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Center for Biomedical Research, Population Council and The Rockefeller University, 1230 York Avenue, New York, New York 10021, USA
(Requests for offprints should be addressed to D J Bernard; Email: dbernard{at}popcbr.rockefeller.edu)
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
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One way in which they meet this challenge is through an acute and robust increase in expression of the ß subunit of dimeric FSH (Fshb), the rate-limiting step in mature hormone synthesis. In rats, increases in Fshb mRNA levels in the pituitary parallel and temporally precede increases in FSH release during the secondary surge (Ortolano et al. 1988, Halvorson et al. 1994). At the same time there is no increase in intra-pituitary FSH content, suggesting that the newly synthesized Fshb mRNA is quickly translated, linked to the glycoprotein hormone 
subunit and constitutively released as mature hormone (Nicol et al. 2004). Arguably, intra-pituitary activins provide the major stimulus for FSH synthesis during this stage of the rodent estrous cycle. Indeed, changes in hormonal milieu (e.g. declines in serum inhibins and intra-pituitary follistatins) following the primary surge provide a permissive environment for locally produced activins to rapidly stimulate increases in Fshb transcription and thereby to generate the secondary surge on estrus morning (Woodruff et al. 1996, Besecke et al. 1997).
Activins selectively regulate FSH production and secretion from gonadotrope cells (Bilezikjian et al. 2004, Gregory & Kaiser 2004). The ligands are members of the TGFB superfamily and come in three forms through the disulfide linking of two related ß subunits: activin A (INHBA-INHBA), activin B (INHBB-INHBB), and activin AB (INHBA-INHBB). Both the Inhba and Inhbb subunit genes are expressed in rat pituitary gland (Halvorson et al. 1994) and in gonadotropes in particular (Roberts et al. 1989), though activin B appears to be the predominant autocrine/paracrine acting form of the ligand in these cells (Corrigan et al. 1991). In primary rat pituitary cultures, recombinant activin A stimulates rapid increases in Fshb mRNA levels and FSH secretion (Carroll et al. 1989, Attardi & Miklos 1990, Weiss et al. 1995), whereas immuno-neutralization of endogenous activin B suppresses FSH release (Corrigan et al. 1991). Moreover, treatment of cycling rats with an activin B monoclonal antibody blunts the secondary surge, strongly implicating the endogenous ligand in the process in vivo (DePaolo et al. 1992).
Like other members of the TGFB superfamily, activins bind to cell-surface receptor serine/threonine kinases (RSK) to produce their effects in target cells (Woodruff 1998). Binding to the type II receptors, ACVR2A or ACVR2B, leads to phosphorylation and activation of the type I receptor, ACVR1B (also known as activin receptor-like kinase 4, ALK4). Recent data suggest that a second type I receptor, ACVR1C (ALK7), may also function as an activin B receptor (Tsuchida et al. 2004). Once activated, both ALK4 and ALK7, which are also RSKs, phosphorylate the intracellular signaling proteins, SMAD2 and SMAD3, on carboxy-terminal serine residues. The SMADs then dissociate from the receptor complex, partner with a co-factor (SMAD4) and translocate to the nucleus where they interact with additional co-regulators to stimulate or repress target gene transcription. Although SMAD-dependent signaling has been most thoroughly characterized, SMAD-independent mechanisms are also important, though these are less well-understood at present and appear to vary between cell types (Attisano & Wrana 2002, Derynck & Zhang 2003, Feng & Derynck 2005).
Whereas activin A can increase Fshb mRNA stability (Carroll et al. 1991), the bulk of its effects on expression appears to be mediated via increases in gene transcription. Indeed, activins can stimulate rapid (immediate-early) increases in Fshb subunit transcription in primary rat pituitary cultures (Weiss et al. 1995) and we observed similar effects in the mouse gonadotrope cell line, LßT2 (Bernard 2004). In this report, we describe part of the molecular pathway through which activins stimulate this response, using LßT2 cells as a model system. The elucidation of this mechanism highlights important species differences in the acute control of FSH synthesis by activins.
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Materials and methods |
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Human recombinant (rh-) activin A and follistatin (FST)-288 were purchased from R&D systems (Minneapolis, MN, USA). Dulbecco’s modified Eagle medium (DMEM) with 4.5 g/l glucose, L-glutamine and sodium pyruvate was from Mediatech (Herndon, VA, USA). Lipofectamine/Plus, Lipofectamine 2000, and gentamycin were purchased from Invitrogen Fetal bovine serum (FBS) was from JRH Biosciences (Lenexa, KS, USA). The anti-SMAD2 (S-20), anti-SMAD3 (FL-425), and anti-SMAD4 (H-552) rabbit polyclonal antibodies were from Santa Cruz Biotechnologies (Santa Cruz, CA, USA). SMAD2 and phospho-SMAD2 rabbit polyclonal antibodies were from Zymed (South San Francisco, CA, USA) and Cell Signaling Technology (Beverly, MA, USA), respectively. The phospho-SMAD3 rabbit polyclonal antibody was a generous gift of Dr Michael Reiss (Robert Wood Johnson Medical School, New Brunswick, NJ, USA). Protease inhibitor tablets (CompleteMini) were purchased from Roche. Aprotinin, leupeptin, pepstatin and phenylmethylsulphonylfluoride (PMSF) were from Sigma. Deoxynucleotide triphosphates (dNTPs), Taq polymerase, and 5 x Passive Lysis Buffer (PLB) were from Promega (Madison, WI, USA).
The –1990/+1mFshb-luc reporter, Smad2 and Smad3 short-hairpin RNAs (shRNAs), constitutively active HA-rat ALK4, Flag-human SMAD2, untagged human SMAD3 Flag-human SMAD3, and Flag-mouse Smad4 expression constructs were described previously (Bernard 2004). Myc-SMAD3 and 6xMyc-SMAD4 were from JJ Lebrun (Royal Victoria Hospital, Quebec, Canada) and Ralf Janknecht (Mayo Clinic, Rochester, MN, USA). Flag-human SMAD2 in pCAGGS was a gift of Dr Elizabeth Roberston (Oxford University, UK). Flag-SMAD3-R74K and –K81R were from Dr C H Heldin (The Ludwig Institute, Uppsala, Sweden). Glutathione S-transferase (GST) -SMAD3 MH1 and GST-SMAD4 MH1 were from Dr. Bert Vogelstein (The Johns Hopkins University School of Medicine, MD, USA).
5' deletion constructs of the mouse Fshb promoter were made using Erase-a-Base reagents following the manufacturer’s instructions (Promega), by PCR or by the sub-cloning of restriction fragments from larger reporters. The SBE mutation (GTCTAGAC to GTCat-GAC) in –1990/+1mFshb-luc and the K88R mutation in Flag-mouse Smad4 were produced by site-directed mutagenesis using the QuikChange protocol (Stratagene).
To generate the human FSHB reporter, salivary genomic DNA was extracted from one of the investigators (DJB) using AquaPure reagents (Biorad) and subjected to PCR using PfuUltra Taq (Stratagene, La Jolla, CA, USA) and the following primer set: (forward) 5'-CGCACGCGTGAACTCAATCAGATCATGTCACGTCACT and (reverse) CGCCTCGAGGAGCTGTAGACTGAATGAAATCTCAGTT. Restriction sites (underlined) were included in the primers to facilitate cloning. The resulting product, corresponding to –1028/+7 of the human FSHB promoter, was digested with MluI/XhoI and ligated into the same sites in pGL3-Basic (Promega) using standard techniques. The 8 base-pair (bp) SBE was introduced into the human promoter using the gene splicing by overlap extension protocol (Horton et al. 1990). Briefly, using the –1028/+7 human reporter as template, two PCRs were performed with the forward primer above and 5'-CTCCAGAGTCTAGACTTTTTCTTTTGTATCTTTAAATCAG and the reverse primer above with 5'-GAAAAAGTCTAGACTCTGGAGTCACAATTAATTTG. The SBE is underlined and italicized in the splicing primers. The resulting amplicons were pooled and a third round of PCR performed with the forward and reverse primers from above. The resulting amplicon was digested with MluI/XhoI and ligated into the same sites in pGL3-Basic. The identities of all constructs were verified by DNA sequencing.
Cell culture and transfection
LßT2 cells were provided by Dr Pamela Mellon (University of California, San Diego, CA, USA) and were cultured as described previously (Bernard 2004). COS7 cells were obtained from Dr Patricia Morris (Population Council, New York, NY, USA) were cultured in DMEM/10%FBS. For the majority of transfection experiments, LßT2 cells were plated in 24-well plates at a density of 2 x 105 cells per well approximately 36 h prior to transfection. For SMAD over-expression studies, cells were plated in 6-well dishes at 1.5 to 2 x 106 cells per well. Cells were transfected with Lipofectamine 2000 (24-well) or Lipofectamine/Plus (6-well) following the manufacturer’s instructions. Reporter plasmids were transfected at 450 ng (24-well) or 1 µg (6-well) per well. Expression plasmids were introduced at 1 µg (6-well) per well. Smad shRNAs were transfected at 100 ng/well at least 24 h prior to ligand treatment. In all experiments, the total amount of DNA transfected was balanced across treatments. COS7 cells were plated in 10-cm dishes and transfected when 70–100% confluent with Lipofectamine and the indicated SMAD expression vectors along with ALK4TD.
Reporter assays
After overnight transfection, LßT2 cells were washed in serum-free DMEM and then treated with 1 nM (25 ng/ml) activin A in DMEM or with DMEM alone (control) for the indicated times. In time-course experiments, the introduction of ligand was staggered so that protein lysates from different treatment groups were collected at the same time. In SMAD over-expression experiments, cells were transfected for 6 h and then fed with growth media. Lysates were collected the following day for luciferase assays. Cells were washed with 1 x phosphate buffered saline (PBS) and lysed in 1 x PLB. Luciferase assays were performed on a Luminoskan Ascent luminometer (Thermo Labsystems, Franklin, MA, USA) using standard reagents. Luciferase values were normalized by protein concentrations determined for the same samples by bicinchoninic acid assay (Pierce). All treatments were performed in triplicate and the data presented are from 2–3 independent experiments.
Electrophoretic mobility shift assays
Nuclear extracts were collected and gel shift experiments were performed as described (Therrien & Drouin 1993), with minor alterations to the protocol. Briefly, following ligand treatment or transfection, cells were washed with 1 x PBS on ice and nuclear extracts collected. Protein concentrations were determined by Bradford assay (BioRad). Nuclear proteins (3–5 µg) were incubated at room temperature with 50 fmol of 5' 32P-ATP (NEN-PerkinElmer) end-labeled double-stranded probes corresponding to –277/–248 of the mouse Fshb promoter in 25 mM HEPES (pH7.2), 150 mM KCl, 5 mM dithiothreitol, 10% glycerol and 500 ng salmon sperm DNA at room temperature for 20 min. Reactions were then run on 5% polyacrylamide gels (44:0.8 acrylamide:bis-acrylamide) in 40 mM Tris–HCl/195 mM glycine (pH 8.5) at 200 volts for 3–5 h at 4 °C. In competition or antibody super-shift experiments, reactions were assembled at room temperature and incubated for 10 min with 500-fold excess unlabeled (cold) competitor or the indicated antibodies prior to the addition of the radio-labeled probe. Gels were dried and exposed to X-ray film.
DNA precipitation
LßT2 cells were plated (1 x 107) in 10-cm dishes. After three days, cells were washed with serum-free DMEM and incubated overnight in the same. The following day, half of the samples were treated with 60 ng/ml activin A for 1 h. Whole cell lysates were then prepared using 300 mM NaCl, 20 mM Tris–HCl (pH 7.5), 1% Triton X-100, 1 mM PMSF, 2 µg/ml leupeptin and aprotinin. One-tenth of each lysate was then used in a DNA precipitation assay with 100 ng biotinylated mouse Fshb SBE probe following published methods (Choy & Derynck 2003). Proteins bound to the probe were purified using streptavidin paramagnetic beads (Promega). Following washes, the proteins were eluted from the beads, separated on 10% NuPage Bis-Tris gels (Invitrogen), transferred to Protran nitrocellulose (Schleicher & Schuell, Keene, NH, USA), and sequentially probed with rabbit anti-phospho-SMAD2, rabbit anti-phospho-SMAD3, and rabbit anti-SMAD4. The details of the western blotting procedures have been described previously (Bernard 2004).
Statistics
The data from replicate experiments were highly similar and were pooled (n = 6 or 9 per treatment) for statistical analyses. Data are presented as fold-change from the control condition (set to 1) in each experiment. In some cases (where indicated), data were log transformed prior to analysis because of unequal variances between treatments. Differences between means were compared using one- or two-way analyses of variance, followed by post-hoc tests where appropriate (Systat 10.2, Richmond, CA, USA). Significance was assessed relative to P < 0.05.
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Results |
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Previously, we showed that activin A stimulates an immediate-early increase in endogenous Fshb transcription in murine LßT2 cells (Bernard 2004). We further showed that a synthetic construct consisting of approximately 2 kb of the mouse Fshb 5' flanking region fused to a luciferase reporter gene (–1990/+1mFshb-luc) was activated by activin A in cell-restricted fashion (i.e. in LßT2 cells, but not in other cell lines; see also (Pernasetti et al. 2001). However, we exposed cells to 20–24 hr of activin A, so it was unclear whether or not the reporter was also acutely (rapidly) regulated. Here, we transfected LßT2 cells with –1990/+1mFshb-luc and treated them with 1 nM activin A for varying time periods. Promoter activity increased significantly with time of exposure to activin A (P < 0.001) and differed significantly from control by 4 h (Fig. 1A
). Reporter activity continued to increase through 30 h, the latest time point measured (data not shown). The promoter-less vector, pGL3-Basic, was not stimulated by brief activin A treatment (Fig. 1B and data not shown).
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To determine cis-regulatory elements mediating the rapid transcriptional response to activin A, LßT2 cells were transfected with mouse Fshb promoter-reporter constructs with progressive truncations from the 5' end and were treated with activin A for 4 h. The results indicated that several promoter regions contributed to the activin A response (Fig. 1B
). All of the reporters were stimulated significantly relative to the promoter-less vector (empty), except for –257/+1. Responses increased progressively 5'of –257, suggesting that several promoter regions (most proximally between –257 and –398) contributed to rapid transcriptional activation of mouse Fshb by activin A.
Rapid activation of the mouse Fshb promoter is Smad-dependent
In other systems, SMAD proteins often broker the rapid actions of activins and TGFBs on transcription through direct DNA binding of SMAD3 and/or SMAD4 to target gene promoters (Yang et al. 2003, Levy & Hill 2005). We observed previously that Smad2 and Smad3 were rapidly phosphorylated and trans-located to the nucleus upon activin A treatment (within 5–10 min) (Bernard 2004), a time frame consistent with their potential roles in immediate-early gene activation. To determine the functional relevance of these proteins, we suppressed endogenous Smad2 or Smad3 protein levels in LßT2 cells by RNA interference (RNAi) using previously validated shRNAs (Bernard 2004). Cells were co-transfected with Smad2 or Smad3 shRNA constructs or empty vector (pBS/U6) and –1990/+1mFshb-luc, and were treated with activin A for 4, 8 or 24 h. As reported previously, both shRNAs inhibited the effects of 24 h activin A (Figs. 2 A, B) (Bernard 2004). Here, we also observed significant antagonism of the activin A responses at 4 and 8 h. Importantly, when correcting for the small (though non-significant) declines in baseline activity, suppression of Smad2 or Smad3 protein levels inhibited activin A-stimulated reporter activity at all time points by 23–27% or 25–43% respectively.
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We screened the activin A-sensitive promoter regions for potential SMAD binding elements (SBE) and observed a perfect 8-bp palindromic element, 5'-GTCTAGAC-3' (Zawel et al. 1998), located at –266/–259 (see Fig. 8A), within the –398/–257 interval (Fig. 1B). This element consists of tandem SMAD boxes, 5'-GTCT-3', in inverted-repeat orientation. To determine whether Smads in LßT2 cells might exert their actions through this element, we first confirmed direct protein binding by gel shift analyses. SMADs 3 and 4, but not SMAD2, bind SBEs through their amino-terminal Mad homology 1 (MH1) domains (Shi et al. 1998, Zawel et al. 1998). We used human GST-SMAD3 MH1 and GST-SMAD4 MH1 fusion proteins and incubated them with a radio-labeled probe corresponding to –277/–248 of the mouse Fshb promoter (hereafter mFshb SBE probe). As predicted, both recombinant proteins interacted directly with the probe (Fig. 3A, lanes 1–3 and 6–8). The abundance of the complexes increased as more recombinant protein was inputted into the assay. In both cases, complexes were disrupted by 500-fold excess unlabeled homologous competitor probe (lanes 4 and 9) and were super-shifted by SMAD-specific antibodies (lanes 5 and 10), confirming that the interactions were specific and that the indicated SMAD proteins were contained within the observed complexes. GST alone did not interact with the mFshb SBE probe (data not shown).
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). Complexes b-e, but not a, were specific and could be competed by incubation with 500-fold excess unlabeled probe (lane 5 and data not shown). One hour of activin A treatment stimulated an increase in the abundance of complexes b and c (lane 4, compare with control cells in lane 2). In addition, overnight incubation of cells with the activin antagonist FST-288 decreased the abundance of these two, but not the other, complexes (lane 3), consistent with previous observations that there is low level endogenous activin (probably activin B) signaling in these cells (Pernasetti et al. 2001). Increased complex abundance could be detected at 30 min post-activin A treatment, peaked at 2–3 h and returned to baseline levels by 24 h (data not shown).
To determine whether or not complexes b and c contained Smads, super-shift experiments were performed with SMAD-specific antibodies (Fig. 3B). Control IgG and anti-SMAD1 (a BMP-regulated SMAD) had no effects on either complex b or c (lanes 6 and 7). In contrast, an anti-SMAD2 antibody disrupted the formation of complex b (lane 8), whereas anti-SMAD3 (lane 9) and anti-SMAD4 (lane 10) antibodies caused super-shifts of both complexes b and c.
To further demonstrate the presence of the indicated Smads in complexes b and c, we over-expressed human SMADs 2 and 3, and mouse Smad4 alone and in combination in heterologous COS7 cells and used nuclear extracts in gel shift analyses with the mFshb SBE probe (Fig. 3B). A complex with similar mobility as complex c in LßT2 cell lysates was detected in control cells transfected with a constitutively active form of the activin type IB receptor (ALK4TD), which generates ligand-like signals in the absence of exogenous ligand (lane 11) (Attisano et al. 1996). Super-shift experiments indicated the presence of endogenous SMAD3 and SMAD4 in this complex (data not shown). Transfection of SMAD2 or Smad4 alone did not significantly alter complex formation (data not shown), whereas the abundance of the complex increased markedly in cells co-transfected with SMAD3 (data not shown, but see Fig. 6). This complex also appeared to include endogenous SMAD4, as it was super-shifted by an anti-SMAD4 antibody (data not shown). The abundance of the complex was further enhanced upon co-transfection of SMADs 3 and 4 (lane 13). Whereas the combination of SMAD2 and Smad4 had no detectable impact on complex formation (lane 12), when the three proteins were co-expressed there was a significant increase in the abundance of two complexes with similar mobility to those of complexes b and c from LßT2 cells (lane 14). An anti-SMAD2 antibody disrupted the formation of complex b from COS7 cells (Fig. 3C, lane 2) and anti-SMAD3 and 4 antibodies super-shifted both complexes (Fig. 3C, lanes 3 and 4). These data were consistent with the observations from LßT2 cells that protein complexes containing Smads 2, 3 and 4 can interact with the SBE at –266/–259 in activin/ALK4-regulated fashion.
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, top two panels). The pSMAD2 antibody detected two proteins in this assay. The smaller of the two likely corresponded to the SMAD2 splice variant, SMAD2
exon3, which we showed previously was expressed in LßT2 cells and could stimulate endogenous mouse Fshb expression (Bernard 2004). Importantly, the amount of Smad4 associating with the biotinylated SBE probe was also increased by activin A (third panel from the top), despite the fact that the total amount of Smad4 in the cell lysates was equivalent in the different treatment conditions (Fig. 3D, bottom panel). Collectively, the gel shift and DNA precipitation data show that activin A stimulated the formation of at least two complexes that could interact with the SBE at –266/–259, one containing pSmad2, pSmad3, and Smad4 (complex b) and the other containing pSmad3 and Smad4 (complex c).
SMADs 2, 3 and 4 synergistically stimulate Fshb transcription
In many systems, over-expression of receptor-regulated (R-) SMADs, like SMADs 2 and 3, can produce effects similar to those of the ligands that activate them, indirectly implicating their functional roles in the underlying signaling pathway (Zhang et al. 1996, Nakao et al. 1997). We employed this approach here to probe the involvement of Smads 2, 3 and 4 in mouse Fshb transcription. To overcome previous difficulties in over-expressing SMAD2 in LßT2 cells (Bernard 2004, Gregory et al. 2005), while avoiding a need to change our culturing conditions (Suszko et al. 2005), we used Flag-tagged full-length human SMAD2 in pCAGGS (Dunn et al. 2004), which contains the chicken ß-actin promoter (Niwa et al. 1991). Using this construct we were able to over-express full-length SMAD2 in LßT2 cells at levels comparable to over-expressed SMAD3 from the weaker CMV promoter (Fig. 4A). As before, exogenous full-length SMAD2 expression from the CMV promoter was comparatively low (compare lanes 2 and 3 to lane 4).
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). SMAD2 alone had no effect, but with SMAD4 was as potent as SMAD3 alone. Remarkably, the combination of SMADs 2, 3 and 4 had very potent transcriptional activity and stimulated the reporter 3 to 5-fold more strongly than SMADs 3 and 4 together. In addition, the combination of SMAD2/3 was as potent as SMAD3/4. These novel data further implicated endogenous Smad2 in transcriptional regulation of the mouse Fshb gene. The SBE is required for activin A- and SMAD2/3/4-stimulated Fshb transcription
We next examined the functional relevance of the SBE for rapid effects of activin A on mouse Fshb transcription. Previous studies showed that a mutation of 2 bp in the center of the SBE of the human SMAD7 promoter, 5'-GTCTAGAC-3' to 5'-GTCatGAC-3', abolished TGFB1-stimulated promoter activity (von Gersdorff et al. 2000). We therefore examined the effects of this 2-bp mutation in the context of the mouse Fshb SBE. Using a radiolabeled wild-type mFshb SBE probe in gel shift analyses, we observed that activin A stimulated the formation of complexes b and c as described earlier (Fig. 5A, compare lanes 2 and 3). A 500-fold molar excess of unlabeled wild-type (WT) probe completely blocked complex formation (lane 4), whereas a competitor probe with the 2-bp mutation (Mut) at positions 4 and 5 of the 8-bp SBE did not (lane 5). Thus, these bp were required for Smad complex binding to the mouse Fshb promoter just as in the SMAD7 promoter.
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). This was particularly true at 4 and 8 h, where activin A failed to significantly stimulate reporter activity from the SBEmut construct. By 24 h, activin A stimulated the mutated promoter, though to a lesser extent than observed with wild-type at 8 or 24 h. Importantly, the same mutation almost completely inhibited the effects of SMAD2/3, SMAD3/4 and SMAD2/3/4 over-expression on mouse Fshb transcription (Fig. 5C). Both SMAD3 and SMAD4 must bind to the SBE for maximal transcriptional activation
We next hypothesized that within the Smad complexes described above, both Smad3 and Smad4 bound to the tandem SBE simultaneously, with each binding to one of the two SMAD boxes (Shi et al. 1998, Chai et al. 2003). We focused on these two Smads because they were present in both complexes b and c in gel-shift analyses (Fig. 3) and because full-length SMAD2 cannot bind SBEs directly owing to a 30 amino acid insertion in its MH1 domain (Dennler et al. 1999, Yagi et al. 1999). If this hypothesis was correct, then disruption of either Smad3 or Smad4 binding should destabilize the protein-DNA interaction and attenuate transcriptional activation. Consistent with this proposition, mutations to single bp within one SMAD box or the other inhibited endogenous Smad complex binding to the SBE (data not shown). To complement these observations, we examined the effects of point mutations in the MH1 ß-hairpins of SMAD3 and SMAD4 on DNA binding and trans-activation of the mouse Fshb promoter.
Conservative substitutions at R74 (R74K) and K81 (K81R) in the ß-hairpin of the MH1 domain were shown previously to inhibit SMAD3-mediated transcriptional activation presumably by disrupting DNA binding, though this was not shown directly (Moren et al. 2000). To confirm that that these mutations indeed disrupt binding, we co-transfected COS7 cells with wild-type (WT) SMAD3, SMAD3-R74K, or SMAD3-K81R alone or in combination with wild-type Smad4. ALK4TD and SMAD2 were co-transfected in all cases. Nuclear extracts were subjected to gel shift analyses with the mFshb SBE probe. SMAD3-WT alone and in combination with Smad4 formed a complex with the DNA probe (Fig. 6, lanes 2 and 5). We could not detect binding of either SMAD3 mutant to the SBE probe when over-expressed alone or with Smad4 (lanes 3, 4, 6, and 7), suggesting that both mutations impaired binding to the SBE. Western blots of the nuclear extracts used in the gel shift analysis indicated comparable levels of pSMAD3 in the different conditions (Fig. 6, bottom panel).
We next examined the functional consequences of these mutations in SMAD3. Wild-type SMAD3, SMAD3-R74K, and SMAD3-K81R were over-expressed alone and in combination with Smad4 in LßT2 cells along with the –1990/+1mFshb-luc reporter. Wild-type SMAD3 stimulated transcription and Smad4 potentiated this effect (Fig. 7A). Neither SMAD3 mutant when expressed alone significantly stimulated reporter activity (Fig. 7A) even though the various forms of SMAD3 were over-expressed at comparable levels (data not shown). Co-expression of Smad4, which had no effect on its own, partially rescued the effect of SMAD3-R74K to approximately 50% of the level seen with wild-type SMAD3/Smad4 and to a level comparable with wild-type SMAD3 expressed alone. There was a trend for Smad4 to partially rescue the effects of SMAD3-K81R, but the fold-stimulation was not as great as wild-type SMAD3 alone or the SMAD3-R74K/Smad4 combination. Stimulation of cells with activin A for approximately 24 h increased transcriptional activation in all cases but did not alter the functional differences seen between the SMAD3 variants (data not shown). Note that the overall level of trans-activation was greater in these experiments than in those described above (Fig. 4B and 5C) likely because the analyses here were in cells from an earlier passage and because less DNA overall was transfected here.
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). However, whereas WT-Smad4 potentiated the effect of over-expressed SMAD3, Smad4-K88R was unable to do so significantly. Further, we observed that Smad4 could partially rescue the trans-activation by SMAD3-R74K (Fig. 7A). We postulated that Smad4 accomplished this by providing some DNA binding ability to the complex, even though it is of reduced affnity relative to complexes with wild-type proteins. Indeed, Smad4-K88R was unable to rescue transcriptional activation by SMAD3-R74K (Fig. 7B). Collectively, the data suggest that both SMADs 3 and 4 must bind directly and simultaneously to the SBE to maximally stimulate mouse Fshb promoter activity. Inter-species differences activin/SMAD activation of Fshb transcription
Finally, we examined whether the mechanisms observed in mice extended to humans. We cloned approximately 1 kb of the human proximal FSHB promoter and compared the nucleotide sequence with that of mouse. The two promoters were highly similar within the first ~330–340 bp (> 72% identity, including gaps) of their respective transcription start sites, but diverged thereafter (Fig. 8A). Interestingly, within the conserved proximal region, an 8-bp gap had to be introduced into the human sequence to facilitate the alignment between the two species and this occurred at exactly the position of the SBE in mouse (Fig. 8A, boxed sequence). Due to the important role of this element in the acute and overall sensitivity of the mouse promoter to activin A (Fig. 5), we hypothesized that the human FSHB promoter might display different activin-response properties. We transfected LßT2 cells with –1028/+7hFSHB-luc or a mouse reporter of similar length, –1195/+1mFshb-luc, and treated them with activin A for 4, 8 or 24 h. Mouse –1195/+1mFshb-luc was stimulated by activin A at all time points (Fig. 8B). In contrast, the human reporter was only significantly up-regulated between 8–24 h. The fold-stimulation observed at 24 h activin A was significantly greater than that observed with the promoter-less vector, pGL3-Basic (data not shown), showing that activin A regulation did depend upon sequences within the human promoter.
To determine whether the lack of the SBE in the human promoter might contribute to its lesser sensitivity to activins, we introduced the 8-bp element into the human reporter (hereafter –1028/+7hFSHB(SBE+)-luc in the location of the gap pictured in Fig. 8A, between bps –276 and –275. Unlike the wild-type promoter, –1028/+7hFSHB(SBE+)-luc responded to activin A within 4 h and was stimulated to a greater extent at 24 h. However, even with this modification, the human promoter was still not as sensitive to activin A as the mouse promoter either acutely (4–8 h) or chronically (24 h). We then compared the sensitivity of the mouse and human promoters to SMAD2/3/4 over-expression. Whereas –1195/+1mFshb-luc was significantly stimulated by SMADs, the wild-type human promoter was not (Fig. 8C). Addition of the SBE to the human promoter conferred SMAD responsiveness, but as with activin A treatment, the overall level of activation was not elevated to that seen in mouse (Fig. 8C).
To determine whether or not the persistent differences between –1195/+1mFshb-luc and –1028/+7hFshb(SBE+)-luc were at least partly attributable to elements upstream of the SBE, we examined the activin A response properties of a truncated mouse promoter (–269/+1mFshb-luc), which contained the SBE and proximal promoter, but lacked more distal elements. This reporter responded to activin A and SMAD2/3/4 more similarly to –1028/+7hFSHB(SBE+)-luc than to –1195/+1mFshb-luc (Figs. 8B and C), suggesting that elements distal to the SBE, which are divergent between mouse and human, contribute to responsiveness of the mouse promoter to activins and SMADs, and this was consistent with results from the 5' deletion experiments (Fig. 1B).
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Discussion |
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SMAD2/3/4 heteromers stimulate mouse Fshb subunit transcription
Previous studies in rat showed that SMADs 3 and 4 could interact with a comparable SBE in the Fshb promoter and that this element was required for SMAD3/4-dependent trans-activation; however, those reports did not show that these interactions were activin-regulated nor did they demonstrate that Smad2 was also contained within protein complexes interacting with this cis-element. In fact, they concluded that Smad2 was not involved in rat Fshb transcriptional regulation (Suszko et al. 2003, 2005, Gregory et al. 2005). Our and others’ previous results implicated SMAD2 in activin A’s stimulation of mouse Fshb transcription and the data presented here more clearly define its role (Dupont et al. 2003, Bernard 2004). First, RNAi experiments show that Smad2 is required for both rapid and chronic effects of activin A on Fshb transcription (Fig. 2A). Second, gel shift and DNA precipitation assays show that activin A stimulates the formation of protein complexes containing pSmad2 that can interact with the SBE (Fig. 3). Third, over-expression experiments indicate that SMAD2 synergizes with SMADs 3 and 4 to stimulate mouse Fshb promoter activity (Fig. 4).
Although the crystal structure of SMAD2/3/4 complexes has not been solved, structures of activated SMAD2:SMAD4 and SMAD3:SMAD4 complexes were determined to be hetero-trimers with stoichiometries of 2:1 (Chacko et al. 2004). Given the high conservation of the trimer interface, however, these structures also predict the assembly of hetero-trimers containing two different R-SMADs. Though we could not definitively determine the precise number of SMAD proteins within the SBE binding complexes for technical reasons, the data we have collected thus far are consistent with the presence of one Smad3 and Smad4 each (data not shown). Based on the available structural data (Chacko et al. 2004), we predict that one Smad2 molecule is similarly contained within the complexes, though additional experimentation is required to show that Smad2/3/4 bind the SBE as hetero-trimers.
The data clearly show that the combination of SMAD2/3/4 has greater transcriptional activation function than that of SMAD3/4. Although we have not yet determined the mechanisms governing their differential effects, there are several possible explanations. For example, SMADs 2, 3, and 4 can all interact with co-activators such as p300/CBP and P/CAF (Feng et al. 1998, Janknecht et al. 1998, Nishihara et al. 1998, Pouponnot et al. 1998, Shen et al. 1998, de Caestecker et al. 2000, Itoh et al. 2000). Therefore, by virtue of having three SMADs in the complex (versus only two), more of these co-activators can potentially be recruited to the mouse Fshb promoter. Alternatively, the overall affinity of activated SMAD complexes for these co-regulatory proteins may be enhanced when SMAD2 is present. It is also possible that SMADs 2 and 3 recruit distinct co-regulators (Chou et al. 2003) and the synergistic activation we observe may be reflective of the combined actions of these different proteins.
SMAD complex binding to the SBE is necessary, but not sufficient for rapid transcriptional activation by activin A
Despite the clear necessity for Smad2/3/4 complexes binding to the SBE at –266/–259, this mechanism does not fully account for rapid actions of activin A on mouse FSHB transcription. First, 5' deletion studies show that cis-elements between –1990 and –399 contribute almost 50% to the response of the –1990/+1reporter to 4 h activin A. The importance of these more distal elements is further underscored by experiments with the human Fshb promoter. The SBE is absent endogenously, but its’ introduction increases both acute (4–8 h) and chronic (24 h) activation by activin A, as well as sensitivity to over-expressed SMADs, but not to levels attained with a mouse promoter-reporter of comparable length (Fig. 8). At the same time, the fold activin A responses of the modified human promoter are very similar to a mouse construct that lacks elements upstream of the SBE (–269/+1). Although highly conserved in their proximal promoters, the two species diverge significantly beyond –330/–340 bp, implicating upstream regulatory elements in activin/Smad responses in mouse. At the same time, the data presented here do not rule out the possibility that differences in the proximal promoters also contribute to inter-species variation in activin A responsiveness.
We have not yet identified the activin-responsive cis-elements within the more distal regions of the mouse promoter or the proteins that interact with them. It seems unlikely, however, that they directly bind Smads because mutation of the proximal SBE almost completely abolishes the effects of over-expressed SMADs (Fig. 5C). Rather, proteins binding more distally may interact with Smads bound to the SBE at –266/–259 to confer greater activin A responsiveness both acutely and chronically. Nonetheless, we cannot reject the possibility that Smads bind to other regions of the mouse Fshb promoter with lower affinity.
Second, some elements within the more proximal promoter may also be required for rapid actions of activins and may interact with Smads. Although we do not observe a statistically significant stimulation of the –257/+1 reporter (which lacks the canonical SBE) by activin A, this region of the promoter does appear to be activated by 4 h relative to the promoter-less vector (Fig. 1B). Within this region of the sheep and mouse promoters two putative Smad boxes (–148/–145 and –115/–112 in mouse) have been described that contribute to activin A-stimulated transcription, though their specific roles in acute regulation were not assessed and no evidence of direct Smad binding to either site was reported (Bailey et al. 2004). We examined the –148/–145 element and observed very low affinity binding of recombinant SMAD3 MH1 and no SMAD4 MH1 binding at all (data not shown). Similarly, using nuclear extracts from control and activin A-treated LßT2 cells we were unable to detect endogenous Smads binding to this element, though specific nuclear protein/DNA complexes were observed (data not shown). Nevertheless, a 2-bp mutation (AGAC to ctAC) decreased, though did not completely block, activin A-stimulated reporter activity at 4 h (data not shown). Thus, regardless of whether or not Smads directly interact with this cis-element, whatever proteins do bind there contributes to acute transcriptional regulation by activins and warrant further investigation.
Third, the formation of Smad2/3/4 complexes and their capacity to bind the SBE is not unique to LßT2 cells (and by extension gonadotropes), though the ability of activins to stimulate Fshb transcription is cell-specific (Pernasetti et al. 2001, Jacobs et al. 2003, Suszko et al. 2003, Bernard 2004). Here, we observe that SMAD2/3/4 complexes that can bind the SBE can be produced through over-expression in COS7 cells or by activin A treatment in T3–1 cells (data not shown), but endogenous Fshb expression and promoter-reporter activity are not observed in these cell lines (Pernasetti et al. 2001, Bernard 2004). Thus, whereas activin-stimulated Smad complexes bind the SBE to acutely regulate transcription, to exert their trans-activation function they appear to do so through interactions with co-factor proteins (Massague & Wotton 2000) and/or by acting in a chromatin environment unique to gonadotrope cells (Yamane et al. 2005). Recently, several nuclear proteins have been implicated in Fshb expression and some appear to participate in activin-regulated transcription (Zakaria et al. 2002, Jacobs et al. 2003, Suszko et al. 2003, Aikawa et al. 2004, Bailey et al. 2004, West et al. 2004). Additional experiments are needed to more clearly define how or if these and other proteins interact with Smad complexes to rapidly regulate transcription.
Acute versus chronic regulation of Fshb transcription by activins
Our data highlight an important distinction in the control of Fshb transcription by activins that has been largely overlooked up to this point; namely that the ligand can both acutely (immediate-early) and chronically (delayed) regulate gene expression and that the underlying mechanisms may be distinct. Whereas acute activin effects in rodents appear to be strongly dependent upon canonical Smad-dependent signaling mechanisms, chronic stimulation may occur somewhat independently of Smads. For example, depletion of Smad2 or Smad3 protein levels by RNAi attenuates but does not completely block effects of 24 h activin A on Fshb transcription (Fig. 2) (Bernard 2004, Suszko et al. 2005). Also, mutation of the SBE in the mouse promoter abrogates the effects of over-expressed SMADs and 4 h activin A, but 24 h activin A can still significantly stimulate transcription (Fig. 5B). Finally, the human FSHB promoter lacks the canonical SBE and is SMAD2/3/4-insensitive, but is stimulated by 24 h activin A (Fig. 8). These latter data suggest that activins chronically regulate the human promoter indirectly, and likely through Smad-independent mechanisms. In mouse, although the SBE and SMADs contribute to both acute and chronic activin A stimulation of promoter activity (Figs. 2, 5 and 8) (Suszko et al. 2003, Bernard 2004, Gregory et al. 2005), Smad-independent signaling, as in human, may contribute to delayed responses. We have not yet determined whether the mechanisms mediating chronic activin A stimulation are conserved between mouse and human.
Nonetheless, the data presented here may provide us with at least part of a molecular explanation for differential patterns of FSH regulation across rodent and human reproductive cycles. As in female rodents, women exhibit concurrent surges of LH and FSH secretion prior to ovulation. In addition, there is a singular rise in FSH levels during the luteal-follicular phase transition. This elevation, like the secondary surge in rodents, is required for ovarian follicular recruitment (Gougeon 1996, Schipper et al. 1998). However, in women, FSH levels begin to rise only 11–12 days after the pre-ovulatory surge and remain elevated for several days before declining during the mid to late follicular phase (Miro & Aspinall 2005). Although the mechanisms controlling singular FSH release in women are not completely understood, existing data suggest a role for inhibins and by extension activins (Welt et al. 2002, Welt 2004, Winters & Moore 2004).
We hypothesize that FSH synthesis and secretion during the follicular phase are chronically, but not acutely regulated because the genetic elements required for rapid, SMAD-dependent actions of activins are not present in the human FSHB promoter (e.g. the SBE and perhaps more distal cis-elements). The lack of the SBE may also contribute to the relatively low levels of FSH during this phase of the cycle (compared with secondary surge levels in rodents) given its contributions to overall activin-sensitivity of the promoter (Figs. 5B and 8B). However, the species-dependent differences we observe may be partly attributable to the fact that LßT2 is a mouse-derived cell line. As a result, co-regulatory proteins that may be needed for human FSHB expression may be lacking in these cells. This concern is somewhat tempered by the observations that the human FSHB gene is both gonadotrope-restricted in its expression and hormonally responsive in the pituitaries of transgenic mice (Kumar et al. 1992, Kumar & Low 1993, 1995). We further hypothesize that Smad-dependent mechanisms observed in rodents may be required because of the unique demands of their short reproductive cycles, specifically the need to rapidly synthesize new FSH for generation of the secondary surge (see Introduction). In contrast, the long time gap between the pre-ovulatory FSH surge and FSH elevation during the luteal-follicular phase transition in women allows ample time to restore depleted intra-pituitary stores, thus obviating a need for acute regulation.
From an evolutionary perspective it is interesting to contemplate whether the loss of the SBE (and other essential cis-elements) precipitated changes in the timing of female reproductive cycles or vice versa. Moreover, it will be important to examine other species to determine: 1) the extent to which acute regulation of the Fshb promoter by activins is related to proximity of the primary and secondary FSH surges, and 2) whether common or distinct signaling mechanisms mediate activin responses across species. Indeed, a comparison of the proximal Fshb promoters in several species, including cow, sheep, pig, human, rat and mouse indicates that the canonical SBE is present only in the rodent genes. Thus, whereas Smads are considered the prototypic signaling proteins for members of the TGFB superfamily, their roles as direct regulators of Fshb transcription may represent a specialization restricted to rodents (and perhaps other species) where the demands of the reproductive cycle require immediate-early gene activation by activins.
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This work was supported by grants from the NICHD to DJB (HD047794 and HD044022). There are no conflicts of interest that would prejudice the impartiality of the presented work.
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Received 3 November 2005
Accepted 18 November 2005
Made available online as an Accepted Preprint 25 November 2005
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 Y. Wang, J. Fortin, P. Lamba, M. Bonomi, L. Persani, M. S. Roberson, and D. J. Bernard Activator Protein-1 and Smad Proteins Synergistically Regulate Human Follicle-Stimulating Hormone {beta}-Promoter Activity Endocrinology, November 1, 2008; 149(11): 5577 - 5591. [Abstract] [Full Text] [PDF]
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N. A. Ciccone, C. T. Lacza, M. Y. Hou, S. J. Gregory, K.-Y. Kam, S. Xu, and U. B. Kaiser A Composite Element that Binds Basic Helix Loop Helix and Basic Leucine Zipper Transcription Factors Is Important for Gonadotropin-Releasing Hormone Regulation of the Follicle-Stimulating Hormone {beta} Gene Mol. Endocrinol., August 1, 2008; 22(8): 1908 - 1923. [Abstract] [Full Text] [PDF]
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P. Lamba, V. Khivansara, A. C. D'Alessio, M. M. Santos, and D. J. Bernard Paired-Like Homeodomain Transcription Factors 1 and 2 Regulate Follicle-Stimulating Hormone {beta}-Subunit Transcription through a Conserved cis-Element Endocrinology, June 1, 2008; 149(6): 3095 - 3108. [Abstract] [Full Text] [PDF]
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D. Coss, C. M. Hand, K. K. J. Yaphockun, H. A. Ely, and P. L. Mellon p38 Mitogen-Activated Protein Kinase Is Critical for Synergistic Induction of the FSH{beta} Gene by Gonadotropin-Releasing Hormone and Activin through Augmentation of c-Fos Induction and Smad Phosphorylation Mol. Endocrinol., December 1, 2007; 21(12): 3071 - 3086. [Abstract] [Full Text] [PDF]
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B. D. Looyenga and G. D. Hammer Genetic Removal of Smad3 from Inhibin-Null Mice Attenuates Tumor Progression by Uncoupling Extracellular Mitogenic Signals from the Cell Cycle Machinery Mol. Endocrinol., October 1, 2007; 21(10): 2440 - 2457. [Abstract] [Full Text] [PDF]
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 L. L. Burger, D. J. Haisenleder, G. M. Wotton, K. W. Aylor, A. C. Dalkin, and J. C. Marshall The regulation of FSHbeta transcription by gonadal steroids: testosterone and estradiol modulation of the activin intracellular signaling pathway Am J Physiol Endocrinol Metab, July 1, 2007; 293(1): E277 - E285. [Abstract] [Full Text] [PDF]
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 K. B Lee, V. Khivansara, M. M Santos, P. Lamba, T. Yuen, S. C Sealfon, and D. J Bernard Bone morphogenetic protein 2 and activin A synergistically stimulate follicle-stimulating hormone {beta} subunit transcription J. Mol. Endocrinol., February 1, 2007; 38(2): 315 - 330. [Abstract] [Full Text] [PDF]
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S. M. McGillivray, V. G. Thackray, D. Coss, and P. L. Mellon Activin and Glucocorticoids Synergistically Activate Follicle-Stimulating Hormone {beta}-Subunit Gene Expression in the Immortalized L{beta}T2 Gonadotrope Cell Line Endocrinology, February 1, 2007; 148(2): 762 - 773. [Abstract] [Full Text] [PDF]
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