| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
a Reproductive Endocrine Unit,
b Department of Medicine and In Vitro Fertilization Unit, Department of Obstetricsand Gynecology, Massachusetts General Hospital, Boston, Massachusetts 02114
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
, ßA, and ßB inhibin/activin subunits, FS, and the four activin receptor subtypes in individual human and mouse oocytes. The resulting expression pattern was further compared to that in human cumulus granulosa cells.
Our results indicate that neither ßA nor ßB mRNA was detectable in any human or mouse oocyte, that subunit was marginally present in some of the human oocytes, and that FS mRNA was detectable in human but not mouse oocytes. On the other hand, inhibin/activin subunit and FS mRNAs were abundantly expressed in cumulus cells. In addition, mRNAs for all four activin receptor subtypes (ActRIA, ActRIB, ActRIIA, and ActRIIB) were easily detectable in both oocytes and granulosa cells and appeared to be differentially expressed in oocytes during nuclear maturation. Finally, RNAs for both zona pellucida 3 and growth-differentiation factor-9, which were originally used as oocyte-specific markers, were detected in human but not mouse cumulus cells, although at lower levels than observed in oocytes. Taken together with previous studies, these results indicate that oocytes may be capable of responding to, but not synthesizing, activin.
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Inhibins and activins are members of the transforming growth factor ß (TGFß) superfamily of growth and differentiation factors. Inhibins are heterodimers containing distantly related and ß (ßA or ßB) subunits, while activins are ß-subunit homodimers [8]. Follistatin (FS) is a structurally unrelated monomeric protein with several alternatively spliced molecular forms (designated FS288 and FS315) that irreversibly bind activin and neutralize activin's biological effects [9, 10].
Like most of the TGFß superfamily, activin signals through two types of closely related transmembrane serine/threonine kinase receptors (designated type I and type II), each represented by two isoforms (ActRIA/ActRIB and ActRIIA/ActRIIB, respectively) [11, 12]. Since a specific inhibin receptor has not yet been identified, and inhibin can bind to the activin type II receptor, it is currently thought that inhibin acts as a naturally occurring antagonist of activin [13], although recent evidence suggests that inhibin may have a separate signal transduction pathway as well [14, 15].
Inhibin is a gonadal hormone that inhibits the synthesis and release of pituitary FSH [8], while both activin and inhibin regulate growth, proliferation, and differentiation through paracrine and/or autocrine actions in a variety of cell types including the gonads [2, 16, 17]. Both activin and inhibin modulate rat ovarian folliculogenesis in vivo [18], while in vitro, activin has been shown to modulate FSH receptor and aromatase induction, inhibin production, and proliferation in granulosa cells [2]. More recently, activin was found to influence follicular organization in vitro. When administered with FSH, activin was found to induce formation of follicular structures from monolayer cultures of immature rat follicles [19]. Interestingly, this induction required the presence of an oocyte, suggesting that activin, acting directly or indirectly, induces oocytes to produce a follicle-organizing factor [6]. In addition, in vitro studies have indicated a role for activin in promoting meiotic and cytoplasmic maturation of the oocyte [20–22] as well as influencing the outcome of in vitro maturation and fertilization of bovine oocytes [23, 24]. Taken together, these studies suggest that oocytes respond to activin in at least two ways: 1) by producing one or more factors that regulate the organization and maturation of developing follicles and 2) by modulating its own maturation. However, it is presently unknown whether oocytes produce activin directly or respond in a paracrine fashion to granulosa-derived activin under in vivo conditions.
To determine whether oocytes can produce inhibin, activin, or FS and whether they possess type I receptors necessary for responding to activin, we examined mRNA expression of inhibin/activin subunits, FS, and the type I and II activin receptor subtypes in oocytes. These results were compared to the mRNA expression pattern in cumulus granulosa cells. To accomplish this largely single-cell analytical procedure, a highly sensitive and specific reverse transcription-polymerase chain reaction (RT-PCR) protocol was developed. Our results indicate that neither human nor mouse oocytes contain detectable levels of activin subunit mRNA but that they express a full complement of activin receptor subtype mRNAs, consistent with the capability of responding to, but not synthesizing, activin.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Germinal vesicle (GV)-stage cumulus-oocyte complexes (COC) were obtained from women undergoing controlled ovarian hyperstimulation for in vitro fertilization. After suppression of endogenous gonadotropin with leuprolide acetate (Lupron; TAP Pharmaceuticals, Chicago, IL; 0.5 mg/day, s.c.), patients were treated with human menopausal gonadotropin (Pergonal; Serono, Randolph, MA) and/or human FSH (Metrodin; Serono) to stimulate follicular development. When the lead follicle was equal to or greater than 16 mm in diameter, hCG (Profasi; Serono, Randolph, MA) was administered, and follicle aspiration was performed 36 h later. GV-stage oocytes isolated from the follicle aspirates that were not required for patient treatment were transported to the laboratory and processed as described below. This protocol was approved by the Massachusetts General Hospital Subcommittee on Human Studies, and informed consent was obtained from all patients.
COC were either processed within 2–3 h of aspiration or cultured for up to 24 h in IVF50 medium (Scandinavian IVF Science, Vero Beach, Florida), under 5% CO2 at 37°C, until spontaneous maturation to metaphase 1 (M1) or metaphase 2 (M2). Before RNA extraction, oocytes were mechanically detached from the bulk of the cumulus mass and treated with hyaluronidase (200 IU/ml; Sigma), and all remaining granulosa cells were removed by repeat pipetting.
Mouse Oocytes and Embryos
Female B6C3F1 mice (22–43 days of age) were superovulated by administration of eCG (5 IU; Professional Compounding Centers of America, Houston, TX), followed by hCG (5 IU; Serono Laboratories, Norwell, MA) 48 h later. Mature (M2) oocytes were collected from the oviducts 16 h after hCG and denuded as described above. Fertilized eggs were obtained from females that were caged with males immediately after hCG induction. Fertilized eggs were cultured in IVF50 medium until the time of embryo hatching (3–5 days; preimplantation embryos). This protocol was approved by the Massachusetts General Hospital Subcommittee on Research Animal Care.
Total RNA Extraction
Individual denuded human oocytes or the detached cumulus mass was extracted with 500 µl of Trizol (GibcoBRL, Grand Island, NY) according to the manufacturer's protocol in the presence of 5 µg Escherichia coli tRNA as a carrier. Mouse oocytes or embryos were similarly treated except that they were pooled into groups of 1–5 before extraction.
RT-PCR
The entire RNA sample extracted from oocytes, or 25–100% of the extracted cumulus RNA, was treated with DNase I (1 U; GibcoBRL) to remove DNA contamination; this was followed by first strand cDNA synthesis in a 20-µl reaction containing 5 mM oligo(dT), 0.5 mM dNTP mix, and 100 U of SuperScript II (GibcoBRL). Complementary DNA-bound RNA was then removed by RNAse H (2 U; GibcoBRL) digestion at 37°C for 20 min. PCR was performed in a 25-µl reaction containing an RT aliquot equivalent to approximately 10% of the RT reaction, 0.2 mM dNTP mix, 0.25–0.5 µM upstream and downstream primers (each), 0.5 µCi [-32P]CTP (3000 Ci/mmol; New England Nuclear, Boston, MA), and 1.25 U Taq polymerase in single-strength PCR buffer A (both from Fisher Scientific, Pittsburgh, PA). After denaturation at 94°C for 3 min, templates were amplified for 22 cycles using a temperature profile of 94°C, 58°C, and 72°C for 30 sec each in GeneAmp PCR System 9600 (Perkin Elmer, Norwalk, CT). Five microliters of single-strength reaction mix containing 1.25 U Taq was then added, and PCR was continued for additional 25 cycles. Ten microliters of the PCR product was resolved by electrophoresis on 5% polyacrylamide gels in single-strength Tris-borate-EDTA buffer and then autoradiographed for 1 h at -80°C. Quantitation of gels was performed by beta-counting radioactive gel bands. For each PCR product, an exponential relationship was observed between increasing RNA concentration and PCR product intensity, although for many targets, the conditions used in these studies were on the plateau phase of the amplification curve to maximize sensitivity.
Most of the primers used in this study for human or mouse FS, , ßA, and ßB inhibin/activin subunits, the four activin receptor subtypes, and ß-actin were previously described and demonstrated to provide single, specific PCR bands whose identities were confirmed by Southern blotting or direct sequencing [25–27]. The sequences of primers not previously utilized are listed in Table 1, and their PCR products' identities were verified by subcloning (TA cloning kit; Invitrogen, San Diego, CA) and sequencing. A cDNA fragment of human growth-differentiation factor-9 (GDF9; [28]) was cloned and sequenced by amplifying human oocyte cDNA with the mouse primers. The human-specific primer set (Table 1) was positioned within this fragment. The FS primer set was designed to detect both alternatively spliced forms of FS; however, only FS315 was significantly amplified when both transcripts were present, although individual cDNAs of either form were successfully amplified. Thus, our results are reported as FS315.
|
All experiments included reactions in which the RT enzyme or cDNA template was omitted to eliminate the possibility of amplification from genomic DNA or contaminating template, and most primer sets crossed introns. In addition, no specific bands were detected in reactions containing the tRNA carrier alone. To confirm the integrity of the RNA templates and the RT-PCR protocol, ß-actin was examined in all samples. Furthermore, zona pellucida 3 (ZP3) [29] and GDF9 [28] were amplified as oocyte-specific markers, while the ovary-specific transcript of the human P450arom was used to ensure the absence of granulosa cell mRNA in the oocyte RNA extracts [30]. For transcripts that were negative in human oocytes, granulosa cell RNA was used as the positive control, while mouse ovary total RNA was used as a positive control sample for mouse oocytes. Mouse embryos were analyzed as a control for the sensitivity of the PCR for targets that were undetectable in oocytes.
Data Analysis
Background, as determined by counting the no-template control lane, was subtracted from each target signal, and the results were grouped as mean ± SEM, collectively or according to the maturation stage of the oocyte. Samples that produced a negative or weak ß-actin signal, or showed contamination in reverse transcriptase- or template-deleted controls, were excluded from analysis. Human oocyte samples that produced a strong P450arom signal were considered to include granulosa cell RNA, and data for these were deleted. Experiments in which mouse ovary RNA failed to produce positive signals were also excluded. Statistical differences between maturation groups were determined using Mann-Whitney U test. Differences were considered significant at p < 0.05. For comparison of human oocytes with cumulus cells and with mouse oocytes and embryos, the number of samples in which each target was detected by autoradiography was reported as a percentage of the total number of samples.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
-subunit mRNA were detected in only 11% of human oocytes. FS315 mRNA was detected in 95% of human oocytes, but the signal intensity was relatively low compared to that in cumulus cells. On the other hand, robust signals were detected for activin receptor subtype transcripts in nearly all oocytes (88–100%). As expected, high levels of ZP3 and GDF9 mRNAs characterized all oocytes. Cumulus cells, on the other hand, expressed all inhibin/activin system targets examined, including the three inhibin subunits, FS315, and all four activin receptors. In these cells the PCR product signal intensity among FS and the inhibin/activin subunits showed the profile Inh- > FS315 = ßA >> ßB, the latter being undetectable in 20% of the samples. Surprisingly, low levels of GDF9 and ZP3 mRNA, which were included as "oocyte-specific" markers, were observed in 40% and 80% (respectively) of the cumulus samples.
|
|
Expression of activin receptor subtype and FS315 mRNA was examined during oocyte meiotic maturation (Fig. 2). All mRNA species examined were detectable at all three nuclear maturational stages. However, the steady-state mRNA levels for ActRIIA, ActRIIB, and ActRIB appeared to vary during maturation. In particular, the ActRIIB PCR product signal intensity was lower (p < 0.05) in M1 oocytes compared to either GV or M2 oocytes, while for ActRIB, the PCR signal intensity was lower in M1 oocytes than in M2 oocytes (p < 0.05). For ActRIIA, the PCR product in GV oocytes appeared to be higher than that in either M1 or M2 oocytes (p = 0.05 and p < 0.05, respectively).
|
Like human oocytes, mouse oocytes did not express inhibin/activin subunits. However, in contrast to findings in human oocytes, FS315 transcripts were not detected in mouse oocytes (Table 2) despite their easy detection in mouse ovary RNA (data not shown). In contrast, 50–75% of hatching-stage blastocysts expressed the activin ßA and ßB subunits and FS315, but not -subunit mRNA. All four activin receptor subtype mRNAs were present in both oocytes and embryos, and their detection frequencies were comparable between mouse oocytes, blastocysts, and human oocytes. As expected, high levels of GDF9 and ZP3 were found in all mouse oocyte samples, but neither was found in embryos.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Inhibin/activin subunit, activin receptor, and FS mRNA levels were also investigated in mouse oocytes so that we could examine these undetectable targets at the greater RNA concentrations afforded by the more readily available mouse oocytes. Interestingly, the only difference between mouse and human oocytes in this analysis was the presence of FS mRNA in human but not mouse oocytes. The mouse primers were located identically to the human primers, and the FS gene is highly conserved between species, so this difference is not likely due to differences in PCR sensitivity. Moreover, FS mRNA was detected in mouse embryos as well as in adult ovaries, both of which served as positive controls. FS has been previously demonstrated to bind activin nearly irreversibly and to neutralize activin's bioactivity [9, 10]. Thus, if the FS mRNA in human oocytes is translated, this difference in FS expression might make human oocytes more resistant to the effects of activin relative to mouse oocytes, or limit the time over which activin can exert its signal.
Using a highly sensitive, single-oocyte RT-PCR approach, we were unable to detect inhibin/activin subunit mRNA in human or mouse oocytes. Since we meticulously removed all cumulus cells before oocyte extraction, our RT-PCR results should reflect transcripts of oocyte origin as opposed to surrounding tissue. This is critical, since we were able to detect evidence of granulosa cell contamination of oocyte mRNA when as few as 5 granulosa cells remained attached to the oocyte before extraction. To ensure that only oocyte mRNA was analyzed in our study, oocyte RNA samples were examined for the ovarian form of aromatase [30], and all samples were deleted in which significant aromatase was detected. Thus, differences between our results and those in previous reports of activin subunit mRNA in oocytes using RT-PCR may be accounted for by residual granulosa cells adhering to oocytes at the time of extraction, since the oocytes were not individually examined before extraction and ovarian control PCR reactions were not performed [31, 32]. Furthermore, our failure to detect inhibin/activin subunit mRNA in oocytes is consistent with previous in situ hybridization studies [33]. On the other hand, the immunohistochemical detection of inhibin/activin subunit proteins in both oocytes and COC [32, 33] might represent activin and/or inhibin from granulosa cells bound to cell surface receptors on oocytes or internalized after receptor binding. Taken together, these results support the concept that granulosa cell-derived activin acts as a paracrine signal in developing oocytes.
Although ActRIIA and ActRIIB mRNAs have been previously demonstrated in mouse and rat oocytes [34–36], we provide evidence for the expression of these receptors in human oocytes, as well as detection of both ActRIA and ActRIB in mouse and human oocytes. Since type I and II receptors are both necessary for activin signal transduction [11], these findings provide support for the ability of oocytes to respond to activin during the maturation process. The recent immunocytochemical identification of type II activin receptors on mouse oocytes further supports this concept [32].
In our experiments, the PCR product from activin receptors appeared to vary significantly across oocyte meiotic maturational stages. ActRIIB and ActRIB mRNA levels were significantly lower at M1 compared to either GV or M2. ActRIIA mRNA, on the other hand, was lower at M1 but remained reduced at M2. To examine these potentially quantitative relationships more closely, we repeated these experiments on a smaller group of human oocytes using a semiquantitative RT-PCR protocol with amplification in the exponential range for each target. We observed a similar receptor mRNA pattern across maturational stages, although it did not reach significance due to the limited number of available oocytes (data not shown). Initially, this increase in mRNA at M2 seemed paradoxical, since RNA transcription ends with oocyte nuclear maturation [37]. However, our RT procedure utilized oligo(dT) to prime cDNA synthesis and therefore presumably detected only polyadenylated transcripts. Moreover, ActRIIA mRNA has been previously demonstrated to undergo cytoplasmic polyadenylation during oocyte maturation and early embryonic development [38]. Therefore, one explanation for higher mRNA levels at M2 is the cytoplasmic polyadenylation of ActRIIB and ActRIB late in oocyte nuclear maturation, resulting in increased detection by our RT method. These observations suggest that an increase in mRNA for both receptor subtypes at M2 might help prepare the oocyte for detection of activin signals during early embryonic development.
In comparison to signals obtained from oocytes, we detected relatively low levels of both ZP3 and GDF9 mRNAs in human cumulus cells. While both ZP3 and GDF9 appear to be oocyte specific in mice [28, 39], several investigators have observed ZP3 mRNA and/or protein in granulosa cells from other species, including humans [40–42]. Thus, ZP3 mRNA detection in human granulosa-lutein cells is consistent with these earlier observations and demonstrates that human granulosa cells, like those from pigs and rabbits, can synthesize ZP3.
Using RT-PCR on RNA recovered from individual human oocytes, the results presented here provide some novel insights into biosynthesis of inhibin/activin subunits, FS, and activin receptors at the mRNA level. Our results suggest that oocytes could respond to an activin signal emanating from surrounding granulosa cells but that oocytes themselves do not make or secrete inhibin or activin. Taken together with the observation that treatment of immature rat follicles with activin and FSH stimulated oocytes to produce a follicle-organizing activity in vitro [6], our results suggest that granulosa cells modulate some aspects of oocyte function by secreting activin, which in turn regulates granulosa cell and follicular development and maturation. The biochemical nature of these activin-responsive signals therefore merits further investigation.
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
2 Correspondence: Yisrael Sidis, Reproductive Endocrine Unit, BHX-5, Massachusetts General Hospital, Boston, MA 02114. FAX: (617) 726–5357; ysidis{at}partners.org
Accepted: May 14, 1998.
Received: February 12, 1998.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
This article has been cited by other articles:
|
|
 |
|
  R. B. Gilchrist, M. Lane, and J. G. Thompson Oocyte-secreted factors: regulators of cumulus cell function and oocyte quality Hum. Reprod. Update, March 1, 2008; 14(2): 159 - 177. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. A. Dragovic, L. J. Ritter, S. J. Schulz, F. Amato, J. G. Thompson, D. T. Armstrong, and R. B. Gilchrist Oocyte-Secreted Factor Activation of SMAD 2/3 Signaling Enables Initiation of Mouse Cumulus Cell Expansion Biol Reprod, May 1, 2007; 76(5): 848 - 857. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Forges, P. Monnier-Barbarino, B. Leheup, and P. Jouvet Pathophysiology of impaired ovarian function in galactosaemia Hum. Reprod. Update, September 1, 2006; 12(5): 573 - 584. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. G Knight and C. Glister TGF-{beta} superfamily members and ovarian follicle development. Reproduction, August 1, 2006; 132(2): 191 - 206. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Mazerbourg and A. J.W. Hsueh Genomic analyses facilitate identification of receptors and signalling pathways for growth differentiation factor 9 and related orphan bone morphogenetic protein/growth differentiation factor ligands Hum. Reprod. Update, July 1, 2006; 12(4): 373 - 383. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Xia, Y. Sidis, A. Mukherjee, T. A. Samad, G. Brenner, C. J. Woolf, H. Y. Lin, and A. Schneyer Localization and Action of Dragon (Repulsive Guidance Molecule b), a Novel Bone Morphogenetic Protein Coreceptor, throughout the Reproductive Axis Endocrinology, August 1, 2005; 146(8): 3614 - 3621. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J.L. Juengel and K.P. McNatty The role of proteins of the transforming growth factor-{beta} superfamily in the intraovarian regulation of follicular development Hum. Reprod. Update, March 1, 2005; 11(2): 144 - 161. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J R V Silva, R van den Hurk, H T A van Tol, B A J Roelen, and J R Figueiredo Gene expression and protein localisation for activin-A, follistatin and activin receptors in goat ovaries J. Endocrinol., November 1, 2004; 183(2): 405 - 415. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T.-T. Wong and Y. Zohar Novel Expression of Gonadotropin Subunit Genes in Oocytes of the Gilthead Seabream (Sparus aurata) Endocrinology, November 1, 2004; 145(11): 5210 - 5220. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K P McNatty, L G Moore, N L Hudson, L D Quirke, S B Lawrence, K Reader, J P Hanrahan, P Smith, N P Groome, M Laitinen, et al. The oocyte and its role in regulating ovulation rate: a new paradigm in reproductive biology Reproduction, October 1, 2004; 128(4): 379 - 386. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Prochazka, L. Nemcova, E. Nagyova, and J. Kanka Expression of Growth Differentiation Factor 9 Messenger RNA in Porcine Growing and Preovulatory Ovarian Follicles Biol Reprod, October 1, 2004; 71(4): 1290 - 1295. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B A Refaat, A O Bahathiq, S Sockanathan, R L Stewart, M Wells, and W L Ledger Production and localization of activins and activin type IIA and IIB receptors by the human endosalpinx Reproduction, August 1, 2004; 128(2): 249 - 255. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 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]
|
|
|
|
|
|
Y. Wang and W. Ge Spatial Expression Patterns of Activin and Its Signaling System in the Zebrafish Ovarian Follicle: Evidence for Paracrine Action of Activin on the Oocytes Biol Reprod, December 1, 2003; 69(6): 1998 - 2006. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. Liu, T. Kuulasmaa, V.-M. Kosma, R. Butzow, T. Vanttinen, C. Hyden-Granskog, and R. Voutilainen Expression of Betaglycan, an Inhibin Coreceptor, in Normal Human Ovaries and Ovarian Sex Cord-Stromal Tumors and Its Regulation in Cultured Human Granulosa-Luteal Cells J. Clin. Endocrinol. Metab., October 1, 2003; 88(10): 5002 - 5008. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. M. Duffy Growth Differentiation Factor-9 Is Expressed by the Primate Follicle Throughout the Periovulatory Interval Biol Reprod, August 1, 2003; 69(2): 725 - 732. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. O. Bahathiq, R. L. Stewart, M. Wells, H. D. Moore, A. A. Pacey, and W. L. Ledger Production of Activins by the Human Endosalpinx J. Clin. Endocrinol. Metab., November 1, 2002; 87(11): 5283 - 5289. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. Welt, Y. Sidis, H. Keutmann, and A. Schneyer Activins, Inhibins, and Follistatins: From Endocrinology to Signaling. A Paradigm for the New Millennium Experimental Biology and Medicine, October 1, 2002; 227(9): 724 - 752. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. A. Pangas, A. W. Rademaker, D. A. Fishman, and T. K. Woodruff Localization of the Activin Signal Transduction Components in Normal Human Ovarian Follicles: Implications for Autocrine and Paracrine Signaling in the Ovary J. Clin. Endocrinol. Metab., June 1, 2002; 87(6): 2644 - 2657. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. E. Drummond, M. T. Le, J.-F. Ethier, M. Dyson, and J. K. Findlay Expression and Localization of Activin Receptors, Smads, and {beta}glycan to the Postnatal Rat Ovary Endocrinology, April 1, 2002; 143(4): 1423 - 1433. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y.-G. Chen, H. M. Lui, S.-L. Lin, J. M. Lee, and S.-Y. Ying Regulation of Cell Proliferation, Apoptosis, and Carcinogenesis by Activin Experimental Biology and Medicine, February 1, 2002; 227(2): 75 - 87. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Newton, Y. Wang, N.P. Groome, and P. Illingworth Inhibin and activin secretion during murine preantral follicle culture and following HCG stimulation Hum. Reprod., January 1, 2002; 17(1): 38 - 43. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Fujiwara, Y. Sidis, C. Welt, G. Lambert-Messerlian, J. Fox, A. Taylor, and A. Schneyer Dynamics of Inhibin Subunit and Follistatin mRNA during Development of Normal and Polycystic Ovary Syndrome Follicles J. Clin. Endocrinol. Metab., September 1, 2001; 86(9): 4206 - 4215. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Zhao, M. A.M. Taverne, G. C. van der Weijden, M. M. Bevers, and R. van den Hurk Effect of Activin A on In Vitro Development of Rat Preantral Follicles and Localization of Activin A and Activin Receptor II Biol Reprod, September 1, 2001; 65(3): 967 - 977. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Kishida and G. V. Callard Distinct Cytochrome P450 Aromatase Isoforms in Zebrafish (Danio rerio) Brain and Ovary Are Differentially Programmed and Estrogen Regulated during Early Development Endocrinology, February 1, 2001; 142(2): 740 - 750. [Abstract] [Full Text] [PDF]
|
|
|
|
