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Endocrinology Vol. 147, No. 9 4351-4362
Copyright © 2006 by The Endocrine Society

Activin Inhibits the Human Pit-1 Gene Promoter through the p38 Kinase Pathway in a Smad-Independent Manner

Chantal de Guise, Annie Lacerte, Shahrzad Rafiei, Rachel Reynaud, Melanie Roy, Thierry Brue and Jean-Jacques Lebrun

Hormones and Cancer Research Unit (C.d.G., A.L., S.R., R.R., M.R., J.-J.L.), Department of Medicine, Royal Victoria Hospital, McGill University, Montreal, Quebec, Canada H3A 1A1; and Department of Endocrinology (R.R., T.B.), Hopital de la Timone, 13005 Marseille, France

Address all correspondence and requests for reprints to: Dr. Jean-Jacques Lebrun, Hormones and Cancer Research Unit, Department of Medicine, Royal Victoria Hospital, 687 Pine Avenue West, Montreal, Quebec, Canada H3A 1A1. E-mail: JJ.Lebrun{at}MUHC.McGill.ca.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The pituitary transcription factor Pit-1 regulates hormonal production from the anterior pituitary gland. However, the mechanisms by which Pit-1 gene expression is regulated in humans are poorly understood. Activin, a member of the TGFß superfamily, acts as a negative regulator of cell growth and prolactin gene expression in lactotrope cells. In this study, we show that activin negatively regulates the human Pit-1 gene promoter. We defined a 117-bp element within the Pit-1 promoter that is sufficient to relay these inhibitory effects. We further investigated the signaling pathways that mediate activin-induced inhibition of Pit-1 gene promoter in pituitary lactotrope cells. We found that the activin effects on Pit-1 gene regulation are Smad independent and require the p38 MAPK pathway. Specifically, blocking p38 kinase activity reverses activin-mediated inhibition of the Pit-1 gene promoter. Together, our results highlight the p38 MAPK pathway as a key regulator of activin function in pituitary lactotrope cells and further emphasizes the critical role played by activin in regulating hormonal production in the pituitary gland.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE PITUITARY GLAND is the primary site of the synthesis, storage, and release of hormones that play a predominant role within the entire human body, and thus, careful regulation of these hormonal levels is essential to maintain organism homeostasis (1). Pituitary tumors account for 15% of intracranial tumors clinically diagnosed (1). The most common type of pituitary adenomas are prolactinomas, tumors of the anterior pituitary prolactin-secreting lactotrope cells (2). Prolactinomas are associated with the release of excess amounts of prolactin hormone (hyperprolactinemia) giving rise to severe clinical syndromes like galactorrhea and gonadal dysfunction leading to infertility (1, 2, 3). To lower serum prolactin levels, therapy with dopamine agonists is currently used for the treatment of hyperprolactinemia (4, 5). However, not all patients are responsive to dopamine agonists, and alternatives need to be developed to suppress hormone hypersecretion and reduce pituitary tumor size without compromising normal pituitary functions. Thus, a better understanding of the signaling pathways that regulate pituitary hormonal production and cell growth is critical to patients with pituitary tumors.

Pituitary organogenesis and maintenance, as well as hormonal production, are under the control of Pit-1, a critical pituitary-specific transcription factor. Pit-1 is required to direct the pituitary-specific expression of prolactin (6, 7, 8, 9), GH (10, 11), and TSH genes (12). Moreover, Pit-1 expression is tightly self-regulated through direct interactions with its own promoter (13). This process of autoregulation of the Pit-1 gene is conserved among rats and humans. The rat Pit-1 promoter contains cAMP-responsive element, which mediates the regulatory effects of glucocorticoids (14), retinoic acid, and thyroid hormone (15). However, these regulatory mechanisms are not conserved in humans, and little is known about regulation of the human Pit-1 gene. The human Pit-1 promoter contains several Pit-1 DNA-binding sites, an octamer-binding site, and a 12-O-tetradecanoylphorbol-13-acetate-responsive element that contribute to Pit-1 gene regulation at the transcriptional level (16). We recently demonstrated that activin and TGFß potently inhibit cell growth and prolactin expression in rat pituitary lactotrope cells (17, 18). Furthermore, our results indicated that activin inhibits these effects on prolactin gene expression through repression of the transcription factor Pit-1 (18). In this study, we investigated the mechanism by which activin regulates human Pit-1 expression.

Activin, a member of the TGFß superfamily of growth factors (19, 20), was initially isolated from gonadal fluid based upon its ability to stimulate FSHß secretion from pituitary gonadotropes (21). Subsequently, activin was shown to regulate cell growth, apoptosis, and differentiation in a large variety of tissues (19). Activin signaling is initiated by ligand binding to two transmembrane-spanning activin type II receptors at the cell surface. This leads to the recruitment and phosphorylation of the activin type I receptor (ALK4) (19, 20). Once activated, ALK4 phosphorylates the main downstream intracellular mediators Smad2 and Smad3 upon two C-terminal serine residues (SxS motif) (22). Once phosphorylated, Smad2 and Smad3 associate with the common-partner Smad4 and translocate to the nucleus where they interact with several transcription factors, coactivators, or corepressors to regulate expression of target genes in a cell- and tissue-specific manner. Although the Smad pathway represents the canonical signaling pathway used by activin and TGFß, other intracellular cascades are known to mediate signaling by these growth factor receptors. In particular, the MAPKs, including ERKs (23, 24), c-Jun N-terminal kinases (JNKs) (25, 26), and p38 kinases (27, 28, 29), have been shown to act downstream of the TGFß receptor complex. Activation of these distinct signaling pathways leads to both Smad-dependent and Smad-independent responses in a cell- and tissue-specific manner (30).

In the present study, we show that activin represses the human Pit-1 gene promoter. We defined a minimal 117-bp element within the Pit-1 gene promoter that is sufficient to relay these inhibitory effects. Our results also indicate that activin regulates Pit-1 expression in a Smad-independent manner. Furthermore, we show that the activin inhibitory effects on Pit-1 expression require the p38 MAPK pathway. Thus, defining the mechanisms by which human Pit-1 expression is regulated will help open new avenues to the development of alternative treatment to prolactinomas.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture
Rat somatolactotrope GH4C1 and Chinese hamster ovarian CHO cells were cultured in DMEM (Hyclone Laboratories, Inc., Logan, UT) in the presence of 10% fetal bovine serum (FBS) (Hyclone), 2 mM L-glutamine (GIBCO, Grand Island, NY), and 5% penicillin/streptomycin antibiotics (GIBCO). All cell lines were incubated at 37 C in an atmosphere of 95% air/5% CO2. For transfection experiments in 24-well plates, GH4C1 cells were seeded at 2 x 105 per well in 1 ml of the appropriate growth medium, and 2 x 106 GH4C1 cells were plated in 10-cm2 petri dishes. CHO cells were used at 2 x 105 per well of 12-well plates for transfection experiments.

Mutagenesis
Mutagenesis of the human Pit-1 promoter has been performed by deletion of six nucleotides of the Pit-1 binding site of the 102/15Luc promoter construct using the QuikChange mutagenesis kit from Stratagene (La Jolla, CA). Oligonucleotides used for the mutagenesis were 5'-GGGAAAAGACTATTAACAAAGGGATTTCCTTGCAG-3' and 5'-CTGCAAGGAAATCCCTTTGTTAATAGTCTTTTCCC-3'. Mutant constructs were confirmed by sequencing and binding efficiency using electrophoretic mobility shift assays. The dominant-negative (DN)Smad2/3 forms were generated by mutation of the C-terminal serine residues within the motif SXS to alanine. Oligonucleotides used for mutagenesis were DNSmad3, 5'-TTACGAATTCATGGACTACAAAGACGACGACGACAAATCGTCCATCCTGCCCTTCACC-3' and 5'-ATTGCTCGAGCTAAGCCACCGCGGAACAGCGGATGCTGGGG-3'; DNSmad2, 5'-TTACGAATTCATGGACTACAAAGACGACGACGACAAATCGTCCATCTTGCCATTCACT-3' and 5'-ATTGCTCGAGTTAGGCCATGGCTGAGCATCGCACTGACGGG-3'. ALK4mL45 was generated by the introduction of three alanines in positions N265, D267, and N268 within the L45 loop of ALK4 as previously described for the type I TGFß mutant receptor (31). Oligonucleotides used for mutagenesis were 5'-GCTGCTGACAATAAAGCCGCTGGCGCCTGGACCCAGCTG-3' and 5'-CAGCTGGGTCCAGGCGCCAGCGGCTTTATTGTCAGCAGC-3'. The fidelity of the mutant constructs was confirmed by sequencing.

Transfection and reporter assays
For luciferase assays in GH4C1 cells, 0.5 µg of the different promoter constructs (human 1321/15Luc, 601/15Luc, 102/140Luc, 102/15Luc, and 3TPLuc) were cotransfected with 0.2 µg of a ß-galactosidase expression vector with lipofectamine and PLUS Reagent (Invitrogen, Carlsbad, CA) in the presence or absence of 0.5 µg of various DNSmad expression plasmids or ALK4 cDNA. Cells were trypsinized 1 d after transfection, split in two, allowed to recover, and serum starved with or without activin (0.5 nM) or TGFß (100 pM) for 18 h. When pretreatment of cells with inhibitors was required, kinase inhibitors were diluted in starvation media and added to the cells 2 h before activin A or TGFß stimulation. Cells were then washed with PBS (pH 7.4) and lysed on ice in 100 µl extraction buffer [1% Triton X-100, 15 mM MgSO4, 4 mM EGTA, 1 mM dithiothreitol (DTT), 25 mM glycylglycine (pH 7.8)] per well of a 24-well plate. The luciferase activity of each sample was measured using 45 µl cell lysate (luminometer from EG&G Berthold, Bad Wildbad, Germany) and normalized to the relative ß-galactosidase activity. CHO cells were transfected using the calcium phosphate method. Briefly, 2 µg of each of the different reporter constructs and 1 µg ß-galactosidase expression vector were transfected using calcium chloride 2.5 mM and a 2x BES (N,N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid)-buffered saline calcium phosphate solution (pH 6.96) (50 mM BES, 280 mM NaCl, and 1.5 mM Na2HPO4). The following day, cells were split, allowed to recover, and serum starved with or without 0.5 nM activin for 18 h. Luciferase assays were performed as described for GH4C1 cells.

Western blot analysis
For short time courses (0–90 min), GH4C1 cells were seeded at 1 x 106 cells per well in six-well dishes in DMEM, 10% FBS. The following day, cells were starved overnight and stimulated or not with 0.5 nM activin A or 100 pM TGFß for different periods of time. Fresh medium was added before each time of stimulation. When pretreatment of cells was required, kinase inhibitors (p38 inhibitors SB202474, SB202190, and PD169316; MEK1/2 inhibitor PD98059; and JNK II inhibitor SP600125) (Calbiochem, EMD Biosciences Inc., San Diego, CA) were diluted in starvation media and added to the cells 2 h before activin A or TGFß stimulation. Kinase inhibitors were prepared in dimethyl sulfoxide (DMSO) (Burdick & Jackson, Muskegon, MI), and controls were done by pretreatment of the cells with equal amounts of DMSO. Cells were lysed on ice with lysis buffer (50 mM HEPES, pH 7.5; 150 mM sodium chloride; 100 mM sodium fluoride; 10 mM sodium pyrophosphate; 5 mM EDTA, pH 8.0; 10% glycerol; 0.5% Nonidet P-40; 0.5% sodium deoxycholate) supplemented with protease inhibitors 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 2 µg/ml pepstatin (BioShop, Burlington, Ontario, Canada), and 1 mM sodium orthovanadate activated with 1% hydrogen peroxide (Sigma-Aldrich, St. Louis, MO). Total cell lysates were vortexed 5 min at 4 C and debris pelleted by centrifugation (140,000 rpm at 4 C for 15 min). Total cell extracts were then separated on polyacrylamide gels, transferred onto nitrocellulose, and incubated with the indicated antibody overnight at 4 C [phospho-p38 (no. 9211), p38 (no. 9212), pERK1/2 (p42/p44) (no. 9106), ERK1/2 (p42/p44) (no. 9102), pJNK/SPAK (no. 9251), and JNK/SPAK (no. 9252) from Cell Signaling Technology, Beverly, MA; Smad2/3 (Sc-8332) and STAT3 (Sc-482) from Santa Cruz Biotechnology, Inc., Santa Cruz, CA; phospho-Smad3 (no. 44-246G) from BioSource International, Camarillo, CA; and phospho-Smad2 antibody, a gift from Dr. Moustakas]. After incubation, the membranes were washed twice for 15 min in washing buffer (50 mM Tris base, pH 7.5; 200 mM sodium chloride; 0.05% Tween 20) and incubated with an appropriate peroxidase-linked secondary antibody (from Santa Cruz; at a 1/10,000 dilution) for 1 h at room temperature. Then, the membranes were washed four times for 15 min, and immunoreactivity was revealed by chemiluminescence (Lumi-Light Plus Western blotting substrate; Roche Molecular Biochemicals, Indianapolis, IN) according to the manufacturer’s instructions. An Alpha Innotech Fluorochem imaging system (Fisher Scientific, Ontario, Canada) was used for the revelation. Proper expression of transfected cDNAs (ALK4wt and ALK4mL45) were assessed by immunoblot using the same cell lysates as used for the luciferase assays and was revealed with a specific anti-ALK4 antibody (gift from Dr. W. Vale).

EMSA
GH4C1 (2 x 106 cells/ml) were seeded in DMEM containing 10% FBS, starved overnight, and stimulated with 0.5 nM activin A for 0–90 min. Nuclear extracts were prepared with a low-salt lysis buffer, buffer A (10 mM HEPES, pH 7.9; 10 mM KCl; 0.1 mM EDTA; 0.1 mM EGTA; 1 mM DTT; supplemented with protease inhibitors 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 2 µg/ml pepstatin; BioShop). For total cell lysates, cells were lysed in 25 µl of 10% Nonidet P-40, and cytoplasmic extracts were centrifuged (14,000 rpm at 4 C for 15 min.). Nuclear pellets were resuspended in a high-salt lysis buffer, buffer C (20 mM HEPES, pH 7.9; 0.4 M NaCl; 1 mM EDTA; 1 mM EGTA; 1 mM DTT; supplemented with protease inhibitors), vortexed 30 min at 4 C, and centrifuged (14,000 rpm at 4 C for 15 min.). Nuclear extracts (5 µg) were incubated with the 32P-labeled probe corresponding to a region of the human Pit-1 promoter that contains the Pit-1-positive autoregulatory binding site (annealed oligonucleotides 5'-AGACTATTAACATGTATAAAGGGATTTCCT-3' and 5'-AGGAAATCCCTTTATACATGTTAATAGTCT-3') for 30 min at room temperature in a binding buffer mix (50% glycerol; 1 M Tris, pH 8.0; 5 M NaCl; 1 M MgCl2; 1 M DTT; supplemented by 1x BSA and 0.025 µg/µl salmon sperm). Extracts were resolved on a 5% nonreducing polyacrylamide gel [prerun in 0.5x Tris-borate-EDTA running buffer for 2 h at 100 V (45 mM Tris, pH 8.0; 45 mM boric acid; 1 mM EDTA, pH 8.0)]. For supershift experiments, 1 µg antibody (anti-Pit-1 Sc-442) from Santa Cruz was added to protein extract 1 h on ice before hybridization.

Statistical analysis
Results are expressed as mean ± SD of three or more separate independent experiments in triplicate. Statistical analysis was assessed by one-way ANOVA or the unpaired t test, as indicated in figure legends, using GrapPad Prism 4 software (GraphPad Software, Inc.). Statistical analyses were meant to compare fold induction (percentage of control) of TGFß/activin-treated samples among themselves within each experiment. Additional post-ANOVA tests were performed when necessary to 1) define linear trend of the dose responses (see Fig. 1Go, post-test for linear trend, GraphPad Prism 4) and 2) compare all data with TGFß-treated control (see Fig. 5Go, A–C, Dunnett’s test GraphPad Prism 4). For all statistical analyses and tests, a P value < 0.05 was considered significant and is indicated on the top of the error bars by an asterisk.


Figure 1
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  		FIG. 1. Activin/TGFß down-regulates Pit-1 gene promoter activity in a dose-dependent manner. GH4C1 cells were transiently transfected with 0.5 µg of the Pit-1 reporter construct –1321/+15Luc together with 0.2 µg of the ß-galactosidase expression plasmid. Cells were treated with increasing concentrations of activin (A) or TGFß (B) for 18 h before being harvested. Luciferase assays were performed and normalized according to the ß-galactosidase activities. Three independent and separate experiments were performed in triplicate. For statistical analysis, ANOVA followed by post-test for linear trend were performed and indicated P = 0.0002 for activin and P < 0.0001 for TGFß.

 

Figure 5
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  		FIG. 5. The Smad pathway is not required for activin/TGFß inhibition of the Pit-1 gene promoter. GH4C1 cells were transiently cotransfected with 0.5 µg of the Pit-1 –102/+15Luc promoter construct or 0.5 µg of the 3TPLuc reporter construct, 0.2 µg of the ß-galactosidase expression plasmid, and 0.5 µg of the different Smad expression plasmids DNSmad2, DNSmad3, and Smad7 (A and B) and ALK4wt or ALK4mL45 (C) as indicated. Cells were then stimulated or not with 0.5 nM activin and assessed for luciferase activity. Transfection of the 3TPLuc reporter construct with the various expression cDNA serves as control to confirm efficiency of these expression plasmids in regulation of activin signaling. Cell lysates were loaded on polyacrylamide gels, transferred onto nitrocellulose, and incubated with the indicated specific antibody to control the expression level of protein. Three independent and separate experiments were performed in triplicate. For statistical analysis, ANOVA (A and B) or unpaired t test (C) were performed: *, P < 0.05 was considered significant.

 

   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Activin and TGFß inhibit the human Pit-1 gene promoter
We recently showed that activin inhibits Pit-1 expression in GH4C1 pituitary cells (18). To determine whether activin regulates Pit-1 at the transcriptional level, we examined the effect of activin on the human Pit-1 gene promoter. For this, we used a reporter construct composed of 1.3 kb of the 5' regulatory sequence of the human Pit-1 fused to the luciferase gene (–1321/+15Luc) (16). This reporter construct was transiently transfected into GH4C1 pituitary cells. GH4C1 are somatolactotrope cells that maintain cell-specific functions and retain the capacity to synthesize and secrete high prolactin levels in a hormone-regulated manner (32, 33). We used this GH4C1 cell line derived from rat pituitary tumor cells as an in vitro model of pituitary tumors (34) because no human pituitary cell line models are available. As shown in Fig. 1AGo, activin significantly inhibited Pit-1 gene promoter activity in a dose-dependent manner. This effect is not restricted to activin, because TGFß was also able to inhibit Pit-1 gene promoter activity in a dose-dependent manner (Fig. 1BGo). These results suggest that members of the activin/TGFß superfamily of growth factors exert a transcriptional effect on Pit-1 gene expression.

A 117-bp fragment of the Pit-1 gene promoter is sufficient to confer the activin/TGFß response
Regulation of the rodent Pit-1 gene transcription has been well studied over the last decade (13, 35, 36). However, other than the binding sites for Pit-1 itself, no other regulatory binding sites identified in rat are conserved in the human Pit-1 promoter. To further explore the promoter sequences mediating activin and TGFß down-regulation of the human Pit-1 gene promoter, we used various 5'-deletion mutants of the (–1321/+15Luc) promoter construct in luciferase assays (Fig. 2AGo). As shown in Fig. 2BGo, our results indicated that activin induced a 50% decrease in luciferase activity with all reporter constructs, including the shorter form (–102/+15Luc). Similar results were obtained when GH4C1 cells stimulated with TGFß, confirming our findings (Fig. 2CGo). These results suggest that the activin-responsive elements are located within the proximal region of the Pit-1 gene promoter between bases –102 and +15.


Figure 2
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  		FIG. 2. A 117-bp fragment of the Pit-1 gene promoter is sufficient to confer the activin/TGFß response. A, Schematic representations of the human Pit-1 promoter constructs. B and C, GH4C1 cells were transiently transfected with 0.5 µg of the indicated truncated human Pit-1 promoter constructs and 0.2 µg of the ß-galactosidase expression plasmid. Cells were treated or not with 0.5 nM activin (B) or 100 pM TGFß (C) for 18 h before being harvested. Luciferase assays were performed and normalized according to the ß-galactosidase activities. Three independent and separate experiments were performed in triplicate. D, Alignments of the conserved Pit-1 promoter sequences from various species. E, Cells were transfected with the mutant Pit-1 promoter, treated with 0.5 nM activin, and assessed for luciferase activity as described previously. Three independent and separate experiments were performed in triplicate. F, Nuclear extracts from GH4C1 cells stimulated with 0.5 nM activin for the indicated periods of time were incubated with a 32P-labeled Pit-1 probe and separated by EMSA (lanes 1–6). In lane 7, a specific anti-Pit-1 antibody was added 1 h before hybridization. The gel was analyzed by autoradiography. G, GH4C1 cells were transfected with various Pit-1 promoter deletion constructs (top) or mutant constructs (bottom), as indicated, treated with 0.5 nM activin, and assessed for luciferase activity as described previously. Three independent and separate experiments were performed in triplicate, and results are expressed in relative luciferase units.

 
The –102/+140Luc construct contains an additional Pit-1-binding site located in the 5'-untranslated region of the gene. This Pit-1-binding site was previously shown to be inhibitory in GH3 cells (16). GH3 is a differentiated neuroendocrine cell line from which GH4C1 cells were derived. However, the Pit-1 inhibitory binding site located between +15 and +40 after the transcriptional start site is not required for the negative regulation of Pit-1 by activin/TGFß, because its removal did not alter the activin/TGFß response (Fig. 2Go, B and C). Together, these results indicate that a 117-bp fragment (–102/+15Luc) of the human Pit-1 gene promoter is sufficient to confer activin/TGFß responsiveness.

Comparison of the 117-bp region of the Pit-1 gene promoter among species reveals a high sequence similarity (86%). As shown in Fig. 2DGo, this suggests that this may represent a major regulatory site for Pit-1 gene expression. This region contains a well-conserved Pit-1-binding site that was previously shown to be critical for Pit-1 to stimulate its own promoter in both rodents (13) and humans (16). To determine whether the activin inhibitory effects on the Pit-1 gene promoter were mediated through modulation of Pit-1 binding to its own autoregulatory site, a deletion mutant was generated in the –102/+15Luc promoter construct (102/15Pit+Del) (Fig. 2DGo). Interestingly, activin was still able to inhibit Pit-1 gene promoter activity of this promoter construct in GH4C1 cells (Fig. 2EGo), suggesting that negative regulation of Pit-1 gene expression by activin is independent of the Pit-1 autoregulatory site. These results were further confirmed by EMSA with nuclear extracts of GH4C1 cells treated or not with activin for various periods of time as indicated in Fig. 2FGo. Using the Pit-1-binding site of the –102/+15Luc promoter construct as a probe, no decrease in Pit-1 binding to its positive autoregulatory site was observed in response to activin treatment (Fig. 2FGo, lanes 1–6). A supershift experiment was performed using a specific anti-Pit-1 antibody and confirmed the presence of the transcription factor Pit-1 in the retarded DNA/protein complex (Fig. 2FGo, lane 7). Together, our data indicate that the activin effect on Pit-1 repression is not mediated through modulation of Pit-1 binding to its autoregulatory site.

Activin effects on Pit-1 gene repression are pituitary cell specific
Pit-1 is specifically expressed in the pituitary; thus, we investigated whether the activin effects on the Pit-1 promoter expression were pituitary cell specific. To assess the specificity of the regulation by activin, the various human Pit-1 promoter constructs described above were transiently transfected in CHO cells that do not express Pit-1 protein. As shown in Fig. 3AGo, activin failed to inhibit Pit-1 gene expression in all human Pit-1 gene promoter constructs. As a positive control, and to demonstrate the activin responsiveness of these cells, CHO cells were transfected with a known activin-responsive promoter construct (3TPLuc) (37). As shown in Fig. 3BGo, activin strongly induced 3TPLuc activity. Overexpression of Pit-1 cDNA in CHO cells increased the basal luciferase activity of the Pit-1 gene promoter (102/+15Luc) but did not change or affect the activin inhibitory effect on this promoter (data not shown). These results indicate that activin regulation of the human Pit-1 gene promoter expression is pituitary cell specific and is not mediated through alteration of Pit-1 autoregulation.


Figure 3
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  		FIG. 3. Activin-dependent down-regulation of Pit-1 promoter expression is pituitary cell specific. A, CHO cells were transiently transfected with 2 µg of the different human Pit-1 promoter constructs together with 1 µg of the ß-galactosidase expression plasmid. The activin response was measured by luciferase assay 18 h after stimulation. B, As positive control for the activin response of those CHO cells, they were transiently transfected with 0.5 µg of the 3TPLuc reporter construct and 0.2 µg of the ß-galactosidase expression plasmid and stimulated with 0.5 nM activin for 18 h before luciferase assay was performed. Three independent and separate experiments were performed in triplicate. C, CHO cells were transfected with various Pit-1 promoter deletion constructs or empty vector, as indicated, treated with 0.5 nM activin, and assessed for luciferase activity as described previously. Three independent and separate experiments were performed in triplicate, and results are expressed in relative luciferase units.

 
Activin/TGFß activates the Smad pathway in pituitary cells
Activin and TGFß biological effects are mediated through different intracellular signaling pathways. Among these, the Smad pathway represents the canonical pathway downstream of activin/TGFß receptors. Activation of this pathway is determined by the ability of the type I receptor to phosphorylate R-Smads after ligand stimulation. The effects of activin/TGFß on the Smad pathway were analyzed in GH4C1 cells. For this, GH4C1 cells were stimulated with activin or TGFß for different periods of time, as indicated in Fig. 4Go, and total cell lysates were analyzed by Western blotting using specific antibodies to phospho-Smad2 and phospho-Smad3. These antibodies recognize the two phosphorylated serine residues in the C-terminal end of the MH2 domain of Smad2/3 (SXS). As shown in Fig. 4Go, A and B, activin and TGFß treatment of GH4C1 cells led to a clear increase in endogenous Smad2 and Smad3 phosphorylation levels in a time-dependent manner (top panels). The membranes were stripped and reprobed with anti-Smad2/3 or anti-Stat3 antibodies for loading controls (Fig. 4Go, A and B, bottom panels). These results indicate that the Smad pathway is functional in GH4C1 cells and is activated in response to activin or TGFß stimulation.


Figure 4
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  		FIG. 4. Activin/TGFß induces Smad2/3 phosphorylation in GH4C1 cells. GH4C1 cells were starved overnight and treated with 0.5 nM activin (A) or 100 pM TGFß (B) (0–90 min). Whole-cell lysates were analyzed by Western blot using specific antibodies against the phosphorylated form of Smad2 or Smad3. Membranes were stripped and reprobed with an anti-Smad2/3 antibody and an anti-STAT3 antibody as a loading control.

 
The Smad pathway is not required for activin/TGFß inhibition of the Pit-1 gene promoter
Having established that activin/TGFß induces phosphorylation of both Smad2 and Smad3 (Fig. 4Go) in GH4C1 cells, we then investigated the relative contribution and involvement of the Smad pathway to activin-mediated inhibition of the human Pit-1 gene expression. For this, we specifically blocked Smad2 or Smad3 signaling using cDNAs encoding for DNSmad2 and DNSmad3 (38). These DN forms of the Smads were generated by mutating the C-terminal serine residues to alanine. As a control, we also used the cDNA encoding for the inhibitory Smad7, which antagonizes activin/TGFß signaling through direct interactions with the activated receptor kinases, thereby preventing Smads access to receptors (39) and by also promoting ubiquitination and degradation of the activated receptor complex (40, 41). Thus, Smad7 is a potent inhibitor of both Smad-dependent and Smad-independent responses.

GH4C1 cells were transiently cotransfected with the –102/+15Luc gene promoter construct and the cDNA encoding for Smad7, DNSmad2, or DNSmad3. Interestingly, whereas overexpression of Smad7 fully reversed the activin inhibitory effect on the Pit-1 gene promoter, overexpression of DNSmad2 or DNSmad3 did not significantly affect activin’s inhibitory effect (Fig. 5AGo). Similar results were obtained with TGFß treatment (data not shown). This suggests that Smad2 and Smad3 are not required for activin/TGFß inhibition of the human Pit-1 gene promoter. To assess the efficiency of our DNSmad constructs in blocking the activin effect, we performed a parallel experiment in GH4C1 cells, in which the DNSmad2 or DNSmad3 cDNAs were cotransfected with the 3TPLuc reporter construct. As seen in Fig. 5BGo, although activin strongly induced 3TPLuc activity in GH4C1cells, this effect was completely reversed in the presence of DNSmad2 or DNSmad3, and Smad7, further validating the efficiency of the mutant Smad constructs.

To confirm that activin-mediated inhibition of the Pit-1 gene promoter is Smad independent, we generated a mutant form of ALK4 in which we mutated the interaction site with the receptor-regulated Smad2 and Smad3. For this, three critical residues located within the L45 loop of the receptor (N265, D267, and N268) were mutated to alanine. A similar mutant construct was previously generated for the TGFß type I receptor (ALK5), leading to inactivation of Smad-dependent TGFß signaling (31). The ability of the activin type I mutant receptor (ALK4mL45) to block Smad-dependent activin responses was first assessed using the 3TPLuc reporter construct. As shown in Fig. 5CGo (left), activin induced a 40-fold induction of the luciferase activity in GH4C1 cells overexpressing the wild-type receptor (ALK4wt). However, this effect was greatly reduced in cells overexpressing ALK4mL45, indicating that the mutant form of the activin receptor, which has lost its ability to interact with the Smads, is unable to transduce activin-mediated 3TPLuc activity. We then tested the mutant receptor with the Pit-1 gene promoter, and as shown in Fig. 5Go (right), overexpression of either ALK4wt or ALK4mL45 led to inhibition of the Pit-1 gene promoter to the same extent, in response to activin treatment. Proper expression of wild-type or mutant ALK4 cDNA was assessed by immunoblot using a specific anti-ALK4 antibody and showed equal expression of the receptors in all lanes (Fig. 5CGo, bottom panels). Combined with our previous results, this indicates that activin-mediated inhibition of Pit-1 gene expression is activin receptor dependent but Smad independent.

Activin/TGFß activates the p38 MAPK pathway in GH4C1 cells
In addition to the canonical Smad pathway, activin/TGFß receptors have been reported to activate other pathways, including several MAPKs, such as ERKs (23, 24), JNKs (25, 26), and p38 kinases (28, 29, 30). Although activation of these pathways by activin/TGFß often acts synergistically with the Smad pathway to relay the biological effects, Smad-independent MAPK-mediated responses have also been reported downstream of these growth factors (25, 26, 30, 32, 33, 42).

Because our results indicate that activin/TGFß-mediated inhibition of the Pit-1 gene promoter is Smad independent, we then sought to determine whether the MAPK pathways (p38, ERK1/2, or JNK1/2) were also activated in pituitary cells in response to activin/TGFß. For this, GH4C1 cells were stimulated with activin or TGFß for varying periods of time, as indicated in Fig. 6Go. Total cell lysates were subsequently analyzed by Western blotting using anti-phospho-p38 (Fig. 6AGo), phospho-ERK1/2 (Fig. 6BGo), phospho-JNK1/2 (Fig. 6CGo), or antibodies directed against nonphosphorylated p38 (Fig. 6AGo), ERK1/2 (Fig. 6BGo), and JNK1/2 (Fig. 6CGo) as loading controls. Interestingly, although no apparent changes were observed in ERK1/2 and JNK1/2 phosphorylation in response to activin or TGFß, both ligands were able to strongly induce phosphorylation of the p38 kinase (Fig. 6AGo, top). Collectively, these findings indicate that activin/TGFß activates the p38 MAPK pathway in pituitary cells, suggesting that this pathway may be involved in activin/TGFß-mediated regulation of Pit-1 gene expression.


Figure 6
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  		FIG. 6. Activin/TGFß activates the p38 kinase pathway in GH4C1 cells but not the MAPK MEK1/ERK1/2 and JNK1/2 pathways. GH4C1 cells were starved overnight and stimulated with 0.5 nM activin or 100 pM TGFß for different periods of time as indicated. A, Total cell lysates (20 µg) were analyzed by immunoblot using specific antibody directed against the phosphorylated form of the p38 kinase. The membrane was stripped and reprobed with an anti-p38 antibody as loading control. B, Similarly, Western blot analysis of whole-cell lysates (20 µg) was done using specific antibody to phospho-ERK1/2. Reprobing the blot with anti-ERK1/2 antibody confirmed equal loading. C, The membrane was immunoblotted with a specific antibody against phospho-JNK1/2 and reprobed with anti-JNK1/2 antibody as loading control.

 
The p38 MAPK pathway is required for activin-induced Pit-1 gene promoter inhibition
To evaluate the contribution of the p38 MAPK pathway to activin-mediated down-regulation of human Pit-1 promoter activity in GH4C1 cells, we used a pharmacological inhibitor (PD169316) to specifically block p38 signaling. As controls, we used a p38 inhibitor inactive analog (SB202474), JNK (SP600125), and MEK (PD98059) specific inhibitors. To first verify the efficiency of the p38 kinase inhibitor, under our experimental conditions, GH4C1 cells were pretreated or not with the various inhibitors 2 h before being stimulated with 0.5 nM activin. Total cell lysates were then analyzed by Western blotting with a specific anti-phospho-p38 antibody or anti-p38 antibody for loading control. As shown in Fig. 7AGo (top), in control cells or cells treated with DMSO alone, activin induced phosphorylation of the p38 kinase as previously shown (Fig. 6AGo). However, in cells treated with the specific p38 kinase inhibitor (PD169316) but not its inactive analog (SB202474), the activin effect on p38 phosphorylation was completely blocked. Treatment of cells with specific inhibitors to JNK (SP600125) or MEK (PD98059) did not affect activin-induced phosphorylation of the p38 kinase. Stripping and reprobing of the membranes with the anti-p38 showed equal loading of proteins in all lanes (Fig. 7AGo, bottom).


Figure 7
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  		FIG. 7. p38 kinase inhibitors antagonize activin-mediated down-regulation of Pit-1 promoter activity. A and B, GH4C1 cells were pretreated 2 h with the indicated protein kinase inhibitors (10 µM SB202474, PD169316, PD98059, and SP600125) before being stimulated or not with 0.5 nM activin for 60 min. Total cell lysates were analyzed by immunoblot using specific antibodies directed against the phosphorylated form of p38 (A) and Smad2 (B). Membranes were stripped and reprobed with an anti-p38 antibody (A) for loading control. C, GH4C1 cells were transiently transfected with 0.5 µg of the Pit-1 –102/+15 promoter construct and 0.2 µg of the ß-galactosidase expression plasmid (left) or 0.5 µg of the 3TPLuc construct and 0.2 µg of the ß-galactosidase expression plasmid (right) before being incubated for 2 h in the presence of the indicated protein kinase inhibitors (10 µM SB202474, SB202190, PD169316, PD98059, and SP600125). After 18 h of stimulation with 0.5 nM activin, cells were harvested and luciferase assays performed and normalized according to the ß-galactosidase activities. Three independent and separate experiments were performed in triplicate. For statistical analysis, ANOVA was performed. *, P < 0.05 was considered significant.

 
Previous reports suggest that p38 kinase inhibitors could also affect activin and TGFß type I receptor activity depending on the cell type (43). Thus, to ensure that the p38 kinase inhibitor (PD169316) did not affect the Smad pathway under our experimental conditions, we examined its effect on activin-induced Smad2 phosphorylation. Total cell lysates from GH4C1 cells treated or not with the different inhibitors as described above were analyzed by Western blot using a specific anti-phospho-Smad2 antibody. As shown in Fig. 7BGo, no significant changes in Smad2 phosphorylation were observed in the presence of the various inhibitors. Thus, these results demonstrate that the p38 kinase inhibitor (PD169316) completely blocks the p38 pathway in response to activin without affecting the Smad pathway.

To then address the role and contribution of the p38 MAPK pathway to activin-induced Pit-1 gene promoter inhibition, GH4C1 cells were transiently transfected with the minimal human Pit-1 gene promoter construct (–102/+15Luc) and pretreated for 2 h with the specific pharmacological inhibitors (at a concentration of 10 µM) before being stimulated with activin. Eighteen hours after stimulation of the cells, luciferase activity was assessed. As shown in Fig. 7CGo, activin induced a potent inhibition of the human Pit-1 gene promoter by 50%, as described in Fig. 2Go. In cells treated with DMSO, p38 kinase inactive analog (SB202474), JNK (SP600125), or MEK (PD98059) inhibitors, the activin-induced Pit-1 gene promoter inhibition was not affected. However, in cells treated with the p38 kinase inhibitor (PD169316), the activin effect was almost completely reversed. Similar results were obtained in experimental conditions using higher concentrations (15, 20, and 25 µM) of the various inhibitors (data not shown). Furthermore, to strengthen our results, we used a second specific p38 kinase inhibitor (SB202190). As shown in Fig. 7CGo, like PD169316, SB202190 p38 kinase inhibitor potently reversed the activin inhibitory effect on Pit-1 gene promoter activity.

All together, these results indicate the activin-mediated inhibition of the human Pit-1 gene promoter in pituitary cells is Smad independent but requires the p38 MAPK pathway.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Strict regulation of pituitary hormonal secretion is essential to ensure proper endocrine functions and prevent the development of pituitary tumors, such as prolactinomas. Excessive production of prolactin by these endocrine tumors leads to severe endocrine and reproductive disorders. We recently found that activin suppresses expression of the pituitary-specific transcription factor Pit-1 (18). However, the mechanisms by which activin inhibits Pit-1 expression remained elusive. In the present study, we show that activin exerts a strong regulatory role upon the human Pit-1 gene promoter. We mapped the region of the Pit-1 promoter that confers this activin responsiveness to a 117-bp fragment, contained within the proximal region of the Pit-1 gene promoter. Furthermore, we show that activin-mediated inhibition of the human Pit-1 gene promoter is Smad independent and relies upon the activation of the p38 MAPK pathway.

The 117-bp proximal region of the human Pit-1 gene promoter that is essential to activin-mediated inhibition of Pit-1 expression corresponds to a –102- to +15-bp fragment surrounding the TATA box. Interestingly, although there is little or no homology between the full-length Pit-1 gene promoter sequences of rodents and humans, this 117-bp fragment is the only region that shows a high level of conservation (86%) between these species. Thus, our findings highlighting this region as critical for activin inhibition of the human Pit-1 gene promoter may also apply to other species and represent a general regulatory mechanism for Pit-1 transcriptional repression.

Regulation of the human Pit-1 gene promoter by activin is pituitary specific, because no activin effects were observed in cells derived from nonpituitary origins. Because Pit-1 itself can autoregulate its own promoter, the possibility that activin could interfere with Pit-1 binding to its positive autoregulatory site was investigated. However, our data clearly indicate that the activin effect is independent of the binding of Pit-1 to this site, suggesting that other pituitary-specific factor(s) are required for activin to mediate its effects. Identification of such factor(s) remains to be determined and will be important to further dissect the mechanisms by which activin regulates Pit-1 expression.

A hallmark of the activin/TGFß signaling pathways is the recruitment and activation of the Smad proteins. Smads represent the canonical intracellular mediators for these growth factors and have been shown to be involved in most of the activin/TGFß biological responses (30). However, accumulating evidence suggests that activin and TGFß also signal through other pathways, such as MAPKs (ERK, p38, and JNK). Our results indicate that both the Smad and p38 MAPK pathways are activated in response to activin in pituitary cells. Smad2, Smad3, and p38 are strongly phosphorylated in response to activin, suggesting that both pathways play a role downstream of activin in pituitary cells. However, our results also indicate that the p38 but not the Smad pathway is required for activin-mediated Pit-1 gene promoter inhibition. Indeed, our results clearly indicate that overexpression of DN forms of Smad2 and Smad3 does not affect the activin effect on Pit-1 gene promoter activity, whereas they strongly reversed the activin effect on another Smad-dependent gene promoter construct (3TPLuc). It is therefore conceivable that activin-induced Smad activation in pituitary cells could lead to other activin effects. This is consistent with our previous studies demonstrating that activin/TGFß-mediated cell growth inhibition and prolactin expression inhibition were dependent on the Smad pathway and the Smad-associated protein menin (17, 18). It will be interesting, in future studies, to investigate whether other TGFß family members, such as the bone morphogenetic proteins, which also transduce their signal through the Smad and p38 pathways, can also regulate Pit-1 gene expression, similar to activin and TGFß. Collectively, these studies suggest that distinct pathways mediate the different activin effects in pituitary cells. Interestingly, prolactin expression is also regulated by Pit-1, indicating that activin inhibits prolactin expression through both Smad-independent Pit-1 repression and Smad-dependent inhibition of the prolactin gene promoter, consistent with the strong and potent inhibitory effect exerted by activin on prolactin production in the pituitary (18).

A previous report from our laboratory showed that the p38 pathway is important to activin-induced cell growth arrest in human breast cancer cells (27). Our current results indicate that this pathway also plays a critical role in mediating the activin effects in the pituitary. The role of the p38 kinase pathway may not be restricted to activin-mediated inhibition of Pit-1 expression. Indeed, various functions have been assigned to the p38 MAPK pathway in the pituitary (44, 45). Furthermore, it was previously shown that activation of the TGFß-activated kinase 1/p38 MAPK pathway but not Smad3 is necessary for activin to induce expression of FSHß in the pituitary (45). Together, these findings further emphasize the role played by the p38 MAPK pathway downstream of activin in the pituitary.

Our study sheds light on the mechanisms by which activin regulates Pit-1 expression in lactotrope cells and demonstrates that the p38 MAPK pathway is critical to activin-mediated repression of the pituitary-specific transcription factor Pit-1. These results are particularly important, because Pit-1 is essential for the proper maintenance and function of the pituitary gland. Elucidation of the intricate signaling pathways that regulate Pit-1 expression within the pituitary will further contribute to the development of new therapies for pituitary tumors and will be of high significance for patients with pituitary prolactinomas, particularly those with therapeutic intolerance or insensitivity to dopamine agonists, for whom surgery remains the last resort.


   Acknowledgments
 
We are thankful to Dr. J. Massague for the 3TP-luc construct, Dr. M. Delhase for the human Pit-1 constructs (–1321/+15 Luc, –601/+15 Luc, –102/+140 Luc, and –102/+15 Luc), and Dr. Y. Eto and Ajinomoto Co., Inc., for activin A. The anti-phospho-Smad2 antibody was a gift from Dr. Moustakas, and the anti-Alk4 antibody was kindly provided by J. Vaughan and Dr. W. Vale. We are thankful to Joanne Ho for critical reading of the manuscript.


   Footnotes
 
J.J.L. is Research Scientist of the National Cancer Institute of Canada supported with funds provided by the Canadian Cancer Society, A.L. is a recipient of a Fonds de Recherche de Sante du Quebec scholarship, and C.d.G. is a recipient of a McGill University Health Center scholarship. This work was supported by a grant from the Canadian Institutes for Health Research (MOP-53141 to J.J.L.).

First Published Online June 1, 2006

Abbreviations: ALK4, Activin type I receptor; ALK4mL45, activin type I mutant receptor; ALK4wt, activin type I wild-type receptor; DMSO, dimethylsulfoxide; DN, dominant-negative; DTT, dithiothreitol; FBS, fetal bovine serum; JNK, c-Jun N-terminal kinase.

Received April 6, 2006.

Accepted for publication May 24, 2006.


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Top
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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