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Departments of Community and Environmental Medicine (M.T.-T., U.L.), Medicine (U.L.), and Developmental and Cell Biology (U.L.), University of California Irvine, Irvine, California 92617
Address all correspondence and requests for reprints to: Dr. Ulrike Luderer, Center for Occupational and Environmental Health, University of California Irvine, 5201 California Avenue, Suite 100, Irvine, California 92617. E-mail: uluderer{at}uci.edu.
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Reactive oxygen species (ROS), such as superoxide anion, hydrogen peroxide, and hydroxyl radical, are produced as byproducts of normal cellular metabolism (10). ROS can react spontaneously with DNA, RNA, protein, and lipids. Oxidative stress occurs when levels of ROS overwhelm the cell’s antioxidant defenses. Electron transport associated with steroidogenesis is an important site of oxygen radical generation in the adrenal gland (11) and testis (12) and likely is also in the ovary. Inadequate protection from ROS that are formed in steroidogenically active granulosa cells could be a potential trigger for follicular atresia. ROS promote apoptosis in various model systems (13), but the role of ROS in apoptosis in ovarian follicles has received limited attention. Although apoptosis induction by treatment with ROS has not been investigated in ovarian follicles, treatment with hydrogen peroxide is toxic to granulosa cells in that it inhibits FSH-stimulated cAMP accumulation and progesterone production (14). Indirect evidence for a proapoptotic role of ROS in ovarian follicles comes from the observation that treatment with several antioxidants prevents apoptosis in cultured follicles (15).
The tripeptide glutathione (GSH, 
-glutamylcysteinylglycine) detoxifies ROS and electrophilic toxicants by spontaneously reacting with them or by providing reducing equivalents for enzymes like glutathione peroxidase and glutathione-S-transferases (16). The reversible glutathionylation of cellular proteins is also an important regulatory mechanism of protein function (17). Under normal conditions, the reduced form of GSH is present far in excess of the oxidized form (GSSG), because of the action of GSH reductase, as well as de novo synthesis of GSH (18). In situations of oxidative stress, the ratio of GSH to GSSG decreases. GSH is synthesized by the successive action of glutamate cysteine ligase (GCL) and GSH synthetase. GCL, the rate-limiting enzyme, is composed of a catalytic (GCLC) and a modifier (GCLM) subunit (18, 19). Gonadotropin hormones regulate ovarian expression of GCL subunit protein and mRNA (20, 21). A gonadotropin stimulus that decreases ovarian apoptosis in vivo (5) also significantly increases ovarian GCLC and GCLM protein levels (20, 21). Depleting GSH by blocking its synthesis induces apoptosis directly (22, 23, 24, 25) in some cell types and sensitizes other cell types to apoptotic stimuli (26, 27). We recently demonstrated that suppression of GSH synthesis in vivo increases atresia of antral follicles in rats (28). Other studies have shown that treatments that enhance cellular GSH concentrations protect against apoptotic stimuli in various cell types (29, 30).
Taken together, these previous studies led us to hypothesize that GSH, under regulation by gonadotropins, serves to protect ovarian follicles against ROS that could cause apoptosis if left unchecked. The present studies used a well-characterized model of follicular apoptosis in which cultured antral follicles undergo apoptosis in the absence of gonadotropin support, and follicles treated with gonadotropin are rescued from apoptosis (3, 4, 5, 31, 32). This model was used to test the specific hypotheses that 1) FSH enhances GSH synthesis in cultured preovulatory follicles, 2) GSH protects follicles against the proapoptotic effects of ROS and, therefore, GSH depletion promotes apoptosis, and 3) intrafollicular ROS rise in the absence of FSH, leading to the initiation of apoptosis.
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Materials and Methods |
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Animals
The 22- to 23-d-old female Sprague Dawley rats weighing 40–51 g were obtained from Charles River Laboratories (Wilmington, MA). Upon arrival, two to three animals per cage were housed in an Association for Assessment and Accreditation of Laboratory Animal Care accredited facility, with free access to deionized water and standard laboratory chow, on a 14-h light, 10-h dark cycle. The experimental protocols were carried out according to the Guide for the Care and Use of Laboratory Animals (33) and were approved by the Institutional Animal Care and Use Committee at the University of California, Irvine.
Preovulatory follicle isolation and culture
Isolation and culture of preovulatory follicles were adapted from previously described methods (5, 31). This whole-follicle culture system preserves the interactions among the oocyte, granulosa cells, and theca cells that are present in vivo. The 25- to 26-d-old female rats were injected sc with 10 IU PMSG in 0.1 ml sterile 0.9% NaCl to promote growth of a cohort of preovulatory follicles (34). PMSG has both FSH and LH activity in the rat (35). Forty-eight hours after the PMSG treatment, the animals were euthanized for collection of ovaries using aseptic technique. Ovaries were placed on ice in serum-free medium pregassed with 95% O2 and 5% CO2. Healthy preovulatory follicles (eight to 12 per ovary) 600–800 µm in diameter were isolated using iris forceps and 270-gauge needles under a dissecting microscope using aseptic technique. To minimize spontaneous apoptosis during follicle preparation, only one ovary was processed at one time, and isolated follicles were kept at 4 C until ready to culture.
Isolated preovulatory follicles were pooled, and three to six follicles per treatment group per experiment were cultured in 2 ml of 1) MEM (Eagle’s MEM, supplemented with 2 mM L-glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin sulfate, and 0.1% fatty acid-free BSA) alone, 2) MEM plus 75 ng/ml ovine FSH (0.33 IU/ml), or 3) 75 ng/ml FSH and 100 µM buthionine sulfoximine (BSO), a specific inhibitor of GCL, the rate-limiting enzyme in GSH synthesis (36), in sterile 20-ml glass scintillation vials. Each vial was gassed with 95% O2 and 5% CO2 every 12 h, sealed with petroleum jelly, covered with Parafilm (Pechiney Plastic Packaging, Menasha, WI), and incubated in a shaking water bath at 37 C for varying durations of time ranging from 2–48 h. Two or three replicate experiments were performed per time point for each end point. Each replicate experiment had its own 0-h control group of follicles that were processed immediately after dissection without culturing and that served as negative controls for apoptosis. The MEM groups served as positive control groups for apoptosis. The dose of ovine FSH was chosen based on a dose-response experiment that compared the amount of oligonucleosomal DNA fragmentation in 0-h control follicles (no apoptosis) with follicles cultured for 24 h with one of five different doses of ovine FSH (5, 25, 75, 200, and 500 ng/ml). DNA fragmentation was assessed by ethidium bromide gel electrophoresis. The results indicated that the antiapoptotic effect of FSH was maximal at 75 ng/ml (data not shown). The dose of BSO was chosen based on preliminary dose-response experiments using cultured granulosa cells collected from preovulatory follicles of PMSG-primed immature rats. Treatments with 100, 250, and 500 µM BSO for 24 h all suppressed GSH below the limit of detection in our assay (0.04 nmol; data not shown). Therefore, the 100-µM dose of BSO was used in the present studies. An additional experiment tested whether the effect of BSO treatment on follicular apoptosis was specifically caused by GSH depletion. There were four experimental groups: 1) 75 ng/ml FSH, 2) FSH plus 100 µM BSO, 3) FSH plus BSO plus 1 mM GSH ethyl ester (GEE, a cell-permeable analog of GSH) (37), or 4) FSH plus BSO plus 5 mM GEE. After 48 h, follicles were processed for GSH assay or terminal dUTP transferase-mediated nick-end-labeling (TUNEL).
At the time of collection, follicles were processed for several assays as follows. For GSH assay, five or six follicles were immediately homogenized on ice in 65 µl 5% sulfosalicylic acid, incubated on ice for 15 min, and centrifuged at 14,000 x g for 2 min at 4 C; supernatants were then stored at –70 C. For TUNEL assay and immunohistochemistry, four follicles were pre-fixed in 4% paraformaldehyde in PBS for 30 min at 4 C, dehydrated in 15% sucrose in PBS for 90 min at 4 C, embedded in Tissue Tek OCT (Sakura Finetek, Torrance, CA), and stored at –70 C until sectioning at 10 µm thickness using a cryostat. For DNA extraction, DNA was extracted from pools of six follicles immediately after termination of culture. For protein extraction, follicles were immediately homogenized in RIPA lysis buffer (1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS in PBS) with protease inhibitors on ice. After incubation on ice for 30 min, lysates were centrifuged at 15,000 x g, and the supernatants were stored at –70 C. The cell viability WST-1 assay and the ROS assays were performed immediately after termination of culture.
Cell viability WST-1 assay
Follicle viability was based on the cleavage of the tetrazolium salt WST-1 to formazan by mitochondrial dehydrogenases of viable follicles. Follicle viability was assessed after 0, 24, or 48 h of culture (n = 12 follicles per treatment group per time point). One follicle from each culture vial was transferred to a well of a 96-well plate containing 200 µl MEM and 20 µl WST-1 reagent. Blank wells contained culture medium and WST-1 reagent with no follicle. The plate was incubated for 4 h at 37 C in a humidified atmosphere of 95% air, 5% CO2. The absorbance at 440 nm was then read using a VersaMax tunable microplate reader (Molecular Devices, Sunnyvale, CA). The average of the blank absorbance values was subtracted from the absorbance value for each well.
Total GSH assay
Supernatants from homogenized follicles were analyzed for total GSH content using a microplate enzymatic assay, modified from Griffith (20, 21, 38). Briefly, 5 µl of sample or GSH standards were pipetted in triplicate with 33 µl deioinized water. The microplate was incubated for 10 min at 30 C. Reaction mixture (162 µl of 0.62 U/ml GSH reductase, 0.26 mM NADPH, and 0.74 mM 5,5'-dithiobis(2-nitrobenzoic acid) in a buffer of 143 mM Na2HPO4 and 6.3 mM EDTA) was added, and color development was monitored every 10 sec for 6.5 min at 412 nm using a Versamax tunable microplate reader (Molecular Devices). The sensitivity of this assay in our hands is 0.04 nmol. The samples from the 24- and 48-h experiments were assayed in one assay, and the samples from the 4- and 12-h experiments were assayed in a second assay. The intraassay coefficients of variation were 10.9 and 6%, respectively. The average GSH content per follicle was calculated as follows. The total number of nanomoles of GSH in the 65-µl sample homogenate was calculated from the concentration determined from the assay (concentrations that were undetectable were arbitrarily set equal to half the limit of detection of 0.008 mM). The nanomoles of GSH were then divided by the number of follicles in the homogenate (five or six follicles per sample that had been cultured in the same vial).
Western blotting
Gel electrophoresis and Western blotting were carried out as previously described (20, 21). Briefly, 20 µg of protein extracts were separated by electrophoresis in 12% Tris-HCl polyacrylamide gels (Bio-Rad, Hercules, CA) and transferred to polyvinylidene difluoride membranes. After transfer, blots were routinely stained with Ponceau Red to verify equal protein loading. Immunostaining was performed using GCLC and GCLM antibodies (39). Each blot was subsequently reprobed using ß-actin antibody (Sigma-Aldrich) as another loading control. After incubation with the appropriate horseradish peroxidase-conjugated secondary antibody, visualization was accomplished by ECL with ECL reagents and exposure to Hyperfilm ECL. Semiquantitative analysis of images was accomplished using a Stratagene molecular documentation and image analysis system with EagleSight software. Densitometry values for GCLC and GCLM bands were normalized to ß-actin values. The average normalized value for the 0-h controls on a given blot was then calculated, and the normalized absorbance for each band was expressed as fold of the average 0-h control value for that blot.
Oligonucleosomal DNA fragmentation detection
DNA was extracted from follicles using the DNeasy tissue kit according to the manufacturer’s instructions. After DNA extraction, 100 ng DNA per sample was 3'-end labeled with [
32P]dideoxynucleotide-ATP (3000 Ci/mmol; Amersham) using terminal transferase enzyme, followed by ethanol precipitation and resuspension in Tris-EDTA buffer according to the method described by Tilly and Hsueh (40).
One hundred nanograms of labeled DNA per lane were loaded onto a 2% agarose gel made with modified Tris-acetate-EDTA buffer. DNA samples were separated by electrophoresis at 50 V for 90 min. After electrophoresis, the gel was placed on Whatman filter paper and dried in a gel dryer at 50 C for 45 min. Dried gels were sealed in plastic bags and placed on Biomax MS film (Eastman Kodak, Rochester, NY) with an intensifying screen for 5 min or longer, depending on the signal intensity. After gel electrophoresis and autoradiographic analysis, the low-molecular-weight portion of each lane of the gel was cut out with a razor blade. The total amount of radiolabel incorporated into the low-molecular-weight DNA portions of the gel was quantified by scintillation counting to estimate the degree of apoptotic DNA fragmentation in any given sample as described (40). The counts per minute for each of the MEM, FSH, and FSH- plus BSO-treated samples were divided by the average of the MEM-positive controls for the same experiment. These fold control values were used for statistical analysis and for presentation of the data.
TUNEL assay for in situ detection of apoptosis
TUNEL was performed using the in situ cell death detection kit as described (41). Briefly, slides were blocked with 3% H2O2 in methanol, permeabilized with 0.1% Triton X-100 in 0.1% sodium citrate, blocked with 3% BSA, and incubated with terminal deoxynucleotidyl transferase solution with fluorescein-labeled dUTP. After washing, sections were again blocked with 3% BSA, washed, and incubated with peroxidase converter solution, followed by color development with diaminobenzidine. Last, slides were counterstained with hematoxylin, dehydrated in graded ethanol and xylene washes, and mounted with Permount. Negative control slides incubated with label solution without terminal transferase and positive control slides pretreated with DNase 1 were included with each experiment.
For quantification of apoptosis, the numbers of TUNEL-positive granulosa cells or theca cells were counted in a blinded manner in five x400 fields per follicle using an Olympus BX60 microscope. The averages of the five granulosa cell counts and the five theca cell counts were calculated for each follicle, and these values were used for statistical analyses.
Immunohistochemistry
Immunohistochemistry was performed essentially as described (21). For antigen retrieval, slides were heated at 80% power in a conventional microwave in 1 mM sodium citrate, 1 mM citric acid buffer for 10 min. The slides were blocked with 3% hydrogen peroxide in methanol for 10 min, 1.5% goat serum in PBS for 20 min, and avidin D and biotin blocking solutions for 15 min each. The slides were incubated with primary anti-cleaved caspase 3 antibody (1:5) in PBS overnight at 4 C, washed, incubated with biotinylated goat antirabbit IgG in PBS for 30 min, washed, incubated with ABC reagents for 30 min, washed, and incubated with diaminobenzidine substrate for 10 min. Finally, the slides were counterstained with hematoxylin, dehydrated in graded ethanol washes, washed in xylene, and mounted with Permount. Negative controls included slides incubated with nonimmune rabbit serum without primary antibody and slides incubated in primary antibody that had been preincubated with activated caspase 3 peptide.
For quantification of caspase 3 activation, the numbers of cleaved caspase 3-positive granulosa cells or theca cells were counted in a blinded manner in five x400 fields per follicle using an Olympus BX60 microscope. The averages of the five granulosa cell counts and the five theca cell counts were calculated for each follicle, and these values were used for statistical analyses.
In situ ROS detection
Preovulatory follicles were subjected to the same treatments as for the previous experiments, except that a hydrogen peroxide-treated group (5% H2O2 for 0.5 h) was added as a positive control for oxidative stress. After 0, 2, 4, 8, 12, 24, or 48 h of culture, treatment media were removed, and the follicles were washed with PBS. The follicles were then incubated with 100 µM 2',7'-dichlorofluorescin diacetate (H2DCFDA) (Molecular Probes, Eugene OR) in MEM for 30 min or with 100 µM dihydrorhodamine 123 (DHR) (Molecular Probes) for 30 min, and were washed again with PBS. H2DCFDA is taken up by cells, where it is converted by esterases to the nonfluorescent compound dichlorofluorescin, which in the presence of ROS is oxidized to the fluorescent dichlorofluorescein (DCF). DHR is a nonfluorescent compound that diffuses into cells, where it is oxidized in the presence of ROS to the cationic and fluorescent compound rhodamine 123 that localizes to mitochondria (42, 43). DCF or DHR fluorescence was viewed and quantified using a Zeiss LSM 511 META laser scanning confocal microscope using a Plan-Neofluor 10x/0.3 objective with Cy2/alexa/fluorescein isothiocyanate excitation and a BP-500–530 filter for emission. The follicle was scanned and the fluorescence intensity was measured for 15 equally spaced layers from the top to the bottom of the follicle (10–25 µm thickness). The area under the curve calculated from the histogram of the 15 DCF or DHR fluorescence intensity measurements per follicle was used for statistical analyses and presentation of data.
Statistical analyses
Data from replicate experiments were pooled for statistical analyses. Differences in GSH concentrations, low-molecular-weight DNA laddering, TUNEL cell counts, activated caspase 3 cell counts, and cell viability among treatment groups (0 h, MEM, FSH, and FSH plus BSO) were evaluated by one-way ANOVA for each time point (4, 12, 24, and 48 h). Differences in ROS measurements after various times in culture (0, 2, 4, 8, and 12 h) were assessed separately for each treatment group (MEM, FSH, and FSH plus BSO) and for each time point by one-way ANOVA. If the overall P value indicated statistical significance (P < 0.05), post hoc least significant difference (LSD) tests were applied to assess differences among groups based on a priori hypotheses. Levene’s test was used to assess homogeneity of variances. If variances were not homogeneous, a square root transformation was applied to the data before ANOVA. If transformation did not result in homogeneous variances, a nonparametric test, the Kruskal-Wallis test, was used to assess the overall effect of treatment. If the latter was statistically significant at P < 0.05, the Mann-Whitney test was then used for intergroup comparisons. SPSS version 11.0 for Macintosh was used for all statistical analyses.
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). Treatment with 75 ng/ml FSH not only prevented the decline in follicular GSH, but also increased GSH concentrations to greater than 0-h control levels at 12, 24, and 48 h of culture (P < 0.05, FSH at 12, 24, and 48 h vs. respective 0-h control; Fig. 1A). By 48 h of FSH treatment, GSH concentrations reached 2.4 ± 0.5 nmol/follicle. Treatment with 100 µM BSO, a specific inhibitor of GCL enzymatic activity, suppressed GSH concentrations to less than 15% of 0-h control levels after 12 h and to less than 10% of 0-h control levels after 24 and 48 h (P < 0.01, BSO at 12, 24, and 48 h vs. respective 0-h control).
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). Cotreatment with BSO did not modify the enhancement of GCLM protein levels by FSH. Levels of GCLC did not differ among treatment groups. GCL subunit protein levels were assessed after 24 h of treatment only.
Treatment with BSO to deplete GSH partially reverses the antiapoptotic effect of FSH on granulosa cells of cultured preovulatory follicles, and supplementation of GSH completely prevents the BSO-induced apoptosis
Preovulatory follicles cultured without gonadotropin support begin to undergo apoptosis within 24 h, whether cultured in the absence (3) or presence (32) of serum. FSH treatment prevents apoptosis in these follicles (3). To investigate whether the enhancement of follicular GSH levels by FSH plays a role in the suppression of apoptosis by FSH, we depleted GSH with BSO in the presence of FSH.
Using TUNEL to localize apoptotic cells within cultured follicles, we observed differential effects on granulosa cells and theca cells of FSH treatment alone compared with combined FSH and BSO treatment (Figs. 2 and 3). Culture in serum-free medium caused a marked, time-dependent increase in both granulosa cell and theca cell apoptosis as judged by TUNEL, with no increase at 4 h, but statistically significant increases at 12, 24, and 48 h (P < 0.001, MEM at 12, 24, and 48 h vs. respective 0 h; Figs. 2 and 3, A and B). FSH treatment brought about a statistically significant decrease in the number of TUNEL-positive granulosa cells at all three later time points (P < 0.001, effect of treatment by ANOVA for 12-, 24-, and 48-h experiments, with FSH different from MEM at P < 0.01 by LSD test). In contrast, FSH treatment enhanced theca cell apoptosis at 24 h (P < 0.001, FSH vs. MEM) but not at 4, 12, or 48 h (Fig. 3B).
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and 3A). The number of TUNEL-positive granulosa cells was significantly increased in FSH- plus BSO-treated groups compared with FSH treatment alone at 24 and 48 h (P = 0.03 and P < 0.001, BSO vs. FSH at 24 and 48 h, respectively; Fig. 3A). The magnitude of granulosa cell apoptosis in the FSH plus BSO group at 48 h was similar to that in the MEM group at 24 h. BSO enhanced the proapoptotic effect of FSH on theca cell apoptosis at 24 h (P = 0.04, BSO vs. FSH at 24 h), but did not enhance apoptosis at 4, 12, or 48 h (Figs. 2 and 3B). To test whether the BSO-induced granulosa cell apoptosis was a specific effect of GSH depletion, we cultured follicles for 48 h with FSH, FSH plus BSO, or FSH plus BSO plus GEE, a cell-permeable analog of GSH. Treatment with 5 mM GEE completely prevented the suppression of follicular GSH concentrations by BSO. GSH concentrations were 2.3 ± 0.3, 0.04 ± 0.0, and 2.4 ± 0.3 nmol/follicle in the FSH, FSH plus BSO, and FSH plus BSO plus 5 mM GEE groups, respectively. The 5 mM GEE dose also completely prevented the BSO-induced increase in granulosa cell apoptosis (Fig. 3C). Both the 1 and 5 mM doses of GEE suppressed theca cell apoptosis in the presence of BSO (Fig. 3C).
We next used an antibody directed against the activated (cleaved) form of caspase 3 as an additional marker of apoptosis. Caspase 3 is activated by both the mitochondrial and the receptor-mediated apoptotic pathways (44). The numbers of activated caspase 3-positive granulosa cells and theca cells increased in a time-dependent manner upon culture in the absence of gonadotropin support. The increase in caspase 3-positive granulosa cells was statistically significant at 4, 12, 24, and 48 h of culture (P < 0.008, MEM vs. respective 0-h controls; Fig. 4B). The increase in caspase 3-positive theca cells was statistically significant at 24 and 48 h (P < 0.001, MEM vs. respective 0-h controls; Fig. 4C). Treatment with FSH caused a significant decline in caspase 3-positive granulosa cells at 12, 24, and 48 h (P < 0.03, FSH vs. MEM). In contrast, FSH increased theca cell caspase 3 activation at 24 and 48 h (P < 0.001, FSH vs. MEM). Combined FSH plus BSO treatment partially overcame the suppressive effect of FSH on activated caspase 3 immunostaining in granulosa cells, and the difference between FSH plus BSO and FSH-treated groups was statistically significant at both 24 and 48 h (P < 0.007; Fig. 4). Thus, activated caspase 3 immunostaining closely paralleled the results with TUNEL (Fig. 3), providing additional evidence that GSH depletion induces apoptosis in granulosa cells of cultured preovulatory follicles.
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). No oligonucleosomal DNA fragmentation was observed in 0-h control follicles (Fig. 5A). Treatment with FSH significantly suppressed oligonucleosomal DNA fragmentation by 67 and 60% at 24 and 48 h, respectively (Fig. 5B; P < 0.001, effect of treatment by ANOVA at 24 and 48 h). Depletion of GSH with BSO partially reversed the antiapoptotic effect of FSH, but this was not statistically significant (P = 0.07 and P = 0.16, BSO vs. FSH at 24 and 48 h, respectively). The absence of a statistically significant effect of GSH depletion on whole-follicle DNA laddering compared with the significant effects observed using two in situ assays for apoptosis, TUNEL and immunostaining for activated caspase 3, highlights the greater sensitivity of the in situ assays.
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; P < 0.001, MEM at 24 and 48 h vs. 0 h). After 24 h of culture, WST-1 cleavage in FSH- and FSH- plus BSO-treated follicles was also decreased compared with 0-h control follicles but did not differ from MEM-treated follicles. Interestingly, at 48 h, FSH-treated follicles had significantly greater WST-1 cleavage than 0- or 48-h MEM controls (P < 0.001, FSH vs. MEM at 0 h; Fig. 5). BSO treatment partially reversed the stimulatory effect of FSH on WST-1 cleavage at 48 h, with WST-1 absorbance in FSH- plus BSO-treated follicles not statistically different from the absorbance in the 0-h controls (Fig. 6). These results again paralleled the effects of BSO and FSH observed on apoptosis, further demonstrating the ability of GSH depletion to reverse the protective effect of FSH on preovulatory follicles.
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Culture in serum-free medium led to a statistically significant increase in DCF fluorescence by 4 h of culture, with levels remaining elevated above baseline 0-h levels at all subsequent time points after the initiation of culture (P < 0.01, MEM-treated at 4, 8, 12, 24, and 48 h of culture vs. 0 h; Fig. 7, A and B). These increases in DCF fluorescence were of similar magnitude as the 8.0 ± 1.8-fold increase observed after 30 min of exposure to 5% hydrogen peroxide. FSH treatment attenuated the rise in ROS production. DCF fluorescence levels at 4 and 24 h after FSH treatment were significantly suppressed compared with MEM treatment at the same time points (P < 0.05, FSH vs. MEM at 4 and 24 h; Fig. 7, A and B). Cotreatment with FSH and BSO appeared to slightly reverse the suppressive effect of FSH on ROS production (Fig. 7). However, the difference between FSH- and FSH- plus BSO-treated groups was not statistically significant at any time point.
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Taken together, the DCF and DHR data show that follicular ROS are significantly increased after 4 h of culture without gonadotropin support and that FSH suppresses ROS production. Both methods showed a partial reversal by BSO of the suppressive effect of FSH on ROS, but this was statistically significant only when DHR was used as a probe for ROS.
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Discussion |
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There is evidence that ROS play roles in the induction of both the death receptor and mitochondrial apoptotic pathways (13). Oxidative stress caused by the addition of exogenous oxidants induces apoptosis in many types of mammalian cells, including hepatocytes (46), epithelial cells (47), and fibroblasts (48). Endogenous production of ROS has also been associated with the induction of apoptosis (24). However, there is controversy as to whether ROS signaling is critical for the induction of apoptosis or is merely a by-product of apoptosis (13, 49). We assessed follicular ROS production using the probes H2DCFDA and DHR (42, 43, 45). Several lines of evidence show that DCF fluorescence indicates production of hydroxyl radical, peroxynitrite, and, in the presence of peroxidases, hydrogen peroxide (50). We observed significant increases in follicle DCF and DHR fluorescence by 4 h of culture in the absence of gonadotropin support (Figs. 7 and 8), which preceded the increase in granulosa cell apoptosis, consistent with the hypothesis that the increase in ROS initiates the apoptotic cascade in granulosa cells. The suppression of follicular ROS by FSH treatment (Figs. 7 and 8), which also inhibited granulosa cell apoptosis, further supports this hypothesis. Depletion of GSH with BSO in the presence of FSH partially reversed the suppression of ROS by FSH (Figs. 7 and 8). Finally, the observations that FSH treatment enhanced follicular levels of the antioxidant GSH (Fig. 1) and that GSH depletion partially reversed the antiapoptotic effects of FSH (Figs. 2–4) provide additional evidence that the rise in ROS initiates the apoptotic cascade in these follicles. There has been discussion in the literature that increases in DCF and DHR fluorescence may not indicate an increase in ROS but rather may be indicative of cytochrome c release, an early event of apoptosis (43, 51). These authors reported that cytochrome c is capable of catalyzing the oxidation of dichlorofluorescin by hydrogen peroxide (51) and of catalyzing the oxidation of DHR by hydrogen peroxide (43). It appears unlikely that the observed increases in DCF and DHR fluorescence in the present studies were a result of cytochrome c release in the presence of no change in cellular hydrogen peroxide levels. We observed no increase in cytochrome c in cytosolic preparations of follicles cultured for 4 h in MEM, compared with 0-h control follicles (data not shown). Activation of caspase 3 occurs almost concomitantly with cytochrome c release (52, 53), and we observed only a very small number of granulosa cells with activated caspase 3 immunostaining at 4 h (Fig. 4). In contrast, we observed large increases in DCF and DHR fluorescence throughout the follicles at 4 h of culture (Figs. 7 and 8).
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). In contrast, clear increases in granulosa cell apoptosis were evident at 12 h of culture in serum-free medium (Fig. 4), before the decline in follicular GSH levels.
Most previous studies that demonstrated antiapoptotic effects of FSH on follicular apoptosis measured oligonucleosomal DNA fragmentation in DNA from whole-follicle or whole-ovary homogenates (3, 4, 5, 55). These studies and the present study showed that FSH decreased apoptosis using this method. Using TUNEL to localize DNA fragmentation and immunostaining to localize activated caspase 3 within cultured preovulatory follicles, we have confirmed a previous report (32) that FSH exerts antiapoptotic effects on granulosa cells and proapoptotic effects on theca cells of cultured preovulatory follicles. In the present study, FSH suppressed granulosa cell apoptosis in preovulatory follicles at 12, 24, and 48 h of culture (Figs. 3A and 4B). In contrast, FSH enhanced theca cell apoptosis in preovulatory follicles at 24 and 48 h but not at 4 or 12 h (Figs. 3B and 4C). Coincident with its suppression of granulosa cell apoptosis, FSH treatment also enhanced follicular GSH concentrations, supporting a role for GSH in mediating the antiapoptotic effect of FSH on granulosa cells. The partial reversal of the antiapoptotic effect of FSH by the GSH synthesis inhibitor BSO (Figs. 3, A and C, and 4B) further supports this hypothesis. The complete prevention of the BSO-induced increase in granulosa cell apoptosis by GSH supplementation (Fig. 3C) demonstrates that this proapoptotic effect of BSO is specifically a result of GSH depletion. The increase in follicular GSH concentrations in response to FSH in the present study was associated with increased protein levels of GCLM, the rate-limiting enzyme in GSH synthesis (Fig. 1). Although protein levels of the catalytic subunit were not increased by FSH treatment, an isolated increase in the modifier subunit likely increased levels of the holoenzyme, which is a heterodimer of the catalytic and modifier subunits (56, 57). We have previously reported that ovarian protein and mRNA levels of GCLM are dramatically up-regulated by 4 h after an ovulatory dose of human chorionic gonadotropin in PMSG-primed rats (21). The present data provide evidence that this isolated increase in GCLM levels in vivo may lead to localized increases in GSH synthesis in preovulatory follicles.
In addition to up-regulating ovarian GSH synthesis, gonadotropins also enhance other antioxidant responses in the ovary. Ovarian mRNA levels of secreted and manganese-containing superoxide dismutase increase after PMSG treatment in rats (15, 58). PMSG treatment also increases ovarian concentrations of vitamin A (59). FSH enhances uptake of ascorbic acid by cultured granulosa cells (60). Up-regulation of these other antioxidant responses in ovarian follicles by FSH may explain why GSH depletion in the present studies did not fully overcome the suppressive effect of FSH on follicular ROS generation or granulosa cell apoptosis in the present study.
Our results are consistent with previous findings that activation of the executioner caspase, caspase 3, is involved in both granulosa and theca cell apoptosis in cultured preovulatory follicles. Our data for activated caspase 3 immunostaining in granulosa and theca cells of preovulatory follicles mirrored our TUNEL data. These data are consistent with previous in vivo observations that procaspase 3 is expressed in granulosa cells of atretic follicles, but not of healthy follicles, in the rat ovary (61) and that activated caspase 3 is expressed in granulosa cells of atretic, but not health,y follicles in the mouse ovary (9). Moreover, a critical role for caspase 3 in granulosa cell apoptosis in the mouse ovary has been demonstrated, because caspase 3 knockout mice have delayed and abnormal granulosa cell apoptosis (9). Because granulosa cells possess FSH receptors, but theca cells do not (62), the effects of FSH on apoptosis in granulosa cells are likely to be direct, whereas the effects of FSH on theca cells are likely to be indirect. FSH treatment in vivo has been shown to suppress apoptotic protease activating factor-1 (APAF-1) protein expression and bax mRNA expression in granulosa cells (6, 8). Because both BAX and apoptotic protease activating factor-1 act upstream of caspase 3 activation in the mitochondrial apoptotic pathway, these data collectively show that FSH acts to inhibit several key points in this pathway in granulosa cells. The present study shows that the full antiapoptotic effect of FSH on granulosa cells cannot occur in the face of follicular GSH depletion. We are unaware of any studies that investigated paracrine signals by which FSH stimulates theca cell apoptosis in cultured follicles. However, FSH has been reported to enhance protein and mRNA expression of granzyme-like proteins in primary human and immortalized rat granulosa cells (63). Granzyme B released from granules within cells can act on adjacent cells to directly activate caspase 8 and other caspases (63), making granzyme-like proteins possible candidates as mediators of the proapoptotic effect of FSH on theca cells of cultured follicles.
In addition to measures of apoptosis, we also assessed follicle health using a cell viability assay. This assay measures metabolic activity, assessed by the cleavage of the tetrazolium salt WST-1 to formazan by mitochondrial dehydrogenases in viable cells. As expected, there was a decline in WST-1 cleavage in follicles cultured for 24 h in serum-free media compared with 0-h control follicles that were not cultured, indicating a decline in follicle viability. Surprisingly, we observed a statistically significant increase in WST-1 cleavage after 48 h of treatment with FSH, compared with 0-h control follicles. This implies that proliferation of follicular cells occurred upon FSH stimulation. However, granulosa cells from preantral, small antral, and preovulatory follicles do not proliferate in response to FSH stimulation alone in vitro (64, 65, 66), and cultured preantral follicles also show minimal growth in response to FSH alone (55). Although FSH does not promote the proliferation of isolated granulosa cells in vitro, it does enhance the growth-promoting effects of other factors, such as transforming growth factor ß (64), activin (65), and growth differentiation factor 9 (66), on granulosa cells. This suggests that FSH may stimulate the release of such paracrine or autocrine factors, which have mitotic actions, from granulosa or theca cells of cultured preovulatory follicles. BSO treatment in the presence of FSH reversed the stimulatory effect of FSH on WST-1 cleavage, consistent with the increased apoptotic granulosa cell death observed in the presence of BSO (Figs. 3 and 4). Another explanation for the increased WST-1 cleavage in the FSH-treated follicles is that FSH stimulated the activity of mitochondrial dehydrogenases. Although it is well established that FSH enhances the expression and activity of the mitochondrial cytochrome P450 cholesterol side chain cleavage enzyme (67, 68), there is little information available about the actions of FSH on other aspects of mitochondrial function. It was recently reported that whole-ovary mitochondrial content, mitochondrial oxygen uptake, mitochondrial electron transfer activities, and mitochondrial indicators of oxidative damage all increased in response to FSH in a hyperstimulated estrous cycle in adult rats (69). Thus, our observations that FSH increased WST-1 cleavage of preovulatory follicles may be related to up-regulation of mitochondrial activity by FSH. It has recently been shown that mitochondrial NADP+-dependent isocitrate dehydrogenase enzymatic activity is suppressed by glutathionylation (70). Glutathionylation of proteins is favored when the cellular redox balance of reduced to oxidized glutathione (GSH:GSSG) shifts toward GSSG (17). This occurs in situations of oxidative stress and GSH depletion (17). ROS have also been shown to induce glutathionylation of proteins even in the absence of a detectable change in GSH:GSSG (17). Thus, FSH could enhance the activity of mitochondrial dehydrogenases by shifting cellular redox balance toward reduced glutathione and decreased glutathionylation.
In summary, we have shown that culture of preovulatory follicles in the absence of gonadotropin support increased follicular ROS levels by 4 h and increased granulosa cell and theca cell apoptosis by 12 h. Treatment with FSH delayed and decreased the production of ROS, enhanced follicular GSH concentrations by 12 h, and suppressed granulosa cell apoptosis by 12 h. Cotreatment with FSH and BSO, a specific inhibitor of GSH synthesis, partially and significantly reversed the suppression of ROS production and granulosa cell apoptosis by FSH. Collectively, our results support the hypotheses that apoptosis in cultured preovulatory follicles is initiated by oxidative stress and that the antioxidant GSH plays a role in mediating the antiapoptotic effect of FSH on granulosa cells of preovulatory follicles. Our results further suggest that abnormalities in follicular glutathione systems could decrease the ability of follicles to respond to oxidative stress or toxicant exposures, causing increased follicular apoptosis and even ovarian failure.
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Acknowledgments |
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Author disclosure summary: M.T.-T. and U.L. have nothing to declare.
First Published Online December 8, 2005
Abbreviations: BSO, Buthionine sulfoximine; DCF, dichlorofluorescein; DHR, dihydrorhodamine 123; ECL, enhanced chemiluminescence; GCL, glutamate cysteine ligase; GCLC, GCL catalytic subunit; GCLM, GCL modifier subunit; GEE, glutathione ethyl ester; GSH, glutathione; GSSG, GSH oxidized form; H2DCFDA, 2',7'-dichlorofluorescin diacetate; LSD, least significant difference; PMSG, pregnant mare serum gonadotropin; ROS, reactive oxygen species; TUNEL, terminal dUTP transferase-mediated nick-end-labeling.
Received October 11, 2005.
Accepted for publication November 30, 2005.
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B (NFB)-independent NFB activation in the gonadotropin regulation of X-linked inhibitor of apoptosis expression during ovarian follicular development in vitro. Endocrinology 143:2732–2740-glutamylcysteine synthetase. J Cell Physiol 182:163–170[CrossRef][Medline]
, significantly different from respective MEM-positive control; 
, significantly different from respective FSH-treated follicles at P < 0.05 by Mann-Whitney test. B, A representative Western blot shows follicular GCLM and GCLC protein levels after 24 h of treatment; ß-actin protein levels are shown as a loading control. The graph shows the mean ± SEM of GCLM and GCLC protein levels normalized to ß-actin and expressed as fold 0-h control levels for the same blot (n = 4 per group). There was a statistically significant effect of treatment on GCLM protein levels (P = 0.03 by Kruskal-Wallis test). *, Significantly different from MEM control by Mann-Whitney test.
