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research-article |
Centre for Diabetes and Endocrinology,3 Rayne Institute, University College London, London WC1E 6JJ, United Kingdom
Department of Metabolic Medicine,4 and Institute of Reproductive and Developmental Biology,5 Imperial College London, London W12 0NN, United Kingdom
ABSTRACT
Insulin receptor signaling regulates female reproductive function acting in the central nervous system and ovary. Female mice that globally lack insulin receptor substrate (IRS) 2, which is a key mediator of insulin receptor action, are infertile with defects in hypothalamic and ovarian functions. To unravel the tissue-specific roles of IRS2, we examined reproductive function in female mice that lack Irs2 only in the neurons. Surprisingly, these animals had minimal defects in pituitary and ovarian hormone levels, ovarian anatomy and function, and breeding performance, which indicates that the central nervous system IRS2 is not an obligatory signaling component for the regulation of reproductive function. Therefore, we undertook a detailed analysis of ovarian function in a novel Irs2 global null mouse line. Comparative morphometric analysis showed reduced follicle size, increased numbers of atretic follicles, as well as impaired oocyte growth and antral cavity development in Irs2 null ovaries. Granulosa cell proliferation was also defective in the Irs2 null ovaries. Furthermore, the insulin- and eCG-stimulated phosphoinositide-3-OH kinase signaling events, which included phosphorylation of Akt/protein kinase B and glycogen synthase kinase 3-beta, were impaired, whereas mitogen-activated protein kinase signaling was preserved in Irs2 null ovaries. These abnormalities were associated with reduced expression of cyclin D2 and increased CDKN1B levels, which indicates dysregulation of key components of the cell cycle apparatus implicated in ovarian function. Our data suggest that ovarian rather than central nervous system IRS2 signaling is important in the regulation of female reproductive function.
insulin, insulin-like growth factor receptor, kinases, oocyte development, ovary
Insulin receptor (INSR) signaling pathways regulate peripheral energy homeostasis by acting in skeletal muscle, adipose tissue, and the liver to control carbohydrate, lipid, and protein metabolism. It is clear that these signals also act in other tissues, for example, in the regulation of pancreatic ß-cell and hypothalamic functions [1]. Insulin receptor signaling has also been implicated in the regulation of female reproductive function through its actions in both the central nervous system (CNS) and ovaries. Female mice that lack insulin receptors in their neurons show impaired fertility with fewer antral follicles and corpora lutea due to reduced release of LH [2]. Insulin also stimulates GnRH secretion in vivo, and intracerebroventricular administration of insulin restores reproductive behavior in diabetic rats [3, 4]. In vitro studies using immortalized GnRH neuronal cell lines have suggested that insulin regulates GnRH expression through the mitogen-activated protein kinase pathway [5]. Therefore, CNS insulin signaling plays a role in regulating female reproductive function in rodents. Insulin signaling has also been shown to have complex roles in ovarian function, including the regulation of ovarian steroidogenesis, follicular development, and granulosa cell proliferation [6–8]. The strong association of insulin resistance with ovarian dysfunction in polycystic ovarian syndrome also suggests a role for insulin signaling in ovarian function [9].
Insulin receptor substrate (IRS) proteins mediate the effects of the insulin receptor on cellular and whole body physiology, including reproduction [10]. Mice that lack Irs1 display profound growth retardation and insulin resistance but have mildly defective reproductive function [11, 12]. Irs3 and Irs4 null mice have minimal metabolic, endocrine, and growth phenotypes [10]. In contrast, mice that lack Irs2 develop diabetes due to a combination of insulin resistance and pancreatic ß cell dysfunction [11]. Female Irs2 null mice are also infertile due to reduced pituitary LH levels and gonadotroph cell numbers, and show reduced gonadotropin-stimulated ovulation and markedly reduced numbers of ovarian follicles and corpora lutea [12]. While CNS IRS2 signaling plays complex roles in energy homeostasis [1], the contribution of CNS IRS2 to female reproductive function has not been established. Ovarian cellular defects and downstream signaling abnormalities in Irs2 null mice have not been characterized. Therefore, we used conditional gene targeting in mice to delete Irs2 in the CNS and examined their reproductive functions. In addition, we generated a novel Irs2 null mouse line and assessed its ovarian morphology, signaling, and cell cycle events.
All of the in vivo studies were performed in accordance to the United Kingdom Home Office Animal Procedures Act (1986) and University College London Animal Ethical Committee Guidelines.
Mice with a floxed allele of Irs2 (Irs2tm1With) and mice that lack Irs2 in the CNS, Irs2tm1With /Irs2tm1With Tg (Nes-cre)1Kln/0 (hereinafter designated as NesCreIrs2KO mice) have been described previously [1]. Mice with a germ-line deletion of Irs2 were also generated using embryonic stem cells that had undergone complete deletion of the Irs2 locus following Cre recombinase treatment during the same targeting strategy [1]. The mice were back-crossed five times onto a C57Bl/6 background prior to analysis. Genotyping was performed as previously described [1]. The mice were maintained on a 12L:12D cycle with free access to water and standard mouse chow (4% fat, RM1; Special Diet Services, Witham, Essex, UK) and housed in specific-pathogen-free barrier facilities.
In Vivo Analyses of Reproductive Function
Estrus cycle characteristics were determined by analysis of vaginal smears for cell type and morphology with Giemsa staining each day for three complete cycles. Fertility was assessed by monitoring daily for 3 mo the numbers of pregnancies, litters, and pups in continuous mating studies with confirmed male breeders and 7–9-week-old females.
Hormone Stimulation, Sample Collection, and Hormone and Peptide Measurements
For the superovulation studies, 8–10-wk-old mice received 10 IU eCG (Folligon; Intervet) by i.p. injection, followed 46 h later by 8 IU hCG (Intervet). Oocytes were flushed from the oviduct 14–16 h after the second injection. For bromodeoxyuridine (BrdU) incorporation studies, the mice received an i.p. injection of 100 mg/kg BrdU (Roche) 2 h before tissue harvesting. Blood samples were collected by tail vein bleeding or by cardiac puncture of terminally anesthetized mice at specific stages of the estrus cycle, and the serum was either stored at –80°C prior to analysis or allowed to coagulate at 4°C overnight, centrifuged, and frozen at –20°C. Pituitaries were harvested, homogenized in 1 M urea in PBS, and stored at –80°C prior to analysis. Serum estrogen and progesterone were measured using immunofluorometric assays (Delfia; Perkin Elmer, Turku, Finland). RIAs for pituitary LH, FSH, and prolactin were performed as previously described [13, 14].
Histomorphometric Analyses and Immunohistochemistry
Ovaries were fixed in 10% buffered formalin, embedded in paraffin, and cut into 5-µm sections. For histomorphometric analysis, sections were stained with hematoxylin-eosin. Follicles, which were classified according to diameter and number of granulosa cell layers, were counted in every tenth section, starting from the fifth section, through the entire ovary. Only those follicles with a clearly visible oocyte nucleus were counted, to avoid double counting. Atretic follicles were defined according to the following morphological criteria: follicle and oocyte shapes, localization of the oocyte in the follicle, presence of a degenerated oocyte, altered zona pellucida, the presence of granulosa cells inside the zona pellucida, the presence of more than three pyknotic nuclei in the granulosa cells, and macrophage invasion. Follicle growth, follicle and oocyte size (diameter), and total follicular and antral cavity area were measured using the LUCIA software (Nikon UK, Kingston-upon-Thames, UK). For immunohistochemistry, sections were deparaffinized and antigen retrieval was performed by microwave boiling of the sections in 10 mM citrate buffer (pH 6.0) twice for 10 min each. Sections were incubated in 20% (v/v) normal goat serum in PBS with 4% (w/v) BSA for 30 min at room temperature, and then with the indicated primary antibodies overnight at 4°C. Detection of bound antibodies was performed using the avidin-biotin complex method (Vectastain ABC Elite kit, Vector Laboratories). Sections were also counterstained with hematoxylin, dehydrated in a graded ethanol series, cleared in xylene, and mounted. The Zymed proliferation cell nuclear antigen (PCNA) and BrdU detection kits were used for PCNA and BrdU staining, respectively.
Mice (8–10 wk of age) were either superovulated as described above or treated with insulin as described previously [15]. In brief, mice were anesthetized and injected i.v. with saline or 5 U of soluble insulin, and the ovaries were removed and snap frozen 10 min later. Tissue preparation and lysis, Western blotting, and quantification were performed as previously described [15]. The blots were stripped and probed for actin when loading controls were required.
Affinity-purified rabbit anti-IRS1 and anti-IRS2 antibodies were from Upstate Biotechnology (Dundee, UK). The anti-AKT, anti-pAKTSer473, anti-glycogen synthase kinase (GSK)3B, anti-pGSK3A/BSer9/21, anti-mitogen-activated protein kinase (MAPK) and anti-pMAPKThr202/Tyr204 antibodies were from Cell Signaling Technology (Beverly, MA). Rabbit polyclonal antibodies to cyclin D2 and CDKN1B (p27KIP1) and mouse anti-actin were from Santa Cruz Biotechnology (Insight Biotechnology, Wembley, UK).
Real-time PCR was performed as previously described using FAM/TAMRA primers (Applied Biosystems, Foster City, CA) [15]. The following primer sets were used: Irs1 Mm00439720_s1, Irs2 (MIRS 396412), and Hprt1 Mm01545399_m1. The relative amounts of mRNA were calculated from an internal standard curve following normalization to the Hprt1 mRNA levels.
All statistical analyses were performed using the GraphPad Prism4 software (GraphPad Software, San Diego, CA), and paired and unpaired t-tests and two-way ANOVA with Bonferroni post-hoc tests were performed as appropriate. P < 0.05 was regarded as statistically significant.
Reproductive Phenotype of NesCreIrs2KO Mice
NesCreIrs2KO mice lack Irs2 expression in the CNS and hypothalamus [1], while their expression of pituitary and ovarian Irs2 mRNA is equivalent to that in control animals, i.e., the relative mRNA levels expressed as percentages of the wild-type (WT) control are: Irs2 in NesCreIrs2KO pituitaries, 98.65 ± 12.5%; Irs2 in NesCreIrs2KO ovaries, 107.4 ± 15.2%; n = 12, P > 0.05. There were no changes in Irs1 expression in the ovary, hypothalamus, and pituitary of NesCreIrs2KO mice (data not shown). Therefore NesCreIrs2KO mice represent a good model for determining the contribution of CNS IRS2 pathways to female reproductive function. The NesCreIrs2KO mice exhibited an extended estrus cycle due to a mildly prolonged diestrus phase (Table 1). Diestrus pituitary prolactin levels were reduced in NesCreIrs2KO mice, whereas the serum estradiol and progesterone and pituitary FSH and LH levels were similar to those of WT mice (Table 1). No differences in the timing of ovulation and the numbers of oocytes recovered were seen when NesCreIrs2KO mice and WT mice were superovulated (total oocytes recovered per animal: WT, 23.6 ± 2.3 vs. NesCreIrs2KO 21.1 ± 2.1, n = 5, P > 0.05). Morphological studies showed that follicles at all developmental stages and corpora lutea were present in these animals and in WT mice, which is consistent with their normal responses to superovulation (Fig. 1, A and B and data not shown). In continuous mating studies, the average duration between litters and the size and number of litters over the study period were similar in NesCreIrs2KO and WT mice (Table 1). NesCreIrs2KO mice were able to suckle and rear their progeny and there were no differences in the postpartum survival rates of these mice compared to WT females (data not shown). Therefore, somewhat surprisingly, deletion of Irs2 in the CNS has a minimal effect on reproductive function in mice.
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A previous analysis of ovarian function in Irs2 null mice was limited to the demonstration of reduced numbers of ovarian follicles and corpora lutea, defective ovarian responses to superovulation, and reduced circulating gonadal steroid levels [12]. There are currently limited options for ovarian-specific gene deletion throughout folliculogenesis. Therefore, we examined in detail the ovarian morphology and ovarian cell proliferation and we analyzed the signaling events and cell cycle components downstream of IRS2 in our novel Irs2 null mouse on a C57Bl/6 background.
After superovulation, reduced numbers of oocytes were recovered from 8–10-week-old Irs2 null mice (total oocytes recovered per animal: WT, 23.6 ± 2.3 vs. Irs2 null 8.3 ± 0.8, n = 5, P < 0.01). Irs2 null ovaries showed increased numbers of atretic follicles, reduced numbers of large antral follicles, and an absence of preovulatory follicles (Fig. 1, C and D). Additional characteristic features of Irs2 null ovaries were the presence of oocytes trapped in corpora lutea (Fig. 1E) and the presence of occasional follicles with double oocytes (Fig. 1F). We explored further the mechanisms underlying these defects.
IRS2 and IRS1 Protein Expression by Ovarian Cells
Strong specific IRS2 staining was seen in oocytes and granulosa cell cytoplasm of the early primordial follicles in the WT mice but not in the Irs2 null animals (Fig. 2, A–D). Less-extensive IRS2 staining was detected in the thecal and stromal cells at all stages of follicular development in the WT mice (Fig. 2, A–C). IRS1 was expressed in a range of ovarian cell types in the WT mice but not in the Irs1 null mice (Fig. 2, E–H). Staining was more intense in the granulosa cells than in thecal cells and increased with follicular growth. In the Irs2 null mice, there was an apparent reduction in the expression of IRS1 protein in the ovary (Fig. 2, I–K). This was confirmed by the demonstration of reduced Irs1 mRNA expression in Irs2 null ovaries (relative Irs1 mRNA levels expressed as percentage of the WT control: 73 ± 2%, n = 3, P < 0.001). These findings suggest that IRS2 plays complex roles in ovarian cellular function. Therefore, we characterized in detail the follicular morphologies of the WT and Irs2 null mice.
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Analysis of Follicular Development in Irs2 Null Ovaries
Morphometric analysis confirmed a marked reduction in the number of follicles of all sizes in Irs2 null ovaries and that 75% of the large antral follicles were atretic in Irs2 null mice (Fig. 3, A and B). When there were fewer than three follicular granulosa cell layers, no differences were observed between the development of these follicles in Irs2 null and WT mice (Fig. 3C). However, the follicles at later stages were reduced in size (Fig. 3C), which suggests that Irs2 null ovaries are defective for granulosa cell proliferation within growing follicles. Complex interactions occur between oocytes and other follicular components, with oocytes promoting follicular growth and directing granulosa cell differentiation [16–19]. In Irs2 null mice, the early oocytes grew as quickly as those in the WT mice but growth was impaired at later stages (Fig. 3D). Antral cavity development and expansion were also markedly attenuated in Irs2 null ovaries (Fig. 3E). After superovulation in the Irs2 null mice there was a minimal increase in antral cavity size (data not shown). These morphological parameters demonstrate that in Irs2 null follicles, three events that are important for normal follicular development from the growing pool to the preovulatory stage are markedly disturbed.
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Granulosa Cell Proliferation in Irs2 Null Follicles
We used PCNA expression and BrdU incorporation to assess further cell proliferation. In the WT ovaries, nuclear PCNA staining was seen in cuboidal granulosa cells in the primary follicles and in most of the granulosa cells of follicles at later stages (Fig. 4, A–D). Thecal cell staining was seen as the follicles developed granulosa cell layers. Oocytes were PCNA positive in the primary follicles onwards (Fig. 4A). In antral follicles, PCNA was expressed in all the granulosa cell layers, with the highest levels seen in those around the oocyte (Fig. 4C). Superovulation induced preovulatory follicles with strong cumulus PCNA staining (Fig. 4D). In contrast, Irs2 null ovaries displayed a marked reduction in granulosa cell PCNA staining in all follicular classes (Fig. 4, E–H). After superovulation, there was little PCNA staining in the Irs2 null ovaries, which suggests that the vast majority of granulosa cells in Irs2 null ovaries do not proceed to a rapid phase of proliferation (Fig. 4H).
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To assess S phase progression, we analyzed BrdU incorporation in the ovaries. In small follicles with two or fewer granulosa cell layers, there were no differences in the numbers of BrdU positive cells in the WT and Irs2 null ovaries. In the WT mice, increased follicle size was associated with a marked increase in BrdU labeling (Fig. 4I). In the large antral follicles, there was a general reduction in BrdU staining but the cumulus cells showed a high frequency of positive staining (Fig. 4J). BrdU labeling was reduced in Irs2 null follicles of all sizes (Fig. 4, K and L). Superovulation increased BrdU labeling in Irs2 null follicles but to a significantly lesser extent to that seen in WT ovaries (percentage of BrdU-positive cells in secondary follicles (WT vs. Irs2 null): without stimulation, 13.8 ± 2.3 vs. 3.6 ± 0.9, n = 6, P < 0.05: after superovulation, 19.3 ± 3.4 vs. 1.7 ± 1.2, n = 6, P < 0.05; and percentage of BrdU-positive cells in the large antral follicles: without stimulation, 26.9 ± 2.1 vs. 9.6 ± 0.3, n = 6, P < 0.05; after superovulation, 33.1 ± 3.3 vs. 17.8 ± 2.8, n = 6, P < 0.05).
Defective PI3 Kinase Signaling Pathways and Abnormal Expression of Cell Cycle Regulators Cyclin D2 and CDKN1B in Irs2 Null Ovaries
IRS2 recruits the PI3 kinase/AKT and MAP kinase pathways, which together mediate the effects of insulin and IGF1 [10]. These downstream signaling events were not examined in the Irs2 null mice described previously [12]. Insulin induced a 5-fold increase in Akt phosphorylation in WT ovaries, which was indicative of PI3 kinase pathway activation, although this was largely abrogated in Irs2 null ovaries (Fig. 5, A and B). Treatment with hCG stimulated AKT phosphorylation in the ovaries of mice of either genotype but both the basal and peak-stimulated AKT phosphorylation were attenuated in Irs2 null ovaries, despite similar levels of AKT expression (Fig. 5, C and D). AKT phosphorylates and inactivates glycogen synthase kinase-3 ß (GSK3B) [20, 21], which in turn phosphorylates cyclin D, thereby targeting it for degradation [22]. Insulin increased GSK3B phosphorylation in the WT ovaries but not in the Irs2 null ovaries (Fig. 5, E and F). In contrast, insulin-stimulated MAP kinase activation was equivalent in the ovaries of both genotypes and hCG-stimulated MAP kinase activation was also largely preserved (Fig. 5, G and H and data not shown).
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We also examined the expression levels of cyclin D2, which is a critical component of the cell cycle apparatus that regulates G1/S phase progression, and the cell cycle inhibitor CDKN1B. The basal expression of cyclin D2 was reduced and that of CDKN1B was elevated in Irs2 null ovaries compared to WT ovaries, which implies an imbalance between these cell cycle regulators (Fig. 5, I–L). Superovulation induction with eCG followed by hCG injection caused a significant reduction in cyclin D2 expression 24 h after hCG treatment (Fig. 5, I and J). This effect was significantly impaired in Irs2 null ovaries. The same treatment induced a significant increase in CDKN1B expression in the WT ovaries but no significant alteration in Irs2 null ovaries, which also demonstrated a higher basal level of CDKN1B (Fig. 5, K and L).
Primary follicles from mice of both genotypes showed no differences in cyclin D2 immunostaining, with a strong pattern of staining seen in most granulosa cells (Fig. 6, A and F). In the subsequent stages of follicular growth, both the numbers of granulosa cells that expressed cyclin D2 and the intensity of staining were reduced in Irs2 null compared to WT ovaries (Fig. 6, B–E and G–J). In WT ovaries, 46 h after the administration of eCG alone, most of the stained cells were granulosa cells surrounding the oocyte (Fig. 6C). In large preovulatory follicles (with or without prior superovulation induction) most of the stained granulosa cell were cumulus cells, whereas the cells in the mural granulosa were mostly unstained (Fig. 6, D and E). These results are consistent with the patterns of PCNA staining and BrdU incorporation described above. In contrast, in the Irs2 null ovaries in which large antral follicles were found, few cells expressed cyclin D2 (Fig. 6, I and J), which suggests that impaired proliferation of cumulus cells contributes to defective ovulation in these mice.
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High levels of CDKN1B expression were found in the corpora lutea of the WT mice, indicating that these cells were no longer dividing (Fig. 6K). In contrast, most of the granulosa cells from growing small and antral follicles did not express CDKN1B (Fig. 6, K–O). In the WT animals, eCG stimulation did not induce the expression of CDKN1B in granulosa and cumulus cells (Fig. 6M). However, in Irs2 null ovaries, some positive staining was observed from the stage of small antral follicles (Fig. 6Q). Moreover, 46 h after eCG stimulation, CDKN1B staining was observed in cumulus cells and in mural and thecal cells in the large antral follicles (Fig. 6R). After superovulation of the WT mice, no CDKN1B staining was seen in the cumulus cells and the majority of the mural granulosa cells (Fig. 6, N and O). This is consistent with the high level of cyclin D2 expression seen 8.5 h after hCG treatment in the cumulus cells in the preovulatory follicles of the WT mice (Fig. 6, D and E). In contrast, in Irs2 null ovaries, the granulosa cells showed positive staining for CDKN1B 46 h after eCG injection (Fig. 6R). After administration of hCG, positive staining for CDKN1B was seen in the cumulus cells, suggesting that proliferation was arrested in these cells (Fig. 6, S and T).
Insulin signaling pathways in the CNS have been implicated in the regulation of female reproductive function [2, 10]. Insulin receptor substrate proteins 1 and 2 are the major downstream effector molecules recruited by the activated insulin receptor [10]. Female Irs2 null mice are infertile, with defects in hypothalamic and ovarian functions, which suggest that IRS2 mediates the effects of the insulin receptor on reproductive function in a number of tissues [12]. Furthermore, since Irs2 null mice display defects in leptin action, IRS2 may integrate the actions of insulin and leptin on energy homeostasis and fertility [12]. Surprisingly, NesCreIrs2KO mice, which lack Irs2 in all their neurons, displayed minimal reproductive abnormalities with almost normal hormone levels, ovarian functions, and fertility rates. There are several possible explanations for these findings. IRS1 may compensate in the CNS of NesCreIrs2KO mice, although we detected no alterations in the expression of Irs1 in the hypothalami of these animals. The timing of deletion of Irs2 is different in Irs2 null and NesCreIrs2KO mice, as the nestin transgene is expressed from E9 onwards, whereas Irs2 expression never occurs in the global null animal. Therefore subtle developmental differences may explain the phenotype. Taken together, these results and the demonstration of marked hyperphagia and obesity in NesCreIrs2KO mice suggest that CNS Irs2 expression is required for the effects of insulin on energy balance but not on reproductive function. Furthermore, we have demonstrated that leptin action in NesCreIrs2KO mice is normal, which suggests that IRS2 is not an obligatory point of convergence for the actions of insulin and leptin [1].
Irs2 was expressed in a number of ovarian cell types, with high levels in the follicular granulosa cells, which suggests that defective insulin and IGF1 actions in these cells may account largely for the infertility phenotype of Irs2 null mice. Analyses of follicular development and growth demonstrated a series of interrelated abnormalities in these animals. While early follicular development and oocyte development were not different between the WT and Irs2 null mice, once the follicles had reached the stage in which growth became gonadotropin-dependent there was a marked decrease in their numbers. There was a significant reduction in the number and size of antral follicles due to reduced antral cavity development and granulosa cell proliferation and an increase in follicular atresia. Normally, in primordial follicles, the oocyte is surrounded by a single layer of nondividing cells that are arrested in G0; these cells can be induced by appropriate stimuli to undergo proliferation. Our studies demonstrate that in primordial and small follicles, cell proliferation is largely preserved in Irs2 null ovaries. However, the rapid phase of follicular growth that occurs when granulosa cells acquire enhanced FSH and LH responsiveness and produce estrogen was abrogated in Irs2 null ovaries. Terminal differentiation (luteinization), which occurs when the LH surge causes granulosa cells in preovulatory follicles to exit the cell cycle, was also abnormal in the Irs2 null ovaries. Taken together, these findings demonstrate disruption of the co-ordinated growth pattern within follicles and loss of the bidirectional communication between oocytes and granulosa cells during follicle development.
The defects seen in our novel Irs2 null mouse model were not as severe as those described in the original knockout mouse. Differences in genetic background are likely to account for this finding, with the current model using mice on a C57Bl/6 background, while the earlier studies were performed in a mixed 129Sv/C57Bl/6 strain. Consistent with this observation are the findings that the metabolic phenotype of Irs2 null mice is strongly influenced by genetic background, and that the fecundity of C57Bl/6 mice is greater than that of 129Sv mice [23].
IRS2 lies downstream of both INSR and IGF1R and is able to recruit the MAP kinase and PI3 kinase signaling cascades, which have been implicated in ovarian function [6–8, 10]. However, we found that MAP kinase expression and phosphorylation in response to insulin or hCG were largely preserved in Irs2 null ovaries. Other signaling components, such as IRS1 and SHC, may be involved in the recruitment and activation of the MAP kinase in Irs2 null ovaries, although normal MAP kinase activity in response to these hormones is insufficient to preserve follicular function in these mice. In contrast, we found significant signaling defects in the PI3 kinase cascade in Irs2 null ovaries, with both insulin and hCG stimulation of AKT being impaired. Previous studies have implicated PI3 kinase/AKT signaling in the maintenance of the preovulatory follicle granulosa layers and have demonstrated that FSH induces a biphasic increase in AKT phosphorylation in rat granulosa cells [24]. We have shown that the metabolic effects of FSH in the mouse cumulus-oocyte complexes are mediated by the PI3 kinase pathway [25]. However, insulin appears to stimulate a larger increase in AKT serine phosphorylation than LH/hCG in the rat ovary in vivo [26]; our data from mouse ovaries are consistent with this observation.
A number of substrates of activated AKT that regulate cell survival and proliferation have been identified, including components of the apoptotic machinery, such as caspase 9 and BAD, which may be involved in IGF1-mediated protection of granulosa cell apoptosis [27, 28]. Members of the FOXO forkhead transcription factor subfamily (e.g., FOXO1a and FOXO3a) are also important substrates of AKT. Mice that lack Foxo3a display follicular activation and depletion of oocytes with secondary infertility, which suggests an important role for this protein in ovarian function [29]. AKT also phosphorylates GSK3B, which may be involved in the regulation of ovarian gene expression by FSH [21, 30]. Therefore, the major roles of FOXO and GSK3B as downstream effectors of AKT signaling involve the co-ordination of cell proliferation and differentiation in the ovary, and defects in these events in Irs2 null ovaries are likely to account for many of the observed abnormalities. The AKT/forkhead signaling cassette has been demonstrated to regulate directly cyclin D2 function in rat granulosa cells, and GSK3B has also been shown to regulate D-type cyclin function [31]. AKT/forkhead signaling also regulates CDKN1B function through both transcriptional and phosphorylation events. Deletion of cyclin D2 in mice results in impaired granulosa cell proliferation, impaired follicular growth, trapped oocytes in the corpora lutea, and ovulatory failure [32, 33]. Mice that lack CDKN1B have normal follicular growth but have impaired luteinization of granulosa cells [34]. In Irs2 null ovaries, we have demonstrated defects in the regulation of both cyclin D2 and CDKN1B due to impaired upstream AKT signaling, which may explain the functional abnormalities of ovarian cellular function in these mice. Reduced AKT activity may lead to the earlier expression of CDKN1B seen in the granulosa and cumulus cells of Irs2 null follicles. Disordered co-ordination of follicular and oocyte functions may also account for the frequent observation of trapped oocytes in Irs2 null follicles.
In summary, we have shown that CNS IRS2 signaling pathways are not required for female reproductive function. In contrast, signaling events downstream of IRS2 in the ovary play critical roles in follicular development and ovulation by regulating key components of the cell cycle apparatus involved in the co-ordination of cell proliferation and differentiation.
ACKNOWLEDGMENTS
We thank P. Smith and A. Parlow of the National Hormone and Pituitary Program and Pituitary Hormones and Antisera Centre for reagents.
FOOTNOTES
1Supported by an MRC Co-operative Component Grant and a Wellcome Trust Functional Genomics Award. 
Correspondence: 2Dominic J. Withers, Centre for Diabetes and Endocrinology, Rayne Institute, 5 University Street, University College London, London WC1E 6JJ, United Kingdom. FAX: 4420 767 96583; e-mail: d.withers{at}ucl.ac.uk
Received: 7 December 2006.
First decision: 23 December 2006.
Accepted: 19 February 2007.
REFERENCES
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