Mol. Hum. Reprod. Advance Access originally published online on March 9, 2007
Molecular Human Reproduction 2007 13(5):287-297; doi:10.1093/molehr/gam008
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Normalization of hormonal imbalances, ovarian follicular dynamics and metabolic effects in follitrophin receptor knockout mice
1 Molecular Reproduction Research Laboratory, Clinical Research Institute of Montreal, (Affiliated to Université de Montréal), Québec, Canada
2 To whom correspondence should be addressed at: Molecular Reproduction Research Laboratory, Clinical Research Institute of Montreal, 110 Pine Avenue West, Montreal (Québec), H2W 1R7, Canada. E-mail: sairamm{at}ircm.qc.ca
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Abstract |
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Genetically modified follitrophin receptor knockout female mice with total FSH-receptor (FSH-R) deletion are sterile and their combined estrogen deficiency-hyperandrogenemic status provides an experimental paradigm to study the effect of hormonal imbalances on ovarian function and metabolic alterations. Elevated LH levels causing hyperandrogenemia perturb normal folliculogenesis. To control diverse pathophysiology associated with hormonal imbalances, we investigated the effects of transplanting a single normal mouse ovary in young mutants. An intact FSH-R signalling system in the graft responded promptly to the up-regulated pituitary gonadotrophins circulating in the host mutant. Resumption of regular estrous cycles validated stimulation of uterine functions. Secretions from the viable functioning grafts partially corrected follicular abnormalities originally present in host ovaries. Stromal hyperplasia responsible for high ovarian LH-receptor and key enzymes in host thecal/interstitial complex and hyperandrogenemia was reduced in host ovaries. Increases in plasma estradiol and reduced LH and free testosterone re-established the negative-feedback system. Reduced android obesity and activation of mammary glands indicated the combined beneficial effects of normalized steroid hormones on target organs. These data provide evidence that ovarian transplantation in mutants corrects estrogen loss and hyperandrogenemia. However, correction of hormonal imbalances is not sufficient to fully restore effects of FSH-R loss in host granulosa cells.
Key words: androgen receptor/hyperandrogenemia/obesity/oocyte/stromal hyperplasia
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionSupplementary dataAcknowledgementReferences |
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The gradual cessation of ovarian function leading up to menopause causes estrogen loss during post-menopausal life, although small but insufficient amounts are still generated at peripheral sites. Differing hormonal profiles have a major impact on healthy ageing of women and quality of life. During reproductive life, ovarian steroid biosynthesis is gonadotrophin-dependent and occurs in thecal and granulosa cell compartments. In the menopausal ovary, follicles undergo atresia, with sparing of androgen-producing theca–interstitial cell components (Shifren and Schiff, 2000). Post-menopausal ovaries are smaller and consist primarily of stromal cells, which retain receptors for LH (Foth and Romer, 2001), that could respond to its rise in circulation secreting testosterone but produce little estrogen (Longcope, 2001; Krug and Berga, 2002). Advancing age coupled with estrogen loss and hyperandrogenemia could lead to insulin resistance, increasing the risks of type II diabetes, hyperlipidemia and cardiovascular disease.
Age-related changes occurring in the reproductive system that affect oocyte quality in middle-aged women are accompanied or preceded by dysfunction of the hypothalamic-pituitary-gonadal axis. We have previously investigated some of these phenomena by targeted disruption of the FSH-Rs that are predominantly expressed in ovarian granulosa cells. In the resulting follitrophin receptor knockout (FORKO) mouse, lack of FSH-R signalling produces ovarian failure resulting in loss of cycles, infertility, estrogen deprivation and structural alterations of the uterus and vagina in female mice as well as other changes in the peripheral systems (Dierich et al., 1998; Danilovich et al., 2000). The lack of estrogen causes a loss of the normal feedback resulting in high pituitary FSH and LH secretion at an early age in null mutants and upon ageing in heterozygous females. The high androgen also induces associated pathophysiological conditions (Danilovich et al., 2000, 2001, 2004). In addition to infertility or early reproductive senescence, these mice exhibit obesity, skeletal abnormality, increased ovarian/uterine tomour incidences, cardiovascular disturbances as well as age-related changes in the central and peripheral nervous systems (Danilovich et al., 2004). We therefore hypothesized that simple organ replacement in the mutants could simulate balanced hormone therapy. Our intent in undertaking this study was 2-fold. First, we wished to explore the behaviour of a normal transplanted ovary when placed in an abnormal hormonal milieu that resembles hyperandrogenemia. Secondly, we asked if such a tissue would continue to function and correct some or all of the abnormalities in the host ovaries and impact on diverse peripheral phenotypes that are manifest in the mutants. Evidence presented here show that both these expectations were partly fulfilled. Correction of prevailing hormonal imbalances and resumption of estrous cycles, beneficial changes in host ovarian structures, normalization of androgens, and amelioration of adiposity indicate restoration of the hypothalamo-pituitary-gonadal axis regulation. These studies could help extend evaluation of replacement therapy after lengthy deprivation or imbalance of sex hormones in females. The study also provides impetus to the consideration of replacement strategies such as the introduction of specific FSH-R forms in granulosa cells of mutant ovaries and their capacity to restore ovarian function.
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Materials and methods |
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Experimental animals
All studies were performed according to the guidelines of the institutional animal care committee. The FORKO mice were obtained (Dierich et al., 1998) by breeding 129T2/SV EmsJ FSH-R male and females of 3–5 months. These provided littermate + / + and – / – females for direct comparison. Animals were housed under controlled temperature and constant light with food and water provided ad libitum. They were genotyped by PCR (Danilovich et al., 2000; Yang et al., 2003). Four-week-old wild-type (WT) females (after the first cycle) were used as ovarian donors for transplantation in 42–45 days old FORKO mutant females. The transplanted (KO-TR) and sham operated (KO-sham) groups had 30 and 25 mice, respectively. Other WT females (42–45 days, n = 21) without surgery served as positive controls.
Transplantation procedure
Ovary collection and transplantations were done simultaneously. WT ovaries were cleaned and kept in fresh sterilized PBS at room temperature till transplantation. Mice were anaesthetized by Pr AErrane [(Isoflurane, USP), Baxter, Toronto, ON, Canada]. Through a small dorso-median transverse incision, one ovary was placed subcutaneously on the left dorso-lateral side of KO-TR mice. During surgery, mice were kept on a warming plate (
37°C) covered with sterile towel. After transplantation the skin incision was sutured with nylon. In KO-sham mice, a similar mass of adipose tissue was placed instead of WT ovaries. After surgery, animals were allowed to recover for at least 7 days. Vaginal smears were then checked for sign of estrous cycles; subsequently, cycle length was calculated by following the duration of each stage.
Collection of samples
KO-TR and WT mice were killed on the morning of proestrous. Acyclic KO-sham mice were killed on the same day. For plasma steroid hormones, blood samples were collected by cardiac puncture using heparin-coated syringes and placed in plastic tubes. Following centrifugation plasma was stored at – 20°C. Grafted and host ovaries and uteri were dissected and weighed. Visceral adipose tissue was weighed and preserved for histology. Grafted ovaries were preserved in 10% formalin for histology. Ovaries in pairs and uterine horns from each WT, KO-TR, and KO-sham groups were snap frozen and stored at – 80°C until extraction for western blotting (WB) and RT–PCR studies. At necropsy, both sides of the abdominal mammary glands (number 4 and 9) from each female were removed for evaluation. The whole-mount preparation protocol was a slight modification of a procedure described by Russo et al. (1989) utilizing acetone for delipidation and toludine blue for coloration. KO-sham mammary glands required longer acetone treatment to remove excess fat surrounding the atrophied mammary tissue.
Steroid hormone radioimmunoassays and gonadotrophin immunoradiometric assay
Immunoassays were performed according to the manufacturer's instructions. The estradiol-17ß and free testosterone (FT) radioimmunoassays of blood plasma samples were performed using Coat-A-Count kits (Diagnostic Products Corp., Los Angeles, CA, USA) with sensitivity of 8 pg ml– 1 and 0.15 pg ml– 1, the intra-assay coefficient of variation (CV) for the radioimmunoassays averaged 6.8% and 5.7%, respectively. The LH immunoradiometric assay (IRMA) of blood plasma samples was performed using Coat-A-Count kit (Diagnostic Products Corp.) with sensitivity of 0.15 mIU ml– 1 and intra-assay CV averaged 11.1%.
Immunohistochemistry and WB
Antibodies: androgen receptor (AR) (N20); cyclin-D2 (Cyc D2), (M-20) and Actin (I-19-R) were rabbit polyclonal IgGs, from Santa Cruz Biotechnology (SantaCruz, CA, USA). Rabbit anti-bovine P450c17 antiserum was kindly donated by Dr A.J.Conley (University of California Davis). The LH-R monoclonal antibody (P1B4) was a gift from Dr J.Wimalasena (Department of Obstetrics and Gynecology, University of Tennessee, Knoxville, TN, USA). Estrogen receptor 
and ß (ER and ERß) rabbit polyclonal IgG were donated by Dr P.Chambon (IGBMC, Strasbourg, France). Specific Zona Pellucida (ZP-A) (ZP2), ZP-B (ZP1) and ZP-C (ZP3) antibodies were kindly provided by Dr U.Eberspaecher (Schering AG Berlin, Germany). Secondary biotinylated goat anti-rabbit and goat anti-mouse were used for rabbit polyclonal and mouse monoclonal antibodies, respectively (ImmunoCruz Staining System, Santa Cruz Biotechnology).
Formalin-fixed, paraffin-embedded sections were used for IHC; sections were rehydrated and exposed to antigen unmasking using citrate buffer (10 mM, pH 6.0). Peroxidase activity was quenched with 3% H2O2. Sections were incubated overnight with the following specific antisera: P450c17, LH-R, Cyclin-D2, AR, ER, ERß, ZP-A, ZP-B and ZP-C at a dilution of 1:5000, 1:1000, 1:50, 1:100, 1:100, 1:100, 1:500, 1:200 and 1:1000, respectively, (except for anti-Cyc D2, ER and ERß, which were incubated for two nights) at 4°C. In negative controls, normal serum was used in the first reaction. The following day, sections were washed and incubated with biotinylated secondary antibody. Signals were amplified with avidin-biotinylated horse-radish peroxidase developed with diamino-benzidine (DAB), counterstained with Mayer's hematoxylin and dehydrated again. Sections were analysed under a light microscope. Immunohistochemistry (IHC) intensity was visually scored from undetectable to strong signal.
For WB, pairs of frozen ovarian tissues were homogenized in lysis buffer (RIPA) containing detergent and protease inhibitor cocktail (20 mM Tris, pH 7.5; 150 mM NaCl; 0.1% Nonidet P-40; 0.5% Sodium deoxycholate; 1 mM EDTA; 0.1% SDS; 0.1 mM phenylmethylsulfonyl fluoride and 5 µg ml– 1 leupeptin). Thirty micrograms of whole tissue proteins were run on sodium dodeyl sulphate-polyacrylamide electrophoresis gels and transferred to the polyvinylidene difluoride membrane for reaction with AR antibody (1:1000). The same membrane was later used to detect actin. After treatment of the blots with 1:10000 dilution of second antibody (HRP- conjugated goat anti-rabbit, Santa Cruz, CA, USA), bands were detected by ECL kit (Amersham Pharmacia Biotech). Band intensities were compared by densitometry.
Reverse transcription–polymerase chain reaction
RNA was extracted from frozen whole ovaries using TRIzol Reagent (Invitrogen Corp.) according to the manufacturer's instructions. RNA samples were then DNase-treated and subjected to semiquantitative RT–PCR previously described by our laboratory (Danilovich et al., 2000). Forward and reverse primers for amplifying AR (251 bp) mRNA were AAT GAG TAC CGC ATG CAC AA (forward) and CTT GAG CAG GAT GTG GGA TT (reverse), based on GenBank sequences (Accession NM_013476
[GenBank]
), and for insulin-like growth factor-I (IGF-1) (250 bp) were ATG AAG CAA AAT GTG CGT TG (forward) and ACC AAA GTC CCT TCG GTG AT (reverse). Each test sample was also simultaneously verified for amplification of actin (243 bp) as an internal control under identical conditions, using primers CCA GAT CAT GTT TGA GAC CTT C (forward) and AGG ATC TTC ATG AGG TAG TCT G (reverse). All RT–PCR results were confirmed by performing Q-PCR.
Statistical analysis
RT–PCR, IHC and WB experiments were done three times. All data were expressed as mean ± SEM and were analysed by one-way ANOVA. A value of P < 0.05 was considered to be statistically significant.
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Results |
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Validation of estrous cycles
The estrous cycle occurring at regular intervals in rodents is an effective indicator of feedback regulation and balanced steroid hormonal effects on reproductive system accessories. Due to the lack of circulating estrogen, null mice never experience sexual maturity and cyclicity is absent. However, with a transplanted normal ovary, 98% KO-TR mice resumed cyclic changes in the vagina as early as 9–10 days post-surgery. Failure in one mouse might be related to surgery. Cornified epithelial cells (data not shown) were distinguished in the vaginal smears at an interval of 5–6 days.
Survival of grafted ovaries and their function in mutant host
Ovarian grafts were distinguishable from the surrounding host subcutaneous (s.c.) tissue; their size did not change after 1 month. Histology of grafted ovaries revealed primordial, primary, secondary follicles and corpora lutea (CLs) (Figure 1A). Antral follicular growth, ovulations and CL formation (Figure 1B) ensued by 3–4 months. The grafted ovaries were functional, as shown by the resumption of estrous cycles in the KO-TR mice (data not shown). We assessed Cyclin D2 (Cyc D2), which is linked to granulosa cell (GC) maturation and function (Sicinski et al., 1996; Robker and Richards, 1998). Expression of Cyc D2 protein indicated that the grafted ovaries remained functional even 3 months after transplantation (Figure 1C and D).
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Effects in host ovaries
We examined the temporal and qualitative effects of ovarian transplantation (OT) on host ovaries with particular interest in the degree of reinstatement of normalcy. KO-TR and KO-sham ovaries were similar in size, and ovarian weights remained unchanged up to 3 months (
1 mg). Most remaining studies in this report show data for this period. Altered host ovarian histology after transplantation was indicated by an apparent reduction in previously abundant interstitial cells and stromal hyperplasia (Figure 1I) (note the atrophic KO-sham ovary, Figure 1G*); presence of some primordial, primary and smaller and bigger secondary follicles was apparent. However, in comparison to WT (Figure 1E) that had CL, maturation of the largest follicle in the host ovary remained arrested before the pre-antral stage.
Response of the uterus
As indicated, the resumption of estrous cycle indicated uterine and vaginal responses in the KO-TR. The previously atrophied uterus (weighing 90% less than WT) was stimulated by reinstated endogenous hormones, and weight increased by 44% over the KO-sham. Uterine horns had less fat tissue unlike KO-sham, which looked pale and thread-like, buried in mass of adipose tissue. Vascularization in the KO-TR uterus was similar to WT. The uterine histology of KO-TR revealed thick myometrial and stromal layers, increased number of endometrial glands and multiple layers in luminal epithelial cells similar to WT (Figure 1F and J); glandular elements of the endometrium were, however, less complex in the KO-TR. The KO-sham uterus lacked all these features following sustained estrogen deprivation (Figure 1H).
Alteration in stromal/interstitial cells
To investigate ovarian LH-R expression and its compartmentalization, we used a specific monoclonal antibody (P1B4) (Indrapichate et al., 1992). The IHC of ovaries revealed dominant LH-R expression in KO-sham (Figure 2B) thecal/interstitial compartments; LH-R was also present in the periphery of blood vessels (Figure 2B), but GCs were negative. In WT ovaries (Figure 2A) thecal/interstitial cells expressed low LH-R, whereas in luteinized cells within the mature follicle, expression was high, indicating the combined effects of FSH-R and LH-R (Figure 2A, CL-*). However, in KO-TR mice only partial correction of the ovaries was evident from a slight reduction in thecal and interstitial cells, but GCs in FSH-R null host ovaries continued to lack LH-R and, as a result, could not undergo luteinization (Figure 2C). Thecal/interstitial cells are major sites of androgen biosynthesis in the ovary regulated by circulating LH. Cytochrome P450c17 (17-hydroxylase/C17-20-lyase, CYP 17) in these cells is the LH-responsive steroidogenic enzyme vital for androgen synthesis (Bogovich and Richards, 1982). Expression of P450c17 was highest in the thecal/interstitial compartment in KO-sham ovaries (Figure 2E, arrow). In the KO-TR ovaries, it was reduced (Figure 2D and F) to the same level as in WT, indicating moderations in LH action in FSH-R null host ovaries. These observations are in accordance with reductions in circulating androgen levels in the host (see below).
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As androgens also exert a direct effect on ovarian function via the AR (Hild-Petito et al., 1991), we examined its expression and cellular localization. There was no change in total AR mRNA in the ovary from all three groups (Figure 3A). However, WB analysis of whole ovaries revealed quantitative differences in AR expression. Interestingly, the total amount of AR protein in KO-sham ovaries that was 3X the WT was not altered in KO-TR ovaries at 3 months (Figure 3B), suggesting that up-regulation in the KO-sham occurred at the translational level. However, IHC revealed AR redistribution in KO-sham and KO-TR according to follicular growth. AR confined to growing follicles in WT (Figure 3C.1) was highly expressed in thecal, interstitial and stromal cells in the KO-sham group (Figure 3C.2). Cellular redistribution in AR expression was apparent in KO-TR ovaries (Figure 3C.3) due to increased folliculogenesis from primordial to primary and secondary follicles and reduction in the thecal/interstitial compartments, stromal cells and atretic follicles. As IGF-I is a potent stimulator of thecal/interstitial cell proliferation, leading to an increase in both the number and the proportion of steroidogenically active thecal/interstitial cells (Duleba et al., 1997), we assessed its expression. IGF-1 mRNA was 3-fold higher in KO-sham ovaries in comparison with age-matched WT, but it was reduced to 1.5-fold in KO-TR ovaries after 3 months of transplantation (P < 0.05) (Supplementary data, Figure S1). Tests on IGF-1 receptor mRNA revealed no change (not shown).
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Zona pellucida glycoprotein expression of grafted/host oocytes
For a single oocyte to grow and attain functional maturity within the follicle, optimal bidirectional communication with the surrounding nurturing cumulus and mural GCs is critical (Eppig et al., 2002). As we have previously shown aberrations in the organization and differential expression of all three ZP glycoproteins that envelop the oocyte cytoplasm (Yang et al., 2003) in FORKO ovaries, it was of interest to examine the extent of repair of these functions at different times. With antibodies currently available to us, we could study the expression of ZP-A and C glycoproteins (but not B, as the aged antibody had lost its antigen recognition) in WT, KO-sham and KO-TR oocytes. Ooplasm of 1-month graft ovaries in a FORKO environment expressed more ZP-A as compared with 3-month graft (Figure 4A and B). ZP-C was highly expressed in 3 months grafts (Figure 4C and D); some cumulus cells were also positive for ZP-C. In comparison to WT oocytes, the periphery of KO-sham oocytes were not smooth and thickness of ZP varied in both groups (5.0 ± 0.3 µm and 3.0 ± 0.1 µm, respectively in secondary follicles, P < 0.003). Grafting a normal ovary induced beneficial changes in the oocytes of KO-TR ovaries (Figure 4G, arrow). Healthy follicles were evident with well growing oocytes (thickness of secondary follicles, 4.0 ± 0.2 µm, P < 0.002). In KO-TR oocytes, ZP-A expression appeared in both ZP and ooplasm (Figure 4G) as compared with ZP-A in ooplasm of KO-sham ovaries (Figure 4F). The visual appearance of ZP-A expression in the KO-TR was similar to WT. In KO-sham oocytes, ZP-C expression, unlike ZP-A, increased in both ooplasm and ZP (Figure 4I) whereas in WT it was restricted to the thin ring of ZP. In KO-TR host ovaries, expression patterns after 3 months became nearly normal, resembling WT, but remained slightly higher (Figure 4J).
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Estrogenic target actions on host ovary and uterus
Estrogen action in target organs is mediated by at least two nuclear receptors ER
and ERß (Couse et al., 1997; Nilsson et al., 1998). Reasoning that sustained estrogen deficiency and androgen dominance might impact ER expression patterns in FORKO ovaries and uteri, we assessed receptors by IHC in both tissues. In comparison to KO-sham, ERß was dominant in KO-TR and WT GCs and a faint staining was also observed in oocytes (Supplementary data, Figure S2A–C; arrows). ER increased in KO-TR uterus and was comparable to WT; nuclear staining was present in endometrial glands and the myometrium (Figure S2D and F), reflecting the stages of the cycle in KO-TR mice. In the KO-sham group, some cytoplasmic staining was seen in the luminal epithelium, indicating diestrus (Figure S2E).
Adjustment of hormonal imbalances in FORKOs
As the data noted above revealed marked improvement in KO-TR target tissues, it was important to know if the imbalances in circulating gonadotrophins and steroid hormonal levels that had prevailed in mutants were also corrected (Table 1). Although both gonadotrophins rise in circulation in FORKOs, we concentrated on assessing LH, as it is responsible for the prevailing hyperandrogenemia (Danilovich et al., 2000; Foth and Romer, 2001). In comparison with KO-sham, LH levels at 3 months post-surgery decreased in the KO-TR group. Aberrations in two major ovarian hormones, estradiol (E2) and FT in blood were also corrected. As measured by radioimmunoassays, estradiol levels in KO-sham were barely detectable. It increased in the KO-TR group after 3 months of transplantation, although it was not equivalent to WT, but the changes were statistically significant compared to KO-sham. Hyperandrogenemia was clearly evident in FORKO mice as FT was high in KO-sham. After 3-months of ovarian grafting, androgen abnormality was abolished in KO-TR females.
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Reduction in visceral adiposity and activation of mammary glands
One of the non-gonadal phenotypes readily obvious in FORKOs is increased adiposity (Danilovich et al., 2000). KO-sham females were 9% heavier than WT and this was reduced to 2% in KO-TR mice in 3 months. All KO-sham mice had more abdominal fat; periuterine fat depot in comparison to WT females was 62% higher. This difference was eliminated in the KO-TR group (Figure 5 top). Histology in WT showed differentiating small (Figure 5A), medium and large adipocytes accompanied with blood vessels. In the KO-sham group, large adipocytes (Figure 5B2) were predominant with moderate medium (Figure 5B1) and relatively fewer small adipocytes indicating effects on cell remodelling. These patterns appeared to be reversed in the KO-TR group; fat tissue was now repopulated by many small adipocyte clusters (Figure 5C1) including the presence of medium sized cells and apparent reduction of large adipocytes (Figure 5C2). Thus the enduring effect of a normal transplanted ovary on abdominal adipocytes was evident.
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Similar effects were also seen in mammary glands fat pads. More adipose tissue surrounded the atrophic mammary gland in KO-sham (not shown), and prolonged solvent treatment was required to reveal glandular structures. Whole mount of glands shown in Figure 5 (D–F) revealed that correction of hormonal deficiency/imbalances in KO-TR was effective in reinstating sufficient duct and terminal end bud (TEB) development.
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Discussion |
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Reproductive success in females requires successful hormone signalling via specific FSH-R localized in ovarian GCs and two studies have demonstrated the loss of fertility in receptor knockout mice (Dierich et al., 1998; Abel et al., 2000; Danilovich et al., 2000). Other phenotypes related to sex hormonal imbalances render these mutants valuable for addressing additional issues of importance to women's health (Danilovich et al., 2004). Although FSH-R expression is predominant in ovarian GCs (Simoni et al., 1997; Dierich et al., 1998), emerging reports note expression at other sites including the oocyte (Patsoula et al., 2001), fallopian tube (Zheng et al., 1996), uterus (Shemesh, 2001) and bone osteoclasts (Sun et al., 2006). The question therefore arises whether various phenotypic abnormalities observed in FORKO mice could be reversed by restoration of tissue-selective FSH-R signalling. The expression of different FSH-R variants from a single large gene (Babu et al., 2000) complicates the design of useful and site-specific replacements. The testing of a transplant approach as in the current study, to our knowledge, is the first report showing that simple transplantation of a normal WT immature ovary (with an intact FSH-R system) into a hormonally abnormal environment of androgen dominance is effective. The graft continued its maturation and function, re-established normal circulating hormonal profiles in the host and partially corrected abnormalities associated with ovarian FSH-R loss. The mechanistic events in FORKOs before (red) and after (green) transplantation are depicted in Figure 6.
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Successful transplantation of a single WT ovary into a young mutant that already had acquired high LH and FSH levels and increasing androgen at 1 month (Balla et al., 2003) demonstrated that both gonadotrophins secreted in mutants were functional, stimulating normal ovarian events in the graft. Resumption of estrous cycles in previously acyclic mutants provides proof that a correct sex hormonal milieu is conducive for normal functioning of the once atrophic host uterine and vaginal musculature and epithelia (Figure 1F, H and J). Ovulation and functional CL continued in grafts. Although favourable effects on host ovaries were evident following the normalized circulating hormone levels (Table 1), the absence of FSH-R in the GCs precluded their complete development, indicating that this requirement for final follicular maturation cannot be surmounted despite amelioration of atresia. Lack of LH-R expression in the GCs of these host ovaries is in accord with observation that FSH-R action is a prerequisite for this induction (Danilovich et al., 2002) and steroidal correction from the graft was not a substitute for it.
Interstitial cells and stromal hyperplasia in FORKO (Figure 1I, asterisk) indicate that elevated LH in mutants contributed to high androgen, causing pathophysiological conditions (Givens et al., 1971; Danilovich et al., 2004). LH regulates thecal cell steroidogenesis by stimulating LH-R expression and activating enzymes including CYP11A, 3ß-hydroxysteroid dehydrogenase (3 ß-HSD) and CYP 17 (Magoffin, 2005). Reduction of LH-R and CYP 17 in KO-TR ovaries contributed to the normalizing androgen status (Figure 2A–F). Excessive androgen production is an end result of many factors, including high IGF-1, which causes a disturbance in the intra-ovarian androgen/estrogen balance and atresia of multiple small follicles in the presence of high FSH (Homburg et al., 1992). Our finding on high expression of IGF-1 mRNA in FORKO ovaries is consistent with these observations (Figure S1); however, IGF-1 receptor status remained unchanged. Partial but significant correction of IGF-1 expression after transplantation indicates that other GC signalling events in addition to a balanced hormonal milieu must be involved in normal IGF-I gene regulation.
High androgens in FORKO sham mice are reminiscent of hyperandrogenemia in women (Judd and Korenman, 1982). In addition to being a precursor for estrogen, androgens act via ARs at two stages in follicular development. Initially, androgen increases recruitment of primordial follicles into the growing pool of developing follicles (Vendola et al., 1999) and FSH induces AR in primary but not mature follicles (Weil et al., 1999). As follicles mature, estrogen levels increase, whereas AR decreases in GCs. GCs express ER-ß (Mowa and Iwanaga, 2000) and AR is repressed by the activation of ER-ß in late antral follicles. The change in the oocyte microenvironment from androgen dominance to estrogen allows progression. Such events, absent in ER-ß mutants, promote follicular atresia in the presence of excess androgen (Cheng et al., 2002). FORKO ovaries are estrogen-deficient and the few follicles that grow do not progress beyond the pre-antral stage, stimulating atresia and AR elevates in the ovary. In addition to aberrant steroidogenesis, the appearance and quality of oocytes are compromised in FORKO mutants (Yang et al., 2003) (Figure 4). After 3 months, oocytes in the grafted ovaries continued to be normal in appearance with expression of zona glycoproteins (Figure 4). However, we cannot say if initial exposure of the graft ovary to a damaging host environment compromised the functional quality of oocytes, as we did not design experiments to verify this. Nevertheless, by 3 months, secretion from grafts was sufficient to repair, in part, previously affected follicular structures within the host, including partial correction in expression of two zona glycoproteins (Figure 4G and J). Although ZP-A deficit and over expression of ZP-C of KO-sham (Figure 4) improve qualitatively in KO-TR ovaries, we do not know if balances between transport and synthesis are restored. Our findings are consistent with data indicating immunoreactive material in both the ooplasm and ZP (Martinez et al., 1996; Eberspaecher et al., 2001). ZP-C is a major component responsible for the initial sperm-oocyte recognition in mouse oocytes. Mice without ZP-C do not shape a ZP matrix at all, even early in oogenesis (Rankin et al., 2001); on the other hand, over expression of ZP-C in KO-sham oocytes (Yang et al., 2003; fig 4I) are insufficient to compensate for deficits in FORKO females. Based on evidence that testosterone stimulates ZP-C protein by gene transcription (Pan et al., 2001) it is tempting to suggest a similar high androgen involvement in the FORKO ovary.
FSH promotes cell division in part by enhancing Cyc D2 transcription in GCs (Sicinski et al., 1996; Burns et al., 2001). High circulating FSH and LH provided by host mice induced Cyc D2 in grafts, stimulating folliculogenesis, ovulation and steroidogenic processes to re-establish negative-feedback mechanisms in host mice. Elevated Cyc D2 found in KO-TR ovaries supports arguments for the direct role of estrogen in its regulation (Burns et al., 2001) and in inducing a limited proliferative effect on host ovarian follicles (data not shown). Our study has also revealed that the relative abundance of genes for example of ER in the uterus and ERß in the ovary is generally in accordance with other reports (Couse et al., 1997; Nilsson et al., 1998). Up regulation of ERß protein in GCs of host ovarian follicles and similar upward changes for ER in the host uterus following transplantation (Supplementary data, Figure S1) provided proof that the steroidal sensitivity of ovarian GCs and uterine epithelium in FORKOs were retained at that age. Based on the observation that all uterine compartments of KO-TR mice underwent adequate stimulation in a host that lacked the FSH-R gene, we can conclude that FSH-R, if present in the uterus (Shemesh, 2001), is not required for this function.
Observations on the adipose tissue assume metabolic significance as they indicate the direct impact of sex hormonal imbalances. An effect of the menopause or hormonal imbalances in PCO women is weight gain and fat redistribution. The abdominal fat mass distribution of KO-sham mice parallels this situation. In a previous study, estrogen replacement reduced excess adipose tissue in FORKO mice, indicating metabolic effects (Sairam et al., 2002). Similarly, grafted ovarian hormones also reduced abdominal adipose tissue in KO-TR mice (Figure 5). Interestingly, normalized circulating hormones could also correct previous developmental abnormalities in the proportions or size of adipocytes. In view of evidence that larger adipocytes are less insulin sensitive (Olefsky and Reaven, 1975) and our recent findings on age-related glucose intolerance and signs of metabolic syndrome in FORKO mutants (Sairam et al., 2006), the current demonstration of the beneficial effect of steroids, as provided in a natural setting, on adipose tissue distribution is notable. Partial, structural and cellular changes noted in the mammary glands and the fat pads of KO-TR mice also demonstrate the combined beneficial effect of estrogen and progesterone (Figure 5).
The choice of the grafting site deserves comment. Although the kidney capsule and the ovarian bursa are two common grafting sites, capsular pressure on the graft limits the expansion of growing follicles (Gosden et al., 1994). Our studies agree with other reports on the use of S.C. space (Weissman et al., 1999) for adequate graft survival and full ovarian function, including ovulation. While our study was being completed, a report on ovarian transplants describing replacement effects in the LH-R knockout mice found that replacing WT ovarian pieces under the bursa of ovariectomized mutants resulted in birth of normal pups, which allowed the authors to argue the redundancy of LH-R expression at sites other than the ovary for maintaining normal fertility (Pakarinen et al., 2005). The intent and design of our study in this report precluded testing the fertility option in the KO-TR. [In other experiments we have placed small pieces of wild-type grafts in the ovarian bursa of mutants in order to restore fertility (data not shown). Cycling mice were caged with fertile males; despite the presence of vaginal plugs indicating cyclicity and restoration of sexual behaviour, fertility was not evident in mutants. It is presumed that the diminutive bursa was incapable of repair to clasp the graft in place and facilitate ovum transport for fertilization. No implantation sites were evident.].
In conclusion, we show the potential for reversal of abnormal endocrine and metabolic functions after OT in an abnormal hormonal milieu. We believe that this approach achieved a balance and re-established feedback relationships that operate under natural conditions (Figure 6) enhancing beneficial effects and reducing adversities. In view of current controversies on the benefits and risks of hormone replacement therapy (writing group for the women's health initiative investigators, Turgeon et al., 2004), we believe that the use of models such as that discussed here would help probe fundamental and mechanistic issues. In particular, consequences of long-term hormonal deprivation or imbalances on potential damage to brain, cardiovascular or bone systems show themselves to be amenable for reversal by transplantation approaches in a form that would mimic natural settings. Although such manoeuvres are not practical in menopausal women, issues related to risks and benefits after long-term deprivation as well as target cellular mechanisms can be studied using this paradigm in models.
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TopAbstractIntroductionMaterials and methodsResultsDiscussionSupplementary dataAcknowledgementReferences |
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Supplementary data are available at http://molehr.oxfordjournals.orgl/
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TopAbstractIntroductionMaterials and methodsResultsDiscussionSupplementary dataAcknowledgementReferences |
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This work was supported by grants from the Canadian Institute of Health Research. Authors are grateful to Drs P. Chambon, J. Wimalasena, A.J. Conley and U. Eberspaecher for providing antibodies used in the work. We also thank Mohini Ramkaran, Annie Vallée and Andre Turgeon for their help in various phases of this study.
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Submitted on October 30, 2006; accepted on January 24, 2007.
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