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Biology of Reproduction 65, 704-709 (2001)
© 2001 Society for the Study of Reproduction, Inc.

Regular Article

Activin Stimulates Proliferation of Rat Ovarian Thecal-Interstitial Cells1

Antoni J. Duleba2,a, Tugce Pehlivanb, Rocco Carbonec, and Robert Z. Spaczynskib

a Yale University School of Medicine, Department of Obstetrics and Gynecology, New Haven, Connecticut 06520 b Karol Marcinkowski University School of Medical Sciences, Department of Gynecology and Obstetrics, 60-535 Poznan, Poland c Yale University School of Medicine, Yale Cancer Center, New Haven, Connecticut 06520

ABSTRACT

There is growing evidence that the function of ovarian theca-interstitial (T-I) cells may be modulated by paracrine actions of activin, inhibin, and follistatin. Furthermore, either dysregulation, dysfunction, or both, of these peptides may play a role in conditions associated with T-I hyperplasia, such as polycystic ovary syndrome (PCOS) and hyperthecosis. This study was designed to evaluate the role of activin, inhibin, and follistatin in the modulation of T-I cell proliferation. Interaction of these peptides with insulin-like growth factor-I (IGF-I), a known stimulator of T-I cell proliferation, was also assessed. Purified rat T-I cells were cultured for 48 h in chemically defined media and with or without activin (3–30 ng/ml), inhibin (3–30 ng/ml), follistatin (100 ng/ml), and/or IGF-I (10 nM). T-I cell proliferation was assessed using radiolabeled thymidine incorporation assay. Activin alone stimulated proliferation of T-I cells in a dose-dependent fashion (by up to 320% above control; P < 0.001), whereas inhibin alone or follistatin alone had no significant effect. Inhibin had also no effect on activin-induced proliferation. Follistatin significantly reduced the stimulatory effects of activin and decreased proliferation by up to 46% (P < 0.01) below the level attained in the presence of activin alone. IGF-I (10 nM), at a dose producing a near-maximal effect, increased proliferation by 175% above control (P < 0.001); insulin (10 nM) increased proliferation by 52% above control (P < 0.03). A combination of IGF-I (10 nM) and activin (30 ng/ml) resulted in a 1090% increase of proliferation above control (P < 0.001); this stimulatory effect was significantly greater than that achieved in the presence of either activin alone or IGF-I alone (P < 0.001). Similarly, a combination of insulin (10 nM) and activin (30 ng/ml) increased proliferation by 506% above control levels. Flow cytometry evaluation revealed that activin increased the proportion of actively dividing cells (in S or G2/M phase of the cell cycle) by 42% (P < 0.02), whereas IGF-I had no effect on the proportion of actively dividing cells. The present findings indicate that an activin-follistatin system may be involved in the regulation of the size of ovarian thecal-stromal compartment. In view of the synergy between activin and IGF-I, and the difference in the effects on the cell cycle distribution, stimulation of T-I proliferation by these agents is likely to be mediated via separate transduction pathways. Excess activin or insufficient follistatin may contribute to T-I hyperplasia.

activin, follistatin, growth factors inhibin, theca cells

INTRODUCTION

Activin, inhibin, and follistatin form an interactive system of glycoproteins that exert a broad range of endocrine, paracrine, and autocrine effects, including modulation of proliferation and differentiation of various tissues. These glycoproteins, originally detected within the ovary, were subsequently identified in various other organs [1–3]. Activin interacts with target cells by binding to type I and type II activin receptors [4]. Inhibin binds to type II activin receptors, possibly in competition with activin; however, a recent study presented evidence for an inhibin-binding protein that is distinct from proteins that bind activin [5]. In many systems, the effects of inhibin are the opposite of those of activin. Follistatin may attenuate the effects of activin by binding to its ß subunit, thus preventing activin from accessing its receptors.

There is growing evidence that an activin-inhibin-follistatin system plays an important role in the regulation of ovarian function, including follicular development, atresia, and steroidogenesis [6–8]. Administration of activin in vivo increases the number of large follicles and induces premature superovulation [9]. Activin also acts directly on granulosa and thecal-interstitial (T-I) cells. In granulosa cell cultures, activin augments aromatase activity, inhibits progesterone production, increases expression of FSH receptors, and stimulates mitogenic activity [10–14]. Steroidogenic effects of activin on granulosa cells are blocked by follistatin [15]. In theca cell cultures, most but not all studies have indicated that activin inhibits androgen production [16–18]. Inhibin has the opposite effects of activin on thecal androgen production and blocks activin-induced effects [16, 19, 20].

The activin-inhibin-follistatin system has been also implicated in the pathophysiology of polycystic ovary syndrome (PCOS) [2, 17, 21–23]. PCOS is characterized by oligo-anovulation and hyperandrogenism associated with increased ovarian androgen production. A linkage between PCOS and follistatin has also been recently demonstrated in an elegant population genetics study [23]. The authors proposed that PCOS may be associated with an increased availability of follistatin and thus attenuated activity of activin. Such a concept is supported by in vitro studies demonstrating inhibition of thecal androgen production by activin [16, 20]. However, these observations are difficult to reconcile with the evidence for an increased availability of activin in follicles from patients with PCOS in comparison to follicles from normal ovaries [2]. This increase of activin may be due in part to decreased expression of the inhibin alpha subunit and follistatin mRNA in PCOS follicles [2]. Ultimately, the role of activin and follistatin in the pathophysiology of PCOS remains unclear.

One of the hallmarks of PCOS is ovarian stromal and thecal hyperplasia [24, 25]. Our recent studies have demonstrated that excessive growth of ovarian T-I cells may be due to proliferative effects of insulin, insulin-like growth factors (IGFs), or both [26–28]. Others have shown that activin may modulate proliferative effects of IGFs in various tissues [29–31]. In view of the above studies, we hypothesized that the activin-inhibin-follistatin system is involved in the regulation of proliferation of ovarian mesenchyme.

MATERIALS AND METHODS

Materials

The following materials were purchased from Sigma Chemical Co. (St. Louis, MO): medium-199 with Hank balanced salt solution (HBSS), medium 199 with HBSS (10x), McCoy 5a medium (modified, without serum), L-glutamine, BSA, trypsin-EDTA (0.05%/0.02%), nitro-blue tetrazolium, 5ß-androstan-3ß-ol-17-one, ß-NAD+, sesame oil, Percoll, recombinant IGF-I, Hoechst 33342, and DiOC5. Collagenase type I (Clostridium histolyticum, CLS1; 146 U/mg) and DNase I (bovine pancreas; 2298 U/mg) were obtained from Worthington Biochemical Co. (Freehold, NY). The following materials were purchased from Grand Island Biological Co. (Grand Island, NY): trypan blue stain (0.4%; wt/vol), antibiotic-antimycotic preparation (penicillin, 10 000 IU/ml; streptomycin, 10 000 µg/ml; amphotericin B, 25 µg/ml), and Dulbecco (1x pH 7.2, without MgCl2 and CaCl2). HEPES was purchased from American Bioanalytical (Natick, MA). Radiolabeled [3H]thymidine was purchased from Amersham Life Sciences Inc. (Arlington Heights, IL).

Human recombinant activin A, human recombinant inhibin A, and human recombinant follistatin were kindly donated by Dr. A.F. Parlow, National Hormone and Pituitary Program (Torrance, CA).

Animals

Immature (25-day-old) female Sprague-Dawley rats were obtained from Taconic Farms (Germantown, NY) and housed with a 12L:12D photoperiod in an air-conditioned environment. Standard rat chow and water were given ad libitum. Starting on the 28th day of age, the animals were injected with 17ß-estradiol (1 mg/0.3 ml sesame oil s.c.) daily for 3 days in order to stimulate ovarian development. This treatment yields ovaries with medium-size follicles and actively dividing T-I cells [32]. On the morning following the last injection (Day 31 of age), the animals were anesthetized with ketamine and xylazine (i.p.) and killed by perfusion with 0.9% saline. All treatments and procedures were in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and a protocol approved by the Yale University Animal Care Committee.

Isolation of T-I Cells and Cell Cultures

Ovaries were dissected, and T-I cells were isolated and purified as described previously [26]. Briefly, ovarian follicles were punctured to release most of the granulosa cells; the remaining ovarian tissues were enzymatically digested, washed, and separated using a Percoll gradient. Each ovary yielded approximately 400 000–600 000 T-I cells. The purity of this preparation was reported previously [26]. Briefly, before plating, 95.4% of cells stained positive for a mesenchymal marker, vimentin; 7.6% of cells stained positive for an epithelial marker, cytokeratin; and 2.2% of cells stained positive for an endothelial marker, factor VIII. The cells were counted using a hemocytometer and viability was determined using a Trypan blue stain exclusion test. Cell viability was in the 85% to 95% range. The cells were cultured in McCoy 5a medium supplemented with L-glutamine (2 mM), BSA (1 mg/ml), penicillin (10 000 IU/ml), streptomycin (10 000 µg/ml), and amphotericin B (25 µg/ml). Cultures were carried out at 37°C in an atmosphere of 5% CO2 in humidified air in 24-well plates (Falcon, Becton Dickinson Labware, Lincoln Park, NJ) or 96-well plates (Corning Glass Works, Corning, NY) for 48 h. The final amount of T-I cells plated was 450 x 103 cells/ml in 24-well plates, and 35 x 103 cells/0.25 ml in 96-well plates.

DNA Synthesis Determination

Radiolabeled [3H]thymidine (1 µCi/well) was added to cultured T-I cells during the last 24 h of culture. At the end of the culture period, the cells were harvested using a multiwell cell harvester (PHD Harvester Model 290, Cambridge Technology, Inc., Watertown, MA). Radioactivity was measured in a liquid scintillation counter, SL 4000 (Intertechnique, Fairfield, NJ). Each treatment was carried out in at least six replicates.

Flow Cytometry Analysis of Cell Cycle

The assessment of the cell cycle distribution was performed by flow cytometry of propidium iodide (PI) stained T-I cells following a modified method described by others [33]. In order to maximize the number of viable cells for flow cytometry analysis, T-I cells were cultured in McCoy 5a medium supplemented with FBS (10%) for 24 h. Subsequently, the cells were washed with serum-free McCoy 5a medium and cultured for 24 h in fresh McCoy 5a serum-free medium with or without activin, IGF-I, or both. At the end of the culture period, the cells were dispersed with the aid of trypsin-EDTA solution, washed, resuspended in 2 ml of ice-cold PBS, and fixed by three stepwise additions of 2 ml each of 95% ice-cold ethanol. The fixed T-I cells were stored at 4°C awaiting flow cytometry analysis. On the day of analysis the T-I cells were spun down and resuspended in ribonuclease solution (1 mg/ml in PBS, pH 7.0) for 30 min at 37°C, and subsequently treated with PI (0.05 mg/ml) for 1 h. Prior to analysis the samples were filtered through 35-micron nylon mesh (TETKO Inc., Elmsford, NY). Flow cytometric analysis was performed with a FACS Vantage flow cytometer (Becton Dickinson Immunocytometry Systems). The cells were excited with an argon laser operating at 488 nm. PI fluorescence was collected in a linear mode through a 630/22 nm band pass filter. At least 15 000 cells were analyzed for each sample. Cell cycle analysis was performed using the Modfit 5.2 multitrapezoid model (Verity Software House, Topsham, ME). The results were expressed as the proliferative index (the sum of percentages of cells in S and G2/M phases of the cell cycle obtained from DNA histograms). Each treatment was carried out in at least four replicates.

General Considerations and Statistical Analysis

In each experiment ovaries were pooled from 10 to 12 rats. Individual experiments were repeated at least twice; results of representative experiments are presented.

Results are presented as the mean ± SEM. Comparisons between the means were performed using ANOVA followed by post-hoc comparisons of individual means using Bonferroni correction. Differences were considered significant at P < 0.05.

RESULTS

DNA Synthesis

DNA synthesis was evaluated by quantifying [3H]thy-midine incorporation. Activin (1–30 ng/ml) produced a dose-dependent increase in DNA synthesis by up to 322% above control levels (Fig. 1). In order to determine whether the effects of activin were modulated by follistatin, T-I cells were cultured with or without follistatin (100 ng/ml), activin (3 or 30 ng/ml), or both. As demonstrated in Figure 2, follistatin alone had no significant effect on DNA synthesis; however, follistatin significantly decreased activin-induced DNA synthesis (by 37% and 46% for activin at 3 and 30 ng/ml, respectively).



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  		FIG. 1. Effect of activin on incorporation of radiolabeled thymidine in cultures of rat T-I cells. The cells were cultured in serum-free medium for 48 h in 96-well plates. Each well contained 0.25 ml of medium. Bars indicate the mean of each treatment, vertical lines indicate the SEM from at least six replicates. * Means significantly different from control (P < 0.05)



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  		FIG. 2. Effects of follistatin on basal and activin-induced DNA synthesis by rat T-I cells. The cells were cultured in serum-free medium for 48 h with or without follistatin (100 ng/ml), activin (3 or 30 ng/ml), or both. Bars indicate the mean of each treatment, vertical lines indicate the SEM from at least six replicates. * Significant difference between means with and without follistatin (P < 0.05)

Figure 3 presents effects of IGF-I and insulin on basal and activin-induced DNA synthesis. IGF-I (10 nM) alone increased DNA synthesis by 175% above the control. Insulin (10 nM) alone increased DNA synthesis by 52% above the control. When the cells were exposed simultaneously to IGF-I and activin, the radiolabeled thymidine incorporation increased by 1090% above control levels, indicating that IGF-I and activin produced a synergistic effect on DNA synthesis. In a similar fashion, insulin in combination with activin increased radiolabeled thymidine incorporation by 506% above control levels.



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  		FIG. 3. Effect of insulin and IGF-I on activin-stimulated DNA synthesis by rat T-I cells. The cells were cultured in serum-free medium with or without activin (30 ng/ml), IGF-I (10 nM), insulin (10 nM), or a combination of these for 48 h. Bars indicate the mean of each treatment and vertical lines indicate the SEM from at least six replicates. * Means significantly different from control (P < 0.05). {dagger} Means significantly different from activin alone (P < 0.05)

Figure 4 summarizes the experiment determining radiolabeled thymidine incorporation in the presence or absence of inhibin (30 ng/ml), IGF-I (10 nM), activin (30 ng/ml), or a combination of these. Inhibin (3–30 ng/ml) had no significant effect on basal DNA synthesis. Furthermore, inhibin had no effect on activin- or IGF-I-induced DNA synthesis. Notably, in the above-tested system there was no positive control for the bioactivity of the inhibin preparation.



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  		FIG. 4. DNA synthesis in the presence or absence of inhibin (30 ng/ml) activin (30 ng/ml), and/or IGF-I (10 nM). The cells were cultured in serum-free medium for 48 h. Bars indicate the mean of each treatment, vertical lines indicate the SEM from at least six replicates. * Means significantly different from control (P < 0.05)

Determination of the Proliferative Index by Flow Cytometry

Figure 5 summarizes the effects of activin (30 ng/ml) and IGF-I (10 nM) on cell cycle progression of cultured T-I cells. Activin alone significantly increased the proportion of proliferating cells, as reflected by a 65% increase in the proliferative index. In contrast, IGF-I had no significant effect on basal or activin-induced proliferative index.



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  		FIG. 5. Effect of activin and IGF-I on proliferative index of rat T-I cells. Cultures were carried out in 24-well plates. The cells were cultured in McCoy 5a medium supplemented with fetal bovine serum (10%) for 24 h. Subsequently, the media were changed and the cells were incubated in serum-free medium with or without activin (30 ng/ml), IGF-I (10 nM), or both, for 24 h. At the end of the culture period the cells were dispersed and stained with propidium iodide for flow cytometry analysis. Bars indicate the mean of each treatment and vertical lines indicate the SEM from at least four replicates for each time point. * Means significantly different from control (P = 0.05)

DISCUSSION

To our knowledge, this is the first report evaluating the effects of activin on proliferation of ovarian mesenchyme. We have shown that 1) activin stimulates proliferation of T-I cells in a dose-dependent fashion; 2) follistatin attenuates activin-induced proliferation; 3) the effects of activin are synergistic with IGF-I and insulin; and 4) activin acts, at least in part, by increasing the proportion of actively dividing cells. The present findings support the notion that an activin-follistatin system may be relevant in the regulation of ovarian growth and folliculogenesis. Indeed, several in vitro studies have shown that activin promoted follicular development, whereas follistatin blocked this effect [34, 35]. Furthermore, administration of follistatin vaccine to heifers resulted in an increase in the number of ultrasonographically detectable follicles [36]. Because follicular development requires proliferation of theca cells, the balance between activin and follistatin may be important in the paracrine and/or autocrine regulation of folliculogenesis.

Activin, inhibin, and follistatin are produced predominantly by granulosa cells [2, 37, 38]. In theca cells, follistatin and the beta A subunit of activin were detected; however, there was no evidence of gene expression for these proteins [37]. The above observations indicate that follistatin and activin may act on theca cells as paracrine agents. Mounting evidence indicates that in the course of follicular development, the relative levels of activin, relative to inhibin, decreases; furthermore, increased production of follistatin in preovulatory follicles further limits effects of activin [39]. These events may be responsible for the regulation of thecal steroidogenesis and growth; specifically, as the follicle matures, the decrease of activin may lead to decreased thecal growth and increased androgen production [16].

The lack of significant fluctuations in the levels of activin and follistatin during the menstrual cycle argues against a significant endocrine role for these peptides in adult women [40, 41]. However, it is possible that activin may be relevant as an endocrine and local regulator of ovarian growth before and during puberty. This concept is supported by the evidence that at puberty, serum follistatin levels are lower than in adulthood [42], indicating thus, that during the period of intense ovarian growth, there may be greater availability of biologically active activin. In addition, the immature ovary may be more sensitive to the effects of activin; for example, activin has been shown to promote folliculogenesis in cultures of follicles from immature, but not adult, mice [43].

Activin and follistatin may also play a role in the pathophysiology of conditions such as hyperthecosis and PCOS. These disorders are characterized by prominent hyperplasia of ovarian stroma and theca [24, 25]. The present study provides arguments for the hypothesis that either increased availability of activin, or decreased follistatin (or both) may contribute to excessive growth of ovarian thecal and stromal compartments. This hypothesis is further supported by the findings of Roberts et al. [2], who demonstrated that expression of follistatin mRNA is lower in granulosa cells from polycystic ovaries. In addition, there is evidence suggesting a linkage between PCOS and the follistatin gene [23]. Excessive growth of ovarian mesenchyme may also be due to a synergy between activin with insulin and IGF-I. Women with PCOS have typically significant hyperinsulinemia and elevated, free, bioavailable IGF-I [44–47]. In these patients, insulin and IGF-I may excessively amplify the proliferative actions of activin.

Synergy between the proliferative effects of activin and insulin/IGF-I indicates that the mechanisms of action of these agents on proliferation of T-I cells are independent. Our previous dose-response experiments have demonstrated that a near-maximal stimulation of proliferation occurs at a dose of 10 nM IGF-I [26]. In the present study, the effect of this dose of IGF-I has been amplified by activin by more than sixfold. Furthermore, our flow cytometry data demonstrate that activin promotes proliferation, at least in part, by increasing the proportion of actively dividing T-I cells (cells in S and G2/M phases of the cell cycle). In contrast, IGF-I has no significant effect on the distribution of cells throughout the cell cycle. A plausible explanation of this observation may be that agents such as IGF-I can accelerate the progression through the cell cycle without affecting the proportion of dividing cells, whereas activin increases the pool of the dividing cells. Thus, activin may act, at least in part, as a competence factor. It should be noted that the T-I cells represent not a pure population of uniform cells, but an enriched preparation of theca externa, theca interna, and stromal cells. It is possible that IGF-I and activin may promote proliferation of separate subpopulations of cells; however, in that case, additive rather than synergistic effects would be more likely. The lack of the effect of IGF-I on the proportion of the dividing cells may be also due to experimental conditions; in this study, effects of IGF-I were observed only at a single time point. Review of the literature indicates that the effects of IGF-I and insulin on the cycling fraction of cells depend on the tissue source; for example, these agents increased the cycling fraction of Swiss 3T3 cells, but not human fibroblasts [48].

Effects of activin on proliferation may be tissue-dependent and possibly species-dependent. In previous studies, activin inhibited proliferation of many cell types, including rat pituitary somatotrophs, human pituitary tumors, insulin-producing INS-1 cells, rat thymocytes, LNCaP prostatic cancer cells, and human hepatocytes [49–54]. In contrast, activin stimulated proliferation of many other cells, including 3T3 fibroblasts, rat vascular smooth muscle, and avian premyocardial cells [30, 31, 52]. The effects of activin on gonadal tissues are also mixed. Thus, activin stimulated proliferation of Sertoli cells but inhibited proliferation of spermatogonia [55]. In the ovary, both stimulation and inhibition of granulosa cell proliferation by activin was reported [10, 56]. Activin also increased proliferation of several human ovarian cancer cell lines but decreased growth of Chinese hamster ovary cells [57, 58]. These divergent observations raise an interesting question: What are the characteristics predisposing given tissues to increase or decrease proliferation in response to activin? Although no clear answer is apparent, it is interesting to note that activin appears to consistently stimulate proliferation in fibroblasts and fibroblast-related tissues, including the T-I cells.

T-I cells, fibroblasts, and other fibroblast-related cells, such as vascular smooth muscle, also have similar proliferative responses to insulin and IGF-I [26, 30, 32, 59, 60]. In a similar fashion to presently observed effects on T-I cells, activin has also been shown to potentiate proliferative effects of IGF-I in cultures of vascular smooth muscle [30].

In summary, we propose that an activin-follistatin system may play a role in the regulation of growth of ovarian thecal and stromal compartments, and that under pathological conditions, dysregulation of this activin-follistatin system may lead to excessive growth of the ovarian mesenchyme.

ACKNOWLEDGMENTS

We acknowledge Dr. A.F. Parlow from the National Hormone and Pituitary Program for his kind donation of human recombinant activin A, human recombinant inhibin A, and human recombinant follistatin.

FOOTNOTES

First decision: 25 January 2001.

1 Flow cytometry studies were performed with support from the Yale Cancer Center Flow Cytometry Shared Resource, U.S. Public Health Service grant CA-16359 from the National Cancer Institute. Back

2 Correspondence: Antoni J. Duleba, Yale University School of Medicine, Department of Obstetrics and Gynecology, 333 Cedar Street, New Haven, CT 06520. FAX: 203 785 7134; antoni.duleba{at}yale.edu Back

Accepted: April 10, 2001.

Received: January 16, 2001.

REFERENCES

  1. Rabinovici J, Spencer SJ, Doldi N, Goldsmith PC, Schwall R, Jaffe RB. Activin-A as an intraovarian modulator: actions, localization, and regulation of the intact dimer in human ovarian cells. J Clin Invest 1992; 89:1528-1536
  2. Roberts VJ, Barth S, el-Roeiy A, Yen SS. Expression of inhibin/activin system messenger ribonucleic acids and proteins in ovarian follicles from women with polycystic ovarian syndrome. J Clin Endocrinol Metab 1994; 79:1434-1439[Abstract]
  3. Erickson GF, Chung DG, Sit A, DePaolo LV, Shimasaki S, Ling N. Follistatin concentrations in follicular fluid of normal and polycystic ovaries. Hum Reprod 1995; 10:2120-2124[Abstract/Free Full Text]
  4. Mathews LS. Activin receptors and cellular signaling by the receptor serine kinase family. Endocr Rev 1994; 15:310-325[CrossRef][Medline]
  5. Chong H, Pangas SA, Bernard DJ, Wang E, Gitch J, Chen W, Draper LB, Cox ET, Woodruff TK. Structure and expression of a membrane component of the inhibin receptor system. Endocrinology 2000; 141:2600-2607[Abstract/Free Full Text]
  6. Hillier SG. Regulatory functions for inhibin and activin in human ovaries. J Endocrinol 1991; 131:171-175[Medline]
  7. Peng C, Ohno T, Khorasheh S, Leung PC. Activin and follistatin as local regulators in the human ovary. Biol Signals 1996; 5:81-89[Medline]
  8. Halvorson LM, DeCherney AH. Inhibin, activin, and follistatin in reproductive medicine. Fertil Steril 1996; 65:459-469[Medline]
  9. Erickson GF, Kokka S, Rivier C. Activin causes premature superovulation. Endocrinology 1995; 136:4804-4813[Abstract]
  10. Miro F, Hillier SG. Modulation of granulosa cell deoxyribonucleic acid synthesis and differentiation by activin. Endocrinology 1996; 137:464-468[Abstract]
  11. Hutchinson LA, Findlay JK, de Vos FL, Robertson DM. Effects of bovine inhibin, transforming growth factor-beta and bovine activin-A on granulosa cell differentiation. Biochem Biophys Res Commun 1987; 146:1405-1412[CrossRef][Medline]
  12. Hillier SG, Miro F. Local regulation of primate granulosa cell aromatase activity. J Steroid Biochem Mol Biol 1993; 44:435-439[CrossRef][Medline]
  13. Nakamura M, Minegishi T, Hasegawa Y, Nakamura K, Igarashi S, Ito I, Shinozaki H, Miyamoto K, Eto Y, Ibuki Y. Effect of an activin A on follicle-stimulating hormone (FSH) receptor messenger ribonucleic acid levels and FSH receptor expressions in cultured rat granulosa cells. Endocrinology 1993; 133:538-544[Abstract]
  14. Cataldo NA, Rabinovici J, Fujimoto VY, Jaffe RB. Follistatin antagonizes the effects of activin-A on steroidogenesis in human luteinizing granulosa cells. J Clin Endocrinol Metab 1994; 79:272-277[Abstract]
  15. Wrathall JH, Knight PG. Effects of inhibin-related peptides and oestradiol on androstenedione and progesterone secretion by bovine theca cells in vitro. J Endocrinol 1995; 145:491-500[Abstract]
  16. Hillier SG, Yong EL, Illingworth PJ, Baird DT, Schwall RH, Mason AJ. Effect of recombinant activin on androgen synthesis in cultured human thecal cells. J Clin Endocrinol Metab 1991; 72:1206-1211[Abstract]
  17. Sawetawan C, Carr BR, McGee E, Bird IM, Hong TL, Rainey WE. Inhibin and activin differentially regulate androgen production and 17 alpha-hydroxylase expression in human ovarian thecal-like tumor cells. J Endocrinol 1996; 148:213-221[Abstract]
  18. Shukovski L, Dyson M, Findlay JK. The effects of follistatin, activin and inhibin on steroidogenesis by bovine thecal cells. Mol Cell Endocrinol 1993; 97:19-27[CrossRef][Medline]
  19. Hsueh AJ, Dahl KD, Vaughan J, Tucker E, Rivier J, Bardin CW, Vale W. Heterodimers and homodimers of inhibin subunits have different paracrine action in the modulation of luteinizing hormone-stimulated androgen biosynthesis. Proc Natl Acad Sci U S A 1987; 84:5082-5086[Abstract/Free Full Text]
  20. Nahum R, Thong KJ, Hillier SG. Metabolic regulation of androgen production by human thecal cells in vitro. Hum Reprod 1995; 10:75-81[Abstract/Free Full Text]
  21. Yamoto M, Minami S, Nakano R. Immunohistochemical localization of inhibin subunits in polycystic ovary. J Clin Endocrinol Metab 1993; 77:859-862[Abstract]
  22. Dewailly D. Physiopathology of polycystic ovary syndrome [in French]. Ann Endocrinol (Paris) 1999; 60:123-130[Medline]
  23. Urbanek M, Legro RS, Driscoll DA, Azziz R, Ehrmann DA, Norman RJ, Strauss 3rd JF, Spielman RS, Dunaif A. Thirty-seven candidate genes for polycystic ovary syndrome: strongest evidence for linkage is with follistatin. Proc Natl Acad Sci U S A 1999; 96:8573-8578[Abstract/Free Full Text]
  24. Hughesdon PE. Morphology and morphogenesis of the Stein-Leventhal ovary and of so called "hyperthecosis". Obstet Gynecol Surv 1982; 37:59-77[Medline]
  25. Lappohn RE, Bogchelman DH. The relation of fertility and ovarian histology after bilateral ovarian wedge resection. Fertil Steril 1989; 52:221-226[Medline]
  26. Duleba AJ, Spaczynski RZ, Olive DL, Behrman HR. Effects of insulin and insulin-like growth factors on proliferation of rat ovarian theca-interstitial cells. Biol Reprod 1997; 56:891-897[Abstract]
  27. Duleba AJ, Spaczynski RZ, Olive DL. Insulin and insulin-like growth factor I stimulate the proliferation of human ovarian theca-interstitial cells. Fertil Steril 1998; 69:335-340[CrossRef][Medline]
  28. Duleba AJ, Spaczynski RZ, Olive DL, Behrman HR. Divergent mechanisms regulate proliferation/survival and steroidogenesis of theca-interstitial cells. Mol Hum Reprod 1999; 5:193-198[Abstract/Free Full Text]
  29. Kojima I, Mogami H, Kawamura N, Shibata H. Activin A inhibits cell-cycle progression induced by insulin-like growth factor-I in Balb/c 3T3 cells. J Endocrinol 1993; 137:99-105[Abstract]
  30. Kojima I, Mogami H, Kawamura N, Yasuda H, Shibata H. Modulation of growth of vascular smooth muscle cells by activin A. Exp Cell Res 1993; 206:152-156[CrossRef][Medline]
  31. Antin PB, Yatskievych T, Dominguez JL, Chieffi P. Regulation of avian precardiac mesoderm development by insulin and insulin-like growth factors. J Cell Physiol 1996; 168:42-50[CrossRef][Medline]
  32. Duleba AJ, Spaczynski RZ, Arici A, Carbone R, Behrman HR. Proliferation and differentiation of rat theca-interstitial cells: comparison of effects induced by platelet-derived growth factor and insulin-like growth factor-I. Biol Reprod 1999; 60:546-550[Abstract/Free Full Text]
  33. Crissman HA, Hirons GT. Staining of DNA in live and fixed cells. Methods Cell Biol 1994; 41:195-209[Medline]
  34. Li R, Phillips DM, Mather JP. Activin promotes ovarian follicle development in vitro. Endocrinology 1995; 136:849-856[Abstract]
  35. Smitz J, Cortvrindt R, Hu Y, Vanderstichele H. Effects of recombinant activin A on in vitro culture of mouse preantral follicles. Mol Reprod Dev 1998; 50:294-304[CrossRef][Medline]
  36. Singh J, Brogliatti GM, Christensen CR, Adams GP. Active immunization against follistatin and its effect on FSH, follicle development and superovulation in heifers. Theriogenology 1999; 52:49-66[CrossRef][Medline]
  37. Roberts VJ, Barth S, el-Roeiy A, Yen SS. Expression of inhibin/activin subunits and follistatin messenger ribonucleic acids and proteins in ovarian follicles and the corpus luteum during the human menstrual cycle. J Clin Endocrinol Metab 1993; 77:1402-1410[Abstract]
  38. Shintani Y, Dyson M, Drummond AE, Findlay JK. Regulation of follistatin production by rat granulosa cells in vitro. Endocrinology 1997; 138:2544-2551[Abstract/Free Full Text]
  39. Hillier SG, Miro F. Inhibin, activin, and follistatin. Potential roles in ovarian physiology. Ann N Y Acad Sci 1993; 687:29-38[Abstract]
  40. Demura R, Suzuki T, Tajima S, Mitsuhashi S, Odagiri E, Demura H, Ling N. Human plasma free activin and inhibin levels during the menstrual cycle. J Clin Endocrinol Metab 1993; 76:1080-1082[Abstract]
  41. Khoury RH, Wang QF, Crowley WF Jr, Hall JE, Schneyer AL, Toth T, Midgley AR, Jr, Sluss PM. Serum follistatin levels in women: evidence against an endocrine function of ovarian follistatin. J Clin Endocrinol Metab 1995; 80:1361-1368[Abstract]
  42. Kettel LM, DePaolo LV, Morales AJ, Apter D, Ling N, Yen SS. Circulating levels of follistatin from puberty to menopause. Fertil Steril 1996; 65:472-476[Medline]
  43. Yokota H, Yamada K, Liu X, Kobayashi J, Abe Y, Mizunuma H, Ibuki Y. Paradoxical action of activin A on folliculogenesis in immature and adult mice. Endocrinology 1997; 138:4572-4576[Abstract/Free Full Text]
  44. Burghen GA, Givens JR, Kitabchi AE. Correlation of hyperandrogenism with hyperinsulinism in polycystic ovarian disease. J Clin Endocrinol Metab 1980; 50:113-116[Abstract]
  45. Dunaif A, Graf M, Mandeli J, Laumas V, Dobrjansky A. Characterization of groups of hyperandrogenic women with acanthosis nigricans, impaired glucose tolerance, and/or hyperinsulinemia. J Clin Endocrinol Metab 1987; 65:499-507[Abstract]
  46. Iwashita M, Mimuro T, Watanabe M, Setoyama T, Matsuo A, Adachi T, Takeda Y, Sakamoto S. Plasma levels of insulin-like growth factor-I and its binding protein in polycystic ovary syndrome. Horm Res 1990; 33:21-26
  47. Homburg R, Pariente C, Lunenfeld B, Jacobs HS. The role of insulin-like growth factor-1 (IGF-1) and IGF binding protein-1 (IGFBP-1) in the pathogenesis of polycystic ovary syndrome. Hum Reprod 1992; 7:1379-1383[Abstract/Free Full Text]
  48. Chen YY, Rabinovitch PS. Mitogen response and cell cycle kinetics of Swiss 3T3 cells in defined medium: differences from human fibroblasts and effects of cell density. Exp Cell Res 1990; 190:145-150[CrossRef][Medline]
  49. Billestrup N, Gonzalez-Manchon C, Potter E, Vale W. Inhibition of somatotroph growth and growth hormone biosynthesis by activin in vitro. Mol Endocrinol 1990; 4:356-362[Abstract]
  50. Danila DC, Inder WJ, Zhang X, Alexander JM, Swearingen B, Hedley-Whyte ET, Klibanski A. Activin effects on neoplastic proliferation of human pituitary tumors. J Clin Endocrinol Metab 2000; 85:1009-1015[Abstract/Free Full Text]
  51. Huotari MA, Palgi J, Otonkoski T. Growth factor-mediated proliferation and differentiation of insulin-producing INS-1 and RINm5F cells: identification of betacellulin as a novel beta-cell mitogen. Endocrinology 1998; 139:1494-1499[Abstract/Free Full Text]
  52. Hedger MP, Drummond AE, Robertson DM, Risbridger GP, de Kretser DM. Inhibin and activin regulate [3H]thymidine uptake by rat thymocytes and 3T3 cells in vitro. Mol Cell Endocrinol 1989; 61:133-138[CrossRef][Medline]
  53. Dalkin AC, Gilrain JT, Bradshaw D, Myers CE. Activin inhibition of prostate cancer cell growth: selective actions on androgen-responsive LNCaP cells. Endocrinology 1996; 137:5230-5235[Abstract]
  54. Zauberman A, Oren M, Zipori D. Involvement of p21(WAF1/Cip1), CDK4 and Rb in activin A mediated signaling leading to hepatoma cell growth inhibition. Oncogene 1997; 15:1705-1711[CrossRef][Medline]
  55. Boitani C, Stefanini M, Fragale A, Morena AR. Activin stimulates Sertoli cell proliferation in a defined period of rat testis development. Endocrinology 1995; 136:5438-5444[Abstract]
  56. Woodruff TK, Lyon RJ, Hansen SE, Rice GC, Mather JP. Inhibin and activin locally regulate rat ovarian folliculogenesis. Endocrinology 1990; 127:3196-3205[Abstract]
  57. Di Simone N, Crowley WF Jr, Wang QF, Sluss PM, Schneyer AL. Characterization of inhibin/activin subunit, follistatin, and activin type II receptors in human ovarian cancer cell lines: a potential role in autocrine growth regulation. Endocrinology 1996; 137:486-494[Abstract]
  58. Gonzalez-Manchon C, Vale W. Activin-A, inhibin and transforming growth factor-beta modulate growth of two gonadal cell lines. Endocrinology 1989; 125:1666-1672[Abstract]
  59. Pfeifle B, Ditschuneit H. Effect of insulin on growth of cultured human arterial smooth muscle cells. Diabetologia 1981; 20:155-158[CrossRef][Medline]
  60. Conover CA, Rosenfeld RG, Hintz RL. Hormonal control of the replication of human fetal fibroblasts: role of somatomedin C/insulin-like growth factor I. J Cell Physiol 1986; 128:47-54[CrossRef][Medline]



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