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Regular Article |
a Laboratory of Reproduction and Lactation, CONICET, 5500 Mendoza, Argentina
b Department of Physiology and Biophysics, University of Illinois at Chicago, Chicago, Illinois 60612
c Department of Morphophysiology, Faculty of Medicine, University of Cuyo, 5500 Mendoza, Argentina
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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androgen receptor, apoptosis, corpus luteum, corpus luteum function, progesterone
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In contrast to what has been demonstrated in ovarian follicles, little information is available concerning the effect of androgens on corpus luteum (CL) function and on the expression of AR in this other ovarian tissue. Rat placenta and ovaries produce androstenedione and testosterone in pregnancy [18]. These aromatizable androgens are extensively involved in supporting luteal progesterone production following their conversion into estrogen [19]. An explanation for this is provided by the presence of estrogen receptors in the rat CL [20]. Androgen-derived estrogen increases the supply of cholesterol substrate needed for steroidogenesis and mediates cell hypertrophy, contributing to the increase in size of the CL at midpregnancy [19]. However, conversion to estrogen is not always required for the action of androgens in the CL. 5
-Dihydrotestosterone, a nonaromatizable androgen, was approximately equipotent to androstenedione and testosterone in stimulating progesterone release from rat and mouse luteal cells in culture [21, 22], and this effect was not reproduced by the addition of estradiol to the culture media [21]. In rats undergoing luteal regression after treatment with the antigestagen RU486, androstenedione was capable of preventing the decrease in serum progesterone concentration and the loss in luteal weight induced by RU486, even in the presence of an aromatase inhibitor or a specific estrogen-receptor blocker [23]. These results suggest that the action of androstenedione on luteal progesterone production and secretion at the time of luteolysis occurs through an androgenic mechanism.
In the present investigation, we examined whether the rat CL expresses the AR gene and whether this expression is developmentally regulated in pregnancy. In addition, because androstenedione is the main androgen produced in pregnant rats [19], and because this androgen strongly stimulates the production of progesterone [21, 23], which is a known survival factor for the CL [24–28], we investigated whether androstenedione is capable of preventing the apoptotic changes associated with luteal regression. We tested this hypothesis in CL obtained from rats after parturition, when structural regression is very advanced [28, 29]. We took advantage of the postpartum ovary being composed of two generations of CL that can be simultaneously studied: the CL of pregnancy, and the newly formed CL derived from the ovulation postpartum [30]. To evaluate further the effect of androstenedione as a survival factor in the rat CL, we also used a previously established in vitro approach in which CL incubated in serum-free conditions accumulate large numbers of cells in different stages of apoptosis and undergo extensive DNA fragmentation [31]. Part of this work was presented at the 2001 Annual Meeting of the Society for the Study of Reproduction [32].
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Adult female rats bred in our laboratory (originally Wistar strain) and weighing 180–220 g were used. The rats were housed under controlled light (lights-on from 0600 to 2000 h) and temperature (22–24°C) conditions and had free access to standard rat chow (Cargill, Mendoza, Argentina) and tap water. To induce pregnancy, female rats were caged individually with fertile males beginning on the afternoon of proestrus. Positive mating was verified on the following morning by identifying sperm or copulation plugs in the vagina. This day was designated as Day 0 of pregnancy. In our laboratory, rats usually give birth on Day 22. Adult, 3-mo-old male rats bred in our laboratory were also used. Animals were handled according to the procedures approved in the Guide for the Care and Use of Laboratory Animals [33].
Experimental Procedure
For the developmental studies, three rats per group were killed at 1300 h on Days 7, 12, 15, 20, and 21 of pregnancy and on Day 4 postpartum. The CL were dissected from the ovaries under a stereoscopic microscope. Prostate, epididymis, and spleen were obtained from three different male rats. All tissues were frozen in liquid nitrogen and stored at -80°C until processed for RNA or protein isolation. For the immunohistochemical studies, whole ovaries from pregnant (Day 15) animals and prostate and epididymis from adult males were obtained and fixed for 1 h at room temperature in a solution of 10% (v/v) phosphate-buffered neutral formalin, dehydrated in ethanol series, cleared in xylene, and embedded in paraffin.
To determine the effect of androstenedione on luteal apoptosis, two groups of six to eight animals were studied after parturition. The pups were removed immediately after parturition, and the rats were injected daily with androstenedione (8 mg/rat s.c.) or vehicle (sunflower seed oil) at 1000 h. Animals were killed by decapitation at 1300 h on Day 4 postpartum. Trunk blood was obtained to determine hormone concentrations. The ovaries were removed and fixed for 1 h at room temperature in a solution of 10% phosphate-buffered neutral formalin, dehydrated in ethanol series, cleared in xylene, and embedded in paraffin for routine hematoxylin-eosin (H&E) staining. Luteal regression was evaluated separately in the two generations of CL by studying the number of nuclei undergoing apoptosis. In two other groups of animals similarly treated, the CL were isolated under a stereoscopic microscope, classified as old (i.e., CL of the previous pregnancy) or new (i.e., newly formed CL after postpartum ovulation), weighed, and used for in vitro incubation.
RNA Isolation and Reverse Transcription-Polymerase Chain Reaction
Total RNA from CL, prostate, and epididymis was purified using Trizol (Life Technologies, Inc., Rockville, MD) according to the manufacturer's instructions. One microgram of total RNA was reverse transcribed using the Advantage RT for PCR kit (Clontech Laboratories, Inc., Palo Alto, CA). For amplification of the reverse transcription (RT) products, the reaction mixture consisted of 1x PCR buffer (ExTaq buffer; Panvera, Madison, WI), 150 µM deoxynucleoside triphosphates, 50 pmol of specific oligonucleotide primers, and 0.8 U of ExTaq (Panvera) in a final volume of 40 µl. The samples were overlaid with light mineral oil, and polymerase chain reaction (PCR) was carried out in a PTC-1000 thermal cycler (MJ Research, Whalthan, MA). Two sets of amplification cycles were separately used for detection of AR mRNA and the mRNA for the ribosomal protein L19, which was used as an internal control. For AR, PCR was carried out for 25 cycles using 93°C for denaturing (45 sec), 69°C for annealing (45 sec), and 72°C for extension (60 sec). For L19, 30 PCR cycles were performed using 93°C for denaturing (45 sec), 65°C for annealing (45 sec), and 72°C for extension (60 sec). Specific primers for AR were based on the sequence of the rat AR gene (5'-CAGCCCCAGCCCAGCGACAGC-3' and 5'-CAGGGTGAGGGGCGGGCAGTAGGA-3') [34], and for L19, the specific primers were 5'-CTGAAGGTCAAAGGGAATGTG-3' and 5'-GGACAGAGTCTTGATATCTC-3' [35]. The predicted sizes of the PCR-amplified products were 423 and 120 base pairs (bp) for AR and L19, respectively. The PCR products were resolved on 1.8% agarose gel (FMC Corp., BioProducts, Rockland, ME), visualized with ethidium bromide using an ultraviolet (UV) transilluminator (UVP, Upland, CA), and photographed using a photodocumentation camera (Fisher Scientific, Pittsburgh, PA).
Immunoblot Analysis
Isolated CL, epididymis, or prostate were homogenized in 2 ml of ice-cold homogenization buffer containing 25 mM Tris-HCl (pH 7.4), 2 mM MgCl2, 1 mM EDTA, 1 mM PMSF, 1 mM dithiothreitol, 1 µM leupeptin, 1 µM pepstatin-A, and 1 µg/ml of aprotinin. Homogenates were assayed for protein content [36], appropriately diluted in 6x concentrated electrophoresis sample buffer, boiled for 10 min, and stored at -80°C until electrophoresed. Equivalent amounts of protein (50 µg) were separated through 8% SDS-PAGE gels and electrotransferred to nitrocellulose membranes (Schleicher & Schuell, Inc., Keene, NH). Immunoblotting was performed by blocking nonspecific binding with 5% (w/v) nonfat milk in Tris-buffered saline (TBS) containing 0.1% (v/v) Tween 20. Blots were then incubated overnight with 1 µg/ml of the polyclonal antibody raised against a peptide mapping at the amino terminus of the AR of human origin (N-20; Santa Cruz Biotechnology, Santa Cruz, CA). The membranes were washed and incubated with a secondary antibody (goat anti-rabbit, 1:1000 dilution) linked to horse radish peroxidase (HRP; Dako Corp., Carpinteria, CA) for 90 min. After extensive washing, blots were developed by chemiluminescence and exposed for 15–120 sec to x-ray films. Data were analyzed using the NIH Image software (National Institutes of Health, Bethesda, MD). The molecular size of immunoreactive bands was estimated by the comigration of a prestained SDS-PAGE molecular mass standard (Benchmark; Gibco/BRL, Gaithersburg, MD).
Immunohistochemistry
Serial paraffin sections (thickness, 5 µm) were mounted on 3-aminopropyltriethoxy-silane-coated slides. The immunohistochemical procedure was performed using a highly sensitive strept ABComplex/HRP detection system (Dako). The sections were deparaffinized with xylene and rehydrated in alcohol solutions of increasing water content, then washed in TBS (0.15 M NaCl and 0.05 M Tris [pH 7.6]). Endogenous peroxidases were blocked in 3% H2O2 for 20 min at room temperature. To reduce nonspecific background staining, the sections were blocked with 3% (v/v) normal goat serum for 20 min. The sections were then incubated with the antiandrogen receptor antibody (N-20; 2 µg IgG/ml) overnight at 4°C in a moist incubation chamber. The sections were washed in TBS and incubated successively with biotinylated goat anti-rabbit IgG (2.5 mg/ml; Dako) for 30 min at room temperature, and strept ABComplex/HRP (1:50 dilution) for 30 min at room temperature. Each incubation was followed by two 10-min washes in TBS. After the final wash, sections were incubated in the dark for 10 min with the diaminobenzidine (DAB) solution (0.05 M Tris buffer [pH 7.6] containing 0.6 mg/ml of DAB and 0.03% [v/v] H2O2) for developing peroxidase activity. After washing with tap water, the sections were dehydrated in serial alcohols to xylene and mounted without counterstaining. In the negative control slides, the primary antibody was replaced with TBS. The tissue samples were observed and photographed with a Nikon Eclipse E 400 microscope (Fryer Company Inc., Huntley, IL).
Counting of Apoptotic Cells
Apoptotic cells were recognized in H&E-stained tissue sections by morphological criteria following the procedure described by Van der Schepop et al. [37] with slight modifications as previously described [27]. Only cells with advanced signs of apoptosis (i.e., containing multiple nuclear fragments) were counted. A microscope with a 100x objective was used, and as many fields as possible were analyzed in each CL for the presence of fragmented nuclei. All the CL in each section were studied, and an average number of apoptotic nuclei per CL was obtained. Results are expressed relative to the control group, in which the number of apoptotic cells was considered to be 100%. This morphometric method for the identification of apoptotic cells was previously validated in the CL by in situ 3' end-labeling [27].
Incubation of CL
The CL (4–6 CL/ml/well in a 24-well tissue-culture plate) were incubated in serum-free medium (McCoy 5A:Ham F12, 1:1; Sigma Chemical Co., St. Louis, MO) containing 25 mM Hepes, 200 IU/ml of penicillin G, 200 µg/ml of streptomycin, and 0.5 µg/ml of amphotericin B at 37°C for 1, 2, 4, 6, or 17 h in an atmosphere of 95% air/5% CO2. After incubation, the CL were immediately frozen in liquid nitrogen and stored at -80°C until genomic DNA isolation. The incubation media was kept at -20°C until the measurement of progesterone concentration.
DNA Fragmentation
The internucleosomal cleavage of DNA was analyzed as follows: At the end of the experiment, CL were immediately removed from the culture media; homogenized in digestion buffer composed of 100 mM NaCl, 10 mM Tris-Cl (pH 8.0), 25 mM EDTA (pH 8.0), 0.5% SDS, and 0.1 mg/ml of proteinase K (Life Technologies); and incubated overnight at 50°C. The DNA was extracted from the digested tissues with phenol/chloroform/isoamyl alcohol (25:24:1, v/v/v). The DNA was precipitated and digested for 1 h at 37°C in 1 µg/ml of ribonuclease solution (deoxyribonuclease-free; Roche, Indianapolis, IN). After extraction and precipitation, an equal amount of DNA (5 µg) was separated by electrophoresis on a 1.8% agarose gel (FMC Corp., BioProducts), impregnated with ethidium bromide, and examined by UV transillumination.
Hormone Assays
Serum prolactin was determined by double-antibody RIA with reagents provided by the National Institute of Diabetes, Digestive, and Kidney Diseases (Bethesda, MD). Results are expressed in terms of the rat prolactin RP-3 standard. The sensitivity of the assay was 0.5 ng/tube. Inter- and intraassay coefficients of variation were less than 10%.
Progesterone concentrations were measured using a commercially obtained kit (Diagnostic Products Corporation, Los Angeles, CA). The sensitivity of the assay was 0.02 ng/ml, and the inter- and intraassay coefficients of variation were 5% and 6%, respectively.
Androstenedione was assayed using an RIA developed in our laboratory with an antiserum raised against 4-androsten-3,17-dione 3-CMO:BSA (Steraloids, Wilton, NH). The antiserum cross-reacted 100% with androstenedione, 8.32% with dehydroisoandrosterone, 5.71% with dehydroepiandrosterone, 0.5% with testosterone, 0.45% with dihydrotestosterone, and less than 0.1% with 5-androstane-3ß,17ß-diol, 5-androstane-3,17ß-diol, corticosterone, cortisol, pregnenolone, 17-hydroxyprogesterone, 20-dihydroprogesterone, progesterone, and 17ß-estradiol. The sensitivity of the assay was 6 pg/tube. The intra- and interassay coefficients of variation were 4.41% and 16.66% respectively.
Statistics
Comparisons between the means of two groups were carried out using the Student t-test. For multiple comparisons, one-way ANOVA followed by the Tukey multiple-range test was used. A level of P < 0.05 was accepted as being statistically significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To determine whether AR mRNA is expressed in the rat CL, we performed RT-PCR to specifically detect the AR mRNA transcript. Shown in Figure 1A is the RT-PCR amplification curve of AR mRNA achieved using 1 µg of total luteal RNA obtained from rats on Day 15 of pregnancy. A linear amplification of AR mRNA was obtained using up to 25 cycles of PCR. Fixing the number of PCR cycles to 25, we next determined that the assay was linear with respect to the amount of input RNA, as shown in Figure 1B. Therefore, subsequent experiments were performed with 25 cycles of amplification of PCR and 1 µg of total RNA.
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To investigate the developmental changes in AR mRNA, CL were isolated on different days of pregnancy. Total RNA was obtained and subjected to RT-PCR with L19 as an internal control. The results shown in Figure 2 indicate that AR mRNA in the CL remains highly expressed throughout gestation but declines significantly before parturition on Day 21.
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To determine the tissue specificity of the transcripts detected in the CL, we also studied AR mRNA expression in two well-known AR-expressing tissues: prostate, and epididymis. The PCR products generated from amplification of cDNA obtained from CL on Day 7 or 21 of pregnancy were of similar size (423 bp) to those generated from prostate and epididymis (Fig. 3A). These results indicate that the same AR transcript is probably expressed in the CL as in the positive control tissues. To study whether the AR mRNA expressed by the rat CL is translated into AR protein, Western blot analysis was performed. As shown in Figure 3B, an AR immunoreactive protein of 120 kDa was identified in CL as well as in prostate and epididymis. However, in contrast to the drop in AR mRNA levels (see Fig. 2), no decline in AR protein was seen on Day 21.
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The distribution of the AR protein within the pregnant rat ovary was assessed by immunohistochemistry. Figure 4A displays positive AR-stained nuclei within a CL obtained from a pregnant (Day 15) rat. Diffuse staining was also observed within the cytoplasm of the luteal cells. We compared the immunostaining found within the CL with that observed in the ovarian follicles and prostate. Early antral follicles displayed positive staining within the granulosa cell layer, whereas no staining was observed in the follicular theca (Fig. 4B). As expected, positive AR immunoreactivity was detected within the nuclei of the prostate epithelial compartment (Fig. 4C). No staining whatsoever was observed in the same tissues when the primary antibody was omitted (Fig. 4, D–F).
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Interference of Luteal Regression by Androstenedione
To study the effect of androstenedione on luteal regression, we used rats whose pups were removed immediately after delivery to prevent establishment of lactation. Under these conditions, the animals undergo extensive luteal regression in both generations of CL by Day 3–4 postpartum [28, 38]. Figure 5 shows that both types of CL express AR mRNA and, thus, are potential target tissues for androgens. When nonlactating rats were treated with androstenedione for 4 days after parturition, the number of apoptotic nuclei within both generations of CL was significantly reduced (Fig. 6, A and B). In addition, the decrease in average weight of the CL seen in control animals by Day 4 postpartum was significantly prevented by treatment with androstenedione (Fig. 6, C and D). The new CL formed after postpartum ovulation were smaller than the CL of previous pregnancy. Despite the tendency for the newly formed CL to be more susceptible to androstenedione than the CL of previous pregnancy, such a difference was not statistically significant. As expected, animals that were treated with androgen displayed very high circulating levels of androstenedione (Fig. 7A). Prolactin, one of the main hormones involved in CL function in rats, did not change with treatment, displaying low circulating levels in both vehicle- and androstenedione-treated animals (Fig. 7B). In contrast, serum progesterone concentration was greatly increased by androstenedione treatment (Fig. 7C).
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We have previously demonstrated that the in vitro induction of luteal cell death under serum-free conditions occurs with similar timing and to a similar extent in both generations of CL observed after parturition in rats [39]. Therefore, for the in vitro study, only one generation of CL was used. As shown in Figure 8A, old CL obtained from nonlactating rats on Day 4 postpartum and incubated in serum-free conditions displayed extensive DNA fragmentation in a time-dependent manner. However, in nonlactating rats that had received a daily dose of androstenedione from Days 1 to 4 postpartum, the in vitro-induced luteal DNA fragmentation was markedly delayed (Fig. 8B). The incubation media was tested for progesterone accumulation (Fig. 8C). The CL from nonlactating rats accumulated little progesterone in the media over time. In contrast, CL obtained from nonlactating rats pretreated with androstenedione accumulated increasing amounts of progesterone over the period of incubation, and levels of progesterone were greater than those in controls at each time point.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Luteal AR Expression
The AR has been located in the CL of several species of mammals, suggesting a role for androgens in modulating luteal function via a classical receptor-mediated pathway. In cycling pigs, luteal cells from the early and midluteal phases stained positive for the AR [40]. Immunohistochemical detection of AR was also shown in CL of rhesus monkeys at different stages of the luteal phase in the menstrual cycle [41]. Whereas regressing CL of the late luteal phase in monkeys stained for AR, fully regressed CL in the early follicular phase of the next cycle did not exhibit receptor staining [42], suggesting a developmental regulation of AR gene expression. In rats, the presence of AR has been demonstrated in ovarian follicles and CL of the estrus cycle [10, 43], yet to our knowledge, no information exists regarding the ovarian expression of AR throughout pregnancy in this species. We have shown, in pregnant rat ovaries, positive AR immunostaining in the granulosa cell layer, but not in the theca cells, of small antral follicles. Our results agree with those of a previous report that documented AR localized predominantly in the granulosa cell nuclei, with the most intense staining in preantral and early antral follicles of cycling rat ovaries [10]. We also established that AR message and protein are expressed throughout gestation in the rat CL. Although luteal AR mRNA declined at the end of pregnancy (see Fig. 2), it was not followed by a concomitant decrease in the expression of AR protein (see Fig. 3B). These results suggest that the decrease in steady-state levels of AR mRNA before parturition is not crucial for normal progression of the luteolytic process in this species.
The AR protein localizes to the nucleus in target cells upon ligand binding [44], but it remains largely in the cytoplasm in the absence of hormone [45]. Some evidence suggests that AR, in addition to its known transcriptional activity, may exhibit some cytoplasmic effect by mimicking the action of growth factors modulating several cellular signaling pathways [46]. For example, treatment of androgen-responsive prostate cancer cells with dihydrotestosterone leads to a rapid and reversible activation of mitogen-activated protein through a mechanism insensitive to AR antagonists [47]. In our study, the AR localized mainly to the nuclei of steroidogenic luteal cells, but diffuse staining was also observed within the cytoplasm, suggesting that AR in the CL may regulate genomic as well as nongenomic processes. Using total extract, a 120-kDa protein was demonstrated in both the CL and the male tissues used as positive controls, prostate and epididymis. As expected, we did not find any immunoreactive protein band in spleen (result not shown), which has been reported to be negative for AR expression [43].
Of particular interest was that AR expression was observed not only in the newly formed CL after postpartum ovulation but also in the old CL of pregnancy in animals killed 4 days after parturition. This indicates that both generations of CL found in postpartum ovaries are potential targets to androgens.
Interference of Luteal Regression by Androstenedione
Luteal regression in the rat is a complex process that involves two stages. The first, named functional regression, is characterized by a rapid decrease in the capacity of the gland to produce and to secrete progesterone. The second, called structural regression, is grossly characterized by a decrease in the weight and size of the CL. This last process is executed long after the initial decline in progesterone secretion and is carried out over several days after parturition [29, 38]. The main mechanism involved in the reduction in size and weight of the CL is the removal of luteal cells by apoptosis and subsequent phagocytosis [27, 48]. In the present study, we have shown that administration of androstenedione to rats after parturition significantly interfered with the process of luteal cell removal. This was demonstrated by 1) the finding that androstenedione prevented the luteal weight loss and the increase in number of apoptotic cells in regressing CL after parturition and 2) the delayed capacity to undergo apoptosis in vitro displayed by the CL obtained from androstenedione-treated animals.
Androgens are involved in both cell death and survival in various tissues. In the prostate, androgens stimulated growth and differentiation and prevented apoptosis [49]. In primary cultures of human neurons, testosterone, acting through the AR, prevented cell death induced by serum deprivation [50]. Conversely, androgens can also promote apoptosis, as demonstrated in breast cancer cells [51] and ovarian granulosa cells [5]. The enhancement of ovarian granulosa cell apoptosis by testosterone strongly contributes to follicular atresia [5]. In sharp contrast to what was demonstrated in ovarian follicles, we have shown in the present study that, within the CL, androstenedione prevents rather than promotes cell death.
Androstenedione has been reported to be a weak androgen; however, it is the major circulating androgen during pregnancy in rats [19]. Whereas during the first week of gestation the ovaries provide androstenedione, it is the placenta that produces this androgen during the second half of pregnancy [19]. Because of its weak androgenic capacity on the AR, androstenedione may mediate its action by intracrine conversion to a more powerful androgen, such as testosterone or 5-dihydrotestosterone, which could impact the AR present in luteal cells. However, the enzyme responsible for the conversion of androstenedione to testosterone in the rat CL remains to be found. Androstenedione may mediate its antiapoptotic effect through intraluteal conversion to estradiol because of the high aromatase activity present in the rat CL [19] and the well-known luteotropic actions of estrogens in pregnancy [19]. However, this possibility was not supported by the finding that administration of 17ß-estradiol to rats after parturition did not mimic the antiapoptotic action of androstenedione (unpublished results). Androstenedione could also act through a nongenomic mechanism, as observed in luteinized human granulosa cells, in which it specifically modifies intracellular calcium concentrations, an effect that can be neither mimicked by testosterone nor blocked by a nuclear receptor antagonist [17]. However, the rapidity with which androstenedione acts in the human granulosa cells in vitro is not consistent with our results, in which androstenedione was administered in vivo and had a long-lasting effect on luteal apoptosis induced in vitro. Despite this consideration, androstenedione might still trigger primarily a short-term, nongenomic mechanism that, in turn, could lead to activation of a complex signal transduction cascade [46] and a subsequent genomic event.
Previous investigations clearly demonstrated that androstenedione is a stimulator of luteal progesterone production and secretion in rodents [21–23]. We have confirmed the strong stimulatory effect of androstenedione on the CL when administered to rats after parturition. Androstenedione, when administered in vivo, increased circulating progesterone to levels similar to those found in pregnancy and enhanced the in vitro capacity of the CL to secrete progesterone. We have previously shown that progesterone is a survival factor in the rat CL. Intraovarian administration of progesterone significantly protected the CL of pregnant rats from functional regression induced by prostaglandin F2 [26]. In in vitro incubated CL as well as in a luteal cell line, progesterone inhibited the expression of 20-hydroxysteroid dehydrogenase, an enzyme that catabolizes progesterone to a metabolite voided of progestational activity and the expression of which is a hallmark of functional luteal regression [24]. In addition, structural luteal regression observed in rats after parturition was strongly reduced by administration of progesterone [28]. The present study clearly shows that androstenedione interferes with luteal regression at least in two facets: stimulating production of progesterone, and inhibiting apoptosis. However, whether progesterone is the final intracrine executor of the protective effect elicited by androstenedione on CL function and survival remains a subject for further investigations.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by grants from the Ministry of Health, Antorchas Foundation, National Agency for Research, and University of Cuyo, Argentina (C.M.T.); The Third World Academy of Sciences, Italy (C.M.T.); and by NIH grants FIRCA 1R03 TW01049-03 (G.G., C.M.T.) and HD11119 (G.G.). 
2 Correspondence: C.M. Telleria, Department of Physiology and Biophysics, University of Illinois, 835 S. Wolcott Ave., M/C 901, Chicago, IL 60612. FAX: 312 996 1414; carlosmt{at}uic.edu
Accepted: December 17, 2001.
Received: November 6, 2001.
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, P < 0.05 and