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Department of Biomedical Sciences, Colorado State University, Fort Collins, Colorado 80523
Address all correspondence and requests for reprints to: T. M. Nett, Department of Biomedical Sciences, Colorado State University, Fort Collins, Colorado 80523. E-mail: terry.nett{at}colostate.edu.
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-hydroxy-progesterone stimulated intracellular Ca2+ mobilization in CHO cells that expressed ovine mPR in Ca2+-free medium (P < 0.05) but not in CHO cells transfected with empty vector. This rise in intracellular Ca2+ is believed to be from the endoplasmic reticulum as intracellular Ca2+ mobilization is absent when mPR transfected cells are first treated with thapsigargin to deplete Ca2+ stores from the endoplasmic reticulum. Isolation, identification, tissue distribution, cellular localization, steroid binding, and a functional response for a unique intracellular mPR in the sheep are presented. |
Introduction |
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The multiple actions of progesterone are believed to be mediated by the binding of progesterone to its specific intracellular or nuclear receptors (nPRs). Progesterone alters transcription of a variety of genes in target tissues involving binding of hormone to its nPR and subsequent modulation of gene expression (6). These nPR-mediated events are relatively slow to occur but produce long-lasting physiological responses often classified as genomic actions of progesterone. However, many effects of progesterone are too rapid to be genomic and are believed to result from binding to membrane progesterone receptors (mPRs). For example, progesterone and its metabolite 17-hydroxy-progesterone induce a rapid, dramatic and long-lasting influx of calcium (Ca2+) in human sperm (7, 8, 9), ultimately resulting in the acrosome reaction (10, 11). In the hypothalamus, progesterone rapidly inhibits pulsatile release of GnRH and consequently decreases release of LH from the pituitary gland (12, 13). Nongenomic effects of acute exposure to progesterone in the uterus include inhibition of transmembrane Ca2+ entry, release of Ca2+ from intracellular stores (14), and membrane hyperpolarization with subsequent activation of K+ channels (15). In spotted sea trout, progestins cause induction of oocyte maturation through a mPR with reduction in cytosolic cAMP and activation of the MAPK pathway within 5 min (16). A recent study with mPRs in human myometrium also demonstrated a reduction of cAMP levels upon ligand activation (17). Furthermore, progesterone inhibits oxytocin binding in the sheep uterus via a nongenomic mechanism (18); whether the response is due to a mPR is not known. Likewise, specific progesterone binding sites exist in ovine CL membrane fractions (19), but the protein responsible has yet to be elucidated. We hypothesized that a mPR was responsible for the progesterone binding in CL membrane fractions. As such, the aims of this study were to isolate and characterize an ovine mPR distinct from the nPR. Our results indicate a mPR primarily expressed in reproductive tissues that is uniquely localized to the endoplasmic reticulum and causes Ca2+ mobilization upon ligand activation.
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sequences (GenBank accession no. AF313616 and AF313620, respectively) as reported by Zhu et al. (20). Sense ovine mPR primer was 5'-TCCCTGCCCCACCCACAGCCATG-3' and the antisense ovine mPR primer was 5'-CAGACACAAACAACTTTACCAGG-3'. Ovine genomic DNA served as template for PCR because the coding region for the human and pig mPR lacks introns. Amplified DNA was analyzed on a 1% agarose gel and visualized via UV light illumination. The PCR product was excised and purified from the agarose using a QIAEX II gel extraction kit (QIAGEN, Valencia, CA) and subsequently ligated into pGEMT-Easy vector (Promega, Madison, WI) per the manufacturer’s instructions. Identity of the cDNA sample was confirmed by sequencing (University of California, Davis, Davis, CA).
RNA preparation and tissue distribution of putative ovine mPR
Numerous tissues were excised from sheep that had been killed with an overdose of sodium pentobarbital and snap frozen in liquid nitrogen until isolation of RNA. Total RNA was isolated using TRI-reagent per the manufacturer’s instructions (Sigma, St. Louis, MO), and each RNA sample was then subjected to the TURBO DNA-free protocol (Ambion, Austin, TX) to ensure absence of DNA contamination. RNA concentration was determined by spectrophotometry, and integrity of RNA verified by 1% agarose gel electrophoresis in the presence of ethidium bromide followed by visualization under UV light. RT-PCR was performed for putative ovine mPR expression via the SuperScript one-step RT-PCR with platinum Taq (Invitrogen, Carlsbad, CA) using 1 µg total cellular RNA per sample. Primers were designed based on homology between pig and sheep mPR sequences and pig ovarian RNA served as a positive control for RT-PCR. Sense primer was 5'-ACCTCCTGCAGGCCAAGTCTG-3' and the antisense primer was 5'-TCCTGGCAAGTGCGGCCCAG-3'. Absence of genomic DNA in RNA preparations was verified by omitting the reverse transcription/platinum Taq mix and substituting Taq DNA polymerase in the reaction for each RNA sample tested.
Isolation of ovine luteal cells
Corpora lutea were collected surgically from superovulated western range ewes on d 10 after ovulation (21), decapsulated, and dissociated into single cells using collagenase (22). Single-cell suspensions were separated into partially purified small and large cell fractions by elutriation (23). Cells were cultured in DMEM, supplemented with 10% fetal bovine serum (FBS) and penicillin-streptomycin (100 IU penicillin and 100 µg/ml streptomycin).
Cellular localization of putative ovine mPR
Fusion proteins consisting of the putative ovine mPR fused to green fluorescent protein (GFP) or hemagglutinin (HA) were used for cellular localization. A C-terminal GFP fusion to the putative ovine mPR was generated by PCR using cDNA containing the full coding sequence of the mPR, with a gene-specific primer that inserted an EcoR1 site upstream of the transcription start site and a gene-specific primer that eliminated the stop codon and substituted a BamH1 site at its 3'-end to create in-frame restriction sites. The PCR product was digested with EcoR1 and BamH1 and ligated into pEGFP-N2 (CLONTECH, Palo Alto, CA) cut with the same enzymes. The result was a fusion protein consisting of the putative ovine mPR and GFP. Identity of the fusion cDNA was confirmed by sequencing (University of California, Davis, Davis, CA). The HA-mPR fusion protein was generated by inserting the coding sequence for the putative ovine mPR into the pKH3 vector. The result was a fusion protein consisting of three HA peptides on the N terminus of the putative ovine mPR. Identity of the fusion cDNA was confirmed by sequencing (University of California, Davis, Davis, CA). To determine whether mPR is present in the cell membrane, Chinese hamster ovary (CHO) cells were plated onto tissue culture dishes containing a no. 0 coverslip (MatTek Cultureware, Ashland, MA) in complete medium [DMEM, supplemented with 10% FBS, nonessential amino acids (8.9 mg/liter L-alanine, 15 mg/liter L-asparagine, 13.3 mg/liter L-aspartic acid, 14.7 mg/liter L-glutamic acid, 7.5 mg/liter glycine, 11.5 mg/liter L-proline, and 10.5 mg/liter L-serine), and penicillin-streptomycin (100 IU penicillin and 100 µg/ml streptomycin)]. The following day cells were transfected with mPR-GFP encoding vector (pEGFP-N2) using the Polyfect transfection procedure (QIAGEN). Luteal cells were treated identically, except transfection was accomplished with Lipofectamine 2000 (Invitrogen). At 24–48 h after transfection, cells were treated with Alexa 594 concanavalin A (Molecular Probes Inc., Eugene, OR) in PBS, which specifically stains the plasma membrane. Cells were fixed with chilled 4% paraformaldehyde and images acquired on a confocal laser scanning microscope using 488- and 543-nm lines of an argon ion laser to excite samples.
For immunocytochemical experiments, CHO cells were plated as previously described and transfected with HA-mPR encoding vector (pKH3) using the Polyfect transfection procedure (QIAGEN). At 24–48 h after transfection, cells were washed with cold PBS, fixed with chilled 4% paraformaldehyde, washed again with cold PBS, and permeabilized with PBS containing 0.3% Triton X-100 for 10 min at room temperature. Cells were again washed with cold PBS and incubated with PBS plus 3% BSA for 1 h at room temperature to minimize nonspecific binding. Cells were incubated with rabbit polyclonal anti-HA antibody (Santa Cruz Biotechnology, Santa Cruz, CA) at 1:250 dilution in PBS plus 3% BSA overnight at 4 C. Cells were washed with cold PBS plus 0.3% Triton X-100 and then incubated with goat antirabbit Alexa 488 antibody (Molecular Probes) at 1:200 dilution in PBS plus 3% BSA for 1 h at room temperature and protected from light. Cells were then washed with cold PBS and images acquired using confocal microscopy similar to mPR-GFP experiments. To determine whether there was nonspecific binding, background fluorescence was determined with HA-mPR-transfected cells subjected to secondary antibody alone.
To determine whether the ovine mPR was localized in the endoplasmic reticulum, CHO cells were cotransfected with mPR-GFP encoding vector (pEGFP-N2) and pDsRed2-ER vector (BD Biosciences, Palo Alto, CA), a red fluorescent protein that is targeted to the endoplasmic reticulum. At 24 h after transfection, cells were fixed with chilled 4% paraformaldehyde and images acquired on a confocal laser-scanning microscope as described previously.
Western analysis
Cos7 (monkey kidney fibroblast) cells grown in complete medium [DMEM, supplemented with 10% FBS and penicillin-streptomycin (100 IU penicillin and 100 µg/ml streptomycin)] were transfected with HA-mPR encoding vector (pKH3) using the Polyfect procedure (QIAGEN). At 24–48 h after transfection, Cos7 cells transfected with HA-mPR and nontransfected cells were washed with cold PBS and cytosolic and membrane fractions prepared as described previously (24) with slight modifications. Briefly, cells were removed from tissue culture plates using a cell scraper and homogenization buffer [100 mM KCl, 5 mM MgCl2, 50 mM Tris-HCl, 1 mM EGTA plus protease inhibitors (pH 7.2)] and collected by centrifugation. Cells were homogenized and cellular debris collected by centrifugation using a tabletop centrifuge at 5000 x g at room temperature for 15 min. Supernatants were collected and spun at 100,000 x g at 4 C for 1 h. Supernatants were removed and kept separate, whereas pellets containing the membrane fractions were resuspended in cold sample buffer [10 mM Tris-HCl, 250 mM sucrose, 1 mM EGTA plus protease inhibitors (pH 7.2)]. Cytosolic proteins were brought to a total volume of 1 ml with distilled water, 100 µl of 0.15% deoxycholate added, and samples vortexed and incubated for 10 min at room temperature. Next, 100 µl of 72% trichloroacetic acid were added, samples vortexed, and incubated on ice for 20 min. Then samples were centrifuged for 15 min at 16,000 x g at room temperature and the pellet washed three times with acetone to remove trichloroacetic acid. The pellet was air dried and then dissolved in cold sample buffer. Protein content was determined for each sample (cytosol and membrane) using the Coomassie Plus protein assay kit (Pierce, Rockford, IL). Equal amounts of protein from each fraction were separated using denaturing PAGE followed by transfer to nitrocellulose membrane for electroblotting. Samples were analyzed for HA-mPR by Western analysis using a monoclonal anti-HA antibody (Roche Diagnostics, Indianapolis, IN) at 1:1000 dilution in 5% nonfat milk made in Tris-buffered saline plus Tween 20. Proteins were detected using chemiluminescence (SuperSignal West Pico chemiluminescent substrate; Pierce).
Receptor binding assays
Crude membrane preparations of CHO cells and CHO cells that expressed ovine mPR were obtained following procedures described (24) with slight modifications. CHO cells were cultured and transfected with an ovine mPR encoding vector (pcDNA3.1+) using the Polyfect procedure (QIAGEN). At 48 h after transfection, mPR transfected and nontransfected CHO cells were washed with cold PBS and cells removed from tissue culture plates using a cell scraper and cold PBS and concentrated by centrifugation. Cells were washed again with cold PBS, cell number determined by hemacytometer count and homogenized in homogenization buffer with a QIAshredder (QIAGEN) per the manufacturer’s instructions. Supernatants were collected and spun at 100,000 x g at 4 C for 1 h. Supernatants were removed and pellets containing the membrane fractions were resuspended in cold sample buffer. An aliquot of the membrane fractions from transfected and nontransfected cells was kept separate and protein concentration determined using the Coomassie Plus protein assay kit (Pierce). Duplicate aliquots of mPR transfected and nontransfected CHO cell membrane fractions were incubated at 4 C for 1 h in a 0.3-ml buffer of 10 mM Tris-HCl, 1.5 mM EDTA, 1 mM dithiothreitol, 10% glycerol (pH 7.6) in the presence of 4 nM 3H-labeled progesterone and digitonin (250 µM). The bound and free tracers were separated by the addition of 0.8 ml ice-cold dextran-coated charcoal [0.3 g defined charcoal and 0.03 g dextran (Sigma) in 100 ml of a buffer of 10 mM Tris-HCl, 1.5 mM EDTA, 1 mM dithiothreitol, and 10% glycerol (pH 7.6)] and incubated on ice for 10 min. After centrifugation at 1100 x g for 15 min at 4 C, 0.9 ml supernatants were carefully removed, mixed with 5 ml of scintillation cocktail, and radioactivity quantified in a Beckman scintillation spectrometer. Nonspecific binding was measured in duplicate in the presence of 4 µM unlabeled progesterone. Additional controls included tubes without cell membrane fractions but with 3H-labeled progesterone and digitonin. 3H-progesterone binding was also measured in the absence or presence of increasing concentrations of several steroids.
For the competition binding measurements, statistical analysis was performed using the Newman-Keuls multiple comparison test in Prism (version 4a, from GraphPad Software, Inc., San Diego, CA), and significance was taken as a value of P < 0.05. When variances were not homogenous, data were transformed by log 10 function.
Measurement of intracellular Ca2+
Intracellular Ca2+ was measured as described by Shlykov and Sanborn (25). Briefly, CHO cells were plated in complete medium onto tissue culture dishes containing a no. 0 coverslip and the following day transfected with a mammalian expression vector encoding mPR (mPR in pcDNA3.1+) or empty vector (pcDNA3.1+). At 24–48 h after transfection, cells were loaded with fura-2-AM (5 µM) (Molecular Probes) at room temperature for 30–35 min in fluorescence buffer [145 mM NaCl, 5 mM KCl, 1 mM Na2HPO4, 0.5 mM MgCl2, 1 mM CaCl2, 10 mM HEPES, 5 mM glucose (pH 7.4)]. After loading, cells were washed twice with fluorescence buffer and used after 35–45 min. For the experiments with extracellular Ca2+ present, Ca2+ concentrations were measured in fluorescence buffer. For the Ca2+-free experiments (extracellular Ca2+ absent), immediately before measuring Ca2+, fluorescence buffer was removed and replaced with Ca2+-free fluorescence buffer supplemented with EGTA (100 µM). Progesterone (0.1–100 nM) or 17-hydroxy-progesterone (100 nM) was added to cells and changes in intracellular free calcium concentration ([Ca2+]i) in individual cells measured at 340 and 380 nm excitation and 510 nm emission wavelengths with an InCyt2 imaging system (Intracellular Imaging Inc., Cincinnati, OH). To verify specificity of the ovine mPR for the progestins tested, cells expressing the ovine mPR were also treated with testosterone (100 nM), estradiol (100 nM), cortisol (100 nM), or RU486 (1–100 nM) and changes in [Ca2+]i measured in individual cells. Also, to investigate whether a nonpermeable analog of progesterone could elicit an increase in intracellular Ca2+, progesterone conjugated to BSA (P4-BSA) (Sigma) was used (1–100 nM based on total conjugate molecular weight). To determine whether Ca2+ release was from the endoplasmic reticulum, cells were incubated with thapsigargin (generous gift from Dr. Sanborn, Colorado State University) (100 nM) in Ca2+-free buffer to deplete Ca2+ stores in the endoplasmic reticulum before treatment with progesterone or 17-hydroxy-progesterone. In each dish, 40–80 individual cells were examined. Where indicated, the responses per dish were averaged and data expressed as the mean 340:380 ratio ± SEM for the average values for n dishes.
Statistical analysis was performed using the Newman-Keuls multiple comparison test in Prism (version 4a; GraphPad Software), and significance was taken as a value of P < 0.05. When variances were not homogenous, data were transformed by log 10 function.
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, respectively (Fig. 1
). Likewise, the putative ovine mPR contains seven-transmembrane domains based on hydrophobicity analysis (DAS-domain prediction and TmPRED). Despite similarity between the putative ovine mPR and human and pig mPR, almost no homology exists between the putative ovine mPR and sheep nuclear PR-A or -B at either nucleotide or amino acid level.
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). Because the coding region for the ovine mPR consists of only one exon, it was critical to ensure no DNA contamination existed in the RNA samples because this would invalidate the tissue distribution results. Ovine mPR expression was detected in CL isolated from sheep at d 10 of their reproductive cycle. Both small and large luteal cells obtained on d 10 of the cycle also expressed mRNA for the ovine mPR (data not shown). Figure 2B displays tissues that tested negative for ovine mPR expression.
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). Expression of mPR-GFP appeared to be in an intracellular tubular network and possibly in the nuclear envelope. Identical localization of mPR-GFP was observed in ovine small (Fig. 3B) and large luteal cells (Fig. 3C), which express endogenous mPR. Likewise, immunocytochemical localization in CHO cells expressing HA-mPR revealed similar subcellular distribution (Fig. 4A) with no HA-mPR detected in the plasma membrane.
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). Western blot analysis confirmed that HA-mPR was present in the membrane fraction of HA-mPR transfected Cos7 cells with an estimated molecular mass of 40 kDa. This 40-kDa band was not detected in the membrane fraction from nontransfected cells or in cytosolic fractions of either transfected or nontransfected cells. To ensure bands were specific for the anti-HA antibody, each protein sample was also subjected to the same Western blot analysis, but the anti-HA antibody was preabsorbed with HA-peptide. Results from these experiments confirmed the specificity of the bands observed (Fig. 4B).
To determine whether mPR-GFP was localized in the endoplasmic reticulum, studies were performed in CHO cells cotransfected with mPR-GFP and pDsRed2-ER, a red fluorescent protein that is targeted to the endoplasmic reticulum. Expression of mPR-GFP yielded nearly complete colocalization with the ER marker as presented in Fig. 5.
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). Only excess unlabeled progesterone and 17-hydroxy-progesterone were able to significantly (P < 0.05) displace binding of radiolabeled progesterone. Estradiol, testosterone, cortisol, and the progesterone antagonist RU486 failed to inhibit binding (Fig. 6).
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-hydroxy-progesterone on intracellular free Ca2+ in cells expressing ovine mPR shows a representative tracing of the results obtained in CHO cells transfected with ovine mPR after addition of progesterone (1 nM) in Ca2+-free medium. Similar results were obtained after treatment with 17-hydroxy-progesterone (100 nM). Both progestins elicited a significant (P < 0.05) increase in [Ca2+]i that occurred approximately 1 min after addition of treatment to CHO cells transfected with ovine mPR, whereas there was no significant increases in [Ca2+]i in cells transfected with empty vector. To verify whether the increase in [Ca2+]i was specific to the progestins, CHO cells transfected with ovine mPR were also treated with testosterone (100 nM), estradiol (100 nM), cortisol (100 nM), RU486 (1–100 nM), or P4-BSA (1–100 nM) in Ca2+-free medium. The increase in [Ca2+]i was not detected after addition of testosterone, estradiol, cortisol, or RU486 (Fig. 7B). Likewise, addition of P4-BSA at 1 or 10 nM did not alter [Ca2+]i, but a slight increase was observed at 100 nM (data not shown). However, even though the P4-BSA was extracted seven times, a small amount of free progesterone was still present in the P4-BSA preparation, which probably accounts for the slight increase with the highest dose of P4-BSA (100 nM) used.
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, pretreatment with thapsigargin caused a depletion of Ca2+ stores in the endoplasmic reticulum and totally abolished the increase in [Ca2+]i induced by progesterone or 17-hydroxy-progesterone. Because the increase in [Ca2+]i occurred at either approximately 30 sec or 2 min after progestin treatment with negligible changes in cells transfected with empty vector, an average of the 340:380 ratio was determined for both mPR-transfected and cells transfected with empty vector from time of treatment to 5 min after treatment. As shown in Fig. 9, the progestin-induced increase in [Ca2+]i was absent in CHO cells transfected with empty vector. Identical results were noted in cells transfected with ovine mPR and treated with vehicle (EtOH, final concentration < 0.02%) (Fig. 9). Only in cells expressing ovine mPR and treated with progesterone (1 nM) or 17-hydroxy-progesterone (100 nM) was a significant increase (P < 0.05) in [Ca2+]i observed.
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) is also expressed mainly in reproductive tissues, particularly in the placenta; testis; ovary; and possibly in the bladder, kidney, and adrenal (20). We, however, did not detect expression of mPR in the kidney, and the adrenal was not examined in the present study. In support of expression in the uterus, the recently reported human mPR homolog was detected in myometrium from pregnant women (17). Another mammalian mPR (rat) homologous to the ovine mPR was recently reported with mRNA expressed in ovary and CL, supporting our tissue expression (28). These researchers also detected expression in adrenal gland, kidney, brain, and lung (28). Currently the differential tissue distribution cannot be reconciled but may be due to interspecies variation or differences in scientific methodology. Despite the presence of seven-transmembrane domains, the ovine mPR does not appear to localize to the plasma membrane based on confocal microscopy experiments with two different mPR fusion proteins. Our initial studies using the mPR-GFP construct distinctly displayed fluorescence around the nucleus, possibly in an intracellular tubular network. These early studies were surprising because we expected localization of the ovine mPR in the plasma membrane due to the seven-transmembrane domains and the plasma membrane localization of the sea trout mPR (16). Experiments were also conducted with other cell lines expressing the mPR-GFP fusion protein to confirm that the localization of mPR-GFP was not due to cell lineage. Each time subcellular localization appeared to be in an intracellular membrane (data not shown). Because the ovine mPR was expressed in the CL, we determined the cellular localization of mPR-GFP in transfected luteal cells. Once again, there was clear perinuclear expression of mPR-GFP. Because GFP is a large protein, it is possible that the tertiary structure of the ovine mPR was altered, which interfered with the final cellular destination in transfected cells. However, immunocytochemical localization of HA-mPR was similar to the mPR-GFP fusion protein. Thus, localization with a plasma membrane marker (concanavalin A) was not evident for either HA-mPR or mPR-GFP. We should note that whereas our data do not suggest plasma membrane localization, this interpretation differs from that of Karteris et al. (17). These researchers concluded that the human mPR homolog is localized to the plasma membrane. Clearly our studies used a different receptor. Thus, there are possible species differences.
It is important to underscore that our data do not eliminate the possibility of plasma membrane expression but certainly suggest prominent localization to an intracellular compartment, likely the endoplasmic reticulum. Interestingly, a similar pattern of intracellular distribution is evident in the paper by Karteris et al. (17). In fact, the authors stated that the mPR localized with nuclear receptor in the cytoplasm. Thus, as in the case of plasma membrane expression, the data presented by Karteris et al. (17) cannot be used to exclude an intracellular site of action such as endoplasmic reticulum membrane. The use of membrane-impermeable progesterone analogs represents one avenue for assessing membrane vs. intracellular sites of action. However, caveats include the presence of free progesterone and a lack of understanding of the stoichiometry with these compounds. Nevertheless, we performed Ca2+ experiments with P4-BSA in CHO cells expressing ovine mPR. No effect was noted at 1 or 10 nM, but a slight increase at 100 nM was observed (data not shown). We are cautious in interpreting the latter because even though the P4-BSA was extracted seven times, free progesterone was still present in the P4-BSA preparation.
Exploiting the HA epitope, Western blots were performed on membrane and cytosolic preparations of cells expressing HA-mPR and nontransfected cells to further elucidate localization of the ovine mPR. A band of approximately 40 kDa, the estimated molecular mass of the ovine mPR, was detected only in the membrane fraction from cells transfected with HA-mPR. It is noteworthy that the membrane preparation used in these experiments contains all cellular membranes, not just the plasma membrane. Additionally, a higher molecular weight band was also detected in the membrane fraction from cells transfected with HA-mPR and appeared to be specific for the anti-HA antibody because detection of both bands was absent when the antibody was preabsorbed with HA peptide (Fig. 4B). It is possible that the higher molecular weight band represents a glycosylated form of the ovine mPR because there are two possible O-linked glycosylation sites at amino acid positions 29 and 34. Furthermore, two bands of similar molecular weight were detected in the membrane fraction from CHO cells transfected with HA-mPR subjected to identical procedures (data not shown).
Confocal microscopy in CHO cells expressing both mPR-GFP and ER-dsRed suggested the notion that ovine mPR is an intracellular transmembrane receptor. Results from these studies displayed nearly complete colocalization of mPR-GFP with the endoplasmic reticulum marker. In support of the ovine mPR cellular localization, Revankar et al. (30) recently reported that GPR-30, a GPCR that binds estradiol is located primarily in the endoplasmic reticulum and elicits Ca2+ mobilization upon estradiol treatment. A functional intracellular receptor requires ligand passage across the plasma membrane. Because progesterone can easily cross the plasma membrane, it seems quite conceivable a functional ovine mPR may be located in the endoplasmic reticulum. In support of this notion is the Ca2+ studies using CHO cells expressing ovine mPR in a Ca2+-free medium in which both progesterone and 17-hydroxy-progesterone caused an increase in intracellular Ca2+ concentrations, suggestive of an intracellular Ca2+ store. Interestingly, when these cells were first treated with thapsigargin, thus depleting the endoplasmic reticulum of Ca2+ stores and then treated with progesterone or 17-hydroxy-progesterone, the rise in intracellular Ca2+ was absent, further suggesting progestin action at the endoplasmic reticulum. Similar results have been noted in luteinized porcine granulosa cells, wherein progesterone caused an increase in intracellular Ca2+ concentrations via Ca2+ mobilization from the endoplasmic reticulum (31). It was also demonstrated that the increase in Ca2+ was due to activation of phospholipase C linked to a pertussis-insensitive G protein, further suggestive of a GPCR, specifically of the Gq family (31). Interestingly, the nuclear PR antagonist, RU-38486 did not inhibit the progesterone-induced increase in [Ca2+]i, suggesting the mPR has a different specificity than the classic nuclear PR (31). Similarly, we did not observe an increase in [Ca2+]i in CHO cells transfected with the ovine mPR upon treatment with RU486 (1–100 nM). Because the ovine mPR is distinct from the nuclear PRs and is expressed in luteal cells, it will be intriguing to determine whether the ovine mPR works through a pathway similar to luteinized porcine granulosa cells to cause Ca2+ mobilization. As such, future research is aimed at deciphering the specific pathway by which the ovine mPR functions.
The present study clearly demonstrates the ovine mPR specifically binds progestins because only progesterone and 17-hydroxy-progesterone significantly displaced binding of 3H-progesterone. Similar to the ovine mPR receptor binding studies, the sea trout mPR also exhibits specific binding for progesterone and 17-hydroxy-progesterone (16). Progesterone, however, appears to be the predominant ligand for the ovine mPR because much higher concentrations of 17-hydroxy-progesterone (1000-fold excess) were needed to compete with 3H-progesterone binding. The other steroids tested did not compete for 3H-progesterone binding, further supporting progesterone as the true ligand for the ovine mPR. It is important to note the membrane fractions used in the present study contain membranes from the plasma membrane as well as the endoplasmic reticulum. Specific progesterone binding sites have been detected in microsomal rich fractions in a variety of mammalian species. Bramley and Menzies (19) reported progesterone binding sites in ovine CL, which are unlike classical PRs in that they are enriched in intracellular membrane fractions and not associated with cytosolic or nuclear fractions. RU486, the nuclear PR antagonist, did not block progesterone binding to intracellular membrane suggestive of a PR unlike the nPR (19). RU486 also did not significantly compete for 3H-progesterone binding in the present study. Similar results have also been reported in microsomal fractions of corpora lutea from cows (32), pigs (33), humans (34), and rats (28). These data provide further support of a PR functionally distinct from the classic nuclear PR and located in an intracellular membrane such as the endoplasmic reticulum.
Because study of the ovine mPR is at the preliminary stages, precise functions of this receptor are not currently known. Most of the research pertaining to mPR has been generated in nonmammalian species, and as such, study of the ovine mPR and other mammalian mPRs should provide for novel insights concerning nongenomic actions of progesterone in mammals. Furthermore, Peluso et al. (35) demonstrated that progesterone receptor membrane component-1 and plasminogen activator inhibitor RNA binding protein interact with each other and play important roles in regulating progesterone’s actions in the ovary. Whether these proteins also interact with the ovine mPR, however, is not currently known. In summary, we have established the groundwork and a firm basis for future studies directed at identifying the unique biological roles of the ovine mPR. The existence of a unique form of progesterone receptor that contains seven-transmembrane domains and resides in the endoplasmic reticulum provides a mechanistically novel method for initiating actions of progesterone. Coupled with the existence of GPR30 binding estradiol and localized in the endoplasmic reticulum, the possibility exists for a new form of steroid receptors that may alter classical steroid dogma regarding the mechanisms by which steroid hormones act.
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Footnotes |
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Disclosure summary: R.L.A., C.M.C., T.A.F., and G.D.N. have nothing to declare. T.M.N. has previously consulted for Boehringer Ingelheim Pharmaceuticals, Inc. and Mylan Laboatories, Inc. and has received lecture fees from the Society of Toxicology. He holds an equity position in Gonex, Inc.
First Published Online June 22, 2006
Abbreviations: [Ca2+]i, Intracellular free calcium concentration; CHO, Chinese hamster ovary; CL, corpus luteum; FBS, fetal bovine serum; GFP, green fluorescent protein; GPCR, G protein-coupled receptor; HA, hemagglutinin; mPR, membrane PR; nPR, nuclear progesterone receptor; P4-BSA, progesterone conjugated to BSA.
Received January 3, 2006.
Accepted for publication June 12, 2006.
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-hydroxyprogesterone. Novel stimulators of calcium influx in human sperm. J Biol Chem 265:1376–1380This article has been cited by other articles:
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C. Stocco, C. Telleria, and G. Gibori The Molecular Control of Corpus Luteum Formation, Function, and Regression Endocr. Rev., February 1, 2007; 28(1): 117 - 149. [Abstract] [Full Text] [PDF]
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P. Thomas, Y. Pang, J. Dong, P. Groenen, J. Kelder, J. de Vlieg, Y. Zhu, and C. Tubbs Steroid and G Protein Binding Characteristics of the Seatrout and Human Progestin Membrane Receptor {alpha} Subtypes and Their Evolutionary Origins Endocrinology, February 1, 2007; 148(2): 705 - 718. [Abstract] [Full Text] [PDF]
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