarian follicular cells. Biochem Biophys Res Commun 324:829–34.[CrossRef][Web of Science][Medline]
Freimann S, Ben-Ami I, Dantes A, et al. (2005) Differential expression of genes coding for EGF-like factors and ADAMTS1 following gonadotropin stimulation in normal and transformed human granulosa cells. Biochem Biophys Res Commun 333:935–43.[CrossRef][Web of Science][Medline]
Fulop C, Kamath RV, Li Y, et al. (1997) Coding sequence, exon-intron structure and chromosomal localization of murine TNF-stimulated gene 6 that is specifically expressed by expanding cumulus cell-oocyte complexes. Gene 202:95–102.[CrossRef][Web of Science][Medline]
Fulop C, Szanto S, Mukhopadhyay D, et al. (2003) Impaired cumulus mucification and female sterility in tumor necrosis factor-induced protein-6 deficient mice. Development 130:2253–61.
Galloway SM, McNatty KP, Cambridge LM, et al. (2000) Mutations in an oocyte-derived growth factor gene (BMP15) cause increased ovulation rate and infertility in a dosage-sensitive manner. Nature Gen 25:279–83.[CrossRef][Web of Science][Medline]
Galway AB, Lapolt PS, Tsafriri A, et al. (1990) Recombinant follicle-stimulating hormone induces ovulation and tissue plasminogen activator expression in hypophysectomized rats. Endocrinology 127:3023–8.
Gardner DK, Pawelczynski M, Trounson AO. (1996) Nutrient uptake and utilization can be used to select viable day 7 bovine blastocysts after cryopreservation. Mol Reprod Dev 44:472–5.[CrossRef][Web of Science][Medline]
Gaytan F, Bellido C, Gaytan M, et al. (2003) Differential effects of RU486 and indomethacin on follicle rupture during the ovulatory process in the rat. Biol Reprod 69:99–105.
Gilchrist RB, Ritter LJ, Armstrong DT. (2004) Oocyte-somatic cell interactions during follicle development in mammals. Anim Reprod Sci 82–83:431–46.
Gomez R, Simon C, Remohi J, et al. (2003) Administration of moderate and high doses of gonadotropins to female rats increases ovarian vascular endothelial growth factor (vegf) and vegf receptor-2 expression that is associated to vascular hyperpermeability. Biol Reprod 68:2164–71.
Gonzalez-Robayna IJ, Alliston TN, Buse P, et al. (1999) Functional and subcellular changes in the A-kinase-signaling pathway: relation to aromatase and Sgk expression during the transition of granulosa cells to luteal cells. Mol Endocrinol 13:1318–37.
Gonzalez-Robayna IJ and Falender AE, et al. (2000) Follicle-stimulating hormone (FSH) stimulates phosphorylation and activation of protein kinase B (PKB/Akt) and serum and glucocorticoid-induced kinase (Sgk): evidence for A kinase-independent signaling by FSH in granulosa cells. Mol Endocrinol 14:1283–300.
Gore-Langton RE and Armstrong DT. (1994) Follicular steroidogenesis and its control. In Knobil E and Neill JD (Eds.). The Physiology of Reproduction Second edition (Raven Press Ltd., New York) 1: pp. 571–627.
Gottsch M, Van Kirk E, Murdoch W. (2002) Role of matrix metalloproteinase 2 in the ovulatory folliculo-luteal transition of ewes. Reproduction 124:347–52.[Abstract]
Goud PT, Goud AP, Qian C, et al. (1998) In-vitro maturation of human germinal vesicle stage oocytes: role of cumulus cells and epidermal growth factor in the culture medium. Hum Reprod 13:1638–44.
Goudet G, Belin F, Bezard J, et al. (1999) Intrafollicular content of luteinizing hormone receptor, {alpha}-inhibin, and aromatase in relation to follicular growth, estrous cycle stage, and oocyte competence for in vitro maturation in the mare. Biol Reprod 60:1120–7.
Granot I and Dekel N. (1994) Phosphorylation and expression of connexin-43 ovarian gap junction protein are regulated by luteinizing hormone. J Biol Chem 269:30502–9.
Gregory L. (1998) Ovarian markers of implantation potential in assisted reproduction. Hum Reprod 13:117–32.
Greve T and Callesen H. (2001) Rendez-vous in the oviduct: implications for superovulation and embryo transfer. Reprod Nutr Dev 41:451–9.[CrossRef][Web of Science][Medline]
Grupen CG, Nagashima H, Nottle MB. (1997) Role of epidermal growth factor and insulin-like growth factor-I on porcine oocyte maturation and embryonic development in vitro. Reprod Fertil Dev 9:571–5.[CrossRef][Medline]
Gudermann T, Birnbaumer M, Birnbaumer L. (1992) Evidence for dual coupling of the murine luteinizing hormone receptor to adenylyl cyclase and phosphoinositide breakdown and Ca2+ mobilization. Studies with the cloned murine luteinizing hormone receptor expressed in L cells. J Biol Chem 267:4479–88.
Gueripel X, Benahmed M, Gougeon A. (2004) Sequential gonadotropin treatment of immature mice leads to amplification of transforming growth factor {beta} action, via upregulation of receptor-type 1, Smad 2 and 4, and downregulation of Smad 6.''. Biol Reprod 70:640–8.
Gui L-M and Joyce IM. (2005) RNA interference evidence that growth differentiation factor-9 mediates oocyte regulation of cumulus expansion in mice. Biol Reprod 72:195–9.
Hagglund AC, Ny A, Leonardsson G, et al. (1999) Regulation and localization of matrix metalloproteinases and tissue inhibitors of metalloproteinases in the mouse ovary during gonadotropin-induced ovulation. Endocrinology 140:4351–8.
Hanrahan JP, Gregan SM, Mulsant P, et al. (2004) Mutations in the genes for oocyte-derived growth factors GDF9 and BMP15 are associated with both increased ovulation rate and sterility in cambridge and belclare sheep (ovis aries). Biol Reprod 70:900–9.
Harper JK. (1994) Gamete and zygote transport. In Knobil E and Neill JD (Eds.). The Physiology of Reproduction(Raven Press, New York) pp. 123–85.
Harris TE, Squires PE, Michael AE, et al. (2001) Human granulosa-lutein cells express functional EP1 and EP2 prostaglandin receptors. Biochem Biophys Res Commun 285:1089–94.[CrossRef][Web of Science][Medline]
Hasegawa J, Yanaihara A, Iwasaki S, et al. (2005) Reduction of progesterone receptor expression in human cumulus cells at the time of oocyte collection during IVF is associated with good embryo quality. Hum Reprod 20:2194–200.
Hashimoto O, Moore RK, Shimasaki S. (2005) Posttranslational processing of mouse and human BMP-15: Potential implication in the determination of ovulation quota. Proc Natl Acad Sci U S A 102:5426–31.
Hazzard TM and Stouffer RL. (2000) Angiogenesis in ovarian follicular and luteal development. Baillieres Best Pract Res Clin Obstet Gynaecol 14:883–900.[CrossRef][Medline]
Hazzard TM, Xu F, Stouffer RL. (2002) Injection of soluble vascular endothelial growth factor receptor 1 into the preovulatory follicle disrupts ovulation and subsequent luteal function in rhesus monkeys. Biol Reprod 67:1305–12.
Hellberg P, Thomsen P, Janson P, et al. (1991) Leukocyte supplementation increases the luteinizing hormone-induced ovulation rate in the in vitro-perfused rat ovary. Biol Reprod 44:791–7.[Abstract]
Hess KA, Chen L, Larsen WJ. (1999) Inter-alpha-inhibitor binding to hyaluronan in the cumulus extracellular matrix is required for optimal ovulation and development of mouse oocytes. Biol Reprod 61:436–43.
Hickey TE, Alvino HJ, Gilchrist RB, et al. (2005) Granulosa cells from polycystic ovaries have increased expression of androgen receptor regulated kallikrein 3 but normal expression of androgen receptor and and two androgen receptor coactivators. Proceedings of eighty-seventh Annual Meeting of the Endocrine SoceitySan Diego, CA, USA pp. 414 Abstract P2–242.
Hillensjo T, Magnusson C, Svensson U, et al. (1981) Effect of luteinizing hormone and follicle-stimulating hormone on progesterone synthesis by cultured rat cumulus cells. Endocrinology 108:1920–4.
Hizaki H, Segi E, Sugimoto Y, et al. (1999) Abortive expansion of the cumulus and impaired fertility in mice lacking the prostaglandin E receptor subtype EP(2). Proc Natl Acad Sci U S A 96:10501–6.
Hoff JD, Quigley ME, Yen SS. (1983) Hormonal dynamics at midcycle: a reevaluation. J Clin Endocrinol Metab 57:792–6.
Holland A, Findlay J, Clements J. (2001) Kallikrein gene expression in the gonadotrophin-stimulated rat ovary. J Endocrinol 170:243–50.[Abstract]
Holmbeck K, Bianco P, Caterina J, et al. (1999) MT1-MMP-deficient mice develop dwarfism, osteopenia, arthritis, and connective tissue disease due to inadequate collagen turnover. Cell 99:81–92.[CrossRef][Web of Science][Medline]
Hong SJ, Chiu PC, Lee KF, et al. (2004) Establishment of a capillary-cumulus model to study the selection of sperm for fertilization by the cumulus oophorus. Hum Reprod 19:1562–9.
Hu Y-C, Wang P-H, Yeh S, et al. (2004) Subfertility and defective folliculogenesis in female mice lacking androgen receptor. Proc Natl Acad Sci U S A 101:11209–14.
Hussein TS, Froiland DA, Amato F, et al. (2005) Oocytes prevent cumulus cell apoptosis by maintaining a morphogenic paracrine gradient of bone morphogenetic proteins. J Cell Sci jcs.02644.
Ikeda S, Nishikimi A, Ichihara-Tanaka K, et al. (2000) cDNA cloning of bovine midkine and production of the recombinant protein, which affects in vitro maturation of bovine oocytes. Mol Reprod Dev 57:99–107.[CrossRef][Web of Science][Medline]
Itoh T, Ikeda T, Gomi H, et al. (1997) Unaltered secretion of beta -amyloid precursor protein in gelatinase A (matrix metalloproteinase 2)-deficient mice. J Biol Chem 272:22389–92.
Jablonka-Shariff A and Olson LM. (1997) Hormonal regulation of nitric oxide synthases and their cell-specific expression during follicular development in the rat ovary. Endocrinology 138:460–8.
Jaffe RC, Donnelly KM, Mavrogianis PA, et al. (1989) Molecular cloning and characterization of a progesterone-dependent cat endometrial secretory protein complementary deoxyribonucleic acid. Mol Endocrinol 3:1807–14.
Jamnongjit M, Gill A, Hammes SR. (2005) Epidermal growth factor receptor signaling is required for normal ovarian steroidogenesis and oocyte maturation. Proc Natl Acad Sci U S A 102:16257–62.
Jansen E, Laven JSE, Dommerholt HBR, et al. (2004) Abnormal gene expression profiles in human ovaries from polycystic ovary syndrome patients. Mol Endocrinol 18:3050–63.
Jasper M and Norman RJ. (1995) Immunoactive interleukin-1 beta and tumour necrosis factor-alpha in thecal, stromal and granulosa cell cultures from normal and polycystic ovaries. Hum Reprod 10:1352–4.
Jensen JT, Schwinof KM, Zelinski-Wooten MB, et al. (2002) Phosphodiesterase 3 inhibitors selectively block the spontaneous resumption of meiosis by macaque oocytes in vitro. Hum Reprod 17:2079–84.
Jensen JT, Zelinski-Wooten MB, Schwinof KM, et al. (2005) The phosphodiesterase 3 inhibitor ORG 9935 inhibits oocyte maturation during gonadotropin-stimulated ovarian cycles in rhesus macaques. Contraception 71:68–73.[CrossRef][Web of Science][Medline]
Jessen T and Odum L. (2003) Role of tumour necrosis factor stimulated gene 6 (TSG-6) in the coupling of inter-alpha-trypsin inhibitor to hyaluronan in human follicular fluid. Reproduction 125:27–31.[Abstract]
Jin S-LC, Richard FJ, Kuo W-P, et al. (1999) Impaired growth and fertility of cAMP-specific phosphodiesterase PDE4D-deficient mice. Proc Natl Acad Sci U S A 96:11998–2003.
Jo M and Curry TE. (2004) Regulation of matrix metalloproteinase-19 messenger RNA expression in the rat ovary. Biol Reprod 71:1796–806.
Jo M, Thomas LE, Wheeler SE, et al. (2004) Membrane type 1-matrix metalloproteinase (MMP)-associated MMP-2 activation increases in the rat ovary in response to an ovulatory dose of human chorionic gonadotropin. Biol Reprod 70:1024–32.
Joyce IM, Pendola FL, O'Brien M, et al. (2001) Regulation of prostaglandin-endoperoxide synthase 2 messenger ribonucleic acid expression in mouse granulosa cells during ovulation. Endocrinology 142:3187–97.
Juengel JL, Hudson NL, Heath DA, et al. (2002) Growth differentiation factor 9 and bone morphogenetic protein 15 are essential for ovarian follicular development in sheep. Biol Reprod 67:1777–89.
Juengel JL, Hudson NL, Whiting L, et al. (2004) Effects of immunization against bone morphogenetic protein 15 and growth differentiation factor 9 on ovulation rate, fertilization, and pregnancy in ewes. Biol Reprod 70:557–61.
Kaivo-Oja N, Mottershead DG, Mazerbourg S, et al. (2005) Adenoviral gene transfer allows smad-responsive gene promoter analyses and delineation of type i receptor usage of transforming growth factor-{beta} family ligands in cultured human granulosa luteal cells. J Clin Endocrinol Metab 90:271–8.
Kalma Y, Granot I, Galiani D, et al. (2004) Luteinizing hormone-induced connexin 43 down-regulation: inhibition of translation. Endocrinology 145:1617–24.[CrossRef][Web of Science][Medline]
Kansra S, Stoll SW, Johnson JL, et al. (2004) Autocrine extracellular signal-regulated kinase (ERK) activation in normal human keratinocytes: metalloproteinase-mediated release of amphiregulin triggers signaling from ErbB1 to ERK. Mol Biol Cell 15:4299–309.
Karagouni EE, Chryssikopoulos A, Mantzavinos T, et al. (1998) Interleukin-1[beta] and interleukin-1[alpha] may affect the implantation rate of patients undergoing in vitro fertilization-embryo transfer. Fertil Steril 70:553–9.[CrossRef][Web of Science][Medline]
Kawamura K, Kumagai J, Sudo S, et al. (2004) Paracrine regulation of mammalian oocyte maturation and male germ cell survival. Proc Natl Acad Sci U S A 101:7323–8.
Kawamura K, Kawamura N, Mulders SM, et al. (2005) Ovarian brain-derived neurotrophic factor (BDNF) promotes the development of oocytes into preimplantation embryos. Proc Natl Acad Sci U S A 102:9206–11.
Kehrl JH and Sinnarajah S. (2002) RGS2: a multifunctional regulator of G-protein signaling. Int J Biochem Cell Biol 34:432–8.[CrossRef][Web of Science][Medline]
Kikukawa Y, Ishikawa M, Sengoku K, et al. (1991) The effect of platelet activating factor on ovulation. Prostaglandins 42:95–104.[CrossRef][Web of Science][Medline]
Kobayashi H, Sun GW, Hirashima Y, et al. (1999) Identification of link protein during follicle development and cumulus cell cultures in rats. Endocrinology 140:3835–42.
Kobayashi H, Suzuki M, Tanaka Y, et al. (2001) Suppression of urokinase expression and invasiveness by urinary trypsin inhibitor is mediated through inhibition of protein kinase C- and MEK/ERK/c-Jun-dependent signaling pathways. J Biol Chem 276:2015–22.
Kokia E, Hurwitz A, Ben-Shlomo I, Adashi E, Yanagishita M. (1993) Receptor-mediated stimulatory effect of IL-1 beta on hyaluronic acid and proteoglycan biosynthesis by cultured rat ovarian cells: role for heterologous cell-cell interactions. Endocrinology 133:2391–4.
Kol S, Ruutiainen-Altman K, Scherzer WJ, et al. (1999) The rat intraovarian interleukin (IL)-1 system: cellular localization, cyclic variation and hormonal regulation of IL-1beta and of the type I and type II IL-1 receptors. Mol Cell Endocrinol 129:115–28.
Lam X, Gieseke C, Knoll M, et al. (2000) Assay and importance of adhesive interaction between hamster (Mesocricetus auratus) oocyte-cumulus complexes and the oviductal epithelium. Biol Reprod 62:579–88.
x PS, Leung K, Ishimaru R, et al. (2003) Roles of cyclic GMP in modulating ovarian functions. Reprod Biomed Online 16:15–23.
Larsen WJ, Chen L, Powers R, et al. (1996) Cumulus expansion initiates physical and developmental autonomy of the oocyte. Zygote 4:335–41.[Web of Science][Medline]
Lash GE and Legge M. (2001) Localization and distribution of platelet activating factor receptors in the mouse ovary and oviduct during the estrous cycle and early pregnancy. Am J Reprod Immunol 45:123–7.[Medline]
Latham KE, Wigglesworth K, McMenamin M, et al. (2004) Stage-dependent effects of oocytes and growth differentiation factor 9 on mouse granulosa cell development: advance programming and subsequent control of the transition from preantral secondary follicles to early antral tertiary follicles. Biol Reprod 70:1253–62.
Laufer N, Tarlatzis BC, Naftolin F. (1984) In vitro fertilization: state of the art. Semin Reprod Endocrinol 2:197–219.[Web of Science]
Lawrence TS, Dekel N, Beers WH. (1980) Binding of human chorionic gonadotropin by rat cumuli oophori and granulosa cells: a comparative study. Endocrinology 106:1114–8.
Lee SL, Sadovsky Y, Swirnoff AH, et al. (1996) Luteinizing hormone deficiency and female infertility in mice lacking the transcription factor NGFI-A (Egr-1). Science 273:1219–21.[Abstract]
Lee D, Pearsall RS, Das S, et al. (2004) Epiregulin is not essential for development of intestinal tumors but is required for protection from intestinal damage. Mol Cell Biol 24:8907–16.
Li L, He S, Sun JM, et al. (2004) Gene regulation by Sp1 and Sp3. Biochem Cell Biol 82:460–71.[CrossRef][Web of Science][Medline]
Lim H, Paria BC, Das SK, et al. (1997) Multiple female reproductive failures in cyclooxygenase 2-deficient mice. Cell 91:197–208.[CrossRef][Web of Science][Medline]
Lorenzo PL, Liu IK, Illera JC, et al. (2001) Influence of epidermal growth factor on mammalian oocyte maturation via tyrosine-kinase pathway. J Physiol Biochem 57:15–22.[Medline]
Loutradis D, Bletsa R, Aravantinos L, et al. (1991) Preovulatory effects of the progesterone antagonist mifepristone (RU486) in mice. Hum Reprod 6:1238–40.
Luciano AM, Lodde V, Beretta MS, et al. (2005) Developmental capability of denuded bovine oocyte in a Co-culture system with intact cumulus-oocyte complexes: role of cumulus cells, cyclic adenosine 3,5-monophosphate, and glutathione. Mol Reprod Dev 71:389–97.[CrossRef][Web of Science][Medline]
Luetteke N, Qiu T, Fenton S, et al. (1999) Targeted inactivation of the EGF and amphiregulin genes reveals distinct roles for EGF receptor ligands in mouse mammary gland development. Development 126:2739–50.[Abstract]
Lydon JP, DeMayo FJ, Conneely OM, et al. (1996) Reproductive phenotpes of the progesterone receptor null mutant mouse. J Steroid Biochem Mol Biol 56:67–77.[CrossRef][Web of Science][Medline]
MacLean JA II, Chen MA, Wayne CM, et al. (2005) Rhox: a new homeobox gene cluster. Cell 120:369–82.[CrossRef][Web of Science][Medline]
Madan P, Bridges PJ, Komar CM, et al. (2003) Expression of messenger RNA for ADAMTS subtypes changes in the periovulatory follicle after the gonadotropin surge and during luteal development and regression in cattle. Biol Reprod 69:1506–14.
Mahi-Brown CA and Yanagimachi R. (1983) Parameters influencing ovum pick-up by oviductal fimbria in the golden hamster. Gamete Res 8:1–10.[Medline]
Maizels ET, Mukherjee A, Sithanandam G, et al. (2001) Developmental regulation of mitogen-activated protein kinase-activated kinases-2 and -3 (MAPKAPK-2/-3) in vivo during corpus luteum formation in the rat. Mol Endocrinol 15:716–33.
Marin Bivens CL, Grondahl C, Murray A, et al. (2004a) Meiosis-activating sterol promotes the metaphase i to metaphase ii transition and preimplantation developmental competence of mouse oocytes maturing in vitro. Biol Reprod 70:1458–64.
Marin Bivens CL, Lindenthal B, O'Brien MJ, et al. (2004b) A synthetic analogue of meiosis-activating sterol (FF-MAS) is a potent agonist promoting meiotic maturation and preimplantation development of mouse oocytes maturing in vitro. Hum Reprod 19:2340–4.
Marsh JM. (1976) The role of cyclic AMP in gonadal steroidogenesis. Biol Reprod 14:130–53.[CrossRef][Web of Science][Medline]
Matsuura T, Sugimura M, Iwaki T, et al. (2002) Anti-macrophage inhibitory factor antibody inhibits PMSG-hCG-induced follicular growth and ovulation in mice. J Assist Reprod Genet 19:591–5.[CrossRef][Web of Science][Medline]
Mazerbourg S, Klein C, Roh J, et al. (2004) Growth differentiation factor-9 signaling is mediated by the type i receptor, activin receptor-like kinase 5. Mol Endocrinol 18:653–65.
McKenzie LJ, Pangas SA, Carson SA, et al. (2004) Human cumulus granulosa cell gene expression: a predictor of fertilization and embryo selection in women undergoing IVF. Hum Reprod 19:2869–74.
McNatty KP, Juengel JL, Reader KL, et al. (2005) Bone morphogenetic protein 15 and growth differentiation factor 9 co-operate to regulate granulosa cell function in ruminants. Reproduction 129:481–7.
Mendoza C, Cremades N, Ruiz-Requena E, et al. (1999) Relationship between fertilization results after intracytoplasmic sperm injection, and intrafollicular steroid, pituitary hormone and cytokine concentrations. Hum Reprod 14:628–35.
Mendoza C, Ruiz-Requena E, Ortega E, et al. (2002) Follicular fluid markers of oocyte developmental potential. Hum Reprod 17:1017–22.
Mikuni M, Pall M, Peterson CM, et al. (1998) The selective prostaglandin endoperoxide synthase-2 inhibitor, NS-398, reduces prostaglandin production and ovulation in vivo and in vitro in the rat. Biol Reprod 59:1077–83.
Minge CE, Ryan NK, Van Der Hoek KH, et al. (2005) Troglitazone regulates peroxisome proliferator-activated receptors and inducible nitric oxide synthase in murine ovarian macrophages. Biol Reprod 105.043729.
Mittaz L, Russell DL, Wilson T, et al. (2004) Adamts-1 is essential for the development and function of the urogenital system. Biol Reprod 70:1096–105.
Mjaatvedt CH, Yamamura H, Capehart AA, et al. (1998) The Cspg2 gene, disrupted in the hdf mutant, is required for right cardiac chamber and endocardial cushion formation. Dev Biol 202:56–66.[CrossRef][Web of Science][Medline]
Montgomery GW, Zhao ZZ, Marsh AJ, et al. (2004) A deletion mutation in GDF9 in sisters with spontaneous DZ twins. Twin Res 7:548–55.[CrossRef][Web of Science][Medline]
Moore RK, Erickson GF, Shimasaki S. (2004) Are BMP-15 and GDF-9 primary determinants of ovulation quota in mammals? Trends Endocrinol Metab 15:356–61.[CrossRef][Web of Science][Medline]
Morris JK and Richards JS. (1995) Luteinizing hormone induces prostaglandin endoperoxide synthase-2 and luteinization in vitro by A-kinase and C-kinase pathways. Endocrinology 136:1549–58.[Abstract]
Morris JK and Richards JS. (1996) An E-box region within the prostaglandin endoperoxide synthase-2 (PGS- 2) promoter is required for transcription in rat ovarian granulosa cells. J Biol Chem 271:16633–43.
Motta PM, Nottola SA, Pereda J, et al. (1995) Ultrastructure of human cumulus oophorus: a transmission electron microscopic study on oviductal oocytes and fertilized eggs. Hum Reprod 10:2361–7.
Mukherjee A, Park-Sarge O, Mayo K. (1996) Gonadotropins induce rapid phosphorylation of the 3',5'-cyclic adenosine monophosphate response element binding protein in ovarian granulosa cells. Endocrinology 137:3234–45.[Abstract]
Mukhopadhyay D, Hascall VC, Day AJ, et al. (2001) Two distinct populations of tumor necrosis factor-stimulated gene-6 protein in the extracellular matrix of expanded mouse cumulus cell- oocyte complexes. Arch Biochem Biophys 394:173–81.[CrossRef][Web of Science][Medline]
Mulac-Jericevic B, Mullinax RA, DeMayo FJ, et al. (2000) Subgroup of reproductive functions of progesterone mediated by progesterone receptor-B isoform. Science 289:1751–4.
Murdoch WJ, Wilken C, Young DA. (1999) Sequence of apoptosis and inflammatory necrosis within the formative ovulatory site of sheep follicles. J Reprod Fertil 117:325–9.
Murdoch WJ, Peterson TA, Van Kirk EA, et al. (1986) Interactive roles of progesterone, prostaglandins, and collagenase in the ovulatory mechanism of the ewe. Biol Reprod 35:1187–94.[Abstract]
Nakamura K, Fujiwara H, Higuchi T, et al. (1997) Integrin [alpha]6 is involved in follicular growth in mice. Biochem Biophys Res Commun 235:524–8.[CrossRef][Web of Science][Medline]
Natraj U and Richards JS. (1993) Hormonal regulation, localization, and functional activity of the progesterone receptor in granulosa cells of rat preovulatory follicles. Endocrinology 133:761–9.
Nef S and Parada LF. (1999) Cryptorchidism in mice mutant for Insl3. 22:295–299 Nature Gen.
Neulen J, Yan Z, Raczek S, et al. (1995) Human chorionic gonadotropin-dependent expression of vascular endothelial growth factor/vascular permeability factor in human granulosa cells: importance in ovarian hyperstimulation syndrome. J Clin Endocrinol Metab 80:1967–71.[Abstract]
Nogueira D, Ron-El R, Friedler S, et al. (2006) Meiotic arrest in vitro by phosphodiesterase 3-inhibitor enhances maturation capacity of human oocytes and allows subsequent embryonic development. Biol Reprod 74:177–84.
Nuttinck F, Reinaud P, Tricoire H, et al. (2002) Cyclooxygenase-2 is expressed by cumulus cells during oocyte maturation in cattle. Mol Reprod Dev 61:93–101.[CrossRef][Web of Science][Medline]
Ny A, Leonardsson G, Hagglund AC, et al. (1999) Ovulation in plasminogen-deficient mice. Endocrinology 140:5030–5.
Ochsner SA, Day AJ, Rugg MS, et al. (2003a) Disrupted function of tumor necrosis factor-{alpha}-stimulated gene 6 blocks cumulus cell-oocyte complex expansion. Endocrinology 144:4376–84.
Ochsner SA, Russell DL, Day AJ, et al. (2003b) Decreased expression of tumor necrosis factor-alpha-stimulated gene 6 in cumulus cells of the cyclooxygenase-2 and EP2 null mice. Endocrinology 144:1008–19.
O'Donnell JB, Hill JL, Gross DJ. (2004) Epidermal growth factor activates cytosolic [Ca2+] elevations and subsequent membrane permeabilization in mouse cumulus-oocyte complexes. Reproduction 127:207–20.
Odor DL and Blandau RJ. (1973) Egg transport over the fimbrial surface of the rabbit oviduct under experimental conditions. Fertil Steril 24:292–300.[Web of Science][Medline]
Ogiwara K, Takano N, Shinohara M, et al. (2005) Gelatinase A and membrane-type matrix metalloproteinases 1 and 2 are responsible for follicle rupture during ovulation in the medaka. Proc Natl Acad Sci U S A 102:8442–7.
Ohnishi J, Ohnishi E, Shibuya H, et al. (2005) Functions for proteinases in the ovulatory process. Biochim Biophys Acta, Proteins Proteomics 1751:95–109.[CrossRef]
Ohta N, Saito H, Kuzumaki T, et al. (1999) Expression of CD44 in human cumulus and mural granulosa cells of individual patients in in-vitro fertilization programmes. Mol Hum Reprod 5:22–8.
Osman P and Dullaart J. (1976) Intraovarian release of eggs in the rat after indomethacin treatment at pro-oestrus. J Reprod Fertil 47:101–3.
Oxberry B and Greenwald G. (1982) An autoradiographic study of the binding of 125 I-labeled follicle- stimulating hormone, human chorionic gonadotropin and prolactin to the hamster ovary throughout the estrous cycle. Biol Reprod 27:505–16.[CrossRef][Web of Science][Medline]
Pall M, Friden BE, Brannstrom M. (2001) Induction of delayed follicular rupture in the human by the selective COX-2 inhibitor rofecoxib: a randomized double-blind study. Hum Reprod 16:1323–8.
Palotie A, Peltonen L, Foidart JM, et al. (1984) Immunohistochemical localization of basement membrane components and interstitial collagen types in preovulatory rat ovarian follicles. Coll Relat Res 4:279–87.[Web of Science][Medline]
Pangas SA, Jorgez CJ, Matzuk MM. (2004) Growth differentiation factor 9 regulates expression of the bone morphogenetic protein antagonist gremlin. J Biol Chem 279:32281–6.
Park J-Y, Richard F, Chun S-Y, et al. (2003) Phosphodiesterase regulation is critical for the differentiation and pattern of gene expression in granulosa cells of the ovarian follicle. Mol Endocrinol 17:1117–30.
Park J-Y, Su Y-Q, Ariga M, et al. (2004) EGF-like growth factors as mediators of LH action in the ovulatory follicle. Science 303:682–4.
Pelletier G, Labrie C, Labrie F. (2000) Localization of oestrogen receptor alpha, oestrogen receptor beta and androgen receptors in the rat reproductive organs. J Endocrinol 165:359–70.[Abstract]
Peng XR, Hsueh AJ, LaPolt PS, et al. (1991) Localization of luteinizing hormone receptor messenger ribonucleic acid expression in ovarian cell types during follicle development and ovulation. Endocrinology 129:3200–7.
Plancha C, Sanfins A, Rodrigues P, et al. (2005) Cell polarity during folliculogenesis and oogenesis. Reprod Biomed Online 10:478–84.[Web of Science][Medline]
Preis KA, Seidel G, Gardner DK. (2005) Metabolic markers of developmental competence for in vitro-matured mouse oocytes. Reproduction 130:475–83.
Procházka R, Kalab P, Nagyova E. (2003) Epidermal growth factor-receptor tyrosine kinase activity regulates expansion of porcine oocyte-cumulus cell complexes in vitro. Biol Reprod 68:797–803.
Procházka R, Srsen V, Nagyová E, et al. (2000) Developmental regulation of effect of epidermal growth factor on porcine oocyte-cumulus cell complexes: nuclear maturation, expansion, and F-actin remodeling. Mol Reprod Dev 56:63–73.[CrossRef][Web of Science][Medline]
Protin U, Schweighoffer T, Jochum W, et al. (1999) CD44-deficient mice develop normally with changes in subpopulations and recirculation of lymphocyte subsets. J Immunol 163:4917–23.
Reich R, Tsafriri A, Mechanic GL. (1985) The involvement of collagenolysis in ovulation in the rat. Endocrinology 116:522–7.
Relucenti M, Heyn R, Correr S, et al. (2005) Cumulus oophorus extracellular matrix in the human oocyte: a role for adhesive proteins. Ital J Anat Embryol 110:219–24.[Medline]
Revelli A, Pacchioni D, Cassoni P, et al. (1996) In situ hybridization study of messenger RNA for estrogen receptor and immunohistochemical detection of estrogen and progesterone receptors in the human ovary. Gynecol Endocrinol 10:177–86.[Web of Science][Medline]
Richard FJ, Tsafriri A, Conti M. (2001) Role of phosphodiesterase type 3A in rat oocyte maturation. Biol Reprod 65:1444–51.
Richards JS. (1994) Hormonal control of gene expression in the ovary. Endocr Rev 15:725–51.
Richards JS, Hernandez-Gonzalez I, Gonzalez-Robayna I, et al. (2005) Regulated expression of adamts family members in follicles and cumulus oocyte complexes: evidence for specific and redundant patterns during ovulation. Biol Reprod 72:1241–55.
Rimon E, Sasson R, Dantes A, et al. (2004) Gonadotropin-induced gene regulation in human granulosa cells obtained from IVF patients: modulation of genes coding for growth factors and their receptors and genes involved in cancer and other diseases. Int J Oncol 24:1325–38.[Web of Science][Medline]
Robert C, Gagne D, Lussier J, et al. (2003) Presence of LH receptor mRNA in granulosa cells as a potential marker of oocyte developmental competence and characterization of the bovine splicing isoforms. Reproduction 125:437–46.[Abstract]
Robker RL and Richards JS. (1998) Hormone-induced proliferation and differentiation of granulosa cells: a coordinated balance of the cell cycle regulators cyclin D2 and p27Kip1. Mol Endocrinol 12:924–40.
Robker RL, Russell DL, Espey LL, et al. (2000) Progesterone-regulated genes in the ovulation process: ADAMTS-1 and cathepsin L proteases. Proc Natl Acad Sci U S A 97:4689–94.
Rodgers RJ, Irving-Rodgers HF, Russell DL. (2003) Extracellular matrix of the developing ovarian follicle. Reproduction 126:415–24.[Abstract]
Rossi G, Macchiarelli G, Palmerini MG, et al. (2006) Meiotic spindle configuration is differentially influenced by FSH and epidermal growth factor during in vitro maturation of mouse oocytes. Hum Reprod 21:1765–70.
Roux PP and Blenis J. (2004) ERK and p38 MAPK-activated protein kinases: a family of protein kinases with diverse biological functions. Microbiol Mol Biol Rev 68:320–44.
Roy SK and Terada DM. (1999) Activities of glucose metabolic enzymes in human preantral follicles: in vitro modulation by follicle-stimulating hormone, luteinizing hormone, epidermal growth factor, insulin-like growth factor I, and transforming growth factor beta1. Biol Reprod 60:763–8.
Russell DL. Infertility in mice with null mutation of the EGR-1 transcription factor. Proceedings of the 35th anual conference of the Society for Reproductive Biology2004Sydney, Australia(CSIRO Publishing, Sydney) Abstract 279.
Russell DL, Salamonsen LA, Findlay JK. (1995) Immunization against the N-terminal peptide of the inhibin alpha 43- subunit (alpha N) disrupts tissue remodeling and the increase in matrix metalloproteinase-2 during ovulation. Endocrinology 136:3657–64.[Abstract]
Russell DL, Doyle KH, Ochsner SA, et al. (2003a) Processing and localization of ADAMTS-1 and proteolytic cleavage of versican during cumulus matrix expansion and ovulation. J Biol Chem 278:42330–9.
Russell DL, Doyle KMH, Gonzales-Robayna I, et al. (2003b) Egr-1 induction in rat granulosa cells by follicle-stimulating hormone and luteinizing hormone: combinatorial regulation by transcription factors cyclic adenosine 3',5'-monophosphate regulatory element binding protein, serum response factor, sp1, and early growth response factor-1. Mol Endocrinol 17:520–33.
Russell DL, Ochsner SA, Hsieh M, et al. (2003c) Hormone-regulated expression and localization of versican in the rodent ovary. Endocrinology 144:1020–31.
Saito H, Hirayama T, Koike K, et al. (1994) Cumulus mass maintains embryo quality. Fertil Steril 62:555–8.[Web of Science][Medline]
Salustri A, Garlanda C, Hirsch E, et al. (2004) PTX3 plays a key role in the organization of the cumulus oophorus extracellular matrix and in in vivo fertilization. Development 131:1577–86.
Salvador LM, Maizels E, Hales DB, et al. (2002) Acute signaling by the LH receptor is independent of protein kinase C activation. Endocrinology 143:2986–94.
Sanfins A, Lee GY, Plancha CE, et al. (2003) Distinctions in meiotic spindle structure and assembly during in vitro and in vivo maturation of mouse oocytes. Biol Reprod 69:2059–67.
Sanfins A, Plancha CE, Overstrom EW, et al. (2004) Meiotic spindle morphogenesis in in vivo and in vitro matured mouse oocytes: insights into the relationship between nuclear and cytoplasmic quality. Hum Reprod 19:2889–99.
Sato F and Marrs RP. (1986) The effect of pregnant mare serum gonadotropin on mouse embryos fertilized in vivo or in vitro. J In Vitro Fertilization Embryo Transfer 3:353–7.[CrossRef]
Sato H, Kajikawa S, Kuroda S, et al. (2001) Impaired fertility in female mice lacking urinary trypsin inhibitor. Biochem Biophys Res Commun 281:1154–60.[CrossRef][Web of Science][Medline]
Sayasith K, Lussier JG, Sirois J. (2005) Role of upstream stimulatory factor phosphorylation in the regulation of the prostaglandin G/H synthase-2 promoter in granulosa cells. J. Biol Chem 280:28885–93.
Sayasith K, Bouchard N, Sawadogo M, et al. (2004) Molecular characterization and role of bovine upstream stimulatory factor 1 and 2 in the regulation of the prostaglandin G/H synthase-2 promoter in granulosa cells. J Biol Chem 279:6327–36.
Schmits R, Filmus J, Gerwin N, et al. (1997) CD44 regulates hematopoietic progenitor distribution, granuloma formation, and tumorigenicity. Blood 90:2217–33.
Schoenfelder M and Einspanier R. (2003) Expression of hyaluronan synthases and corresponding hyaluronan receptors is differentially regulated during oocyte maturation in cattle. Biol Reprod 69:269–77.
Schultz RM, Montgomery RR, Belanoff JR. (1983) Regulation of mouse oocyte meiotic maturation: Implication of a decrease in oocyte cAMP and protein dephosphorylation in commitment to resume meiosis. Dev Biol 97:264–73.[CrossRef][Web of Science][Medline]
Seger R, Hanoch T, Rosenberg R, et al. (2001) The ERK signaling cascade inhibits gonadotropin-stimulated steroidogenesis. J Biol Chem 276:13957–64.
Segi E, Haraguchi K, Sugimoto Y, et al. (2003) Expression of messenger RNA for prostaglandin E receptor subtypes EP4/EP2 and cyclooxygenase isozymes in mouse periovulatory follicles and oviducts during superovulation. Biol Reprod 68:804–11.
Sekiguchi T, Mizutani T, Yamada K, et al. (2002) Transcriptional regulation of the epiregulin gene in the rat ovary. Endocrinology 143:4718–29.
Sela-Abramovich S and Chorev E, et al. (2005) Mitogen-activated protein kinase mediates luteinizing hormone-induced breakdown of communication and oocyte maturation in rat ovarian follicles. Endocrinology 146:1236–44.[CrossRef][Web of Science][Medline]
Sharma SC and Richards JS. (2000) Regulation of AP1 (Jun/Fos) factor expression and activation in ovarian granulosacells: relation of JunD and Fra2 to terminal differentiation. J Biol Chem 275:33718–28.
Shimada M, Nishibori M, Isobe N, et al. (2003) Luteinizing hormone receptor formation in cumulus cells surrounding porcine oocytes and its role during meiotic maturation of porcine oocytes. Biol Reprod 68:1142–9.
Shindo T, Kurihara H, Kuno K, et al. (2000) ADAMTS-1: a metalloproteinase-disintegrin essential for normal growth, fertility, and organ morphology and function. J Clin Invest 105:1345–52.[Web of Science][Medline]
Shitsukawa K, Andersen CB, Richard FJ, et al. (2001) Cloning and characterization of the cyclic guanosine monophosphate-inhibited phosphodiesterase PDE3A expressed in mouse oocyte. Biol Reprod 65:188–96.
Shukovski L and Tsafriri A. (1994) The involvement of nitric oxide in the ovulatory process in the rat. Endocrinology 135:2287–90.[Abstract]
Simon C, Tsafriri A, Chun S, et al. (1994) Interleukin-1 receptor antagonist suppresses human chorionic gonadotropin-induced ovulation in the rat. Biol Reprod 51:662–7.[Abstract]
Singh B and Armstrong D. (1997) Insulin-like growth factor-1, a component of serum that enables porcine cumulus cells to expand in response to follicle-stimulating hormone in vitro. Biol Reprod 56:1370–5.[Abstract]
Singh B, Zhang X, Armstrong D. (1993) Porcine oocytes release cumulus expansion-enabling activity even though porcine cumulus expansion in vitro is independent of the oocyte. Endocrinology 132:1860–2.
Sirois J and Richards JS. (1992) Purification and characterization of a novel, distinct isoform of prostaglandin endoperoxide synthase induced by human chorionic gonadotropin in granulosa cells of rat preovulatory follicles. J Biol Chem 267:6382–8.
Sirois J and Richards JS. (1993) Transcriptional regulation of the rat prostaglandin endoperoxide synthase 2 gene in granulosa cells. Evidence for the role of a cis- acting C/EBP beta promoter element. J Biol Chem 268:21931–8.
Sirois J, Simmons D, Richards J. (1992) Hormonal regulation of messenger ribonucleic acid encoding a novel isoform of prostaglandin endoperoxide H synthase in rat preovulatory follicles. Induction in vivo and in vitro. J Biol Chem 267:11586–92.
Sirois J, Sayasith K, Brown KA, et al. (2004) Cyclooxygenase-2 and its role in ovulation: a 2004 account. Hum Reprod Update 10:373–85.
Smith MF, Gutierrez CG, Ricke WA, et al. (2005) Production of matrix metalloproteinases by cultured bovine theca and granulosa cells. Reproduction 129:75–87.
Smitz J, Cortvrindt R, Hu Y. (1998) Epidermal growth factor combined with recombinant human chorionic gonadotrophin improves meiotic progression in mouse follicle-enclosed oocyte culture. Hum Reprod 13:664–9.
Somfai T, Kikuchi K, Onishi A, et al. (2004) Relationship between the morphological changes of somatic compartment and the kinetics of nuclear and cytoplasmic maturation of oocytes during in vitro maturation of porcine follicular oocytes. Mol Reprod Dev 68:484–91.[CrossRef][Web of Science][Medline]
Sriraman V and Richards JS. (2004) Cathepsin L gene expression and promoter activation in rodent granulosa cells. Endocrinology 145:582–91.
Sriraman V, Sharma SC, Richards JS. (2003) Transactivation of the progesterone receptor gene in granulosa cells: evidence that Sp1/Sp3 binding sites in the proximal promoter play a key role in luteinizing hormone inducibility. Mol Endocrinol 17:436–49.
Sriraman V, Rudd MD, Lohmann SM, et al. (2005) Cyclic GMP dependent protein kinase II is induced by LH and PR dependent mechanisms in granulosa cells and cumulus oocyte complexes of ovulating follicles. Mol Endocrinol me.2005–0317.
Standaert FE, Zamora CS, Chew BP. (1991) Quantitative and qualitative changes in blood leukocytes in the porcine ovary. Am J Reprod Immunol 25:163–8.[Medline]
Stanton H, Rogerson FM, East CJ, et al. (2005) ADAMTS5 is the major aggrecanase in mouse cartilage in vivo and in vitro. 434:648–52.
Steinbauch M and Loeb J. (1961) Isolation of an alpha2-globulin from human plasma. Nature 192:1196.[Medline]
Sterneck E, Tessarollo L, Johnson PF. (1997) An essential role for C/EBPbeta in female reproduction. Genes Dev 11:2153–62.
Stickens D, Behonick DJ, Ortega N, et al. (2004) Altered endochondral bone development in matrix metalloproteinase 13-deficient mice. Development 131:5883–95.
Stouffer RL. (2002) Pre-ovulatory events in the rhesus monkey follicle during ovulation induction. Reprod Biomed Online 4:1–4.
Stouffer RL. (2003) Progesterone as a mediator of gonadotrophin action in the corpus luteum: beyond steroidogenesis. Hum Reprod Update 9:99–117.
Stouffer RL and Duffy DL. (2003) Luteinizing hormone acts directly at granulosa cells to stimulate periovulatory processes: modulation of luteinizing hormone effects by prostaglandins. Endocrine 22:249–56.[CrossRef][Web of Science][Medline]
Su Y-Q, Wigglesworth K, Pendola FL, et al. (2002) Mitogen-activated protein kinase activity in cumulus cells is essential for gonadotropin-induced oocyte meiotic resumption and cumulus expansion in the mouse. Endocrinology 143:2221–32.
Su Y-Q, Denegre JM, Wigglesworth K, et al. (2003) Oocyte-dependent activation of mitogen-activated protein kinase (ERK1/2) in cumulus cells is required for the maturation of the mouse oocyte-cumulus cell complex. Dev Biol 263:126–38.[CrossRef][Web of Science][Medline]
Sun GW, Kobayashi H, Suzuki M, et al. (2002) Link protein as an enhancer of cumulus cell-oocyte complex expansion. Mol Reprod Dev 63:223–31.[CrossRef][Web of Science][Medline]
Sun GW, Kobayashi H, Suzuki M, et al. (2003) Follicle-stimulating hormone and insulin-like growth factor i synergistically induce up-regulation of cartilage link protein (Crtl1) via activation of phosphatidylinositol-dependent kinase/Akt in rat granulosa cells. Endocrinology 144:793–801.
Sutton ML, Gilchrist RB, Thompson JG. (2003) Effects of in-vivo and in-vitro environments on the metabolism of the cumulus-oocyte complex and its influence on oocyte developmental capacity. Hum Reprod Update 9:35–48.
Sutton-McDowall ML, Gilchrist RB, Thompson JG. (2004) Cumulus expansion and glucose utilisation by bovine cumulus-oocyte complexes during in vitro maturation: the influence of glucosamine and follicle-stimulating hormone. Reproduction 128:313–9.
Suzuki T, Sasano H, Kimura N, et al. (1994) Immunohistochemical distribution of progesterone, androgen and oestrogen receptors in the human ovary during the menstrual cycle: relationship to expression of steroidogenic enzymes. Hum Reprod 9:1589–95.
Suzuki T, Sasano H, Takaya R, et al. (1998) Leukocytes in normal-cycling human ovaries: immunohistochemical distribution and characterization. Hum Reprod 13:2186–91.
Suzuki M, Kobayashi H, Tanaka Y, et al. (2004) Reproductive failure in mice lacking inter-alpha-trypsin inhibitor (ITI) - ITI target genes in mouse ovary identified by microarray analysis. J Endocrinol 183:29–38.
Szoltys M, Slomczynska M, Tabarowski Z. (2003) Immunohistochemical localization of androgen receptor in rat oocytes. Folia Histochem Cytobiol 41:59–64.[Medline]
Tadakuma H, Okamura H, Kitaoka M, et al. (1993) Association of immunolocalization of matrix metalloproteinase 1 with ovulation in hCG-treated rabbit ovary. J Reprod Fertil 98:503–8.
Tajima K, Dantes A, Yao Z, et al. (2003) Down-regulation of steroidogenic response to gonadotropins in human and rat preovulatory granulosa cells involves mitogen-activated protein kinase activation and modulation of DAX-1 and steroidogenic factor-1. J Clin Endocrinol Metab 88:2288–99.
Takahashi T, Morrow JD, Wang H, et al. (2006) Cyclooxygenase-2 derived prostaglandin E2 directs oocyte maturation by differentially influencing multiple signaling pathways. J Biol Chem in press.
Talbot P, Shur BD, Myles DG. (2003) Cell adhesion and fertilization: steps in oocyte transport, sperm-zona pellucida interactions, and sperm-egg fusion. Biol Reprod 68:1–9.
Tamura K, Tamura H, Kumasaka K, et al. (1998) Ovarian immune cells express granulocyte-macrophage colony-stimulating factor (GM-CSF) during follicular growth and luteinization in gonadotropin-primed immature rodents. Mol Cell Endocrinol 142:153–63.[CrossRef][Web of Science][Medline]
Tanghe S, Van Soom A, Mehrzad J, et al. (2003) Cumulus contributions during bovine fertilization in vitro. Theriogenology 60:135–49.[CrossRef][Web of Science][Medline]
Tesarik J, Pilka L, Drahorad J, et al. (1988) The role of cumulus cell-secreted proteins in the development of human sperm fertilizing ability: implication in IVF. Hum Reprod 3:129–32.
Tirone E, D'Alessandris C, Hascall VC, et al. (1997) Hyaluronan synthesis by mouse cumulus cells is regulated by interactions between follicle-stimulating hormone (or epidermal growth factor) and a soluble oocyte factor (or transforming growth factor beta 1). J Biol Chem 272:4787–94.
Tomic D, Miller KP, Kenny HA, et al. (2004) Ovarian follicle development requires Smad3. Mol Endocrinol 18:2224–40.
Topilko P, Schneider-Maunoury S, Levi G, et al. (1998) Multiple pituitary and ovarian defects in Krox-24 (NGFI-A, Egr-1)- targeted mice. Mol Endocrinol 12:107–22.[CrossRef][Web of Science][Medline]
Tsafriri A. (1995) Ovulation as a tissue remodelling process. Proteolysis and cumulus expansion. Adv Exp Med Biol 377:121–40.[Medline]
Tsafriri A, Abisogun AO, Reich R. (1987) Steroids and follicular rupture at ovulation. J Steroid Biochem 27:359–63.[CrossRef][Web of Science][Medline]
Tsafriri A, Lieberman ME, Ahren K, et al. (1976a) Dissociation between LH-induced aerobic glycolysis and oocyte maturation in cultured Graafian follicles of the rat. Acta Endocrinol 81:362–6.
Tsafriri A, Lieberman ME, Koch Y, et al. (1976b) Capacity of immunologically purified FSH to stimulate cyclic AMP accumulation and steroidogenesis in Graafian follicles and to induce ovum maturation and ovulation in the rat. Endocrinology 98:655–61.
Tsafriri A, Bicsak TA, Cajander SB, et al. (1989) Suppression of ovulation rate by antibodies to tissue-type plasminogen activator and alpha 2-antiplasmin. Endocrinology 124:415–21.
Tsafriri A, Chun S-Y, Zhang R, et al. (1996) Oocyte maturation involves compartmentalization and opposing changes of cAMP levels in follicular somatic and germ cells: studies using selective phosphodiesterase inhibitors. Dev Biol 178:393–402.[CrossRef][Web of Science][Medline]
Tsafriri A, Cao X, Ashkenazi H, et al. (2005) Resumption of oocyte meiosis in mammals: On models, meiosis activating sterols, steroids and EGF-like factors. Mol Cell Endocrinol 234:37–45.[CrossRef][Web of Science][Medline]
Tullet JMA, Pocock V, Steel JH, et al. (2005) Multiple signaling defects in the absence of RIP140 impair both cumulus expansion and follicle rupture. Endocrinology 146:4127–37.
Turley EA, Noble PW, Bourguignon LYW. (2002) Signaling properties of hyaluronan receptors. J Biol Chem 277:4589–92.
Ujioka T, Russell DL, Okamura H, et al. (2000) Expression of regulator of G-protein signaling protein-2 gene in the rat ovary at the time of ovulation [In Process Citation]. Biol Reprod 63:1513–7.
Vaday GG, Franitza S, Schor H, et al. (2001) Combinatorial signals by inflammatory cytokines and chemokines mediate leukocyte interactions with extracellular matrix. J Leukoc Biol 69:885–92.
Van der Hoek KH, Maddocks S, Woodhouse CM, et al. (2000) Intrabursal injection of clodronate liposomes causes macrophage depletion and inhibits ovulation in the mouse ovary. Biol Reprod 62:1059–66.
Van Soom A, Tanghe S, De Pauw I, et al. (2002) Function of the cumulus oophorus before and during mammalian fertilization. Reprod Domest Anim 37:144–51.[CrossRef][Web of Science][Medline]
Vanderhyden BC, Caron PJ, Buccione R, et al. (1993) Developmental pattern of the secretion of cumulus expansion-enabling factor by mouse oocytes and the role of oocytes in promoting granulosa cell differentiation. Dev Biol 140:307–17.[CrossRef]
Vanderhyden BC, Macdonald EA, Nagyova E, et al. (2003) Evaluation of members of the TGFbeta superfamily as candidates for the oocyte factors that control mouse cumulus expansion and steroidogenesis. Reproduction 61:55–70.[Medline]
Varani S, Elvin JA, Yan C, DeMayo J, et al. (2002) Knockout of pentraxin 3, a downssream target of growth differentiation factor-9, causes female subfertility. Mol Endocrinol 16:1154–67.
Vermeiden J, Roseboom T, Goverde A, et al. (1997) An artificially induced follicle stimulating hormone surge at the time of human chorionic gonadotrophin administration in controlled ovarian stimulation cycles has no effect on cumulus expansion, fertilization rate, embryo quality and implantation rate. Hum Reprod 12:1399–402.
Vu TH, Shipley JM, Bergers G, et al. (1998) MMP-9/gelatinase B is a key regulator of growth plate angiogenesis and apoptosis of hypertrophic chondrocytes. Cell 93:411–22.[CrossRef][Web of Science][Medline]
Wang X and Greenwald G. (1993a) Human chorionic gonadotropin or human recombinant follicle-stimulating hormone (FSH)-induced ovulation and subsequent fertilization and early embryo development in hypophysectomized FSH-primed mice. Endocrinology 132:2009–16.
Wang X and Greenwald G. (1993b) Hypophysectomy of the cyclic mouse. I. Effects on folliculogenesis, oocyte growth, and follicle-stimulating hormone and human chorionic gonadotropin receptors. Biol Reprod 48:585–94.[Abstract]
Wang H, Wen Y, Polan ML, et al. (2005) Exogenous granulocyte-macrophage colony-stimulating factor promotes follicular development in the newborn rat in vivo. Hum Reprod 20:2749–56.
Wang LJ, Brannstrom M, Cui KH, et al. (1997) Localisation of mRNA for interleukin-1 receptor and interleukin-1 receptor antagonist in the rat ovary. J Endocrinol 152:11–7.
Webb RJ, Marshall F, Swann K, et al. (2002) Follicle-stimulating hormone induces a gap junction-dependent dynamic change in [cAMP] and protein kinase A in mammalian oocytes. Dev Biol 246:441–54.[CrossRef][Web of Science][Medline]
White R, Leonardsson G, Rosewell I, et al. (2000) The nuclear receptor co-repressor Nrip1 (RIP140) is essential for female fertility. Nat Med 6:1368–74.[CrossRef][Web of Science][Medline]
Wiersma A and HirschB, Tsafriri A, et al. (1998) Phosphodiesterase 3 inhibitors suppress oocyte maturation and consequent pregnancy without affecting ovulation and cyclicity in rodents. J Clin Invest 102:532–7.[Web of Science][Medline]
Wilson CL, Heppner KJ, Labosky PA, et al. (1997) Intestinal tumorigenesis is suppressed in mice lacking the metalloproteinase matrilysin. Proc Natl Acad Sci U S A 94:1402–7.
Wong WY and Richards JS. (1992) Induction of prostaglandin H synthase in rat preovulatory follicles by gonadotropin-releasing hormone. Endocrinology 130:3512–21.
Wu R, Van der Hoek KH, Ryan NK, et al. (2004) Macrophage contributions to ovarian function. Hum Reprod Update 10:119–33.
Wu Y, Wu J, Lee DY, et al. (2005) Versican protects cells from oxidative stress-induced apoptosis. Matrix Biol 24:3–13.[Web of Science][Medline]
Wulff C, Wiegand SJ, Saunders PTK, et al. (2001) Angiogenesis during follicular development in the primate and its inhibition by treatment with truncated Flt-1-Fc (vascular endothelial growth factor TrapA40). Endocrinology 142:3244–54.
Wulff C, Wilson H, Wiegand SJ, et al. (2002) Prevention of thecal angiogenesis, antral follicular growth, and ovulation in the primate by treatment with vascular endothelial growth factor Trap R1R2. Endocrinology 143:2797–807.
Xu F, Hazzard TM, Evans A, et al. (2005) Intraovarian actions of anti-angiogenic agents disrupt periovulatory events during the menstrual cycle in monkeys. Contraception 71:239–48.[CrossRef][Web of Science][Medline]
Yan C, Wang P, DeMayo J, et al. (2001) Synergistic roles of bone morphogenetic protein 15 and growth differentiation factor 9 in ovarian function. Mol Endocrinol 15:854–66.
Yang W-L, Godwin AK, Xu X-X. (2004) Tumor necrosis factor-{alpha}-induced matrix proteolytic enzyme production and basement membrane remodeling by human ovarian surface epithelial cells: molecular basis linking ovulation and cancer risk. Cancer Res 64:1534–40.
Yang S-H, Son W-Y, Yoon S-H, et al. (2005) Correlation between in vitro maturation and expression of LH receptor in cumulus cells of the oocytes collected from PCOS patients in HCG-primed IVM cycles. Hum Reprod 20:2097–103.
Yeung WS, Lee KF, Koistinen R, et al. (2006) Roles of glycodelin in modulating sperm function. Mol Cell Endocrinol 250:149–56.[CrossRef][Web of Science][Medline]
Yokoo M, Tientha P, Kimura N, et al. (2002) Localisation of the hyaluronan receptor CD44 in porcine cumulus cells during in vivo and in vitro maturation. Zygote 10:317–26.[CrossRef][Web of Science][Medline]
Yoshino M, Mizutani T, Yamada K, et al. (2002) Early growth response gene-1 regulates the expression of the rat luteinizing hormone receptor gene. Biol Reprod 66:1813–9.
Yoshioka S, Ochsner S, Russell DL, et al. (2000) Expression of tumor necrosis factor-stimulated gene-6 in the rat ovary in response to an ovulatory dose of gonadotropin. Endocrinology 141:4114–9.
Young KA, Tumlinson B, Stouffer RL. (2004) ADAMTS-1/METH-1 and TIMP-3 expression in the primate corpus luteum: divergent patterns and stage-dependent regulation during the natural menstrual cycle. Mol Hum Reprod 10:559–65.
Yun YW, Yuen BH, Moon YS. (1987) Effects of superovulatory doses of pregnant mare serum gonadotropin on oocyte quality and ovulatory and steroid hormone responses in rats. Gamete Res 6:109–20.
Zackrisson U, Mikuni M, Wallin A, et al. (1996) Ovary and ovulation: cell-specific localization of nitric oxide synthases (NOS) in the rat ovary during follicular development, ovulation and luteal formation. Hum Reprod 11:2667–73.
Zeleznik AJ, Saxena D, Little-Ihrig L. (2003) Protein kinase B is obligatory for follicle-stimulating hormone-induced granulosa cell differentiation. Endocrinology 144:3985–94.[CrossRef][Web of Science][Medline]
Zelinski-Wooten MB, Hutchison JS, Hess DL, et al. (1998) A bolus of recombinant human follicle stimulating hormone at midcycle induces periovulatory events following multiple follicular development in macaques. Hum Reprod 13:554–60.
Zhang Y and Dufau ML. (2003) Dual mechanisms of regulation of transcription of luteinizing hormone receptor gene by nuclear orphan receptors and histone deacetylase complexes. J Steroid Biochem Mol Biol 85:401–14.[CrossRef][Web of Science][Medline]
Zhang X, Jafari N, Barnes RB, et al. (2005) Studies of gene expression in human cumulus cells indicate pentraxin 3 as a possible marker for oocyte quality. Fertil Steril 83:1169–79.[CrossRef][Web of Science][Medline]
Zhu X, Gilbert S, Birnbaumer M, et al. (1994) Dual signaling potential is common among Gs-coupled receptors and dependent on receptor density. Mol Pharmacol 46:460–9.[Abstract]
Zhuo L, Yoneda M, Zhao M, et al. (2001) Defect in SHAP-hyaluronan complex causes severe female infertility. A study by inactivation of the bikunin gene in mice. J Biol Chem 276:7693–6.
Zimmermann RC, Xiao E, Husami N, et al. (2001) Short-term administration of antivascular endothelial growth factor antibody in the late follicular phase delays follicular development in the rhesus monkey. J Clin Endocrinol Metab 86:768–72.
Zimmermann RC, Xiao E, Bohlen P, et al. (2002) Administration of antivascular endothelial growth factor receptor 2 antibody in the early follicular phase delays follicular selection and development in the rhesus monkey. Endocrinology 143:2496–502.
Zimmermann RC, Hartman T, Kavic S, et al. (2003) Vascular endothelial growth factor receptor 2-mediated angiogenesis is essential for gonadotropin-dependent follicle development. J Clin Invest 112:659–69.[CrossRef][Web of Science][Medline]
Zuelke KA and Brackett BG. (1992) Effects of luteinizing hormone on glucose metabolism in cumulus-enclosed bovine oocytes matured in vitro. Endocrinology 131:2690–6.
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