Synergy between Activin A and Gonadotropin-Releasing Hormone in Transcriptional Activation of the Rat Follicle-Stimulating Hormone-ß Gene

Susan J. Gregory, Charlemagne T. Lacza, Alissa A. Detz, Shuyun Xu, Laura A. Petrillo and Ursula B. Kaiser

Division of Endocrinology, Diabetes and Hypertension, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts 02115

Address all correspondence and requests for reprints to: Dr. Ursula B. Kaiser, Brigham and Women’s Hospital and Harvard Medical School, Division of Endocrinology, Diabetes and Hypertension, 221 Longwood Avenue, Boston, Massachusetts 02115. E-mail: ukaiser{at}partners.org.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Both activin and GnRH can independently stimulate expression of the FSHß subunit gene. In this study, we used the gonadotrope-derived LßT2 cell line to investigate the potential interaction between activin and GnRH in regulating the transcriptional activity of the rat FSHß gene promoter. Activin A and GnRH synergistically enhanced rat FSHß transcriptional activity. Overexpression of SMAD3 (mediator of decapentaplegic-related protein 3), but not of SMAD2, increased transcriptional activation of the rat (r) FSHß gene promoter, which was further enhanced by the combined overexpression of SMAD3 and 4 (3+4). The stimulatory effects of SMAD3 overexpression were localized to –472/–256 of the rFSHß gene promoter, and activin- and GnRH-responsive proteins were shown to bind to region –284/–252. Sequence analysis identified a consensus palindromic SMAD-binding site at –266/–259 of the rFSHß gene promoter. Mutation of two bases located in the center of this palindrome effectively abrogated SMAD4 binding, markedly reduced SMAD3 and 3+4 stimulation of the rFSHß gene promoter, and significantly decreased the synergistic enhancement of promoter activity by both activin A and GnRH, and SMAD3 and GnRH. Blockade of the MAPK-signaling pathway did not significantly affect the response to combined stimulation with activin and GnRH. In contrast, interference with SMAD3 signaling caused a significant reduction in activin and GnRH synergy. The results indicate that SMAD3 plays an important role in the synergistic effects of activin and GnRH and demonstrate that this synergy is mediated by a palindromic cis-element located at –266/–259 of the rFSHß gene promoter.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
ACTIVIN IS PRODUCED in multiple tissues throughout the body and has diverse effects both during development and in the adult. One of the major physiological functions of activin is the regulation of the reproductive axis through its ability to stimulate FSH synthesis and secretion. Activin is a known activator of FSHß gene expression, stimulating the transcriptional activity of the FSHß subunit gene (1) and stabilizing levels of FSHß mRNA (2, 3). A member of the TGFß superfamily, it is formed by the dimerization of ß-subunits (ßA or ßB) to form activin A (ßAßA), activin B (ßBßB), or activin AB (ßAßB) (4, 5, 6). No significant differences in the potency or ability to stimulate FSH release of the different activin subtypes have been detected (6). Both activin ßA- and ßB-subunits have been detected in the pituitary gland of prepubertal rats, and although expression of the activin ßB-subunit remained constant, expression of the activin ßA-subunit was significantly reduced in the adult (7). Previous studies in adult rats have shown activin ßB expression in gonadotrope cells (8, 9, 10), suggesting that activin B is produced by gonadotropes and may act in an autocrine/paracrine fashion to regulate FSH production and secretion (11).

Like other members of the TGFß superfamily, activin signaling is mediated by two subtypes of membrane-bound serine/threonine protein kinase receptors. The activin ligand binds initially to the activin type II receptor and subsequently induces dimerization with the activin type I receptor. The type I receptor is phosphorylated upon formation of this heteromeric complex and recruits members of the SMAD (mediator of decapentaplegic-related protein) transcription factor family (12, 13). SMAD2 and SMAD3 are the principal signal transduction molecules associated with activin signaling/action. They are characterized as receptor-regulated SMAD proteins (R-SMAD) because of their ability to interact with and be phosphorylated by the activin type I receptor. After phosphorylation, they associate with the common mediator SMAD4 and translocate to the nucleus as multimeric complexes. In the nucleus, the SMAD proteins regulate gene transcription (12, 14). Both SMAD3 and -4 proteins have been shown to bind to a palindromic SMAD-binding sequence (5'-GTCTAGAC-3') (15). The binding of SMAD3 to this palindromic element has been further characterized. An 11-amino acid ß-hairpin in the MH1 region of SMAD3 becomes embedded in the major groove of the DNA to contact the GTCT sequence (16). However, there is no evidence for direct binding of SMAD2 to this DNA sequence (15, 17). A unique 30-amino acid residue (exon 3) located just before the ß-hairpin in the MH1 domain of SMAD2 has been shown to interfere with DNA binding (16, 18). Despite this, SMAD2 is functionally active in many systems and interacts with transcriptional cofactors that can stimulate transcriptional activity and stabilize protein-DNA associations (19, 20, 21, 22, 23).

In addition to the stimulatory effects of activin on FSH, GnRH is also a potent stimulator of FSH synthesis and release (24, 25). GnRH is synthesized and secreted from specialized hypothalamic neurons in a pulsatile manner and stimulates the seven-transmembrane domain, G protein-coupled GnRH receptor (GnRHR) present on the cell surface of gonadotropes (25, 26). Activation of phospholipase C by the GnRHR generates the production of inositol 1,4,5-triphosphate (IP3) and diacylglycerol and results in mobilization of calcium and activation of protein kinase C (27). Downstream signaling events include the activation of MAPK cascades, including ERK, jun-N-terminal kinase, and p38MAPK (27). However, the specific mechanisms by which GnRH can stimulate FSHß gene transcription are not yet fully understood.

Interactions between activin and GnRH-signaling pathways have been demonstrated in {alpha}T3–1 and LßT2 gonadotrope-derived cell lines. GnRH-stimulated transcriptional activity of the mouse GnRHR gene promoter was significantly enhanced when {alpha}T3–1 cells were cotreated with activin and GnRH in combination (28, 29). Immunocytochemical studies identified that treatment of {alpha}T3–1 and LßT2 cells with activin induced the nuclear translocation of SMAD3. Interestingly, nuclear translocation of SMAD proteins could also be detected after treatment with GnRH (29), suggesting that GnRH-signaling pathways can also lead to SMAD activation.

Although both activin and GnRH alone have been found to stimulate FSH secretion, the effect of a potential interaction between these two key regulatory factors on FSHß gene transcription has not been examined. The present study was undertaken to investigate potential interactions between activin and GnRH in regulating transcriptional activity of the rat FSHß gene promoter and to elucidate the cis-elements and trans-factors mediating this response.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Activin A and GnRH Synergistically Enhance Transcriptional Activation of the Rat FSHß Gene Promoter
Unlike other gonadotrope-derived cell lines, LßT2 cells are able to synthesize and secrete FSH in response to both activin A and GnRH stimulation (30, 31). Due to these characteristics as well as their mature phenotype, LßT2 cells were used to examine the effect of potential interactions between activin A and GnRH on the rat (r)FSHß gene promoter. Transient transfections of –472/+15 rFSHßLuc or the empty vector pXP2 (used as a negative control) were performed, and the level of luciferase activity was measured as a direct indicator of transcriptional activity (Fig. 1Go). A pSV-ß-galactosidase reporter was cotransfected as an internal control to correct for transfection efficiency, and all values were expressed as luciferase/ß-galactosidase. Treatment with activin A alone induced a 3.9 ± 0.1-fold increase in –472/+15 rFSHßLuc activity, when compared with activity in the untreated group (P < 0.05). Similar results were observed when cells were treated with GnRH agonist and a 6.9 ± 1.0-fold increase in luciferase activity resulted (P < 0.01). Cotreatment with activin A and GnRH agonist further enhanced luciferase activity, by 22.0 ± 1.1-fold over activity in untreated cells (P < 0.01). No statistically significant effect of activin or GnRH agonist was detected in cells transfected with empty vector alone. The marked increase in luciferase activity in response to this cotreatment demonstrates a synergistic effect of activin and GnRH in the enhancement of transcriptional activation of the rFSHß gene promoter.



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Fig. 1. Activin A and GnRH Act Both Independently and Synergistically to Enhance Transcriptional Activation of the rFSHß Gene Promoter

LßT2 cells were cotransfected with –472/+15 rFSHßLuc or pXP2, and pSV-ß-galactosidase. Cells were assigned to one of four treatment groups: 1) no treatment; 2) activin A (100 ng/ml); 3) GnRH agonist (100 nM); or 4) activin A (100 ng/ml) + GnRH agonist (100 nM). Luciferase activity was normalized to ß-galactosidase activity. Results are shown as mean ± SEM of three independent experiments, each performed in triplicate. A statistically significant difference between the mean luciferase values for each treatment group is denoted by different letters (P < 0.01, except for the difference between a and b where P < 0.05).

 
SMAD3, But Not SMAD2, Stimulates Transcriptional Activation of the rFSHß Gene Promoter
Activin is known to be a potent stimulator of FSH synthesis and secretion both in vivo (32, 33) and in vitro (1, 31, 34). Activin signaling is mediated by the SMAD family of transcription factors. Typically, activin binds to its receptor at the cell surface and activates members of the SMAD family of proteins, which are phosphorylated and translocate to the nucleus in partnership with the common mediator SMAD4 to effect transcriptional activation of target genes (12, 35). To investigate whether members of the SMAD family could activate the rFSHß gene promoter in the gonadotrope-derived LßT2 cell line, we performed transient transfection studies using expression vectors encoding SMAD proteins (Fig. 2AGo). Cells were cotransfected with –2000/+698 rFSHßLuc, pSV-ß-galactosidase, and expression vectors for the SMAD proteins (SMAD2, -3, or -4), either alone or in combination with the common mediator SMAD4 (SMAD2+4, SMAD3+4). We investigated the role of SMAD2 and -3 proteins because they have been shown previously to be activin responsive in other systems. Overexpression of SMAD3 significantly stimulated transcriptional activation of the rFSHß gene promoter, by 35.3 ± 9.5-fold compared with the empty control vector (P < 0.05). Moreover, when SMAD3 and -4 were expressed in combination, there was a further increase in luciferase activity, by 80.1 ± 16.4-fold (P < 0.01). In contrast, overexpression of SMAD2, either alone or in combination with SMAD4, or SMAD4 alone, did not have a statistically significant effect on luciferase activity (SMAD2, 0.5 ± 0.1-fold; SMAD2+4, 6.0 ± 0.7-fold; SMAD4, 3.1 ± 0.4-fold; P > 0.05). To ensure that the transfection amounts of SMAD2 and SMAD3 were not limiting, we performed transient transfection studies with –472/+15 rFSHßLuc and progressively increasing amounts (0, 0.1, 0.5, 1, 2, and 4 µg) of the SMAD2 and SMAD3 expression vectors. In the case of SMAD3, a progressive increase in stimulation of –472/+15 rFSHßLuc activity was seen with increasing amounts of transfected plasmid (data not shown). A response was observed with as little as 0.5 µg of SMAD3 expression vector and was maximal with 2 µg of SMAD3 expression vector. In contrast, for SMAD2, no increase in –472/+15 rFSHßLuc activity was seen at any concentration of transfected SMAD2 plasmid, not even with 4 µg (data not shown). These results are consistent with previous data (36) and extend those studies by taking into consideration the effects of transfection efficiency by the expression and correction for pSV-ß-galactosidase activity.



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Fig. 2. Overexpression of SMAD3, Alone or in Combination with SMAD4, Increases Activation of the rFSHß Gene Promoter

A, LßT2 cells were cotransfected with –2000/+698 rFSHßLuc, pSV-ß-galactosidase, and the pCS2 expression vectors encoding SMAD2, -3, or -4 alone or in combination (SMAD2+4 or 3+4). Cotransfection with the empty pCS2 vector was used to normalize the total amount of DNA transfected in each group and as a negative control. Luciferase activity was normalized to ß-galactosidase activity. Results are shown as mean ± SEM of three independent experiments, each performed in triplicate. Fold stimulation was calculated relative to the luciferase activity in the group transfected with the empty pCS2 vector alone. *, Statistically significant difference between luciferase activity in cells transfected with SMAD3 compared with the empty vector pCS2 (P < 0.05). +, A further increase in activity in cells transfected with SMAD3+4 compared with SMAD3 alone (P < 0.05). B, Western blot analyses were performed using whole-cell lysates from LßT2 cells transfected with pCS2, SMAD2, -3, or -4 expression vectors. Membranes were blotted with SMAD3, SMAD2/3, and actin antibodies. Arrows indicate bands of the correct molecular weight for the respective proteins. *, Complex representing SMAD2.

 
In the absence of a functional effect of SMAD2 on the rFSHß gene promoter, we performed Western blot analysis to confirm that overexpression of the SMAD proteins could be detected after transfection of the corresponding SMAD expression vectors (Fig. 2BGo). An increase in the cellular content of SMAD3 protein in LßT2 cells transiently transfected with the SMAD3 expression vector was clearly detected after immunoblot with an antibody to SMAD3. Western blot analysis of whole-cell extract from LßT2 cells transiently transfected with the SMAD2 expression vector identified an increased expression of the higher molecular weight SMAD2 protein in these cells, when compared with cells transfected with empty (pCS2), SMAD3, or SMAD4 expression vectors. Actin was used as a control to ensure equal loading of each lane. These results demonstrate that SMAD3, but not SMAD2, is the key member of the SMAD family of transcription factors that can stimulate the activity of the rFSHß gene promoter in the gonadotrope-derived LßT2 cell line.

Identification of a SMAD3-Responsive Element in the rFSHß Gene Promoter
SMAD3 is the primary member of the TGFß/activin-signaling pathway capable of potently stimulating activation of the rFSHß gene promoter. The specific region of the rFSHß gene promoter that mediates SMAD3 responsiveness was identified by coexpressing 5'- and 3'-deletion constructs with the SMAD3 expression vector in transient transfection studies (Fig. 3Go). A robust response to SMAD3 overexpression was detected in cells transfected with –2000/+698 rFSHßLuc (50.1 ± 13.9-fold) and –472/+15 rFSHßLuc (87.6 ± 30.6-fold) (P < 0.05). This response was significantly reduced by further 5'-deletion, i.e. –256/+15 rFSHßLuc, –140/+15 rFSHßLuc and –50/+15 rFSHßLuc (6.8 ± 1.0-fold; 2.1 ± 0.6-fold, and 3.8 ± 0.9-fold, respectively). In fact, the mean luciferase activity derived from these three latter 5'-deletion constructs in the presence of overexpressed SMAD3 did not differ statistically from the activity in cells transfected with the empty pCS2 vector as control (P > 0.05). These data clearly demonstrate that the stimulatory effects of SMAD3 overexpression are localized between –472 and –256 of the rFSHß gene promoter.



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Fig. 3. –472/–256 of the rFSHß Gene Promoter Mediates SMAD3-Responsive Transcriptional Activation

LßT2 cells were cotransfected with 5'- and 3'-deletion constructs of rFSHßLuc (as indicated) or the empty vector pXP2, the SMAD3 expression vector or the empty control vector pCS2, and pSV-ß-galactosidase. Luciferase activity was normalized to ß-galactosidase activity. Results are shown as mean ± SEM of three independent experiments, each performed in triplicate. Fold stimulation by SMAD3, when compared with the empty vector pCS2, is shown to the right of the histogram for each construct. *, Statistically significant difference in the fold stimulation of luciferase activity by SMAD3 compared with the empty vector pXP2 (P < 0.05).

 
LßT2 Cell Nuclear Protein Binding to –284/–252 of the rFSHß Gene Promoter Is Increased after Stimulation with Activin A and GnRH
SMAD3 and -4 bind to specific DNA sequences of the mouse GnRHR gene (29, 37), early mesoendodermal response gene Mix.2 (38), and mouse goosecoid (21) promoters and are capable of recruiting transcriptional cofactors (21, 37, 38). SMAD proteins have been shown to bind to specific GC-rich motifs, and a palindromic SMAD-binding consensus element has been identified (15) that was able to bind human SMAD3 and -4 proteins in vitro. Sequence analysis of region –472 to –256 of the rFSHß gene promoter identified a putative homologous palindromic SMAD-binding site (GTCTAGAC) at –266/–259 (Fig. 4AGo).



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Fig. 4. Identification of a Consensus SMAD-Binding Site within –284 to –252 of the rFSHß Gene Promoter and Characterization of LßT2 Cell Nuclear Protein Binding

A, Sequence analysis revealed a putative palindromic consensus SMAD-binding site at –266/–259 of the rFSHß gene promoter. To further characterize whether nuclear proteins from the gonadotrope-derived LßT2 cells could bind to this site, an oligonucleotide spanning –284/–252 of the rFSHß gene promoter was generated (referred to as rFSHß-P). B, EMSA was performed using 32P-end-labeled rFSHß-P as probe and nuclear extracts from LßT2 cells that were treated with or without activin A for the indicated time periods. rFSHß-P in the absence of nuclear extract (NNE) was used as a negative control (lane 1). Four protein-DNA complexes were identified using nuclear extracts from LßT2 cells that had been treated with or without activin A (lanes 2–7, complexes a, b1, b2 and c). C, EMSA was performed using 32P-end-labeled rFSHß-P as probe and nuclear extracts from LßT2 cells treated with or without GnRH agonist, activin A, or activin A + GnRH agonist for the indicated time periods. In lane 1, rFSHß-P was incubated in the absence of nuclear extract (NNE). Four protein-DNA complexes were again identified using nuclear extracts from LßT2 cells (lanes 2–11, complexes a, b1, b2, and c). These complexes were similar to those detected in Fig. 4B; however, the gel was run over a shorter period of time, and the bands present in complex a are not as well resolved as those in Fig. 4C.

 
EMSAs were performed to determine whether this region of the rFSHß gene promoter could bind nuclear proteins extracted from LßT2 cells. An oligonucleotide probe corresponding to –284/–252 of the rFSHß gene promoter (rFSHß-P; Fig. 4AGo) was incubated with nuclear proteins extracted from LßT2 cells treated with activin A for 0, 1, 4, 12, 24, and 48 h (Fig. 4BGo). A lane in which nuclear extract was omitted was used as a negative control. Four protein-DNA complexes could be consistently identified (labeled a, b1, b2, and c). Complex a was variably resolved into a single complex (Fig. 4CGo) or into several closely migrating complexes (Fig. 4BGo). The gel in Fig. 4CGo was run over a shorter period of time, and the bands present in complex a are not as well resolved as those in Fig. 4BGo. Complexes b1 and b2 also migrated closely to each other (Fig. 4Go, B and C). The intensity of complexes a, b1, and b2 increased after treatment with activin A (Fig. 4BGo), reaching a maximal intensity at 12–24 h, suggesting that these bands contain activin-responsive proteins. A similar increase in the intensity of complex c was also detected after treatment with activin A, albeit to a lesser extent than the increases seen in complexes a, b1, and b2.

The functional interaction between activin A and GnRH that was seen in Fig. 1Go was further examined by extracting nuclear proteins from LßT2 cells treated with GnRH agonist for 0, 30 min, 1 h, 2 h, or 4 h with or without 24 h stimulation with activin A. Once again, four major protein-DNA complexes (a, b1, b2, and c, Fig. 4CGo) were detected by EMSA when rFSHß-P was used as probe. The intensity of complexes a, b1, and b2 were not affected after GnRH treatment. Complex c, however, progressively increased in intensity, reaching a peak after 4 h of GnRH treatment, suggesting that complex c may contain GnRH-responsive proteins. Nuclear proteins extracted from cells treated with both activin and GnRH resulted in a further increase in the intensity of the protein-DNA complexes. Bands a, b1, and b2 all increased in intensity after treatment with activin and GnRH, when compared with the intensity of protein-DNA complexes from cells treated with GnRH alone.

These results indicate binding of multiple nuclear proteins from LßT2 cells to region –284/–252 of the rFSHß gene promoter. Binding of nuclear proteins to this sequence was enhanced by treating LßT2 cells with activin A or GnRH alone. Moreover, protein-DNA binding was further amplified by the combined treatment with activin A and GnRH.

SMAD3 and -4 Bind to –284/–252 of the rFSHß Gene Promoter
In the previous experiment, nuclear proteins from LßT2 cells were shown to bind to –284/–252 of the rFSHß gene promoter containing a palindromic SMAD-binding site. However, the identity of the proteins was not established. To determine whether members of the SMAD family of proteins were present in the DNA-protein complexes, EMSA was carried out and supershift studies were performed using antibodies specific to SMAD2, -3, or -4 (Fig. 5AGo).



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Fig. 5. Identification of SMAD3 and 4 Binding to rFSHß-P by EMSA and Magnetic Separation Techniques

A, EMSA was performed using 32P-end-labeled rFSHß-P as probe and nuclear extracts from LßT2 cells treated with or without activin A for 12 h. Protein-DNA complexes a, b1, b2, and c were identified. A 500-fold excess of unlabeled rFSHß-P (lane 4; SI, specific inhibitor) or nonspecific LHSF oligonucleotide (lane 5; NS, nonspecific inhibitor) was added as a competitor. Binding reactions were coincubated with an antibody raised against an epitope common to both SMAD2 and -3 proteins (lane 6), with two different antibodies raised against the SMAD3 protein (lane 7: SC, Santa Cruz antibody; lane 8: ZY, Zymed antibody) or with an antibody to SMAD4 (lanes 9–11). SS indicates a supershifted complex formed in the presence of the SMAD4 antibody. *, Indicates the presence of SMAD4 peptide, included to preabsorb the SMAD4 antibody (lane 10). B, Magnetic separation of proteins binding to biotinylated rFSHß-P was performed using nuclear extracts from LßT2 cells that were either treated with activin A for 24 h (center panel) or transfected with the SMAD3 expression vector (right panel). A nonspecific biotinylated probe, corresponding to –120/–90 of the rFSHß gene promoter was used as a negative control (left panel). The arrow indicates the band corresponding to SMAD3.

 
Four protein-DNA complexes could again be identified when nuclear extracts from LßT2 cells treated with or without activin A were incubated with rFSHß-P as probe (lanes 2 and 3). This banding pattern was similar to that detected in previous EMSA experiments (Fig. 4Go, B and C). However, the intensity of all complexes was decreased in this experiment. To verify the specificity of protein-DNA binding, a 500-fold excess of unlabeled rFSHß-P was added to the DNA binding reaction to act as a specific competitor. Nuclear protein binding to the probe was completely abolished by the addition of unlabeled rFSHß-P (lane 4), whereas incubation with an unrelated nonspecific DNA oligonucleotide (LH steroidogenic factor-1 binding site, or LHSF) did not block protein-DNA interactions (lane 5). These competition data confirm that LßT2 cell nuclear proteins bind specifically to –284/–252 of the rFSHß gene promoter.

To establish which, if any, of the SMAD family of proteins were capable of binding to this region of the rFSHß gene promoter, we incubated antibodies to SMAD2/3, SMAD3, and SMAD4 with the protein-DNA binding mixture. An antibody specific for SMAD2/3 (lane 6) and two different antibodies raised against SMAD3 did not have any effect on the pattern of protein-DNA binding (lanes 7 and 8). In contrast, in the presence of the SMAD4 antibody, two clear supershifted complexes could be detected (lane 9), and the intensity of complex b2 was substantially reduced. Specificity of the antibody for the SMAD4 protein was ensured by preabsorbing the antiserum with the peptide to which it was raised, before addition to the protein-DNA binding mixture (lane 10). In addition, the antibody was also incubated with rFSHß-P in the absence of nuclear proteins (lane 11). No supershifted bands were detected in either of these controls, indicating that complex b2 is a specific SMAD4-DNA binding complex.

SMAD3 overexpression had a clear functional effect in transient transfection experiments, significantly stimulating the activity of the rFSHß gene promoter (Figs. 2AGo and 3Go). However, SMAD3 binding could not be identified in the supershift studies. Other studies using the same antibodies have also reported SMAD4 binding to consensus SMAD-binding sites. However, they were also unable to demonstrate a supershifted complex when the SMAD3 antibody was incubated with nuclear extract and the SMAD-binding element of the mouse GnRHR gene promoter as probe (37). This may be attributable to a poor ability of these antibodies to recognize the native form of the protein. To address this possibility, we incubated nuclear proteins from cells treated with activin A for 24 h with biotinylated rFSHß-P. Using magnetic separation techniques, the protein-DNA complexes were isolated and the proteins therein were run on a denaturing SDS-PAGE gel. Western blot analysis using the same SMAD3 antibody as in gel shift studies (Fig. 5AGo, lane 8) identified SMAD3 protein after magnetic separation of proteins bound to rFSHß-P (Fig. 5BGo; arrow indicates the SMAD3 protein band). A nonspecific biotinylated probe, corresponding to –120/–90 of the rFSHß gene promoter, was used as a negative control. No SMAD3 protein was isolated by the magnetic separation technique using the nonspecific –120/–90 as probe. Cell lysates from cells transfected with the SMAD3 expression vector were used as a positive control and confirmed SMAD3 overexpression in transfected cells (Fig. 5BGo).

The Palindromic SMAD-Binding Element at –266/–259 Binds SMAD4 and Mediates SMAD3 and SMAD3+4 Responsiveness of the rFSHß Gene Promoter
To further localize the SMAD-binding cis-element within the rFSHß gene promoter, SMAD4 binding to rFSHß-P was examined in greater detail. Sequential mutations of the 8-bp palindromic SMAD-binding sequence and the 5'- and 3'-flanking regions were generated in the rFSHß-P oligonucleotide (Fig. 6AGo). These mutants, designated M1–M6, were used as competitors in EMSA studies in which the SMAD4 complex was supershifted (Fig. 6BGo). In the presence of SMAD4 antibody, two supershifted complexes could again be clearly detected (Fig. 6BGo, lane 4). Formation of the supershifted complexes was prevented by the addition of excess unlabeled rFSHß-P (lane 5). Competition with excess M1 and M6 oligonucleotides also prevented SMAD4 binding to rFSHß-P, so that no supershifted bands were detected (lanes 6 and 11). However, mutations within the palindromic SMAD-binding site significantly impaired the ability of these mutants to bind to SMAD4; thus, supershifted bands were still detected even in the presence of 500-fold excess M2–M5 (lanes 7–10). SMAD4 binding was most effectively abrogated by M3, which contains a mutation of 2 bp located within the center of this palindrome (lane 8). These data demonstrate that the SMAD-binding site at –266/–259 of the rFSHß gene promoter can indeed bind SMAD4 and that 2-bp mutations within any part of this element significantly impair SMAD4 binding capability.



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Fig. 6. The Palindromic Consensus SMAD-Binding Site Is Functionally Important for the Transcriptional Activation of the rFSHß Gene Promoter

A, Sequential 2-bp mutations of the putative SMAD-binding site at –266/–259 of the rFSHß gene promoter were generated in the rFSHß-P oligonucleotide. Mutants were designated M1–M6. B, M1–M6 were used as competitors for SMAD4 binding in EMSA studies in which the SMAD4 complex was supershifted. Protein-DNA complexes a, b1, b2, and c are again indicated, as is the supershifted complex (SS) formed in the presence of SMAD4 antibody. A 500-fold excess of rFSHß-P (wild type) or M1–M6 was added as indicated (lanes 5–11). C, LßT2 cells were cotransfected with wild-type –472/+15 rFSHßLuc, –472/+15 M3 rFSHßLuc, or the pXP2 empty vector in conjunction with pCS2, SMAD3, or SMAD3+4 expression vectors. Luciferase activity was normalized to expression of pSV-ß-galactosidase, and measurements are expressed as fold stimulation, calculated for each luciferase reporter as the fold increase in promoter activity in the presence of SMAD3 ± SMAD4 when compared with the corresponding control group transfected with empty pCS2 vector. Results are mean ± SE from multiple experiments. *, A statistically significant difference in the fold stimulation of promoter activity compared with the empty vector pXP2 (P < 0.05).

 
To address the functional significance of the palindromic SMAD-binding element, site-directed mutagenesis was performed, and a 2-bp sequence mutation corresponding to M3 (Fig. 6AGo) was introduced into –472/+15 rFSHßLuc (generating –472/+15 M3 rFSHßLuc). LßT2 cells were cotransfected with either wild-type –472/+15 rFSHßLuc or –472/+15 M3 rFSHßLuc along with SMAD3 ± SMAD4 expression vectors. Luciferase activity was normalized to expression of pSV-ß-galactosidase, and the fold increase in promoter activity for each luciferase reporter in the presence of SMAD3 ± SMAD4 above that in the control group transfected with empty pCS2 vector was calculated (Fig. 6CGo). Transcriptional activity of –472/+15 rFSHßLuc was significantly increased when SMAD3 was overexpressed, either alone or in combination with SMAD4 (SMAD3, 63.5 ± 18.4-fold; SMAD3+4, 202.0 ± 91.4-fold; P < 0.05). In contrast, the stimulatory effects of SMAD3 or SMAD3+4 overexpression were significantly reduced in –472/+15 M3 rFSHßLuc (SMAD3, 10.3 ± 7.2; SMAD3+4, 20.1 ± 9.3; Fig. 6CGo). Indeed, no statistically significant difference was detected between –472/+15 M3 rFSHßLuc and the empty vector pXP2 in the presence of overexpressed SMAD3 or SMAD3+4. These data demonstrate that mutation of two nucleotides within the center of the SMAD-binding site, which prevents SMAD4 binding, markedly reduces stimulation of the rFSHß gene promoter by SMAD3 and 3+4.

Mutation of the SMAD-Binding Element at –266/–259 Prevents the Synergistic Stimulation of the rFSHß Gene Promoter by Activin A and GnRH, and by SMAD3 and GnRH
Mutation of the SMAD-binding element in –472/+15 rFSHßLuc significantly reduced the ability of SMAD3, alone or in combination with SMAD4, to transcriptionally activate the rFSHß gene promoter. To determine whether this mutation was also able to prevent the synergistic stimulation of the rFSHß gene promoter by activin A and GnRH, we transfected LßT2 cells with the wild-type –472/+15 rFSHßLuc, mutant –472/+15 M3 rFSHßLuc, or pXP2 (Fig. 7AGo). After transfection, cells were treated with activin A, GnRH agonist, or both. Luciferase activity was normalized to expression of pSV-ß-galactosidase, and the fold increase in promoter activity above that in the untreated control group was calculated for each luciferase reporter relative to pXP2. Activin A treatment induced a modest activation of luciferase activity for the wild-type –472/+15 rFSHßLuc (1.4 ± 0.1-fold) that did not reach statistical significance in this study. GnRH significantly stimulated luciferase activity by 3.2 ± 0.4-fold. When cells were stimulated with activin A and GnRH agonist in combination, the fold induction in luciferase activity was further enhanced (5.5 ± 1.0-fold; P < 0.01, when compared with activin A and GnRH agonist treatments alone). Mutation of the SMAD-binding element prevented the synergistic stimulation of the rFSHß gene promoter by the combined treatments of activin A and GnRH agonist. Cells transfected with the mutant –472/+15 M3 rFSHßLuc demonstrated a smaller, but not significantly different, increase in luciferase activity after activin A treatment (1.2 ± 0.1-fold). Stimulation with GnRH agonist induced a 2.5 ± 0.5-fold increase in transcriptional activity, again smaller but not significantly different from the response of wild-type –472/+15 rFSHßLuc. However, in contrast to the results from LßT2 cells transfected with wild-type –472/+15 rFSHßLuc, the combined treatment with activin A and GnRH agonist did not enhance transcriptional activity of the mutant construct above that detected in cells treated with GnRH alone (activin A + GnRH, 2.8 ± 0.5-fold; GnRH, 2.5-fold ± 0.5; P > 0.05). These data demonstrate that SMAD binding is necessary for the synergistic stimulation of the rFSHß gene promoter by activin A and GnRH.



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Fig. 7. The Palindromic SMAD-Binding Site Is Functionally Important for the Synergistic Stimulation of the rFSHß Gene Promoter by Activin A and GnRH Agonist, and by SMAD3 and GnRH Agonist

A, LßT2 cells were cotransfected with wild-type –472/+15 rFSHßLuc, –472/+15 M3 rFSHßLuc or pXP2, and pSV-ß-galactosidase. Cells were assigned to one of four treatment groups: 1) no treatment; 2) activin A (100 ng/ml); 3) GnRH agonist (100 nM); or 4) activin A (100 ng/ml) + GnRH agonist (100 nM). Luciferase activity was normalized to expression of pSV-ß-galactosidase, and measurements are expressed as fold stimulation when compared with the untreated group. All values were normalized to pXP2. Results are shown as mean ± SEM from multiple independent experiments as indicated. Statistically significant differences between fold stimulation values for treatment groups are denoted by different letters (P < 0.05). B, LßT2 cells were transfected with wild-type –472/+15 rFSHßLuc, –472/+15 M3 rFSHßLuc, and pSV-ß-galactosidase in conjunction with pCS2 or SMAD3 expression vectors. Cells were assigned to one of two treatment groups: 1) no treatment, or 2) 4 h GnRH agonist (100 nM). Luciferase activity was normalized to expression of pSV-ß-galactosidase, and measurements are expressed as fold stimulation when compared with the untreated group. Results are shown as mean ± SEM from four experiments. Statistically significant differences between treatment groups are denoted by different letters (P < 0.05).

 
To further investigate the involvement of activin-signaling pathways in the synergistic stimulation of the rFSHß gene promoter by activin and GnRH, we examined whether SMAD3 and GnRH could act in a similar synergistic manner to enhance transcriptional activity of this promoter. We transfected LßT2 cells with the wild-type –472/+15 rFSHßLuc, mutant –472/+15 M3 rFSHßLuc, or pXP2 in combination with the pCS2 or SMAD3 expression vectors (Fig. 7BGo). After transfection, cells were either left untreated, or treated with GnRH agonist. Cotransfection of the wild-type –472/+15 rFSHßLuc and the SMAD3 expression vector induced a significant activation of luciferase activity, when compared with that of the control group (98.8 ± 7.3-fold; P < 0.01). GnRH stimulated luciferase activity by 6.6 ± 2.3-fold (P < 0.05) compared with the untreated group. When cells transfected with wild-type –472/+15 rFSHßLuc and the SMAD3 expression vector were stimulated with GnRH agonist, the fold induction in luciferase activity was further enhanced (263.9 ± 33.0-fold; P < 0.01) compared with transcriptional activation by SMAD3 or GnRH agonist alone. Mutation of the SMAD-binding element significantly reduced the synergistic stimulation of the rFSHß gene promoter by SMAD3 and GnRH agonist. Cells transfected with the mutant –472/+15 M3 rFSHßLuc demonstrated a significant reduction in luciferase activity after SMAD3 overexpression, when compared with SMAD3 stimulation of the wild-type –472/+15 rFSHßLuc (16.0 ± 2.4-fold vs. 98.8 ± 7.3-fold, respectively; P < 0.01). Stimulation with GnRH agonist induced a 5.0 ± 1.3-fold increase in transcriptional activity, which was not significantly different from the response of wild-type –472/+15 rFSHßLuc to GnRH. A significant reduction in transcriptional activity was observed when cells transfected with the mutant –472/+15 M3 rFSHßLuc in combination with the SMAD3 expression vector and treated with GnRH agonist were compared with the wild-type –472/+15 rFSHßLuc (SMAD3 + GnRH, 57.6 ± 15.1-fold vs. 263.9 ± 33.0-fold, respectively; P < 0.01). Taken together, these data demonstrate that SMAD3 and GnRH synergistically stimulate transcriptional activity of the rFSHß gene promoter. Furthermore, mutation of the SMAD-binding element of this promoter significantly reduced stimulation by SMAD3 as well as synergy between SMAD3 and GnRH. These data indicate that SMAD binding plays an important role in the synergistic activation of the rFSHß gene promoter by activin A/SMAD3 and GnRH.

Role of the MAPK Pathway in the Synergistic Stimulation of the rFSHß Gene Promoter by Activin A and GnRH
Mutation of the SMAD-binding element of –472/+15 rFSHßLuc significantly reduced synergistic activation of the rFSHß gene promoter by activin A/SMAD3 and GnRH (Fig. 7Go, A and B). However, in both cases, transcriptional activation was not completely abolished. In particular, although markedly reduced, some synergy between SMAD3 and GnRH persisted. To investigate the potential involvement of GnRH-stimulated intracellular signaling pathways in this synergistic stimulation, we transiently transfected LßT2 cells with –472/+15 rFSHßLuc or pXP2 and pSV-ß-galactosidase (Fig. 8Go). Cells were treated 23 h after transfection with a MEK (MAPK kinase 1/2) inhibitor, U0126, or dimethylsulfoxide (DMSO, the resuspension agent). Cells were then treated with activin A, GnRH agonist, or both. Luciferase activity was normalized to expression of pSV-ß-galactosidase, and the fold increase in promoter activity above that in the untreated control group was calculated for each luciferase reporter. Cells treated with the MEK inhibitor did not demonstrate any change in the response to activin A (activin A + DMSO, 3.2 ± 0.4-fold; activin A + MEK inhibitor, 2.6 ± 0.4-fold, respectively; P > 0.05). However, not unexpectedly, addition of the MEK inhibitor significantly reduced the stimulation of the rFSHß gene promoter by GnRH agonist compared with cells that were not treated with MEK inhibitor (2.7 ± 0.3-fold vs. 5.3 ± 0.3-fold; P < 0.01). The response to combined treatment of activin A and GnRH agonist was also reduced in the presence of the MEK inhibitor (9.5 ± 2.2-fold vs. 15.0 ± 2.9-fold), although this decrease did not reach statistical significance.



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Fig. 8. Effects of Inhibition of MAPK Activity on Stimulation of the rFSHß Gene Promoter by GnRH Agonist and Activin

LßT2 cells were cotransfected with wild-type –472/+15 rFSHßLuc or pXP2 and pSV-ß-galactosidase. Cells were treated with 10 µM of a MEK inhibitor (UO126) or with vehicle only (DMSO). Within these two groups, cells were then assigned to one of the following treatment groups: 1) no treatment; 2) activin A (100 ng/ml); 3) GnRH agonist (100 nM); or 4) activin A (100 ng/ml) + GnRH agonist (100 nM). Luciferase activity was normalized to expression of pSV-ß-galactosidase, and measurements are expressed as fold stimulation when compared with the untreated group. Results are shown as mean ± SEM from three experiments. Statistically significant differences between treatment groups are denoted by different letters (P < 0.05).

 
These data demonstrate that the MAPK signaling pathway plays a role in GnRH stimulation of the rFSHß gene promoter. The MAPK signaling pathway may also contribute to the synergistic stimulation of the rFSHß gene promoter by activin and GnRH, although it does not appear to be the principal transcriptional regulator mediating this response.

Role of SMAD3 in the Synergistic Stimulation of the rFSHß Gene Promoter by Activin A and GnRH
To further investigate the role of SMAD3 in activin-induced stimulation of the rFSHß gene promoter and in the synergistic stimulation of this promoter by activin A and GnRH, we transiently transfected LßT2 cells with –472/+15 rFSHßLuc and pSV-ß-galactosidase, in combination with a dominant-negative SMAD3 expression vector or the corresponding empty expression vector pRK5, used as a control (Fig. 9Go). The dominant-negative SMAD3 protein is truncated at its C terminus and encodes amino acids 1–381 of the SMAD3 protein (39). Cells were treated with activin A, GnRH agonist, or both. In cells transfected with the wild-type –472/+15 rFSHßLuc and the empty expression vector pRK5, activin A treatment induced a significant activation of luciferase activity, above that of the untreated control group (2.3 ± 0.2-fold; P < 0.01). A similar increase in transcriptional activity was also detected after GnRH treatment (2.1 ± 0.3-fold; P < 0.01). When cells transfected with wild-type –472/+15 rFSHßLuc and pRK5 were stimulated with activin A and GnRH agonist in combination, the fold induction in luciferase activity was further enhanced, compared with the untreated group (5.1 ± 0.5-fold; P < 0.01), with activin alone (P < 0.01), or with GnRH alone (P < 0.01).



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Fig. 9. A Dominant-Negative SMAD3 Interferes with Synergistic Stimulation of the rFSHß Gene Promoter by Activin A and GnRH Agonist

LßT2 cells were cotransfected with wild-type –472/+15 rFSHßLuc and pSV-ß-galactosidase and a dominant-negative SMAD3 expression vector or the corresponding pRK5 empty expression vector. Cells were assigned to one of four treatment groups: 1) no treatment; 2) activin A (100 ng/ml); 3) GnRH agonist (100 nM); or 4) activin A (100 ng/ml) + GnRH agonist (100 nM). Luciferase activity was normalized to expression of pSV-ß-galactosidase. This experiment was repeated multiple times, and results are shown as mean ± SEM from a representative experiment. Statistically significant differences between treatment groups are denoted by different letters (P < 0.05).

 
Cells transfected with the wild-type –472/+15 rFSHßLuc and the dominant-negative SMAD3 expression vector demonstrated a modest reduction in basal activity of the FSHß gene promoter in cells that received no treatment, when compared with cells transfected with the pRK5 empty vector. However, this decrease did not reach statistical significance. Cells transfected with the wild-type –472/+15 rFSHßLuc and the dominant-negative SMAD3 expression vector, and treated with activin A or GnRH alone, demonstrated a statistically significant reduction in promoter activity when compared with cells transfected with the pRK5 empty vector (P < 0.05). In both these groups, transcriptional activity was not significantly different from the baseline levels in untreated cells transfected with the empty vector, pRK5. A similar decrease in transcriptional activity was also detected after treatment with activin and GnRH combined, with a significant reduction in transcriptional activity when compared with cells transfected with the empty vector (P < 0.01). Although some synergy was still present between activin and GnRH, the overall magnitude of the response in the presence of the dominant-negative SMAD3 was not significantly different from the transcriptional activity induced by activin or GnRH alone, in cells transfected with the empty vector.

These data again suggest that SMAD3 plays a role in both activin- and GnRH-induced activation of the rFSHß gene promoter. Furthermore, inhibition of SMAD3 signaling with a dominant-negative SMAD3 significantly reduced the ability of activin and GnRH to synergistically stimulate the rFSHß gene promoter.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
This study demonstrates that activin A and GnRH act synergistically to enhance transcriptional activity of the rFSHß gene promoter. The murine gonadotrope-derived LßT2 cell line was used in this study to investigate the interaction between activin and GnRH in regulating transcriptional activity of the rFSHß gene promoter. This cell line has previously been reported to express the FSHß subunit and to synthesize and release FSH in response to both activin and GnRH (30, 31). Moreover, the LßT2 cell line endogenously expresses activin receptors and the SMAD family of transcription factors (31, 40).

We have demonstrated that overexpression of SMAD3, but not of SMAD2, can robustly stimulate FSHß transcription. SMAD2 contains an extra 30 amino acid residues located just before the ß-hairpin loop in the MH1 domain; this region is not present in the SMAD3 protein and has been shown to impair the ability of SMAD2 to bind to DNA and, in certain systems, prevents SMAD2 transactivation of specific gene promoters (17, 18). Although previous studies have shown that SMAD2 can be phosphorylated by activin stimulation in LßT2 cells (40, 41), direct interactions of the SMAD2 protein and the FSHß gene promoter have not been reported.

Mapping studies localized SMAD3 activation of the rFSHß gene to region –472/–256. Sequence analysis identified a consensus SMAD-binding site at position –266/–259. This site is identical to an 8-bp palindromic sequence identified in the vestigial promoter that has been shown to bind human SMAD3 and 4 and to confer TGFß responsiveness to a minimal promoter (15). Four protein-DNA complexes were identified to bind to a region of the rFSHß gene promoter containing this palindromic SMAD-binding sequence (Fig. 4Go, B and C). An increase in the intensity of the protein-DNA complexes after activin treatment suggests that proteins contained within these complexes are responsive to activin. The intensity of these complexes reached a peak after 12–24 h of activin treatment, consistent with the functional data in which treatment with activin A stimulated a maximal induction in FSHß gene transcription at 24 h (data not shown). Previous studies have also shown a peak in FSHß mRNA transcripts 12–24 h after activin stimulation in LßT2 cells (40). Interestingly, we also detected a further increase in the intensity of the protein-DNA complexes after cotreatment with activin A and GnRH agonist. This increase in protein-DNA binding indicates that cross-talk between the activin and GnRH-signaling pathways acts to increase protein binding to region –284/–252 of the rFSHß gene promoter. Indeed, GnRH has been previously shown to stimulate activin-signaling pathways in the gonadotrope and to induce nuclear translocation of the SMAD proteins (29). In combination, these data suggest that SMAD proteins are likely part of both the activin- and GnRH-responsive complexes.

Both SMAD3 and -4 were shown to bind to –284/–252 of the rFSHß gene promoter. Mutation of the palindromic SMAD-binding element located within this region confirmed the functional importance of this element in both SMAD3 and 3+4 responsiveness and in the synergistic activation of the rFSHß gene promoter by activin A and GnRH, and by SMAD3 and GnRH. Although both Figs. 1Go and 7AGo show a clear synergistic enhancement in transcriptional activity after treatment with activin and GnRH, the overall magnitude of all responses was reduced in Fig. 7AGo. This may be due to the increased passage number of LßT2 cells used in Fig. 7AGo, which has been previously observed to decrease the responsiveness of the cells (36). Our data are consistent with this previous study, in which the same SMAD-binding element was identified and shown to be important for SMAD3- and activin-mediated FSHß gene transcription (36). In our study, mutation of this SMAD-binding element also prevented the synergistic enhancement in rFSHßLuc activity after treatment with activin A and GnRH agonist or SMAD3 and GnRH agonist, demonstrating the importance of SMAD binding for this synergistic response.

A similar synergism between activin and GnRH was demonstrated previously in the {alpha}T3–1 gonadotrope-derived cells (29). When these cells were transfected with the mouse GnRHR gene promoter, activin A augmented the GnRH-mediated transcriptional activation of this promoter, an effect that could be abrogated by the mutation of a consensus SMAD-binding element (29). The role of activin as an important mediator of GnRH responsiveness has also been demonstrated in studies in which follistatin, a protein that can neutralize the effects of activin, was shown to inhibit GnRH responsiveness of the ovine FSHß gene promoter (31). Treatment of primary pituitary cell cultures from transgenic mice expressing the ovine FSHß subunit gene promoter with activin and GnRH also resulted in an augmented activin response (42). This was corroborated in vivo, where the combined administration of activin A and GnRH agonist to female rats enhanced both FSH release and the levels of FSHß subunit mRNA when compared with rats treated with either GnRH or activin alone (43). In our study, an intact SMAD-binding element was essential for the synergistic effects of both activin and GnRH, and SMAD3 and GnRH, suggesting that cross-talk between these signaling pathways is dependent upon SMAD binding.

There are several possible mechanisms by which activin and GnRH may interact. First, activin can act through SMAD-independent pathways. Activin has been shown to activate both p38/MAPK and jun-N-terminal kinase/MAPK (44, 45) and to up-regulate GnRHR expression (28), which may act to augment GnRH responsiveness. Second, GnRH has been shown to stimulate translocation of the SMAD3 protein in LßT2 cells (29). For translocation to occur, the SMAD proteins must be phosphorylated at the conserved carboxy-terminal domain to relieve the autoinhibition by their amino-terminal domains, which enables SMAD4 association, nuclear translocation, and signaling (46). In the case of activin signaling, the activin type I receptor mediates phosphorylation of the SMAD proteins at the carboxy-terminal SSXS motif. However, there is evidence to suggest that ERK/MAPK and p38/MAPK-signaling pathways can induce phosphorylation of SMAD3 at phosphorylation sites in the middle linker region, which can also promote nuclear translocation of SMAD3 and -4 (47, 48). Because GnRH is known to activate ERK and p38, it is possible that SMAD proteins may become phosphorylated in response to GnRH stimulation through this mechanism, thereby inducing the subsequent nuclear translocation of these proteins and enabling them to activate the FSHß gene promoter. We further investigated whether signaling cross-talk between the MAPK family of proteins and activin played a significant role in activin and GnRH synergistic stimulation of the FSHß gene promoter using a MEK inhibitor (Fig. 8Go). As previously shown, a significant decrease in GnRH-stimulated activity of the rFSHß gene promoter was detected when MAPK signaling was blocked. A similar, but not significant, decrease in activin and GnRH synergism was also detected when the MAPK pathway was inhibited. This reinforces the current understanding that the MAPK signaling pathway is involved in GnRH stimulation of the FSHß gene promoter. Furthermore, it suggests that cross-talk between the MAPK signaling pathway and activin signaling pathways (e.g. SMAD3) may play a role in activin and GnRH synergism. However, because the response to activin and GnRH was not significantly reduced, or completely prevented, this suggests that there are other mechanisms by which activin and GnRH can interact.

Third, it is possible that GnRH can stimulate cofactors that may interact with the SMAD proteins to enhance FSHß gene transcription. SMAD proteins are known to associate with many different cofactors, which can either enhance (36, 37, 49) or repress (50, 51) SMAD-stimulated transcriptional activity. SMAD proteins have been shown to recruit general coactivators such as p300 and cAMP reponse element binding protein (CREB)-binding protein that do not bind to the DNA but interact with the MH2 domain of the SMAD protein to increase transcription of target genes. Furthermore, interactions with DNA-binding proteins such as FAST-1 that lack a transactivation domain but enhance binding stability, and transcription factors such as AP-1 that can bind to DNA and activate transcription on their own have been demonstrated (22). A pituitary-specific Pitx2 isoform can also bind to a region 29–60 bp downstream of the SMAD-binding element in the rFSHß gene promoter and has been shown to play a role in activin-mediated FSHß transcriptional activity, suggesting that it may be an important coregulator of activin signaling (36). In addition, homeodomain proteins Pbx1 and Prep1 have been shown to bind to the ovine FSHß gene promoter in association with SMAD2, -3, and -4 proteins (52). Thus, it is possible that additional proteins also bind to sequences surrounding the SMAD-binding element. These may not only stabilize SMAD binding but also mediate activin and GnRH responses. Therefore, in the absence of SMAD binding, GnRH-responsive stimulatory cofactors may be unable to activate or enhance transcriptional activity. EMSA provided evidence of a GnRH-responsive complex that bound to –284/–252 of the rFSHß gene. It is possible that these GnRH-responsive proteins are important cofactors that contribute to the synergistic interaction between activin and GnRH. In light of the incomplete reduction in activin and GnRH synergy after blockade of the MAPK-signaling pathway, the interaction of GnRH-responsive coactivators with SMAD3 seems a likely mechanism of action for activin and GnRH synergy. Indeed, when the SMAD3 signaling pathway was blocked using a dominant-negative SMAD3 protein, activin and GnRH synergy was significantly decreased. Transcriptional activation of the rFSHß gene promoter by activin and GnRH alone was also reduced when SMAD3 signaling was blocked, suggesting that SMAD3 is important not only for activin signaling, but that it may also play a role in GnRH signaling. This is consistent with the previous demonstration that GnRH can induce nuclear translocation of the SMAD proteins in gonadotrope-derived cell lines (29).

In summary, we have demonstrated that activin and GnRH act in a synergistic manner to regulate the transcriptional activity of the rFSHß gene promoter in the gonadotrope-derived LßT2 cell line. We report that SMAD3, but not SMAD2, can stimulate the activity of the rFSHß gene promoter. Furthermore, we have localized the stimulatory effects of SMAD3 to region –472/–256 and shown that a consensus palindromic SMAD-binding site located at –266/–259 of the rFSHß gene promoter specifically binds SMAD proteins and is necessary for both SMAD3- and 3+4-induced transcriptional activity as well as synergy between activin and GnRH in activation of the rFSHß gene promoter. Our results also indicate that SMAD3 plays a central role in mediating these synergistic effects. These data provide important evidence that cross-talk between the activin and GnRH signaling pathways can enhance rFSHß expression at a transcriptional level.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Des-Gly10,[D-Ala6]-LHRH-ethylamide (GnRH agonist) was obtained from Sigma Chemical Co. (St. Louis, MO). Recombinant human activin A was obtained from R&D Systems (Minneapolis, MN). A MEK inhibitor (UO126) was purchased from Promega Corp. (Madison, WI). A polyclonal SMAD 2/3 antibody was generated in goats, mapping at the carboxy terminus of SMAD3 of human origin. A polyclonal SMAD3 antibody was generated in goats to a peptide mapping within the amino-terminal domain of SMAD3 of human origin. A polyclonal SMAD4 antibody was generated in goats to a peptide mapping at the carboxy terminus of SMAD4 of human origin. All antibodies react with mouse, rat, and human SMAD proteins and were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). A blocking peptide that reacts with the goat antibody to SMAD4 was also purchased from Santa Cruz. A polyclonal SMAD2/3 antibody generated in rabbits to amino acids 186–273 of the human SMAD2 was purchased from Upstate (Waltham, MA). A polyclonal SMAD3 antibody generated in rabbits to a 20-amino acid synthetic peptide derived from a central portion of the linker domain of SMAD3 was purchased from Zymed Laboratories, Inc. (South San Francisco, CA). A horseradish peroxidase-conjugated mouse monoclonal IgG1 actin (C2) antibody generated to an amino acid sequence mapping to the carboxy terminus of human actin was purchased from Santa Cruz. All oligonucleotides were prepared by Life Technologies (Gaithersburg, MD). The LßT2 cells were generously donated by Dr. Pamela Mellon (University of California, San Diego, CA).

Reporter Plasmids and Expression Vectors
rFSHßLuc constructs contain specific segments (as indicated) of the rFSHß gene fused upstream of the luciferase reporter gene in pXP2 (53, 54). –2000/+698 rFSHßLuc, –472/+15 rFSHßLuc, –256/+15 rFSHßLuc, –140/+15 rFSHßLuc, and –50/+15 rFSHßLuc have been described previously (53, 55). A –472/+15 M3 rFSHßLuc mutant construct was generated using the QuikChange Site-Directed Mutagenesis kit (Stratagene, La Jolla, CA; AG replacement of CT at –264/–263 in the palindromic SMAD-binding element). The sense and antisense oligonucleotide primers used to introduce the mutations correspond to the M3 mutant described in Fig. 6AGo. The identity of the mutant reporter construct was confirmed by sequencing using the dideoxynucleotide chain-termination method. An expression vector encoding ß-galactosidase driven by the simian virus 40 early promoter (pSV-ß-galactosidase from Promega) was used in transfection studies as an internal standard and control. SMAD2, -3, and -4 expression vectors in pCS2 were a kind gift from Dr. Malcolm Whitman (Harvard Medical School, Boston, MA) (38). The dominant-negative C-terminal truncated SMAD3 expression vector was a gift from Dr. Rik Derynck (University of California at San Francisco, San Francisco, CA) (39).

Cell Culture and Transient Transfection
LßT2 (mouse gonadotrope-derived) cells were maintained in monolayer culture in high-glucose DMEM (Invitrogen, Carlsbad, CA) supplemented with 10% (vol/vol) fetal bovine serum (Omega Scientific, Tarzana, CA), 100 U/ml penicillin, and 100 µg/ml streptomycin sulfate (Invitrogen) at 37 C in humidified 5% CO2-95% air. Cells were transiently transfected with one of the rFSHßLuc constructs, either alone or in combination with a SMAD expression vector, by electroporation. In all transfections, pSV-ß-galactosidase was included to serve as an internal standard and control. For each experiment, LßT2 cells were suspended in 0.4 ml of Dulbecco’s PBS plus 5 mM glucose containing the plasmid DNA to be transfected. In each experiment, the total amount of DNA transfected was standardized between groups by cotransfecting with the empty expression vector when necessary. The cells were exposed to a single electrical pulse of 0.24 V from a total capacitance of 960 µF using a gene pulser apparatus from Bio-Rad Laboratories (Hercules, CA). After electroporation, cells were plated in 10% FBS-containing medium in six-well plates.

Cells cotransfected with the SMAD expression vectors were maintained in culture for 48 h after transfection. The medium was changed after 24 h and at 48 h. The cells were harvested and luciferase assays were performed as previously described (56) using an LB 953 Autolumat luminometer (EG&G Berthold, Nashua, NH) set to measure for 20 sec with no delay.

For LßT2 cells transiently transfected with the rFSHßLuc construct alone, or in combination with the dominant-negative SMAD3 expression vector, cells were maintained in DMEM containing 10% FBS for 24 h; 24 h after transfection the medium was changed to a 1% FBS-containing DMEM, and three wells were designated to one of the following treatment groups: 1) no treatment (medium alone for 24 h); 2) 100 ng/ml activin A for 24 h; 3) 100 nM GnRH agonist for 4 h; or 4) 100 ng/ml activin A for 24 h and 100 nM GnRH agonist for the final 4 h. Cells were harvested and luciferase assays were performed 48 h after transfection. To determine the optimal doses of GnRH and activin A, and the optimal time course required for maximal luciferase stimulation, dose response and time course experiments were performed (data not shown). GnRH stimulation of transcriptional activity was maximal after 4 h stimulation with 100 nM GnRH agonist. For activin A, transcriptional activity was maximal after 24 h stimulation with 100 ng/ml activin A.

For experiments using the MAPK inhibitor, cells were first maintained in DMEM containing 10% FBS. The medium was changed to DMEM containing 1% FBS 23 h after transfection, and the cells were split into two groups, group A and group B. The MAPK inhibitor (UO126) was resuspended according to the manufacturer’s guidelines using DMSO. Wells from group A were used as a control and were treated with DMSO. Wells from group B were treated with 10 µM UO126 for 1 h before addition of any further treatment and throughout the remainder of the experimental period. Wells from both groups were designated to one of the following treatment groups: 1) no treatment (medium for 24 h); 2) 100 ng/ml activin A for 24 h; 3) 100 nM GnRH agonist for 4 h, or 4) 100 ng/ml activin A for 24 h and 100 nM GnRH agonist for the final 4 h.

Cells were harvested and luciferase assays were performed 48 h after transfection. ß-Galactosidase activity was assayed at 410 nm in a DU640 spectrophotometer (Beckman Coulter, Inc., Fullerton CA) using standard colorimetric protocols (56). Luciferase activity was normalized to ß-galactosidase activity.

Preparation of Nuclear Extract
LßT2 cells were grown to approximately 60% confluency in 100 x 20 mm tissue culture dishes (Corning, Inc., Corning, NY) in DMEM containing 10% FBS. The medium was then changed to a 1% FBS-containing DMEM before treatment with 100 ng/ml activin A for 0 or 30 min, or 1 h, 4 h, 12 h, 24 h, or 48 h, followed by microextraction of nuclear protein. Nuclear proteins were also extracted from cells treated with GnRH agonist (100 nM) for 0 or 30 min or 1 h, 2 h, or 4 h, either with or without 24 h activin A treatment (100 ng/ml). All cells were subsequently washed with 12 ml Dulbecco’s PBS and harvested for nuclear protein microextraction following the method of Therrien and Drouin (57).

EMSA
A double-stranded oligonucleotide corresponding to –284/–252 of the rFSHß gene promoter (rFSHß-P) containing a putative palindromic SMAD-binding element was used as a probe for EMSA. Complementary oligonucleotides were annealed in an annealing buffer containing 0.1 M NaCl, 10 mM TRIS-HCl (pH 8.0), and 1 mM EDTA (pH 8.0) and labeled at the 5'-end with [{gamma}-32P]ATP (PerkinElmer Life Sciences, Boston, MA) by T4 polynucleotide kinase (New England Biolabs, Beverly, MA). Nuclear extracts (5 µg) were incubated for 1 h on ice with a 200,000 cpm probe in DNA binding buffer [0.01 µg/µl salmon sperm DNA, 2.15 mM phenylmethyl sulfonyl fluoride, 5 mM dithiothreitol, 20 mM HEPES (pH 7.9), 60 mM KCl, 5 mM MgCl2, 1 mg/ml BSA, and 5% (vol/vol) glycerol]. For competition studies, unlabeled DNA was added 1 h before the addition of the {gamma}-32P-labeled DNA probe. rFSHß-P was used as unlabeled competitor in the competition studies. Sequential 2-bp mutations of rFSHß-P (M1–M6) and an unrelated consensus SF1 binding sequence [LHSF (58)] were also used as competitors. For supershift experiments, antibodies to SMAD2/3 (Santa Cruz), SMAD3 (Santa Cruz and Zymed), and SMAD4 (Santa Cruz) were added to the binding mixture 1 h before the addition of the {gamma}-32P-labeled DNA probe. The protein-DNA complexes were resolved by gel electrophoresis on 5% low-ionic strength nondenaturing PAGE in 0.5x Tris-borate-EDTA buffer (45 mM Tris base, 45 mM boric acid, 1 mM EDTA, pH 8.3). The gels were then dried and exposed to radiographic film for analysis.

Magnetic Separation of DNA Binding Proteins
rFSHß-P and an oligonucleotide corresponding to –120/–90 of the rFSHß gene promoter were 5'-biotinylated and used as probes to isolate DNA-binding proteins in LßT2 cell nuclear extract. Nuclear extract (10 µg) was incubated for 2 h on ice with 160 pmol of double-stranded probe in DNA binding buffer [0.01 µg/µl salmon sperm DNA, 2.15 mM phenylmethyl sulfonyl fluoride, 5 mM dithiothreitol, 20 mM HEPES (pH 7.9), 60 mM KCl, 5 mM MgCl2, 1 mg/ml BSA, and 5% (vol/vol) glycerol]. 400 µg Dynabeads MyOne streptavidin (DynAl Biotech, Lake Success, NY) were washed three times in DNA binding buffer before being added to the reaction vessel. The binding mixture was incubated for an additional 30 min on a rocking platform at 4 C. DNA-protein complexes bound to the dynabeads were immobilized using a magnetic particle concentrator (MPC-S, DynAl). The beads were washed four times in 400 µl of binding buffer to remove excess unbound protein, and the eluted proteins were analyzed by SDS-PAGE Western blot analysis.

Western Blot Analysis
Whole-cell lysates were extracted from LßT2 cells 48 h after transient transfection of SMAD2, -3, or -4 expression vectors. Cells were washed with 12 ml of Dulbecco’s PBS and harvested in a total of 250 µl of RIPA buffer [1x PBS, 1% Igepal CA-630 (Sigma-Aldrich, St. Louis, MO), 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate, 0.1 mg/ml PMSF, 1 mg/ml aprotinin, and 1 mM sodium orthovanadate]. Cell lysates were microcentrifuged at 10,000 x g for 10 min at 4 C, and the supernatant was collected and stored at –80 C. Protein lysate (5 µg) was incubated with 2x sodium dodecyl sulfate gel loading buffer containing 200 mM dithiothreitol at 100 C for 3 min and electrophoresed on 7.5% SDS-PAGE at 80 V for 20 min, and then at 150 V for 1 h. Proteins were transferred to Immobilon-P transfer membrane (Millipore Corp., Bedford, MA) using Tris-glycine transfer buffer (12 mM Tris base, 96 mM glycine, 20% methanol) for 1 h at 4 C. Membranes were subsequently incubated in blocking solution containing 3% nonfat milk and 2% BSA (Sigma-Aldrich) in Tris-buffered saline (10 mM Tris, pH 8.0; 150 mM NaCl) with 0.05% Tween-20 (TBS-T; Fisher Scientific, Hampton, NH). After blocking, the membranes were incubated with one of the following antibodies overnight at 4 C: SMAD3 (2 µg/ml, Zymed); SMAD 2/3 (1:1000, Upstate), or actin (1:500, Santa Cruz). Membranes were washed three times in TBS-T before being incubated for 1 h at room temperature in goat antirabbit IgG horseradish peroxidase (HRP) diluted to a concentration of 1:5000 with blocking solution. The actin antibody was HRP conjugated and did not require incubation with a secondary antibody. Blots were washed three times with TBS-T and incubated with supersignal west pico chemiluminescent substrate (Pierce Chemical Co., Rockford, IL) for the detection of HRP. Western blots were analyzed after exposure to light-sensitive Biomax ML film (Eastman Kodak Co., Rochester, NY).

Statistical Analysis
Transfections were performed in triplicate and repeated a minimum of three times. For all transfection experiments, ß-galactosidase activity was used as an internal standard to correct for cell transfection efficiency. Data were expressed as luciferase/ß-galactosidase activity. ANOVA was performed to determine whether there was a statistically significant difference between treatment groups, and Fisher’s protected least significant difference post hoc test (PLSD) was used to make pairwise comparisons.


    ACKNOWLEDGMENTS
 
We thank Dr. Malcolm Whitman for the SMAD expression vectors and Dr. Pamela Mellon for the LßT2 cells.


    FOOTNOTES
 
This work was supported by National Institutes of Health Grant RO1 HD33001 (to U.B.K.).

First Published Online September 16, 2004

Abbreviations: DMSO, Dimethylsulfoxide; GnRHR, GnRH receptor; HRP, horseradish peroxidase; LHSF, LH steroidogenic factor-1 binding site; MEK, MAPK kinase; SMAD, mediator of decapentaplegic-related protein; TBS-T, Tris-buffered saline-Tween 20.

Received for publication December 8, 2003. Accepted for publication September 8, 2004.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

  1. Weiss J, Guendner MJ, Halvorson LM, Jameson JL 1995 Transcriptional activation of the follicle-stimulating hormone ß subunit gene by activin. Endocrinology 136:1885–1891[Abstract]
  2. Carroll RS, Corrigan AZ, Vale W, Chin WW 1991 Activin stabilizes follicle-stimultaing hormone-ß messenger ribonucleic acid levels. Endocrinology 129:1721–1726[Abstract]
  3. Attardi B, Winters SJ 1993 Decay of follicle-stimulating hormone-ß messenger RNA in the presence of transcriptional inhibitors and/or inhibin, activin, or follistatin. Mol Endocrinol 7:668–680[Abstract]
  4. Ling N, Ying SY, Ueno N, Shimasaki S, Esch F, Hotta M, Guillemin R 1986 Pituitary FSH is released by a heterodimer of the ß-subunits from the two forms of inhibin. Nature 321:779–782[Medline]
  5. Ling N, Ying SY, Ueno N, Shimasaki S, Esch F, Hotta M, Guillemin R 1986 A homodimer of the ß-subunits of inhibin A stimulates the secretion of pituitary follicle stimulating hormone. Biochem Biophys Res Commun 138:1129–1137[Medline]
  6. Mason AJ, Berkemeier LM, Schmelzer CH, Schwall RH 1989 Activin B: precursor sequences, genomic structure and in vitro activities. Mol Endocrinol 3:1352–1358[Abstract]
  7. Wilson ME, Handa RJ 1998 Activin subunit, follistatin, and activin receptor gene expression in the prepubertal female rat pituitary. Biol Reprod 59:278–283[Abstract/Free Full Text]
  8. Roberts V, Meunier H, Vaughan J, Rivier J, Rivier C, Vale W, Sawchen P 1989 Production and regulation of inhibin subunits in pituitary gonadotropes. Endocrinology 124:552–554[Abstract]
  9. Fernandez-Vazquez G, Kaiser UB, Albarracin CT, Chin WW 1996 Transcriptional activation of the gonadotropin-releasing hormone receptor gene by activin A. Mol Endocrinol 10:356–366[Abstract]
  10. Roberts VJ, Peto CA, Vale W, Sawchenko PO 1992 Inhibin/activin are costored with FSH and LH in secretory granules of the rat anterior pituitary gland. Neuroendocrinology 56:214–224[Medline]
  11. Corrigan AZ, Bilezikjian LM, Carroll RS, Bald LN, Schmelzer CH, Fen BM, Mason AJ, Chin WW, Schwall RH, Vale W 1991 Evidence for an autocrine role of activin B within rat anterior pituitary cultures. Endocrinology 128:1682–1684[Abstract]
  12. Massague J, Chen Y-G 2000 Controlling TGF-ß signaling. Genes Dev 14:627–644[Free Full Text]
  13. Derynck R, Zhang YE 2003 Smad-dependent and Smad-independent pathways in TGF-ß family signalling. Nature 425:577–584[CrossRef][Medline]
  14. Derynck R, Zhang Y, Feng X-H 1998 Smads: transcriptional activators of TGF-ß responses. Cell 95:737–740[Medline]
  15. Zawel L, Dai JL, Buckhaults P, Zhou S, Kinzler KW, Vogelstein B, Kern SE 1998 Human SMAD3 and SMAD4 are sequence-specific transcription activators. Mol Cell 1:611–617[Medline]
  16. Shi Y, Wang Y-F, Jayaraman L, Yang H, Massague J, Pavletich NP 1998 Crystal structure of Smad MN1 domain bound to DNA: insights on DNA binding in TGF-ß signaling. Cell 94:585–594[Medline]
  17. Dennler S, Itoh S, Vivien D, ten Dijke P, Huet S, Gauthier J-M 1998 Direct binding of Smad3 and Smad4 to critical TGFß-inducible elements in the promoter of human plasminogen activator inhibitor-type 1 gene. EMBO J 11:3091–3100[CrossRef]
  18. Yagi K, Goto D, Hamamoto T, Takenoshita S, Kato M, Miyazono K 1999 Alternatively spliced varient of Smad2 lacking exon 3. J Biol Chem 2:703–709[CrossRef]
  19. Chen X, Weisberg E, Fridmacher V, Watanabe M, Naco G, Whitman M 1997 Smad4 and Fast-1 in the assembly of activin-responsive factor. Nature 389:85–89[CrossRef][Medline]
  20. Liu F, Pouponnot C, Massague J 1997 Dual role of the Smad4/DPC4 tumor suppressor in TGFß-inducible transcriptional complexes. Genes Dev 11:3157–3167[Abstract/Free Full Text]
  21. Labbe E, Silvestri C, Hoodless PA, Wrana JL, Attisano L 1998 SMAD2 and SMAD3 positively and negatively regulate TGFß-dependent transcription through the forkhead DNA-binding protein FAST2. Mol Cell 2:109–120[Medline]
  22. Massague J, Wotton D 2000 Transcriptional control by the TGF-ß/Smad signaling system. EMBO J 19:1745–1754[Abstract/Free Full Text]
  23. Miyazawa K, Shinozaki M, Hara T, Furuya T, Miyazono K 2002 Two major Smad pathways in TGF-ß superfamily signalling. Genes Cells 7:1191–1204[Abstract/Free Full Text]
  24. Gharib SD, Wierman ME, Shupnik MA, Chin WW 1990 Molecular biology of the pituitary gonadotropins. Endocr Rev 11:177–199[Medline]
  25. Padmanabhan V, Karsch FJ, Lee JS 2002 Hypothalamic, pituitary and gonadal regulation of FSH. Reprod Suppl 59:67–82[Medline]
  26. Sealfon SC, Weinstein H, Millar RP 1997 Molecular mechanisms of ligand interaction with the gonadotropin-releasing hormone receptor. Endocr Rev 18:180–205[Abstract/Free Full Text]
  27. Shacham S, Harris D, Ben-Shlomo H, Cohen I, Bondil D, Przedecki F, Lewy H, Ashkenazi IE, Seger R, Naor Z 2001 Mechanisms of GnRH receptor signaling on gonadotropin release and gene expression in pituitary gonadotrophs. Vitam Horm 63:63–90[CrossRef][Medline]
  28. Norwitz ER, Xu S, Jeong K-H, Bedecarrats GY, Winebrenner LD, Chin WW, Kaiser UB 2002 Activin A augments GnRH-mediated transcriptional activation of the mouse GnRH receptor gene. Endocrinology 143:985–997[Abstract/Free Full Text]
  29. Norwitz ER, Xu S, Spiryda LB, Park JS, Jeong K-H, McGee EA, Kaiser UB 2002 Direct binding of AP-1 (Fos/Jun) proteins to a SMAD binding element facilitates both gonadotropin-releasing hormone (GnRH)- and activin-mediated transcriptional activation of the mouse GnRH receptor gene. J Biol Chem 277:37469–37478[Abstract/Free Full Text]
  30. Graham KE, Nusser KD, Low MJ 1999 LßT2 gonadotrope cells secrete follicle stimulating hormone (FSH) in response to activin A. J Endocrinol 162:R1–R5
  31. Pernasetti F, Vasilyev VV, Rosenburg SB, Bailey JS, Huang HJ, Miller WL, Mellon PL 2001 Cell-specific transcriptional regulation of follicle-stimulating hormone-ß by activin and gonadotropin-releasing hormone in the LßT2 pituitary gonadotrope cell model. Endocrinology 142:2284–2295[Abstract/Free Full Text]
  32. Carroll RS, Kowash PM, Lofgren JA, Schwall RH, Chin WW 1991 In vivo regulation of FSH synthesis by inhibin and activin. Endocrinology 129:3299–3304[Abstract]
  33. Kochman K, Gajewska A 1996 Biosynthesis of gonadotropins in vivo. Acta Neurobiol Exp (Warsz) 56:753–756[Medline]
  34. Weiss J, Harris PE, Halvorson LM, Crowley WF, Jameson JL 1992 Dynamic regulation of follicle-stimulating hormone-ß messenger ribonucleic acid levels by activin and gonadotropin-releasing hormone in perifused rat pituitary cells. Endocrinology 131:1403–1408[Abstract]
  35. Welt C, Sidis Y, Keutmann H, Schneyer A 2002 Activins, inhibins, and follistatins: from endocrinology to signaling. A paradigm for the new millennium. Exp Biol Med 227:724–752[Abstract/Free Full Text]
  36. Suszko MI, Lo DJ, Suh H, Camper SA, Woodruff S 2003 Regulation of the rat follicle stimulating hormone ß-subunit promoter by activin. Mol Endocrinol 17:318–332[Abstract/Free Full Text]
  37. Ellsworth BS, Burns AT, Escudero KW, Duval DL, Nelson SE, Clay CM 2003 The gonadotropin releasing hormone (GnRH) receptor activating sequence (GRAS) is a composite regulatory element that interacts with multiple classes of transcription factors including SMADS, AP-1 and a forkhead DNA binding protein. Mol Cell Endocrinol 29:93–111[CrossRef]
  38. Yeo CY, Chen X, Whitman M 1999 The role of FAST-1 and Smads in transcriptional regulation by activin during early Xenopus embryogenesis. J Biol Chem 274:26583–26590
  39. Zhang Y, Feng X, We R, Derynck R 1996 Receptor-associated Mad homologues synergize as effectors of the TGF-ß response. Nature 383:168–172[CrossRef][Medline]
  40. Dupont J, McNeilly J, Vaiman A, Canepa S, Combarnous Y, Taragnat C 2003 Activin signaling pathways in ovine pituitary and LßT2 gonadotrope cells. Biol Reprod 68:1877–1887[Abstract/Free Full Text]
  41. Bernard DJ 2004 Both SMAD2 and SMAD3 mediate activin-stimulated expression of the follicle-stimulating hormone ß subunit in mouse gonadotrope cells. Mol Endocrinol 18:606–623[Abstract/Free Full Text]
  42. Huang H-J, Sebastian J, Strahl BD, Wu JC, Miller WL 2001 Transcriptional regulation of the ovine follicle-stimulating hormone-ß gene by activin and gonadotropin-releasing hormone (GnRH): involvement of two proximal activator protein-1 sites for GnRH stimulation. Endocrinology 142:2267–2274[Abstract/Free Full Text]
  43. Gajewska A, Siawrys G, Bogacka I, Przala J, Lerrant Y, Counis R, Kochman K 2002 In vivo modulation of follicle-stimulating hormone release and ß subunit gene expression by activin A and the GnRH agonist buserelin in female rats. Brain Res Bull 58:475–480[CrossRef][Medline]
  44. Zhang L, Wang W, Hayashi Y, Jester JV, Birk DE, Gao M, Liu C-Y, Kao WW-Y, Karin M, Xia Y 2003 A role for MEK kinase 1 in TGF-ß/activin-induced epithelium movement and embryonic eyelid closure. EMBO J 22:4443–4454[Abstract/Free Full Text]
  45. Kaiser UB, Sabbagh E, Katzenellenbogen RA, Conn PM, Chin WW 1995 A mechanism for the differential regulation of gonadotropin subunit gene expression by gonadotropin-releasing hormone. Proc Natl Acad Sci USA 92:12280–12284[Abstract]
  46. Hata A, Lo RS, Wotton D, Lagna G, Massague J 1997 Mutations increasing autoinhibition inactivate tumour suppressors Smad2 and Smad4. Nature 388:82–87[CrossRef][Medline]
  47. Hayashida T, Decaestecker M, Schnaper HW 2003 Cross-talk between ERK MAP kinase and Smad signaling pathways enhances TGF-ß-dependent responses in human mesangial cells. FASEB J 11:1576–1578
  48. Furukawa F, Matsuzaki K, Mori S, Tahashi Y, Yoshida K, Sugano Y, Yamagata H, Matsushita M, Seki T, Inagaki Y, Nishizawa M, Fujisawa J, Inoue K 2003 p38 MAPK mediates fibrogenesis signal through Smad3 phosphorylation in rat myofibroblasts. Hepatology 38:879–889[CrossRef][Medline]
  49. Yamamura Y, Hua X, Bergelson S, Lodish HF 2000 Critical role of Smads and AP-1 complex in transforming growth factor-ß-dependent apoptosis. J Biol Chem 275:36295–36302[Abstract/Free Full Text]
  50. Chen CR, Kang Y, Siegel PM, Massague J 2002 E2F4/5 and p107 as Smad cofactors linking the TGFß receptor to c-myc repression. Cell 110:19–32[Medline]
  51. Chen F, Ogawa K, Liu X, Stringfield TM, Chen Y 2002 Repression of Smad2 and Smad3 transactivating activity by association with a novel splice variant of CCAAT-binding factor C subunit. Biochem J 364:571–577[CrossRef][Medline]
  52. Bailey JS, Rave-Harel N, McGuillivray SM, Coss D, Mellon PL 2004 Activin regulation of the follicle-stimulating hormone ß-subunit gene involves Smads and the TALE homeodomain proteins Pbx1 and Prep1. Mol Endocrinol 18:1158–1170[Abstract/Free Full Text]
  53. Cocolakis E, Lemay S, Ali S, Lebrun J-J 2001 The p38 MAPK pathway is required for cell growth inhibition of human breast cancer cells in response to activin. J Biol Chem 276:18430–18436[Abstract/Free Full Text]
  54. Nordeen SK 1988 Luciferase reorter gene vectors for analysis of promoters and enhancers. Biotechniques 6:454–458[Medline]
  55. Zakaria MM, Jeong K-H, Lacza C, Kaiser UB 2002 Pituitary homeobox 1 (Ptx1) activates the rat follicle-stimulating hormone ß (rFSHß) gene promoter. Mol Cell Endocrinol 16:1840–1852[CrossRef]
  56. Edlund T, Walker MD, Barr PJ, Rutter WJ 1985 Cell-specific expression of the rat insulin gene: evidence for role of two distinct 5' flanking elements. Science 230:912–916[Medline]
  57. Therrien M, Drouin J 1993 Cell-specific helix-loop-helix factor required for pituitary expression of the pro-opiomelanocortin gene. Mol Cell Biol 13:2342–2353[Abstract]
  58. Halvorson LM, Kaiser UB, Chin WW 1996 Stimulation of luteinizing hormone ß gene promoter activity by the orphan nuclear receptor, steroidogenic factor-1. J Biol Chem 12:6645–6650[CrossRef]