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Prostaglandin (PG) F₂ Receptor Expression and Signaling in Human Endometrium: Role of PGF₂ in Epithelial Cell Proliferation

Medical Research Council Human Reproductive Sciences Unit, Centre for Reproductive Biology, Edinburgh EH16 4SB, United Kingdom

Address all correspondence and requests for reprints to: Dr. Henry N. Jabbour, Medical Research Council Human Reproductive Sciences Unit, Centre for Reproductive Biology, The Chancellor’s Building, 49 Little France Crescent, Edinburgh EH16 4SB, United Kingdom. E-mail: h.jabbour{at}hrsu.mrc.ac.uk.

	Abstract

Top Abstract Introduction Subjects and Methods Results Discussion References

 
Prostaglandin (PG) F₂, a member of the prostanoid bioactive lipid family, is secreted by human endometrium throughout the menstrual cycle and is present in both menstrual fluid and medium of endometrial explants in culture. PGF₂ mediates its effects through a seven-transmembrane G-protein-coupled receptor (FP). The aim of this study was to examine the temporal expression, signaling, and role of FP receptor in the human endometrium. Quantitative RT-PCR analysis demonstrated highest expression of FP receptor in the mid- to late-proliferative phase, compared with early-proliferative and secretory phase endometrium. In situ hybridization studies localized FP receptor mRNA expression to the epithelial cell compartment during the mid- to late-proliferative phase. Moreover, treatment of endometrial tissue with 1–100 nM PGF₂ induced a concentration-dependent increase in inositol phosphate mobilization, indicating functional FP receptor expression. The Ishikawa human endometrial epithelial cell line was used to investigate further the signaling and role of PGF₂ in endometrial epithelial cells. Ishikawa cells endogenously express the FP receptor, and treatment with 1–100 nM PGF₂ elicits a concentration-dependent increase in inositol phosphate release. Moreover, treatment of Ishikawa cells with 100 nM PGF₂ induced phosphorylation of ERK1/2 that was abolished when cells were cotreated with 50 µM PD98059 (MAPK kinase inhibitor) or 10 µM U73122 [phospholipase C (PLC) inhibitor]. Treatment of Ishikawa cells with PGF₂ for 24 h induced a significant concentration-dependent increase in Ishikawa cell proliferation. Coincubation of the cells with 50 µM PD98059 or 2 µM U73122 demonstrated that PLC inhibition significantly reduced PGF₂-induced proliferation, whereas MAPK kinase inhibition had no effect. In summary, these studies demonstrate increased FP receptor expression in endometrial epithelial cells during the proliferative phase of the menstrual cycle and identify a role for PGF₂ in epithelial cell proliferation via a PLC-dependent pathway.

	Introduction

Top Abstract Introduction Subjects and Methods Results Discussion References

 
PROSTAGLANDIN (PG) F₂ is a prostanoid belonging to the eicosanoid family of biologically active lipids (1). PGs are synthesized from arachidonic acid by a combination of cyclooxygenase (COX) and specific synthase enzymes (2). To date, there are two identified isoforms of the COX enzyme: constitutively expressed COX-1 that generates PGs for normal physiological function; and COX-2, an early response gene whose expression can be rapidly induced (3). PGs mediate their actions via seven-transmembrane G-protein-coupled receptors (GPCRs) specific to each prostanoid. The GPCR for PGF₂ (FP) has been cloned in humans and couples to the G protein Gq, phospholipase C (PLC) activation, and release of inositol phosphates and diacylglycerol (4).

Overexpression of COX-2 and secretion of PGs, such as PGE₂, have been associated with enhanced epithelial cell proliferation and resistance to apoptosis (5, 6). In human endometrium, COX-2 expression is localized to epithelial cells with maximal expression levels during the proliferative phase (7, 8, 9, 10, 11). In addition to PGE₂, PGF₂ is a major metabolite of COX enzymes in human endometrium that can be measured from both human endometrial explants in culture and samples of menstrual fluid (12, 13, 14). Moreover, in vitro studies using separated human endometrial epithelial cells demonstrate higher PGF₂ secretion by epithelial cells extracted from proliferative, rather than secretory, phase tissue (15). In the context of pathological disease, levels of PGF₂ and PGE₂ in menstrual fluid of women suffering from menorrhagia (excessive bleeding during menstruation) are significantly higher than PG levels from women with normal menstrual bleeding (16). Together, these observations implicate PGs, including PGF₂, in both normal and pathological human endometrial function and outline a key regulatory function for PGF_2a in endometrial epithelial cells.

To date, many studies have identified the myometrium as a target for PGF₂. The FP receptor is highly expressed in human myometrium (17), and nanomolar concentrations of PGF₂ induce contractions of myometrial strips in vitro (18). Evidence that the myometrium is a major target for PGF₂ is further supported by observations in FP receptor knockout mice, where loss of myometrial contractility and failed induction of parturition are the major phenotypic changes (19). Within the endometrium, however, little is known about the role of epithelial-cell-derived PGF₂ or its possible sites of action within the functionalis layer.

The aims of this study were to investigate the temporal pattern and site of expression of FP receptor in the human endometrium across the menstrual cycle. Moreover, the Ishikawa cell line was used to investigate further the signaling pathways of PGF₂ and its potential role in proliferation of human endometrial epithelial cells.

	Subjects and Methods

Top Abstract Introduction Subjects and Methods Results Discussion References

 
Subjects and source of endometrial tissue

Endometrial biopsies (n = 12) at different stages of the menstrual cycle were collected, with an endometrial suction curette (Pipelle, Laboratoire CCD, Paris, France), from women with regular menstrual cycles (25–35 d). In addition, full-thickness endometrial biopsies (n = 18) at all stages of the menstrual cycle (n = 3 from early-, mid-, and late-proliferative; and n = 3 from early-, mid-, and late-secretory) were collected from women undergoing hysterectomy for benign gynecological indications. Shortly after pipelle suction or hysterectomy, tissue was either snap-frozen in dry ice and stored at -70 C (for RNA extraction), fixed in neutral buffered formalin and wax embedded [for in situ hybridization (ISH) studies], or placed in ice-cold RPMI 1640 (for in vitro culture; Life Technologies, Inc., Paisley, UK) supplemented with 2 mmol L-glutamine, 100 U penicillin, and 100 µg/ml streptomycin (all from PAA Laboratories Ltd., Teddington, UK). All subjects reported regular menstrual cycles (cycle length, 25–35 d), and no women had received a hormonal preparation in the 3 months preceding biopsy collection. Biopsies were dated according to stated last menstrual period and confirmed by histological assessment according to criteria of Noyes and co-workers (20). Furthermore, circulating estradiol and progesterone concentrations at the time of biopsy were consistent for both stated last menstrual period and histological assignment of menstrual cycle stage. Ethical approval was obtained from Lothian Research Ethics Committee, and written informed patient consent was obtained before biopsy collection.

Reagents, cell culture, and treatments

PGF₂ (Sigma-Aldrich Corp., Poole, UK) was stored at -20 C as a 10-mM stock solution in ethanol. PD98059 (50 mM stock) and U73122 (4 mM stock) (Calbiochem, Nottingham, UK) were stored at -20 C in dimethylsulfoxide (DMSO). Standard reagents, unless otherwise specified, were obtained from Sigma-Aldrich Corp. Ishikawa cells (European Collection of Cell Culture, Centre for Applied Microbiology, Wiltshire, UK) were grown in DMEM/HAMS F12 medium with Glutamax (Life Technologies, Inc.) containing 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% fetal calf serum (PAA Laboratories Ltd.). The day before each experiment was started, Ishikawa cells were cultured in serum-free medium containing 3 µg/ml indomethacin.

Taqman quantitative RT-PCR and standard RT-PCR

RNA was extracted from endometrial biopsies and Ishikawa cells with Tri-reagent (Sigma-Aldrich Corp.), following the manufacturer’s guidelines. Once extracted and quantified, RNA samples were reverse-transcribed using MgCl₂ (5.5 mM), deoxynucleotide triphosphates (0.5 mM each), random hexamers (2.5 µM), ribonuclease inhibitor (0.4 U/µl), and multiscribe reverse transcriptase (1.25 U/µl; all from PE Applied Biosystems, Warrington, UK). The mix was aliquoted into individual tubes (16 µl/tube) with template RNA (4 µl/tube of 100 ng/µl RNA). After mixing by brief centrifugation, samples were incubated for 90 min at 25 C, 45 min at 48 C, and 5 min at 95 C. Thereafter, cDNA samples were stored at -20 C. A tube with no reverse transcriptase was included to control for any DNA contamination.

To measure cDNA expression, a reaction mix was prepared containing Taqman buffer (5.5 mM MgCl₂, 200 µM deoxy-ATP, 200 µM deoxy-CTP, 200 µM deoxy-GTP, 400 µM deoxyuridine 5-triphosphate), ribosomal 18S forward and reverse primers (50 nM) and probe (50 nM), forward and reverse primers for FP receptor (300 nM), FP receptor probe (100 nM), AmpErase UNG (0.01 U/µl), and AmpliTaq Gold DNA Polymerase (0.025 U/µl); PE Applied Biosystems). After mixing, 48 µl was aliquoted into separate tubes and 2 µl/replicate (40 ng) of cDNA added and mixed before placing duplicate 24-µl samples into a 96-well Taqman PCR plate (PE Applied Biosystems). A no-template control (containing water) was included in triplicate. Wells were sealed with optical caps, and the PCR reaction was carried out using an ABI Prism 7700. FP receptor primers and probe for quantitative PCR were designed using the PRIMER express program (PE Applied Biosystems). The sequence of the FP receptor primers and probe were: forward, 5'-GCAGCTGCGCTTCTTTCAA-3'; reverse, 5'-CACTGTCATGAAGATTACTGAAAAAAATAC-3'; probe (FAM-labeled, 6-carboxyfluorescein), 5'-CACAACCTGCCAGACGGAAAACCG-3'. The ribosomal 18S primers and probe sequences were: forward, 5'-CGGCTACCACATCCAAGGAA-3'; reverse, 5'-GCTGGAATTACCGCGGCT-3'; probe, (VIC-labeled), 5'-TGCTGGCACCAGACTTGCCCTC-3'. Data were analyzed and processed using Sequence Detector v1.6.3 (PE Applied Biosystems) as instructed by the manufacturer. Briefly, the software calculates the reaction cycle number at which sample fluorescence reaches a determined level for both 18S control and FP receptor. This difference in cycle number is the relative abundance of FP receptor in each sample; and, by comparing with an internal positive control, relative expression can be determined. Results are expressed as mean relative expression vs. the internal positive standard ± SEM.

To assess FP receptor expression in Ishikawa cells, standard RT-PCR was carried out on RNA extracted from Ishikawa cells as above. Reverse transcription (RT) was carried out using the same protocol as quantitative PCR, then 2 µl cDNA was added to standard PCR mix containing forward 5'-CACAACCTGCCGACGGAAAACCG-3' and reverse 5'-CGACGCCTGAATTTTATAGTCTCGATG-3' primers. These primers are at position 207 and 697 bp of the coding region of the FP receptor and were designed to amplify a fragment of 490 bp. To amplify by PCR, sample mix was denatured at 94 C for 2 min and subjected to 35 cycles of 94 C for 30 sec, 63 C for 30 sec, and 72 C for 40 sec, with a final extension step of 72 C for 7 min. After amplification, samples were cooled to 4 C, and 10 µl of the PCR mix was visualized on a 1% agarose gel. Plasmid pcDNAI/Amp, containing the human FP receptor cDNA [generously donated by Mark Abramovitz, Merck Frosst Canada Ltd., Kirkland, Quebec, Canada (4)], was used as a positive control; and Ishikawa RNA, subjected to RT in the absence of reverse transcriptase (RTase) enzyme, was used as a negative control.

ISH

FP receptor oligonucleotide double-fluorescein isothiocyanate (FITC)-labeled cDNA probes were obtained from Biognostik GmbH (Gottingen, Germany). Sections (5-µm) were cut onto Gelatin-coated Superfrost slides (BDH Laboratory Supplies, Butterworth, UK) from full-thickness human uterine biopsies collected across the menstrual cycle (n = 18). Tissue was dewaxed in xylene, rehydrated using increasing concentrations of ethanol, then proteinase-K-digested [100 µg/ml in 100 mM Tris-HCl (pH 7.6) containing 50 mMEDTA], for 15 min at 37 C, to enhance cDNA probe access. After washing in diethylpyrocarbonate-H₂O, sections were prehybridized for 4 h at 30 C with hybridization mixture (50 µl; supplied with probe) before adding cDNA probe (6 U/ml hybridization mix) and incubating overnight at 30 C. Posthybridization washes of 1x saline sodium citrate for 5 min (twice) and 0.1x saline sodium citrate at 42 C for 15 min (twice) were carried out, and the FITC-labeled probe was detected using standard immunocytochemical reagents (TSA Biotin System; NEN Life Science Products, Hounslow, UK). Endogenous peroxidase activity was first blocked with 3% H₂O₂ in methanol for 30 min, before blocking nonspecific binding with the supplied buffer for 30 min. Conjugated anti-FITC-HRP (Roche Molecular Biochemicals, Lewes, UK) was added in blocking buffer, and the sections were incubated for 60 min. After washing, biotinyltyramide amplification reagent was applied to each slide and incubated for 15 min. Streptavidin-HRP was applied after washing, incubated for 30 min, and washed, and probe localization was visualized with diaminobenzidine. Control oligonucleotide double-FITC-labeled cDNA probe, containing the same proportion of cysteine (C) and guanine (G) bases as the FP receptor probe, was included to assess background hybridization. All treatments were carried out at room temperature unless otherwise specified.

Protein assay and Western blotting

To measure ERK phosphorylation, Ishikawa cells were seeded in 6-well plates at 2.5 x 10⁵ cells/well and serum-starved overnight (3 separate experiments were set up for each treatment). Cells were stimulated with 100 nM PGF₂ in serum-free medium containing 3 µg/ml indomethacin for 10 and 30 min. Inhibitors (50 µM PD98059 or 10 µM U73122) were added, 1 h before PGF₂, with control cells receiving equal vehicle concentrations. After incubation with PGF₂, the reaction was stopped on ice, and Ishikawa cells were lysed in 250 µl/well lysis buffer [150 mM NaCl; 50 mM Tris-HCl (pH 7.4); 10 mM EDTA; 0.6% Noridet P40; 1 mM Na₃VO₄; 10% glycerol; 10 µg/ml pepstatin; 1 mM phenylmethylsulfonylfluoride] for 30 min. Lysate was collected (by scraping the wells), clarified by centrifugation at 15,300 x g for 10 min at 4 C, before storing at -20 C. Protein concentration was determined using the modified Lowry method (D2 Protein Assay kit; Bio-Rad Laboratories, Inc., Hemel Hempstead, UK). For Western analysis, 20 µg protein in standard loading buffer [25 mM Tris-HCl (pH 6.8); 0.8% SDS; 1% 2-mercaptoethanol; 4% glycerol; 0.01% bromophenol blue] was loaded, per lane, on a 4–12% Tris glycine gel and separated by SDS-PAGE (40 mA per gel, for 90 min). Proteins were transferred to a nitrocellulose membrane (Millipore Corp. (UK) Ltd., Watford, UK) that was then blocked in TNS-Tween (50 mM Tris-HCl, 150 mM NaCl, and 0.05% vol/vol Tween 20) containing 5% albumin wt/vol, for 1 h, before probing with antibodies. Sequentially, the membrane was probed with anti-phospho-ERK1/2 antibody (1:1000 with 5% albumin wt/vol; Autogen Bioclear, Calne, UK), antirabbit alkaline-phosphatase-conjugated IgG (1:30000; Sigma, UK), and detected using a fluorescence detection system (ECF; Amersham Biosciences UK Ltd., Little Chalfont, UK). Between all treatments, membranes were washed three times with TNS-Tween. To control for equal loading, the membranes were reprobed with anti-ERK (1:1000 with 5% albumin wt/vol; Autogen Bioclear), followed by antimouse alkaline-phosphatase-conjugated IgG (1:30,000; Sigma), and visualized using a luminol detection system. Changes in ERK1/2 phosphorylation were then quantified, relative to total ERK expression, and plotted as mean fold-increase above basal expression ± SEM.

Total inositol phosphate assays

Measurement of total inositol phosphate production was carried out using a standard protocol (21). Briefly, tissue samples (n = 4) or Ishikawa cells (1 x 10⁵ cells/well in 6-well plates; 4 separate experiments) were incubated with inositol-free DMEM containing 1% dialyzed heat-inactivated FCS and 0.5 µCi/well myo-[³H]inositol (Amersham Pharmacia Biotech) for 48 h. Medium was removed, and cells were incubated, for 1 h at 37 C in 1 ml buffer (140 mM NaCl, 20 mM HEPES, 4 mM KCl, 8 mM glucose, 1 mM MgCl₂, 1 mM CaCl₂, 1 mg/ml BSA, 10 mM LiCl) with 10 µM U73122 or 0.1% DMSO (vehicle). Subsequently, 1–100 nM PGF₂ in DMEM was added for 1 h at 37 C (control cells received DMEM alone) before terminating reactions on ice, aspirating medium, and permeablizing cells in 500 µl ice-cold 10-mM formic acid for 30 min on ice. Total [³H] inositol phosphate was separated from the formic acid cell extracts on AG 1-X8 anion exchange resin (Bio-Rad Laboratories, Inc.) and eluted with a 1-M ammonium formate/0.1-M formic acid solution. The associated radioactivity was determined by liquid scintillation counting and plotted, relative to protein concentrations determined using a D2 Protein Assay kit (Bio-Rad Laboratories, Inc.).

Proliferation assay

Proliferation of Ishikawa cells in response to PGF₂ was investigated (in five independent experiments) using a BrdU incorporation ELISA (Roche Diagnostics GmbH, Mannheim, Germany). For each experiment, Ishikawa cells were seeded in 96-well plates at 5 x 10³ cells/well and, once adhered, starved overnight to synchronize cell cycle. Medium was replaced with fresh medium containing 1–100 nM PGF₂, and cells were incubated for 24 h (control cells received medium alone). Cells were then labeled with 10 µM BrdU for 4 h and fixed, and BrdU incorporation was assessed using a standard ELISA technique. BrdU incorporation in Ishikawa cells is presented as a percentage of untreated cells and plotted as mean ± SEM (n = 5).

To ascertain the signaling pathway mediating PGF₂-induced proliferation, Ishikawa cells were pretreated for 1 h with either 50 µM PD98059 or 2 µM U73122 (control cells received 0.1% DMSO vehicle) before the addition of 100 nM PGF₂. As above, 10 µM BrdU was added for 4 h, and proliferation was assessed by ELISA. Results for each inhibitor are expressed as percentage of control and plotted as average ± SEM (n = 4 for each treatment).

Statistics

Where appropriate, data were subjected to statistical analysis with ANOVA and Fisher’s protected least significant differences tests (Statview 4.0; Abacus Concepts Inc., Piscataway, NJ), and statistical significance was accepted when P value was less than 0.05.

	Results

Top Abstract Introduction Subjects and Methods Results Discussion References

 
The temporal pattern of expression of FP receptor was assessed by Taqman quantitative RT-PCR using RNA samples of human endometrium collected across the menstrual cycle (Fig. 1). FP receptor expression was significantly higher in mid- to late-proliferative endometrium (0.40 ± 0.02; P < 0.05), when compared with early-proliferative (0.06 ± 0.02), early-secretory (0.09 ± 0.01), and late-secretory (0.05 ± 0.01) phases of the menstrual cycle.

View larger version (11K): [in this window] [in a new window]   Figure 1. Quantitative RT-PCR expression of FP receptor in normal human endometrium from early-proliferative (EP), mid- to late-proliferative (MP/LP), early-secretory (ES), and late-secretory (LS) phases of the menstrual cycle (n = 3 for each group; *, P < 0.05). Results are expressed as relative FP receptor mRNA expression levels (control sample = 1) and are plotted as mean ± SEM.

View larger version (184K): [in this window] [in a new window]   Figure 2. ISH for FP receptor in (A) early-proliferative, (B) mid-proliferative, (C) late-proliferative, (D) early-secretory, (E) mid-secretory, and (F) late-secretory human endometrium. FP receptor mRNA is localized to epithelial cells (G denotes glandular epithelium) in mid- and late-proliferative endometrium (B and C, respectively) and perivascular cells across the cycle (V denotes blood vessel; B, D, and E). Scale bar, 50 µm.

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View larger version (12K): [in this window] [in a new window]   Figure 3. Fold increase in inositol phosphate release in human endometrium treated with 1–100 nM PGF2 for 1 h at 37 C (n = 4; *, P < 0.05). Results are expressed as mean ± SEM fold-increase, relative to control tissue.

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View larger version (21K): [in this window] [in a new window]   Figure 4. RT-PCR for FP receptor in Ishikawa cells. Lane 1, DNA ladder; lanes 2 and 3, Ishikawa cell mRNA reverse-transcribed in the presence (lane 2) or absence (lane 3) of RTase, respectively; lane 4, a fragment of FP receptor amplified by PCR from pcDNAI/Amp plasmid containing the full FP receptor cDNA (positive control).

View larger version (17K): [in this window] [in a new window]   Figure 5. Fold increase in inositol phosphate release in Ishikawa cells treated with 1–100 nM PGF2, for 1 h at 37 C, in the absence or presence of 10 µM U73122 (n = 4; *, P < 0.05, compared with control values). U73122 was added to the cells, 1 h before stimulation with PGF2. Results are expressed as mean ± SEM fold increase, relative to control tissue.

View larger version (33K): [in this window] [in a new window]   Figure 6. ERK1/2 phosporylation in Ishikawa cells after treatment with 100 nM PGF2, for 10 and 30 min, in the presence or absence of 10 µM U73122. A, Western blot analysis of phosphorylated ERK1/2 and total ERK1/2 expression; B, mean ± SEM increase in phosphorylated ERK1/2 band intensity standardized to total ERK1/2 band intensity (n = 3 for each treatment group; *, P < 0.05, compared with untreated cells).

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View larger version (20K): [in this window] [in a new window]   Figure 7. Effect of PGF2 on Ishikawa cell proliferation. A, BrdU incorporation for 4 h in Ishikawa cells after 20 h of incubation with 1–100 nM PGF2. Data are presented as mean ± SEM percentage increase in BrdU incorporation from untreated control cells (n = 5; *, P < 0.05, compared with control cells). B, BrdU incorporation in Ishikawa cells after treatment with 100 nM PGF2, in the absence (Control) or presence of 50 µM PD98059 or 2 µM U73122 (*, P < 0.05, compared with 100 nM PGF2 alone). Data are presented as mean ± SEM percentage increase in BrdU incorporation from untreated control cells (n = 4 for each group).

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	Discussion

Top Abstract Introduction Subjects and Methods Results Discussion References

 
In the human uterus, PGF₂ induces potent myometrial contractions (18); and the loss of this effect, with resultant failure of parturition, is the major phenotype in the FP receptor knockout mouse (19). Studies have shown that myometrial cells release large amounts of PGF₂ in response to inflammatory stimuli and are likely to be the source of PGF₂ that induces myometrial contraction in vivo (22). Within the uterus, PGF₂ is also synthesized in the functionalis layer of human endometrium, particularly from epithelial cells derived from proliferative phase biopsies (15). However, the target cells and function of endometrial-epithelial-cell-derived PGF₂ within the human endometrium remains to be elucidated.

FP receptor expression in the human endometrium is temporally regulated, with the highest level of expression detected in proliferating endometrium. Moreover, expression of the FP receptor is localized predominantly to the epithelial cells. The FP receptors in the human endometrium are functional, because treatment with exogenous PGF₂ activates the PLC pathway and release of inositol phosphate. Taken together, the data indicate that PGF₂ in the human endometrium acts in an autocrine/paracrine manner, during the proliferative phase, to regulate epithelial cell function in vivo.

The factors that regulate expression of the FP receptor during the proliferative phase in the human endometrium are not clear. However, the temporal expression pattern for FP receptor mimics estrogen concentrations in women with normal reproductive cycles (23), suggesting a positive correlation between estrogen and FP receptor levels. Although regulation of the FP receptor by estrogen in the human endometrium has not been investigated, steroid hormones have been shown to affect expression of the FP receptor in other model systems. In the sheep corpora luteum and rat uterus, estrogen can promote FP receptor expression, whereas progesterone suppresses expression (24, 25). Analysis of the FP receptor promoter regions in the rat and bovine genes identifies a GC-rich promoter sequence containing several putative transcription initiation sites but no classic estrogen-response elements (26, 27). Estrogen may, nevertheless, be regulating FP receptor expression through an alternate signaling pathway downstream of the estrogen receptor. Similar observation has been reported for estrogen-induced COX-2 gene expression in human mammary epithelial cells (28). In this system, estrogen elicits PKC-mediated activation of p38 and ERK1, leading to activator protein-1 activity and transcription initiation through the cAMP response element.

In the human endometrium, production of PGF₂ by separated epithelial cells is greatest in the proliferative phase of the menstrual cycle (15), and this study confirms a similar temporal pattern of expression of the FP receptor in these cells. This temporal up-regulation of PGF_2a synthesis and expression of the FP receptor during the proliferative phase suggests that PGF₂ may be associated with promotion of glandular epithelial cell proliferation. To this end, the Ishikawa human endometrial epithelial cell line was used in this study to ascertain the potential role of PGF₂ in proliferation. Ishikawa cells were shown to express the FP receptor and to activate inositol phosphate release and ERK1/2 phosphorylation. Phosphorylation of ERK1/2 by PGF₂ in these cells is activated through a PLC-dependent signaling pathway, given that coincubation of the cells with U73122 reduced ERK1/2 phosphorylation. Recent data demonstrate that GPCRs, in addition to activating G-proteins, also activate MAPK cascades that are potent regulators of cell differentiation, development, and apoptosis (29). These kinases also play a central role in mitogenesis signaling, given that impeding their function prevents cell proliferation in response to a number of growth-stimulating agents (30). ERK activation in response to PGF₂ has already been demonstrated in luteal (31) and granulosa (32) cells, although the exact role of PGF₂-induced ERK phosphorylation in these systems is not clarified.

In the Ishikawa cells, PGF₂ is capable of inducing a concentration-dependent increase in proliferation. The induction of proliferation in these cells is dependent on activation of the PLC pathway, because inhibition of this pathway abolished the PGF₂-mediated proliferation. Previous studies have outlined the potential role of PLC in proliferation; and, in the NIH-3T3 mouse fibroblast cell line, PGF₂-induced DNA synthesis is dependent on PLC activation (33). Loss of the FP receptor expression from epithelial cells during the secretory phase of the cycle is suggestive of the loss of the proliferative potential of PGF_2a during the latter phase of the menstrual cycle. Although ERK activation was elucidated in Ishikawa cells, PGF₂-induced proliferation is not mediated through this pathway, because proliferation in response to PGF₂ was not reduced in the presence of the MEK inhibitor. This suggests that ERK activation by PGF₂ may promote other functions in endometrial epithelial cells. One potential role is the promotion of epithelial cell survival and inhibition of apoptosis. Inhibition of the ERK pathway has been shown previously to induce G1 arrest, decreased expression of antiapoptotic genes, promotion of caspase enzyme activity, and induction of programmed cell death by apoptosis (34).

In addition to epithelial cells, FP receptor expression is localized in the perivascular cells within the human endometrium across the menstrual cycle. This expression is most likely localized within the smooth-muscle layer of the perivascular region (35), where FP possibly mediates vasopressor effects that have been associated with PGF₂ function in the vasculature (36).

In summary, the data presented in this manuscript demonstrate functional expression of FP receptors in epithelial cells of the human endometrium during the proliferative phase of the menstrual cycle. In addition, PGF₂ induces proliferation of human endometrial epithelial cells via a PLC signaling pathway. Future studies will outline further the diversity of signaling pathways and phenotypic effects that may be associated with the role of PGF₂ in the glandular epithelial cells of the human endometrium.

	Acknowledgments

 
We thank Prof. Hilary Critchley for kindly allowing us access to endometrial wedge biopsies and Sheila Wright for technical assistance.

	Footnotes

 
Abbreviations: COX, Cyclooxygenase; DMSO, dimethylsulfoxide; FITC, fluorescein isothiocyanate; FP, PGF₂ receptor; GPCR, G-protein-coupled receptor; ISH, in situ hybridization; MEK, MAPK kinase; PG, prostaglandin; PLC, phospholipase C; RT, reverse transcription.

Received August 30, 2002.

Accepted December 30, 2002.

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