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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.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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 
T3–1 and LßT2 gonadotrope-derived cell lines. GnRH-stimulated transcriptional activity of the mouse GnRHR gene promoter was significantly enhanced when T3–1 cells were cotreated with activin and GnRH in combination (28, 29). Immunocytochemical studies identified that treatment of 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.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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). 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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). 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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). 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. 3). 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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) was incubated with nuclear proteins extracted from LßT2 cells treated with activin A for 0, 1, 4, 12, 24, and 48 h (Fig. 4B). 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. 4C) or into several closely migrating complexes (Fig. 4B). The gel in Fig. 4C was run over a shorter period of time, and the bands present in complex a are not as well resolved as those in Fig. 4B. Complexes b1 and b2 also migrated closely to each other (Fig. 4, B and C). The intensity of complexes a, b1, and b2 increased after treatment with activin A (Fig. 4B), 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. 1 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. 4C) 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. 5A).
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, 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. 2A and 3). 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. 5A, lane 8) identified SMAD3 protein after magnetic separation of proteins bound to rFSHß-P (Fig. 5B; 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. 5B).
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. 6A). These mutants, designated M1–M6, were used as competitors in EMSA studies in which the SMAD4 complex was supershifted (Fig. 6B). In the presence of SMAD4 antibody, two supershifted complexes could again be clearly detected (Fig. 6B, 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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) 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. 6C). 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. 6C). 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. 7A). 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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). 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. 7, 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. 8). 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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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. 9). 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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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.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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. 4, 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. 1 and 7A show a clear synergistic enhancement in transcriptional activity after treatment with activin and GnRH, the overall magnitude of all responses was reduced in Fig. 7A. This may be due to the increased passage number of LßT2 cells used in Fig. 7A, 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 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. 8). 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.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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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. 6A. 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 [
-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 -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 -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.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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