Abstract
Involvement of peroxisome proliferator-activated receptor-γ (PPAR-γ ) activation and bone morphogenetic protein (BMP) signaling in regulating cell proliferation and hormonal production of pituitary tumors has been reported, although the underlying mechanism remains poorly understood. Here, we investigated regulatory roles of PPARα and PPARγ in gonadotropin transcription and cell mitosis modulated by pituitary activin/BMP systems using a mouse gonadotropinoma cell line Lβ T2, which expresses activin/BMP receptors, transcription factor Smads, PPARα , and PPARγ . In Lβ T2 cells, BMP signaling shown by Smad1/5/8 phosphorylation and Id-1 transcription was readily activated by BMPs. A PPARγ agonist, pioglitazone significantly reduced BMP-induced DNA synthesis by Lβ T2; whereas the PPARα agonist, fenofibric acid, did not. In accordance with the effects on cell mitosis, pioglitazone but not fenofibric acid significantly decreased BMP-induced Id-1-Luc activation. Neither fenofibric acid nor pioglitazone affected activin signaling detected by (CAGA)9-Luc activity. Both PPARα and PPARγ ligands directly suppressed transcriptional activities of FSHβ , LHβ , and GnRHR. Activation of PPARα and PPARγ increased mRNA levels of follistatin, but did not affect the expression of follistatin-related gene. Thus, PPAR agonists not only directly suppress gonadotropin transcription and BMP signaling, but also inhibit the biological actions of activins which facilitate gonadotropin transcription through upregulating follistatin expression. In addition, pioglitazone increased BMP ligands mRNA, but decreased activin-β B mRNA in Lβ T2 cells. Collectively, PPAR activation differentially regulates gonadotrope cell proliferation and gonadotropin transcription in a ligand-dependent manner.
Introduction
Peroxisome proliferator-activated receptors (PPARs) that include PPARα , PPARβ /δ, and PPARγ belong to the family of nuclear hormone receptors, which are related to thyroid hormone, retinoic acid, androgen, and estrogen receptors (Desvergne & Wahli 1999). PPARγ is activated after the binding of natural ligands such as polyunsaturated fatty acids and prostaglandin metabolites (Willson et al. 2000). PPARγ is also activated by synthetic ligands thiazolidine-diones, e.g. rosiglitazone, pioglitazone, or troglitazone (Lehmann et al. 1995). The binding of thiazolidinediones to their receptors increases insulin sensitivity, and most bind primarily to PPARγ in adipose cells. PPARγ is expressed abundantly in adipose tissue, colon, and hemopoietic cells, moderately in kidney, liver and small intestine, and in small amounts in skeletal muscle (Gurnell 2005). PPARγ was originally described as having a primary role in adipocyte differentiation, but more recent studies have implicated it in processes of inflammation, atherosclerosis, cell cycle control, apoptosis, and carcinogenesis (Auwerx 1999). PPARs act as a transcription factor when heterodimerized with the retinoid X receptor and bound to the proper ligands, including natural and synthetic ligands.
Recent studies have revealed that PPARγ is expressed in pituitary adenomas and PPARγ ligands suppress proliferation and hormonal production of pituitary tumors. Heaney et al.(2002) first demonstrated that PPARγ expression is restricted to adrenocorticotropin (ACTH)-secreting cells of the normal human anterior pituitary. Moreover, PPARγ is also expressed in human ACTH-secreting pituitary tumors causing Cushing’s disease and the activation of PPARγ potently inhibits cell proliferation as well as ACTH secretion of the pituitary adenomas (Heaney et al. 2002). These results support the role for PPARγ as a novel target for treating patients with Cushing’s disease. They further reported that pituitary PPARγ is abundantly expressed in human prolactin-, growth hormone-producing, and nonfunctioning pituitary tumors, and that PPARγ agonists potently inhibit tumor growth and hormonal secretion of these pituitary adenomas (Heaney et al. 2003). These results indicated a possible role for PPARγ as a novel molecular target for treating patients with various pituitary tumors that are unresponsive to current dopamine agonists and/or somatostatin receptor antagonists. However, the underlying mechanism has yet to be elucidated.
Bone morphogenetic proteins (BMPs) belong to the transforming growth factor-β (TGF-β ) superfamily. BMPs were originally identified as the active components in bone extracts capable of inducing bone formation at ectopic sites. A variety of physiological BMP actions in many endocrine tissues, including the ovary (Otsuka et al. 2000, 2001, Miyoshi et al. 2007), pituitary (Otsuka & Shimasaki 2002, Takeda et al. 2003), thyroid (Suzuki et al. 2005), and adrenal (Suzuki et al. 2004, Kano et al. 2005, Inagaki et al. 2006), have been recently clarified. We previously reported the characteristics regarding human follicle-stimulating hormone (FSH)-producing adenomas compared with nonfunctioning adenomas, in which the levels of follistatin mRNA were decreased in FSH-producing adenomas (Takeda et al. 2003). There is increasing evidence that locally produced BMPs, which can also be functionally regulated by follistatin, play critical roles in differentiation of pituitary gonadotrope. For instance, BMP-4 promotes pituitary prolactinoma pathogenesis through a Smad–estrogen receptor crosstalk (Paez-Pereda et al. 2003), whereas BMP-4 inhibits corticotropic pathogenesis and Cushing’s disease in adult pituitary tumor cells (Giacomini et al. 2006). Thus, the pituitary BMP system seems likely to act as a regulator not only for organogenesis and differentiation process of pituitary cells, but also for transformation and tumorigenesis of the differentiated pituitary cells.
In the present study, we investigated the mechanism by which PPAR agonists regulate transcription of gonadotropins using Lβ T2 cell line derived from mouse gonadotrope tumors that express activin/BMP ligands, type I and type II BMP receptors and Smad signaling molecules. Here, we uncovered that PPAR activation plays a differential role in controlling gonadotropin transcription and tumorigenesis of pituitary gonadotrope cells by regulating an endogenous BMP/activin system.
Materials and Methods
Reagents and supplies
Dulbecco’s modified Eagle’s medium (DMEM), penicillin–streptomycin solution, and dimethylsulfoxide (DMSO) were purchased from Sigma–Aldrich Co. Ltd. Recombinant human BMP-2, BMP-4, BMP-6 and BMP-7, and activin A were purchased from R&D Systems Inc. (Minneapolis, MN, USA). Pioglitazone and fenofibric acid were provided from Takeda Chemical Industries (Osaka, Japan) and Kaken Pharmaceutical Co. Ltd (Tokyo, Japan) respectively. Lβ T2 cells were provided by Dr Pamela L Mellon, University of California San Diego. Plasmids of Id-1-Luc were provided from Dr Tetsuro Watabe and Dr Kohei Miyazono, Tokyo University, Japan. Plasmids of oFSHβ-Luc which contains 4741 bp of ovine FSHβ promoter, luteinizing hormone β (LHβ )-Luc which contains 1800 bp of rat LHβ promoter, and gonadotropin-releasing hormone receptor (GnRHR)-Luc which contains 1227 bp of mouse GnRHR promoter were gifted by Dr Pamela L Mellon and Dr William L Miller, North Carolina State University.
Cell culture and thymidine incorporation assay
Lβ T2 cells were grown in DMEM containing 10% fetal calf serum (FCS) supplemented with penicillin–streptomycin at 37° C in an atmosphere of 5% CO2 in air. Cells were grown in 10 cm dishes until they became confluent and were transferred into 12-well plates at a concentration of 3 × 104 viable cells/well. After preculture, medium was replaced with fresh DMEM containing 1% FCS and indicated combinations of BMPs and PPAR ligands were added to the culture medium. After 24-h culture, 0.5 μCi/ml [methyl-3H]thymidine (Amersham Pharmacia) was added, and incubated for 3 h at 37 ° C. The incorporated thymidine was detected as we previously reported (Takeda et al. 2004). Cells were then washed with PBS, incubated with 10% ice-cold trichloroacetic acid for 60 min at 4 ° C, and solubilized in 0.5 M NaOH, and radioactivity was determined with a liquid scintillation counter (TRI-CARB 2300TR, Packard Co., Meriden, CT, USA).
Transient transfection and luciferase assay
Lβ T2 cells were grown in 10 cm plates until they became confluent and then replaced into 24-well plates. After 24-h preculture, the cells (~60% confluent) were transiently transfected with 500 ng of each luciferase reporter plasmid and 50 ng cytomegalovirus-β-galactosidase plasmid (pCMV-β-gal) using FuGENE 6 transfection reagent (Roche Molecular Biochemicals) for 24 h. Cells were treated with BMPs, activins, and PPAR ligands in fresh DMEM containing 1% FCS for 24 h and the cells were washed with PBS and lysed with Cell Culture Lysis Reagent (Toyobo, Osaka, Japan). Luciferase activity and β-galactosidase (β-gal) activity of the cell lysate were measured by luminescencer-PSN (ATTO, Tokyo, Japan), as reported previously (Miyoshi et al. 2006). The data were shown as the ratio of luciferase to β-gal activity.
RNA extraction, RT-PCR, and quantitative real-time PCR analysis
To prepare total cellular RNA, Lβ T2 cells were cultured in 12-well plate (5 × 105 viable cells) and treated with indicated concentrations of growth factors and PPAR ligands in fresh DMEM containing 1% FCS. After 24-h culture, the medium was removed, and total cellular RNA was extracted using TRIzol reagent (Invitrogen Corp.), quantified by measuring absorbance at 260 nm, and stored at −80 ° C until assay. Total RNAs of anterior pituitary, ovary, and kidney tissues were extracted from adult male Wistar rats and BALBc mice using TRIzol reagent. The expression of BMP receptors, binding proteins, and Smad mRNAs were detected by RT-PCR analysis. The extracted RNA (1.0 μg) was subjected to an RT reaction using the First-Strand cDNA synthesis system (Invitrogen Corp.) with random hexamer (2 ng/μl), reverse transcriptase (200 U), and deoxy-NTP (0.5 mM) at 42 ° C for 50 min and 70 ° C for 10 min. Subsequently, hot-start PCR was performed using MgCl2 (1.5 mM), deoxy-NTP 0.2 mM), and 2.5 U Taq DNA polymerase (Invitrogen Corp.) under the same condition. Oligonucleotides used for RT-PCR were custom-ordered from Kurabo Biomedical Co. (Osaka, Japan). PCR primer pairs were selected from different exons of the corresponding genes as follows: activin receptor-like kinase-2 (ALK-2), 639–661 and 1100–1121 (from GenBank accession no. NM_007394); ALK-3, 426–445 and 916–935 (from NM_009758); ALK-4, 130–150 and 637–657 (from NM_007395); ALK-6, 598–617 and 1034–1053 (from NM_007560); activin type II receptor (ActRII), 611–630 and 1083–1102 (from NM_007396); ActRIIB, 130–154 and 377–400 (NM_007397); BMP type II receptor (BMPRII), 13–32 and 515–534 (from NM_007561); Smad1, 878–899 and 1193–1214 (from NM_008539); Smad2, 111–131 and 535–555 (from NM_010754); Smad3, 474–494 and 828–848 (from NM_016769); Smad4, 712–732 and 1082–1102 (from NM_008540); Smad5, 636–658 and 1062–1083 (from NM_008541); Smad6, 1081–1101 and 1527–1547 (from NM_008542); Smad7, 1912–1932 and 2328–2348 (from AJ000550); Smad8, 271–291 and 538–558 (from AF175408); follistatin, 361–385 and 525–550 (from NM_008046); follistatin-related gene (FLRG), 632–651 and 910–929 (from AF276238); PPARα , 601–622 and 932–953 (from NM_011144); PPARγ , 350–371 and 721–742 (from NM_011146); and a housekeeping gene, ribosomal protein L19 (RPL19), 373–393 and 547–567 (from NM_009078). Aliquots of PCR products were electrophoresed on 1.5% agarose gels and visualized after ethidium bromide staining. For the quantification of follistatin and FLRG mRNA levels, real-time PCR was performed using LightCycler-FastStart DNA Master SYBR Green I system (Roche Diagnostic Co.) under conditions of annealing at 60 ° C with 4 mM MgCl2, following manufacturer’s protocol. Accumulated levels of fluorescence were analyzed by the second derivative method after the melting curve analysis (Roche Diagnostic), and then the expression levels of target genes were standardized by RPL19 level in each sample.
Western immunoblot analysis
Cells (2 × 105 viable cells) were precultured in 12-well plates in DMEM containing 10% FCS for 24 h. After preculture, medium was replaced with serum-free fresh DMEM, and then indicated concentrations of BMPs and PPAR ligands were added to the culture medium. After stimulation with growth factors for 1 h, cells were solubilized in 100 μl RIPA lysis buffer (Upstate Biotechnology, Inc., Lake Placid, NY, USA) containing 1 mM Na3VO4, 1 mM sodium fluoride, 2% sodium dodecyl sulfate, and 4% β-mercaptoethanol. The cell lysates were then subjected to SDS–PAGE/immunoblotting analysis using anti-phospho-Smad1/5/8 antibody (Cell Signaling Technology Inc., Beverly, MA, USA).
Immunofluorescence microscopy
Cells were precultured in serum-free DMEM and then cells at ~50% confluency were treated with BMPs. After 1-h stimulation, cells were fixed with 4% formaldehyde in PBS, permeabilized with 0.5% Triton X-100 in PBS at room temperature, and washed thrice with PBS. The cells were then incubated with anti-phospho-Smad1/5/8 antibody for 1 h and washed thrice with PBS. Cells were then incubated with Alexa Flour 488 anti-rabbit IgG (Invitrogen Corp.) in a humidified chamber for 1 h and washed with PBS, and then stained cells were visualized under a fluorescent microscope.
cDNA array analysis
GEArray system (SuperArray Bioscience Corp., Frederick, MD, USA) that includes 96 genes of mouse TGF-β and BMP signaling pathway was used for analyzing the expression pattern of BMP signaling system in Lβ T2 cells. As reported previously (Miyoshi et al. 2006), extracted total RNAs (3.0 μg) were used as templates to generate biotin-16-dUTP-labeled cDNA probes according to manufacturer’s instruction. The cDNA probes were denatured and hybridized at 60 ° C with the cDNA array membranes, which were washed and exposed to X-ray films with use of chemiluminescent substrates. To analyze the array results, we scanned the X-ray film and the image was inverted as grayscale TIFF files. The spots were digitized and analyzed using GEArray analyzer software (SuperArray Bioscience Corp.) and the data were normalized by subtraction of the background as the average intensity levels of three spots containing plasmid DNA of pUC18. The averages of two spots of glyceraldehyde-3-phosphate dehydrogenase and four spots of cyclophilin A (PPIA) were used as positive controls to compare the membranes. Using these standardized data, we compared the signal intensity of the membranes using the GEArray analyzer program (SuperArray Bioscience Corp.).
Statistical analysis
All results are shown as means ± s.e.m. of data from at least three separate experiments, each performed with triplicate samples. Differences between groups were analyzed for statistical significance using ANOVA (StatView 5.0 software, Abacus Concepts Inc., Berkeley, CA, USA). P < 0.05 were accepted as statistically significant.
Results
We first examined the mRNA expression of components of the activin/BMP system by RT-PCR analysis in Lβ T2 cells. The expression of BMP type I receptors, including ALK-2 (also called ActRIA), ALK-3 (BMPRIA), and ALK-4 (ActRIB); BMP type II receptors, including ActRII, ActRIIB, and BMPRII; binding proteins, including FLRG and follistatin; and Smad signaling molecules, including Smad1/2/3/4/5/8; and inhibitory Smads, Smad6/7 were detected (Fig. 1). ALK-6 (BMPRIB) mRNA expression was not detected in Lβ T2 cells. PPARγ mRNA expression was detected at comparatively higher levels than PPARα in Lβ T2 cells, which parallels the expression pattern of PPARs in normal anterior pituitary tissues in rat. In contrast, mouse ovary and kidney tissues expressed PPARα and PPARγ mRNA at similar levels (Fig. 1).
As shown in Fig. 2, a PPARγ agonist, pioglitazone, but not the PPARα agonist, fenofibric acid, suppressed cellular DNA synthesis shown by thymidine incorporation at a high dose of 10 μM. Cell mitosis induced by BMP-2 and BMP-4 was suppressed by pioglitazone, but not fenofibric acid (Fig. 2). To assess the mechanism by which PPAR ligands affect BMP signaling in Lβ T2 cells, intracellular signaling activity of BMP was examined using BMP–Smad1/5/8 pathway-specific reporter, Id-1-Luc. As shown in Fig. 3, BMP-2, but not activin A, increased Id-1-Luc activity, while PPAR ligands had no specific effects on Id-1-Luc activity. BMP-2 induction of Id-1-luc was specifically reduced by the PPARγ agonist but not by the PPARα agonist in a dose-dependent manner. Western blots analysis demonstrated rapid phosphorylation of Smad1/5/8 protein induced by BMP-2, BMP-4, BMP-6, and BMP-7 (Fig. 4). Immunofluorescence study clearly demonstrated nuclear localization of the phosphorylated Smad1/5/8 molecules by treatments with BMP-2 and BMP-4 (Fig. 4). Smad phosphorylation induced by BMP-2 and BMP-4 was not affected even in the presence of PPAR agonists (Fig. 4), suggesting that the PPAR activation inhibits BMP signal transduction at the downstream of Smad1/5/8 phosphorylation in Lβ T2 cells.
We next examined the transcriptional regulation of FSHβ , LHβ , and GnRHR in Lβ T2 cells. As shown in Fig. 5, activin A, but not BMP-2, stimulated transcription of FSHβ , LHβ , and GnRHR. Importantly, in striking contrast with the established concept that activin does not stimulate LHβ mRNA expression or LH synthesis in primary pituitary cells, activin did stimulate LHβ transcription in Lβ T2 cells. Similar data were previously reported by Pernasetti et al.(2001).
It was of note that PPAR agonists suppressed transcriptional activities of FSHβ , LHβ , and GnRHR in Lβ T2 cells (Fig. 5). To assess the mechanism by which PPAR agonists affect activin signaling in Lβ T2 cells, we used the activin–Smad2/3-specific reporter (CAGA)9-Luc. PPARα and PPARγ agonists had no direct effects on (CAGA)9-Luc activity. In addition, neither PPARα nor PPARγ agonists affected (CAGA)9-Luc activation induced by exogenously added activin A (Fig. 6). Therefore, it is likely that PPARα and PPARγ directly inactivate transcription of FSHβ , LHβ , and GnRHR without affecting Smad2/3 signaling in Lβ T2 cells.
Interestingly, follistatin expression was increased in a dose-dependent manner by PPAR agonists (Fig. 7). In contrast, expression of FLRG, another activin-binding protein, was not altered by PPAR agonists. Activin A induced FLRG expression in Lβ T2 cells (Fig. 7). Thus, PPAR agonists not only directly suppress gonadotropin transcription and BMP signaling, but also indirectly inhibit the biological actions of activins/BMPs through the upregulation of follistatin expression. BMP ligands, including BMP-2, BMP-4, BMP-6 and BMP-7, as well as the inhibin-α and activin-β B subunits are expressed in Lβ T2 cells (Otsuka & Shimasaki 2002). In the present study, changes in the expression of BMP ligands were investigated by cDNA array analysis (Fig. 8). In the presence of PPARγ agonists, the expression of BMP ligands, including BMP-2, BMP-3, BMP-4, BMP-6, BMP-7, and BMP-8a, were increased, while that of activin-β B subunit was decreased in Lβ T2 cells.
Discussion
In the present study, the effects of PPARα and PPARγ activation on transcriptional regulation of gonadotropins were investigated in Lβ T2 cells, which express activin/BMP receptors, Smad2/3 and Smad1/5/8, and PPARα and PPARγ . A PPARγ agonist, pioglitazone, but not a PPARα agonist, fenofibric acid, significantly reduced BMP-induced DNA synthesis by Lβ T2. In accordance with the effects on cell mitosis, PPARγ but not PPARα activation inhibited the BMP signaling evidenced by Id-1-Luc activity (Fig. 9).
Heaney et al.(2003) demonstrated an interesting observation using Lβ T2 cells subcutaneously injected into nude mice. Tumor weights increased and plasma LH levels were markedly decreased in mice treated with high-dose ( > 20 mg/kg per day) of rosiglitazone, although low-dose (5 mg/kg per day) rosiglitazone was insufficient for inhibiting the tumor growth. These results confirm the potent effects of PPARγ activation on anti-proliferation in vivo and demonstrate the potential role of PPARγ ligands for therapy in gonadotrope and nonfunctioning pituitary tumors (Heaney 2003).
PPARγ acts as a transcription factor, and its involvement in adipocyte differentiation led to studies on anti-proliferative effects of PPARγ activation on various neoplasms (Lefebvre et al. 1998). The anti-proliferative effects of PPARγ ligands are related to activation of cyclin-dependent kinase inhibitors (p21 and p27), reduction of cyclins D1 and E, and induction of apoptosis via activation of caspase 3 and inhibition of anti-apoptotic proteins (Bcl-2 and Bcl-XL), although recent data suggest that the suppression of cell growth by rosiglitazone may be, at least in part, via a PPARγ-independent pathway (Palakurthi et al. 2001, Emery et al. 2006). In addition to the inhibitory effects of PPARγ agonist on BMP-induced Lβ T2 cell mitosis, both PPARα and PPARγ agonists directly reduced transcriptional activities of FSHβ , LHβ , and GnRHR. These ligands also increased mRNA levels of an activin-binding protein, follistatin. Thus, PPARα and PPARγ activation differentially regulates gonadotrope cell proliferation and gonadotropin transcription in a ligand-dependent manner (Fig. 9).
Recent reports have suggested that BMPs are directly involved in regulating gonadotrope function. Huang et al.(2001) reported that BMP-6 and BMP-7 stimulate FSH synthesis and secretion in a gonadotrope cell line Lβ T2. BMP-15 selectively stimulates FSH biosynthesis and secretion by rat primary pituitary cells without affecting LH secretion or GnRHR transcription (Otsuka & Shimasaki 2002). Lee et al.(2007) reported an interesting observation that BMP-2 and activin A synergistically and independently stimulate FSHβ transcription in Lβ T2 cells. In their study, BMP-2 enhanced FSHβ transcription using cells transfected with mouse 1.9 kb FSHβ reporter construct (Lee et al. 2007). However, BMP-2 effect on FSHβ transcriptional activation was negligible compared with activin A actions in our present study using ovine 4.7 kb FSHβ promoter. This difference could be due to the assay sensitivity or the possible actions of 5′-distal FSHβ regulatory region on modulating BMP-responsive FSHβ transcription.
Alexander et al.(1995) study have shown that a binding protein for activin/BMP, follistatin is expressed in more than half of nonfunctioning adenomas as well as in normal human pituitary tissues. Interestingly, follistatin expression was not detected in other functioning adenomas, including prolactinomas, growth hormone-, and ACTH-producing adenomas in the same study (Alexander et al. 1995). Penabad et al.(1996) also showed that follistatin protein is detected in nonadenomatous gonadotropes but barely in gonadotropinomas, in which follistatin immunoreactivity is co-localized with FSHβ subunit in the cytoplasm.
Since FSH-producing pituitary adenomas express the activin-β B subunit, impairment of activin neutralization due to reduced follistatin may result in increased cell mitosis and FSH hypersecretion in gonadotropinomas. In this regard, we previously showed the critical difference between FSH-producing adenoma and nonfunctioning adenoma with respect to the level of follistatin mRNA (Takeda et al. 2003). Namely, in nonfunctioning adenomas, the expression of follistatin is abundant and thereby the activation of activin/BMP signaling is insufficient, resulting in less production of FSH despite the existence of activins and BMPs. On the contrary, in gonadotropinomas, the effect of follistatin is negligible and thereby activins can bind their receptor and transduce the signaling effectively to stimulate FSH synthesis and secretion (Takeda et al. 2003).
Regarding the pituitary BMP system, BMP-2 and BMP-4 play a key role for the initial development of the anterior pituitary (Scully & Rosenfeld 2002). BMP-4 is required during the first stage of pituitary organogenesis for the proliferation of the Rathke’s pouch, which gives rise to Pit-1 lineage cells including lactotrope cells. Moreover, over-expression of noggin or a dominant-negative ALK-3 in the anterior pituitary leads to the arrest of the development of Pit-1-expressing lineage (Scully & Rosenfeld 2002). During the second stage of pituitary organogenesis, an inhibition of BMP-2, a ventrodorsal gradient, by an FGF-8, a dorso-ventral gradient, leads to differentiation of corticotrope cells (Dasen & Rosenfeld 2001). Overexpression of FGF-8 in the developmental pituitary results in defect of Pit-1 lineage cells with enhanced differentiation of corticotrope cells, implying that inhibition of the BMP signaling is necessary for corticotrope development (Kioussi et al. 1999).
BMP-4 not only governs pituitary organogenesis, but also plays a key role in the pathogenesis of differentiated pituitary lineages. It is reported that BMP-4 is overexpressed in the lactotrope adenoma derived from dopamine type-2 receptor (D2R) null mouse as well as human prolactinomas (Paez-Pereda et al. 2003). Of particular interest, a binding protein for BMPs, noggin expression is conversely down-regulated in the prolactinoma from the D2R null mouse (Paez-Pereda et al. 2003), suggesting a new concept that BMP-4 promotes cell proliferation in lactotoropes in conjunction with Smad–estrogen receptor interaction under the influence of its binding protein. BMP-4 also inhibits ACTH secretion and cell proliferation of corticotropinoma cells (Giacomini et al. 2006). In addition, anti-proliferative effect of retinoic acid on corticotropinoma seems to be mediated by the BMP-4 action. Thus, BMP-4 promotes pituitary prolactinoma through a Smad–estrogen receptor crosstalk (Paez-Pereda et al. 2003), while BMP-4 inhibits corticotrope pathogenesis, i.e. Cushing’s disease (Giacomini et al. 2006).
As seen in the imbalanced expression of BMP-4 and noggin by lactotrope adenomas (Paez-Pereda et al. 2003), the intrapituitary regulation of activin/BMPs by follistatin may be involved in the functional transition between nonfunctioning adenomas and FSH-producing adenomas in vivo. Based on our earlier data, the expression levels of BMP-6 and BMP-7 were lowered while that of activin-β B subunit was increased in FSH-producing adenomas (Takeda et al. 2003), implying involvement of the pituitary activin/BMP system in development of gonadotropinomas. Interestingly, PPARγ activation increased expression of BMP-6 and BMP-7 mRNAs while reduced that of activin-β B mRNA in Lβ T2 cells in the present results, suggesting the reminiscent pattern shown in nonfunctioning adenomas rather than FSH-producing adenomas.
It seems likely that PPAR agonists not only directly suppress gonadotropin transcription and BMP signaling, but also inhibit effects of endogenous activins on gonadotropin transcription through upregulating follistatin expression. Therefore, one may consider that PPARα and PPARγ might be involved in differentiation of gonadotropinoma from nonfunctioning adenomas by modulating follistatin transcription in the pituitary adenomas. In contrast to the changes in follistatin expression, mRNA levels of FLRG, which has been reported to regulate biological actions of activins A, B and AB, and BMP-2 (Tsuchida 2004), were not altered by PPAR activation in Lβ T2 cells. Arai et al.(2003) reported that FLRG plays important roles in the regulation of reproductive events, especially in the placenta and decidua during the latter half of pregnancy. In their study, FLRG mRNA was weakly detected in the rat pituitary, as shown in Lβ T2 cells. The finding that FLRG, but not follistatin, expression was increased by activin also provides the possibility of differential regulation of transcription between FLRG and follistatin in pituitary gonadotrope as shown in other tissues such as placenta and skin (Wankell et al. 2001, Wang et al. 2003). Roles of FLRG in gonadotrope cells need to be further elucidated in the future study.
Collectively, the present study demonstrates PPAR regulation of gonadotropin transcription and tumorigenesis of pituitary gonadotrope cells partly through regulating the endogenous BMP/activin/follistatin system (Fig. 9). PPAR activation in gonadotropes may play a key role in determining capability of gonadotropins production in the pituitary, which may be linked to differentiation between nonfunctioning and FSH-producing adenomas.
Expression of BMP system and peroxisome proliferator-activated receptors (PPARs) in Lβ T2 cells. The expression of mRNAs encoding BMP type I and II receptors, binding proteins, Smads, and a housekeeping gene RPL19 was examined by RT-PCR in total cellular RNA extracted from Lβ T2 cells. Expression pattern of PPARα and PPARγ mRNA from Lβ T2 cells was compared with that from rat anterior pituitary, mouse ovary, and mouse kidney tissues by RT-PCR. Aliquots of PCR products were electrophoresed on 1.5% agarose gel, visualized by ethidium bromide staining and shown as representative of those obtained from three independent experiments. MM indicates molecular weight marker.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR activation on DNA synthesis by Lβ T2 cells. After preculture, cells (3 × 104 viable cells) were treated with indicated concentrations of fenofibric acid (FA, a PPARα agonist), pioglitazone (PIO, a PPARγ agonist), and BMP-2 and BMP-4. After 24-h culture, 0.5 μCi/well of [methyl-3H]thymidine was added and incubated for 3 h at 37 ° C. Cells were then washed, precipitated by trichloroacetic acid and solubilized in NaOH and its radioactivity was counted. Results show the mean ± s.e.m. of data performed with triplicate incubations. *P < 0.05 and **P < 0.01 versus control or between the indicated groups.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR activation on BMP signaling in Lβ T2 cells. After preculture, cells were transiently transfected with 500 ng Id-1-Luc and 50 ng cytomegalovirus-β-galactosidase plasmid (pCMV-β-gal). Cells were then treated with indicated concentrations of PPAR agonists (PPARα , FA: fenofibric acid; PPARγ , PIO: pioglitazone) and BMP-2/activin A (100 ng/ml) for 24 h. Cells were washed with PBS, lysed, and the luciferase and β-galactosidase (β gal) activities were measured by luminometer. Results were shown as the ratio of luciferase to β gal activity and graphed as mean ± s.e.m. of data performed with triplicate treatments. *P < 0.05 and **P < 0.01 versus control or between the indicated groups.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR agonists on Smad1/5/8 phosphorylation in Lβ T2 cells. After 24-h preculture, cells (2 × 105 viable cells) were treated with BMP-2, BMP-4, BMP-6 and BMP-7, and activin A (100 ng/ml) in combination with PPAR agonists (PPARα , fenofibric acid; PPARγ , pioglitazone; 10 μM) in serum-free conditions. Following 1-h stimulation with growth factors, cells lysates were obtained and then subjected to SDS–PAGE/immunoblotting (IB) analysis using anti-phospho-Smad1/5/8 antibody. Immunofluorescent (IF) studies were also performed using pSmad1/5/8 antibody on BMP-treated cells for 1 h.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR agonists on gonadotropin and GnRHR transcription by Lβ T2 cells. Cells were precultured in 12-well plates and then transiently transfected with 500 ng FSHβ-Luc, LHβ-Luc and GnRHR-Luc, and 50 ng cytomegalovirus-β-galactosidase plasmid (pCMV-β-gal). Cells were treated with indicated concentrations of PPAR agonists (PPARα , FA: fenofibric acid; PPARγ , PIO: pioglitazone) and activin A/BMP-2 (100 ng/ml) for 24 h. Cells were washed with PBS, lysed, and the luciferase and β-galactosidase (β gal) activities were measured by luminometer. Results were shown as the ratio of luciferase to β gal activity and graphed as mean ± s.e.m. of data performed with triplicate treatments. *P < 0.05 versus control.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR activation on activin signaling in Lβ T2 cells. After preculture, cells were transiently transfected with 500 ng (CAGA)9-Luc and 50 ng cytomegalovirus-β-galactosidase plasmid (pCMV-β-gal). Cells were then treated with indicated concentrations of PPAR agonists (PPARα , FA: fenofibric acid; PPARγ , PIO: pioglitazone) and activin A/BMP-2 (100 ng/ml) for 24 h. Cells were washed with PBS, lysed, and the luciferase and β-galactosidase (β gal) activities were measured by luminometer. Results were shown as the ratio of luciferase to β gal activity and graphed as means ± s.e.m. of data performed with triplicate treatments. *P < 0.05 and **P < 0.01 versus control.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR agonists on mRNA expression of follistatin (FS) and follistatin-related gene (FLRG) in Lβ T2 cells. After preculture, cells (4 × 105 viable cells) were treated with indicated concentrations of PPAR agonists (PPARα , FA: fenofibric acid; PPARγ , PIO: pioglitazone) and activin A/BMP-2 (100 ng/ml). Total cellular RNAs were extracted and subjected to RTreaction. Levels of steady-state mRNAs of FS and FLRG were analyzed by quantitative real-time PCR and standardized by level of RPL19 in each sample. Changes of the mRNA levels by treatments were graphed. Results show the mean ± s.e.m. of data performed with triplicate treatments. *P < 0.05 and **P < 0.01 versus control.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
Effects of PPAR activation on the expression pattern of BMP ligands in Lβ T2 cells. Total cellular RNAs (3 μg) were extracted from Lβ T2 cells cultured for 24 h in the absence (−) or presence ( + ) of a PPARγ agonist, pioglitazone (10 μM). The RNAs were used as templates to generate biotin-16-dUTP-labeled cDNA probes for GEArray membranes (SuperArray Bioscience Corp.) that include human TGF-β and BMP signaling molecules. The cDNA probes were denatured and hybridized with the cDNA array membranes and then the membranes were washed and exposed to X-ray films using chemiluminescent substrate. The spots on the X-ray films were scanned, digitized, and the signal intensities were numerically converted. The signal intensities of the spots on the membranes obtained from two separate experiments were analyzed using the GEArray analyzer program (SuperArray Bioscience Corp.) after subtraction of the background levels of pUC18 DNA. The average of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as positive control to compare the membranes.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
A possible mechanism by which PPAR regulates gonadotropin transcription and cell proliferation in gonadotrope cells. Lβ T2 cells express BMP type I and type II receptors, and transcription factor Smads. Neither PPARα nor PPARγ affects activin signaling, whereas both PPAR agonists suppress transcriptional activities of gonadotropins (Gn) directly. Activation of PPARγ , but not PPARα , specifically suppresses BMP signaling of Lβ T2 cells, leading to inhibition of cell mitosis. PPARs enhance follistatin mRNA expression that sequesters biological actions of activin.
Citation: Journal of Endocrinology 194, 1; 10.1677/JOE-07-0138
We thank Dr Pamela L Mellon for Lβ T2 cells, Drs William L Miller and Pamela L Mellon for the oFSHβ-Luc, LHβ-Luc, and GnRHR-Luc plasmids. We thank Dr R Kelly Moore for helpful discussion and critical reading of the manuscript. We also thank Drs Tetsuro Watabe and Kohei Miyazono for providing Id-1-Luc plasmid. This work was supported in part by Grants-in-Aid for Scientific Research, Yamaguchi Endocrine Research Association, Takeda Science Foundation, The Tokyo Biochemical Research Foundation, Sakakibara Memorial Research Grant, The Ichiro Kanehara Foundation, and Kato Memorial Bioscience Foundation. The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.
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