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BOR - Papers in Press, published online ahead of print March 3, 2004.
Biol Reprod 2004, 10.1095/biolreprod.103.016113
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BIOLOGY OF REPRODUCTION 71, 130–138 (2004)
DOI: 10.1095/biolreprod.103.016113
© 2004 by the Society for the Study of Reproduction, Inc.

Ovary

The Follicle-Deplete Mouse Ovary Produces Androgen1

Loretta P. Mayer3, Patrick J. Devine3, Cheryl A. Dyer4, and Patricia B. Hoyer2,3

Department of Physiology,3 University of Arizona, Tucson, Arizona 85724 Department of Biological Sciences,4 Northern Arizona University, Flagstaff, Arizona 86011


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The follicle-depleted postmenopausal ovary is enriched in interstitial cells that produce androgens. This study was designed to cause follicle depletion in mice using the industrial chemical, 4-vinylcyclohexene diepoxide (VCD), and characterize the steroidogenic capacity of cells in the residual ovarian tissue. From a dose-finding study, the optimal daily concentration of VCD was determined to be 160 mg/kg. Female B6C3F1 immature mice were treated daily with vehicle control or VCD (160 mg kg–1 day–1, 15 days, i.p.). Ovaries were removed and processed for histological evaluation. On Day 15 following onset of treatment, primordial follicles were depleted and primary follicles were reduced to about 10% of controls. On Day 46, primary follicles were depleted and secondary and antral follicles were reduced to 0.7% and 2.6% of control, respectively. Seventy-five percent of treated mice displayed disruptions in estrous cyclicity. All treated mice were in persistent diestrus (acyclic) by Day 58. Plasma FSH levels were increased (P < 0.05) relative to controls on Day 37 and had plateaued by Day 100. Relative to age-matched cyclic controls, by Day 127, the significant differences in VCD-treated mice included reduced ovarian and uterine weights, elevated plasma LH and FSH, and reduced plasma progesterone and androstenedione. Furthermore, plasma 17ß-estradiol levels were nondetectable. Unlike controls, immunostaining for LH receptor, and the high density lipoprotein receptor (SR-BI), was diffuse in ovarian sections from VCD-treated animals. Ovaries from Day 120 control and VCD-treated animals were dissociated and dispersed cells were placed in culture. Cultured cells from ovaries of VCD-treated animals produced less LH-stimulated progesterone than control cells. Androstenedione production was nondetectable in cells from cyclic control animals. Conversely, cells from VCD-treated animals produced androstenedione that was doubled in the presence of insulin and LH (1 and 3 ng/ml). Collectively, these data demonstrate that VCD-mediated follicle depletion results in residual ovarian tissue that may be analogous to the follicle-deplete postmenopausal ovary. This may serve as a useful animal model to examine the dynamics of follicle loss in women as ovarian senescence ensues.

aging, interstitial cells, steroid hormones, theca cells


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
At birth, the mammalian ovary contains a finite number of primordial follicles. The majority of these will never mature to ovulation. Rather, >99% undergo a process of degeneration, designated atresia. During the atretic process, granulosa cells and oocytes are eliminated via physiological cell death, apoptosis. In contrast, the theca interna cells contained in follicles undergoing atresia transition into interstitial cells [1]. Thus, as the ovary continues to lose follicles by atresia, there is an increase in the number of theca-derived interstitial cells, which are responsible for the androgenic contribution of the climacteric ovary. At that time, the cellular composition of the ovary shifts to an interstitial and stromal cell-rich tissue [2]. Therefore, residual ovarian interstitial and stromal tissue likely provides a physiological contribution by producing androgens during the perimenopausal transition and in postmenopause in women [3]. As suggested by Liu et al. [4], the postmenopausal ovary may contribute to an androgen excess, which may or may not impact many pathologies associated with menopause.

Previous studies have determined that repeated treatment of rats and mice with the industrial chemical 4-vinylcyclohexene diepoxide (VCD) selectively destroys primordial and primary ovarian follicles [5–8]. Following 15 days of VCD treatment in rats (80 mg kg–1 day–1, i.p.) there was a 50% loss of primordial follicles relative to controls [6]. The unhealthy appearance of follicles in VCD-treated ovaries was morphologically and ultrastructurally similar to follicles undergoing natural atresia in control animals [6, 9]. Additionally, VCD treatment activated several intracellular signaling pathways associated with apoptosis in isolated small preantral follicles [10–12]. For example, stimulation of the apoptotic branch of the Bcl2 family of proto-oncogenes culminated in activation of the protease caspase-3 [10]. Taken together, the findings demonstrate that VCD induces selective loss of small preantral (primordial and primary) follicles by accelerating apoptotic events associated with the normal process of atresia.

Two long-term studies were conducted in mice and rats (age 28 days) treated daily for 30 days with the parent compound 4-vinylcyclohexene (VCH, mice) or the active diepoxide metabolite (VCD, rats). The effects on reproductive function were evaluated at different time points for up to 360 days after the onset of treatment [9, 13]. Circulating levels of FSH were significantly elevated relative to control cyclic animals on Day 120 (rats) and Day 240 (mice), indicating the loss of hormonal negative feedback on the pituitary from the ovary. Acyclicity and ovarian atrophy had occurred by Day 360 in both species, whereas control animals were still cycling. Thus, premature ovarian failure via depletion of the primordial follicle pool can be induced in mice and rats by repeated injections with VCH or VCD. The long lag between dosing and ovarian failure in these studies resulted because primordial follicle loss had not been complete when treatment with those doses of chemicals was stopped on Day 30. Thus, a residual follicle pool (11% mice, 31% rats) was still present for recruitment for follicular development and ovulation. Studies performed over 2 yr of dosing of mice and rats by the National Toxicology Program showed no overt, systemic, or generalized toxic effects of the industrial chemical VCH or the active diepoxide metabolite VCD [14, 15]. However, there was a reduction in ovarian follicles in both studies.

It has been difficult to investigate the physiologic function of the ovarian interstitial compartment separate from the functional units, follicles and corpora lutea, due to a lack of experimental animal models that possess a follicle-deplete ovary. To better understand the function of interstitial cells, it would be desirable to accelerate atresia, causing the ovary to become prematurely depleted of follicles. Such an animal would be useful for studying endocrine function selectively in the residual interstitial cell-enriched ovarian tissue. Therefore, the studies presented here were designed to use increased concentrations of VCD to develop a novel VCD treatment protocol that rapidly and optimally accelerates follicle loss to provide a model of premature ovarian senescence in mice for subsequent characterization of the endocrine status of the residual ovarian (interstitial) tissue.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals

Immature female B6C3F1 mice (21 days) were obtained from Harlan Laboratories (Indianapolis, IN) and The Jackson Laboratories (Bar Harbor, ME), housed in plastic cages, and maintained on 12L:12D cycles at 22 ± 2°C. Animals were allowed to acclimate to the animal facilities for 1 wk prior to initiation of treatment. Food and water were available ad libitum. All experiments and killing by exsanguination under anesthesia were approved by the University of Arizona and Northern Arizona University Institutional Animal Care and Use Committees and conformed to the Guide for the Care and Use of Experimental Animals.

Treatment

The treatment protocol to use VCD to induce follicular depletion has a patent pending and VCD-treated mice are commercially available through the Jackson Laboratory (Bar Harbor, ME). Immature mice (Day 28) were randomly selected for each treatment group, weighed, and injected daily with VCD (80–320 mg/kg, 15 days, i.p.; Sigma-Aldrich, St. Louis, MO) or sesame oil vehicle control (n = 6/group per VCD concentration). In another experiment, animals were injected daily with vehicle control or VCD (160 mg/kg, 15 days, i.p.). On Days 8, 10, 12, 14, 15, 30, 37, 46, 58, 120, and 127, animals were weighed, killed, and the ovaries and uteri collected and weighed (n = 6/group per treatment day) Note: on all but Day 120, adrenals, kidneys, spleen, and livers were also collected and weighed. Estrous cycles of each animal were monitored daily at 0800 h by vaginal cytology from the date of vaginal opening to persistent diestrus in VCD-treated mice and from date of vaginal opening to Day 120 in sesame oil vehicle control mice. In animals on Days 8–58 and Day 127, one ovary from each animal was prepared for histological evaluation (n = six animals/group). The contralateral ovary from three animals per group was prepared for immunostaining and confocal microscopy. On Day 120, ovaries were collected for disassociation and cell culture incubations. Each control group contained ovarian cells from animals in estrus (four), diestrus-1 (three), diestrus-2 (six), and proestrus (two) at the time of killing (ovaries/group; n = three groups). Whole blood was collected by cardiac puncture and the plasma separated and stored at –20°C. Additionally, on Day 100 following initiation of treatment, whole-blood samples were collected by retro-orbital puncture and plasma was separated and stored at –20°C.

Caspase-3 activity was measured in three separate groups of animals (three replicates). In each replicate, six animals/group were treated as described above, and on Day 10, both ovaries from each animal were collected for isolation of small preantral follicles and analysis of caspase-3 activity.

For cell culture experiments, an additional 61 mice were treated (27 control, 34 VCD, 160 mg kg–1 day–1, 15 days, i.p.), and on Day 37, both ovaries from three control and four VCD-treated mice were collected and prepared for single-cell suspensions. On Day 120, an additional 54 mice were treated and both ovaries from eight control and 10 VCD-treated mice were collected for preparation of cell suspensions for culture (experiment repeated twice, n = 3). Each control replicate contained cells from the following number of ovaries collected from mice in the respective estrus stages: three–four (diestrus-1), six (diestrus-2), two (proestrus), and four (estrus).

Histology and Follicle Counting

The ovary was trimmed free of fat and placed in Bouin fixative (2 h), transferred to 70% ethanol, paraffin-embedded and serially sectioned (4– 5 µm), mounted, and stained with hematoxylin and eosin. Follicles were counted in every 20th section to avoid double counting of small preantral follicles (25–100 µm in diameter). Each ovary produced 10–13 sections [6, 7, 16]. Follicles were classified as primordial (oocyte surrounded by a single layer of flattened granulosa cells), small primary (oocyte surrounded by a single layer of <20 cuboidal cells with no oocyte growth), large primary (enlarged oocyte surrounded by a single layer of 21–26 cuboidal cells), secondary (oocyte surrounded by multiple layers of granulosa cells), or antral (follicles containing a fluid-filled antrum) and counted [16, 17]. In evaluation of antral follicle numbers, all were observed because they are >250 µm in diameter and every 20th section (5 µm/section) represents a distance of 100 µm.

Assessment of Hepatic Function

Hepatocellular histopathology and enzymatic activity of circulating aspartate aminotransferase (AST) and alanine aminotransferase (ALT) were determined by the Diagnostic Laboratory at the Arizona Health Sciences Center.

Follicle Isolation and Caspase-3 Protease Activity Measurement

Small preantral follicles, primordial and primary (25–100 µm in diameter), were prepared by gentle enzymatic dissociation of ovaries and sorting with micropipettes as previously described [16]. In each replicate, follicles were collected and pooled from both ovaries of six mice in each treatment (control or VCD; n = three replicates). Following isolation, follicles were washed twice with Medium 199 (M199) and a whole-cell homogenate was prepared. The cleavage activity of caspase-3 in isolated ovarian small preantral follicles was measured as described by Hu et al. [10]. Briefly, the enzymatic reaction was carried out at 37°C in protease assay buffer (20 mM HEPES, 100 mM NaCl, 10 mM dithiothreitol, 1 mM EDTA, 0.1% [w/v] 3-3 ([3-cholamidopropyl] dimethylammonio)-1-propanesulfonate, and 10% sucrose, pH 7.2). Cellular protein (60–180 µg) was incubated with 50 µM of caspase-3 substrate, Ac-DEVD-7-amino-4-methylcoumarin (AMC) for 60 min. Substrate cleavage was detected by measurement of the fluorescence of free AMC at 460 nm emission upon excitation at 380 nm.

Dispersed Cell Preparation and Culture

Ovaries were prepared as described by Dyer and Curtiss [18]. Briefly, tissue was trimmed, cut into small pieces, and incubated at 37°C in a shaking water bath for 90 min with 0.1 ml/ovary of a collagenase/DNase solution in M199 that contained 4 mg/ml of collagenase (Clostridium histolyticum, CLS type I, 144 units/mg; Worthington Biochemical Corp., Lakewood, NJ), 10 µg/ml of deoxyribonuclease (bovine pancreas, 2100 units/mg; Life Technologies, Rockville, MD), and 10 mg/ml of bovine serum albumin (Sigma, St. Louis, MO). Cells were pelleted by centrifugation at 500 x g for 5 min at room temperature, washed, resuspended in McCoys 5a medium and plated at 30 000–50 000 cells per well in a 96-well plate in 0.25 ml of serum-free McCoys 5a modified medium (Life Technologies, Rockville, MD) with penicillin and streptomycin, human high density lipoprotein (HDL; a gift from Dr. Carole L. Banka, La Jolla Institute for Molecular Medicine, San Diego, CA) at 100 µg/ml and incubated in a 95% air, 5% CO2 humidified incubator at 37°C [19]. The next day, luteinizing hormone (LH; NIDDK-oLH-25) at 1, 3, 10, 30, or 100 ng/ml ± bovine insulin (Gibco BRL, Rockville, MD) at 1 µg/ml were added to the media. Cells were incubated for an additional 48 h and supernatants were collected. Cells and media were stored at –20°C. The DNA content of each well was determined using the CyQuant DNA assay (Molecular Probes, Eugene, OR). Values for steroid levels from cell cultures were determined for each treatment well and were corrected for differences in DNA content by dividing individual steroid values for each well by the DNA content (µg) value for that well as determined by the CyQuant DNA assay.

Hormone Assays

Progesterone and androstenedione were assayed in culture supernatants in a fluid-phase assay in which free 3H-steroid was separated from that bound to antiserum by absorption onto activated charcoal [18]. Six wells were incubated for each treatment concentration within each treatment group and assayed individually. Plasma levels of 17ß-estradiol, androstenedione, and progesterone were determined by Coat-A-Count RIAs (Diagnostic Products Inc., Los Angeles, CA). Sensitivity and inter- and intraassay coefficients of variation were 17ß-estradiol 1.4 pg/ml, 4.6% and 14%; androstenedione, 0.04 ng/ml, 8.4% and 5.7%; progesterone, 0.02 ng/ ml, 6.0% and 4.7%, respectively. Plasma FSH and LH were measured by RIA according to instructions with kits from the National Hormone and Pituitary Distribution Program. All gonadotropin samples were assayed in duplicate. Sensitivity and inter- and intraassay coefficients of variation were FSH, 200 pg/ml, 2.7% and 6.7%; LH, 86 pg/ml, 5.3% and 2.5%, respectively. The results of all RIAs were calculated by four-parameter logistic analysis using the software AssayZap (BioSoft, Ferguson, MO).

Immunofluorescence and Confocal Microscopy

Ovaries were placed in 4% formalin (2 h) and transferred to 70% EtOH, paraffin embedded, sectioned (4–5 µm), and allowed to air dry for several hours before deparaffinization prior to staining. Slides were blocked with 5% BSA (Gamma Biological, Houston, TX) for 5 min at room temperature. Tissue was incubated with anti-LH receptor (LHr) 200 mg/ml (a gift from J. Wimalansena, University of Tennessee, Knoxville, TN [20]) for 18 h at 4°C. Specificity of the LHr antibody was validated using Western blotting in rat ovarian tissue. The expected band at 93 kDa was observed along with two minor distinct bands of about 60 and 70 kDa, which have been reported to represent proteolytic fragments of the receptor in rat prostate and testis [21]. Sections were incubated with a 1: 75 dilution of biotinylated goat anti-mouse antibody (Vector Lab, Burlingame, CA) in PBS with 1% BSA for 1 h at room temperature, followed by Cy5-conjugated Streptavidin at 15 µg/ml in PBS (Jackson Labs, West Grove, PA) for 1 h at room temperature. To reduce background staining from RNA, slides were incubated with RNase at 3 mg/ml in PBS (Sigma) for 1 h. A polyclonal rabbit antibody was raised against the last 14 amino acids of the mouse HDL receptor (SR-BI) as described [22] and was purified by passing it over a protein A column. Specificity of the antibody was determined by Western blotting, wherein a single band of the apparent molecular mass of 82 kDa was observed. To detect SR-BI, slides were prepared as described above and sections were incubated with rabbit anti-SR-BI at a concentration of 80 µg/ml overnight at 4°C. Sections were incubated with a 1:75 dilution of biotinylated goat anti-rabbit antibody (Vector Lab) in PBS with 1% BSA for 1 h at room temperature and further processed as described above for the LH receptor staining. Genomic DNA was detected in all cells with YoYo-1 at 0.5 µM in PBS (Molecular Probes) incubated for 10 min. Slides were repeatedly washed between incubations with PBS, cover slipped with aqueous mounting medium, and stored in the dark at 4°C until viewed on a Leica confocal microscope (Leica Microsystems AG, Wetzlar, Germany) at 491/509 and 650/670 nm with an Argon/Krypton laser. Images were acquired at 40x magnification. In each experiment, an immunonegative section was performed for each ovary by omitting the primary antibody.

Data Analysis

The effect of VCD treatment on follicle number was determined by Student t-test between two group means for parametric data, and for nonparametric data, by Kruskal-Wallis, each with significance set at P < 0.05. Tissue weights and plasma hormone concentrations were averaged for each treatment and the means (±SEM) in control versus treated animals were analyzed for significant differences by one-way analysis of variance (ANOVA) with significance set at P < 0.05. The differences in steroid concentrations of cell cultures among treatments were analyzed using a one-way repeated measures ANOVA with values averaged for each treatment and the means (±SEM) reported. Post hoc tests (Tukey-Kramer) were used where appropriate. Tests for homogeneity of variance (Bartlett) and normality (Shapiro-Wilk) were routinely performed to ensure that the assumptions of the ANOVA were met. Experiments involving follicle isolations were repeated with separate groups of mice (n = 6/group) for independent replicates. Comparison between treated and control groups used the unpaired Student t-test. The correlation between antral follicle numbers and plasma FSH concentrations was determined by regression analysis with significance set at P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Characterization of the VCD-Treated Ovary

In a detailed dose-response study, VCD treatment (80– 320 mg kg–1 day–1 effective dose) resulted in a significant reduction (P < 0.05) in both primordial and primary follicle pools relative to controls on Day 15 after the initiation of treatment (Fig. 1). Histological evaluation demonstrated that treatment with VCD (160 mg kg–1 day–1, ≤15 days, i.p.) selectively reduced the number of ovarian primordial and primary follicles so that, by Day 37 after the initiation of treatment, none remained (data not shown). At earlier time points (Days 8–15), only these populations were reduced while larger follicles were unaffected (data not shown). By Day 46 (31 days after completion of the 15-day dose schedule), there was also a substantial reduction (P < 0.05) in the number of secondary and antral follicles to 0.7% and 2.6% of the level seen in control animal ovaries, respectively (Fig. 2). A dose of 160 mg/kg was the optimal dose at which primordial follicle loss was accelerated to the greatest extent without affecting larger follicles or other tissues. Therefore, the remainder of the dosing studies utilized VCD at 160 mg kg–1 day–1 for 15 days.



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  		FIG. 1. Follicle depletion in VCD-treated mice. Ovaries were collected from VCD-treated (80–320 mg kg–1 day–1 effective dose) or vehicle control mice on Day 15 following onset of treatment and processed for histological evaluation as described in Materials and Methods. A = 80 mg/ kg, 1 dose/day; B = 80 mg/kg, 2 doses/day; C = 160 mg/kg, 1 dose/day; D = 240 mg/kg, 1 dose/day; E = 80 mg/kg, 3 doses/day; F = 160 mg/ kg, 2 doses/day. Values represent the mean total number of follicles counted in every 20th section of each ovary ± SEM, n = 6. Data were analyzed for significant differences using the Kruskal-Wallis test for nonparametric data (*, different from control, P < 0.05)



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  		FIG. 2. Effect of VCD on various follicle types. Ovaries were collected from VCD-treated (160 mg kg–1 day–1, 15 days) or vehicle control mice on Day 46 following onset of treatment and processed for histological evaluation as described in Materials and Methods. Values are the mean total number of follicles counted in every 20th section of each ovary ± SEM, n = 6. Data were analyzed for significance using the Kruskal-Wallis test for nonparametric data (*, different from control, P < 0.05)

VCD Treatment and Increased Caspase-3 Activity

In a previous study in rats, daily VCD treatment at 80 mg kg–1 day–1 for 10 days caused activation of proapoptotic caspase-3 protease in targeted small preantral follicles during the induction of apoptosis [10]. To confirm that VCD also stimulates this enzyme and apoptosis in small follicles in mice, caspase-3 activity was measured in isolated small preantral follicles on Day 10 following initiation of VCD treatment (160 mg kg–1 day–1). Small preantral follicles were isolated and caspase-3 activity was measured 4 h after the final VCD treatment. Relative to controls, there was an increase in caspase-3 cleavage activity (control, 5.4 ± 0.40 arbitrary units [AU]; VCD-treated, 7.7 ± 0.08 AU; P < 0.05, n = 3).

Onset of Irregular Estrous Cyclicity and Increased FSH

VCD treatment resulted in a gradual disruption of estrous cyclicity. All control animals had regular cycles at all time points. The mean cycle length in control animals was 4.49 ± 0.48 days. VCD-treated animals with cycle lengths that fell outside this range were determined to be irregular. The percentage of VCD-treated mice with irregular cycles was 29% on Day 15, 50% on Day 37, 75% on Day 46, and 100% on Day 58 after the onset of 15 days of treatment. On the last day of VCD treatment (Day 15), there was no increase in plasma FSH levels in VCD-treated animals relative to controls. However, by Day 37 following the onset of treatment, plasma FSH levels were increased (P < 0.05) in VCD-treated animals, relative to controls (VCD, 11.2 ± 5.7 SEM; control, 1.7 ± 0.06 SEM ng/ml; Fig. 3). The increase in FSH in VCD-treated mice continued on Day 46 (14.3 ± 1.3 SEM ng/ml) and Day 58 (20.2 ± 1.2 SEM ng/ ml) to attain a plateau by Days 100 (25.8 ± 0.2 SEM ng/ ml), 120 (25.9 ± 1.1 SEM ng/ml), and 127 (27.5 ± 2.2 SEM ng/ml). There was no change in plasma levels of FSH in the control animals at any time point (Day 37, 1.7 ± 0.4 SEM; Day 46, 2.1 ± 0.2 SEM; Day 58, 1.8 ± 0.04 SEM; Day 100, 5.7 ± 0.9 SEM; Day 120, 4.1 ± 0.06 SEM; Day 127, 4.1 ± 0.07 SEM ng/ml). There was an inverse relationship (P < 0.05) between the number of antral follicles and the increase in circulating levels of FSH on Days 15– 46 (Fig. 3).



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  		FIG. 3. Effect of VCD treatment on circulating levels of FSH and antral follicle loss. A) Plasma was collected from VCD-treated or vehicle control mice on Days 15–127 for determination of FSH content (data analyzed by ANOVA; * P < 0.05 different from control). The Tukey-Kramer post hoc test was used and different letters are different from one another (P < 0.05, n = 6). B) Ovaries were collected from VCD-treated mice on Days 15, 30, 37, and 46 and processed for histological evaluation. Antral follicles were counted in every 20th section and a regression analysis was performed to correlate circulating levels of FSH with antral follicle number (r2 = 0.87, P < 0.05, n = 6 per group at each time point)

Uterine and Ovarian Weights in VCD-Treated Mice

On Day 120 after the onset of VCD treatment, ovarian atrophy could be observed in VCD-treated mice compared with controls. Furthermore, there was a marked absence of ovarian follicles and corpora lutea that were seen in control ovaries (Fig. 4). There was a reduction (P < 0.05) in ovarian (20% of control) and uterine (50% of control) weights in VCD-treated mice relative to controls (ovaries, control 0.053, VCD 0.011 mg/g body weight; uteri, control 0.40, VCD 0.23 mg/g body weight).



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  		FIG. 4. Effect of VCD treatment on ovarian morphology. Micrograph of sections of representative ovaries stained with hematoxylin and eosin collected from vehicle control (A) and VCD-treated (B) animals on Day 120. Bar in (A) = 500 µm

Effect of VCD Treatment on Tissue Weights of Nonreproductive Organs

There was no effect of VCD treatment on body weight or adrenal, kidney, or spleen relative to body weight at any time point (data not shown). There was a modest increase (P < 0.05) in liver weight on Day 15 (10% above control) and Day 37 (15% above control), which had returned to control levels by Day 46. As a measure of liver function in VCD-treated animals on Days 10, 15, and 46, hepatic enzymes AST and ALT were within the normal range and there was no difference in histology between livers from treated and control animals (data not shown).

Plasma Levels of LH, FSH, and Steroids in VCD-Treated Mice

On Day 127, plasma LH and FSH levels were greater (P < 0.05) in VCD-treated mice relative to cyclic controls (Table 1). On Day 127, plasma levels of progesterone and androstenedione were reduced (P < 0.05) in VCD-treated mice relative to controls and 17ß-estradiol levels were undetectable (Table 1). With the drop in 17ß-estradiol to nondetectable levels and maintenance of circulating androstenedione, the relative ratio of androgen to estrogen in plasma of VCD-treated mice was increased compared with control.


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  		TABLE 1. Effect of VCD treatment on plasma gonadotropins and steroids.*

Steroidogenic Capacity of VCD-Treated Ovarian Tissue

Immunohistochemistry was used to examine control and VCD-treated ovaries for protein distribution of LH receptor (LHr) and the HDL receptor, SR-BI. The location of LHr expression by immunofluorescence (red staining) was observed in theca interna and interstitial cells in control ovaries (Fig. 5A). In ovaries from VCD-treated animals, LHr staining was more diffuse throughout the residual ovarian tissue than seen in controls (Fig. 5B). There was no immunofluorescence when the primary antibody was omitted (Fig. 5C).



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  		FIG. 5. Effect of VCD treatment on tissue distribution of receptors for LH and SR-BI. Ovaries from VCD-treated and vehicle control mice were collected on Day 127 and processed for immunostaining and confocal microscopy as described in Materials and Methods. Sections were immunostained (red) for LHr (A, B) or SR-BI (D, E) in controls (A, D) and VCD-treated (B, E) ovaries. Green staining shows visualization of genomic DNA in all cells to assist in identification of ovarian structures. Immunonegative sections (C, F) show no staining when primary antibody is omitted. Magnification x40. Bar in (A) = 50 µm

The strongest staining for SR-BI protein (the HDL receptor), which is indicative of active cellular steroidogenesis, was observed in theca interna and interstitial cells in the control ovarian sections (Fig. 5D). In contrast, in the ovarian section of a VCD-treated animal, SR-BI was uniformly dispersed and stained at a lower intensity (Fig. 5E). SR-BI immunostaining was also observed in the oocyte. No immunofluorescence was seen in sections stained without primary antibody (Fig. 5F).

Cell Culture Steroidogenic Production

To confirm the production of 17ß-estradiol in ovarian cultures of VCD-treated animals, ovarian cells on Day 37 were incubated with cholesterol substrate (HDL) and/or a combination of LH (10 ng/ml) and FSH (10 ng/ml). The 17ß-estradiol content of the media was significantly less in VCD-treated ovarian cells when compared with controls (31.28 ± 2.31 SEM pg/ml-µg DNA, VCD versus 72.27 ± 2.35 SEM pg/ml-µg DNA, control).

Cyclicity in control animals in the Day 120 cell culture experiment was determined by vaginal cytology (Days 100–120). At the time of tissue collection, six animals were in estrus, five in diestrus-1, nine in diestrus-2, and three in proestrus. To maintain consistency, cells from animals in different stages of the cycle were pooled for the control cultures as described in Materials and Methods.

Ovarian luteal cells make progesterone. Furthermore, in the rodent, progesterone is the substrate used by theca and interstitial cells to produce androstenedione via the {Delta}4 pathway [1]. To determine if there was substrate for androstenedione synthesis, progesterone content of the media from cultured cells from control and VCD-treated ovaries was measured. Ovarian cells from Day 120 control and VCD-treated animals produced progesterone in response to LH stimulation (Fig. 6). In addition to interstitial tissue, control ovaries contained theca, granulosa, and luteal cells as observed in histological sections. Cells from these ovaries produced progesterone and were stimulated (P < 0.05) by LH (≥1 ng/ml). Cells from VCD-treated ovaries produced 20%–40% less (P < 0.05) progesterone than cells from control ovaries.



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  		FIG. 6. LH-stimulated production of progesterone in vitro. Whole ovarian dispersates were obtained from VCD-treated and vehicle control mice. On Day 120 following onset of treatment, cells were prepared and cultured with 0, 1, 3, 10, or 30 ng/ml LH, and human HDL (100 µg/ml), ± insulin (1 µg/ml). Progesterone production was determined after 48 h of culture as described in Materials and Methods. In each well, progesterone content of medium was normalized to cellular DNA content. Data were analyzed by repeated-measures ANOVA and are represented as group mean values ± SEM (n = 3 separate experiments; * P < 0.05; Tukey-Kramer post hoc tests were performed and letters are different, P < 0.05, within groups)

In control cell cultures, androstenedione production was below the limit of detection. Therefore, only results are shown for cells from ovaries of VCD-treated animals (Fig. 7). Cells from these animals produced androstenedione, ranging from 10–13 ng/ml-µg DNA per culture well, with no apparent sensitivity to LH. Interstitial cell androgen production is responsive to insulin [23]. Therefore, insulin was added to culture media to test its impact on progesterone and androstenedione production (Figs. 6 and 7). The amount of progesterone produced by control cells and cells from VCD-treated animals was unaffected by the presence of insulin (Fig. 6). However, insulin stimulated (P < 0.05) androstenedione production in cells from VCD-treated animals at LH concentrations of 1 and 3 ng/ml (Fig. 7).



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  		FIG. 7. LH-stimulated production of androstenedione in vitro by ovarian cell cultures from VCD-treated animals. Whole ovarian dispersates were obtained from VCD-treated animals. On Day 120 following onset of treatment, cells were prepared and cultured with 0, 1, 3, 10, or 30 ng/ml LH and human HDL (100 µg/ml) ± insulin (1 µg/ml). Androstenedione production was determined after 48 h of culture as described in Materials and Methods. In each well, androstenedione content of medium was normalized to cellular DNA content. Data were analyzed by repeated-measures ANOVA and are represented as group mean values ± SEM (n = 3 separate experiments; * P < 0.05)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
4-Vinylcyclohexene is an industrial chemical used in the manufacture of rubber tires, polyesters, coatings, and plastics [24]. Its diepoxide metabolite, 4-vinylcyclohexene diepoxide (VCD), has been extensively characterized for its ability to cause selective loss of small preantral (primordial and primary) follicles in ovaries of rats and mice after repeated dosing [5–9, 16, 25]. Follicle loss appears to involve apoptosis, as VCD treatment in rats at 80 mg kg–1 day–1 has been shown to activate several apoptotic intracellular signaling pathways in isolated, small preantral follicles. Specifically, VCD increased the expression of proapoptotic Bad, the Bax/BclxL ratio on the mitochondrial membrane, leakage of mitochondrial cytochrome c into the cytosol, caspase-3 cleavage activity, and activation of the JNK and p38 branch of the MAPK signaling pathway [10–12]. Collectively, these past studies have led to the conclusion that VCD treatment causes follicle loss via acceleration of the natural process of atresia [6, 8, 10–12].

In the present study, repeated treatment of mice with VCD at a higher concentration (160 mg kg–1 day–1) caused an accelerated selective depletion of ovarian primordial and primary follicles. Without the precursor follicle populations for recruitment, eventually all growing follicles (preantral and antral), also became depleted. As antral follicles became reduced, there was an associated increase in circulating FSH levels. This was likely due to a concomitant reduction in ovarian 17ß-estradiol and inhibin, which provided a negative feedback effect on FSH release [26]. The result is a follicle-deplete, ovary-intact animal. As a result of follicle depletion, relative to age-matched cyclic controls, there was an increase in FSH and LH, a decrease in progesterone and androstenedione, and 17ß-estradiol fell to nondetectable levels. Thus, because 17ß-estradiol dropped below the limits of detection and androstenedione was only reduced by one third, the endocrine milieu became relatively androgen enriched in the VCD-treated animals. There is a greater androgen to estrogen ratio in postmenopausal women [2–4, 27–31], although there has been some debate as to the contribution of the postmenopausal ovary to androgen production [32]. Whereas follicle loss has not been spontaneous in VCD-treated animals, the endocrine profile is similar to that in postmenopausal women (low estrogens, high gonadotropins). Thus, a mouse that retains residual ovarian tissue may be useful for the assessment of androgen contributions to events associated with postmenopausal pathologies such as cardiovascular disease, diabetes, osteoporosis, and Alzheimer disease [4, 33–35].

No evidence of generalized VCD toxicity in tissues other than the ovary was seen. The modest initial increase in liver weight returned to levels similar to that of the untreated controls by Day 46. This increased liver weight was likely due to an increased expression of the enzymes associated with xenobiotic metabolism that are commonly induced by exposure to foreign chemicals, such as VCD. Notably, the liver enzymes AST and ALT were not increased in plasma, and no hepatocellular lesions developed. Therefore, the reduction in uterine and ovarian weights was the only irreversible tissue effect observed. Whereas it cannot be conclusively ruled out that VCD caused a direct effect on the ovaries, the evidence in other tissues presented here and results from previous studies would support that this is not the case [6, 9–12, 14, 15]. The decrease in ovarian weight was most likely the direct result of VCD-induced follicle depletion and premature ovarian failure, whereas the decrease in uterine weight resulted from the withdrawal of ovarian 17ß-estradiol and its well-known tropic effects on this target tissue [36].

Ovarian interstitial cells are large and polyhedral in shape and contain extensive lipid stores, LH receptor (LHr), SR-BI, and steroidogenic enzymes to facilitate androgen production [1, 37]. Interstitial cells are derived from theca cells in atretic follicles [1]. Staining for LH receptor and SR-BI was concentrated in theca interna as well as in interstitial cells in the control ovaries. This distribution of those markers for androgenic cells demonstrates that these phenotypic characteristics are retained as theca interna transition into interstitial cells. Although VCD selectively targeted primordial and primary follicles, ultimately follicles of all sizes were lost. Residual ovarian tissue in VCD-treated ovaries contained LHr and SR-BI protein and displayed the ability to synthesize progesterone and androstenedione. Therefore, this tissue demonstrated phenotypic properties suggestive of interstitial/stromal cells [1, 38].

Circulating levels of progesterone were higher in cyclic controls as compared with VCD-treated animals on Day 127. Likewise, in vitro progesterone production was greater in cultures from control as compared with VCD-treated animals on Day 120. Whereas the ovaries of control animals were enriched in corpora lutea, ovaries from VCD-treated mice were devoid of them. The greater progesterone production in vivo and in vitro reflects this circumstance.

The rodent adrenal does not produce androgen [39]. Therefore, circulating androstenedione levels that persisted after VCD-induced ovarian failure are derived from the ovary. In support of this conclusion, cells from VCD follicle-deplete ovaries produced androstenedione in culture. On Day 127, circulating androstenedione levels were higher in cyclic controls as compared with VCD-treated animals, yet in vitro, androstenedione production of dispersed cells was detectable in cultures from VCD-treated but not control animals. Circulating androstenedione levels can be provided by ovarian theca interna as well as interstitial cells. Because VCD-treated animals were follicle deplete, circulating androstenedione would have been solely from interstitial cells, thus, lower than cyclic controls. In contrast, in cultured ovarian cells, unlike VCD-treated ovaries, androstenedione was undetectable in cyclic controls. Control animal cell dispersates contain a heterogeneous mixture of interstitial, theca interna, granulosa, and luteal cells. Because cultures were established on the basis of total cell number, cultures from VCD-treated animals contained a much greater percentage of androgen-producing (theca/interstitial) cells than those from controls. Therefore, relative androgen production in this homogeneous mixture of cells was detectable.

Androgen output of ovarian cells isolated from VCD-treated animals was increased in the presence of insulin and LH (1 and 3 ng/ml). This is in agreement with an earlier report from isolated theca/interstitial cells cultured from immature rats [23]. The combined in vivo and in vitro results reported here support the conclusion that VCD-induced follicle depletion results in an interstitial cell-enriched ovary that produces androgen in response to LH when insulin is also present. Insulin resistance is often observed in hyperandrogenic postmenopausal women that have increased plasma levels of insulin [40–42]. Although insulin-resistant tissues such as muscle are insensitive to insulin, ovarian production of androgen in these women is stimulated by insulin [33, 42]. These findings are similar to the scenario in postmenopausal hyperandrogenic women in which circulating LH levels are increased and residual ovarian tissue is insulin sensitive [33, 41, 42]. The physiological impact of elevated androgens that are unopposed by estrogens in postmenopausal women has not been widely studied [4, 36, 43–45]. Thus, the follicle-deplete, ovary-intact mouse model developed here will be ideal to investigate this circumstance.

Rodent ovaries do not become senescent until very late in life. Clearly, a rodent model developed in a younger mouse would have great potential for studies related to ovarian senescence in humans. Other animal models have been used for studies related to ovarian senescence in women. However, there are drawbacks in each case. The most commonly used is the ovariectomized animal [46]. In that case, ovarian senescence is abrupt and no residual ovarian tissue remains. There are several transgenic mouse animal models, such as mice with mutations in the FSH receptor and growth differentiation factor 9 and other transforming growth factor beta super family ligands [26, 46]. However, in such cases, fetal development of the reproductive system has been affected from the beginning and normal reproductive function has never been achieved. Finally, nonhuman primates provide the most suitable model for humans [46]. Yet these animals are not economically feasible for large-scale experimentation and they reach ovarian senescence well along in their life expectancy. Thus, the VCD-treated, follicle-deplete mouse may prove to be a useful alternative to these other models.

In summary, VCD-induced premature ovarian failure in mice leads to the development of a follicle-deplete, interstitial or stromal cell-rich ovary that resembles the endocrine status and function of the postmenopausal ovary. Because follicle depletion occurs over time, this may serve as an approach for studying the dynamics of follicle loss in women as ovarian senescence ensues (perimenopause). Recent clinical findings have demonstrated a need to find better ways of evaluating the risks versus benefits of long-term steroid hormone replacement [47–49]. Such an approach may be worthwhile for studies designed as a comparison for women who have undergone menopause by the natural process.


   ACKNOWLEDGMENTS
 
The authors wish to thank Dr. Carole L. Banka, the La Jolla Institute for Molecular Medicine, for her comments and kind gift of human HDL. At the University of Arizona, Dr. Xaioming Hu and Dr. Dun Lu, for their assistance with the caspase-3 activity assay; Patty Christian, for the confocal image analysis; Sam Marion, for hormone RIAs; and Kathila Rajapaksa, for assistance with animal handling, are appreciated. At Northern Arizona University, Matt Freyer, Rachel Allred, Gina Buss, Tracy Layton, Zach Robinson, Rachel Steagall, and Stefanie Raymond-Whish are thanked for the culture steroid assays and animal handling, and Dawn Nice and Angie Trujillo are acknowledged for production of the SR-BI antibody.


   FOOTNOTES
 
1 Supported by NIH grants RO1-ES08979, RO1-ES09246, RO1-AG21948, and Center Grant ES-06694 to P.B.H. and American Heart Association Fellowship to L.P.M. Back

2 Correspondence: Patricia B. Hoyer, Department of Physiology, The University of Arizona, 1501 North Campbell Ave., Tucson, AZ 85724-5051. FAX: 520 626 2382; hoyer{at}u.arizona.edu Back

Received: 13 February 2003.

First decision: 3 March 2003.

Accepted: 24 February 2004.


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