Activin Responsiveness of the Murine Gonadotropin-Releasing Hormone Receptor Gene Is Mediated by a Composite Enhancer Containing Spatially Distinct Regulatory Elements
Brian D. Cherrington,
Todd A. Farmerie,
Clay A. Lents,
Jeremy D. Cantlon,
Mark S. Roberson and
Colin M. Clay
Animal Reproduction and Biotechnology Laboratory (B.D.C., T.A.F., C.A.L., J.D.C., C.M.C.), Department of Biomedical Sciences, Colorado State University, Fort Collins, Colorado 80523; and Department of Biomedical Sciences (M.S.R.), Cornell University, Ithaca, New York 14853
Address all correspondence and requests for reprints to: Colin M. Clay, Colorado State University, Department of Biomedical Sciences/Animal Reproduction and Biotechnology Laboratory, 1683 Campus Delivery, Ft. Collins, Colorado 80523. E-mail: Colin.Clay{at}colostate.edu.
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ABSTRACT
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The promoters of mouse and rat GnRH receptor (GnRHR) genes differ markedly in regard to activin regulation. Activin stimulates the mouse GnRHR promoter, although it has no impact on that of the rat. To test whether this difference was due to a single nucleotide change in the rat GnRHR activating sequence (GRAS) homolog, we tested a mouse promoter with the rat GRAS homolog and a rat promoter with intact mouse GRAS. The single change in GRAS eliminated activin responsiveness of the mouse GnRHR promoter; however, intact mouse GRAS did not confer activin responsiveness to the rat promoter. Thus, although necessary, GRAS is not sufficient for activin responsiveness of the murine GnRHR promoter. Use of chimeric rat and mouse promoters led to the identification of a 36-bp region residing immediately downstream of GRAS that is necessary for activin responsiveness of the mouse GnRHR gene promoter. Scanning mutagenesis of the 36-bp region localized the functional boundaries of the key regulatory element to adjacent TAAT motifs. The presence of tandem TAAT motifs, the core binding site for multiple members of the homeodomain family of binding proteins, raised the possibility that this region represented a binding site for a homeodomain protein. This region displayed specific binding to a recombinant homeodomain of LHX2. We suggest that GRAS and the downstream activin regulatory element together define a unique and complex activin/TGFß-responsive "enhanceosome" whose functional attributes depend on the binding of multiple classes of transcription factors at spatially distinct sites in the promoter of the murine GnRHR gene.
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INTRODUCTION
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THE INTERACTION OF GnRH with specific, high-affinity receptors located on gonadotrope cells of the anterior pituitary gland is central to the regulation of reproductive function in mammals. The pulsatile discharge of GnRH from hypothalamic neurons not only stimulates but also is obligatory for synthesis and secretion of LH (1, 2). Given the pivotal role of GnRH in reproduction, much effort has been expended toward understanding the physiological consequences of regulation of GnRH and its cognate pituitary receptor. A member of the superfamily of hepathelical G protein-coupled receptors, the GnRH receptor (GnRHR) is coupled to activation of G
q/11, which initiates multiple intracellular responses including LH secretion and increased expression of the common
- and unique LHß-subunit genes and the GnRHR gene itself (1, 3, 4). It is clear that relative changes in GnRH secretion from the hypothalamus are important determinants of LH secretion; however, changes in the number of pituitary receptors for GnRH are also implicated as an important mechanism underlying the regulation of gonadotropin secretion (5, 6, 7). Thus, both hypothalamic secretion of GnRH and pituitary concentration of GnRHRs are targets for regulation. In regard to the latter, of the multiple inputs that have been implicated in affecting changes in GnRHR numbers, perhaps the most dramatic are those associated with estradiol-17ß, GnRH, and activin (8, 9, 10, 11).
A member of the TGFß family of signaling molecules, activin represents homo- or heterodimeric complexes of the different inhibin ß-subunits and is implicated in a number of physiological processes ranging from embryonic development and tissue patterning to the synthesis and secretion of FSH (12, 13, 14). As with other members of the TGFß family, intracellular signaling by activin is initiated by binding to a type II receptor at the plasma membrane and subsequent recruitment and phosphorylation of a type I receptor (12, 13). Functionally, this complex acts as a serine/threonine kinase and phosphorylates the activin/TGFß restricted Smad proteins (Smad2 or 3) (12, 13, 15). These Smad proteins then typically associate with the common Smad partner (Smad4) and translocate from the cytoplasm to the nucleus. In the nucleus, Smads bind to specific DNA regulatory elements to alter the rate of transcription of target genes; however, because Smad proteins alone display relatively weak DNA binding, transcriptional regulation by these proteins is often achieved via multiprotein complexes that contain both Smad and non-Smad protein partners (12, 13, 15).
Within the anterior pituitary gland, the activin ß-subunits are produced by gonadotrope cells as is the activin binding protein follistatin (12, 16). The latter is considered to be a primary modulator of the biological effects of activin and thus prevents activin binding to its cognate receptor (17, 18, 19). In addition to gonadotrope expression, follistatin is also produced by pituitary folliculostellate cells (20, 21). Thus, although initially described as an endocrine regulator of pituitary function, autocrine and paracrine mechanisms are likely more important components of activin signaling in the pituitary gland (22, 23). In this regard, it is clear that regulation of FSH production is a central biological role of activin in the pituitary (17). The effects of activin on FSH are evident as both increased secretion and enhanced expression of the FSH ß-subunit gene (17, 24, 25). In addition to its effects on FSH, multiple studies have established that activin regulates GnRHR expression (26, 27, 28, 29, 30). At issue then are the mechanisms that account for activin regulation of FSHß and GnRHR gene expression. To address the latter, we have examined transcriptional activity of the GnRHR gene promoter in the gonadotrope-derived
T31 cell line (31). Given that
T31 cells, like gonadotropes, express activin and activin receptors, we used an indirect approach based on the ability of follistatin to bind and inactivate activin to localize activin responsiveness of the murine GnRHR gene to an element termed the GnRHR activating sequence or GRAS (26).
Consistent with its ability to mediate activin responsiveness, GRAS has been shown to bind members of the Smad family of transcription factors (27, 28). However, the functional phenotype of GRAS depends not only on Smad binding but also activator protein-1 (AP-1) and FoxL2, a member of the winged-helix or forkhead family of transcription factors (27, 28). Thus, as is the case with many activin/TGFß response elements, GRAS is a composite enhancer whose functional activity is dependent on the binding of a Smad protein complex and multiple non-Smad protein partners (27, 28). Furthermore, elimination of any one of these binding components eliminates the functional activity of GRAS (27). In light of this, we predicted that a single nucleotide difference between mouse GRAS and the rat GRAS homolog (32) would deprive the latter of activin responsiveness. Consistent with this prediction, we find that the rat GnRHR promoter is not responsive to activin. However, GRAS alone cannot account for the functional divergence in activin responsiveness of the proximal promoters of the mouse and rat GnRHR genes. This observation raised the possibility that, while necessary, GRAS may not be sufficient for activin responsiveness of the mouse GnRHR gene. Consistent with this possibility, we find that activin responsiveness of the mouse GnRHR gene requires not only GRAS but also a previously undefined element located approximately 15 bp downstream of GRAS that we term the downstream activin regulatory element (DARE). Based on the ability of DARE to interact with a recombinant homeodomain, we suggest that activin regulation of the murine GnRHR gene is mediated by a complex "enhanceosome" (33) that includes spatially distinct Smad and homeodomain binding componentsa scenario reminiscent of the mechanism recently described for activin regulation of the rat and ovine FSH ß-subunit genes (24, 34).
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RESULTS
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Adenoviral Delivery of Follistatin Is Effective in Attenuating Transcriptional Activity of the GnRHR Gene Promoter
As
T31 cells produce endogenous activin B, these cells essentially exist in a constitutively activin-stimulated state (26). As such, we have used an indirect approach based on the addition of the activin binding protein follistatin to functionally map activin/TGFß regulation of the GnRHR gene promoter, an approach that localized activin responsiveness of the GnRHR gene to GRAS (26). Unfortunately, this approach requires a continual supply of purified or recombinant follistatin. As an alternative, we tested the efficacy of adenoviral delivery of human follistatin (AdCAFS288, a generous gift from Dr. Wylie Vale, Salk Institute, La Jolla, CA) (35) in a transfection/infection paradigm. In our application,
T31 cells were transfected with either the mouse or rat 600 GnRHR promoters fused to luciferase, the mouse 600 promoter containing a loss of function mutation in GRAS (36), or three copies of GRAS fused to the rat prolactin minimal promoter (37). Three hours after transfection, cells were infected with 1000 MOI (multiplicity of infection) of AdCAFS288 or the same MOI of an adenoviral construct expressing green fluorescent protein (GFP). Consistent with earlier work using soluble recombinant follistatin (26), infection with AdCAFS288 led to an approximately 50% reduction in activity of the 600 wild-type mouse GnRHR promoter and a greater than 90% attenuation in activity of the 3XGRAS-LUC vector as compared with cells infected with Ad-GFP (Fig. 1
). Thus, adenoviral delivery of a follistatin cDNA is an effective paradigm for removal of activin input to
T31 cells. As expected, transcriptional activity of the 600 promoter containing mutated GRAS was unaffected by AdCAFS288. The proximal promoter of the rat GnRHR gene was not affected by follistatin expression.

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Fig. 1. Adenoviral Delivery of Follistatin Attenuates Transcriptional Activity of the Proximal Promoter of the Mouse But Not Rat GnRHR Genes
The indicated luciferase expression vectors were transiently transfected with pRSV-LacZ into T31 cells. Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity, and values are expressed as fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells. Values represent the mean ± SEM of triplicate samples in three separate transfections. The relative positions of GRAS, AP-1, and SF-1 elements are indicated in the mouse and rat promoters.
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A Single Nucleotide Change in GRAS Does Not Account for the Functional Divergence in Activin Responsiveness of the Mouse and Rat GnRHR Gene Promoters
Despite greater than 90% homology, the proximal promoters of the mouse and rat GnRHR genes are functionally divergent in regard to activin responsiveness; whereas the mouse promoter is activin responsive, the rat proximal promoter is not (Fig. 1
). A direct comparison of these promoters reveals perfect conservation of sequence in the steroidogenic factor 1 (SF-1) and AP-1 binding sites; however, the rat GRAS homolog differs from mouse by a single nucleotide transition (Fig. 2
). Given the requirement for GRAS in mediating activin regulation of the mouse GnRHR gene, we hypothesized that this single nucleotide change in the GRAS element would account for the divergence in activin responsiveness of the rat and mouse GnRHR promoters. To test this hypothesis, chimeric promoters were constructed in which the 1-bp difference in the GRAS element was reciprocally exchanged between the two promoters, essentially creating a rat GnRHR promoter with an "intact" mouse GRAS (rat-GRAS-repair) and a mouse GnRHR promoter with the rat GRAS homolog. We reasoned that this 1-bp exchange would render the rat promoter activin responsive, whereas the activin response of the mouse promoter would be lost. Consistent with this prediction, the transcriptional activity of the mouse promoter containing the rat GRAS homolog was unaffected by follistatin (Fig. 2
). Surprisingly, however, activin responsiveness was not conferred on the rat promoter containing intact mouse GRAS.

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Fig. 2. A Single Nucleotide Change in GRAS Does Not Account for the Functional Divergence in Activin Responsiveness of the Mouse and Rat GnRHR Genes
T31 cells were transfected with pRSV-LacZ and luciferase vectors containing approximately 600 bp of proximal promoter from either the mouse or rat GnRHR genes, the mouse promoter containing the rat GRAS homolog, or the rat promoter containing mouse GRAS (rat-GRAS-repair). Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity, and values are expressed as fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells. Values represent the mean ± SEM of triplicate samples in three separate transfections.
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A 36-bp Region Located between GRAS and AP-1 Is Necessary for Activin Responsiveness of the Mouse GnRHR Gene Promoter
The data in Fig. 2
are subject to several interpretations. First, the rat promoter contains an element that actively represses the functional activity of GRAS. Alternatively, the rat promoter is lacking an additional regulatory element that is necessary for activin responsivenessan element that would, presumably, be present in the mouse promoter. We reasoned that both possibilities would be testable by constructing a series of chimeric promoters in which progressive regions of proximal promoter were reciprocally exchanged between the mouse promoter and rat-GRAS-repair promoter. In short, we were screening for a promoter exchange that presented as both a loss and gain of function. The initial set of chimeric promoters represented exchanges at approximately 500, 250, and 150 bp between the wild-type mouse and rat promoters (Fig. 3
). There was no impact of exchanging the distal 100-bp regions on the follistatin response of either the mouse or rat-GRAS-repair promoters (Fig. 3
). In contrast, placement of approximately 250 or 350 bp of distal mouse sequence was sufficient to confer follistatin responsiveness to the proximal rat promoter. Thus, these exchanges defined a region between 500 and 250 in the mouse promoter that leads to a gain of activin regulation of the rat promoter. The converse result was observed when rat sequence was exchanged into the mouse promoter; either 250 or 350 bp of distal rat sequence eliminated follistatin response. As such, the critical regulatory information must reside between 500 and 250. Importantly, the contribution of at least a portion of this sequence was previously evaluated using a series of scanning mutations that placed the recognition site for NotI in 12 "block replacements" progressively from 500 to 365 (37). Only one of these (BR11) affected the follistatin/activin response of the mouse GnRHR gene promoter and, in fact, served to functionally identify GRAS (37). Thus, this previous work allowed us to direct additional exchanges to the promoter regions residing downstream of GRAS between approximately 365 and 250. Also, it was notable that this region contains several potential regulatory elements including a composite binding site for Oct-1 and nuclear factor Y (NF-Y) originally termed SURG-1 (sequence underlying GnRH regulation) that contributes to GnRH responsiveness of the murine promoter (38). Thus, we first focused on this 36-bp region and constructed chimeric promoters in which the homologous sequence was reciprocally exchanged between the mouse GnRHR promoter and the rat-GRAS-repair promoter (Fig. 4
). This 36-bp reciprocal exchange revealed the desired phenotype, i.e. both a gain of function (follistatin responsiveness) of the rat-GRAS-repair promoter and a loss of function of the mouse promoter (Fig. 4
). Thus, both GRAS and one or more regulatory elements arrayed within a 36-bp region located between GRAS and AP-1 would appear to be necessary for activin responsiveness of the mouse GnRHR gene.

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Fig. 3. Activin Response Maps to a Region between 500 and 250 of the Mouse GnRHR Gene Promoter
T31 cells were transfected with pRSV-LacZ, 600 wild-type mouse promoter, rat-GRAS-repair or the indicated chimeras of the mouse and rat GnRHR promoters. Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity, and values are expressed as fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells. Values represent the mean ± SEM of triplicate samples in three separate transfections.
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Fig. 4. A 36-bp Region between GRAS and AP-1 Is Necessary for Activin Responsiveness of the Mouse GnRHR Promoter and Sufficient to Confer Activin Responsiveness on the Rat-GRAS-Repair Promoter
T31 cells were transfected with pRSV-LacZ, the wild-type mouse and rat GnRHR promoters, or the indicated chimeras of the mouse and rat promoters. Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity, and values are expressed as fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells. Values represent the mean ± SEM of triplicate samples in three separate transfections. *, Value is different (P < 0.05) from fold change of wild-type 600 mouse GnRHR promoter.
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The DARE Localizes to an 18-bp Sequence between 365 and 348 in the Proximal Promoter of the Mouse GnRHR Gene
The data in Fig. 4
suggest the presence of an additional regulatory element located between 375 and 340 that cooperates with GRAS to confer activin responsiveness. To refine the functional boundaries of this accessory element(s), we next expanded the scanning mutagenesis approach that was used to define GRAS and placed the 8-bp recognition motif for NotI between 372 and 340 in the context of the mouse GnRHR promoter. In all, four separate mutations (BR13-BR16) were constructed (Fig. 5
). Although BR13 and BR16 had no effect on activin responsiveness (follistatin repression), the follistatin response of the mouse promoter was eliminated by placement of NotI between 355 and 348 (BR15) and attenuated by BR14 (365 to 358) (Fig. 5
). Thus, functional mapping localizes a novel DARE to a region between 365 and 348 in the mouse GnRHR gene promoter. Finally, it is interesting to note that, like mutations in GRAS, the BR14 and BR15 mutations decrease "basal" promoter activity to a level indistinguishable from that seen with follistatin treatment of the wild-type mouse GnRHR promoter (Fig. 5
).

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Fig. 5. The DARE Localizes to an 18-bp Sequence between 365 and 348 in the Proximal Promoter of the Mouse GnRHR Gene
T31 cells were transfected with pRSV-LacZ, the wild-type mouse or rat GnRHR promoters, or mouse GnRHR promoters containing a series of scanning NotI mutations. Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity, and values are expressed as both adjusted light units (left panel) or fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells (right panel). Values represent the mean ± SEM of triplicate samples in three separate transfections. *, Value is different (P < 0.05) from fold change of wild-type 600 mouse GnRHR promoter.
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NF-Y Does Not Account for the Functional Properties of DARE
A canonical CCAAT motif located within the 36-bp exchange region has been shown to bind the CCAAT box binding factor NF-Y (39, 40). Interestingly, the BR15 mutation effectively disrupts this motif and also disrupts activin responsiveness; however, BR16 also eliminates the core CCAAT motif but has no effect on the follistatin response of the mouse promoter. Thus, it seems unlikely that this motif or NF-Y accounts for the functional properties of DARE. Nevertheless, we used EMSA to directly test the ability of DARE to displace NF-Y binding to an established NF-Y binding site in the LH ß-subunit gene (41). Consistent with Keri et al. (41), a specific bound complex was evident with the LHß probe and was shifted by anti-NF-Y antiserum (Fig. 6
). Homologous competition displaced binding to the radioactive probe in the supershifted complex. In contrast, whereas the BR14 oligonucleotide displayed a limited capacity to displace NF-Y binding to the radioactive probe, essentially no competition was evident with increasing concentrations of nonradioactive DARE, or DARE containing the BR15 and BR16 mutations. Thus, based on both the functional and EMSA data, it would not appear that the CCAAT box or NF-Y contributes to the functional activity of DARE. Finally, in addition to the NF-Y containing complex, a faster migrating complex was evident with LHß probe (indicated by an asterisk); however, as with NF-Y, the ability of the DARE competitors to displace binding at this complex was markedly reduced compared with the homologous competitor.

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Fig. 6. NF-Y Does Not Interact with the CCAAT Motif Located in DARE
An established NF-Y binding site from the LHß subunit gene was radiolabeled and incubated with T31 nuclear protein alone or in the presence of either IgG or NF-Y antibody and subjected to electrophoresis in a nondenaturing polyacrylamide gel. Specificity of binding in the supershifted NF-Y containing complex was assessed by adding increasing concentrations (10x, 50x, 100x) of nonradioactive homologous DNA, DARE, or DARE containing the indicated block replacement mutations.
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DARE Is Capable of Binding the LHX2 Homeodomain
In addition to the CCAAT motif, the region encompassed by DARE contains several TAAT motifs that are the core DNA binding sites of multiple homeodomain DNA binding proteins. Thus, we reasoned that DARE may represent a binding site for a homeobox binding protein. Consistent with this possibility, we find that an in vitro translated homeodomain of the LIM-homeodomain protein LHX2 is capable of binding DARE in EMSA (Fig. 7
). A supershift resulting from the inclusion of rabbit anti-LHX2 homeodomain antiserum (42), but not an equal volume of normal rabbit serum (NRS), was used to confirm the identity of the bound complex. Also, consistent with the functional mapping, the LHX2 complex was displaced by homologous competition but not by BR15.

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Fig. 7. DARE Is Capable of Binding in Vitro Translated LHX2 Homeodomain
Radioactively labeled DNA encompassing DARE (shaded sequence) was incubated with in vitro translated homeodomain of LHX2 either alone or in the presence of a 1:10 or 1:100 dilution of either NRS or LHX2 homedomain antiserum. In the final two lanes, the inclusion of a 100x molar excess of either DARE (homologous competition) or DARE containing the BR15 mutation in the binding reaction was used to assess the specificity of the bound complex.
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Disruption of the Paired TAAT Motifs in DARE Eliminates Activin Responsiveness of the Murine GnRHR Gene Promoter
The EMSA data suggest that DARE may represent a binding site for one or more homeodomain containing proteins. If correct, then we reasoned that the paired TAAT motifs disrupted by BR14 and BR15 would be key to the functional attributes of this element. To address this issue, a series of murine GnRHR promoters were constructed that contained 2-bp transversion mutations that spanned the entire BR14 and BR15 regions (designated µ14.15 and µ15.15; Fig. 8
). Although there was some attenuation in basal activity, the follistatin response of promoters containing the µ14.1, µ14.2, and µ14.3 mutations was not affected (Fig. 8
). In contrast, mutation of the TAAT motif in BR14 (µ14.4 and µ14.5) led to a significant reduction in both basal activity and the follistatin response as compared with the wild-type mouse promoter. Interestingly, mutation of the four bases of sequence (µ15.1 and µ15.2) residing between the two TAAT motifs had little effect on either basal promoter activity or activin responsiveness. However, as was the case for µ14.4 and µ14.5, disruption of the core TAAT motif in the BR15 region (µ15.3 and µ15.4) reduced both basal activity and the follistatin response of the mouse GnRHR promoter. Based on these data, the TAAT motifs in the BR14 and BR15 regions both contribute to the functional activity of DARE. We next constructed a promoter that contains both the µ14.4 and µ15.4 mutations (µ14.4/15.4), thus effectively disrupting both TAAT sites in a single construct. Consistent with a combined role for both TAAT motifs, the µ14.4/15.4 promoter displayed the lowest basal activity and a complete loss of the follistatin response. Finally, because the sequence of the rat GnRHR promoter in the BR14 and BR15 regions diverges from the mouse at five nucleotides, we next sought to determine whether any of these differences may partially account for the lack of functional activity of the rat DARE homolog. Toward this end, we constructed a mouse promoter in which the A residue in BR14 was converted to G (mouse 600, rat 14.4). Consistent with the dinucleotide transversions in µ14.4 and µ14.5, both basal activity and follistatin response of this construct were attenuated (Fig. 8
). In contrast, conversion of the three nucleotides in BR15 to the homologous rat sequence (mouse 600, rat 15.1/2) had little effect on either basal promoter activity or follistatin responsiveness, a result consistent with the µ15.1 and µ15.2 scanning mutants.

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Fig. 8. The TAAT Motifs in DARE Are Critical for Activin Regulation of the Mouse GnRHR Promoter
T31 cells were transfected with pRSV-LacZ, the wild-type mouse or rat GnRHR promoters or mouse GnRHR promoters containing a series of 2-bp mutations or rat-to-mouse substitutions. Three hours after transfection, cells were infected with either AdCAFS288 or Ad-GFP. Cells were harvested 48 h after infection, and cellular lysates were assayed for luciferase and ß-galactosidase activity. Luciferase values were corrected for ß-galactosidase activity and normalized to the level of expression of the 600 mouse GnRHR promoter in each transfection, and values are expressed as both normalized adjusted light units (left panel) or fold change in adjusted luciferase activity in Ad-CAFS288 vs. Ad-GFP infected cells (right panel). Values represent the mean ± SEM of triplicate samples in three separate transfections. *, Value is different (P < 0.05) from fold change of wild-type 600 mouse GnRHR promoter.
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DISCUSSION
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The TGFß superfamily is important for tissue patterning and development in multiple organ systems including the reproductive axis (43). Specifically, sexual differentiation, primordial germ cell, gonadal, and pituitary development are regulated by TGFß family members (43, 44). In the developing pituitary, signaling gradients of bone morphogenetic proteins (BMPs) activate unique expression patterns of transcription factors that mediate a dorsal to ventral patterning of the hormone-producing cell types (44, 45). Of particular importance to reproductive function is the gonadotrope, a cell type that is uniquely characterized by expression of GnRHRs and production of LH and FSH. The GnRHR mediates the primary stimulatory input to gonadotropes, whereas FSH is essential for later stages of ovarian follicular growth in females and regulation of Sertoli cell function in males (3, 46). Activin is also produced by gonadotropes and is thought to affect changes in FSH and GnRHR expression via autocrine and paracrine mechanisms (26, 47). In the case of FSHß, removal of activin input through follistatin treatment results in a decrease in FSHß mRNA and FSH (17). The production of mice deficient in the activin receptor type II has provided in vivo confirmation for the key role of activin in regulation of FSH synthesis and secretion (48). As with the production of FSH, multiple reports have demonstrated activin regulation of GnRHR expression (26, 27, 28, 29, 30).
Consistent with their role in mediating intracellular signaling by TGFß family members, activin regulation of both the FSH ß-subunit and GnRHR genes is at least partially dependent on Smad proteins (27, 28, 34). In the case of the murine GnRHR gene, the effects of Smads are mediated by binding at the distal end of GRAS (27, 28). However, as with many activin/TGFß-responsive elements, GRAS is a composite regulatory element whose functional activity is dependent not only on Smad binding but also AP-1 and a forkhead DNA binding protein (27, 28). Elimination of any one of these binding components leads to a complete loss of the functional activity of GRAS and activin responsiveness of the GnRHR gene promoter (27). Thus, functional mapping localized activin responsiveness to GRAS. As such, we were surprised to find that the single nucleotide divergence in the rat GRAS homolog could not account for the lack of activin responsiveness of the proximal promoter of the rat GnRHR gene. In fact, an activin-responsive phenotype of the rat promoter only emerged with the inclusion of both intact GRAS and an additional 36 bp of downstream sequence from the mouse GnRHR promoter. This comparative analysis and construction of mouse/rat chimeric promoters provided the key observations leading to the definition of a new regulatory element we have termed DARE. As is the case for GRAS, mutation of DARE leads to a loss of activin responsiveness of the mouse GnRHR gene promoter. Thus, neither GRAS nor DARE alone is sufficient for activin regulation. Rather, this functional phenotype requires both elements and, presumably, a unique configuration of multiple DNA binding proteins to form a complex activin/TGFß-responsive enhanceosome.
The presence of a core TAAT sequence is characteristic of DNA binding sites defined for a number of homeodomain proteins (49). The conservation of this signature motif in the promoter regions disrupted by the two loss of function mutations (BR14 and BR15) that defined the boundaries of DARE raised the possibility that DARE represents a binding site for a homeodomain protein. Consistent with this possibility, DARE was capable of interacting with the DNA binding domain of the LIM-homeodomain protein LHX2 in EMSA. Furthermore, 2-bp scanning mutagenesis established that the activin-responsive properties of DARE are dependent on a contribution of both the distal and proximal TAAT motifs. As such, DARE may serve as a binding site for at least two homeodomain proteins. Alternatively, DARE might represent a binding site for homodimers or heterodimers of paired-like homeodomain proteins; however, the 4-bp separation of the TAAT motifs would not place this element in the more prototypical P2 or P3 binding sites characterized for the bicoid or Pax family of proteins (49). Finally, our mutagenesis studies suggest that the lack of functional activity of the rat DARE homolog is at least partially due to disruption of the more distal TAAT motif located in the BR14 region.
It is important to note that DARE partially overlaps with an element termed SURG-1 (5' CTAATTGGA 3') that was previously defined as contributing to GnRH responsiveness of the GnRHR promoter (38) and, more recently, shown to interact with NF-Y and Oct-1 (40). However, it seems unlikely that these proteins account for the contribution of DARE to activin responsiveness because the BR16 mutation effectively eliminates the binding sites defined for both NF-Y and Oct-1 (40) but has no effect on the follistatin response of the mouse promoter. Thus, although we certainly do not dispute a functional contribution of NF-Y/Oct-1 binding at SURG-1, it would appear that this event is independent of the contribution of DARE to activin responsiveness. In further support of this notion is the fact that the NF-Y/Oct-1 binding site is conserved in the rat promoter; however, substitution of mouse DARE with the rat DARE homolog eliminated activin responsiveness of the mouse promoter.
Our data suggest that both GRAS and DARE participate in conferring activin responsiveness to the proximal promoter of the murine GnRHR gene. As such, this functional phenotype appears to require the formation of a Smad-containing protein complex at GRAS and, potentially, a homeodomain protein complex at DARE. At issue, ultimately, is the mechanism(s) by which these spatially separate complexes cooperate to yield a functional activin/TGFß-responsive enhanceosome. It is certainly possible that components of the GRAS and DARE protein complexes directly interact. Smad proteins are clearly capable of interacting with a wide array of transcription factors including AP-1, forkhead DNA binding proteins, and homeodomain proteins (27, 50, 51). For example, paired-like homeodomain proteins of the Mix family and forkhead transcription factors have been shown to recruit activated Smads to distinct promoter elements (51). Thus, activin-mediated protein-protein interactions may be necessary to assemble the key binding components at GRAS and DARE and, ultimately, recruitment of the appropriate coactivator. If correct, such a mechanism would be strikingly similar to what was recently described for activin signaling to the FSHß subunit gene in the gonadotrope derived LßT2 cell line. Specifically, Bailey et al. (34) reported that activin responsiveness of the ovine FSHß gene promoter is partially dependent on a functional interaction between TALE homeodomain (Pbx1 and Prep1) and Smad proteins. Similarly, activin regulation of the rat FSHß subunit gene requires both Smads and Pitx2, a member of the bicoid-related homeodomain proteins (24).
Although activin is certainly capable of regulating expression of the GnRHR gene (26, 27, 29, 30), Kumar et al. (48) recently reported that GnRHR expression was maintained in mice made deficient in the type II activin receptor gene. As such, it is difficult to reach an unambiguous conclusion as to the precise physiological role of activin in affecting GnRHR gene expression. Several points are, however, important to consider. First, the pituitary phenotype in those embryos in which the knockout was lethal was not reported. Second, there is evidence that type II receptors for TGFß family members can be somewhat promiscuous in their interaction with type I receptors (52, 53). Thus, it is possible that components of activin signaling are retained in the activin receptor II null mice. Third, given that multiple TGFß family members, including BMP2, BMP4, and more recently BMP6 and BMP7, are involved in pituitary morphogenesis and development (45, 54, 55) it is possible that the key inputs to the GnRHR gene include not only activin but also other members of the TGFß superfamily. There is evidence for intrapituitary expression of BMPs and BMP receptors and, in fact, expression of these proteins by gonadotropes themselves (53, 56). At present, the complete array of TGFß family members expressed by the
T31 cell line is unknown; however, follistatin is capable of binding both activin and BMPs (57). Thus, although we have shown that activin alone is capable of increasing transcriptional activity of the murine GnRHR promoter (26), it is possible that the effects of follistatin addition or overexpression are not confined to removing only activin input. As such, it would not seem judicious to dismiss a role for TGFß family signaling in regulating transcriptional activity of the GnRHR gene. For example, the role of both TGFß family signaling and homeodomain proteins in initiating cell-specific gene programs makes the emerging properties of the activin-responsive enhanceosome in the GnRHR gene an intriguing candidate for activation of GnRHR gene expression in the developing pituitary.
In summary, we have taken advantage of the divergence in activin responsiveness between the proximal promoters of the mouse and rat GnRHR genes to identify a new regulatory element in the mouse GnRHR promoter we have termed DARE. Both DARE and GRAS are necessary for activin/TGFß responsiveness of the murine GnRHR gene promoter. We have not yet established the specific identity of the DARE binding components; however, it appears likely that this component(s) represents one or more members of the homeodomain protein family. Thus, GRAS and DARE together define a unique and complex activin/TGFß-responsive enhanceosome whose functional attributes are dependent on the binding of multiple classes of transcription factors at spatially distinct regulatory elements located in the proximal promoter of the murine GnRHR gene.
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MATERIALS AND METHODS
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Materials
T31 cells were generously provided by Dr. Pamela Mellon (University of California San Diego). Dr. Wylie Vale (Salk Institute, La Jolla, CA) provided the adenoviral follistatin (AdCAFS288) and GFP (Ad-GFP) expression vectors (35), whereas purified LHX2 homeodomain and LHX2 antiserum (58) were kindly provided by Dr. Mark Roberson (Cornell University, Ithaca, NY). The NF-Y antibody was obtained from Rockland Inc. (Gilbertsville, PA). Oligonucleotides were obtained from Invitrogen (Carlsbad, CA) and Colorado State University Macromolecular Core Facility (Fort Collins, CO) (see Table 1
for oligonucleotide sequences). DNA sequencing was conducted by Macromolecular Services at the University of California Davis. Restriction enzymes and DNA modifying enzymes were obtained from Fermentas (Hanover, MD) and New England Biolabs (Beverly, MA). Hyperfilm and G-25 Microspin columns were obtained from Amersham Biosciences (Piscataway, NJ). The amplification and purification of Ad-CAFS288 and Ad-GFP were performed as previously described (59).
Transfections and Adenovirus Infections
Culture of
T31 cells and transient transfections using SuperFect (Qiagen, Valencia, CA) were performed as previously described (26, 27), except that 0.8 µg of the test luciferase plasmid and 0.2 µg of RSV-ß-galactosidase transfection efficiency control vector were used and after 3-h incubation of cells with SuperFect/DNA mixture, media was replaced with growth media containing 1000 MOI of AdCAFS288 or the same MOI of an adenoviral construct expressing GFP. Cells were harvested 48 h post infection and assayed for luciferase activity (26, 27). Values were normalized for transfection efficiency by dividing the luciferase activity by ß-galactosidase activity. Within a transfection, all treatments and vectors were tested in triplicate, and all transfections were repeated at least three times using different plasmid preparations. Values are presented as the mean + SEM.
EMSA
EMSA were conducted as previously described (26, 36, 60, 61). Briefly,
T31 nuclear extracts (27, 36) were incubated with Dignam D buffer (20 mM HEPES, 20% glycerol, 0.1 M KCl, 0.4 mM EDTA), polydeoxyinosinic deoxycytidylic acid (2 µg), radiolabeled probe (100,000 cpm), and appropriate unlabeled competitor for 20 min at room temperature. For the NF-Y supershift, the established NF-Y binding site from the bovine LHß-subunit gene (41) served as the radioactive probe, and 2 µg of anti-NF-Y antibody or the same mass of a nonspecific IgG was added after the 20-min incubation and then incubated for an additional 10 min. Complexes were resolved by electrophoresis in pre-run (100 V for 1 h) 5% polyacrylamide gels in 0.5x Tris/glycine buffer (36) and visualized by autoradiography. In Fig. 7
, purified LHX2 homeodomain protein (58) was incubated at 4 C in Dignam buffer D [20 mM HEPES (pH 7.9), 20% glycerol (vol/vol), 0.1 M KCl, 0.2 mM EDTA, 0.5 mM dithiothreitol] with 2 µg of polydeoxyinosinic deoxycytidylic acid for 10 min. After incubation, radiolabeled probe (100,000 cpm) and appropriate amount of unlabeled competitor were added and incubated for 20 min. For supershift of LHX2 homeodomain protein, 1 µl of rabbit anti-LHX2 homeodomain antiserum or NRS was added and incubated for an additional 10 min. Reactions were electrophoresed in Tris-glycine buffer for 22.5 h at 35 mA in 6% polyacrylamide gels. Gels were pre-run at 100 V for 1 h. Gels were transferred to blotting paper, dried, and exposed to Hyperfilm MP for approximately 16 h at 70 C. In all EMSA experiments, oligonucleotides were labeled using polynucleotide kinase and [
-32P]ATP. Double-stranded DNA probes were purified by centrifugation through a G-25 Microspin column.
Vector Construction
The plasmid pMGR-600Luc, with approximately 600 bp of the 5'-flanking region from the murine GnRHR gene fused to the cDNA encoding luciferase in the pGL3-Basic vector (Promega, Madison, WI) (60), and 3XGRAS-Luc (36, 37) have been described previously. An analogous promoter vector for the rat was generated by digesting 1100 rat GnRHR (pLuc1.1GnRH-R BstEII, provided by R. Counis, Université Pierre et Marie Curie, Paris, France) with XbaI/NcoI and ligating into the NheI/NcoI sites of pGL3-Basic to produce pRGR-600Luc.
The 600 vectors containing 1-bp exchange in the GRAS element were constructed by sequential rounds of PCR. Overlapping primers RMGs and RMGas (in which the rat GRAS sequence is replaced by that of the mouse) were used in separate reactions with pGL3 flanking primers GL2 and RV3 and wild-type 1100 rat GnRHR promoter template to generate downstream (RMGs and GL2) and upstream (RGMas and RV3) fragments. Products were gel-isolated and used as template in a second PCR using GL2 and RV3, and the product was subcloned into pGEM-T Easy (Promega). Presence of the mutation was confirmed by sequence analysis (Davis Sequencing, Davis, CA). A 600 promoter fragment of rat GnRHR promoter with murine GRAS was then excised by XbaI/NcoI digestion and cloned into the NheI/NcoI sites of pGL3-Basic (rat-GRAS-repair). A similar approach was performed using MRGs and MRGas and murine 600 GnRHR promoter to generate mouse GnRHR with the rat GRAS sequence, the final product being excised from pGEM-TEz with KpnI/NcoI and subcloned into pGL3-Basic at KpnI/NcoI (prGRAS/MGRLuc).
The 500 exchanges between the 5' end of the mouse and rat promoters were created by reciprocal exchanges of KpnI/MfeI (600/500) and MfeI/NcoI (500/8) promoter fragments of pMGR-600Luc and rat-GRAS-repair vectors, ligated into pGL3-Basic at KpnI/NcoI to generate pr(500)MGR and pm(500)RGR.
For the 250 exchanges, the fusion point selected represents a sequence conserved in both species where half-sites for the blunt-cutting restriction enzymes Ecl136 I and SmaI abut. Upstream fragments (600/250) were generated by PCR using templates pMGR-600Luc and rat-GRAS-repair, with primers RV3 and PIas, which incorporates at its 5' end a full Ecl136 I site. The downstream fragments were created by PCR from pMGR-600Luc (oligonucleotides GL2 and mPIs) and pRGR-600Luc (GL2 and rPIs), incorporating a SmaI site at their upstream end. PCR products were cloned into pGEM-T Easy, and sequence confirmed by sequence analysis. Fragments excised with KpnI and Ecl136 I (upstream) and SmaI/NcoI (downstream), and ligated together into pGL3-Basic, the blunt SmaI and Ecl136 I half-sites combining to regenerate the native sequence, generating pr(250)MGR and pm(250)RGR. The 150 exchanges used a similar approach, only PCR primers PIIas and RV3 (upstream) or PIIs and GL2 (downstream) were used to generate promoter fragments from pMGR-600Luc and rat-GRAS-repair that were cloned into pGEM-T Easy, excised, and joined in pGL3-Basic at compatible BclI and BglII sites derived from the PIIs and PIIas to generate pr(150)MGR and pm(150)RGR.
The 36-bp exchange clones were made by sequential rounds of PCR, as with the GRAS exchanges. Using pMGR-600Luc template, upstream (primers RV3 and SW1as) and downstream (GL2 and SW1s) fragments of the mouse promoter with 36 bp of rat sequence substituted between GRAS and AP-1 elements were generated by PCR. These were gel-isolated and used as template in a second round of PCR, using RV3 and GL2. The same approach generated the analogous rat vector, using rat-GRAS-repair with RV3 and SW2as (upstream) and GL2 and SW2s (downstream). Products of the second round of PCR were subcloned into pGEM-T Easy and transferred to PGL3 using KpnI/NcoI sites to generate pSW1MGRLuc and pSW2mGRAS/RGRLuc.
The block replacement construct pBR13 was described previously (36). Block replacement constructs pBR14, pBR15, and pBR16 were made by combining three pieces. An upstream fragment was generated by PCR using an oligonucleotide that incorporated a HpaI site immediately downstream of the GRAS element (HPAas), using pMGR-600Luc as template. This product was cloned into pGEM-T Easy and excised with SacI/HpaI. A downstream fragment was attained by cutting pSW1MGRLuc (mouse promoter with 36 bp of the rat) with SspI, just upstream of the AP-1 element, and NcoI. The sequence between GRAS and AP-1 was generated by annealing sense and antisense oligonucleotides in which native sequence is replaced with the 8-bp NotI restriction site (BR14s/BR14as, BR15s/BR15as, BR16s/BR16as), and the three promoter pieces were combined by either sequential or concurrent ligation and inserted into SacI/NcoI sites of pGL3-Basic.
The 2-bp mutants (14.15 and 15.14) were made by directly ligating an upstream PCR fragment generated with RV3 and the appropriate downstream primer (14.1n, 14.2n, 14.3n, 14.4n, 14.5n, 15.1n, 15.2n, 15.3n, 15.4n) to the SspI/NcoI fragment of pSW1MGRLuc, described above, cutting with SacI/NcoI and ligating into pGL3-Basic. For the TAAT motif double mutant (14.4/15.4) and mouse-to-rat substitutions (600Rat14.4, 600Rat15.1/2) the analogous upstream PCR product (RV3 to 14.4/5, R/M14.4A, or R/M15.1/2) added an Eco RV half-site. It was cloned into pGEM-T Easy, then excised with SacI/EcoRV, and ligated with the SspI/NcoI fragment of pSW1MGRLuc into pGL3-Basic.
Statistical Analysis
In every transfection, each treatment and vector was analyzed in triplicate, and the experiments were replicated three times using different plasmid preparations. Data are expressed as means ± SEM. Students t test was used to compare the difference between cells infected with Ad-GFP and Ad-CAFS288 within a vector. In Figs. 4
, 5
, and 8
, data were analyzed by ANOVA, and, when the f-test was significant (P < 0.05), means were separated by Tukeys honest significant difference or Duncans Multiple Range Test.
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FOOTNOTES
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First Published Online January 6, 2005
Abbreviations: AP-1, Activator protein-1; BMP, bone morphogenetic protein; DARE, downstream activin regulatory element; GFP, green fluorescent protein; GnRHR, GnRH receptor; GRAS, GnRHR activating sequence; MOI, multiplicity of infection; NF-Y, nuclear factor Y; NRS, normal rabbit serum; SF-1, steroidogenic factor-1; SURG-1, sequence underlying GnRH regulation.
Received for publication May 27, 2004.
Accepted for publication December 27, 2004.
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