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Biology of Reproduction 60, 929-936 (1999)
©Copyright 1999 Society for the Study of Reproduction, Inc.

Effects of Glucocorticoids on Maturation of Pig Oocytes and Their Subsequent Fertilizing Capacity In Vitro1

Jyuer-Ger Yanga, Wei-Yi Chena, and P. Shirley Li2,a

a Department of Physiology, College of Medicine, National Cheng Kung University, Tainan, Taiwan 70101, Republic of China


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The aim of this study was to assess the possible role of glucocorticoids in the maturation of pig oocytes and their subsequent fertilizing capacity in vitro. Pig cumulus-enclosed oocytes collected from prepubertal gilts were cultured in Waymouth MB752/1 medium supplemented with sodium pyruvate (50 µg/ml), LH (0.5 µg/ml), FSH (0.5 µg/ml), and estradiol-17ß (1 µg/ml) in the presence or absence of cortisol or dexamethasone (DEX) for 24 h; they then were cultured without hormonal supplements in the presence or absence of cortisol or DEX for an additional 16–24 h. Treatment of cumulus-enclosed or denuded oocytes with increasing concentrations of cortisol or DEX for 48 h resulted in a dose-response inhibition of germinal vesicle breakdown (GVB). Increasing duration (12–48 h) of treatment with DEX (10 µg/ml) led to a time-dependent inhibition of GVB, which achieved statistical significance by 12 h. The addition of DEX (10 µg/ml) to maturation medium immediately after culture or at 12 h, 24 h, or 36 h after culture also decreased the percentage of oocytes with GVB. When oocytes were exposed to DEX for 48 h, the maturation rate was reduced. The degree of this reduction was dependent on DEX, and a concentration of DEX higher than 0.1 µg/ml was needed. The inhibitory effect of DEX on the maturation of oocytes was prevented by the glucocorticoid receptor antagonist RU-486. Exposure of oocytes to DEX for 40 h did not prevent sperm penetration, affect the incidence of polyspermy, or decrease the ability of oocytes to form a male pronucleus. The intracellular concentration of glutathione (GSH) in cumulus-enclosed oocytes was 4.4 mM per oocyte. Exposure of oocytes to DEX (0.01–10 µg/ml) had no effect on GSH concentration. These results demonstrate that glucocorticoids directly inhibit the meiotic but not cytoplasmic maturation of pig oocytes in vitro. This inhibitory effect is not mediated through a decrease in the level of intracellular GSH.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Glucocorticoids potentially affect normal gonadal function by acting at any one or more of the following levels in hypothalamic-pituitary-gonadal axis: 1) the hypothalamus (to decrease the synthesis and release of GnRH); 2) the anterior pituitary gland (to inhibit the synthesis and release of gonadotropins); 3) the testis/ovary (to modulate steroidogenesis and/or gametogenesis directly). It is thought that the principal deleterious actions of the glucocorticoids occurs at the hypothalamus and the anterior pituitary gland, creating a state of hypogonadotropic hypogonadism. This view is supported by a huge body of data, including observations that administration of synthetic glucocorticoids can significantly decrease hypothalamic GnRH release [1–3] and can inhibit the GnRH-stimulated release of LH and FSH from the pituitary [3–7]. However, recent studies have identified glucocorticoid receptors in ovarian and testicular cells [8–11] and have clearly shown that glucocorticoids have direct actions on gonadal steroidogenesis, both in vivo and in vitro [8, 12–17]. Another mechanism by which the hypothalamic-pituitary-adrenal (HPA) axis may influence reproductive function is by a direct effect of glucocorticoids on the target tissues of sex steroids [18].

It is well known that mammalian oocytes are maintained in a stage of meiotic arrest until shortly before ovulation. It is only after the preovulatory surge of LH that the oocyte resumes meiotic maturation and becomes fertilizable. To date, the direct effects of glucocorticoids on oocyte maturation have only been studied in several species of fish, such as Atlantic croaker [19]. Although no comparable data are yet available for mammals and the presence of glucocorticoid receptors on the oocyte has not been reported, it has been shown that in women undergoing ovarian follicular aspiration for in vitro fertilization, the concentration of cortisol measured in the follicular fluid is about 2-fold higher in follicles containing morphologically mature as opposed to immature oocytes [20], implying a positive correlation between the local cortisol concentration and oocyte maturity in humans. With respect to the male, no direct evidence has been reported to involve glucocorticoids in germ cell development. However, results from some studies show the presence of glucocorticoid receptor mRNA in pachytene spermatocytes and spermatids of the seminiferous tubules [10, 21]. With Western blot analysis, rodent epididymal sperm containing the glucocorticoid receptor have also been reported [22].

Glutathione (GSH) is a major intracellular thiol that has important biological functions during cellular proliferation, amino acid transport, and DNA and protein synthesis; and it protects cells against oxidation [23]. The synthesis of GSH during oocyte maturation appears to be a prerequisite for sperm chromatin decondensation and hence for male pronucleus formation after sperm penetration of mouse [24], hamster [25], and pig [26] oocytes. An inability by porcine oocytes to synthesize sufficient GSH during maturation in vitro reduces their ability to form a male pronucleus after fertilization [26]. The addition of cysteine to maturation medium increases the GSH content of in vitro-matured porcine oocytes [27], whereas addition of low-molecular weight thiols, such as cysteamine and ß-mercaptoethanol, to culture medium enhances cysteine-mediated GSH synthesis in in vitro-produced 6- to 8-cell bovine embryos [28] and increases synchronous pronuclear formation and normal embryonic development in porcine oocytes matured and fertilized in vitro [29].

The present study was conducted to investigate whether glucocorticoids affect the in vitro maturation (IVM) of pig oocytes and their subsequent fertilizing capacity as determined by germinal vesicle breakdown (GVB), intracellular GSH content, sperm penetration, and male pronucleus formation.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Culture Media

The basic medium used for the manipulation of oocytes consisted of Waymouth MB752/1 (with L-glutamine but no sodium bicarbonate; Gibco, Grand Island, NY) supplemented with sodium pyruvate (50 µg/ml; Sigma Chemical Co., St. Louis, MO), fetal calf serum (FCS; 5%; Hyclone Labs., Logan, UT), penicillin (100 U/ml; Sigma), streptomycin sulfate (50 µg/ml; Sigma), and fungizone (0.625 µg/ml; Gibco) as described previously [30] with slight modifications. The IVM medium consisted of basic medium supplemented with 0.5 µg/ml porcine LH (USDA-pLH-B2), 0.5 µg/ml porcine FSH (USDA-pFSH-I1), and 1 µg/ml estradiol-17ß (Sigma). The sperm preincubation medium (pH 7.8) consisted of modified Medium 199 (mM199, with Earle's salts; Gibco) containing calcium lactate (2.92 mM; Merck, Darmstadt, Germany), sodium pyruvate (0.91 mM), glucose (3.05 mM; Merck), Hepes (25 mM; Sigma), penicillin (50 U/ml), and streptomycin sulfate (50 µg/ml) and supplemented with 10% (v:v) FCS [31]. The fertilization medium consisted of mM199 (pH 7.4) supplemented with 10 mM caffeine-sodium benzoate (Sigma) and 10% FCS [31].

Collection of Oocytes

Ovaries were collected from prepubertal gilts at a local slaughterhouse and transported to the laboratory within 2 h in 0.85% (w:v) NaCl containing penicillin (250 U/ml), streptomycin sulfate (250 µg/ml), fungizone (0.625 µg/ml), and sucrose (0.25 M; Merck) at room temperature. Oocytes were aspirated from antral follicles (3–6 mm in diameter) with a 20-gauge needle fixed to a 5-ml disposable syringe and washed twice in basic medium and once in IVM medium. Only oocytes surrounded by at least 3–4 uniform layers of compact cumulus cells were selected, using a low-power (x20–30) dissecting microscope (Olympus Co., Tokyo, Japan) for the experiments. All oocytes from a batch of ovaries used on each day of experimental set-up were collected and pooled before distribution to all treatments within an experiment.

IVM

Groups of 10 cumulus-enclosed oocytes (CEOs) were cultured in a 100-µl droplet of IVM medium covered with paraffin oil for 24 h at 39°C in an atmosphere of 5% CO2 in air [31]. The CEOs were then washed three times in basic medium and transferred to a 100-µl droplet of basic medium, and cultured for an additional 24-h period. Cortisol (hydrocortisone sodium succinate; Solu-Cortef, Upjohn Co., Taipei, Taiwan), dexamethasone (DEX; dexamethasone phosphate; Narn Guang Chemical Co., Tainan, Taiwan), and RU486 (Mifepristone; Roussel-Uclaf, Paris, France) were added at the beginning of culture or as indicated to the IVM medium at the concentration indicated in the Results section. In some experiments, denuded oocytes (DOs; without layers of cumulus cells) were used for culturing.

Sperm Preparation

The sperm-rich fractions of ejaculates were obtained from a Duroc boar by the gloved-hand method. Semen samples were washed three times with 0.85% NaCl (w:v) containing 100 mg/L BSA (fraction V; Sigma). Washed spermatozoa were subsequently diluted to 2 x 108 cells/ml in the sperm preincubation medium (pH 7.8) and incubated for 90 min at 39°C in an atmosphere of 5% CO2 in air.

In Vitro Fertilization (IVF)

After 40 h of culture for maturation, oocytes with an expanded cumulus mass were washed three times in fertilization medium and then placed into a droplet of 50 µl of fertilization medium covered with paraffin oil, which was pregassed in a CO2 incubator at 39°C for more than 3 h. The sperm concentration (2 x 108 cells/ml) in preincubated medium was diluted to 2 x 106 cells/ml with mM199 at pH 7.8 and supplemented with 10% FCS, and 50 µl of diluted spermatozoa was added to the 50-µl droplet containing oocytes. Oocytes were cultured with spermatozoa for 8 h at 39°C in an atmosphere of 5% CO2 in air.

Assessment of Nuclear Maturation, Sperm Penetration, and Pronuclear Formation

At the end of the culture (48 h after IVM or 8 h after IVF), oocytes were freed from cumulus cells by agitation using a narrow-bore glass pipette in PBS (pH 7.4) and washed three times with PBS. The DOs were mounted, fixed for 48 h in acetic alcohol (methanol and acetic acid, 3:1, v:v), stained with 1% (w:v) orcein in 45% (v:v) acetic acid, and then examined under a Nomarski differential interference microscope (Olympus) for assessment of nuclear maturation. Resumption of meiosis was assessed by the breakdown of the germinal vesicle. Oocytes with a polar body and a metaphase plate were classified as being in metaphase (M) II stage and were regarded as matured. Oocytes were designated as penetrated when they had one or more swollen sperm heads or a male pronucleus and its corresponding sperm tail. Those oocytes with more than one swollen sperm nucleus or male pronucleus were considered to be polyspermic. Only oocytes containing male and female pronuclei with an intact nuclear membrane were considered as having formed male and female pronuclei.

Assay of GSH

At the end of IVM, oocytes were stripped of surrounding cumulus cells by repeated pipetting with a narrow-bore glass pipette in PBS and washed three times with PBS. For each replicate, groups of 25–30 oocytes in 5 µl of PBS from each treatment were placed in microtubes and frozen at -20°C for 18 h. The frozen samples were thawed once at room temperature, 5 µl of 1.25 M phosphoric acid (Sigma) was added to the tube, and then oocytes were ruptured by repeated agitation using a narrow-bore glass pipette. The tubes containing samples were stored at -20°C until assayed. Blanks containing 5 µl of PBS without oocytes were similarly prepared.

GSH was determined by the DTNB (5,5'-dithiobis-[2-nitrobenzoic acid])-GSSG (glutathione disulfide) reductase recycling assay [32]. Briefly, 700 µl of 0.33 mg NADPH/ml (Sigma) in 0.2 M sodium phosphate buffer containing 10 mM EDTA (stock buffer, pH 7.2; Merck), 100 µl of 6 mM DTNB (Sigma) in the stock buffer, and 190 µl of water were added into the microtubes containing thawed samples. Twenty-five microliters of 100 IU/ml glutathione reductase (Sigma) was added with mixing to initiate the reaction. The rate of 5-thio-2-nitrobenzoic acid (TNB) formation was followed at 412 nm with a spectrophotometer (DU-640; Beckman Instruments, Inc., Fullerton, CA) and recorded at 0.5 and 2 min after the addition of glutathione reductase. Both the assay reagent blank and GSH (Sigma) standards (30–240 ng/200 µl) were assayed under the same conditions. The amount of GSH in each sample was determined by comparison with a standard curve prepared at the same time. This amount was divided by the number of oocytes in the sample to obtain the total GSH content per oocyte. To calculate the mean cell volume, the diameter of a total of 200 denuded oocytes was measured by means of an inverted phase-contrast microscope (Olympus) with an ocular micrometer (0.01 mm). The mean diameter was 117.85 ± 0.14 µm, and calculation by the equation V = 4 {pi}r3/3 indicated that the mean cell volume of oocytes was 857.01 pl. Then, the GSH concentration per oocyte, estimated by the total GSH content per oocyte and the mean cell volume, was determined as described by Calvin et al. [24].

Statistical Analysis

Percentages for all variables were compared using the STATVIEW (Abacus Concepts, Inc., Berkeley, CA) program for ANOVA and Fisher's protected least-significant difference test. All percentage data were subjected to arc sine transformation before statistical analysis. Oocyte GSH levels were analyzed from eight replicates in four experiments by ANOVA via the STATVIEW program. A value of p < 0.05 was considered to be statistically significant.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Effects of Glucocorticoids on Oocyte Nuclear Maturation

To test whether glucocorticoids could play a role in the meiotic maturation process, oocytes were cultured in the presence or absence of various concentrations of cortisol or DEX, a synthetic glucocorticoid. The oocytes were evaluated after 48 h of culture, and the percentages of oocytes with GVB are presented in Figures 1 and 2. Cortisol had a significant (p < 0.05) effect in inhibiting GVB (Fig. 1). The effect was concentration-dependent, with the minimal effective dose at 0.1 µg/ml. The inhibition of GVB increased with the dose, up to a maximum of 10 µg/ml. DEX at concentrations of 0.1, 1, 5, 10, and 50 µg/ml blocked (p < 0.05) GVB in a dose-dependent manner (Fig. 2).



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  		FIG. 1. Effect of the dose of cortisol on the maturation of pig oocytes. The results are expressed as the percentage of oocytes with GVB after 48 h of culture in the presence or absence of increasing concentrations of cortisol (0.01–10 µg/ml). Values were obtained from three independent experiments and are expressed as mean ± SEM. Bars with different letters differ significantly (p < 0.05). In parentheses: number of oocytes examined.



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  		FIG. 2. Effect of the dose of DEX on the maturation of pig oocytes. The results are expressed as the percentage of oocytes with GVB after 48 h of culture in the presence or absence of increasing concentrations of DEX (0.001–50 µg/ml). Values were obtained from two independent experiments and are expressed as mean ± SEM. Bars with different letters differ significantly (p < 0.05). In parentheses: number of oocytes examined.

Denuded oocytes were then used to study the possible effect of cumulus cells on the glucocorticoid influence on oocyte maturation (Fig. 3). DEX at 0.1–50 µg/ml for 48 h suppressed GVB in a dose-dependent manner. This effect reached a maximum at 10 µg/ml.



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  		FIG. 3. Effect of the dose of DEX on the maturation of denuded pig oocytes. The results are expressed as the percentage of oocytes with GVB after 48 h of culture in the presence or absence of increasing concentrations of DEX (0.1–50 µg/ml). Values were obtained from 3 to 9 independent experiments and are expressed as mean ± SEM. Bars with different letters differ significantly (p < 0.05). In parentheses: number of oocytes examined.

In the time-course experiment (Fig. 4A), oocytes were incubated for 0, 12, 24, 36, and 48 h in the presence of 10 µg DEX/ml, washed thoroughly, and then incubated in the culture medium without DEX for an additional 48, 36, 24, 12, and 0 h, respectively. Increasing duration (12–48 h) of treatment with DEX led to a time-dependent inhibition of GVB, achieving statistical significance (p < 0.001) by 12 h. The addition of DEX after 12–36 h of culture was as effective in inhibiting GVB as it was at the beginning of culture (Fig. 4B).



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  		FIG. 4. Effect of DEX (10 µg/ml) during various periods on nuclear maturation (GVB) of pig oocytes evaluated after 48 h of culture. A) Oocytes were cultured with DEX for 0, 12, 24, 36, or 48 h; washed thoroughly; and then cultured in culture medium without DEX for an additional 48, 36, 24, 12, and 0 h, respectively. B) Oocytes were cultured with DEX added at the beginning of culture, or 12 h, 24 h, 36 h, or 48 h after culture. The results are expressed as mean ± SEM from three independent experiments. Values with different letters differ significantly (p < 0.001). In parentheses: number of oocytes examined.

Effect of Glucocorticoid on IVM and IVF

To determine whether DEX affects IVM and IVF, oocytes were cultured for 40 h with various concentrations (0.01–10 µg/ml) of DEX before being transferred to the fertilization medium. Exposure of oocytes to DEX suppressed the maturation rates in a dose-dependent manner (Table 1). At 8 h after insemination, 71% of the in vitro-matured oocytes had been penetrated by spermatozoa and had already resumed meiotic progression from the MII stage, and 41% had male pronuclear formation (Table 1). Exposure of oocytes to DEX for 40 h did not prevent sperm penetration, affect the incidence of polyspermy, or decrease the ability of oocytes to form a male pronucleus (Table 1).


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  		TABLE 1. Effect of DEX on in vitro maturation, sperm penetration, and male pronuclear formation in pig oocytes at 8 h after insemination.

Effect of RU486 on Glucocorticoid Inhibition ofNuclear Maturation

The glucocorticoid receptor antagonist RU486 [33] was used to determine whether the effects described above were mediated by the glucocorticoid receptor. Oocytes were cultured for 48 h with DEX (10 µg/ml) in the presence or absence of RU486 (50 µM). The results are presented in Figure 5. RU486 antagonized the inhibitory effect of DEX on GVB and maturation rate.



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  		FIG. 5. Effect of RU486 on dexamethasone (DEX) actions in pig oocytes. Oocytes were cultured with or without DEX (10 µg/ml) in the presence or absence of RU486 (50 µM) for 48 h. The results are expressed as mean ± SEM from three independent experiments. p < 0.001 for a vs. b. In parentheses: number of oocytes examined.

Effect of Glucocorticoids on GSH Concentration

Whether suppression of GVB by glucocorticoids could be accounted for by changes in intracellular GSH levels was further evaluated. CEOs were cultured for 48 h with various concentrations of DEX. Four samples (30 oocytes/sample) of each treatment from two replications were assayed. The intracellular concentration of GSH in oocytes treated with medium alone was 4.4 ± 1.6 mM per oocyte. Exposure of oocytes to DEX had no effect on GSH concentration. Results from the two experiments in which oocytes were treated with DEX at concentrations of 0.01, 0.1, 1, and 10 µg/ml were 6.22 ± 0.65, 6.37 ± 1.38, 6.26 ± 1.17, and 5.19 ± 0.7 mM GSH/oocyte, respectively.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The studies reported here demonstrated that cortisol or DEX is able to significantly inhibit meiotic maturation in cumulus-enclosed pig oocytes in a dose- and time-dependent manner and that it can directly affect meiotic maturation in denuded oocytes. This inhibitory effect is prevented by the glucocorticoid receptor antagonist RU-486, thus suggesting that resumption of meiosis and maturation of the oocytes result essentially from glucocorticoid receptor-mediated events. The exposure of oocytes to DEX does not decrease the ability of oocytes to form a male pronucleus at 8 h after insemination. This finding suggests that DEX may not inhibit the cytoplasmic maturation of oocytes matured in vitro. In the literature, a number of studies support the concept that an inability by porcine oocytes to synthesize sufficient GSH during maturation in vitro reduces their ability to form a male pronucleus after fertilization [26]. This, and the fact that DEX was seen in this report not to affect GSH levels in oocytes, led us to speculate that its action in the oocyte is not mediated by this intracellular thiol.

In our studies, the minimum effective inhibitory dose of cortisol was at 100 ng/ml, which is similar to that found in pigs after ACTH administration or application of an acute stressor [34], but above the circulating levels under nonstressed conditions. Since > 90% of circulating glucocorticoids are bound by transcortin [35, 36], the present observations may represent an overestimate of the in vivo sustained stressed conditions, and the effect seen is probably more pharmacological than physiological. Thus, the influence of glucocorticoids in normal oocyte maturation may be minimal. However, perturbations of the HPA axis known to result in excessive production of glucocorticoids, or the administration of large amounts of glucocorticoids for therapeutic reasons, has been shown to alter the normal activity of ovarian follicular development. Interruption of folliculogenesis under these conditions may be explained, at least in part, by a direct glucocorticoid action on oocyte maturation.

The studies reported here provide evidence that glucocorticoids can act directly on oocytes. To date, the direct effects of glucocorticoids on oocyte maturation have been investigated in several species of fish. In these species, cortisol or corticosterone has been reported to promote final maturation of the oocyte [19, 37]. In selected species, such maturational effects appear to reflect the direct actions of the corticosteroid per se [37], whereas in others, they seem to result from a synergy with the 20ß-hydroxymetabolites of progesterone [19]. The apparent discrepancy between the above data and the present results may be explained by the differences in experimental animal and/or conditions used. Although no comparable data are yet available for other mammals, it has been shown that in women undergoing ovarian follicular aspiration for IVF, the concentration of cortisol measured in the follicular fluid is approximately 2-fold higher in follicles containing mature vs. immature oocytes [20], thus implying a positive correlation between the local cortisol concentration and oocyte maturity in humans.

In the present study, a significant inhibitory action of DEX on GVB was demonstrated. The addition of DEX late in the meiotic progression (during the last 12 h of culture) also resulted in a remarkable decrease in maturation of the oocytes during a 48-h culture period. Christmann et al. [38] reported that pig oocytes show heavy chromatin condensation at 12 h after culture. By 24 h, most oocytes had entered MI. Matured MII oocytes were observed mainly after 35 h in culture and increased gradually to reach 85% of the total at 45–48 h in culture [39]. Therefore, the effect of glucocorticoids on the maturation of oocyte appears to be time-dependent. In our study, DEX was also able to act directly on denuded oocytes. It is known that the inhibition of oocyte maturation is usually mediated by the cells of the cumulus oophorus, and that the relief of inhibition induced by gonadotropins or by several growth factors is a result of their effect on the cumulus cells and not on the oocyte itself [40]. Therefore, our data suggest that pig oocytes possess specific glucocorticoid receptors. In fact, we have observed that glucocorticoid receptor mRNA was expressed in denuded pig oocytes (data not shown). In this regard, it has been already observed that glucocorticoid receptors are present in rat primary spermatocytes and that the expression of these receptors in the seminiferous tubule of the rat testis appears to be confined to stages I–III and XIII–XIV of the spermatogenic cycle [11]. Moreover, the inhibitory action by DEX was prevented in CEOs by RU486. Glucocorticoid receptors have been identified in granulosa cells [9], and glucocorticoids have been shown to exert direct effects on ovarian steroidogenesis, both in vivo and in vitro [12, 14,17]. However, RU-486 also blocks progesterone receptors [33]. Studies in human granulosa cells indicate that progesterone regulates granulosa cell proliferation and differentiation in an autocrine-paracrine manner [41]. Another study of porcine granulosa cells with the synthetic progesterone R-5020 showed that the involvement of progesterone in its own production was minimal [42]. Although the concentration of progesterone in culture media during maturation in vitro was not measured in this study, a recent study has found that pig oocyte-cumulus complexes under paraffin oil were exposed to extremely low concentrations of progesterone during maturation [43]. Therefore, our results indicate that the effects of DEX on meiotic maturation in pig oocytes are mediated through the glucocorticoid receptor.

Recent reports indicate that GSH is an important factor for male pronuclear formation of pig oocytes [44]. Supplementation of cysteine, which possibly served as a substrate for GSH, into maturation medium improved both the GSH concentration and the incidence of male pronuclear formation [27]. Our GSH studies indicate that DEX did not affect the intracellular GSH concentration during oocyte maturation in CEOs. On the other hand, exposure of oocytes to DEX for 40 h had no effect on the ability of oocytes to form a male pronucleus after IVF. It is thus possible that GSH may not be involved in the inhibitory effect of DEX on meiotic maturation. Other mechanisms to delay the resumption of meiosis may exist. The oocytes used in this study were from large antral follicles at various stages of growth and atresia. The possibility remains that DEX can affect oocytes from secondary or primary follicles or exerts specific effects on the oocytes in the follicle destined for ovulation. It is unclear whether the ability of DEX to suppress oocyte maturation in vitro can explain its direct action on oocyte maturation in vivo. The physiological significance of this result remains to be confirmed.

In the present study, culture of CEOs with DEX before insemination did not prevent sperm penetration and did not affect the incidence of polyspermic penetration. It is believed that the release of cortical granules during fertilization prevents further sperm penetration at the level of the zona pellucida (zona reaction) and plasma membrane (vitelline block), although there are differences in the relative efficiencies of the zona reaction and of the vitelline block among species [45]. For the pig, it has been reported that the exocytosis of cortical granules occurs slowly and incompletely during IVF [46], resulting in a high incidence of polyspermy after IVF [47, 48]. Our findings suggest that this slow and incomplete exocytosis of cortical granules in vitro is apparently not affected markedly by DEX treatment before IVF.

It is of interest to consider the potential mechanisms by which glucocorticoids inhibit meiotic maturation in pig oocytes. The meiotic division in oocytes is a protracted process that is naturally arrested at the diplotene of the first prophase, which corresponds to the G2 phase of the cell cycle. Meiotically arrested oocytes are referred to as immature oocytes. Resumption of meiosis in these oocytes is known as oocyte maturation and entails a G2-to-M transition. The transition from G2 to M phase requires the cells to initiate complex processes including GVB, chromatin condensation, and reorganization of the cytoskeleton and progression to the MII stage, at which they undergo a second arrest [49]. The occurrence of these events is regulated by a substantial increase in the oocyte's cytosolic kinase activity [50, 51]. An important component of this activity is cyclin B-p34cdc2 kinase, also called maturation/metaphase-promoting factor (MPF) [52, 53]. Active MPF is responsible for the onset of M phase in all eukaryotic cells including oocytes [54, 55]. MPF is able to phosphorylate many of the proteins involved in nuclear membrane formation, chromatin condensation, and microtubular reorganization [56–58]. In the present study, glucocorticoids inhibited nuclear membrane breakdown. These results indicate that glucocorticoids influence resumption of meiosis. Since oocytes containing one polar body and a metaphase plate were regarded as matured, the observation that glucocorticoids suppressed the rate of maturation of pig oocytes suggests that glucocorticoids may prevent chromatin condensation or reorganization of the cytoskeleton. There is also some stereospecificity involved since glucocorticoids do not inhibit overall maturation. It is reported that inhibition of endogenous proteases prevented both chromatin condensation and nuclear membrane disintegration in pig oocytes [59]. Also inhibitors of endogenous proteases were found to inhibit initiation of maturation in cow [60], rat [61], amphibian [62], and starfish [63] oocytes. It is therefore possible that glucocorticoids effectively block proteolysis during culture of oocytes in vitro. As MPF is responsible for induction of chromatin condensation and GVB during M phase, proteolysis could be the first step in posttranslational modification of MPF, promoting its ability to condense chromatin. Glucocorticoids may also block MPF kinase activity and thus the ability of oocytes to enter into M phase. Fluctuation of MPF kinase activity during meiotic maturation has been reported in pig oocytes [39]. In addition, activation of p34cdc2 kinase depends upon both its association with its cyclin B regulatory subunit [64, 65] and a subsequent change in the phosphorylated state of key tyrosine and threonine residues on the catalytic p34cdc2 subunit [66]. The changes in concentration of these molecules during meiotic maturation have been reported in pig oocytes [67]. There is a possibility that glucocorticoids may decrease the levels of p34cdc2 and cyclin B or change the association between the two subunits in pig oocytes during meiotic progression from G2 to MII. Moreover, glucocorticoids may modify MPF activation by inhibiting dephosphorylation at threonine-14 and tyrosine-15 or by inhibiting phosphorylation at threonine-161 of p34, consequently blocking MPF activity. Investigations are in progress to determine the role that glucocorticoids play in the nuclear meiotic events that lead to the induction of GVB and meiotic maturation.

In conclusion, we have shown that glucocorticoids can influence meiotic but not cytoplasmic maturation of pig oocytes in vitro. These observations raise the possibility that glucocorticoids in vivo may act directly on the ovary to modulate oocyte maturation. Additional studies are necessary to clarify the mechanisms involved in the suppressing effect on the maturation of pig oocytes.


   ACKNOWLEDGMENTS
 
The authors are grateful to Tai-Yu Products Corp., Hsinying Factory, for providing pig ovaries and to Dr. Douglas J. Bolt, USDA Animal Hormone Program, for providing porcine LH and FSH. Special thanks are also given to Ms. Tzy-Ling Chen for her assistance with ovary collections.


   FOOTNOTES
 
1 This study was supported by a grant (No. NSC86–2314-B006–083) from the National Science Council, Republic of China. Back

2 Correspondence. FAX: 886 6 236 2780; lifupi{at}mail.ncku.edu.tw Back

Accepted: November 17, 1998.

Received: June 30, 1998.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Fonda ES, Rampacek GB, Kraeling RP. The effect of adrenocorticotropin or hydrocortisone on serum luteinizing hormone concentrations after adrenalectomy in the prepubertal gilt. Endocrinology 1984; 114:268–273.[Abstract]
  2. Dubey AK, Plant TM. A suppression of gonadotropin secretion by cortisol in castrated male rhesus monkeys (Macaca mulatta) mediated by the interruption of hypothalamic gonadotropin-releasing hormone release. Biol Reprod 1985; 33:423–431.[Abstract]
  3. Rosen H, Jameel ML, Barkan AL. Dexamethasone suppresses gonadotropin-releasing hormone (GnRH) secretion and had direct pituitary effects in male rats: differential regulation of GnRH receptor and gonadotropin responses to GnRH. Endocrinology 1988; 122:2873–2880.[Abstract]
  4. Li PS, Wagner WC. In vivo and in vitro studies on the effect of adrenocorticotropic hormone or cortisol on the pituitary response to gonadotropin-releasing hormone. Biol Reprod 1983; 29:25–37.[Abstract]
  5. Li PS. Effect of cortisol or adrenocorticotropic hormone on luteinizing hormone secretion by pig pituitary cells in vitro. Life Sci 1987; 41:2493–2501.[CrossRef][Medline]
  6. Brann WB, Putnam CD, Mahesh VB. Corticosteroid regulation of gonadotropin and prolactin secretion in the rat. Endocrinology 1990; 126:159–166.[Abstract]
  7. Li PS. Actions of corticotropin-releasing factor or cortisol on follicle-stimulating hormone secretion by isolated pig pituitary cells. Life Sci 1993; 53:141–151.[CrossRef][Medline]
  8. Evain D, Morera AM, Saez JM. Glucocorticoid receptors in interstitial cells of the rat testis. J Steroid Biochem 1976; 7:1135–1139.[CrossRef][Medline]
  9. Schreiber JR, Nakamura K, Erickson GF. Rat ovary glucocorticoid receptor: identification and characterization. Steroids 1982; 39:569–584.[CrossRef][Medline]
  10. Levy FO, Ree AH, Eikvar L, Govindan MV, Jahnsen T, Hasson V. Glucocorticoid receptor and glucocorticoid effects in rat Sertoli cells. Endocrinology 1989; 124:430–436.[Abstract]
  11. Schultz R, Isola J, Parvinen M, Honkaniemi J, Wikstrom AC, Gustafsson JA, Pelto-Huikko M. Localization of the glucocorticoid receptor in testis and accessory sexual organs of male rat. Mol Cell Endocrinol 1993; 95:115–120.[CrossRef][Medline]
  12. Hsueh AJW, Erickson GF. Glucocorticoid inhibition of FSH-induced estrogen production in cultured rat granulosa cells. Steroids 1978; 32:639–649.[CrossRef][Medline]
  13. Welsh TH, Bambino TH, Hsueh AJW. Mechanism of glucocorticoid-induced suppression of testicular androgen biosynthesis in vitro. Biol Reprod 1982; 27:1138–1146.[Abstract]
  14. Ben-Rafall Z, Benadiva CA, Garcia CJ, Flickinger GL. Cortisol stimulation of estradiol and progesterone secretion by human granulosa cells is independent of follicle-stimulating hormone effect. Fertil Steril 1988; 49:813–816.[Medline]
  15. Hales DB, Payne AH. Glucocorticoid-mediated repression of P450scc mRNA and de novo synthesis in cultured Leydig cells. Endocrinology 1989; 124:2099–2104.[Abstract]
  16. Li PS. Effect of cortisol on testosterone production by immature pig Leydig cells. J Steroid Biochem Mol Biol 1991; 38:205–212.[CrossRef][Medline]
  17. Michael AE, Pester LA, Curtis P, Shaw RW, Edwards CRW, Cooke BA. Direct inhibition of ovarian steroidogenesis by cortisol and the modulatory role of 11ß-hydroxysteroid dehydrogenase. Clin Endocrinol 1993; 38:641–644.[Medline]
  18. Rabin DS, Johnson EO, Brandon DD, Liapi C, Chrousos GP. Glucocorticoids inhibit estradiol-mediated uterine growth: possible role of the uterine estradiol receptor. Biol Reprod 1990; 42:74–80.[Abstract]
  19. Patino R, Thomas P. Induction of maturation of Atlantic croaker oocytes by 17{alpha}, 20ß, 21-trihydroxy-4-pregnen-3-one in vitro: consideration of some biological and experimental variables. J Exp Zool 1990; 255:97–109.[CrossRef][Medline]
  20. Fateh M, Ben-Rafall Z, Benadiva CA, Mastroianni L, Flickinger GL. Cortisol levels in human follicular fluid. Fertil Steril 1989; 51:528–541.
  21. Levy FO, Eikvar L, Piotrowska B, Yen O, Jahnsen T, Hansson V. Characterization and cellular localization of glucocorticoid receptors in rat seminiferous tubules: detection of glucocorticoid receptor mRNA. Ann NY Acad Sci 1987; 513:487–489.
  22. Kaufmann SH, Wright WW, Okret S, Wikstrsm A-CH, Gustafsson JA, Shaper NL, Shaper JH. Evidence that rodent epididymal sperm contain the Mr ~94,000 glucocorticoid receptor but lack the Mr ~90,000 heat shock protein. Endocrinology 1992; 130:3074–3084.[Abstract]
  23. Meister A, Anderson ME. Glutathione. Annu Rev Biochem 1983; 52:711–760.[CrossRef][Medline]
  24. Calvin HI, Grosshans K, Blake EJ. Estimation and manipulation of glutathione levels in prepubertal mouse ovaries and ova: relevance to sperm nucleus transformation in the fertilized egg. Gamete Res 1986; 14:265–275.[CrossRef]
  25. Perreault SD, Barbee RR, Slott VL. Importance of glutathione in the acquisition and maintenance of sperm nuclear decondensing activity in maturing hamster oocytes. Dev Biol 1988; 125:181–186.[CrossRef][Medline]
  26. Yoshida M. Role of glutathione in the maturation and fertilization of pig oocytes in vitro. Mol Reprod Dev 1993; 35:76–81.[CrossRef][Medline]
  27. Yoshida M, Ishigaki K, Nagai T, Chikyu M, Pursel VG. Glutathione concentration during maturation and after fertilization in pig oocytes: relevance to the ability of oocytes to form male pronucleus. Biol Reprod 1993; 53:173–178.[Abstract]
  28. Takahashi M, Nagai T, Hamano S, Kuwayama M, Okamura N, Okano A. Effect of thiol compounds on in vitro development and intracellular glutathione content of bovine embryos. Biol Reprod 1993; 49:228–232.[Abstract]
  29. Grupen CG, Nagashima H, Nottle MB. Cysteamine enhances in vitro development of porcine oocytes matured and fertilized in vitro. Biol Reprod 1995; 53:173–178.
  30. Hirao Y, Nagai T, Kubo M, Miyano T, Miyake M, Kato S. In vitro growth and maturation of pig oocytes. J Reprod Fertil 1994; 100:333–339.[Abstract]
  31. Funahashi H, Day BN. Effects of the duration of exposure to hormone supplements on cytoplasmic maturation of pig oocytes in vitro. J Reprod Fertil 1993; 98:179–185.[Abstract]
  32. Anderson ME. Determination of glutathione and glutathione disulfide in biological samples. Method Enzymol 1985; 113:548–555.[Medline]
  33. Horwitz KB. The molecular biology of RU 486. Is there a role for antiprogestins in the treatment of breast cancer. Endocr Rev 1992; 13:146–163.[CrossRef][Medline]
  34. Juniewicz PE, Johnson BH. Ability of cortisol and progesterone to mediate the stimulatory effect of adrenocorticotropic hormone upon testosterone production by the porcine testis. Biol Reprod 1984; 30:134–142.[Abstract]
  35. Westphal U. Steroid protein interactions: concentrations and binding affinities of corticosteroid binding globulin in sera of men, monkey, rat, rabbit and guinea pig. Arch Biochem Biophys 1967; 118:556–567.[CrossRef][Medline]
  36. Thompson EB, Lippman ME. Mechanism of action of glucocorticoids. Metabolism 1974; 23:159–202.[CrossRef][Medline]
  37. Goswami S, Sundararaj BL. In vitro maturation and ovulation of oocytes of the catfish, Heteropneustes fossilis (Bloch): effects of mammalian hypophyseal hormones, catfish pituitary homogenate, steroid precursors and metabolites, and gonadal and adrenocortical steroids. J Exp Zool 1974; 178:467–478.[CrossRef]
  38. Christmann L, Jung T, Moor RM. MPF components and meiotic competence in growing pig oocytes. Mol Reprod Dev 1994; 38:85–90.[CrossRef][Medline]
  39. Naito K, Toyoda Y. Fluctuation of histone H1 kinase activity during meiotic maturation in porcine oocytes. J Reprod Fertil 1991; 93:467–473.[Abstract]
  40. Dekel N, Beers WH. Rat oocyte maturation in vitro: relief of cyclic AMP inhibition by gonadotropins. Proc Natl Acad Sci USA 1978; 75:4369–4373.[Abstract/Free Full Text]
  41. Chaffkin LM, Luciano AA, Peluso JJ. Progesterone as an autocrine/paracrine regulator of human granulosa cell proliferation. J Clin Endocrinol Metab 1992; 75:1404–1408.[Abstract]
  42. Romero G, Garmey JC, Veldhuis JD. The involvement of inositol phosphoglycan mediators in the modulation of steroidogenesis by insulin and insulin-like growth factor-1. Endocrinology 1993; 132:1561–1568.[Abstract]
  43. Funahashi H, Cantley TC, Stumpf TT, Terlouw SL, Day BN. In vitro development of in vitro-matured porcine oocytes following chemical activation or in vitro fertilization. Biol Reprod 1994; 50:1072–1077.[Abstract]
  44. Niwa K. Effectiveness of in vitro maturation and in vitro fertilization techniques in the pig. J Reprod Fertil Suppl 1993; 48:49–59.[Medline]
  45. Yanagimachi R. Mammalian fertilization. In: Knobil E, Neill JD, Ewing LL, Greenwald GS, Markert CL, Pfaff DW (eds.), The Physiology of Reproduction, vol. 1. New York: Raven Press; 1988: 135–185.
  46. Cran DG, Cheng WTK. The cortical reaction in pig oocytes during in vivo and in vitro fertilization. Gamete Res 1988; 13:41–51.
  47. Wang WH, Niwa K, Okuda K. In-vitro generation of pig oocytes matured in culture by frozen-thawed ejaculated spermatozoa. J Reprod Fertil 1991; 93:491–496.[Abstract]
  48. Zheng YS, Sirard MA. The effect of sera, bovine serum albumin and follicular cells on in vitro maturation and fertilization of porcine oocytes. Theriogenology 1992; 37:779–790.
  49. Norbury C, Nurse P. Animal cell cycles and their control. Annu Rev Biochem 1992; 61:441–470.[CrossRef][Medline]
  50. Crosby IM, Osborn JC, Moor RM. Changes in protein phosphorylation during the maturation of mammalian oocytes in vitro. J Exp Zool 1984; 229:459–466.[CrossRef][Medline]
  51. Kastrop PMM, Bevers MM, Destru OHJ, Kruip ThAM. Changes in protein synthesis and phosphorylation patterns during bovine oocyte maturation in vitro. J Reprod Fertil 1990; 90:305–310.[Abstract]
  52. Brizuela L, Draetta G, Beach D. Activation of human cdc2 protein as a histone H1 kinase is associated with complex formation with the p62 subunit. Proc Natl Acad Sci USA 1989; 86:4362–4366.[Abstract/Free Full Text]
  53. Draetta G, Beach D. Activation of cdc2 protein kinase during mitosis in human cells: cell cycle-dependent phosphorylation and subunit rearrangement. Cell 1988; 54:17–26.[CrossRef][Medline]
  54. Murray AW. Creative blocks: cell-cycle checkpoints and feedback control. Nature 1992; 359:599–604.[CrossRef][Medline]
  55. Nurse P. Universal control mechanism regulating onset of M-phase. Nature 1990; 344:503–507.[CrossRef][Medline]
  56. Gurley LR, D'Anna JAA, Barham SS, Deaven LL, Tobey RA. Histone phosphorylation and chromatin structure during mitosis in Chinese hamster cells. Eur J Biochem 1978; 84:1–15.[Medline]
  57. Peter M, Sanghera JS, Pelech SL, Nigg EA. Mitogen activated protein kinases phosphorylate nuclear lamins and display sequence specificity overlapping that of mitotic protein kinase p34cdc2. Eur J Biochem 1992; 205:287–294.[Medline]
  58. Verde F, Dogterom M, Stelzer E, Karsenti E, Leibler S. Control of microtubule dynamics and length by cyclin A-dependent and cyclin B-dependent kinases in Xenopus eggs extracts. J Cell Biol 1992; 118:1097–1108.[Abstract/Free Full Text]
  59. Kubelka M, Motlik J, Fulka J Jr, Procházka R, Rimkevicová Z, Fulka J. Time sequence of germinal vesicle breakdown in pig oocytes after cycloheximide and p-aminobenzamide block. Gamete Res 1988; 19:423–431.[CrossRef][Medline]
  60. Jagiello G, Ducayen M, Goonan W, Downey S. Inhibition of bovidae oocyte meiosis by serine protease inhibitor. Proc Soc Exp Biol Med 1978; 157:550–532.[Medline]
  61. Fleming AD, Khalil W, Armstrong DT. Porcine follicular fluid does not inhibit maturation of rat oocytes in vitro. J Reprod Fertil 1983; 69:665–670.[Abstract]
  62. Guerrier P, Moreau M, Dorée M. Inhibition de la réinitiation de la méiose des ovocytes de Xenopus laevis par trois antiproteases naturelles, l'antipaine, la chymostatise et la leupeptine. CR Acad Sci Paris 1977; 284:317–319.
  63. Clark TG, Kanatnai H. Effect of some inhibitors of proteolytic enzymes on maturation of starfish oocytes. Biol Bull 1975; 149:423.
  64. Murray AW, Kirschner MW. Cyclin synthesis drives the early embryonic cell cycle. Nature 1989; 339:275–280.[CrossRef][Medline]
  65. Murray AW, Solomon MJ, Kirschner MW. The role of cyclin synthesis and degradation in the control of maturation promoting factor activity. Nature 1989; 339:280–286.[CrossRef][Medline]
  66. Solomon MJ, Glotzer M, Lee TH, Philippe M, Kirschner MW. Cyclin activation of p34cdc2. Cell 1990; 63:1013–1024.[CrossRef][Medline]
  67. Naito K, Hawkins C, Yamashita M, Nagahama Y, Aoki F, Kohmoto K, Toyoda Y, Moor RM. Association of p34cdc2 and cyclin B1 during meiotic maturation in porcine oocytes. Dev Biol 1995; 168:627–634.[CrossRef][Medline]



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