Peroxisome Proliferator-Activated Receptor Suppresses 11?-Hydroxysteroid Dehydrogenase Type 2 Gene Expression in Human Placental Trophoblas
http://www.100md.com
内分泌学杂志 2005年第3期
Canadian Institutes of Health Research Group in Fetal and Neonatal Health and Development, Children’s Health Research Institute and Lawson Health Research Institute, Departments of Obstetrics and Gynecology and Physiology and Pharmacology, University of Western Ontario, London, Ontario, Canada N6A 4G5
Address all correspondence and requests for reprints to: Dr. K. Yang, Children’s Health Research Institute, Room A5-132, Victoria Research Laboratories-Westminster Campus, 800 Commissioners Road East, London, Ontario, Canada N6A 4G5. E-mail: kyang@uwo.ca.
Abstract
Accumulating evidence suggests that the human placental enzyme 11?-hydroxysteroid dehydrogenase type 2 (11?-HSD2) plays a key role in fetal development by controlling fetal exposure to maternal glucocorticoids. Recently, the nuclear peroxisome proliferator-activated receptor (PPAR) has been found to be the most abundantly expressed PPAR subtype in the human placenta, but its function in this organ is unknown. Given that PPAR-null mice exhibited placental defects and consequent intrauterine growth restriction, the present study was undertaken to examine the hypothesis that PPAR regulates human placental function in part by targeting 11?-HSD2. Using cultured human trophoblast cells as a model system, we demonstrated that 1) the putative PPAR agonist carbaprostacyclin (cPGI2) reduced 11?-HSD2 activity as well as 11?-HSD2 expression at both protein and mRNA levels; 2) GW610742 (a selective PPAR agonist) mimicked the effect of cPGI2, whereas indomethacin (a known ligand for PPAR and PPAR) had no effect; 3) the cPGI2-induced down-regulation of 11?-HSD2 mRNA did not require de novo protein synthesis; 4) cPGI2 suppressed HSD11B2 promoter activity, but did not alter the half-life of 11?-HSD2 mRNA; and 5) the inhibitory effect of cPGI2 on HSD11B2 promoter activity was abrogated in trophoblast cells cotransfected with a dominant negative PPAR mutant. Taken together, these findings suggest that activation of PPAR down-regulates HSD11B2 gene expression in human trophoblast cells, and that this effect is mediated primarily at the transcriptional level. Thus, the present study reveals 11?-HSD2 as an additional target for PPAR and identifies a molecular mechanism by which this nuclear receptor may regulate human placental function.
Introduction
GLUCOCORTICOIDS ARE ESSENTIAL for normal fetal organ growth and maturation (1). However, excessive glucocorticoid exposure in utero leads to intrauterine growth restriction (IUGR) and may predispose to type 2 diabetes and hypertension later in life (2, 3). The placental enzyme 11?-hydroxysteroid dehydrogenase type 2 (11?-HSD2) is believed to play a key role in protecting the fetus from exposure to high levels of maternal glucocorticoid by converting maternal cortisol to its inactive metabolite, cortisone (4). Indeed, placental 11?-HSD2 activity is correlated with birth weight (5) and is attenuated in pregnancies complicated with IUGR (6). Moreover, IUGR is a characteristic feature of apparent mineralocorticoid excess syndrome, an autosomal recessive disorder in which point mutations in the HSD11B2 gene render the enzyme inactive (7). Thus, the precise control of placental 11?-HSD2 expression and activity appears to be critical to normal fetal development.
Previous studies have demonstrated that retinoic acid and forskolin increase, whereas nitric oxide, progesterone, and catecholamines decrease 11?-HSD2 activity and mRNA levels in JEG-3 choriocarcinoma cells and cultured human placental trophoblasts (8, 9, 10, 11, 12). Furthermore, placental 11?-HSD2 activity, but not mRNA, is reduced by prostaglandins, leukotriene B4, and calcium (13, 14). In addition, a series of elegant studies by Albrecht, Pepe, and colleagues (15, 16, 17, 18) has implicated estrogen as a critical factor in maintaining the developmental pattern of expression of 11?-HSD2 in the baboon placenta. More recently, we and others have provided evidence that hypoxia is a negative regulator of human placental 11?-HSD2 expression and activity (19, 20). However, our understanding of placental 11?-HSD2 regulation is incomplete, particularly at the level of HSD11B2 gene transcription.
There is a good correlation between placental 11?-HSD2 activity and mRNA in both uncomplicated pregnancies and those complicated by IUGR (5, 21), suggesting that regulation of 11?-HSD2 in the human placenta may occur at a transcriptional level. Indeed, we have demonstrated recently that glucocorticoids stimulate HSD11B2 gene transcription in cultured human trophoblast cells (22). Peroxisome proliferator-activated receptors (PPARs) are members of the nuclear receptor superfamily. Three distinct PPAR subtypes have been identified, namely, PPAR, PPAR (also known as PPAR?), and PPAR, with unique tissue distributions and physiological functions. They act as regulatory transcription factors that heterodimerize with retinoid X receptors (RXR), bind to peroxisome proliferator-responsive elements (PPREs) located in the regulatory region of target genes, and modulate gene expression in response to ligand activation (23). Although PPAR and PPAR have been studied extensively, relatively less is known about PPAR (24). PPAR is widely expressed, but some tissues, such as the brain, skin, colon, and placenta, express higher levels (25). Recent studies have implicated PPAR in embryo implantation (26), epidermal maturation (27), colon cancer (28), preadipocyte proliferation (29), and obesity (30). Homozygous PPAR-null mice exhibit placental defects and reduced birth weight (31, 32), indicating a pivotal role for this nuclear receptor in murine placental function and fetal development. However, the role of PPAR in human fetal development is unknown. Given the particularly high level of PPAR expression in the human placenta (25), we hypothesized that this nuclear receptor may regulate human placental function in part by targeting 11?-HSD2. In the present study we tested this hypothesis using cultured human placental trophoblast cells as a model system. We present the first evidence that the activation of PPAR suppresses HSD11B2 gene transcription, leading to reduced 11?-HSD2 activity and expression in human trophoblast cells.
Materials and Methods
Reagents and supplies
[1,2,6,7-N-3H]Cortisol (80 Ci/mmol) was purchased from DuPont Canada, Inc. (Markam, Canada). Nonradioactive cortisol and cortisone were obtained from Steraloids, Inc. (Wilton, NH). Cycloheximide (CHX) and 5,6-dichlorobenzimidazole 1?-D-ribfuranoside (DRB) were purchased from Sigma-Aldrich Canada Ltd. (Oakville, Canada). Carbaprostacyclin (cPGI2) was obtained from Cayman Chemical Co. (Ann Arbor, MI), and GW610742 was a gift from GlaxoSmithKline (Research Triangle Park, NC). Polyester-backed thin layer chromatography plates were obtained from Fisher Scientific Ltd. (Nepean, Canada). All solvents used were purchased from VWR Canlab (Mississauga, Canada). Cell culture supplies were obtained from either Invitrogen Life Technologies, Inc. (Burlington, Canada) or Fisher Scientific. General molecular biology reagents were purchased from Invitrogen Life Technologies, Inc., or Pharmacia Canada, Inc. (Baie d’Urfe, Canada). Oligonucleotides were synthesized by a Pharmacia Gene Assembler and purified with NAP-50 columns (Pharmacia Biotech, Piscataway, NJ) according to the manufacturer’s instructions.
Placental trophoblast cell cultures
Placental trophoblast cells were isolated using a modification of the method of Kliman (33) as previously described (20). Ethics approval for procurement of human placentas was obtained from the University of Western Ontario ethics board for health sciences research involving human subjects. Briefly, human placentas were obtained from uncomplicated pregnancies at term after elective cesarean section. Villous tissues were dissected free from fetal membranes and blood vessels, rinsed in 0.9% NaCl2, and digested with 0.125% trypsin and 0.02% deoxyribonuclease I (Sigma-Aldrich Canada Ltd.) in DMEM containing 0.05% streptomycin and gentamicin (Invitrogen Life Technologies, Inc.) three times for 30 min each time. The placental cells were loaded onto a 5–70% Percoll gradient at step increments of 5% Percoll and centrifuged at 2500 x g for 20 min to separate different cell types. Cytotrophoblasts between the density markers of 1.049 and 1.062 g/ml were collected and plated in either 24-well plates (for enzyme activity assay) or 35-mm dishes (for Western blot analysis) at a density of 1.35 x 106 cells/ml in medium 199 containing 10% fetal calf serum (Invitrogen Life Technologies, Inc.). The cells were maintained at 37 C in humidified 5% CO2-95% air (20% O2) for 48 h. We have shown previously that the isolated cytotrophoblasts will differentiate into syncytiotrophoblasts over 48 h of culture under the conditions used in the present study. After 48 h, the trophoblast cells (in triplicate) were treated for 24 h (or as indicated otherwise) with various compounds in medium containing 2% fetal calf serum. Controls, also performed in triplicate, received an equivalent volume of vehicle (ethanol or dimethylsulfoxide).
RT-PCR
To verify the expression of PPAR in cultured human placental trophoblasts, the relative abundance of PPAR mRNA was assessed by a standard RT-PCR, as described previously (11). Briefly, total RNA was extracted from cultured trophoblast cells as well as human placental tissues using TRIzol reagent (Invitrogen Life Technologies, Inc.) according to the manufacturer’s instructions. One microgram of total RNA was reverse transcribed using a standard oligo(deoxythymidine) primer in a total volume of 20 μl. An aliquot (2 μl) of the RT reaction was then subjected to a standard PCR (94 C, 30 sec; 55 C, 30 sec; 72 C, 45 sec; 30 cycles) using sequence-specific primers (forward primer, 5'-GGT GAA TGG CCT GCC TCC CTA CAA; reverse primer, 5'-CAC AGA ATG ATG GCC GCA ATG AAT), which correspond to nucleotides 1029–1052 and 1409–1386 in the published human PPAR cDNA (GenBank accession no. BC007578), respectively. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as control. The same PCR conditions were used for GAPDH, except that the annealing temperature was 56 C, and 28 cycles were used. The primers for GAPDH (forward primer, 5'-ATC ACT GCC ACC CAG AAG AC; reverse primer, 5'-TGT GAG GGA GAT GCT CAG TG) correspond to nucleotides 565–584 and 1126–1145 in the published GAPDH cDNA (GenBank accession no. M17701). A fraction of the PPAR and GAPDH RT-PCR products was subjected to electrophoresis on a 1.2% agarose gel, stained with ethidium bromide, and photographed.
Assay of 11?-HSD2 activity: radiometric conversion assay
The level of 11?-HSD2 activity in intact cells at various time points and after different treatment regimens was determined by measuring the rate of cortisol to cortisone conversion, as described previously (20). Briefly, the cells were incubated for 1 h at 37 C in serum-free medium containing approximately 50,000 cpm [3H]cortisol and 100 nM unlabeled cortisol. At the end of the incubation, medium was collected, and steroids were extracted. The extracts were dried, and the residues were resuspended. A fraction of the resuspension was spotted on a thin layer chromatography plate that was developed in chloroform/methanol (9:1, vol/vol). The bands containing the labeled cortisol and cortisone were identified by UV light of the cold carriers, cut out into scintillation vials, and counted in Scintisafe Econo 1 (Fisher Scientific). The rate of cortisol to cortisone conversion was calculated, and the blank values (defined as the amount of conversion in the absence of cells) were subtracted and expressed as a percentage of control. Results are shown as the mean ± SEM.
Protein extraction and Western blot analysis
Cells were lysed with cold lysis buffer (100 mM NaCl, 50 mM sodium fluoride, 0.1% sodium dodecyl sulfate, 1% sodium deoxycholate, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 0.1 mM phenylmethylsulfonylfluoride, 1 mM orthovanadate, and 50 mM Tris-HCl, pH 7.5) for 5 min at room temperature. The total cell lysates were collected with a cell scraper, vortexed vigorously, and centrifuged at 10,000 x g for 20 min at 4 C. The supernatant was collected, and the protein content was determined by the Bradford method using a protein assay kit (Bio-Rad Laboratories, Inc., Missasauga, Canada) with BSA as a standard.
Western blot analysis was conducted as described previously (20). Briefly, 30 μg protein extracts were subjected to a standard 12% SDS-PAGE. After electrophoresis, proteins were transferred to nitrocellulose using a Bio-Rad Mini Transfer Apparatus. 11?-HSD2 protein was detected on the nitrocellulose filter using an enhanced chemiluminescence Western blotting analysis system (Pharmacia Biotech) following the manufacturer’s instructions. Briefly, the nitrocellulose filter was blocked overnight at 4 C with 10% Blotto in 0.1% Tween 20 in Tris-buffered saline (TTBS), and incubated with primary antibody (HUH23; 0.25 μg/ml in TTBS) for 1 h at room temperature. The primary antibody was a polyclonal rabbit antihuman 11?-HSD2 antibody (gift from Dr. Z. Krozowski, Baker Heart Research Institute, Melbourne, Victoria, Australia) (34). After three 5-min washes with TTBS, the filter was incubated with horseradish peroxidase-labeled second antibody and developed in enhanced chemiluminescence detection reagents. The filter was then exposed to x-ray film (Eastman Kodak Co., Rochester, NY) for 1–5 min. Densitometry was performed on the radiographs, and the level of 11?-HSD2 protein was expressed as arbitrary units.
Assessment of 11?-HSD2 mRNA: real-time quantitative RT-PCR
To determine whether changes in 11?-HSD2 activity after the different treatment regimens were associated with alterations in 11?-HSD2 mRNA levels, the relative abundance of 11?-HSD2 mRNA in trophoblast cells was assessed by a two-step real-time quantitative RT-PCR (qRT-PCR), as described previously (22).
Briefly, total RNA was extracted from cultured cells using an RNeasy Mini Kit (Qiagen, Mississauga, Canada) coupled with on-column deoxyribonuclease digestion with the ribonuclease-free deoxyribonuclease set (Qiagen) according to the manufacturer’s instructions. One half microgram of total RNA was reverse transcribed in a total volume of 20 μl using the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA) following the manufacturer’s instructions. For every RT reaction set, one RNA sample was set up without reverse transcriptase enzyme to provide a negative control. Gene transcript levels of 18S ribosomal RNA (rRNA; housekeeping gene) and 11?-HSD2 were quantified separately by TaqMan assays using the TaqMan Universal PCR Master Mix (Applied Biosystems) and the universal thermal cycling parameters (2 min at 50 C and 10 min at 95 C, followed by 40 cycles of 15 sec at 95 C and 1 min at 60 C) on an ABI PRISM 7900HT Sequence Detection System (Applied Biosystems). Levels of 18S rRNA were assessed using TaqMan rRNA control reagents (Applied Biosystems), and levels of 11?-HSD2 mRNA were determined using custom-designed TaqMan assays. Primers (60 nM each) and TaqMan MGB probe (200 nM) for human 11?-HSD2 were designed using Primer Express software (Applied Biosystems; Table 1), and their optimal concentrations were determined following guidelines developed for sequence detection systems by Applied Biosystems.
TABLE 1. Primer and probe sequences for qRT-PCR
Levels of 18S rRNA and 11?-HSD2 mRNA in each RNA sample were quantified using the relative standard curve method (Applied Biosystems). Briefly, standard curves for 18S rRNA and 11?-HSD2 were generated by performing a dilution series of the untreated control cDNA. For each RNA sample, the relative amounts of 18S rRNA and 11?-HSD2 mRNA were determined, and the ratio of 11?-HSD2 mRNA to 18S rRNA was calculated. For each experiment, the amount of 11?-HSD2 mRNA at any given time point under various treatment conditions is expressed relative to the amount of transcript present in the untreated control.
Assessment of de novo protein synthesis involvement
Trophoblast cells were cultured as outlined above and pretreated with the translational inhibitor, CHX (10 μM), for 1 h before the addition of cPGI2 (10 μM). At the end of 24-h treatment, the cells were harvested for RNA isolation and qRT-PCR analysis.
Assessment of 11?-HSD2 mRNA stability
The trophoblast cells were cultured as outlined above and treated with cPGI2 (10 μM) for 12 h. Transcription was then stopped with DRB (25 μM), and cells were harvested at discrete times (0–12 h) thereafter for RNA isolation and qRT-PCR analysis.
Transient transfection and reporter gene assay
To determine whether activation of PPAR altered the rate of HSD11B2 gene transcription, a standard reporter gene assay was performed, as previously described (22). Briefly, the isolated trophoblast cells were plated on 24-well plates and cultured under standard conditions for 48 h. The cells were then cotransfected with 0.8 μg/well pGL3-HSD11B2P (+330 bp) and 0.2 μg/well of a cytomegalovirus promoter (pCMV) ?-galactosidase plasmid (Promega Corp., Madison, WI), or with 0.8 μg/well pGL3-Basic and 0.2 μg/well pCMV-?-galactosidase (negative control). The HSD11B2 promoter construct was generated from the pGL3-HSD11B2P (+4.5 kb) (22) with the use of an unique XhoI site located 330 bp upstream of the translational start codon (Fig. 6A). Trophoblast cells were also cotransfected similarly with a dominant negative PPAR mutant construct (gift from Dr. Paul Grimaldi, Centre de Biochimie, Universite de Nice-Sophia Antipolis, Parc Valrose, Nice, France) (35). All transfections were carried out in serum-free medium 199 for 1 h using Transfast transfection reagent (Promega Corp.) at a ratio of 2:1 (transfection reagent to DNA) according to the manufacturer’s instructions. At the end of transfection, cells were gently overlaid with fresh medium containing 10% serum and incubated for 4 h. The medium was then replaced with fresh medium containing 2% serum, and the cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, luciferase and galactosidase activities were analyzed using the Luciferase Assay System (Promega Corp.) and the ?-Galactosidase Enzyme Assay System (Promega Corp.), respectively. Luciferase activity was measured using a Lumat LB 9507 luminometer (EG&G Berthold, Bad Wildbad, Germany) and normalized against ?-galactosidase activity. Each transfection was performed in triplicate, and a total of three to five independent experiments were carried out.
FIG. 6. Effects of cPGI2 on HSD11B2 promoter activity. A, Schematic representation of the reporter gene construct. Trophoblast cells were transfected with a luciferase reporter gene construct driven by the HSD11B2 promoter and pCMV ?-galactosidase (B) or were cotransfected with a dominant negative PPAR mutant construct (C), as described in Materials and Methods. Four hours after transfection, the cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, luciferase and ?-galactosidase activities were determined, and the ratio of luciferase activity to that of ?-galactosidase was calculated. Data are expressed as the mean ± SEM (n = 4). *, P < 0.05 vs. control.
Statistical analyses
Results are presented as the mean ± SEM of three to five independent experiments (i.e. tissues from different patients), as indicated. Statistical analyses of 11?-HSD2 activity data were performed using one-way ANOVA, followed by Tukey’s post hoc test. 11?-HSD2 protein, mRNA, and promoter activity data were analyzed by a standard t test. Significance was set at P < 0.05. Calculations were performed using SPSS software version 9.0 (SPSS, Inc., Chicago, IL).
Results
Expression of PPAR mRNA in cultured human placental trophoblasts
To verify the expression of PPAR in our cultured human placental trophoblast cells, we determined the presence of PPAR mRNA by a standard RT-PCR. As shown in Fig. 1, PPAR mRNA was detected in total RNA samples from both placental tissues and cultured syncytiotrophoblast cells. Consistent with the predominant syncytiotrophoblast localization, higher levels of PPAR mRNA were detected in cultured trophoblast cells than in placental tissues.
FIG. 1. Expression of PPAR mRNA in cultured placental trophoblast cells. Total RNA was extracted from placental villi at term and from cultured trophoblast cells. One microgram of RNA was used in a standard RT-PCR to amplify mRNA for human PPAR and GAPDH (used as an internal control), as described in Materials and Methods. A fraction of the RT-PCR products was subjected to electrophoresis on a 1.2% agarose gel. This figure shows the results of one representative experiment.
Effects of cPGI2 on 11?-HSD2 activity
To study the effects of cPGI2, a putative PPAR agonist, on placental 11?-HSD2 activity, isolated human trophoblast cells were treated with 10 μM cPGI2 for various times (6, 12, and 24 h). This treatment resulted in a time-dependent decrease in 11?-HSD2 activity, such that a significant reduction (P < 0.05) occurred after 6 h of treatment (Fig. 2A). Furthermore, the cells were also treated with various concentrations of cPGI2 (1–20 μM), and this led to a concentration-dependent decrease in 11?-HSD2 activity, with a maximal effect at 10 μM cPGI2 (Fig. 2B).
FIG. 2. Time- and concentration-dependent effects of cPGI2 on 11?-HSD2 activity. A, Time-dependent effects of cPGI2 on 11?-HSD2 activity. Trophoblast cells were incubated with or without 10 μM cPGI2 for various periods of time (6, 12, and 24 h). B, Concentration-dependent effects of cPGI2 on 11?-HSD2 activity. Trophoblast cells were treated for 24 h with increasing concentrations of cPGI2 (1–20 μM). At the end of treatment, the level of 11?-HSD2 activity in intact cells was determined by a standard radiometric conversion assay, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments, each performed in triplicate. *, P < 0.05 vs. control.
Mechanisms of cPGI2-induced decrease in 11?-HSD2 activity
To determine the signal transduction pathway by which cPGI2 decreases placental 11?-HSD2 activity, we treated the cells with H89, an inhibitor of protein kinase A (PKA). We reasoned that if the cPGI2-induced decrease in 11?-HSD2 activity was due to activation of the classical prostacyclin receptor (IP) treatment with H89 would abolish the effect of cPGI2. Although H89 alone caused a 40% reduction in the level of 11?-HSD2 activity, it failed to abrogate the inhibitory effect of cPGI2 (Fig. 3A). This suggested that the effect of cPGI2 on 11?-HSD2 was probably mediated through activation of PPAR.
FIG. 3. Effects of cPGI2 on 11?-HSD2 activity: involvement of PPAR. A, Effects of a PKA inhibitor. Trophoblast cells were treated for 24 h with cPGI2 (10 μM), H89 (10 μM), or the combination of cPGI2 and H89. B, Effects of indomethacin and a selective PPAR agonist. Trophoblast cells were treated for 24 h with indomethacin (10 μM), cPGI2 (10 μM), or GW610742 (10 μM). C, Effects of 9-cis-RA. Trophoblast cells were treated for 24 h with RA (100 nM), a submaximal concentration of cPGI2 (5 μM), or the combination of RA and cPGI2. At the end of treatment, the level of 11?-HSD2 activity in intact cells was determined by a radiometric conversion assay, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments, each performed in triplicate. *, P < 0.05 vs. control.
To provide additional support for the involvement of PPAR in down-regulating placental 11?-HSD2 activity, we used a selective PPAR agonist, GW610742. Treatment of the trophoblast cells with GW610742 resulted in a similar decrease (to that observed with cPGI2) in 11?-HSD2 activity. In addition, indomethacin, a known ligand for both PPAR and PPAR (36), had no effect (Fig. 3B).
Having established the involvement of PPAR in mediating the cPGI2-induced down-regulation of placental 11?-HSD2 activity, we conducted the next experiment to elucidate the molecular mechanisms by which cPGI2 reduces 11?-HSD2 activity. Given that PPARs, like many other members of the nuclear receptor superfamily, exert their effects by forming obligate heterodimers with the RXR, we sought to determine whether cPGI2 and the RXR ligand 9-cis-retinoid acid (9-cis-RA) would exert synergistic effects on 11?-HSD2 activity. As shown in Fig. 3C, 9-cis-RA alone did not alter 11?-HSD2 activity. Moreover, combined treatment with 9-cis-RA and a submaximal effective concentration of cPGI2 (5 μM) did not result in an additional decrease in 11?-HSD2 activity (Fig. 3C).
Effects of cPGI2 on 11?-HSD2 protein and mRNA
To determine whether the cPGI2-induced decrease in 11?-HSD2 activity was a result of reduced 11?-HSD2 expression, levels of 11?-HSD2 protein and mRNA were assessed by Western blot analysis and qRT-PCR, respectively. As shown in Fig. 4, treatment of trophoblast cells for 24 h with 10 μM cPGI2 led to a similar reduction in the levels of both 11?-HSD2 protein and mRNA.
FIG. 4. Effects of cPGI2 on 11?-HSD2 expression. Trophoblast cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, levels of 11?-HSD2 protein (B) and mRNA (C) were determined by Western blot analysis and qRT-PCR, respectively, as described in Materials and Methods. One representative autoradiograph is shown in A. Each bar represents the mean ± SEM of four independent experiments. *, P < 0.05 vs. control.
Effect of CHX on the cPGI2-induced decrease in 11?-HSD2 mRNA
To determine whether the cPGI2-induced decrease in 11?-HSD2 mRNA required de novo protein synthesis, trophoblast cells were treated with cPGI2 in the absence and presence of CHX, a protein synthesis inhibitor. We found that cPGI2 was equally effective in reducing 11?-HSD2 mRNA in the absence and presence of CHX, indicating that de novo protein synthesis was not required (Fig. 5A).
FIG. 5. Effects of protein and mRNA synthesis inhibitors on the cPGI2-induced decrease in 11?-HSD2 mRNA. A, Effects of CHX. Trophoblast cells were pretreated with CHX (10 μM) for 1 h, then treated in the absence or presence of cPGI2 (10 μM) for 24 h. At the end of treatment, total cellular RNA was isolated, and the steady state level of 11?-HSD2 mRNA was assessed by qRT-PCR, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments. *, P < 0.05 vs. control. B, Effects of DRB. Trophoblast cells were pretreated with () or without () 10 μM cPGI2 for 12 h. The cells were then treated with 25 μM DRB in the absence or presence of 10 μM cPGI2 (defined as time zero). At the indicated time points thereafter, total cellular RNA was isolated, and the steady state level of 11?-HSD2 mRNA was assessed by qRT-PCR, as described in Materials and Methods. Each data point is expressed as a percentage of the maximum determined at time zero. Data from one representative experiment are shown.
Effects of cPGI2 on 11?-HSD2 mRNA stability
As a first step in elucidating the molecular mechanisms by which cPGI2 decreases 11?-HSD2 mRNA, we assessed the half-life of 11?-HSD2 mRNA by a standard mRNA decay assay using 25 μM DRB, an inhibitor of mRNA synthesis. As shown in Fig. 5B, cPGI2 did not alter the half-life of 11?-HSD2 mRNA.
Effects of cPGI2 on HSD11B2 gene transcription
To determine whether cPGI2 reduces the rate of HSD11B2 gene transcription, trophoblasts were transiently transfected with a luciferase construct containing a 330-bp 5'-flanking region of the human HSD11B2 gene (Fig. 6A). Treatment with cPGI2 (10 μM) resulted in a significant decrease in HSD11B2 promoter activity (Fig. 6B). Moreover, the cPGI2-induced repression of HSD11B2 promoter activity was prevented in cells cotransfected with a dominant negative PPAR mutant (Fig. 6C), thus corroborating the idea that the effects of cPGI2 on placental 11?-HSD2 are mediated by the activation of PPAR.
Discussion
Placental 11?-HSD2 plays a pivotal role in controlling the precise level of fetal exposure to maternal glucocorticoids (4), and it also modulates local actions of glucocorticoids within the placenta (37, 38). Thus, 11?-HSD2 is an important regulator of placental function and fetal development. Indeed, a proper level of 11?-HSD2 in the human placenta appears to be critical for normal fetal development, because attenuated expression of this enzyme is associated with pregnancies complicated by IUGR (6, 21). Moreover, IUGR is a characteristic feature of 11?-HSD2 deficiency (7). Although a variety of hormones, cytokines, and intracellular signaling molecules as well as hypoxia (8, 9, 10, 11, 12, 13, 19, 20, 22) are known to modulate placental 11?-HSD2, the regulation of this important enzyme is incompletely understood.
Abundant PPAR mRNA and protein are expressed in human trophoblast cells (25, 39), but the function of this nuclear receptor in the human placenta is unknown. The present study was designed to investigate the role of PPAR in regulating 11?-HSD2 in cultured human trophoblast cells. As a first step, we sought to verify the expression of PPAR in our cultured cells. Using RT-PCR, we demonstrated that PPAR mRNA was not only present, but also expressed at a higher level, in purified trophoblast cells compared with whole tissue extracts. This observation is consistent with the previously reported predominant localization of PPAR to the syncytiotrophoblast layer of human placenta (39).
We then examined the effect of cPGI2, a putative PPAR agonist, on 11?-HSD2 activity. We showed that treatment of trophoblast cells with cPGI2 led to a time- and concentration-dependent decrease in the level of 11?-HSD2 activity. Furthermore, the cPGI2-induced decrease in 11?-HSD2 activity was a consequence of reduced 11?-HSD2 expression, because levels of both 11?-HSD2 protein and mRNA were decreased after cPGI2 treatment. Given that the actions of cPGI2 can be mediated by the classical G protein-coupled membrane IP receptor (40) or the ligand-activated nuclear receptor PPAR (41), we conducted a series of experiments to determine which of the two distinct signaling pathways was responsible for mediating the cPGI2-induced decrease in 11?-HSD2 activity.
With respect to the IP receptor-mediated signaling pathway, it is believed that binding of cPGI2 to the G protein-coupled IP receptor stimulates the production of cAMP, which, in turn, binds to the regulatory subunit of PKA, releasing the active catalytic subunit. Activated PKA modulates the function of various nuclear transcription factors that bind to DNA sequences present in the regulatory regions of cAMP-responsive genes (40). We showed previously that forskolin (an activator of PKA) increased 11?-HSD2 activity and mRNA in human placental trophoblast cells (10), implicating PKA as a positive regulator of placental 11?-HSD2. Given the opposite effects of cPGI2 and forskolin, we concluded that the cPGI2-induced reduction in 11?-HSD2 activity was unlikely to be due to the IP receptor activation. To provide experimental evidence for our contention, we treated trophoblast cells with the PKA inhibitor H89. Consistent with our previous report of stimulatory effects of forskolin on placental 11?-HSD2, treatment of trophoblast cells with H89 alone resulted in a reduction of 11?-HSD2 activity. This suggested that the PKA pathway might be involved in maintaining high levels of 11?-HSD2 expression in trophoblast cells. However, H89 was ineffective in blocking the inhibitory effect of cPGI2. Taken together, these findings suggested that the cPGI2-induced down-regulation of placental 11?-HSD2 was probably mediated by PPAR.
To provide additional evidence for the involvement of PPAR, we examined the effects of GW610742, a selective PPAR agonist, on 11?-HSD2 activity. Treatment of trophoblast cells with GW610742 resulted in a similar decrease in the level of 11?-HSD2 activity. In contrast, indomethacin, a promiscuous ligand for both PPAR and PPAR (36), had no effect. Taken together, these results indicated that the cPGI2-induced decrease in 11?-HSD2 activity was a result of activation of PPAR.
PPARs are ligand-activated transcription factors that exert their effects by forming heterodimers with RXR, binding to PPREs in the regulatory regions of target genes, and inducing ligand-dependent transactivation. Gene targeting studies in mice have demonstrated a critical role for RXR in placentation and fetal development (42, 43). Moreover, PPAR and RXR have been shown to colocalize in the syncytiotrophoblastic layer of the term human placenta (39). Therefore, we determined whether these two cognate receptors would interact to regulate 11?-HSD2 in cultured trophoblast cells. We treated trophoblast cells with the pan-RXR ligand 9-cis-RA in the presence and absence of the putative PPAR agonist cPGI2. Our results showed that 9-cis-RA alone had no effect on 11?-HSD2 activity. Moreover, 9-cis-RA did not augment the inhibitory effect of a submaximal concentration of cPGI2 on 11?-HSD2 activity, suggesting that the cPGI2-induced decrease in 11?-HSD2 activity was unlikely to be due to interactions between PPAR and RXR through a PPRE. Consistent with this idea, there are no apparent PPRE consensus motifs in the 330-bp 5'-flanking region of human HSD11B2 gene. Recently, PPAR and PPAR have been reported to repress gene transcription by restricting the binding of specific transcription factors to respective response elements (44) or by competing with critical coactivators necessary for transcriptional activation (45). In addition, there is evidence that PPAR exerts transcriptional repression via interactions with corepressor proteins (46, 47). Pertinent to our findings, both PPAR and PPAR have been shown to interact with specificity protein-1 (Sp1), causing transcriptional suppression of vascular endothelial growth factor receptor-2 gene (48) and thromboxane receptor gene (49), respectively. Given that the human HSD11B2 gene contains two Sp1-binding sites within the 330-bp 5'-flanking region (50, 51), it is tempting to speculate that PPAR may repress HSD11B2 gene transcription via an interaction with Sp1 in human trophoblast cells. Additional studies are currently being conducted to examine this possibility.
The observed decrease in the level of 11?-HSD2 mRNA in cultured trophoblast cells after treatment with cPGI2 may be attributed to a direct effect of cPGI2 on 11?-HSD2 or an indirect effect involving the synthesis of an intermediary protein. To determine whether the cPGI2-induced reduction of 11?-HSD2 mRNA required de novo protein synthesis, we treated trophoblast cells with cPGI2 in the absence and presence of CHX, a protein synthesis inhibitor. Although CHX alone decreased the basal level of 11?-HSD2 mRNA (a nonspecific effect probably caused by a decrease in the levels of various proteins that are required to sustain the general cell transcriptional machinery), it did not block the cPGI2-induced decrease in 11?-HSD2 mRNA, suggesting that de novo protein synthesis was not required.
In theory, a decrease in the steady state mRNA level of a given gene can be achieved by reducing the rate of gene transcription and/or by decreasing the mRNA stability. To determine which of these mechanisms was responsible for the observed decrease in 11?-HSD2 mRNA, we treated trophoblast cells with the mRNA synthesis inhibitor DRB in the absence and presence of cPGI2 and determined the rate of 11?-HSD2 mRNA decay by qRT-PCR. Under the conditions of the present study, cPGI2 did not affect the half-life of 11?-HSD2 mRNA, suggesting that the cPGI2-induced decrease in 11?-HSD2 mRNA was probably mediated at the level of HSD11B2 gene transcription. To provide direct evidence for this contention, we examine the effect of cPGI2 on HSD11B2 promoter activity. Our results showed that cPGI2 reduced HSD11B2 promoter activity, and that this effect was abolished in trophoblast cells expressing a dominant negative PPAR mutant. Taken together, these data suggest that activation of PPAR suppresses HSD11B2 gene transcription. Thus, our present findings provide additional evidence for PPAR-mediated transcriptional suppression.
The demonstration of PPAR-induced suppression of placental HSD11B2 gene expression will probably have far-reaching implications for our understanding of the role of this enzyme in normal and pathological pregnancies. We believe that the precise level of 11?-HSD2 in the human placenta is tightly controlled by a balance between stimulatory factors, such as retinoic acids (11), glucocorticoids (22), and activators of the PKA pathway (8), and inhibitory factors, such as prostaglandins (13), hypoxia (19, 20), and PPAR. Thus, it is conceivable that aberrant activities of PPAR, at the level of receptor itself, its ligands, and/or its coregulatory proteins (i.e. coactivators and corepressors), in the human placenta may lead to an altered expression of 11?-HSD2 and, consequently, abnormal fetal development. Indeed, PPAR-null mice exhibited placental defects and IUGR phenotype (32). Given the previous findings that the attenuated placental 11?-HSD2 expression in human pregnancies complicated with IUGR was not due to imprinting or mutations in the HSD11B2 gene (21), it is attempting to speculate that overactivation of PPAR may be a contributing factor in these pathological pregnancies.
In conclusion, the present study demonstrates that cPGI2 decreases 11?-HSD2 activity and expression in cultured human placental trophoblast cells, and that this effect is probably mediated by the nuclear receptor PPAR, which functions to repress HSD11B2 gene transcription. If PPAR reduces HSD11B2 gene expression in the human placenta in vivo, our findings would provide a molecular mechanism by which this nuclear receptor may regulate human placental function and, consequently, fetal development.
Acknowledgments
We thank Dr. Daniel Hardy for technical assistance, and Dr. Joseph Torchia for insightful suggestions.
References
Cole TJ, Blendy JA, Monaghan AP, Krieglstein K, Schmid W, Aguzzi A, Fantuzzi G, Hummler E, Unsicker K, Schutz G 1995 Targeted disruption of the glucocorticoid receptor gene blocks adrenergic chromaffin cell development and severely retards lung maturation. Genes Dev 9:1608–1621
Seckl JR 1994 Glucocorticoids and small babies. Q J Med 87:259–262
Seckl JR 2001 Glucocorticoid programming of the fetus; adult phenotypes and molecular mechanisms. Mol Cell Endocrinol 185:61–71
Yang K 1997 Placental 11?-hydroxysteroid dehydrogenase: barrier to maternal glucocorticoids. Rev Reprod 2:129–132
Stewart PM, Rogerson FM, Mason JI 1995 Type 2 11?-hydroxysteroid dehydrogenase messenger ribonucleic acid and activity in human placenta and fetal membranes: its relationship to birth weight and putative role in fetal adrenal steroidogenesis. J Clin Endocrinol Metab 80:885–890
Shams M, Kilby MD, Somerset DA, Howie AJ, Gupta A, Wood PJ, Afnan M, Stewart PM 1998 11?-Hydroxysteroid dehydrogenase type 2 in human pregnancy and reduced expression in intrauterine growth restriction. Hum Reprod 13:799–804
Kitanaka S, Tanae A, Hibi I 1996 Apparent mineralocorticoid excess due to 11?-hydroxysteroid dehydrogenase deficiency: a possible cause of intrauterine growth retardation. Clin Endocrinol (Oxf) 44:353–359
Pasquarette MM, Stewart PM, Ricketts ML, Imaishi K, Mason JI 1996 Regulation of 11?-hydroxysteroid dehydrogenase type 2 activity and mRNA in human choriocarcinoma cells. J Mol Endocrinol 16:269–275
Sun K, Yang K, Challis JR 1997 Differential regulation of 11?-hydroxysteroid dehydrogenase type 1 and 2 by nitric oxide in cultured human placental trophoblast and chorionic cell preparation. Endocrinology 138:4912–4920
Sun K, Yang K, Challis JR 1998 Regulation of 11?-hydroxysteroid dehydrogenase type 2 by progesterone, estrogen, and the cyclic adenosine 5'-monophosphate pathway in cultured human placental and chorionic trophoblasts. Biol Reprod 58:1379–1384
Tremblay J, Hardy DB, Pereira LE, Yang K 1999 Retinoic acid stimulates the expression of 11?-hydroxysteroid dehydrogenase type 2 in human choriocarcinoma JEG-3 cells. Biol Reprod 60:541–545
Sarkar S, Tsai SW, Nguyen TT, Plevyak M, Padbury JF, Rubin LP 2001 Inhibition of placental 11?-hydroxysteroid dehydrogenase type 2 by catecholamines via -adrenergic signaling. Am J Physiol 281:R1966–R1974
Hardy DB, Pereria LE, Yang K 1999 Prostaglandins and leukotriene B4 are potent inhibitors of 11?-hydroxysteroid dehydrogenase type 2 activity in human choriocarcinoma JEG-3 cells. Biol Reprod 61:40–45
Hardy DB, Dixon SJ, Narayanan N, Yang K 2001 Calcium inhibits human placental 11?-hydroxysteroid dehydrogenase type 2 activity. Biochem Biophys Res Commun 283:756–761
Pepe GJ, Waddell BJ, Stahl SJ, Albrecht ED 1988 The regulation of transplacental cortisol-cortisone metabolism by estrogen in pregnant baboons. Endocrinology 122:78–83
Baggia S, Albrecht ED, Pepe GJ 1990 Regulation of 11?-hydroxysteroid dehydrogenase activity in the baboon placenta by estrogen. Endocrinology 126:2742–2748
Pepe GJ, Davies WA, Dong KW, Luo H, Albrecht ED 1999 Cloning of the 11?-hydroxysteroid dehydrogenase (11?-HSD)-2 gene in the baboon: effects of estradiol on promoter activity of 11?-HSD-1 and -2 in placental JEG-3 cells. Biochim Biophys Acta 1444:101–110
Pepe GJ, Burch MG, Albrecht ED 2001 Estrogen regulates 11?-hydroxysteroid dehydrogenase-1 and -2 localization in placental syncytiotrophoblast in the second half of primate pregnancy. Endocrinology 142:4496–4503
Alfaidy N, Gupta S, DeMarco C, Caniggia I, Challis JR 2002 Oxygen regulation of placental 11?-hydroxysteroid dehydrogenase 2: physiological and pathological implications. J Clin Endocrinol Metab 87:4797–4805
Hardy DB, Yang K 2002 The expression of 11?-hydroxysteroid dehydrogenase type 2 is induced during trophoblast differentiation: effects of hypoxia. J Clin Endocrinol Metab 87:3696–3701
McTernan CL, Draper N, Nicholson H, Chalder SM, Driver P, Hewison M, Kilby MD, Stewart PM 2001 Reduced placental 11?-hydroxysteroid dehydrogenase type 2 mRNA levels in human pregnancies complicated by intrauterine growth restriction: an analysis of possible mechanisms. J Clin Endocrinol Metab 86:4979–4983
van Beek JP, Guan H, Julan L, Yang K 2004 Glucocorticoids stimulate the expression of 11?-hydroxysteroid dehydrogenase type 2 in cultured human placental trophoblast cells. J Clin Endocrinol Metab 89:5614–5621
Desvergne B, Wahli W 1999 Peroxisome proliferator-activated receptors: nuclear control of metabolism. Endocr Rev 20:649–688
Lee CH, Olson P, Evans RM 2003 Minireview: lipid metabolism, metabolic diseases, and peroxisome proliferator-activated receptors. Endocrinology 144:2201–2207
Mukherjee R, Jow L, Croston GE, Paterniti Jr JR 1997 Identification, characterization, and tissue distribution of human peroxisome proliferator-activated receptor (PPAR) isoforms PPAR2 versus PPAR1 and activation with retinoid X receptor agonists and antagonists. J Biol Chem 272:8071–8076
Lim H, Gupta RA, Ma WG, Paria BC, Moller DE, Morrow JD, DuBois RN, Trzaskos JM, Dey SK 1999 Cyclo-oxygenase-2-derived prostacyclin mediates embryo implantation in the mouse via PPAR. Genes Dev 13:1561–1574
Matsuura H, Adachi H, Smart RC, Xu X, Arata J, Jetten AM 1999 Correlation between expression of peroxisome proliferator-activated receptor ? and squamous differentiation in epidermal and tracheobronchial epithelial cells. Mol Cell Endocrinol 147:85–92
Gupta RA, Tan J, Krause WF, Geraci MW, Willson TM, Dey SK, DuBois RN 2000 Prostacyclin-mediated activation of peroxisome proliferator-activated receptor in colorectal cancer. Proc Natl Acad Sci USA 97:13275–13280
Hansen JB, Zhang H, Rasmussen TH, Petersen RK, Flindt EN, Kristiansen K 2001 Peroxisome proliferator-activated receptor (PPAR)-mediated regulation of preadipocyte proliferation and gene expression is dependent on cAMP signaling. J Biol Chem 276:3175–3182
Wang YX, Lee CH, Tiep S, Yu RT, Ham J, Kang H, Evans RM 2003 Peroxisome-proliferator-activated receptor activates fat metabolism to prevent obesity. Cell 113:159–170
Peters JM, Lee SS, Li W, Ward JM, Gavrilova O, Everett C, Reitman ML, Hudson LD, Gonzalez FJ 2000 Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor ?(). Mol Cell Biol 20:5119–5128
Barak Y, Liao D, He W, Ong ES, Nelson MC, Olefsky JM, Boland R, Evans RM 2002 Effects of peroxisome proliferator-activated receptor on placentation, adiposity, and colorectal cancer. Proc Natl Acad Sci USA 99:303–308
Kliman HJ, Nestler JE, Sermasi E, Sanger JM, Strauss III JF 1986 Purification, characterization, and in vitro differentiation of cytotrophoblasts from human term placentae. Endocrinology 118:1567–1582
Krozowski Z, MaGuire JA, Stein-Oakley AN, Dowling J, Smith RE, Andrews RK 1995 Immunohistochemical localization of the 11?-hydroxysteroid dehydrogenase type II enzyme in human kidney and placenta. J Clin Endocrinol Metab 80:2203–2209
Bastie C, Luquet S, Holst D, Jehl-Pietri C, Grimaldi PA 2000 Alterations of peroxisome proliferator-activated receptor activity affect fatty acid-controlled adipose differentiation. J Biol Chem 275:38768–38773
Lehmann JM, Lenhard JM, Oliver BB, Ringold GM, Kliewer SA 1997 Peroxisome proliferator-activated receptors and are activated by indomethacin and other non-steroidal anti-inflammatory drugs. J Biol Chem 272:3406–3410
Sun K, Yang K, Challis JR 1998 Glucocorticoid actions and metabolism in pregnancy: implications for placental function and fetal cardiovascular activity. Placenta 19:353–360
Burton PJ, Waddell BJ 1999 Dual function of 11?-hydroxysteroid dehydrogenase in placenta: modulating placental glucocorticoid passage and local steroid action. Biol Reprod 60:234–240
Wang Q, Fujii H, Knipp GT 2002 Expression of PPAR and RXR isoforms in the developing rat and human term placentas. Placenta 23:661–671
Namba T, Oida H, Sugimoto Y, Kakizuka A, Negishi M, Ichikawa A, Narumiya S 1994 cDNA cloning of a mouse prostacyclin receptor. Multiple signaling pathways and expression in thymic medulla. J Biol Chem 269:9986–9992
Forman BM, Chen J, Evans RM 1997 Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors and . Proc Natl Acad Sci USA 94:4312–4317
Sapin V, Dolle P, Hindelang C, Kastner P, Chambon P 1997 Defects of the chorioallantoic placenta in mouse RXR null fetuses. Dev Biol 191:29–41
Wendling O, Chambon P, Mark M 1999 Retinoid X receptors are essential for early mouse development and placentogenesis. Proc Natl Acad Sci USA 96:547–551
Delerive P, De Bosscher K, Besnard S, Vanden Berghe W, Peters JM, Gonzalez FJ, Fruchart JC, Tedgui A, Haegeman G, Staels B 1999 Peroxisome proliferator-activated receptor negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-B and AP-1. J Biol Chem 274:32048–32054
Li M, Pascual G, Glass CK 2000 Peroxisome proliferator-activated receptor -dependent repression of the inducible nitric oxide synthase gene. Mol Cell Biol 20:4699–4707
Shi Y, Hon M, Evans RM 2002 The peroxisome proliferator-activated receptor , an integrator of transcriptional repression and nuclear receptor signaling. Proc Natl Acad Sci USA 99:2613–2618
Lee CH, Chawla A, Urbiztondo N, Liao D, Boisvert WA, Evans RM, Curtiss LK 2003 Transcriptional repression of atherogenic inflammation: modulation by PPAR. Science 302:453–457
Meissner M, Stein M, Urbich C, Reisinger K, Suske G, Staels B, Kaufmann R, Gille J 2004 PPAR activators inhibit vascular endothelial growth factor receptor-2 expression by repressing Sp1-dependent DNA binding and transactivation. Circ Res 94:324–332
Sugawara A, Uruno A, Kudo M, Ikeda Y, Sato K, Taniyama Y, Ito S, Takeuchi K 2002 Transcription suppression of thromboxane receptor gene by peroxisome proliferator-activated receptor- via an interaction with Sp1 in vascular smooth muscle cells. J Biol Chem 277:9676–9683
Agarwal AK, White PC 1996 Analysis of the promoter of the NAD+ dependent 11?-hydroxysteroid dehydrogenase (HSD11K) gene in JEG-3 human choriocarcinoma cells. Mol Cell Endocrinol 121:93–99
Nawrocki AR, Goldring CE, Kostadinova RM, Frey FJ, Frey BM 2002 In vivo footprinting of the human 11?-hydroxysteroid dehydrogenase type 2 promoter: evidence for cell-specific regulation by Sp1 and Sp3. J Biol Chem 277:14647–14656(Laura Julan, Haiyan Guan,)
Address all correspondence and requests for reprints to: Dr. K. Yang, Children’s Health Research Institute, Room A5-132, Victoria Research Laboratories-Westminster Campus, 800 Commissioners Road East, London, Ontario, Canada N6A 4G5. E-mail: kyang@uwo.ca.
Abstract
Accumulating evidence suggests that the human placental enzyme 11?-hydroxysteroid dehydrogenase type 2 (11?-HSD2) plays a key role in fetal development by controlling fetal exposure to maternal glucocorticoids. Recently, the nuclear peroxisome proliferator-activated receptor (PPAR) has been found to be the most abundantly expressed PPAR subtype in the human placenta, but its function in this organ is unknown. Given that PPAR-null mice exhibited placental defects and consequent intrauterine growth restriction, the present study was undertaken to examine the hypothesis that PPAR regulates human placental function in part by targeting 11?-HSD2. Using cultured human trophoblast cells as a model system, we demonstrated that 1) the putative PPAR agonist carbaprostacyclin (cPGI2) reduced 11?-HSD2 activity as well as 11?-HSD2 expression at both protein and mRNA levels; 2) GW610742 (a selective PPAR agonist) mimicked the effect of cPGI2, whereas indomethacin (a known ligand for PPAR and PPAR) had no effect; 3) the cPGI2-induced down-regulation of 11?-HSD2 mRNA did not require de novo protein synthesis; 4) cPGI2 suppressed HSD11B2 promoter activity, but did not alter the half-life of 11?-HSD2 mRNA; and 5) the inhibitory effect of cPGI2 on HSD11B2 promoter activity was abrogated in trophoblast cells cotransfected with a dominant negative PPAR mutant. Taken together, these findings suggest that activation of PPAR down-regulates HSD11B2 gene expression in human trophoblast cells, and that this effect is mediated primarily at the transcriptional level. Thus, the present study reveals 11?-HSD2 as an additional target for PPAR and identifies a molecular mechanism by which this nuclear receptor may regulate human placental function.
Introduction
GLUCOCORTICOIDS ARE ESSENTIAL for normal fetal organ growth and maturation (1). However, excessive glucocorticoid exposure in utero leads to intrauterine growth restriction (IUGR) and may predispose to type 2 diabetes and hypertension later in life (2, 3). The placental enzyme 11?-hydroxysteroid dehydrogenase type 2 (11?-HSD2) is believed to play a key role in protecting the fetus from exposure to high levels of maternal glucocorticoid by converting maternal cortisol to its inactive metabolite, cortisone (4). Indeed, placental 11?-HSD2 activity is correlated with birth weight (5) and is attenuated in pregnancies complicated with IUGR (6). Moreover, IUGR is a characteristic feature of apparent mineralocorticoid excess syndrome, an autosomal recessive disorder in which point mutations in the HSD11B2 gene render the enzyme inactive (7). Thus, the precise control of placental 11?-HSD2 expression and activity appears to be critical to normal fetal development.
Previous studies have demonstrated that retinoic acid and forskolin increase, whereas nitric oxide, progesterone, and catecholamines decrease 11?-HSD2 activity and mRNA levels in JEG-3 choriocarcinoma cells and cultured human placental trophoblasts (8, 9, 10, 11, 12). Furthermore, placental 11?-HSD2 activity, but not mRNA, is reduced by prostaglandins, leukotriene B4, and calcium (13, 14). In addition, a series of elegant studies by Albrecht, Pepe, and colleagues (15, 16, 17, 18) has implicated estrogen as a critical factor in maintaining the developmental pattern of expression of 11?-HSD2 in the baboon placenta. More recently, we and others have provided evidence that hypoxia is a negative regulator of human placental 11?-HSD2 expression and activity (19, 20). However, our understanding of placental 11?-HSD2 regulation is incomplete, particularly at the level of HSD11B2 gene transcription.
There is a good correlation between placental 11?-HSD2 activity and mRNA in both uncomplicated pregnancies and those complicated by IUGR (5, 21), suggesting that regulation of 11?-HSD2 in the human placenta may occur at a transcriptional level. Indeed, we have demonstrated recently that glucocorticoids stimulate HSD11B2 gene transcription in cultured human trophoblast cells (22). Peroxisome proliferator-activated receptors (PPARs) are members of the nuclear receptor superfamily. Three distinct PPAR subtypes have been identified, namely, PPAR, PPAR (also known as PPAR?), and PPAR, with unique tissue distributions and physiological functions. They act as regulatory transcription factors that heterodimerize with retinoid X receptors (RXR), bind to peroxisome proliferator-responsive elements (PPREs) located in the regulatory region of target genes, and modulate gene expression in response to ligand activation (23). Although PPAR and PPAR have been studied extensively, relatively less is known about PPAR (24). PPAR is widely expressed, but some tissues, such as the brain, skin, colon, and placenta, express higher levels (25). Recent studies have implicated PPAR in embryo implantation (26), epidermal maturation (27), colon cancer (28), preadipocyte proliferation (29), and obesity (30). Homozygous PPAR-null mice exhibit placental defects and reduced birth weight (31, 32), indicating a pivotal role for this nuclear receptor in murine placental function and fetal development. However, the role of PPAR in human fetal development is unknown. Given the particularly high level of PPAR expression in the human placenta (25), we hypothesized that this nuclear receptor may regulate human placental function in part by targeting 11?-HSD2. In the present study we tested this hypothesis using cultured human placental trophoblast cells as a model system. We present the first evidence that the activation of PPAR suppresses HSD11B2 gene transcription, leading to reduced 11?-HSD2 activity and expression in human trophoblast cells.
Materials and Methods
Reagents and supplies
[1,2,6,7-N-3H]Cortisol (80 Ci/mmol) was purchased from DuPont Canada, Inc. (Markam, Canada). Nonradioactive cortisol and cortisone were obtained from Steraloids, Inc. (Wilton, NH). Cycloheximide (CHX) and 5,6-dichlorobenzimidazole 1?-D-ribfuranoside (DRB) were purchased from Sigma-Aldrich Canada Ltd. (Oakville, Canada). Carbaprostacyclin (cPGI2) was obtained from Cayman Chemical Co. (Ann Arbor, MI), and GW610742 was a gift from GlaxoSmithKline (Research Triangle Park, NC). Polyester-backed thin layer chromatography plates were obtained from Fisher Scientific Ltd. (Nepean, Canada). All solvents used were purchased from VWR Canlab (Mississauga, Canada). Cell culture supplies were obtained from either Invitrogen Life Technologies, Inc. (Burlington, Canada) or Fisher Scientific. General molecular biology reagents were purchased from Invitrogen Life Technologies, Inc., or Pharmacia Canada, Inc. (Baie d’Urfe, Canada). Oligonucleotides were synthesized by a Pharmacia Gene Assembler and purified with NAP-50 columns (Pharmacia Biotech, Piscataway, NJ) according to the manufacturer’s instructions.
Placental trophoblast cell cultures
Placental trophoblast cells were isolated using a modification of the method of Kliman (33) as previously described (20). Ethics approval for procurement of human placentas was obtained from the University of Western Ontario ethics board for health sciences research involving human subjects. Briefly, human placentas were obtained from uncomplicated pregnancies at term after elective cesarean section. Villous tissues were dissected free from fetal membranes and blood vessels, rinsed in 0.9% NaCl2, and digested with 0.125% trypsin and 0.02% deoxyribonuclease I (Sigma-Aldrich Canada Ltd.) in DMEM containing 0.05% streptomycin and gentamicin (Invitrogen Life Technologies, Inc.) three times for 30 min each time. The placental cells were loaded onto a 5–70% Percoll gradient at step increments of 5% Percoll and centrifuged at 2500 x g for 20 min to separate different cell types. Cytotrophoblasts between the density markers of 1.049 and 1.062 g/ml were collected and plated in either 24-well plates (for enzyme activity assay) or 35-mm dishes (for Western blot analysis) at a density of 1.35 x 106 cells/ml in medium 199 containing 10% fetal calf serum (Invitrogen Life Technologies, Inc.). The cells were maintained at 37 C in humidified 5% CO2-95% air (20% O2) for 48 h. We have shown previously that the isolated cytotrophoblasts will differentiate into syncytiotrophoblasts over 48 h of culture under the conditions used in the present study. After 48 h, the trophoblast cells (in triplicate) were treated for 24 h (or as indicated otherwise) with various compounds in medium containing 2% fetal calf serum. Controls, also performed in triplicate, received an equivalent volume of vehicle (ethanol or dimethylsulfoxide).
RT-PCR
To verify the expression of PPAR in cultured human placental trophoblasts, the relative abundance of PPAR mRNA was assessed by a standard RT-PCR, as described previously (11). Briefly, total RNA was extracted from cultured trophoblast cells as well as human placental tissues using TRIzol reagent (Invitrogen Life Technologies, Inc.) according to the manufacturer’s instructions. One microgram of total RNA was reverse transcribed using a standard oligo(deoxythymidine) primer in a total volume of 20 μl. An aliquot (2 μl) of the RT reaction was then subjected to a standard PCR (94 C, 30 sec; 55 C, 30 sec; 72 C, 45 sec; 30 cycles) using sequence-specific primers (forward primer, 5'-GGT GAA TGG CCT GCC TCC CTA CAA; reverse primer, 5'-CAC AGA ATG ATG GCC GCA ATG AAT), which correspond to nucleotides 1029–1052 and 1409–1386 in the published human PPAR cDNA (GenBank accession no. BC007578), respectively. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as control. The same PCR conditions were used for GAPDH, except that the annealing temperature was 56 C, and 28 cycles were used. The primers for GAPDH (forward primer, 5'-ATC ACT GCC ACC CAG AAG AC; reverse primer, 5'-TGT GAG GGA GAT GCT CAG TG) correspond to nucleotides 565–584 and 1126–1145 in the published GAPDH cDNA (GenBank accession no. M17701). A fraction of the PPAR and GAPDH RT-PCR products was subjected to electrophoresis on a 1.2% agarose gel, stained with ethidium bromide, and photographed.
Assay of 11?-HSD2 activity: radiometric conversion assay
The level of 11?-HSD2 activity in intact cells at various time points and after different treatment regimens was determined by measuring the rate of cortisol to cortisone conversion, as described previously (20). Briefly, the cells were incubated for 1 h at 37 C in serum-free medium containing approximately 50,000 cpm [3H]cortisol and 100 nM unlabeled cortisol. At the end of the incubation, medium was collected, and steroids were extracted. The extracts were dried, and the residues were resuspended. A fraction of the resuspension was spotted on a thin layer chromatography plate that was developed in chloroform/methanol (9:1, vol/vol). The bands containing the labeled cortisol and cortisone were identified by UV light of the cold carriers, cut out into scintillation vials, and counted in Scintisafe Econo 1 (Fisher Scientific). The rate of cortisol to cortisone conversion was calculated, and the blank values (defined as the amount of conversion in the absence of cells) were subtracted and expressed as a percentage of control. Results are shown as the mean ± SEM.
Protein extraction and Western blot analysis
Cells were lysed with cold lysis buffer (100 mM NaCl, 50 mM sodium fluoride, 0.1% sodium dodecyl sulfate, 1% sodium deoxycholate, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 0.1 mM phenylmethylsulfonylfluoride, 1 mM orthovanadate, and 50 mM Tris-HCl, pH 7.5) for 5 min at room temperature. The total cell lysates were collected with a cell scraper, vortexed vigorously, and centrifuged at 10,000 x g for 20 min at 4 C. The supernatant was collected, and the protein content was determined by the Bradford method using a protein assay kit (Bio-Rad Laboratories, Inc., Missasauga, Canada) with BSA as a standard.
Western blot analysis was conducted as described previously (20). Briefly, 30 μg protein extracts were subjected to a standard 12% SDS-PAGE. After electrophoresis, proteins were transferred to nitrocellulose using a Bio-Rad Mini Transfer Apparatus. 11?-HSD2 protein was detected on the nitrocellulose filter using an enhanced chemiluminescence Western blotting analysis system (Pharmacia Biotech) following the manufacturer’s instructions. Briefly, the nitrocellulose filter was blocked overnight at 4 C with 10% Blotto in 0.1% Tween 20 in Tris-buffered saline (TTBS), and incubated with primary antibody (HUH23; 0.25 μg/ml in TTBS) for 1 h at room temperature. The primary antibody was a polyclonal rabbit antihuman 11?-HSD2 antibody (gift from Dr. Z. Krozowski, Baker Heart Research Institute, Melbourne, Victoria, Australia) (34). After three 5-min washes with TTBS, the filter was incubated with horseradish peroxidase-labeled second antibody and developed in enhanced chemiluminescence detection reagents. The filter was then exposed to x-ray film (Eastman Kodak Co., Rochester, NY) for 1–5 min. Densitometry was performed on the radiographs, and the level of 11?-HSD2 protein was expressed as arbitrary units.
Assessment of 11?-HSD2 mRNA: real-time quantitative RT-PCR
To determine whether changes in 11?-HSD2 activity after the different treatment regimens were associated with alterations in 11?-HSD2 mRNA levels, the relative abundance of 11?-HSD2 mRNA in trophoblast cells was assessed by a two-step real-time quantitative RT-PCR (qRT-PCR), as described previously (22).
Briefly, total RNA was extracted from cultured cells using an RNeasy Mini Kit (Qiagen, Mississauga, Canada) coupled with on-column deoxyribonuclease digestion with the ribonuclease-free deoxyribonuclease set (Qiagen) according to the manufacturer’s instructions. One half microgram of total RNA was reverse transcribed in a total volume of 20 μl using the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA) following the manufacturer’s instructions. For every RT reaction set, one RNA sample was set up without reverse transcriptase enzyme to provide a negative control. Gene transcript levels of 18S ribosomal RNA (rRNA; housekeeping gene) and 11?-HSD2 were quantified separately by TaqMan assays using the TaqMan Universal PCR Master Mix (Applied Biosystems) and the universal thermal cycling parameters (2 min at 50 C and 10 min at 95 C, followed by 40 cycles of 15 sec at 95 C and 1 min at 60 C) on an ABI PRISM 7900HT Sequence Detection System (Applied Biosystems). Levels of 18S rRNA were assessed using TaqMan rRNA control reagents (Applied Biosystems), and levels of 11?-HSD2 mRNA were determined using custom-designed TaqMan assays. Primers (60 nM each) and TaqMan MGB probe (200 nM) for human 11?-HSD2 were designed using Primer Express software (Applied Biosystems; Table 1), and their optimal concentrations were determined following guidelines developed for sequence detection systems by Applied Biosystems.
TABLE 1. Primer and probe sequences for qRT-PCR
Levels of 18S rRNA and 11?-HSD2 mRNA in each RNA sample were quantified using the relative standard curve method (Applied Biosystems). Briefly, standard curves for 18S rRNA and 11?-HSD2 were generated by performing a dilution series of the untreated control cDNA. For each RNA sample, the relative amounts of 18S rRNA and 11?-HSD2 mRNA were determined, and the ratio of 11?-HSD2 mRNA to 18S rRNA was calculated. For each experiment, the amount of 11?-HSD2 mRNA at any given time point under various treatment conditions is expressed relative to the amount of transcript present in the untreated control.
Assessment of de novo protein synthesis involvement
Trophoblast cells were cultured as outlined above and pretreated with the translational inhibitor, CHX (10 μM), for 1 h before the addition of cPGI2 (10 μM). At the end of 24-h treatment, the cells were harvested for RNA isolation and qRT-PCR analysis.
Assessment of 11?-HSD2 mRNA stability
The trophoblast cells were cultured as outlined above and treated with cPGI2 (10 μM) for 12 h. Transcription was then stopped with DRB (25 μM), and cells were harvested at discrete times (0–12 h) thereafter for RNA isolation and qRT-PCR analysis.
Transient transfection and reporter gene assay
To determine whether activation of PPAR altered the rate of HSD11B2 gene transcription, a standard reporter gene assay was performed, as previously described (22). Briefly, the isolated trophoblast cells were plated on 24-well plates and cultured under standard conditions for 48 h. The cells were then cotransfected with 0.8 μg/well pGL3-HSD11B2P (+330 bp) and 0.2 μg/well of a cytomegalovirus promoter (pCMV) ?-galactosidase plasmid (Promega Corp., Madison, WI), or with 0.8 μg/well pGL3-Basic and 0.2 μg/well pCMV-?-galactosidase (negative control). The HSD11B2 promoter construct was generated from the pGL3-HSD11B2P (+4.5 kb) (22) with the use of an unique XhoI site located 330 bp upstream of the translational start codon (Fig. 6A). Trophoblast cells were also cotransfected similarly with a dominant negative PPAR mutant construct (gift from Dr. Paul Grimaldi, Centre de Biochimie, Universite de Nice-Sophia Antipolis, Parc Valrose, Nice, France) (35). All transfections were carried out in serum-free medium 199 for 1 h using Transfast transfection reagent (Promega Corp.) at a ratio of 2:1 (transfection reagent to DNA) according to the manufacturer’s instructions. At the end of transfection, cells were gently overlaid with fresh medium containing 10% serum and incubated for 4 h. The medium was then replaced with fresh medium containing 2% serum, and the cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, luciferase and galactosidase activities were analyzed using the Luciferase Assay System (Promega Corp.) and the ?-Galactosidase Enzyme Assay System (Promega Corp.), respectively. Luciferase activity was measured using a Lumat LB 9507 luminometer (EG&G Berthold, Bad Wildbad, Germany) and normalized against ?-galactosidase activity. Each transfection was performed in triplicate, and a total of three to five independent experiments were carried out.
FIG. 6. Effects of cPGI2 on HSD11B2 promoter activity. A, Schematic representation of the reporter gene construct. Trophoblast cells were transfected with a luciferase reporter gene construct driven by the HSD11B2 promoter and pCMV ?-galactosidase (B) or were cotransfected with a dominant negative PPAR mutant construct (C), as described in Materials and Methods. Four hours after transfection, the cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, luciferase and ?-galactosidase activities were determined, and the ratio of luciferase activity to that of ?-galactosidase was calculated. Data are expressed as the mean ± SEM (n = 4). *, P < 0.05 vs. control.
Statistical analyses
Results are presented as the mean ± SEM of three to five independent experiments (i.e. tissues from different patients), as indicated. Statistical analyses of 11?-HSD2 activity data were performed using one-way ANOVA, followed by Tukey’s post hoc test. 11?-HSD2 protein, mRNA, and promoter activity data were analyzed by a standard t test. Significance was set at P < 0.05. Calculations were performed using SPSS software version 9.0 (SPSS, Inc., Chicago, IL).
Results
Expression of PPAR mRNA in cultured human placental trophoblasts
To verify the expression of PPAR in our cultured human placental trophoblast cells, we determined the presence of PPAR mRNA by a standard RT-PCR. As shown in Fig. 1, PPAR mRNA was detected in total RNA samples from both placental tissues and cultured syncytiotrophoblast cells. Consistent with the predominant syncytiotrophoblast localization, higher levels of PPAR mRNA were detected in cultured trophoblast cells than in placental tissues.
FIG. 1. Expression of PPAR mRNA in cultured placental trophoblast cells. Total RNA was extracted from placental villi at term and from cultured trophoblast cells. One microgram of RNA was used in a standard RT-PCR to amplify mRNA for human PPAR and GAPDH (used as an internal control), as described in Materials and Methods. A fraction of the RT-PCR products was subjected to electrophoresis on a 1.2% agarose gel. This figure shows the results of one representative experiment.
Effects of cPGI2 on 11?-HSD2 activity
To study the effects of cPGI2, a putative PPAR agonist, on placental 11?-HSD2 activity, isolated human trophoblast cells were treated with 10 μM cPGI2 for various times (6, 12, and 24 h). This treatment resulted in a time-dependent decrease in 11?-HSD2 activity, such that a significant reduction (P < 0.05) occurred after 6 h of treatment (Fig. 2A). Furthermore, the cells were also treated with various concentrations of cPGI2 (1–20 μM), and this led to a concentration-dependent decrease in 11?-HSD2 activity, with a maximal effect at 10 μM cPGI2 (Fig. 2B).
FIG. 2. Time- and concentration-dependent effects of cPGI2 on 11?-HSD2 activity. A, Time-dependent effects of cPGI2 on 11?-HSD2 activity. Trophoblast cells were incubated with or without 10 μM cPGI2 for various periods of time (6, 12, and 24 h). B, Concentration-dependent effects of cPGI2 on 11?-HSD2 activity. Trophoblast cells were treated for 24 h with increasing concentrations of cPGI2 (1–20 μM). At the end of treatment, the level of 11?-HSD2 activity in intact cells was determined by a standard radiometric conversion assay, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments, each performed in triplicate. *, P < 0.05 vs. control.
Mechanisms of cPGI2-induced decrease in 11?-HSD2 activity
To determine the signal transduction pathway by which cPGI2 decreases placental 11?-HSD2 activity, we treated the cells with H89, an inhibitor of protein kinase A (PKA). We reasoned that if the cPGI2-induced decrease in 11?-HSD2 activity was due to activation of the classical prostacyclin receptor (IP) treatment with H89 would abolish the effect of cPGI2. Although H89 alone caused a 40% reduction in the level of 11?-HSD2 activity, it failed to abrogate the inhibitory effect of cPGI2 (Fig. 3A). This suggested that the effect of cPGI2 on 11?-HSD2 was probably mediated through activation of PPAR.
FIG. 3. Effects of cPGI2 on 11?-HSD2 activity: involvement of PPAR. A, Effects of a PKA inhibitor. Trophoblast cells were treated for 24 h with cPGI2 (10 μM), H89 (10 μM), or the combination of cPGI2 and H89. B, Effects of indomethacin and a selective PPAR agonist. Trophoblast cells were treated for 24 h with indomethacin (10 μM), cPGI2 (10 μM), or GW610742 (10 μM). C, Effects of 9-cis-RA. Trophoblast cells were treated for 24 h with RA (100 nM), a submaximal concentration of cPGI2 (5 μM), or the combination of RA and cPGI2. At the end of treatment, the level of 11?-HSD2 activity in intact cells was determined by a radiometric conversion assay, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments, each performed in triplicate. *, P < 0.05 vs. control.
To provide additional support for the involvement of PPAR in down-regulating placental 11?-HSD2 activity, we used a selective PPAR agonist, GW610742. Treatment of the trophoblast cells with GW610742 resulted in a similar decrease (to that observed with cPGI2) in 11?-HSD2 activity. In addition, indomethacin, a known ligand for both PPAR and PPAR (36), had no effect (Fig. 3B).
Having established the involvement of PPAR in mediating the cPGI2-induced down-regulation of placental 11?-HSD2 activity, we conducted the next experiment to elucidate the molecular mechanisms by which cPGI2 reduces 11?-HSD2 activity. Given that PPARs, like many other members of the nuclear receptor superfamily, exert their effects by forming obligate heterodimers with the RXR, we sought to determine whether cPGI2 and the RXR ligand 9-cis-retinoid acid (9-cis-RA) would exert synergistic effects on 11?-HSD2 activity. As shown in Fig. 3C, 9-cis-RA alone did not alter 11?-HSD2 activity. Moreover, combined treatment with 9-cis-RA and a submaximal effective concentration of cPGI2 (5 μM) did not result in an additional decrease in 11?-HSD2 activity (Fig. 3C).
Effects of cPGI2 on 11?-HSD2 protein and mRNA
To determine whether the cPGI2-induced decrease in 11?-HSD2 activity was a result of reduced 11?-HSD2 expression, levels of 11?-HSD2 protein and mRNA were assessed by Western blot analysis and qRT-PCR, respectively. As shown in Fig. 4, treatment of trophoblast cells for 24 h with 10 μM cPGI2 led to a similar reduction in the levels of both 11?-HSD2 protein and mRNA.
FIG. 4. Effects of cPGI2 on 11?-HSD2 expression. Trophoblast cells were treated with 10 μM cPGI2 for 24 h. At the end of treatment, levels of 11?-HSD2 protein (B) and mRNA (C) were determined by Western blot analysis and qRT-PCR, respectively, as described in Materials and Methods. One representative autoradiograph is shown in A. Each bar represents the mean ± SEM of four independent experiments. *, P < 0.05 vs. control.
Effect of CHX on the cPGI2-induced decrease in 11?-HSD2 mRNA
To determine whether the cPGI2-induced decrease in 11?-HSD2 mRNA required de novo protein synthesis, trophoblast cells were treated with cPGI2 in the absence and presence of CHX, a protein synthesis inhibitor. We found that cPGI2 was equally effective in reducing 11?-HSD2 mRNA in the absence and presence of CHX, indicating that de novo protein synthesis was not required (Fig. 5A).
FIG. 5. Effects of protein and mRNA synthesis inhibitors on the cPGI2-induced decrease in 11?-HSD2 mRNA. A, Effects of CHX. Trophoblast cells were pretreated with CHX (10 μM) for 1 h, then treated in the absence or presence of cPGI2 (10 μM) for 24 h. At the end of treatment, total cellular RNA was isolated, and the steady state level of 11?-HSD2 mRNA was assessed by qRT-PCR, as described in Materials and Methods. Each data point is expressed as a percentage of the control value, and each bar represents the mean ± SEM of four independent experiments. *, P < 0.05 vs. control. B, Effects of DRB. Trophoblast cells were pretreated with () or without () 10 μM cPGI2 for 12 h. The cells were then treated with 25 μM DRB in the absence or presence of 10 μM cPGI2 (defined as time zero). At the indicated time points thereafter, total cellular RNA was isolated, and the steady state level of 11?-HSD2 mRNA was assessed by qRT-PCR, as described in Materials and Methods. Each data point is expressed as a percentage of the maximum determined at time zero. Data from one representative experiment are shown.
Effects of cPGI2 on 11?-HSD2 mRNA stability
As a first step in elucidating the molecular mechanisms by which cPGI2 decreases 11?-HSD2 mRNA, we assessed the half-life of 11?-HSD2 mRNA by a standard mRNA decay assay using 25 μM DRB, an inhibitor of mRNA synthesis. As shown in Fig. 5B, cPGI2 did not alter the half-life of 11?-HSD2 mRNA.
Effects of cPGI2 on HSD11B2 gene transcription
To determine whether cPGI2 reduces the rate of HSD11B2 gene transcription, trophoblasts were transiently transfected with a luciferase construct containing a 330-bp 5'-flanking region of the human HSD11B2 gene (Fig. 6A). Treatment with cPGI2 (10 μM) resulted in a significant decrease in HSD11B2 promoter activity (Fig. 6B). Moreover, the cPGI2-induced repression of HSD11B2 promoter activity was prevented in cells cotransfected with a dominant negative PPAR mutant (Fig. 6C), thus corroborating the idea that the effects of cPGI2 on placental 11?-HSD2 are mediated by the activation of PPAR.
Discussion
Placental 11?-HSD2 plays a pivotal role in controlling the precise level of fetal exposure to maternal glucocorticoids (4), and it also modulates local actions of glucocorticoids within the placenta (37, 38). Thus, 11?-HSD2 is an important regulator of placental function and fetal development. Indeed, a proper level of 11?-HSD2 in the human placenta appears to be critical for normal fetal development, because attenuated expression of this enzyme is associated with pregnancies complicated by IUGR (6, 21). Moreover, IUGR is a characteristic feature of 11?-HSD2 deficiency (7). Although a variety of hormones, cytokines, and intracellular signaling molecules as well as hypoxia (8, 9, 10, 11, 12, 13, 19, 20, 22) are known to modulate placental 11?-HSD2, the regulation of this important enzyme is incompletely understood.
Abundant PPAR mRNA and protein are expressed in human trophoblast cells (25, 39), but the function of this nuclear receptor in the human placenta is unknown. The present study was designed to investigate the role of PPAR in regulating 11?-HSD2 in cultured human trophoblast cells. As a first step, we sought to verify the expression of PPAR in our cultured cells. Using RT-PCR, we demonstrated that PPAR mRNA was not only present, but also expressed at a higher level, in purified trophoblast cells compared with whole tissue extracts. This observation is consistent with the previously reported predominant localization of PPAR to the syncytiotrophoblast layer of human placenta (39).
We then examined the effect of cPGI2, a putative PPAR agonist, on 11?-HSD2 activity. We showed that treatment of trophoblast cells with cPGI2 led to a time- and concentration-dependent decrease in the level of 11?-HSD2 activity. Furthermore, the cPGI2-induced decrease in 11?-HSD2 activity was a consequence of reduced 11?-HSD2 expression, because levels of both 11?-HSD2 protein and mRNA were decreased after cPGI2 treatment. Given that the actions of cPGI2 can be mediated by the classical G protein-coupled membrane IP receptor (40) or the ligand-activated nuclear receptor PPAR (41), we conducted a series of experiments to determine which of the two distinct signaling pathways was responsible for mediating the cPGI2-induced decrease in 11?-HSD2 activity.
With respect to the IP receptor-mediated signaling pathway, it is believed that binding of cPGI2 to the G protein-coupled IP receptor stimulates the production of cAMP, which, in turn, binds to the regulatory subunit of PKA, releasing the active catalytic subunit. Activated PKA modulates the function of various nuclear transcription factors that bind to DNA sequences present in the regulatory regions of cAMP-responsive genes (40). We showed previously that forskolin (an activator of PKA) increased 11?-HSD2 activity and mRNA in human placental trophoblast cells (10), implicating PKA as a positive regulator of placental 11?-HSD2. Given the opposite effects of cPGI2 and forskolin, we concluded that the cPGI2-induced reduction in 11?-HSD2 activity was unlikely to be due to the IP receptor activation. To provide experimental evidence for our contention, we treated trophoblast cells with the PKA inhibitor H89. Consistent with our previous report of stimulatory effects of forskolin on placental 11?-HSD2, treatment of trophoblast cells with H89 alone resulted in a reduction of 11?-HSD2 activity. This suggested that the PKA pathway might be involved in maintaining high levels of 11?-HSD2 expression in trophoblast cells. However, H89 was ineffective in blocking the inhibitory effect of cPGI2. Taken together, these findings suggested that the cPGI2-induced down-regulation of placental 11?-HSD2 was probably mediated by PPAR.
To provide additional evidence for the involvement of PPAR, we examined the effects of GW610742, a selective PPAR agonist, on 11?-HSD2 activity. Treatment of trophoblast cells with GW610742 resulted in a similar decrease in the level of 11?-HSD2 activity. In contrast, indomethacin, a promiscuous ligand for both PPAR and PPAR (36), had no effect. Taken together, these results indicated that the cPGI2-induced decrease in 11?-HSD2 activity was a result of activation of PPAR.
PPARs are ligand-activated transcription factors that exert their effects by forming heterodimers with RXR, binding to PPREs in the regulatory regions of target genes, and inducing ligand-dependent transactivation. Gene targeting studies in mice have demonstrated a critical role for RXR in placentation and fetal development (42, 43). Moreover, PPAR and RXR have been shown to colocalize in the syncytiotrophoblastic layer of the term human placenta (39). Therefore, we determined whether these two cognate receptors would interact to regulate 11?-HSD2 in cultured trophoblast cells. We treated trophoblast cells with the pan-RXR ligand 9-cis-RA in the presence and absence of the putative PPAR agonist cPGI2. Our results showed that 9-cis-RA alone had no effect on 11?-HSD2 activity. Moreover, 9-cis-RA did not augment the inhibitory effect of a submaximal concentration of cPGI2 on 11?-HSD2 activity, suggesting that the cPGI2-induced decrease in 11?-HSD2 activity was unlikely to be due to interactions between PPAR and RXR through a PPRE. Consistent with this idea, there are no apparent PPRE consensus motifs in the 330-bp 5'-flanking region of human HSD11B2 gene. Recently, PPAR and PPAR have been reported to repress gene transcription by restricting the binding of specific transcription factors to respective response elements (44) or by competing with critical coactivators necessary for transcriptional activation (45). In addition, there is evidence that PPAR exerts transcriptional repression via interactions with corepressor proteins (46, 47). Pertinent to our findings, both PPAR and PPAR have been shown to interact with specificity protein-1 (Sp1), causing transcriptional suppression of vascular endothelial growth factor receptor-2 gene (48) and thromboxane receptor gene (49), respectively. Given that the human HSD11B2 gene contains two Sp1-binding sites within the 330-bp 5'-flanking region (50, 51), it is tempting to speculate that PPAR may repress HSD11B2 gene transcription via an interaction with Sp1 in human trophoblast cells. Additional studies are currently being conducted to examine this possibility.
The observed decrease in the level of 11?-HSD2 mRNA in cultured trophoblast cells after treatment with cPGI2 may be attributed to a direct effect of cPGI2 on 11?-HSD2 or an indirect effect involving the synthesis of an intermediary protein. To determine whether the cPGI2-induced reduction of 11?-HSD2 mRNA required de novo protein synthesis, we treated trophoblast cells with cPGI2 in the absence and presence of CHX, a protein synthesis inhibitor. Although CHX alone decreased the basal level of 11?-HSD2 mRNA (a nonspecific effect probably caused by a decrease in the levels of various proteins that are required to sustain the general cell transcriptional machinery), it did not block the cPGI2-induced decrease in 11?-HSD2 mRNA, suggesting that de novo protein synthesis was not required.
In theory, a decrease in the steady state mRNA level of a given gene can be achieved by reducing the rate of gene transcription and/or by decreasing the mRNA stability. To determine which of these mechanisms was responsible for the observed decrease in 11?-HSD2 mRNA, we treated trophoblast cells with the mRNA synthesis inhibitor DRB in the absence and presence of cPGI2 and determined the rate of 11?-HSD2 mRNA decay by qRT-PCR. Under the conditions of the present study, cPGI2 did not affect the half-life of 11?-HSD2 mRNA, suggesting that the cPGI2-induced decrease in 11?-HSD2 mRNA was probably mediated at the level of HSD11B2 gene transcription. To provide direct evidence for this contention, we examine the effect of cPGI2 on HSD11B2 promoter activity. Our results showed that cPGI2 reduced HSD11B2 promoter activity, and that this effect was abolished in trophoblast cells expressing a dominant negative PPAR mutant. Taken together, these data suggest that activation of PPAR suppresses HSD11B2 gene transcription. Thus, our present findings provide additional evidence for PPAR-mediated transcriptional suppression.
The demonstration of PPAR-induced suppression of placental HSD11B2 gene expression will probably have far-reaching implications for our understanding of the role of this enzyme in normal and pathological pregnancies. We believe that the precise level of 11?-HSD2 in the human placenta is tightly controlled by a balance between stimulatory factors, such as retinoic acids (11), glucocorticoids (22), and activators of the PKA pathway (8), and inhibitory factors, such as prostaglandins (13), hypoxia (19, 20), and PPAR. Thus, it is conceivable that aberrant activities of PPAR, at the level of receptor itself, its ligands, and/or its coregulatory proteins (i.e. coactivators and corepressors), in the human placenta may lead to an altered expression of 11?-HSD2 and, consequently, abnormal fetal development. Indeed, PPAR-null mice exhibited placental defects and IUGR phenotype (32). Given the previous findings that the attenuated placental 11?-HSD2 expression in human pregnancies complicated with IUGR was not due to imprinting or mutations in the HSD11B2 gene (21), it is attempting to speculate that overactivation of PPAR may be a contributing factor in these pathological pregnancies.
In conclusion, the present study demonstrates that cPGI2 decreases 11?-HSD2 activity and expression in cultured human placental trophoblast cells, and that this effect is probably mediated by the nuclear receptor PPAR, which functions to repress HSD11B2 gene transcription. If PPAR reduces HSD11B2 gene expression in the human placenta in vivo, our findings would provide a molecular mechanism by which this nuclear receptor may regulate human placental function and, consequently, fetal development.
Acknowledgments
We thank Dr. Daniel Hardy for technical assistance, and Dr. Joseph Torchia for insightful suggestions.
References
Cole TJ, Blendy JA, Monaghan AP, Krieglstein K, Schmid W, Aguzzi A, Fantuzzi G, Hummler E, Unsicker K, Schutz G 1995 Targeted disruption of the glucocorticoid receptor gene blocks adrenergic chromaffin cell development and severely retards lung maturation. Genes Dev 9:1608–1621
Seckl JR 1994 Glucocorticoids and small babies. Q J Med 87:259–262
Seckl JR 2001 Glucocorticoid programming of the fetus; adult phenotypes and molecular mechanisms. Mol Cell Endocrinol 185:61–71
Yang K 1997 Placental 11?-hydroxysteroid dehydrogenase: barrier to maternal glucocorticoids. Rev Reprod 2:129–132
Stewart PM, Rogerson FM, Mason JI 1995 Type 2 11?-hydroxysteroid dehydrogenase messenger ribonucleic acid and activity in human placenta and fetal membranes: its relationship to birth weight and putative role in fetal adrenal steroidogenesis. J Clin Endocrinol Metab 80:885–890
Shams M, Kilby MD, Somerset DA, Howie AJ, Gupta A, Wood PJ, Afnan M, Stewart PM 1998 11?-Hydroxysteroid dehydrogenase type 2 in human pregnancy and reduced expression in intrauterine growth restriction. Hum Reprod 13:799–804
Kitanaka S, Tanae A, Hibi I 1996 Apparent mineralocorticoid excess due to 11?-hydroxysteroid dehydrogenase deficiency: a possible cause of intrauterine growth retardation. Clin Endocrinol (Oxf) 44:353–359
Pasquarette MM, Stewart PM, Ricketts ML, Imaishi K, Mason JI 1996 Regulation of 11?-hydroxysteroid dehydrogenase type 2 activity and mRNA in human choriocarcinoma cells. J Mol Endocrinol 16:269–275
Sun K, Yang K, Challis JR 1997 Differential regulation of 11?-hydroxysteroid dehydrogenase type 1 and 2 by nitric oxide in cultured human placental trophoblast and chorionic cell preparation. Endocrinology 138:4912–4920
Sun K, Yang K, Challis JR 1998 Regulation of 11?-hydroxysteroid dehydrogenase type 2 by progesterone, estrogen, and the cyclic adenosine 5'-monophosphate pathway in cultured human placental and chorionic trophoblasts. Biol Reprod 58:1379–1384
Tremblay J, Hardy DB, Pereira LE, Yang K 1999 Retinoic acid stimulates the expression of 11?-hydroxysteroid dehydrogenase type 2 in human choriocarcinoma JEG-3 cells. Biol Reprod 60:541–545
Sarkar S, Tsai SW, Nguyen TT, Plevyak M, Padbury JF, Rubin LP 2001 Inhibition of placental 11?-hydroxysteroid dehydrogenase type 2 by catecholamines via -adrenergic signaling. Am J Physiol 281:R1966–R1974
Hardy DB, Pereria LE, Yang K 1999 Prostaglandins and leukotriene B4 are potent inhibitors of 11?-hydroxysteroid dehydrogenase type 2 activity in human choriocarcinoma JEG-3 cells. Biol Reprod 61:40–45
Hardy DB, Dixon SJ, Narayanan N, Yang K 2001 Calcium inhibits human placental 11?-hydroxysteroid dehydrogenase type 2 activity. Biochem Biophys Res Commun 283:756–761
Pepe GJ, Waddell BJ, Stahl SJ, Albrecht ED 1988 The regulation of transplacental cortisol-cortisone metabolism by estrogen in pregnant baboons. Endocrinology 122:78–83
Baggia S, Albrecht ED, Pepe GJ 1990 Regulation of 11?-hydroxysteroid dehydrogenase activity in the baboon placenta by estrogen. Endocrinology 126:2742–2748
Pepe GJ, Davies WA, Dong KW, Luo H, Albrecht ED 1999 Cloning of the 11?-hydroxysteroid dehydrogenase (11?-HSD)-2 gene in the baboon: effects of estradiol on promoter activity of 11?-HSD-1 and -2 in placental JEG-3 cells. Biochim Biophys Acta 1444:101–110
Pepe GJ, Burch MG, Albrecht ED 2001 Estrogen regulates 11?-hydroxysteroid dehydrogenase-1 and -2 localization in placental syncytiotrophoblast in the second half of primate pregnancy. Endocrinology 142:4496–4503
Alfaidy N, Gupta S, DeMarco C, Caniggia I, Challis JR 2002 Oxygen regulation of placental 11?-hydroxysteroid dehydrogenase 2: physiological and pathological implications. J Clin Endocrinol Metab 87:4797–4805
Hardy DB, Yang K 2002 The expression of 11?-hydroxysteroid dehydrogenase type 2 is induced during trophoblast differentiation: effects of hypoxia. J Clin Endocrinol Metab 87:3696–3701
McTernan CL, Draper N, Nicholson H, Chalder SM, Driver P, Hewison M, Kilby MD, Stewart PM 2001 Reduced placental 11?-hydroxysteroid dehydrogenase type 2 mRNA levels in human pregnancies complicated by intrauterine growth restriction: an analysis of possible mechanisms. J Clin Endocrinol Metab 86:4979–4983
van Beek JP, Guan H, Julan L, Yang K 2004 Glucocorticoids stimulate the expression of 11?-hydroxysteroid dehydrogenase type 2 in cultured human placental trophoblast cells. J Clin Endocrinol Metab 89:5614–5621
Desvergne B, Wahli W 1999 Peroxisome proliferator-activated receptors: nuclear control of metabolism. Endocr Rev 20:649–688
Lee CH, Olson P, Evans RM 2003 Minireview: lipid metabolism, metabolic diseases, and peroxisome proliferator-activated receptors. Endocrinology 144:2201–2207
Mukherjee R, Jow L, Croston GE, Paterniti Jr JR 1997 Identification, characterization, and tissue distribution of human peroxisome proliferator-activated receptor (PPAR) isoforms PPAR2 versus PPAR1 and activation with retinoid X receptor agonists and antagonists. J Biol Chem 272:8071–8076
Lim H, Gupta RA, Ma WG, Paria BC, Moller DE, Morrow JD, DuBois RN, Trzaskos JM, Dey SK 1999 Cyclo-oxygenase-2-derived prostacyclin mediates embryo implantation in the mouse via PPAR. Genes Dev 13:1561–1574
Matsuura H, Adachi H, Smart RC, Xu X, Arata J, Jetten AM 1999 Correlation between expression of peroxisome proliferator-activated receptor ? and squamous differentiation in epidermal and tracheobronchial epithelial cells. Mol Cell Endocrinol 147:85–92
Gupta RA, Tan J, Krause WF, Geraci MW, Willson TM, Dey SK, DuBois RN 2000 Prostacyclin-mediated activation of peroxisome proliferator-activated receptor in colorectal cancer. Proc Natl Acad Sci USA 97:13275–13280
Hansen JB, Zhang H, Rasmussen TH, Petersen RK, Flindt EN, Kristiansen K 2001 Peroxisome proliferator-activated receptor (PPAR)-mediated regulation of preadipocyte proliferation and gene expression is dependent on cAMP signaling. J Biol Chem 276:3175–3182
Wang YX, Lee CH, Tiep S, Yu RT, Ham J, Kang H, Evans RM 2003 Peroxisome-proliferator-activated receptor activates fat metabolism to prevent obesity. Cell 113:159–170
Peters JM, Lee SS, Li W, Ward JM, Gavrilova O, Everett C, Reitman ML, Hudson LD, Gonzalez FJ 2000 Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor ?(). Mol Cell Biol 20:5119–5128
Barak Y, Liao D, He W, Ong ES, Nelson MC, Olefsky JM, Boland R, Evans RM 2002 Effects of peroxisome proliferator-activated receptor on placentation, adiposity, and colorectal cancer. Proc Natl Acad Sci USA 99:303–308
Kliman HJ, Nestler JE, Sermasi E, Sanger JM, Strauss III JF 1986 Purification, characterization, and in vitro differentiation of cytotrophoblasts from human term placentae. Endocrinology 118:1567–1582
Krozowski Z, MaGuire JA, Stein-Oakley AN, Dowling J, Smith RE, Andrews RK 1995 Immunohistochemical localization of the 11?-hydroxysteroid dehydrogenase type II enzyme in human kidney and placenta. J Clin Endocrinol Metab 80:2203–2209
Bastie C, Luquet S, Holst D, Jehl-Pietri C, Grimaldi PA 2000 Alterations of peroxisome proliferator-activated receptor activity affect fatty acid-controlled adipose differentiation. J Biol Chem 275:38768–38773
Lehmann JM, Lenhard JM, Oliver BB, Ringold GM, Kliewer SA 1997 Peroxisome proliferator-activated receptors and are activated by indomethacin and other non-steroidal anti-inflammatory drugs. J Biol Chem 272:3406–3410
Sun K, Yang K, Challis JR 1998 Glucocorticoid actions and metabolism in pregnancy: implications for placental function and fetal cardiovascular activity. Placenta 19:353–360
Burton PJ, Waddell BJ 1999 Dual function of 11?-hydroxysteroid dehydrogenase in placenta: modulating placental glucocorticoid passage and local steroid action. Biol Reprod 60:234–240
Wang Q, Fujii H, Knipp GT 2002 Expression of PPAR and RXR isoforms in the developing rat and human term placentas. Placenta 23:661–671
Namba T, Oida H, Sugimoto Y, Kakizuka A, Negishi M, Ichikawa A, Narumiya S 1994 cDNA cloning of a mouse prostacyclin receptor. Multiple signaling pathways and expression in thymic medulla. J Biol Chem 269:9986–9992
Forman BM, Chen J, Evans RM 1997 Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors and . Proc Natl Acad Sci USA 94:4312–4317
Sapin V, Dolle P, Hindelang C, Kastner P, Chambon P 1997 Defects of the chorioallantoic placenta in mouse RXR null fetuses. Dev Biol 191:29–41
Wendling O, Chambon P, Mark M 1999 Retinoid X receptors are essential for early mouse development and placentogenesis. Proc Natl Acad Sci USA 96:547–551
Delerive P, De Bosscher K, Besnard S, Vanden Berghe W, Peters JM, Gonzalez FJ, Fruchart JC, Tedgui A, Haegeman G, Staels B 1999 Peroxisome proliferator-activated receptor negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-B and AP-1. J Biol Chem 274:32048–32054
Li M, Pascual G, Glass CK 2000 Peroxisome proliferator-activated receptor -dependent repression of the inducible nitric oxide synthase gene. Mol Cell Biol 20:4699–4707
Shi Y, Hon M, Evans RM 2002 The peroxisome proliferator-activated receptor , an integrator of transcriptional repression and nuclear receptor signaling. Proc Natl Acad Sci USA 99:2613–2618
Lee CH, Chawla A, Urbiztondo N, Liao D, Boisvert WA, Evans RM, Curtiss LK 2003 Transcriptional repression of atherogenic inflammation: modulation by PPAR. Science 302:453–457
Meissner M, Stein M, Urbich C, Reisinger K, Suske G, Staels B, Kaufmann R, Gille J 2004 PPAR activators inhibit vascular endothelial growth factor receptor-2 expression by repressing Sp1-dependent DNA binding and transactivation. Circ Res 94:324–332
Sugawara A, Uruno A, Kudo M, Ikeda Y, Sato K, Taniyama Y, Ito S, Takeuchi K 2002 Transcription suppression of thromboxane receptor gene by peroxisome proliferator-activated receptor- via an interaction with Sp1 in vascular smooth muscle cells. J Biol Chem 277:9676–9683
Agarwal AK, White PC 1996 Analysis of the promoter of the NAD+ dependent 11?-hydroxysteroid dehydrogenase (HSD11K) gene in JEG-3 human choriocarcinoma cells. Mol Cell Endocrinol 121:93–99
Nawrocki AR, Goldring CE, Kostadinova RM, Frey FJ, Frey BM 2002 In vivo footprinting of the human 11?-hydroxysteroid dehydrogenase type 2 promoter: evidence for cell-specific regulation by Sp1 and Sp3. J Biol Chem 277:14647–14656(Laura Julan, Haiyan Guan,)