Mol. Hum. Reprod. Advance Access originally published online on October 11, 2005
Molecular Human Reproduction 2005 11(9):615-621; doi:10.1093/molehr/gah215
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Characterization of 17ß-hydroxysteroid dehydrogenase type 4 in human ovarian surface epithelial cells
1Department of Reproductive Medicine and Surgery, 2Department of Molecular Pharmacology and 3Department of Gynecology, Graduate School of Medical Sciences, Kumamoto University, Kumamoto, Japan
4 To whom correspondence should be addressed at: Department of Reproductive Medicine and Surgery, Graduate School of Medical Sciences, Kumamoto University, Honjo 1-1-1, Kumamoto 860-8556, Japan. E-mail: yumikonagayoshi{at}fc.kuh.kumamoto-u.ac.jp
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Abstract |
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The human ovarian surface epithelium (hOSE) is a single layer of mesothelial-type primitive epithelial cells that are potential estrogen targets. It has been reported that hOSE cells can produce estrogen. However, the mechanisms that regulate estrogen level(s) in hOSE cells are not yet known. To elucidate the enzymes involved in these reactions, we examined gene expression of 17ß-hydroxysteroid dehydrogenases (17ß-HSDs) in primary hOSE (POSE) and OSE2a cells using RT–PCR. We found that POSE cells and cells of the immortalized hOSE line, OSE2a, bidirectionally converted estrone (E1) and 17ß-estradiol (E2). Both cell types expressed mRNA for 17ß-HSD type 1 (17ß-HSD1), suggesting that the enzyme is involved in the E1 to E2 conversion. Interestingly, both cells expressed 17ß-HSD4 mRNA but not 17ß-HSD2 mRNA. We prepared an antibody against the carboxyl terminal of 17ß-HSD4 (anti-17ß-HSD4 antibody), which recognized the 80 and 48 kDa proteins in POSE and OSE2a cells based on immunoblot analysis. Furthermore, immunohistochemical study revealed the presence of 17ß-HSD4 in hOSE cells in the human ovary. These results suggest that 17ß-HSD4 is involved in estrogen inactivation and may protect against an excessive accumulation of E2 in hOSE cells.
Key words: estrogen/ovarian surface epithelial cells/ovary/17ß-HSD4
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Introduction |
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The human ovarian surface epithelium (hOSE) that surrounds each ovary is a sheet of cuboidal mesothelial cells thought to arise from coelomic epithelium with retention of both epithelial and mesenchymal potential (Auersperg et al., 1984
; Okamura and Katabuchi, 2001, 2005). The hOSE cells participate in formation of the ovarian cortex and share embryonic origins with granulosa cells (GCs) (Auersperg et al., 2001). In adult life, hOSE cells play an important part in the cyclic rupture of the Graafian follicle and formation of the corpus luteum. Unlike GCs, which proliferate, differentiate into lutein cells and die as the corpus luteum regresses, hOSE cells continually proliferate and recolonize the ovarian surface in the wake of each ovulation (Murdoch, 1996; Katabuchi and Okamura, 2003). The hOSE cells have been reported to be a possible source of ovarian epithelial neoplasms under the hypothesis that injury and repair of surface epithelial cells after cyclic ovulation can result in cells that accumulate multiple genetic alterations with development of oncogenic phenotypes (Fathalla, 1971; Godwin et al., 1992).
The hOSE has unique features that are absent from the immediately adjacent pelvic mesothelium, suggesting that local factors in the ovarian cortex may play a role in modifying the growth and morphology of this important cell type. Extracellular signals to which hOSE cells are likely to respond include steroid hormones, polypeptide growth factors, extracellular matrix components and cell adhesion molecules. Cultured hOSE cells are reported to produce growth factors, cytokines, cell adhesion molecules and proteolytic enzymes (Auersperg et al., 2001).
Several studies have suggested that locally produced estrogen is involved in development of epithelial endometriosis within the highly estrogenic ovarian environment (Noble et al., 1996; Nisolle and Donnez, 1997; Ohtake et al., 1999). Expression of estrogen receptors alpha (ER
) and beta (ERß) mRNA in hOSE cells has been reported in normal ovaries and in various hOSE cell lines (Brandenberger et al., 1998; Lau et al., 1999), suggesting that estrogen regulates their function and growth.
Recent studies have demonstrated that estrogenic activity in placenta and endometrium is regulated by tissue-specific estrogen bioavailability (Mustonen et al., 1998; Madsen et al., 2004). The last step of estrogen synthesis is catalysed by 17ß-hydroxysteroid dehydrogenases (17ß-HSDs), which convert less active 17-keto-steroids to active 17ß-hydroxy-forms. To date, 12 different isozymes of 17ß-HSDs have been identified (Adamski and Jakob, 2001; Luu-The, 2001; Mindnich et al., 2004), differing in tissue distribution, catalytic preferences, substrate specificity, and subcellular localization and mechanisms of regulation. In this study, we investigated intracellular control of estrogen activity in hOSE cells and characterized isozymes of 17ß-HSDs in hOSE cells both at the mRNA and protein level.
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Materials and methods |
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Cell and tissue collection and preparation
The hOSE cells were prepared from ovaries of 12 women undergoing abdominal total or radical hysterectomy at Kumamoto University Hospital as approved by the University Ethics Committee. Informed consent was obtained before collecting hOSE cells. Patients’ characteristics are summarized in Table I. All ovaries were grossly normal, and no pathologic lesions were observed on histological examination. Preparation of hOSE cells was performed by published methods (Nakamura et al., 1994
). Briefly, after collagenase digestion under aseptic conditions, hOSE cells were scraped with a surgical blade. The cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM)/F-12 medium (1:1 mixture) (Invitrogen, Carlsbad, CA, USA) containing 20% fetal bovine serum (FBS; Thermo Trace, Carlsbad, CA, USA), 100 U/ml penicillin G (Invitrogen) and 100 µg/ml streptomycin (Invitrogen) in a humidified 5% CO2 incubator. After a few days of incubation, viable hOSE cells were detached from the dish with 0.125% trypsin and 0.01% EDTA and collected by centrifugation. Cells from each dish were resuspended in medium and split into four new culture dishes.
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At seeding after dispersal with trypsin-EDTA, the viable cell number was 0.1–2.7 x 106 per ovary. The cells were subsequently passaged up to three times when cell number was too small for the following examination. It takes approximately 4 days to be confluent. The lifespan was approximately several weeks at longest and varied among the samples. However, in some cases, the cells did not reach subconfluent, and these cells were not employed for the following experiments. We employed these primary, passaged, non-immortalized hOSE cells for the following studies as POSE cells. Because the number of POSE cells was small and the lifespan in culture was limited, we had established immortalized hOSE cell lines using simian virus 40 (SV40) large T antigen, as previously reported (Nitta et al., 2001). One cell line, OSE2a, was derived from a reproductive-age patient, and these cells showed luteinizing hormone-dependent growth (Tashiro et al., 2003). They retained an epithelial character as shown by morphological, immunocytochemical and ultrastructural studies that yielded the same findings as those of POSE cells in vivo (Nakamura et al., 1994). OSE2a cells were cultured in DMEM/F-12 medium (1:1 mixture) containing 10% FBS, 100 U/ml penicillin G and 100 µg/ml streptomycin in a humidified 5% CO2 incubator. THP-1 was obtained from the Health Science Research Resources Bank (Number JCRB0112.1; THP-1, Osaka, Japan). The THP-1 cell line, which was established from human acute monocytic leukaemia cells, reportedly expresses 17ß-HSD4 (Jakob et al., 1995). THP-1 cells were cultured in RPMI 1640 medium with 10% FBS, 50 U/ml penicillin G and 100 µg/ml streptomycin at 37°C in a humidified 5% CO2 incubator.
Estrogen production in POSE and OSE2a cells
Subconfluent POSE cells were trypsinized and seeded at 1 x 105 cells per well in four chamber microtiter plates (Nunc, Nunclon, Denmark) in DMEM/F-12 medium with 1% FBS. The POSE cells from a patient were used for a single experiment.
As for OSE2a cells, confluent cells were subcultured in four-well microtiter plates at 1 x 105 cells per well in serum-free DMEM/F-12 medium.
When the cells became confluent, of 50 ng/ml androstenedione, E1 or E2 (Sigma-Aldrich, St. Louis, MO, USA) dissolved in propyleneglycol containing 10% ethanol was added to the medium as substrate, and the cells were incubated for an additional 24 h. The culture medium in the four-well plates was collected and stored at –20°C until measurement of estrogen was performed.
Total RNA isolation and RT–PCR
Cultured cells were washed twice with phosphate-buffered saline (PBS), and total RNA was prepared using an RNeasy Mini kit (Qiagen, Valencia, CA, USA). Total RNA was quantified by measuring absorbance at 260 nm. Single-stranded cDNA was reverse transcribed from 2 µg total RNA with a Superscript II reverse transcriptase (Invitrogen) at a final concentration of 2.5 U in buffer containing 25 mM Tris–HCl, pH 8.3, 37.5 mM KCl, 4 mM MgCl2, 4 mM dithiothreitol (DTT), 0.5 mM dNTPs and 2.5 µM oligo (deoxythymidine) primer (Invitrogen). Incubation was performed at 42°C for 50 min, followed by incubation at 70°C for 15 min Total RNA was digested with ribonuclease H (Life Technologies, St. Paul, MN, USA) at 37°C for 20 min. PCR was performed using 0.5 mL Easy Start (Molecular BioProducts, San Diego, CA, USA) containing Taq DNA polymerase (Invitrogen). PCR primers were designed based on published sequences. PCR was carried out after denaturation of the sample at 95°C for 5 min. Each cycle consisted of denaturation at 95°C for 45 s, annealing for 45 s and extension at 72°C for 45 s. After amplification, the final 7 min extension step was carried out at 97°C. Primer sequences, annealing temperature and PCR cycle number and the expected size of each product are summarized in Table II. PCR amplification was performed in a PC-707 thermocycler (Program Temp Control System PC-707, Astec, Tokyo, Japan). PCR products were separated by electrophoresis on a 1.5% agarose gel and visualized by ethidium bromide staining.
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DNA sequencing
PCR products were purified using QIAquick PCR Purification Kit (Qiagen), according to the manufacturer’s recommended protocol. Purified PCR products were quantified using an RNA/DNA calculator (Amersham Pharmacia Biotech, Piscataway, NJ, USA). Each PCR product (100 ng) was sequenced on both strands using a thermal cycling method with fluorescent dye labelled-dideoxynucleotide terminators and Taq polymerase (Big Dye Terminator Cycle Sequencing Ready Reaction Kit; Applied Biosystems, Foster, CA, USA). The sequences were obtained using an automated DNA sequencer (ABI PRISM 310; Applied Biosystems). A database search was performed for obtained sequences using the Basic Local Alignment Search Tool (BLAST) 2.0 program (http://www.ncbi.nlm.nih.gov). All PCR products were sequenced to confirm that the cDNAs of interest had been amplified.
Protein extraction
Both cell types were cultured for 7 days at 37°C in DMEM/F-12 medium containing 10% FBS. Cells were washed two times in PBS, pH 7.2, lysed for 30 min on ice in a homogenization buffer containing 60 mM Tris–HCl, pH 7.5, 180 mM NaCl, 6 mM EDTA, 18 mM Na4P2O7, 60 mM NaF, 1.2% NP-40, 1 mM DTT, 1 mM phenylmethylsulphonyl fluoride (PMSF), 10 µg/ml leupeptin and 1 µg/ml pepstatin A and clarified by centrifugation at 14 000 x g for 20 min at 4°C. Human endometrial tissue, the positive control for 17ß-HSDs, was obtained from a surgically removed uterus for a gynaecological indication with permission from the patient and homogenized in a homogenization buffer as described above with a Teflon homogenizer and centrifuged at 14 000 x g for 5 min at 4°C to obtain supernatants.
Preparation of anti-17ß-HSD4 antibody
A carboxyl-terminal peptide (C-peptide) corresponding to residues 724–736 (QKLQMILKDYAKL) of 17ß-HSD4 was synthesized, purified by high-performance liquid chromatography and coupled to keyhole limpet hemocyanin (KLH) in collaboration with Transgenic (Kumamoto, Japan). Polyclonal antibody to 17ß-HSD4 was prepared by immunizing rabbits with KLH-coupled C-peptide. Antiserum showed a high titer (up to 1:729 000) against the C-peptide by enzyme-linked immunosorbent assay (ELISA). The immunoglobulin G (IgG) fraction was prepared with a HiTrap Protein A HP affinity column (Amersham Biosciences, Piscataway, NJ, USA). The antibody was further purified by antigen-affinity chromatography. In control experiments, the antibody was preabsorbed with C-peptide (1:50) overnight at 4°C.
Immunoblot analysis
Protein concentrations of cell extracts and supernatant of endometrial tissues were determined by Bradford’s method (Bradford, 1976) with bovine serum albumin (BSA) as standard. The same amounts of proteins were treated with sodium dodecyl sulphate (SDS) sample buffer (Laemmli, 1970) and boiled for 2 min. Samples containing 20 µg protein were subjected to SDS–polyacrylamide gel electrophoresis (SDS–PAGE) in 10% acrylamide and transferred to a polyvinylidene difluoride (PVDF) membrane (Millipore, Billerica, MA, USA) in 25 mM Tris, 193 mM glycine and 20% methanol by semidry method (Trans-blot®, Bio-Rad Laboratories, Hercules, CA, USA). The membrane was incubated overnight with the anti-17ß-HSD4 antibody (1:500) or anti-17ß-HSD4 antibody preabsorbed with C-peptide. After washing, the membrane was incubated with horseradish peroxidase-conjugated goat anti-rabbit IgG antibody (1:10,000). Immunoreactive proteins were detected using an enhanced chemiluminescence detection kit (Amersham Biosciences), according to the manufacturer’s protocol. A suitable antibody to 17ß-HSD1 was not available for immunoblot analysis.
Immunohistochemistry
Paraffin-embedded sections were deparaffinized in xylene and hydrated in a series of graded ethanol solutions, then microwaved in 10 mM citrate, pH 6.0, for 30 min for antigen retrieval (Shi et al., 1997). Sections were incubated in 3% H2O2 in methanol for 30 min to quench endogenous peroxidase activity. Slides were immersed, tissue face down, in 50 µL 2% BSA with antibody against affinity-purified anti-17ß-HSD4 antibody (1:200), anti-17ß-HSD4 antibody preabsorbed with C-peptide or control serum and incubated at room temperature in a humid chamber for 30 min. Following three consecutive washes in PBS for 5 min each, the sections were incubated at room temperature for 30 min with a peroxidase-conjugated goat anti-rabbit IgG antibody (EnVision+ System, DAKO, Carpinteria, CA, USA). Peroxidase activity was visualized by incubation with 3,3'-diaminobenzidine (DAB) reaction reagent (DAKO) for several seconds. The resulting slides were washed with water, dehydrated by a series of graded ethanols and mounted. Photographs were obtained under light microscopy VANOX-S (OLYMPUS, Tokyo, Japan).
Hormone assays
E1 and E2 were measured by Mitsubishi Kagaku Bio-clinical Laboratories using Direct estrone kit (DBC, London, Ontario, Canada) and DPC estradiol kit (Diagnostic Products, LA, CA, USA), respectively. Detectable limits in these assay conditions were 8 pg/ml. The cross reactivities of the estradiol kit to E1 and the estrone kit to E2 were reported by their laboratories to be below 1.03% and 0.7%, respectively. We also checked the cross reactivities by ourselves, 50 ng/ml E1 in the E2 kit and 50 ng/mL E2 in the E1 kit. The concentration of E1 in the culture medium was below detectable levels just after we added 50 ng/ml E2, and E2 was not detected just after 50 ng/ml E1 was added in the culture medium. Hormone assays were performed in duplicate or triplicate.
Statistical analysis
Values were expressed as the mean ± SD. Statistical analysis was performed using Student’s t-test. In the case of multiple comparisons, significances of individual differences were assessed using the Scheffe test followed by a one-way analysis of variance (ANOVA). All of the analyses were completed using Statview 5.0.1 (SAS Institute, Cary, NC, USA). P < 0.05 was considered statistically significant.
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Results |
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Estrogen production in POSE and OSE2a cells
We first examined whether POSE cells could produce E1 and E2 in the presence of corresponding substrate. Among 12 patients, we studied four preparations of POSE cells from the different individuals (n = 4; patients 1–4 in Table I) followed by statistical analysis. Values are represented as mean ± SD. Concentrations of E1 and E2 in culture medium containing 1% FBS were below detectable limits. The concentrations of E1 and E2 were below detectable level (8 pg/ml, respectively) just after (0 h) the addition of 50 ng/ml of E2 and E1, respectively. E2 concentration in culture medium was increased to 3.4 ± 0.6 ng/ml (P < 0.05), when cells were incubated with 50 ng/ml E1 for 24 h. When cells were incubated with 50 ng/ml E2, E1 concentration was increased to 2.3 ± 0.6 ng/ml (P < 0.05). These results indicated minimum variability in estrogen production in response to E1 and E2 despite different clinical conditions of the patients from whom the cells were obtained.
E2 production in OSE2a cells subsequently increased with increasing the incubation time over 24 h (Figure 1A). When cells were incubated with 50 ng/ml E2 as substrate, E1 concentration was also increased up to 12 h. POSE cells also showed time-dependent conversion of both E2 to E1 and E1 to E2, respectively (data not shown). The number of POSE cells was too limited to compare the kinetics with OSE2a cells. These results clearly showed that bidirectional conversion between E1 and E2 occurred in POSE cells and in OSE2a cells and that this conversion was time dependent. Estrogen was not produced by POSE and OSE2a cells when 50 ng/ml androstenedione was added as substrate (data not shown). These results suggested that OSE2a cells mimicked the character of POSE cells and that OSE2a cells were useful for further studies of the physiological functions of hOSE.
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Identification of mRNA for 17ß-HSD subtypes in POSE and OSE2a cells
As conversion between E1 and E2 is conducted by different isozymes in various tissues, we focused on the isozymes responsible for conversion between E1 and E2 in hOSE cells. First, we examined isozyme expression in POSE cells from five patients (n = 5; patients 5–9 in Table I) and OSE2a cells by RT–PCR. When we used a primer pair for 17ß-HSD1, a 143 base pair (bp) band was detected in all five POSE cell preparations, OSE2a cells and placenta were employed as a positive control (Figure 2A). We could not detect PCR product for 17ß-HSD2 in any hOSE cell types, whereas it was clearly detected in endometrium (Figure 2B). Instead, a 749 bp PCR product corresponding to 17ß-HSD4 was amplified in all hOSE cell types at levels comparable with those obtained with endometrium (Figure 2B). Direct sequencing of PCR products from five POSE cell preparations and OSE2a cells confirmed that 143 and 749 bp fragments corresponded with 17ß-HSD1 and 17ß-HSD4, respectively.
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Immunoblot analysis of 17ß-HSD4 in POSE and OSE2a cells
For the next step, we developed an anti-17ß-HSD4 antibody to identify enzyme at the protein level. When immunoblot analysis was carried out with cell extracts from POSE cells from three patients (n = 3; patients 10–12 in Table I) and OSE2a cells, two proteins with apparent molecular masses of 80 and 48 kDa were recognized by an affinity-purified anti-17ß-HSD4 antibody (Figure 3A). None of these proteins were detected when antibody was preabsorbed with C-peptide used as an antigen (Figure 3B). The antibody reacted with proteins with similar size in THP-1 cells and endometrium, both of which were employed as positive controls for 17ß-HSD4 expression (Figure 3C). The 80 kDa protein was more abundant in OSE2a cells than in endometrium. In contrast, the 48 kDa protein was most abundant in endometrium among all samples. The antibody cross-reacted with proteins of apparent lower molecular masses than 80 kDa (Figure 3A and C). It is not clear at present whether these extra bands were because of non–specific interactions with the antibody or degraded forms of 17ß-HSD1. We could not examine the expression of 17ß-HSD1 at the protein level, because a suitable antibody to 17ß-HSD1 was not available.
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Immunohistochemical staining
In control experiments, anti-17ß-HSD4 antibody immunostained endometrium strongly (Figure 4B), whereas no immunostaining was observed using preabsorbed antibody (Figure 4C). In human ovary, hOSE cells were strongly immunostained with anti-17ß-HSD4 antibody (Figure 4E). Immunostaining was not observed when consecutive sections were immunostained with preabsorbed antibody (Figure 4F). Furthermore, the ovarian stroma was slightly immunostained with anti-17ß-HSD4 antibody. OSE2a cells were also immunostained with anti-17ß-HSD4 antibody (data not shown). From these results, we concluded that 17ß-HSD4 is expressed in hOSE cells in the human ovary as well as OSE2a cells.
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Discussion |
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Various reports suggest that hOSE cells are capable of synthesizing E2 (Ivarsson et al., 2001
). In this study, E2 was not produced in POSE and OSE2a cells when androstenedione was added as substrate for aromatase (p450arom), whereas E2 was converted from E1. These results suggest that E2 production in quiescent hOSE cells depends on the presence of extracellular E1.
17ß-HSD1 was the first 17ß-HSD to be purified and cloned (Luu-The et al., 1989). In various tissues, 17ß-HSD1 preferentially catalyses E1 to E2 conversion. Recent reports suggest that increased E2 production in endometrial cells contributes to the pathogenesis and development of endometrial hyperplasia and adenocarcinoma (Utsunomiya et al., 2001). Our findings strongly suggest that the conversion of E1 to E2 which are catalysed by 17ß-HSD1 is a natural feature for hOSE.
There is accumulating evidence that conversion between E2 and E1 is conducted by 17ß-HSD isozyme(s) in intact and tumour cells. It has been suggested that conversion from E1 to E2 is conducted by 17ß-HSD1 and that conversion from E2 to E1 is conducted by 17ß-HSD2 and 17ß-HSD4. It has been reported that 17ß-HSD1 and 17ß-HSD2 mainly regulate E2 tissue levels, modulating estrogenic action in human breast tissue (Söderqvist et al., 1998) and endometrium (Casey et al., 1994). Several reports have suggested that substantial changes in estrogen metabolism occur in malignant breast tissue (Poutanen et al., 1992; Spiers et al., 1998) and in endometrial abnormalities (Utsunomiya et al., 2001). Although 17ß-HSD2 has been known as the major enzyme for E2 to E1 conversion in estrogen target tissues including placenta and endometrium (Mustonen et al., 1998; Madsen et al., 2004), we did not detect mRNA for 17ß-HSD2 in POSE or OSE2a cells under our experimental conditions. Interestingly, 17ß-HSD4 mRNA was clearly detected in both hOSE cells, suggesting that 17ß-HSD4 is the catalyst in hOSE cells.
Immunoblot analysis using affinity-purified anti-17ß-HSD4 antibody detected two major proteins with apparent molecular masses of 80 and 48 kDa in POSE and OSE2a cells, as well as in THP-1 and endometrium. Furthermore, immunohistochemistry with anti-17ß-HSD4 antibody clearly showed that 17ß-HSD4 was present in OSE cells in human ovary sections in vivo, as well as in OSE2a cells. Given the absence of 17ß-HSD2 in hOSE cells, it is highly possible that 17ß-HSD4 plays the major role in conversion of E2 to E1. To our knowledge, this is the first report on the presence of 17ß-HSD4 in hOSE cells. The ovarian stroma was slightly immunostained with anti-17ß-HSD4 antibody. The physiological meanings of this result are not clear at present. The expression of gene encoding several enzymes involved in gonadal steroidogenesis has been reported in the ovarian stroma (Barbieri, 1992; Jabara et al., 2003). Therefore, it is possible that 17ß-HSD4 plays some physiological roles in ovarian stroma.
Initially, 17ß-HSD4 was reported as an enzyme converting E2 to E1 in porcine endometrium (Adamski et al., 1992). Its physiological function may be protection against excessive E2 in estrogen target tissues, such as endometrium. It has been reported that 17ß-HSD4 is widely distributed throughout non-steroidogenic tissues and cells including THP-1 cells (Martel et al., 1992; Jakob et al., 1995). In these tissues, 17ß-HSD4 has been reported to be involved in ß-oxidation of very long chain fatty acids (van Grunsven et al., 1998, 1999; Baes et al., 2000), branched fatty acids (van Veldhoven et al., 1996; Dieuaide-Noubhani et al., 1997) and bile acid synthesis (Dieuaide-Noubhani et al., 1996). Inactive mutations in the 17ß-HSD4 gene lead to a fetal form of Zellweger syndrome (de Launoit and Adamski, 1999). It is not clear at present whether or not 17ß-HSD4 is involved in lipid metabolism in hOSE cells. It has been suggested that 17ß-HSD4 is capable of converting E2 to E1 by in vitro studies (Dinkel et al., 2002). Taken together with this results, it is highly possible that 17ß-HSD4 is involved in conversion of E2 to E1 in intact hOSE cells. One report (Husen et al., 2000) stated that expression of 17ß-HSD4 decreased in endometrial cell lines HEC-1A and RL95-2 when the cells were treated with FBS. It has also been reported that progesterone, peroxisome proliferators and dexamethasone are able to induce 17ß-HSD4 expression in endometrium and THP-1 cells (Jakob et al., 1995; Kaufmann et al., 1995). However, these effects were not observed in OSE2a cells in our preliminary experiments (unpublished data). Regulation of 17ß-HSD4 expression in hOSE cells is an important project for future study.
Human 17ß-HSD4 gene (HSD17B4) encodes an 80 kDa protein containing three catalytically active domains: the N-terminal short chain alcohol dehydrogenase reductase (SDR), the central hydratase domain and the C-terminal sterol carrier protein 2 (SCP2)-like domain (Figure 5) (Leenders et al., 1998). Full-length 17ß-HSD4 is easily cleaved between the SDR and hydratase domains plus SCP2-like domain in peroxisomes (Markus et al., 1995; Corton et al., 1996; Jiang et al., 1996, 1997). Based on the protein structure of human 17ß-HSD4, the immunoreactive 48 kDa protein we observed appears to be the hydratase plus SCP2-like domain cleaved from the 80 kDa protein. Although 17ß-HSD4 is a member of the SCP2 gene family, including SCP2 and SCPx (Ohba et al., 1994), the function of the SCP2-like domain is still controversial. It may be required for import of 17ß-HSD4 into peroxisomes, or it may act as a molecular chaperone in peroxisomes to protect intact enzyme from denaturation (Bun-ya et al., 2000). It has been reported that antibody against the N-terminal portion of porcine 17ß-HSD4 reacts with an 80 and a 32 kDa protein in porcine myometrium (Kaufmann et al., 1995). Recently, one group (Brown et al., 2004) developed an antibody against equine 17ß-HSD4 and found that the antibody reacted with an 80 and a 45 kDa protein in equine preovulatory follicles. Taken together with this results, the 45 kDa protein in equine follicles must be the cleaved fragment from the full-length 17ß-HSD4. In addition, we found that the proportion of the 80 and the 48 kDa protein differed among the examined cells and endometrium, suggesting that the cleavage is under tissue-specific control. Elucidation of the role of the 48 kDa protein is necessary for future study. Our anti-17ß-HSD4 antibody is suitable for immunoblot and immunohistochemical analyses and useful for these studies.
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In conclusion, hOSE cells can convert E1 and E2 bidirectionally using 17ß-HSD1 and 17ß-HSD4, respectively. These results suggest that the physiology of hOSE cells are controlled under both extracellular and intracellular estrogen activation/inactivation.
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Acknowledgements |
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We are grateful to Dr. Shinji Fukuda and Miss. Naoko Uemura (Kumamoto University, Japan) for their excellent and technical support and Dr. Masahiro Shono (Kumamoto University, Japan) for critical comments.
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Submitted on May 5, 2005; revised on June 26, 2005; accepted on July 19, 2005
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