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Monash Institute of Reproduction and Development (J.J.B., D.M.d.K., A.E.O’C., J.R.M.) and Department of Anatomy and Cell Biology (J.J.B., N.G.W.), Monash University; and Prince Henry’s Institute of Medical Research (P.G.F.), Clayton, Victoria 3168, Australia
Address all correspondence and requests for reprints to: John R Morrison, Monash Institute of Reproduction and Development, 27–31 Wright Street, Clayton, Victoria 3168, Australia. E-mail:john.morrison{at}med.monash.edu.au.
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Abstract |
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Introduction |
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In the female, activin has been demonstrated to have mitogenic effects on granulosa cells (the female equivalent of Sertoli cells) in the ovary, especially in combination with FSH (10, 11). In cultured testis fragments obtained from d 9 rats, treatment with activin A and FSH resulted in a synergistic increase in Sertoli cell proliferation (compared with untreated fragments or fragments treated with FSH alone) as determined by 5-bromo-2'deoxyuridine incorporation (8). No significant effect was observed in fragments isolated from d 3 or d 18 rats. Further investigation of the window of activin responsiveness led to the demonstration that in testis fragments, Sertoli cells become significantly responsive to activin at some time between d 5 and 7 pp and become unresponsive to activin at some time between d 11 and 13 pp (12). Contrasting with these findings, treatment of d 3 testis fragment cultures with activin A resulted in a significant decrease in Sertoli cell proliferation in both the presence and absence of FSH (9). Treatment of these d 3 cultures with the activin binding-protein follistatin also suppressed Sertoli cell proliferation, suggesting a role for endogenous activin or bone morphogenetic proteins (which are also antagonized by follistatin) (9). Of note, treatment of d 3 testis fragments with activin was found to induce proliferation of gonocytes (9), and at later stages of development, activin has been demonstrated to promote spermatogonial proliferation (13), and to maintain mitochondrial differentiation in spermatocytes (14), suggesting a multifaceted role for activin during testis development.
A direct role for activin A in the regulation of Sertoli cell development would require evidence of the receptor to be present in, and the ligand available to Sertoli cells. Immunohistochemical analysis demonstrates expression of the ßA and ßB subunits of activin/inhibin in both the Sertoli cells and isolated interstitial cells in d 12 rats (15), whereas medium conditioned by cultured primary Sertoli cells isolated from d 21 rats or Sertoli derived TM-4 cells contains activin-like activity (16). These findings led to a widespread assumption that, within the seminiferous tubule, activins are produced by Sertoli cells as autocrine factors. However, peritubular myoid cells have also been demonstrated to express high levels of activin (17). Furthermore, most primary Sertoli cell cultures contain significant degrees of contamination by peritubular cells (18, 19), whereas the TM-4 Sertoli cell line is known to be highly mutated (20) and hence is unlikely to be representative of normal Sertoli cells. It is also quite likely that the profile of activin expression in the testis may change significantly after Sertoli cells differentiate and during postnatal development.
The activin receptors are divided into two subtypes, type II receptors (ActRIIa and ActRIIb) are ligand binding receptors, which then recruit type I receptors that initiate the signal transduction cascade. Within the testis, there is a relatively widespread expression of activin receptors. ActRIIa message has been demonstrated by Northern blot in isolated pachytene spermatocytes, round spermatids, Sertoli cells, and at low levels in isolated Leydig cells (21). In situ hybridization has confirmed the expression of ActRIIa message in spermatocytes, spermatids, and Sertoli cells, but not Leydig cells (22). ActRIIb message on the other hand has been demonstrated by in situ hybridization in primary spermatocytes and Sertoli cells (23). A transient up-regulation of ActRIIa has been reported in Sertoli cells between d 7 and 9 pp (12). No studies appear to have addressed the specific expression of activin type I receptors in the testis.
While testis fragment cultures are very effective for investigating the action of exogenous factors on testis development, it is difficult to elucidate precise local actions. Since testis fragments contain Sertoli cells, peritubular cells, Leydig cells, testicular macrophages, blood vessels, gonocytes, and spermatogonia, it is possible that a hormone or growth factor applied to such a culture system will exert effects on many different cell types, in some cases inducing expression of further paracrine factors that may affect Sertoli cells. Furthermore, because activin is produced by the testis, it is difficult to delineate the effects of endogenous activin from activin applied to the fragment (16, 17, 24).
To investigate the role of activin A on Sertoli cell development in the absence of other cell types, we have used a high purity, proliferative phase Sertoli cell culture method that has been recently developed in our laboratory. These methods clearly demonstrate that activin A is capable of directly inducing mitosis in Sertoli cells isolated from d 6 and d 9 rat pups. Sertoli cells isolated from d 3 rat pups are initially unresponsive to activin A but become responsive after 8 d in culture. A functional role for activin on Sertoli cells is further supported by the finding that activin A induces secretion of both inhibin A and B by the Sertoli cell. Furthermore, we show that Sertoli cells isolated from d 6 rats produce undetectable amounts of activin A, whereas peritubular cells from the same age produce significant quantities, suggesting that peritubular cells are a major source of activin during early postnatal testicular development.
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Materials and Methods |
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Sertoli cells were isolated according to previously described methods (25). Briefly, testes were removed, decapsulated, and diced. The resultant fragments of seminiferous cords were rinsed in DMEM/F12 medium (no. 12400-024, Life Technologies, Inc., Rockville, MD), then digested with 3 x 105 U/liter collagenase type II (no. C6885, Sigma) in DMEM/F12 at 22 C for 1 h, washed and then treated with 0.05% wt/vol hyaluronidase (no. H2126, Sigma) in DMEM/F12 for 1 h at 22 C. The seminiferous cords were then rinsed and plated down onto culture dishes coated with 1 µg/cm2 laminin (no. L2020, Sigma) in DMEM/F12 containing insulin, transferrin and selenium (ITS concentrate no. 41400-045, Life Technologies, Inc.), and penicillin/streptomycin (Life Technologies, Inc.). After 24 h of culture, medium was changed and some cultures were exposed to media supplemented with recombinant human (rh) activin A (R&D Systems, Minneapolis, MN; 0–100 ng/ml), rhFSH (Organon, 0–390 IU/liter), rh inhibin A (0–100 ng/ml) (26), and/or bovine follistatin (0–200 ng/ml) (27) alone or in combinations.
Suspended peritubular cells were recovered from the supernatant following collagenase digestion of d 6 pp rat seminiferous cords as previously described (28). Suspended cells were centrifuged at 2000 x g and rinsed once before being seeded onto culture vessels in DMEM/F12 with penicillin, streptomycin, and 10% fetal calf serum. Cells were grown to confluence and passaged once before use.
When required, cells were passaged by treatment with a volume of trypsin/versene solution containing 0.1% wt/vol trypsin (T2021, Sigma), 0.5 mM tetrasodium EDTA (E6511, Sigma) in PBS for 5 min at 37 C followed by 2 vol 0.05% wt/vol soybean trypsin inhibitor (no. T6522, Sigma) in DMEM/F12. Recovered cells were counted on a hemocytometer using trypan blue exclusion to determine viability. Viable cells were rinsed and replated at 4 x 104 cells/cm2.
Proliferation assay
[Methyl-3H]-thymidine incorporation was used as a measure of proliferative activity. [Methyl-3H]-thymidine (2.0 Ci/mmol, 3 x 106cpm/ml media, Amersham Biosciences, Castle Hill, New South Wales, Australia) was added to the culture medium for 6 h. Cells were harvested using a Micromate 196 cell harvester (Packard Instruments, Meriden, CT), and incorporated radionucleotide counted using a liquid scintillation counter. All [3H]-thymidine incorporation assays were performed in quadruplicate in at least three separate experiments.
Inhibin and activin assays
Inhibin A and B concentrations were measured using specific ELISAs (29, 30) according to the manufacturer’s instructions (Oxford Bio-Innovations, Oxfordshire, UK) with some modifications. Standard (inhibin A, WHO 91/784; inhibin B, WHO 96/78) and samples were diluted in unconditioned culture media containing the same additives as were used in the culture. Samples were then treated as per the manufacturer’s protocol. Duplicate samples were added to the plates and incubated overnight at room temperature. The plates were washed and alkaline phosphatase-conjugated inhibin detection antibody was added for 3 h at room temperature. After washing, the alkaline phosphatase activity was detected using an amplification kit (ELISA Amplification System, Life Technologies, Inc.) as per the manufacturer’s instructions; the substrate was then incubated for 2 h at room temperature. Inhibin A assays were all performed on the same plate; the intraplate % coefficient of variation (%CV) was 9% and the limit of detection for the assay was 4 pg/ml. Inhibin B assays were performed in three plate; the intraplate %CV was 8%, the interplate %CV was 9% and the limit of detection was 5 pg/ml. Samples were diluted out in a dose-dependent manner and were parallel to the standard curve in both assays (data not shown).
Activin A concentrations were measured using a specific ELISA (31) according to the manufacturer’s instructions (Oxford Bio-Innovations) with some modifications. The standard used was rh activin A as previously described (32). Standards and samples were diluted in unconditioned culture media containing the same additives as were used in the culture. Six percent sodium dodecyl sulfate in 0.05 M PBS solution was added (3% final concentration) followed by boiling for 3 min to denature proteins and remove the interference of follistatin and other known sources of error. The samples were allowed to cool before the addition of H2O2 (2% final concentration) and subsequent 30 min incubation. A total of 25 µl 20% BSA/0.1 M Tris/5% Triton X-100/0.9% NaCl/0.1%NaN3 was added to each well before the addition of the treated samples. Duplicates were added to the E4 (anti-ßA subunit) monoclonal antibody coated plate and incubated overnight at room temperature. The plate was washed and the detection antibody (biotinylated-E4) was added for 2 h at room temperature. After washing, alkaline phosphatase linked to streptavidin was added to the wells and incubated at room temperature for 1 h. After further washes, the alkaline phosphatase activity was detected using an amplification kit (ELISA Amplification System, Life Technologies, Inc.) as per the manufacturer’s instructions; the substrate was then incubated for 1 h at room temperature. All samples were measured in a single plate, the intraplate %CV was 7%, and the limit of detection for the assay was 0.01 ng/ml. Media samples were diluted out in a dose-dependent manner and were parallel to the standard. One hundred percent recoveries were obtained from spiked samples (Okuma, Y., D. M. de Kretser, and M. P. Hedger, manuscript in preparation).
RNA isolation
Cells for RNA isolation were cultured in laminin coated six-well culture dishes (Falcon, North Ridge, New South Wales, Australia). Cells were harvested by treatment with trypsin/versene for 5 min at 37 C, counted on a hemocytometer, and 2 x 106 cells pelleted at 500 x g. Poly A mRNA was isolated using oligo-(deoxythymidine)25 magnetic dynabeads (DynAl, Lake Success, CA) according to the manufacturer’s instructions. mRNA was eluted into 10 mM Tris-HCl (pH 7.5) and stored at -70 C.
Reverse transcription (RT)
RT was performed using Superscript II (Life Technologies, Inc.) according to the manufacturer’s instructions, primed with oligo-(deoxythymidine)10–15 (Pharmacia, Uppsala, Sweden). For every sample, a no RT control was performed in which all incubations and buffers were identical but no Superscript enzyme was added. This control verified the absence of contaminating genomic DNA in PCRs (data not shown).
PCR primers
PCR primers (sequences shown in Table 1
) were designed using primer3 (http://www-genome.wi.mit.edu/cgi-bin/primer/primer3_www.cgi) to previously described sequences for glyceraldehyde-3-phosphate dehydrogenase (GAPDH, accession no. NM_017008.1), ß-actin (NM031144.1), activin receptor type I (ActRI; L19341.1), activin receptor type IIa (ActRIIa; L10639.1), activin receptor type IIb (ActRIIb; M87067.1), follistatin (NM012561.1), and FSH receptor (FSHr; L02842.1). Primers were synthesized by SigmaGenosys Australia (Castle Hill, New South Wales, Australia). For each primer set, annealing temperature was optimized by performing PCR on identical replicates in a gradient block thermocycler (PCR express with 96-well gradient block, Hybaid, Ashford, UK). PCR was performed in a 25-µl solution containing 0.5 U of Taq polymerase (Pharmacia), 10 mM Tris-HCl, 1.5 mM MgCl2, 50 mM KCl, 0.2 mM deoxynucleotide triphosphates (Biotech, Roseville, New South Wales, Australia), 1 mM forward and reverse primers, and 1 µl of cDNA from the RT reaction. The initial denaturation step was at 94 C for 3 min, followed by 35 cycles of denaturation for 30 sec at 94 C, annealing for 30 sec at 54–66 C (described below), and extension for 1–1.5 min (depending upon product size) at 72 C. The reaction was subjected to a final extension for 3 min at 72 C before being electrophoresed at 90 V on a 1% agarose gel stained with ethidium bromide. The annealing temperature used was the highest temperature at which a large amount of specific PCR product could be seen when run on an agarose gel. For all primer sets tested, PCR at this temperature yielded no products of size other than that expected.
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Real-time PCR
For quantitative PCR, a real-time fluorometric capillary based thermocycler was used (LightCycler, Roche Diagnostics, Basel, Switzerland). PCR was performed in a 10-µl solution containing SYBR green PCR master mix (1x, Roche), MgCl2 (concentrations determined empirically; final concentrations used shown in Table 1
), 0.1 mM primers, and 0.2 µl of cDNA from the RT reaction. To minimize pipetting error, solutions were diluted such that 4 µl was the smallest volume pipetted at any time. PCR was performed as follows: 10 min initial denaturation at 95 C, followed by 45 cycles of 95 C denaturation for 10 sec, annealing at 59–63 C (see Table 1) for 5 sec and extension at 72 C for 5–20 sec (see Table 1). DNA content was measured by fluorometry at the end of each extension step. At the end of the PCR, melting curve analysis was performed to verify product specificity by increasing temperature from 66–95 C at 0.1 C per second, measuring fluorescence constantly. In each reaction, a standard curve was prepared from serial dilutions of an arbitrary standard.
PCR products were quantified using the LightCycler software. These values were controlled for RT efficiency and cDNA loading by normalizing with an endogenous control (GAPDH). Normalized values were then calibrated against values for untreated control Sertoli cells to give a rational value.
Radiolabeled activin A binding
Sertoli cells were plated at 0.25, 0.5, 1, and 2 x 106 cells/well in 24-well plates and maintained as described above in the presence or absence of FSH and/or activin A for 3 d before assay.
Recombinant human carrier-free activin A (1 µg, R&D Systems) was iodinated by a chloramine-T method (33) using Na125I (ICN Australia, Seven Hills, New South Wales, Australia), yielding [125I]activin of specific activity 72 µCi/µg. Cell cultures were washed twice with binding assay buffer [(DMEM/F12 medium supplemented with 50 mM HEPES, 0.1% BSA, 0.4 mM EDTA, and 50 ng/ml phenylmethylsulfonylfluoride (Sigma)]. Monolayers of Sertoli cells were exposed to activin tracer at 32,000 cpm per 0.25 ml assay buffer/well (
40 pM), in the absence or presence of unlabeled activin (0.003–10 nM) for 4 h at 26 C, with rotary mixing at 30 cycles/min. Nonspecific binding to duplicate Sertoli cell cultures that had received each hormone treatment was determined using an excess of unlabeled activin (10 nM), and was subtracted from total binding data to assess specific binding. After removal of the assay mixture, cultures were placed on ice and washed three times with ice-cold PBS, then each monolayer was dissolved in 0.10 ml of 0.1% Triton X-100 in PBS, and bound [125I]activin in the recovered lysate and a single rinse was pooled for subsequent 
radioactivity determination. Binding data for Sertoli cells plated at different cell densities were corrected for cell number, determined by counting cells in duplicate cultures that received the same treatments in parallel.
Statistical analyses
All statistical analyses involved one-way ANOVA followed by Tukey’s post hoc test. Differences between groups were deemed significant if P < 0.05.
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Results |
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). In the presence of a maximal stimulating dose of FSH, activin A synergized with FSH in a dose-dependent manner by inducing Sertoli cell division to reach a plateau at 50 ng/ml representing a 24-fold increase in thymidine uptake compared with cells treated with FSH alone (Fig. 1B). The effects of activin were blocked by addition of the activin antagonist, follistatin.
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). Inhibin B secretion was significantly stimulated (P < 0.001) by both FSH (1.5-fold) and activin A (2.3-fold) independently, whereas the combination of these factors resulted in a 13-fold increase (Fig. 2B). After 3 d of culture, there were no significant differences in the numbers of Sertoli cells per well between treatment groups (data not shown).
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; follistatin protein could not be determined because the follistatin RIA (34) does not detect rat follistatin]. To verify that follistatin was not responsible for the increased Sertoli cell proliferation in activin A treated cultures, we assessed the influence of various doses of follistatin on thymidine incorporation by Sertoli cells. Follistatin did not significantly affect Sertoli cell thymidine incorporation over a wide range of concentrations (10–200 ng/ml) in either the presence and absence of FSH (Fig. 3B).
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ActRI expression was significantly increased in response to combined FSH and activin A and this increase could be inhibited by follistatin. No significant changes in ActRI expression were observed in any other treatment groups (Fig. 4A). ActRIIa expression was not changed by FSH or activin A alone, but was significantly suppressed in response to FSH and activin A in combination. This action of FSH and activin A was completely reversed by the addition of follistatin to the culture. In contrast, ActRIIa expression increased significantly in response to the combination of activin A and follistatin, but when FSH was added, this increase was suppressed (Fig. 4B).
ActRIIb expression was significantly suppressed in response to FSH and activin A in combination, and the addition of follistatin reversed this change (Fig. 4C). FSH receptor expression was not significantly altered by any treatment (Fig. 4D). Follistatin expression was significantly increased in response to activin A (P < 0.001) and the response to FSH and activin A in combination was significantly (P < 0.001) greater than activin A alone. Again, the addition of follistatin to the various activin A treated cultures suppressed the increases (Fig. 4E).
Sertoli cells express activin receptors, and expression is increased by treatment with FSH and activin A
Binding of [125I]activin A to Sertoli cells (1 x 106 cell/well) was determined after 3 d of culture in the presence of FSH. Scatchard analysis of the binding data revealed a population of high affinity binding sites (Kd of 100 pM; 280 sites/cell) and a population of low affinity binding sites (Kd of 32 nM; 11,000 sites/cell; Fig. 5A).
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). Treatment of cultures with activin A also caused a small but insignificant increase in the specific binding of activin, but the combination of FSH with activin increased binding 8-fold above untreated Sertoli cells (P < 0.01; Fig. 5B).
Responsiveness of Sertoli cells to activin A is age related
To investigate the previously reported age-dependent activin response (8), we isolated Sertoli cells from d 3, 6, and 9 pp rats and cultured them with FSH and/or activin A for 2 or 8 d before measuring thymidine incorporation and mRNA expression. When treated with FSH, Sertoli cells collected from d 3 pups and cultured for 2 d showed a significant increase in thymidine uptake. Activin A, alone or in combination with FSH, did not affect thymidine uptake in these cultures (Fig. 6A). In contrast, when cultured for 8 d, these d 3 Sertoli cells had acquired responsiveness to activin A, and showed an augmented response to treatment with activin and FSH together, becoming indistinguishable in their response from Sertoli cells collected from d 6 pups. Sertoli cells collected from d 6 (Fig. 6B) and d 9 (Fig. 6C) rats were responsive to activin A after both 2 and 8 d in culture.
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, A–C). When cultured for a 8 d, d 3 Sertoli cells, which acquire activin A responsiveness over this period, had significant increases in both ActRIIa and ActRIIb expression compared with the more moderate increases in expression seen by d 6 Sertoli cells over this culture period (Fig. 7, B and C). Day 6 Sertoli cells did also show a significant increase in ActRI expression over this longer culture period (Fig. 7A).
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Discussion |
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Our initial experiments indicated that activin A alone was capable of stimulating d 6 Sertoli cell proliferation to a degree similar to that observed for the classical Sertoli cell mitogen FSH. In combination, activin A and FSH synergistically stimulated Sertoli cell proliferation, as has been previously reported in testis fragment cultures by Boitani and co-workers (8, 12). We demonstrate that synergy between activin A and FSH is not due to inhibin or follistatin exerting a mitogenic effect on Sertoli cells. When we investigated the expression of mRNA encoding follistatin and receptors for activin and FSH, it appeared that the changes in expression observed would resist the synergy between activin and FSH rather than reinforcing it. Direct measurement of activin receptors using binding of radiolabeled activin; however, demonstrated that treatment with activin and FSH caused an increase in the number of activin receptors per cell. Taken together, this data suggests that treatment with activin and FSH cause activin receptor complexes to be stabilized rather than inducing increased expression. This effect could account, at least in part, for the dramatic increase in Sertoli cell mitosis induced by FSH and activin together.
Further to the stimulation of Sertoli cell proliferation, activin A showed a capacity to induce expression of inhibins (Fig. 2), especially in the presence of FSH. In humans, Sertoli cell proliferation is marked in the late fetal stages and during the first 2 yr of life (35), which correlates well with circulating inhibin B levels (36). Following birth, there is a significant elevation of serum inhibin B, which relates to increased gonadotropin levels; in hypogonadotropic patients, in whom Sertoli cell division is suppressed, inhibin B levels remain low, but rise in response to treatment with exogenous FSH (37, 38). Taken together, previous data indicate that inhibin B is a good marker for the Sertoli cells response to the mitogenic activity of FSH (39). The data presented in this paper indicate that activin A is similarly capable of inducing both proliferation, and inhibin B (which in vivo would lead to decreased FSH secretion by the pituitary). Activin A can thus contribute both to the generation of Sertoli cells, and the feedback loop that keeps FSH secretion in check.
Given the concentrations of activin A required to elicit a response in Sertoli cells, we hypothesized (as others have) that activin A may be primarily a paracrine or autocrine factor. To investigate this, we measured the secretion of activin A by both Sertoli and peritubular cells in vitro. We found that Sertoli cell secretion of activin A was undetectable, whereas peritubular cells secreted relatively large quantities (Fig. 8), suggesting a primarily paracrine (rather than autocrine) role during testicular development. This finding could explain the discrepancies between the findings of Fragale et al. (12) and the present study; Fragale et al. (12) showed that, in testis fragment cultures, exogenous activin only elicits a Sertoli cell proliferative response when applied in combination with FSH. In this paper, we show that isolated Sertoli cells are capable of proliferating in response to activin alone, albeit at lower levels than in combination with FSH. If peritubular cells are a major source of activin A in the testis, then testis fragments necessarily secrete endogenous activin A, which is likely to impact upon Sertoli cell proliferation even in the absence of exogenous factors. It is likely, therefore, that Sertoli cells in testis fragments are not significantly responsive to exogenous activin A because they are already stimulated by endogenous activin A.
Previous studies have demonstrated that Sertoli cells contain activin receptors, that the testis produces activin and that activin may affect Sertoli cell development. Our data provide strong evidence that activin A is capable of directly inducing Sertoli cell development. Given the observed synergy between activin A and FSH, the age-dependent responsiveness of Sertoli cells to activin and the high levels of activin A secreted by peritubular cells, we suggest that in vivo, activin A provides a means for peritubular cells to exert paracrine control over seminiferous epithelium development at the end of the Sertoli cell proliferative phase. This hypothesis is consistent with the notion that activin promotes proliferation and suppresses differentiation of gonocytes during postnatal testis development (9). Together, these mechanisms would allow coordinated development of gonocytes and Sertoli cells, preventing premature gonocyte differentiation.
During the window of activin responsiveness, minor changes to the rate of Sertoli cell proliferation could have major consequences in defining the number of Sertoli cells present in the adult. Furthermore, because activin A is also capable of inducing expression of inhibin by Sertoli cells, it may contribute to the pituitary-gonadal feedback loop in a similar manner to FSH. Because Sertoli cell number is a key regulator of sperm production and fertility, this mechanism could provide a further level of control over testis growth and subsequent fertility.
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Acknowledgments |
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Footnotes |
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Abbreviations: ActRIIa and ActRIIb, Activin receptor types I and II; %CV, % coefficient of variation; pp, post partum; rh, recombinant human; RT, reverse transcription.
Received June 6, 2002.
Accepted for publication October 4, 2002.
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J. J. Buzzard, K. L. Loveland, M. K. O'Bryan, A. E. O'Connor, M. Bakker, T. Hayashi, N. G. Wreford, J. R. Morrison, and D. M. de Kretser Changes in Circulating and Testicular Levels of Inhibin A and B and Activin A During Postnatal Development in the Rat Endocrinology, July 1, 2004; 145(7): 3532 - 3541. [Abstract] [Full Text] [PDF]
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 Y. Xia, Y. Sidis, and A. Schneyer Overexpression of Follistatin-Like 3 in Gonads Causes Defects in Gonadal Development and Function in Transgenic Mice Mol. Endocrinol., April 1, 2004; 18(4): 979 - 994. [Abstract] [Full Text] [PDF]
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J. J. Buzzard, N. G. Wreford, and J. R. Morrison Thyroid Hormone, Retinoic Acid, and Testosterone Suppress Proliferation and Induce Markers of Differentiation in Cultured Rat Sertoli Cells Endocrinology, September 1, 2003; 144(9): 3722 - 3731. [Abstract] [Full Text] [PDF]
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 K. D. Carpenter, K. Hayashi, and T. E. Spencer Ovarian Regulation of Endometrial Gland Morphogenesis and Activin-Follistatin System in the Neonatal Ovine Uterus Biol Reprod, September 1, 2003; 69(3): 851 - 860. [Abstract] [Full Text] [PDF]
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