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REPRODUCTION-DEVELOPMENT |
Division of Hormone Research, Department of Cell Biology, Pharmacology, and Neuroscience (M.G., Z.-X.Y., N.B., M.C., V.P.), Georgetown University Medical Center, Washington, DC 20007; and the Chemical Industry Institute of Toxicology Centers for Health Research (J.C.C.), Research Triangle Park, North Carolina 27709
Address all correspondence and requests for reprints to: Dr. V. Papadopoulos, Division of Hormone Research, Department of Cell Biology, Georgetown University Medical Center, 3900 Reservoir Road, Washington, DC 20007. E-mail: . papadopv{at}georgetown.edu
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and PPARß/
, but not for PPAR
. In vivo treatment of mice with PPs resulted in the reduction of both testis PBR mRNA and circulating testosterone levels, in agreement with the proposed role of PBR in steroidogenesis. By contrast, liver PBR mRNA levels were increased, in agreement with the proposed role of PBR in cell growth/tumor formation in nonsteroidogenic tissues. However, PPs did not inhibit testosterone production and testis PBR expression in PPAR-null mice. These results suggest that the antiandrogenic effect of PPs is mediated by a PPAR-dependent inhibition of Leydig cell PBR gene expression. |
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Most PPs elicit similar responses in rodent livers, including changes in peroxisome proliferation, cell proliferation, and hepatocarcinogenesis. Concurrent with the increase in number and size of peroxisomes, there is transcriptional induction of a group of peroxisomal enzymes responsible for ß-oxidation of fatty acids, the cytochrome P450 CYP4A enzymes found in the endoplasmic reticulum, and the cytosolic fatty acid-binding and acyl-CoA-binding proteins (2, 3, 4). These morphological and biochemical changes are associated with alterations in lipid metabolism (6, 13, 14). Thus, the transcriptional and morphological changes reflect an adaptive response that maintains the homeostasis of cellular lipids. These adaptive responses have been shown to be mediated by a family of receptors called PP-activated receptors (PPARs) (4), which are members of the nuclear receptor superfamily. PPARs exist as three separate gene products: , ß/, and in various species (3, 4). PPARs function as ligand-activated transcription factors mediating the effects of PPs on the expression of genes implicated in lipid metabolism (3, 4). Differences have been reported in responsiveness to PPs among isoforms, levels of expression during development, and amounts in various tissues (3, 4). Interestingly, Leydig cells were found to express the PPAR and PPARß/ proteins (15, 16).
In addition to their effect on liver, PPs also induce pathological changes in the reproductive system. They exert subtle effects on fertility and litter size (17, 18, 19) and not-so-subtle effects on the male reproductive system, such as testicular atrophy manifested in young animals by damage to the Sertoli cells and detachment of germ cells from the seminiferous tubule epithelium (20, 21). Because of the effect of phthalates on development, Mayer and Sanders (22) suggested that they might also have an effect on steroid synthesis. In the mouse, treatment with di-2-ethylhexyl phthalate (DEHP) led to a decrease in testicular testosterone (23), as did the proximate metabolite of DEHP, mono(2-ethylhexyl)phthalate (MEHP) (24). In later studies, Jones et al. (25) demonstrated that pretreatment with MEHP and mono-n-octyl phthalate, for 2 h, inhibited LH-stimulated testosterone secretion in primary cultures of rat Leydig cells. The findings that treatment with phthalates led to decreased testosterone levels in LH- stimulated isolated Leydig cells (25) and that coadministration of testosterone with DEHP prevented DEHP-induced testicular toxicity (26) suggest that Leydig cells may be the primary target of phthalates in the testis.
Treatment of rats with the PP ciprofibrate resulted in increased levels of peroxisomal ß-oxidation and catalase genes in Leydig cells (27). No changes in peroxisomal volume or activities were shown in the seminiferous tubule compartment. In addition, clofibrate was found to alter cell growth and gene expression and inhibit progesterone synthesis in human choriocarcinoma JEG-3 cells (28).
In the studies described below, we investigated the direct effects of PPs on Leydig cell function (steroid synthesis) and examined the mechanism underlying these effects. Our results demonstrate that PPs exhibit antiandrogenic properties through inhibition of the hormone-induced rate-determining step in steroid biosynthesis, the transport of cholesterol into mitochondria. This effect is mediated by the PP-activated and PPAR-mediated transcriptional suppression of the peripheral-type benzodiazepine receptor (PBR) gene encoding a high-affinity mitochondrial cholesterol-binding protein (29, 30) involved in the regulation of cholesterol transport across the mitochondrial membranes (31).
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-32P]deoxy-CTP (specific activity, 3000 Ci/mmol), [32P]nicotinamide adenine dinucleotide (specific activity, 800 Ci/mmol), [1,2,6,7-N-3H]testosterone (specific activity, 93.9 Ci/mmol), [7-N-3H]pregnenolone (specific activity, 22.6 Ci/mmol), and [1,2,6,7-N-3H]progesterone (specific activity, 94.1 Ci/mmol) were obtained from NEN Life Science Products (Wilmington, DE). Translabeled L-methionine, [35S]: Lcysteine, [35S] (35S-met/cys; specific activity>1000 Ci/mmol) was obtained from ICN Pharmaceuticals, Inc. (Costa Mesa, CA). PK 11195 and aminoglutethimide were obtained from Research Biochemicals International (Natick, MA). Nitrocellulose (0.45 µm) was from Hoefer Scientific (San Francisco, CA). Trilostane (4,5-epoxy-17-hydroxy-3-keto-5-androstan-2-carbonitrile) was donated by Stegram Pharmaceuticals (Sussex, UK). 22R-Hydroxycholesterol, dibutyryl-cAMP, testosterone, progesterone, dimethylsulfoxide (DMSO), bezafibrate and DEP, DEHP, and methycellulose were purchased from Sigma-Aldrich Corp. (St. Louis, MO). WY-14,643 was from BIOMOL Research Laboratories, Inc. (Plymouth Meeting, PA) or ChemSyn Science Labs (Lenexa, KS). MEHP was a gift of Dr. B. Davis (National Institute of Environmental Health Sciences, Research Triangle Park, NC). Cell culture supplies were purchased from Life Technologies, Inc. (Grand Island, NY), and cell culture plasticware was from Corning, Inc. (Corning, NY). Electrophoresis reagents and materials were supplied from Bio-Rad Laboratories, Inc. (Richmond, CA). Oligonucleotides were synthesized and purified by Bio-Synthesis, Inc. (Lewsville, TX). All other chemicals used were of analytical grade and were obtained from various commercial sources.
Animal studies
All experimental protocols were reviewed and approved by the Animal Care and Use Committee of the Chemical Industry Institute of Toxicology (CIIT) Centers for Health Research. Twelve-week-old male PPAR(-/-)-null and wild-type SV129 mice, generated and characterized as previously described (32), were obtained from the CIIT breeding colony. Mice were treated by daily gavage for 7 consecutive days with 1 g/kg·d DEHP or 50 mg/kg·d WY-14,643 in 1% methycellulose. The animals were killed the morning after the last day of the treatment. Sera for testosterone measurements and livers and testes for determination of PBR mRNA levels were collected and stored frozen at -70 C.
Cell culture and treatments
MA-10 mouse Leydig tumor cells were a gift from Dr. Mario Ascoli (University of Iowa). Cells were plated at low density for 24 h in modified Waymouth’s MB 752/1 medium containing 20 mM HEPES, 1.2 g/l NaHCO3, and 15% horse serum (pH 7.4), as previously described (33). After 24 h, the media were changed, and fresh media containing the indicated amounts of various compounds or their solvents were added for 24, 48, or 72 h. At the end of the incubation, the cells were washed with serum-free medium and stimulated for 2 h with saturating amounts of hCG (1 nM) in serum-free medium. This concentration of hCG was previously shown (33) to maximally stimulate steroid synthesis by these cells. In some experiments, the cAMP analog dibutyryl-cAMP (1 mM) and the cholesterol derivative 22R-hydroxycholesterol (10 µM) were used instead of hCG. At the end of the 2-h incubation period, the cell media were saved for progesterone determination, and the cells were either dissolved in 0.1 N NaOH for protein determination or processed for protein (Western) or mRNA (Northern) blot analysis or other biochemical assays.
R2C cells from ATCC (Manassas, VA), derived from rat Leydig tumors, were grown in modified Waymouth’s MB752/1 medium containing 20 mM HEPES, 1.2 g/liter NaHCO3, and 10% horse serum, pH 7.4, as previously described (34). Cells were treated with the indicated PPs, following the protocol described above for the MA-10 cells. Because these cells produce steroids in a constitutive, hormone-independent manner, the cells were incubated for 2 h, in the presence of media alone, at the end of the treatment.
Testicular interstitial cells were prepared by collagenase dissociation (33) of testes obtained from adult Sprague Dawley rats (300 g). Leydig cells were further purified using discontinuous Percoll gradient centrifugation as previously described. The preparations obtained contained 75–85% Leydig cells, as shown by the histochemical staining for 3ß-hydroxysteroid dehydrogenase (33). Isolated rat Leydig cells were cultured for 48 h with various concentrations of the indicated PPs, under the same conditions as described above for the MA-10 cells. At the end of the incubation, the cells were washed with serum-free medium and stimulated for 2 h with saturating amounts of hCG (1 nM) in serum-free medium. Media were collected for testosterone measurements.
Measurement of Leydig cell catalase activity
MA-10 and R2C cells, cultured in 150-mm culture dishes, were treated as described above, for 48 h, with or without 100 µM of either bezafibrate or MEHP. At the end of the incubation, cells were collected, and catalase activity was measured by spectrophotometry as described by Aebi (35).
Measurement of cholesterol transport in mitochondria
For these studies, MA-10 cells, cultured in 150-mm culture dishes, were treated as described above, for 48 h, with or without the indicated concentrations of various PPs. At the end of the preincubation, cells were washed with serum-free medium and treated with 1 nM hCG in the presence of 0.76 mM aminoglutethimide, a specific inhibitor of the P450scc (P450 side-chain cleavage). After 20 min incubation time, media were removed, the cells were resuspended in PBS, and mitochondria were isolated in buffers containing aminoglutethimide as described previously (36). The rate of pregnenolone formation, reflecting the amount of cholesterol available to the P450scc, was determined, as previously described, in aminoglutethimide-free buffer (36) in the presence of 1 µM trilostane, an inhibitor of 3ß-hydroxysteroid dehydrogenase enzyme (37). Pregnenolone formation was measured as an index of P450scc activity.
Radioimmunoassays
Progesterone, testosterone, and pregnenolone were measured by means of RIA (33, 36) using antibodies specific for each steroid (ICN Pharmaceuticals, Inc.). The data were analyzed using the MultiCalc software from EGG-Wallace, Inc. (Gaithersburg, MD).
Cell viability and mitochondrial integrity
Cell viability was analyzed using the trypan blue exclusion method. At the end of the incubation, the cells were washed three times with PBS and incubated for 15 min with 0.1% trypan blue stain. After washing, stained cells (dead cells) were counted by light microscopy. Cell viability and mitochondrial integrity were also evaluated, following the above-described cell treatment protocol, by measuring the levels of cellular 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) reduction (38) using the TACS MTT proliferation assay (Trevigen, Inc., Gaithersburg, MD).
Total protein labeling
A total of 106 MA-10 cells were treated with and without 300 µM bezafibrate or its solvent (DMSO) for 48 h. Cells were then translabeled with 35S-met/cys (0.6 mCi/well) in met/cys free medium for 4 h. Total protein labeling was examined, as we previously described (39).
Measurement of DNA damage
The possibility that PFDA may cause DNA damage to the cells was examined using the poly(ADP-ribose) polymerase assay (40 ; Trevigen, Inc.). Cells were treated with and without 300 µM bezafibrate or its solvent (DMSO) for 48 h. Cells were collected and resuspended in 50 mM Tris [pH 8.0] buffer supplemented with 25 mM MgCl2 and 0.1 mM phenylmethylsulfonylfluoride. Cells were sonicated and centrifuged, and the supernatants were used to determine the poly(ADP-ribose) polymerase activity, as we previously described (39).
Radioligand binding assays
Radioligand binding assays were performed as previously described (33). In brief, cells treated with the indicated amounts of PPs were scrapped from culture flasks into PBS, dispersed by trituration, and centrifuged at 1200 x g for 5 min. The cells were resuspended in PBS to a final concentration of 5–10 µg protein/100 µl. Saturation binding studies were performed on 10 µg protein for the MA-10 cells in a final vol of 300 µl in the presence of the radioligand at the indicated concentrations. Nonspecific binding was determined in the presence of 6 µM of the homologous nonradioactive ligand. After incubation at 4 C for 2 h, the assays were stopped by filtration through GF/B filters (Brandel, Gaithersburg, MD), equilibrated in 0.1% polyethyleneimine, and washed with 40 ml ice-cold PBS. Radioactivity trapped on the filters was determined by liquid scintillation counting. The dissociation constant and binding capacity were defined after Scatchard analysis of the specific binding using the LIGAND program (41). In some experiments, the direct effect of PFDA on PK 11195 ligand binding to isolated MA-10 Leydig cell mitochondria was examined.
SDS-PAGE and immunoblot analysis
Mitochondria were isolated as we previously described (33, 39). Mitochondrial proteins were fractionated by one-dimensional SDS-PAGE and electrotransferred onto nitrocellulose. The nitrocellulose was subjected to immunoblot analysis using anti-PBR (1:1000) antiserum and goat antirabbit IgG alkaline phosphatase-conjugated secondary antibody followed by the substrate nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl-phosphate for detection (39). Densitometric analysis of the immunoreactive protein bands was performed using the Sigmagel software (Jandel Scientific, San Rafael, CA).
RNA (Northern) blot analysis
Total cellular and tissue RNA was isolated by the acid guanidinium thiocyanate-phenol-chloroform extraction method (42) using the RNASTAT reagent (Tel-Test Inc., Friendswood, TX). RNA electrophoresis, transfer, probe labeling, and membrane hybridization against the [32P]cDNA probe for PBR were performed as previously described (39). Screen-enhanced autoradiography was performed by exposing X-OMAT AR films (Eastman Kodak Co., Rochester, NY) to the blots at -80 C for 48 h. Densitometric analysis of the immunoreactive protein bands was performed using the Sigmagel software (Jandel Scientific).
Measurement of PBR mRNA stability
To examine the effects of bezafibrate on PBR mRNA stability, we treated MA-10 cells for 48 h with 100 µM of bezafibrate and then measured the decay of PBR mRNA by incubating the cells with actinomycin D (10 µg/ml) for an additional 15, 30, 60, and 180 min (39). Total cellular RNA was isolated as described above, and PBR mRNA levels were measured by RNA (Northern) blot analysis.
RT-PCR
Total RNA was isolated from MA-10 Leydig cells using a commercial kit (RNASTAT 60). RT-PCR was used to detect PPAR expression in MA-10 Leydig cells. The first cDNA strand was synthesized from 10 µg total RNA using specific primers for PPAR, PPARß/, and PPAR. The following primers were used: for PPAR, (5'-GGTCAAGGCCCGGGTCATACTCGCAGG-3' and 5'-TCAGTACATGTCTCTGTAGATCTCT-3'); for PPARß/, (5'-GTCATGGAACAGCCACAGGAGGAGA-CCCCT-3' and 5'-GGGAGGAATTCTGGGAGAGGTCTGCACAGC-3'); and for PPAR, (5'-GAGATGCCATTCTGGCCCACCAACTTCGG-3' and 5'-TATCATAAATAAGCTTCAATCGGATGGTTC-3'). RT-PCR reactions were performed using the Access RT-PCR system (Promega Corp., Madison, WI) according to the manufacturer’s instructions. RT reactions were performed at 48 C for 1 h, and they were terminated by heating at 94 C for 2 min. Amplification was carried out by 40 cycles at 94 C for 1 min, 55 C for 2 min, and 72 C for 3 min, followed by an extension step at 72 C for 7 min. Total mouse liver RNA was used as an internal control (Ambion, Inc., Austin, TX). The resulting products were purified on 2% low-melting-point agarose gel and sequenced. The ABI PRISM dyes terminator cycle sequencing ready reaction kit (PE Applied Biosystems, Foster City, CA) and a PE Applied Biosystems sequencer were used for sequencing at the Lombardi Cancer Center Sequencing Core Facility (Georgetown University). DNA sequences were analyzed by using the Entrez and BLAST program against GenBank Database.
Transient transfections
MA-10 cells were allowed to grow to 70–80% confluence for the transfection experiments. The MA-10 cells were transiently transfected using the electroporation method. Briefly, each Genepulser cuvette (0.4-cm gap; Bio-Rad Laboratories, Inc.) contained 8 x 106 cells in 350 µl antibiotic-free complete Waymouth’s growth medium, plus 50 µl of 0.1x Tris-EDTA containing the plasmid preparations. The cells were cotransfected with 30 µg of the pPBR (-1219/+36) CAT (chloramphenicol acetyltransferase) construct, and 5 µg per transfection of the pSV promoter-ß-galactosidase plasmid (Stratagene, La Jolla, CA) to check transfection efficiency. The 1219-bp promoter region fragment, which contained the first 36 bp of exon 1 of the rat PBR gene, subcloned into the polylinker of the pCAT-basic (Promega Corp.) was provided by Dr. Krueger (Ref. 43 ; Georgetown University). As control, MA-10 cells were also transiently cotransfected with 30 µg of the pCAT-basic plasmid (Promega Corp.) and 5 µg per transfection of the pSV promoter-ß-galactosidase plasmid (Stratagene). Cells in electroporation cuvettes were electroshocked using 330 V and a capacitance of 950 µFd generated from Genepulser II (Bio-Rad Laboratories, Inc.). The cells were kept at room temperature for 10 min before plating onto six-well dishes and further allowed to grow in complete Waymouth’s medium for 16 h. The media were changed into serum-free Waymouth’s, and cells were treated with or without 100 µM bezafibrate, MEHP, DEP, or DMSO. The cells were harvested, 24 h later, for CAT-ELISA (5-Prime 
3-Prime, Inc., Boulder, CO) and ß-galactosidase (Promega Corp.) activity measurements.
Protein measurement
Protein was measured by the method of Bradford (44), using BSA as a standard.
Statistics
Statistical analysis was performed by one-way ANOVA and unpaired Student’s t test using the INSTAT 3.00 package from GraphPad Software, Inc. (San Diego, CA).
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). This effect was statistically significant at doses of 10-5 M and above (ANOVA, P < 0.001). Bezafibrate also significantly inhibited progesterone biosynthesis after 24 or 72 h of pretreatment (Table 1). DMSO, at the dilutions used, had no effect on hCG-stimulated progesterone production.
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The results presented above on the MA-10 mouse tumor cell line were confirmed using isolated adult rat Leydig cells (Fig. 1B
). The 48-h pretreatment with bezafibrate resulted in the inhibition of the subsequent hCG-stimulated testosterone production, with an IC50 close to 10-4 M, similar to that seen in MA-10 cells. The effect of bezafibrate on MA-10 and isolated rat Leydig cells is not unique to this PP. It was also seen with other members of this diverse family of compounds, uch as MEHP, the proximate metabolite of the phthalate ester plasticizer DEHP, and the experimental hypolipidemic drug WY-14,643 (Fig. 1, C and D). The inactive diethylphthalate (DEP) was ineffective. Moreover, the inhibitory effect of PPs on Leydig cell steroidogenesis was not restricted to hormone-dependent cell models, given that it was also found in the R2C rat Leydig tumor cell line that constitutively produces high levels of steroids. Bezafibrate and MEHP inhibited the formation of progesterone in R2C cells, with IC50 values of 30 and 100 µM, respectively (Fig. 1, E and F).
Exposure of bezafibrate up to 10-3 M for 48 h did not affect MA-10 cell proliferation or viability, as shown by the fact that the MTT values were constant, showing neither increase (proliferation) nor decrease (cell death) over the control value upon treatment (Table 2 and data not shown). These results also suggest that bezafibrate up to 10-3 M does not affect mitochondrial integrity, because the MTT assay is based on MTT reduction by the mitochondrial diaphorase enzyme, an index of mitochondrial integrity (38). Although pretreatment with 300 µM bezafibrate for 48 h led to inhibition of the hCG-stimulated steroid synthesis by 60% (Fig. 1), it did not affect total protein synthesis, which can be used as an additional index of toxicity (Table 2). Moreover, 300 µM bezafibrate did not induce DNA damage in the cells (Table 2), as determined by measuring the activity of the poly(ADP- ribose) polymerase, an enzyme involved in DNA repair.
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), an index of peroxisome proliferation.
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, A and B). We then examined whether bezafibrate affected the first step in steroid biosynthesis, the transformation of cholesterol into pregnenolone by the P450 scc enzyme located within the inner mitochondria membrane. MA-10 cells were pretreated for 48 h with 100 µM bezafibrate, MEHP, or DEP and then incubated in the presence of 22R-hydroxycholesterol (10 µM), the soluble form of cholesterol that readily enters the inner mitochondrial membrane. Progesterone production was identical in both control and bezafibrate-pretreated cells (Fig. 2A), suggesting that the site of action of bezafibrate is not at the P450scc or other steroidogenic enzyme level, but rather at an earlier step of the pathway. However, MEHP inhibited cholesterol-supported steroid formation (Fig. 2B), suggesting that MEHP acts on many steps of the steroidogenic machinery. Thus, the inhibitory effect of bezafibrate, as well as one of the MEHP sites of action on steroidogenesis, seems to be attributable to an impairment of the multistep process of cholesterol mobilization, transport into mitochondria, and loading onto P450scc in response to hCG stimulation.
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) (P < 0.001 by Student’s t test). These data indicate that bezafibrate and MEHP inhibited the transfer or loading of cholesterol to the inner mitochondrial membrane P450scc.
PPs inhibit PBR ligand binding capacity, protein, and mRNA expression
Bezafibrate treatment of the cells induced a significant (P < 0.001 by Student’s t test) decrease in the number of PBR binding sites (Bmax, maximal binding capacity) in MA-10 cells measured using the high-affinity PBR ligand PK 11195 in saturation binding studies analyzed by Scatchard analysis (Fig. 3A). The effect of bezafibrate on PBR Bmax was time (Table 4)- and dose-dependent, with an IC50 of approximately 100 µM (data not shown). This effect was also observed after 100 µM MEHP treatment, when the number of PK 11195 binding sites decreased by 50% (data not shown). No effect on receptor affinity was observed (data not shown). As with progesterone synthesis, when a 48-h pretreatment with 100 µM bezafibrate was followed by washing and further incubation of the cells in bezafibrate-free medium for 48 h, PBR levels returned to control levels (data not shown). Bezafibrate, when tested at concentrations up to 100 µM, did not displace radiolabeled PK 11195 in direct ligand binding studies performed using isolated MA-10 mitochondria. However, MEHP used at a concentration of 100 µM displaced 20% of the radiolabeled PBR drug ligand PK 11195 (data not shown).
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). Densitometric analysis suggested a 60% decrease of the 18-kDa PBR protein expression in bezafibrate-treated cells (Fig. 3B). In addition, bezafibrate (Fig. 3C) and MEHP (Fig. 3D), which were both tested at 100-µM concentrations, reduced PBR mRNA levels by 60 and 50%, respectively, whereas the solvent DMSO and the inactive phthalate DEP had no significant effect.
PPs do not accelerate PBR mRNA decay but suppress PBR gene transcription
The results presented above suggest that bezafibrate and MEHP exert a direct effect on PBR mRNA stability or transcription. We tested whether bezafibrate affects PBR mRNA stability, by exposing the cells to 100 µM bezafibrate for 48 h and then measuring the decay of PBR mRNA in cells incubated with actinomycin D for various time periods up to 180 min. The results suggested that bezafibrate did not affect either PBR or the housekeeping gene cyclophilin mRNA decay (data not shown). Thus, we extended these studies by examining the effects of bezafibrate treatment on PBR gene transcription. The -1219/+36 DNA region upstream from the transcription start site of the PBR gene (43) has been shown previously to contain the regulatory elements necessary and sufficient for expression of PBR in MA-10 mouse Leydig tumor cells. The possible role of promoter elements present within this region in the down-regulation of PBR gene transcription by bezafibrate was examined using transient transfection experiments. MA-10 mouse Leydig tumor cells transfected with the PBR (-1219/+36)-chloramphenicol acetyltransferase (CAT) construct were treated with 100 µM bezafibrate or the solvent DMSO. Bezafibrate treatment caused a reduction of approximately 50% in the expression of CAT activity (Fig. 4A). Treatment of the cells with 100 µM MEHP, but not DEP or DEHP, also resulted in a approximately 50% reduction of CAT activity (Fig. 4B). These results suggest that the inhibitory effects of bezafibrate and MEHP on PBR expression are attributable to suppression of PBR gene transcription.
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-null) mice and PPARß/ mRNAs in MA-10 cells (Fig. 5A). The primer pairs amplified internal fragments of 1.6 kb and 130 bp size for the PPAR and PPARß/ genes, respectively. Nucleotide sequencing of the amplified fragments indicated that the amplified PPAR and PPARß/ MA-10 gene fragments exhibit 94 and 100% nucleotide sequence homology to the previously described rat PPAR (45) and rat PPARß/ (46, 47) gene products, respectively. PPAR mRNA was not detectable in MA-10 cells.
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mRNA, we investigated the role of this receptor in mediating the effects of PPs on Leydig cell PBR gene transcription and steroid synthesis. Wild-type PPAR(+/+) and PPAR(-/-) null mice were treated with the MEHP precursor DEHP or WY-14,643. WY-14,643 was chosen because it is well characterized in terms of its ability to bind and activate PPAR isoforms (4) and to induce responses in the liver and kidney through a PPAR-dependent mechanism and does induce effects in the testis, including alterations in the levels of an estrogen metabolism enzyme (estrogen dehydrogenase) and induction of Leydig cell tumors in certain rat strains. Moreover, WY-14,643 inhibited both MA-10 and rat Leydig cell steroid formation (Fig. 1, C and D). Treatment of wild-type mice with PPs resulted in reduction of circulating testosterone levels (Fig. 5B) (P < 0.05 for DEHP and WY-14,643 by Student’s t test), validating the in vitro effects of PPs observed in MA-10, R2C, and normal rat Leydig cells. Compared with the wild-type controls, circulating testosterone levels were decreased in PPAR(-/-) null control mice (P < 0.01 by Student’s t test), suggesting a positive constitutive role for PPAR in maintaining Leydig cell steroid formation (Fig. 5B). Surprisingly, treatment with PPs restored circulating testosterone to normal levels, indicating that these compounds may also act either through PPARß/ or through another unknown mechanism. Testis PBR mRNA levels in wild-type mice were decreased after treatment with either DEHP or WY-14,643, roughly in parallel with the testosterone levels (P < 0.05 by Student’s t test for both treatments). In contrast, liver PBR mRNA levels were increased in wild-type mice after treatment with WY-14,643 (P < 0.001 by Student’s t test) (Fig. 5C), indicating a tissue-specific regulation of PBR expression by PPs. The inhibitory effects of PPs on testis PBR mRNA levels were reversed in PPAR(-/-) null mice, suggesting that the inhibitory effect of PPs on PBR gene expression is attributable to PPAR (Fig. 5C). In addition, the stimulatory effect of WY-14,643 on liver PBR mRNA was absent in liver tissue from PPAR(-/-) null mice. |
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and PPARß/ in transactivation assays, bezafibrate, which acts preferentially on PPARß/ in Xenopus, is more efficient on the rodent PPAR isoform similarly to MEHP and WY-14,643 (4). The data presented here clearly demonstrate that PPs inhibit, in a dose- and time-dependent manner, hCG-stimulated progesterone production in MA-10 mouse Leydig tumor cells. Moreover, PPs inhibited, with the same potency, the hCG-stimulated testosterone production in isolated rat Leydig cells, indicating that the results seen with the MA-10 cells were not specific to a Leydig tumor cell line. In addition, PPs also inhibited steroid formation by the R2C Leydig cells, a constitutively steroid-producing tumor cell line (34). These results suggest that normal and tumoral Leydig cells from mice and rats respond in a similar manner to PPs. Moreover, the finding that PPs inhibit the hormone-induced and constitutively sustained steroidogenesis with similar IC50 values indicates that these compounds act on a common regulatory component of the steroidogenic pathway.
Steroidogenesis is regulated by trophic hormones that trigger three responses: 1) changes in the state of phosphorylation of specific proteins via the cAMP-protein kinase; 2) induction of protein synthesis; and 3) stimulation of lipid synthesis (49, 50). One or all of these hormone-induced changes will trigger the transport of cholesterol from sites of storage or synthesis to the inner mitochondrial membrane, where C27 scc takes place via an enzymatic reaction. Detailed studies have shown that the reaction catalyzed by P450scc is not the rate-limiting step in the synthesis of steroid hormones; rather, it is the transport of the precursor, cholesterol, from intracellular sources to the inner mitochondrial membrane and the subsequent loading of cholesterol in the P450scc active site that is thought to be rate-limiting (49, 50). This hormone-dependent transport mechanism was shown to be mediated by cAMP and to be localized in the mitochondrion (49, 50). Pregnenolone is formed and leaves the mitochondrion to undergo enzymatic transformation in the endoplasmic reticulum, which gives rise to the final steroid products. Thus, the rate-determining step in the regulation of steroidogenesis and the primary site of acute hormone action is the process of cholesterol delivery to P450scc across the mitochondrial membranes.
PPs inhibited cAMP-stimulated progesterone synthesis by MA-10 cells. However, bezafibrate did not affect the 22R-hydroxycholesterol-supported steroid formation, whereas MEHP inhibited this step of the pathway. 22R-hydroxycholesterol is a hydrosoluble form of cholesterol that enters the inner mitochondria membrane readily and an intermediate of the P450scc reaction. This result suggests that bezafibrate did not act on the P450scc or other steroidogenic enzymes involved in the biosynthesis of the final steroid product. In contrast, our data indicate that MEHP exhibits activity at multiple sites in the steroidogenic pathway, including steroidogenic enzymes.
The effect of PPs was reversible, suggesting that they do not exert a permanent toxic effect on Leydig cells at the concentrations and times used. These results were corroborated by a series of assays measuring mitochondrial integrity, total protein synthesis, DNA, and cell viability. These tests clearly showed that bezafibrate did not exert any toxic effect on the cells under the conditions used. Data on the nontoxic effect of MEHP on Leydig cell viability and mitochondrial integrity were recently reported (51). Moreover, a recent detailed morphologic analysis of the MEHP-treated cells failed to demonstrate any morphological alterations of the cells that would indicate toxicity compared with control, with the potential exception of the increase in number of lipid droplets in treated cells (51). Moreover, the MTT assays showed no significant increase in cell numbers at all concentrations used, suggesting that PPs do not trigger Leydig cell proliferation, in contrast with their well-described proliferative effects in liver. In conclusion, PPs exert a specific antisteroidogenic effect on Leydig cells. If this effect of PPs is conserved among species, concern should be raised about the risk in human populations exposed long-term to PPs. Thus, characterization of the molecular determinants involved in the response of Leydig cells to PPs will be useful in developing a predictive model of risk to humans.
PPs have been well characterized for their effect on liver peroxisome proliferation (27). Interestingly, we did not observe any significant effect of PPs on MA-10 and R2C Leydig cell catalase activity examined either by an enzymatic assay or histochemistry (Dees, J. H., and V. Papadopoulos, unpublished results). These results are in agreement with previous studies by Reddy and colleagues (27), who failed to show any change on Leydig cell catalase immunolabeling in rats treated with fibrates and our detailed morphometric electron microscopy studies on MA-10 cells treated with MEHP (51).
As noted above, bezafibrate did not directly affect P450scc or other steroidogenic enzymes, but rather substrate (cholesterol) availability was limiting. Previous studies indicated that MEHP treatment induced a buildup of cytoplasmic lipid droplets that could be attributable to reduced utilization, release of lipids from the cells, or both (51). Because hCG has been shown to reduce the number of lipid droplets in Leydig cells in vivo (52) in a time frame close to the hCG-induced steroid production, cholesterol esters might form at least part of the content of these lipid droplets. Deesterified cholesterol from these lipid droplets is part of the substrate used for testosterone formation by Leydig cells. The fact that the number of the lipid droplets was found to be 3-fold greater in the MEHP-treated cells (51) suggests that most of the cytoplasmic substrate was not transported to the mitochondria and, instead, accumulated in the droplets. Taken together, the data from the bezafibrate and MEHP studies suggest that the site of action of PP is at the level of cholesterol transport into the mitochondria.
Biochemical studies using the inhibitor of P450scc, aminoglutethimide, demonstrated that all PPs tested inhibited the transfer of cholesterol to the P450scc. Because PPs did not affect Leydig cell mitochondrial integrity, we examined their effect on the expression of molecular entities involved in cholesterol transport in steroidogenic cells, such as PBR (31) and the steroidogenesis acute regulatory protein StAR (53). Recent observations revealed that the PP PFDA inhibits rat Leydig cell steroidogenesis by an action on PBR (39). These earlier studies demonstrated that PFDA, which decreased PBR drug ligand binding and protein and mRNA levels, did not affect PBR transcription, but rather accelerated PBR mRNA decay (39). This action of PFDA on PBR mRNA stability indicated a new mechanism of action of PPs, distinct from the classic transcription-mediated regulation of target genes, and this effect may be attributable to the halogen part of this PP.
PBR is a high-affinity, cholesterol-binding protein (29, 30) found in a multimeric complex located at the junctions between outer and inner mitochondrial membranes (contact sites). In this location, the PBR complex could function as a pore, allowing the translocation of cholesterol from the outer to the inner mitochondrial membrane (54, 55). Given the importance of PBR in cholesterol transport, we focused our studies on examining the effect of PPs on PBR expression and function. We demonstrated that a decrease in PBR expression, as determined by decreases in PBR ligand binding and protein and mRNA levels after treatment with the PPs bezafibrate, MEHP, and WY-14,463, closely correlated with reduced hCG-stimulated steroid formation by Leydig cells. A close comparison of the data shown in Tables 1 and 4 indicates that there is a close temporal correlation between the decrease in steroid synthesis and the decrease in PBR levels. Moreover, the fact that bezafibrate inhibited both steroidogenesis and PBR expression levels with the same IC50 suggests a cause-effect relationship. This cause-effect relationship between the effect of bezafibrate on PBR expression and steroid synthesis was further supported by the observation that both PBR levels and steroid synthesis returned to normal on termination of the treatment. Moreover, we did not see any direct effect of bezafibrate on PBR ligand binding, which excludes the possibility that this compound may act as a PBR ligand. However, part of the effect of MEHP could be attributable to a direct effect of the ligand-receptor interaction.
It should be noted that, although this study focuses on the effects of PPs on PBR expression, we cannot dismiss that any other protein involved in intracellular lipid homeostasis and/or cholesterol transport into mitochondria might be a target for PPs. Preliminary studies failed to show an effect of bezafibrate on the 30-kDa mature StAR protein levels in MA-10 cell extracts (unpublished), in agreement with our previous data on the effect of PFDA on StAR levels (39). Although these data suggest that the StAR protein is not a target of PPs in Leydig cells, experiments on the effect of PPs on StAR biosynthesis and processing have to be performed before concluding on the role of StAR in the PP-induced inhibition of the hormone-stimulated steroidogenesis.
The presence of both positive and negative regulatory transcriptional elements has been identified in the sequence upstream of the first exon of the rat PBR gene (43). We examined whether PPs affect PBR mRNA stability or expression through altered regulation of one or more of these transcriptional elements. In vitro transfection studies of the PBR promoter into Leydig cells indicated that PPs affect PBR gene expression directly. Further studies on PBR mRNA stability indicated that bezafibrate and MEHP did not act by accelerating the PBR mRNA decay, further indicating that these compounds act by suppressing PBR gene transcription. Taken together, these data suggest multiple mechanisms of action of PPs in the testis on PBR suppression of expression.
In rodents, PPs are reported to have major effects on the expression of several genes implicated mainly in lipid metabolism (reviewed in Refs. 2, 3, 4). PPs activate specific receptors, called PPARs, which belong to the nuclear hormone receptor gene superfamily (3, 4). So far, three distinct PPARs have been characterized: , ß/, and (3, 4). PPARs heterodimerize with RXR, the 9-cis-retinoic acid receptor, and bind to DNA on sequence-specific response elements called PPREs (PP response elements), located in the promoter regions of target genes, thereby modulating their transcriptional activity (3, 4). While the mechanisms of positive regulation of gene expression by PPs via PPAR-RXR interacting with positive PPRE sequences are fairly well understood (3, 4), the mechanisms of negative gene regulation by PPs and PPAR-RXR still remain to be thoroughly resolved.
Recent publications identified PPAR and PPARß/ immunoreactive proteins in Leydig cells of the testis (15, 16). We confirmed these findings by showing that MA-10 cells contain PPAR and PPARß/ mRNAs but not mRNA for PPAR. In addition, Leydig cells are known to respond to retinoids and express the RXR (56), which potentiates gene regulation by PPAR (4). Why different members of the PPAR family coexist is unknown. But one PPAR could control the expression of the others (57). In summary, these findings show that Leydig cells contain the components necessary to respond to PPs.
The in vitro effects of PPs on Leydig cell steroidogenesis were confirmed in mice in vivo. Treatment of the mice for 7 d with PPs resulted in the reduction of testis PBR mRNA levels and circulating testosterone levels. In contrast, WY-14,643 treatment induced an increase in liver PBR mRNA levels, indicating a tissue specificity of PBR expression in response to PPs. Although striking, this opposite effect of PPs on testis and liver PBR mRNA expression fits well with the proposed roles for this receptor. First, considering the proposed role for PBR in cholesterol transport and steroidogenesis, the decreased testicular PBR levels predict the decrease in testosterone formation by Leydig cells. Second, PBR is also expressed at high levels in various tumors of the breast (58), brain (59), and liver (60). Moreover, the increased expression of PBR in breast cancer cells correlates with increased cell proliferation, the acquisition of an aggressive phenotype (58), and the ability of these cells to grow tumors in severe combined immunodeficient mice (61). In these instances, PBR seems to be localized around or within the nucleus and to mediate the import of cholesterol into the nucleus (58). Because PPs have been well characterized for their hepatocyte-growth-stimulating and hepatocarcinogenic effects (1, 2), an increase in PBR levels may be linked to PP-induced accelerated cell growth.
Recent construction and characterization of a mouse that lacks a functional PPAR gene confirmed the role of PPAR in the regulation of genes involved in hepatic peroxisome proliferation (32). In the present study, we report that PP-induced decreases in testis PBR mRNA levels, and liver PBR mRNA induction by WY-14,643 observed in wild-type mice were not found in PPAR-null mice. In the same tissues (liver and testis), the PPs used induce the expression, in wild-type mice, of two marker genes regulated by PPAR, acyl-CoA-oxidase and Cyp4a. However, both of these gene products are not induced by PPs in PPAR-null mice (Refs. 32, 62 , and 63 ; and Valles, E. G., and J. C. Corton, manuscript in preparation). These results suggest that the PP-induced regulation of PBR gene transcription is mediated by PPAR. Interestingly, the circulating testosterone levels were reduced in untreated PPAR-null (compared with wild-type) mice, suggesting that the presence of this nuclear receptor is required for normal steroid formation. These data, however, bring up an interesting phenomenon; on one hand, PPAR is required for normal steroid synthesis (positive effect); and on the other hand, it mediates an inhibitory transcriptional effect in the presence of PPs. One possible explanation is that the absence of PPAR might affect the expression of other gene products that play a role in steroid biosynthesis. Alternatively, the function and/or DNA-binding properties of transcription factors binding to the DNA close to PPRE might be also affected by the absence of PPAR.
Treatment of the PPAR-null mice with PPs restored testosterone formation and PBR mRNA to normal levels, suggesting that PPs may also act through PPAR-independent pathways. In support of this hypothesis, recent studies demonstrated that part of the toxic effect of DEHP on testis was retained in PPAR-null mice (62).
PP-responsive genes contain PPREs consisting of a near-perfect direct repeat (DR) of two copies of the sequence TGA/TCCT separated by 1 bp (DR-1). However, only half of the nucleotides within each repeat sequence are strictly conserved, whereas others vary considerably (3). Hence, additional cis and trans parameters, other than those dictated by the DR and/or PPAR-RXR heterodimer, are presumably involved in binding and transactivation of PP-regulated genes. We looked for specific PPRE elements within the proximal PBR gene promoter and identified a PPRE-like element in the -410/-380 region of the PBR promoter. This element has homology with previously identified PPREs, including the PPRE present in the human transferrin promoter that mediates an inhibitory signal on transferrin gene expression. Studies on the role of this PPRE-like element in the PP-induced inhibition of PBR expression are in progress in our laboratory.
Although the effects of PPs on various species have been well documented, humans are believed to be relatively unresponsive to PP exposure (64, 65, 66). However, the following finding suggested that humans retain responsiveness to PPs. The human liver PPAR is activated in vitro by a range of PPs shown to be active in rodents (67, 68). Decreases in circulating cholesterol and triglyceride levels after fibrate therapy (64) were attributed to increases in apolipoprotein A-II levels mediated by PPAR (69); and clofibrate was found to alter cell growth and gene expression and to inhibit progesterone synthesis in human choriocarcinoma JEG-3 cells (28). However, humans may possess an altered spectrum of responses to PP exposure that do not include the rodent responses related to hepatocarcinogenesis. It should be noted that the studies performed in humans are of limited value because of the short follow-up period. Thus, concern remains about the true risk in human populations exposed long-term to hypolipidemic drugs and other PPs. As noted above, liver tumor promotion by PPs in rodents may be directly related to their ability to induce proteins involved in lipid transport and metabolism (3, 4), such as PBR. Some of these proteins have been shown to be carriers of phenol and phthalate derivatives (70). Phenolic antioxidants and phthalate plasticizers are widely used in the food and packaging industries. Levels of phthalates have been determined, in food samples, to reach 20 mg/kg (71). Humans consume as much as 0.5 mg/kg body weight/day of phthalates (72), which have been found in human urine (73, 74). Considering that the level of contamination of PPs is high in industrial areas (75), together with the chemical characteristics of these compounds that favor their accumulation in lipid environment and tissues rich in lipid, PPs are obviously putative health hazards. The results presented herein on the PPs MEHP and bezafibrate, together with our previously reported findings on PFDA (39), define distinct mechanisms of action by which PPs decrease the levels of PBR mRNA, including inhibition of either transcriptional or posttranscriptional (39) events. The resulting decrease in PBR protein, which controls the amount of cholesterol transported to the P450scc, leads to a reduction of the amount of steroids formed by Leydig cells. These changes could ultimately affect spermatogenesis and fertility.
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1 Current address: Laboratory of Histology and Embryology, School of Medicine, University of Athens, Athens 11527, Greece. 
2 Current address: Endocrinologie Moléculaire de la Reproduction, UPRES-A Centre National de la Recherche Scientifique 6026, Université de Rennes I, 35042 Rennes Cedex, France.
Abbreviations: Bmax, Maximal binding capacity; CAT, chloramphenicol acetyltransferase; DEHP, di-2-ethylhexyl phthalate; DEP, diethylphthalate; DMSO, dimethylsulfoxide; DR, direct repeat; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; hCG, human chorionic gonadotropin; MEHP, mono(2-ethylhexyl)phthalate; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide; PBR, peripheral-type benzodiazepine receptor; PP, peroxisome proliferator; PPAR, PP-activated receptor; PFDA, perfluorodecanoic acid; PPRE, PP-response element; RXR, 9-cis-retinoic acid receptor or retinoid X receptor; scc, side-chain cleavage; StAR, steroidogenesis acute regulatory protein.
Received January 25, 2002.
Accepted for publication March 19, 2002.
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