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Gamete Biology |
Department of Anatomy3
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Institute of Reproduction and Development,4 Monash University, Clayton, Victoria 3800, Australia
Prince Henry's Institute of Medical Research,5 Clayton, Victoria 3168, Australia
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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cumulus cells, estradiol, gamete biology, in vitro fertilization, oocyte development
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The Aromatase knockout (ArKO) mouse, which lacks the capacity to synthesize endogenous estrogen, provides an opportunity to explore the nature of the role of estrogen in folliculogenesis and oocyte development. Female ArKO mice are infertile, primarily as a result of anovulation. Elevated serum FSH, LH, and testosterone levels correlate with a progressive degeneration of the ovary and its structures, including cystic and hemorrhagic follicles, a gradual loss of secondary and antral follicles, interstitial cell hyperplasia, and increased collagen deposition [6]. In 12- to 14-wk-old ArKO mice, however, follicles up to the large antral stage have been observed, containing apparently healthy oocytes. This therefore presents the question as to whether the young ArKO ovary is capable of responding to a protocol of superovulation induction and whether the observed oocytes are capable of sustaining fertilization and subsequent development. In this current study, we have characterized ArKO follicle and oocyte developmental competence using the techniques of in vitro maturation, fertilization, and culture to blastocyst. Our results demonstrate that 1) estrogen is indirectly required for ovulation as a negative regulator of gonadotroph production and 2) estrogen is not essential to maintain reproductive fitness of immature mammalian oocytes.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Littermate ArKO, wild-type (WT), and heterozygote (Het) mice were maintained on soy-free mouse chow (GlenForrest Stockfeeders, Western Australia). Animals were genotyped as previously described [7]. Two age groups were utilized for each experiment: 4 (prepubertal) and 7 (adult) wk of age. Two replicates of the first experiment (superovulation) were performed with 10 mice per group for each mouse genotype and age. Three replicates of the second experiment (in vitro maturation, fertilization, and culture) were carried out, 10 mice per genotype per age group. All experiments were conducted with appropriate Monash Medical Centre Animal Ethics Committee approval.
Superovulation Protocol
Mice were superovulated with an intraperitoneal (i.p.) injection of 5 IU eCG (5 IU/ml; Folligon, Intervet, Castle Hill, NSW, Australia) in 0.1 ml saline (0.9% NaCl; British Drug House, Kilsyth, VIC, Australia), followed 28 h later with an i.p. injection of 5 IU hCG (5 IU/ml; Chorulon, Intervet) in 0.1 ml saline (0.9% NaCl; British Drug House). Twelve hours after hCG injection, mice were killed by cervical dislocation. Both ovaries were excised and fixed immediately. Both oviducts were also excised and placed in 35-mm culture dishes (Falcon; Becton Dickinson Labware, Franklin Lakes, NJ) containing 2–4 ml prewarmed Hepes-buffered (Calbiochem, La Jolla, CA) potassium simplex optimized medium supplemented with amino acids (KSOMaa) handling medium [8] + 3 mg/ml BSA (Pentex Crystallized; Miles, Kankakee, IL), and opened with watchmakers forceps to release the cumulus-oocyte complexes (COCs), which were visualized using a stereoscopic microscope (Leica Wild 32; Heerbrugg, Switzerland) fitted with a warm stage at 37°C to reduce temperature changes. The number of COCs present in the oviducts of each mouse was recorded.
Once it had been established that ArKO ovaries did not respond to our superovulation regimen, the protocol was modified to remove the hCG injection. Mice were killed by cervical dislocation 48 h after the injection of eCG; the ovaries were excised and placed in 35-mm culture dishes containing 2–4 ml prewarmed Hepes-KSOMaa. The ovarian capsule was removed by blunt dissection and the ovaries were systematically punctured with a 29-gauge 0.33 x 12.7-mm ultrafine needle to release the immature COCs, which were visualized under a stereoscopic microscope fitted with a warm stage at 37°C.
Media and Culture Conditions
Potassium simplex optimized medium supplemented with amino acids (KSOMaa) was employed for handling, maturing, and culturing ArKO oocytes/zygotes [8]. Modified Tyrode 6 (MT6) culture medium was employed to capacitate sperm [9]. The media utilized for all experiments were filter-sterilized using 0.22-µm filters (Millepore Corporation, Bedford, MA). Experiments were repeated using charcoal-stripped, phenol-red-free media to ensure all potential sources of estrogenic-like activity were removed.
Oocytes/zygotes were cultured in groups of 10–15 per 20-µl microdrop of culture medium contained in 60-mm culture dishes (Falcon; Becton Dickinson Labware), which was overlaid with 7 ml light mineral oil (Sigma, St. Louis, MO) to prevent medium evaporation and to reduce pH fluctuations. All prepared culture dishes were equilibrated in a humidified atmosphere of 5% CO2 in air at 37°C for 4 h before use (Heraeus CO2 incubator; Radiometer Pacific, Copenhagen, Denmark). All cultures were performed under equivalent conditions.
Preparation of Sperm
Proven stud heterozygote male mice from the ArKO colony were killed by cervical dislocation and their epididymides were placed in a 35-mm culture dish containing 2–4 ml prewarmed Hepes-KSOMaa. An incision was made in the middle of the epididymis with a 19-gauge needle x 
inch (Terumo, Elkton, MD). The epididymis was then placed in a loosely capped 5-ml tube containing equilibrated MT6 culture medium plus 3 mg/ ml BSA and incubated in a humidified atmosphere of 5% CO2 in air at 37°C for 2 h to allow the sperm to disperse and capacitate.
Oocyte Maturation Assessment
COCs were collected using a hand-drawn sterile glass pipette (Chase Instruments, Glen Falls, NY), briefly incubated in Hepes-KSOMaa containing 50 IU hyaluronidase (Sigma), and gently aspirated up and down to remove the cumulus cells. The oocytes were then rinsed in Hepes- KSOMaa to visualize the maturation status of the oocyte. The appearance of a nucleus within the cytoplasm of the oocyte in the absence of a polar body was used to indicate the germinal vesicle stage (prophase I) of development; the absence of a nucleus and a polar body identified germinal vesicle breakdown/metaphase I; and the presence of a polar body in the perivitelline space indicated metaphase II. Immature COCs were transferred to 60-mm culture dishes containing KSOMaa culture media supplemented with 10% fetal calf serum (Gibco BRL, Life Technologies Ltd., Auckland, New Zealand), 0.28% eCG, and 0.06% hCG and incubated as described above for 18 h. Maturation status was assessed at 6, 12, and 18 h.
Only the oocytes deemed mature were utilized for in vitro fertilization. Mature oocytes were rinsed in Hepes-KSOMaa and then MT6 before being transferred to 60-mm culture dishes containing microdrops of medium containing capacitated sperm. Spontaneous fertilization was allowed to progress for 2 h under the culture conditions described above. Oocytes were then rinsed and transferred to preequilibrated KSOMaa in 60-mm culture dishes and cultured for a further 4 h. Successful fertilization was assessed by the appearance of two pronuclei and the extrusion of a second polar body into the perivitelline space. Those oocytes failing to fertilize were removed from culture, leaving only zygotes in the culture dish. Zygotes were cultured for a further 4 days, at which time development to blastocyst was assessed and recorded.
Staining for Nuclear Chromatin
Approximately 30% of each type of oocyte recognized as immature, in vitro matured, and zygotes were randomly selected and stained in 2–3 ml Hoechst 33342 bisbenzimide (Sigma) overnight. Groups of 1–3 of these oocytes/zygotes were then transferred to a glass slide containing a minute drop of light mineral oil and covered with a 22- x 22-mm glass cover slip (Menzel-glaser; Saarbrockener, Braunschweigh, Germany). Chromatin was visualized using a fluorescence microscope fitted with an ultraviolet lamp and excitation fluorescence 460-nm blue filter (Leitz; Fluovert FU, Wetlar, Germany). The maturation/fertilization status of the oocytes was assessed based on the appearance of the chromatin within the germinal vesicle of the immature oocyte, the metaphase plate and polar body of the mature oocyte and within the pronuclei and second polar body of the fertilized oocytes.
Ovarian Histology
Ovaries were fixed in 10% formalin (pH 7.6) for 24 h at room temperature [10], then transferred to plastic cassettes (Tissue-Tek, Miles, Elkhart, IN) and processed manually by washing with saline followed by dehydration through graded ethanol steps to a 1-h incubation in heated (45°C) paraffin wax and embedding in paraffin. The paraffin blocks were immersed in mollifex (BDH Laboratory Supplies, Poole, England) for 1–2 min, and then placed on ice for 2–3 min before sectioning at 3 µm (Leica Wild 2030). Sections mounted on glass slides were dried overnight at 37°C. Sections were stained using a modified Masson Trichrome stain [11] and cover slipped using DPX (BDH Lab Supplies, Poole, England).
Statistics
Data were analyzed statistically by analysis of variance (ANOVA) followed by Tukey-Kramer post tests for multiple comparisons. Bartlett test was used to check for homogeneity of variances. A P value of less than 0.05 was considered to be biologically significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cumulus-oocyte complexes (COCs) were not recovered from the oviducts of 4- or 7-wk-old ArKO mice following exogenous administration of eCG and hCG, whereas both WT and Het mice superovulated as expected with no difference in ovulation rate between genotypes. However there was a significant decrease in ovulation rates between the 4- and 7-wk-old WT and Het mice (P < 0.001) (Table 1). Interestingly, cumulus cells not containing oocytes were recovered from the oviducts of 4-wk-old (but not 7-wk-old) ArKO mice, indicating ovulation events involved in the breakdown of the follicle wall may still be present.
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Ovarian Morphology
The ovarian morphology of superovulated 4-wk-old mice revealed that all stages of follicular development from primordial to antral stages were evident within the ovaries of all three genotypes. However, only WT and Het ovaries contained corpora lutea. Het and ArKO ovaries both presented with large cystic follicles, which is evidence of atresia. Oocytes within these atretic follicles were small, with pyknotic or absent nuclei, as reported previously in nonsuperovulated ArKO mice [6] (data not shown).
By 7 wk of age, conversely, there were pronounced differences in the ovarian morphology of superovulated mice. Wild-type ovaries, as expected, displayed an increased number of tightly packed follicles, ranging from primordial to preovulatory, together with corpora lutea. The Het ovaries exhibited a similar complement of developing follicles, but with an increase in the number of atretic follicles. The ArKO ovaries, in contrast, displayed aberrant morphology with large numbers of cystic follicles showing signs of atresia, containing shrunken, pyknotic, or anuclear oocytes, with only a few healthy-looking follicles. These follicles were loosely packed with an uneven, minimal, or absent granulosa cell layer (data not shown).
Retrieval of Oocytes
The number of immature oocytes harvested from 4-wk- old WT, Het, and ArKO following eCG stimulation did not differ significantly (P > 0.05), with an approximate average of 16 oocytes per mouse (Table 1). A noticeable difference between the ArKO, WT, and Het oocytes, however, was the presence or absence of surrounding cumulus cells. While WT and Het immature oocytes released from ovary perforations were all cumulus enclosed, the majority of 4-wk- old ArKO immature oocytes were deficient in cumulus cells. To compensate for this deficiency, cumulus cells were cultured together with immature oocytes in vitro to provide factors that may be beneficial to the oocyte during in vitro maturation. Oocytes retrieved from ArKO ovaries at 7 wk of age were not denuded, as observed for the 4-wk-old ArKO mice, but rather were enclosed within a typical cumulus complex.
Oocyte In Vitro Maturation, Fertilization, and Blastocyst Rates
Oocyte maturation rates were not significantly different between the three genotypes or across both age groups (P > 0.05), with almost 75% of the immature oocytes progressing to metaphase II in vitro (Table 2). Mature oocyte morphology from the three genotypes was also not different (Fig. 1). Chromatin staining confirmed the arrest of these oocytes at the second meiotic division, with chromatin staining clearly present in the oocyte and the polar body (data not shown).
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Only oocytes deemed mature (appearance of first polar body evident in perivitelline space; Fig. 1) were inseminated. Similar to in vitro maturation rates, in vitro fertilization rates did not differ between WT, Het, and ArKO oocytes (Table 2), with approximately 67% of the mature oocytes fertilizing.
Only those oocytes recognized as fertilized (presence of two pronuclei and a second polar body in the perivitelline space) were cultured to blastocysts. Blastocyst development rates for the WT, Het, and ArKO also did not differ significantly (Table 2), with approximately 65% of the zygotes developing to the blastocyst stage. WT, Het, and ArKO blastocyst morphology was also similar (Fig. 2).
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All in vitro maturation, fertilization, and blastocyst culture experiments repeated in charcoal-stripped, phenol-red- free media recorded equivalent results to those reported above (data not shown).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The current study demonstrates that the ovaries of 4- and 7-wk-old ArKO mice cannot be superovulated to produce COCs. However, when manually retrieved, oocyte in vitro maturation, fertilization, and blastocyst development rates did not differ between ArKO, WT, or Het mice or between the two age groups (4 and 7 wk old) examined, suggesting estrogen is not essential to maintain oocyte developmental competence.
The ArKO nonresponsiveness to a typical superovulation protocol is perhaps not surprising, given the chronically elevated levels of gonadotrophs reported in adult ArKO females [6, 13]. The ArKOs aged 12–14 wk in the Fisher et al. [13] study and the ArKOs aged 10–12 and 21– 23 wk and 1 yr in the Britt et al. [6] study all consistently displayed serum LH and FSH levels 2- to 10-fold higher than the WT at corresponding ages. Previous studies investigating the role of the ER signaling mechanism found that a dysfunctional HPG axis leads to overstimulation of LH production, which in turn results in hemorrhagic and cystic ovarian conditions (similar to that observed in this study and others) that are incapable of complete follicle development and growth [14, 15]. Adult 
ERKO mice fail to spontaneously ovulate [14, 16], and this remains so following superovulation [14]. However, ovulatory capacity can be restored in the superovulated adult ERKOs if they are first treated with gonadotroph antagonists that result in the restoration of gonadotroph and steroid levels to normal values in these mice [14]. Prepubertal ERKOs have normal gonadotroph and steroid levels and are capable of ovulation following superovulation [14]. In contrast with the adult ERKOs, adult ßERKOs can ovulate spontaneously, although at reduced rates, which are not improved by superovulation [17]. These mice have a normal HPG axis [18]; thus, estrogen is able to regulate the release of gonadotrophs in the ßERKO by acting on the ER in the hypothalamus. It is likely that the actions of estrogen through the ERß receptor in the ovary is contributory to the process of ovulation but that the intraovarian role of estrogen in the process of follicle development to ovulation is much less important than its role as a feedback regulator of the HPG axis [17]. Observations of the ovulatory capacity in both ERKOs further support the hypothesis that a normal HPG axis is required for the maintenance of normal gonadotroph and steroid levels, which is essential to the process of ovulation. Further support is provided by observations in the ßERKO model, where LH levels are elevated compared with other ERKOs [19, 20]. The ßERKO also fails to ovulate spontaneously [19]. The ßERKO ovarian morphology has also been reported to be significantly perturbed, exhibiting follicle transdifferentiation to structures resembling seminiferous tubules of the testis [19], a phenomenon also observed in ArKO ovaries [6]. The HPG axis in the ßERKO has been reported to be dysfunctional, and the very high LH levels in this mouse indicate that the absence of estrogen action on both the hypothalamic ER and the ovarian ERß leads to exacerbation of ovarian pathology. The recent report by Couse and colleagues [20] demonstrates that it is ER that is essential to maintain proper function of the HPG axis.
Sexually immature ArKO females, however, were observed to partially respond to superovulation. Prepubertal (4-wk-old) ArKO oviducts did contain cumulus cells following eCG and hCG administration, suggesting these ovaries may have been responding to the exogenous gonadotrophs with the typical events involved in follicle wall breakdown to release the contents of the follicles. The absence of oocytes within these cumulus complexes indicates the oocytes within the final-stage tertiary follicles were atretic or their interaction with the intrafollicular cumulus cells was compromised in some way. There have been no reports on the endocrinology of prepubertal ArKOs to date; however, as the cyclic release of gonadotrophs does not occur until the onset of puberty, the prepubertal gonadotroph levels and thus the HPG axis in the ArKO is assumed to be normal; hence, elevated gonadotrophs are unlikely to be responsible for the attenuated response of 4-wk-old ArKO ovaries. Furthermore, the observation that cumulus cells, not COCs, were present in the oviducts of the 4-wk- old superovulated ArKOs suggests that some ovulatory response has occurred. That immature oocytes recovered from eCG-stimulated ovaries in the second experiment were deficient in cumulus cells suggests that there may be some pathology in the coupling of cumulus cells to the oocyte. Certainly, in states of low to nil estrogen, reminiscent of the ArKO mouse, gap junction formation has been reported to be compromised, and exogenous administration of 17ß-estradiol to ovariectomized or immature rats stimulated the formation of gap junctions [21–23]. Therefore, it remained a possibility that superovulated prepubertal ArKOs did ovulate COCs; however, due to the tenuous relationship between the oocyte and its cumulus cells, the oocyte was retained in the bursa while the cumulus cells were picked up by the cilia on the fimbria and transported into the oviduct. Examination of ArKO bursa, however, demonstrated that oocytes were not being retained in this compartment (data not shown). Hence, the reason(s) behind the tenuous coupling of cumulus cells to the oocyte in the 4-wk-old superovulated ArKOs is unclear and remains to be clarified, but evidently, estrogen plays some indirect role in this event.
Surprisingly, inspection of ovaries from 4-wk-old superovulated ArKO mice did not reveal any CLs, as may be expected concomitant with cumulus cells being released into the oviduct. Transition of a follicle into a CL involves a multitude of remodeling events incorporating morphological, endocrinological, and biochemical changes. However, the vital importance of active, locally produced estrogen in the generation of healthy and fully functioning CLs has been demonstrated by others, e.g., Lund et al. [24]. Following administration of the aromatase inhibitor, arimidex, to proestrous ewes, Lund and colleagues reported that resultant CLs were smaller, contained fewer granulosa cells, were deficient in large steroidogenic cells, contained less progesterone than CLs from control animals, and were without the usual phagocyte influx into luteal tissues. The competency of the CLs from these ewes with attenuated estradiol biosynthesis was evidently significantly compromised. Similarly, given that ArKO mice are subject to lifelong estrogen deficiency, we postulate that the estrogen- regulated local events comprising the remodeling of tissue to form a CL are more severely compromised, and identifiable CLs are not generated.
Ovulation can be compared with an inflammatory response, in which cells of the immune system are recruited to the ovary and participate in the process of tissue remodeling. It has been established that estrogen contributes to regulating recruitment of these cells to the site of tissue remodeling involved in each ovulation event, particularly macrophage, neutrophilic granulocyte, and T lymphocyte populations [25, 26]. ER transcripts have been identified in mature peripheral T lymphocytes (reviewed in [27]), which have also been recovered from follicular fluid [28]. The tissue distribution of macrophages and neutrophils varies between preovulatory and ovulating follicles and is associated with increased numbers periovulation, suggesting they are actively participating in the ongoing tissue remodeling [25]. It is very likely, therefore, that the loss of endogenous estrogen in ArKO mice will have a significant effect on these populations of cells and their role in tissue remodeling concomitant with ovulation. Certainly, the chronic loss of estrogen in ArKO mice has been demonstrated to result in thymic regression and impaired thymocyte development [29]; however, the direct and indirect effects of this loss on follicle development in terms of cells of the immune system remain to be elucidated.
Treatment of 7-wk-old ArKOs with a gonadotroph antagonist in an attempt to restore normal circulating levels of gonadotrophs and androgens before superovulation treatment would be a valuable further experiment to explore the above hypothesis. In the event that ArKO mice fail to ovulate, the role of estrogen in the induction of LH- and FSH- receptor expression by the cumulus and granulosa cells should be examined. FSH and LH are key players in the processes of follicle recruitment and selection [30]. LH and FSH receptors are expressed by granulosa cells, and in response to the binding of estrogen to ERs, gonadotroph receptor numbers are significantly increased [31, 32]. A study determining whether LH and FSH receptors are present and in normal numbers in the follicles of the estrogen-deficient ArKO may reveal another potential cause for the failure of the ArKO to ovulate. If this is the case, it should also be noted that correcting LH- and FSH-receptor deficiency may not be as simple as supplying the mouse with exogenous estrogen. In the study carried out by Toda et al. [33], it was reported that administration of exogenous estrogen did not restore fertility to the ArKO and it was suggested that the ERs in these mice were unhealthy due to the presence of hemorrhagic and cystic follicles in ArKO ovaries. Thus, as well as examining ArKO follicles for functional LH and FSH receptors, the presence of functional ERs should also be investigated.
The present study provides further evidence in support of the hypothesis that estrogen is required to maintain normal circulating levels of gonadotrophs through its regulation via the HPG axis and that normal gonadotroph levels are required for complete follicle development and the process of ovulation. Prolonged exposure to very high levels of LH and androgens as a result of the failure of estrogen as a negative-feedback regulator of the HPG axis results in pathological ovarian morphology that deteriorates further with age. Therefore, estrogen indirectly maintains normal ovarian morphology that is required for complete follicle development and eventual ovulation.
The finding that ArKO oocytes were capable of maturation, fertilization, and development in similar numbers to WTs and Hets was somewhat surprising, as it has been reported that an androgen-dominated environment has a negative impact on oocyte maturation in vitro and subsequent embryo development [34]. In vitro maturation studies have yet to be performed in the ERKOs, so the relative importance of the action of estrogen on ER and ERß to the process of oocyte maturation is not known. However, evidence from the ERKOs would suggest that functional estrogen signaling mechanisms are not required to confer maturational and developmental competence to the oocyte [17, 35]. The results in the present study support this hypothesis with the findings that exposure to estrogen at least up to the early antral stages of development is not essential for the oocyte to be developmentally competent. If there is indeed a need for estrogen for developmental competence, it is likely to be required at extremely low levels, as the experiments outlined in the present study were conducted in both complete and charcoal-stripped/phenol-red-free media under oil, which very rapidly depletes any available steroid in the culture medium [36, 37].
Comparative in vitro studies that complement this use of the ArKO mouse model have also investigated the consequences of estrogen depletion to follicle growth and oocyte maturation. Hu et al. [38] cultured mouse preantral follicles in vitro in the presence of Arimidex, an aromatase inhibitor. While Arimidex treatment impaired the fertilization rate of oocytes, there were no concomitant adverse effects on follicular development, in vitro ovulation rates, nor preimplantation embryo development.
The hypothesis that estrogen was essential for oocyte developmental competence was proposed due to the expression of mRNA for both estrogen receptors in mouse unfertilized oocytes [39] and COCs [39, 40] and in postimplantation mouse embryos from Days 8 to 18 of gestation [39] and human COCs and oocytes [41]. The estrogen-responsive finger protein, efp, demonstrated a similar expression pattern [40], indicating estrogen was active during these developmental windows. Data from those studies also suggested that the effects of estrogen may be direct via receptor-mediated mechanisms, or indirect, such as through the action of estrogen-regulated growth-promoting factors [39, 40]. Contrary to this supposition, however, the results from the current study demonstrate that estrogen action is not a requirement for oocyte developmental competency to the blastocyst stage. There are several reports in the literature regarding the role of estrogen and in particular its action via ERs on the development and implantation of preimplantation embryos [39, 42, 43]. These reported the presence of ER mRNA in the oocyte [39] and the presence of the ER in the nucleus of mouse blastocysts and in all blastocyst cell types [44] as well as the presence of ER protein in the blastocyst [43]. In the study of Hou et al. [44], it was also reported that embryos treated with estrogen in vitro have significantly higher implantation rates when transferred to foster mothers. It was thus suggested from these findings that estrogen action on ER at the blastocyst stage may aid in implantation of preimplantation embryos. Although immature oocytes from the ArKO are capable of maturation and complete embryo development to the implantation stage, it remains to be determined whether these blastocysts are capable of implanting and developing to normal offspring. Experiments involving the transfer of ArKO blastocysts to WT fosters would address this question.
In conclusion, the role of estrogen in ovulation appears to be to regulate cyclic gonadotroph release via its action on the ERs in the hypothalamus. In doing so, estrogen maintains adequate ovarian conditions for growth and maturation of an ovulatory follicle and also aids in the initiation of recruitment, selection, and ovulation by triggering the secretion of the appropriate levels of gonadotrophs and stimulating adequate expression of FSH and LH receptors as well as ERs. Apparently, however, estrogen is not directly required for the production and growth of oocytes capable of maturation and complete preimplantation development. Whether the blastocysts developed from these oocytes are capable of complete development to live offspring when transferred to foster WTs remains to be elucidated.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Margaret Jones, PHIMR, Level 4, Block E, 246 Clayton Road, Clayton VIC 3168, Australia. FAX: 61 3 9594 6376; Margaret.Jones{at}phimr.monash.edu.au
Received: 11 August 2003.
First decision: 30 August 2003.
Accepted: 23 December 2003.
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REFERENCES |
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and ß in reproductive tissues. Reproduction 2003 125:143-149[Abstract] but not ERß. Mol Endocrinol 2003 17:1039-1053This article has been cited by other articles:
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  I. Segers, T. Adriaenssens, W. Coucke, R. Cortvrindt, and J. Smitz Timing of Nuclear Maturation and Postovulatory Aging in Oocytes of In Vitro-Grown Mouse Follicles with or Without Oil Overlay Biol Reprod, May 1, 2008; 78(5): 859 - 868. [Abstract] [Full Text] [PDF]
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 A. A. Murray, A. K.E. Swales, R. E. Smith, M. D. Molinek, S. G. Hillier, and N. Spears Follicular growth and oocyte competence in the in vitro cultured mouse follicle: effects of gonadotrophins and steroids Mol. Hum. Reprod., February 1, 2008; 14(2): 75 - 83. [Abstract] [Full Text] [PDF]
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N. Kawano and M. Yoshida Semen-Coagulating Protein, SVS2, in Mouse Seminal Plasma Controls Sperm Fertility Biol Reprod, March 1, 2007; 76(3): 353 - 361. [Abstract] [Full Text] [PDF]
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J. M. A. Emmen, J. F. Couse, S. A. Elmore, M. M. Yates, G. E. Kissling, and K. S. Korach In Vitro Growth and Ovulation of Follicles from Ovaries of Estrogen Receptor (ER){alpha} and ER{beta} Null Mice Indicate a Role for ER{beta} in Follicular Maturation Endocrinology, June 1, 2005; 146(6): 2817 - 2826. [Abstract] [Full Text] [PDF]
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