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Department of Physiological Sciences, Eastern Virginia Medical School, Norfolk, Virginia 23507
Address all correspondence and requests for reprints to: Diane M. Duffy, Department of Physiological Sciences, Eastern Virginia Medical School, 700 Olney Road, Lewis Hall, Norfolk, Virginia 23507. E-mail: duffydm{at}evms.edu.
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PGE2 production by the periovulatory follicle is initiated by the ovulatory gonadotropin surge, which induces the expression of PG synthesis enzymes by the granulosa cells. Arachidonic acid is synthesized de novo, taken up by the cell, or cleaved from membrane phospholipids by phospholipase A2. Cyclooxygenase (COX)-1 or COX-2 then converts arachidonic acid into PGH2, which is then converted to PGE2 through the activity of PGE synthase. Gonadotropin control of COX-2 expression by granulosa cells is well established (9, 10, 11); gonadotropins can also induce granulosa cell expression of phospholipase A2 (12, 13) and PGE synthase (14). In rodents and domestic animals, follicular fluid PGE2 levels rise rapidly after expression of COX-2 is detected (9, 10, 15), supporting the hypothesis that induction of COX-2 by the ovulatory gonadotropin surge is the rate-limiting step in follicular PGE2 production and ovulation (10). COX-2 expression by the granulosa cells of primate periovulatory follicles is also induced by the ovulatory gonadotropin surge (11). However, gonadotropin-stimulated COX-2 mRNA expression precedes maximal follicular fluid PGE2 levels by 24 h in primate periovulatory follicles. Although gonadotropin-induced granulosa cell COX-2 expression is necessary for PGE2 production, COX-2 may not catalyze the rate-limiting step controlling the elevation in follicular PGE2 levels necessary for follicle rupture in primates.
Although it is known that PGE2 can be metabolized to biologically inactive 15-keto PGE2 derivatives through the activity of an NAD+-dependent 15-hydroxy PG dehydrogenase (PGDH) and then further metabolized by a combination of enzymatic and nonenzymatic processes (16), the expression and activity of PGDH in the primate periovulatory follicle has not been examined. The lung is the major site of PGDH activity; PGs are generally released into the blood and metabolized as they pass through the lungs (17). However, examples do exist of PG-regulated tissues in which PG metabolism by PGDH plays a key physiologic role. For example, placental PGs likely play a key role in the initiation of parturition in several mammalian species; decreased PGDH expression and increased PG concentrations have been observed at term (18, 19). Because the pattern of COX-2 expression cannot, in itself, explain the pattern of follicular fluid PGE2 concentrations seen throughout the primate periovulatory interval, we hypothesized that PGE2 may be produced and then rapidly metabolized by PGDH during the middle portion of the periovulatory interval. This rapid metabolism of PGE2 may delay the intrafollicular rise in PGE2 required for ovulation and contribute to the longer periovulatory interval in primates.
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Materials and Methods |
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Granulosa cells, follicular fluid, and whole ovaries were obtained from adult female cynomolgus macaques at Eastern Virginia Medical School (Norfolk, VA). All animal protocols and experiments were approved by the Eastern Virginia Medical School Animal Care and Use Committee and were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Monkeys were maintained in temperature-controlled rooms on a 12-h light, 12-h dark cycle. Animals were fed commercial monkey chow and had water available at all times. Adult females with regular menstrual cycles were checked daily for menstruation; the 1st d of menstruation was designated d 1 of the menstrual cycle. Blood samples were obtained under ketamine chemical restraint (10 mg/kg body weight) by femoral or saphenous venipuncture, and serum was stored at –20 C. Aseptic surgeries were performed in a dedicated surgical suite under isoflurane anesthesia and involved either laparoscopy or midline laparotomy.
A controlled ovarian stimulation model developed for the collection of multiple oocytes for in vitro fertilization was used to obtain monkey granulosa cells and follicular fluid (n = 4–7/time point) (20). Beginning within 3 d of initiation of menstruation, recombinant human FSH (45 IU twice daily, Serono Reproductive Biology Institute, Rockland, MA) was administered for 6 d, followed by twice daily administration of 45 IU of recombinant human FSH plus 30 IU r-hLH (Serono Reproductive Biology Institute) for 2 d to stimulate the growth of multiple follicles. The GnRH antagonist Antide (0.5 mg/kg body weight; Serono Reproductive Biology Institute) was also administered daily to prevent an endogenous ovulatory LH surge. Adequate follicular development was monitored by serum estradiol levels and ultrasonography (21). Follicular aspiration was performed before (0 h) or 12, 24, and 36 h after administration of 1000 IU r-hCG (d 9; Serono Reproductive Biology Institute). In spontaneous menstrual cycles, follicle rupture in monkeys occurs approximately 40 h after the ovulatory gonadotropin surge (22), so these times span the periovulatory interval. Previous studies in rhesus monkeys verified ovulation sites on ovaries and oocytes in the oviducts after this protocol (20, 23); preliminary experiments in cynomolgus monkeys confirmed ovulation sites in response to this protocol (not shown). To obtain undiluted follicular fluid as well as granulosa cells, each follicle was pierced with a 22-gauge needle, and the aspirated contents of all follicles larger than 4 mm in diameter were pooled.
Tissue preparation
Monkey granulosa cells and follicular fluid were obtained from follicular aspirates as described previously (24). Briefly, aspirates were subjected to centrifugation to pellet the oocytes and granulosa cells; the resulting supernatant (i.e. follicular fluid) was removed and stored at –80 C. Oocytes were mechanically removed, and a granulosa cell-enriched population of the remaining cells was obtained by Percoll gradient centrifugation (24). Viability of granulosa cell-enriched preparations averaged 80% as assessed by trypan blue exclusion. Cells were either used immediately for cell culture or were frozen in liquid nitrogen and stored at –80 C for preparation of total RNA or cell lysates. Whole ovaries were bisected, maintaining at least two periovulatory follicles greater than 4 mm in diameter on each piece. Ovarian tissue was fixed in 4% paraformaldehyde and embedded in paraffin.
Real-time RT-PCR
PGDH mRNA levels were analyzed by real-time RT-PCR using a Roche LightCycler (Indianapolis, IN). Total RNA was obtained from granulosa cells using Trizol reagent (Invitrogen, Rockville, MD) and was stored at –80 C. Total RNA was incubated with DNase, and RT was performed as described previously (25). PCR was performed using the FastStart DNA Master SYBR Green I kit (Roche) following manufacturer’s instructions using 0.5 mM of each primer, 4.0 mM MgCl2, and an annealing temperature of 57 C. PGDH and ß-actin mRNA content of each sample were determined in independent assays. PCR primers for PGDH (up, 5'-TCCAGTGCGATGTGGC; down, 5'-GCAACGGGCATGAGTC) and ß-actin (up, 5'-ATCCGCAAAGACCTGT; down, 5'-GTCCGCTAGAAGCAT) were designed based on the human sequences (accession nos. NM_00860 and NM_00101, respectively) using LightCycler Probe Design software (Roche). PCR products were sequenced (Microchemical Core Facility, San Diego State University, CA). PCR products were 256 (accession no. AB059653) and 270 (accession no. AY765990) bp in length for PGDH and ß-actin, respectively. The nucleic acid identity between the monkey and human sequences was 98.8% for PGDH and 97.4% for ß-actin; the monkey PGDH product shared 100% sequence identity with the previously reported monkey PGDH sequence (accession no. AB059653). At least 5 log dilutions of the sequenced PCR product was included in each assay and used to generate a standard curve. All data were expressed as the ratio of PGDH mRNA to ß-actin mRNA for each sample. Intra- and interassay coefficients of variation were less than 10%.
Western blotting for PGDH
Granulosa cells were thoroughly lysed on ice in PBS containing 0.5% sodium dodecyl sulfate and 0.1% Triton X-100, mixed with denaturing sample buffer, heated to 95 C for 10 min, and loaded onto 4–20% gradient polyacrylamide Tris-HCl gels (Bio-Rad, Hercules, CA). Each experiment included at least four lanes of serial-diluted granulosa cell lysate; detection and densitometric analysis of the protein of interest in these samples was used to generate a standard curve for semiquantitative analysis of granulosa cell lysates. Proteins were transferred to polyvinylidene difluoride membranes (Immobilon, Millipore, Billerica, MA), and Western blotting proceeded as previously reported for monkey progesterone receptors (26), except that the urea denaturation step was omitted. The anti-PGDH primary antibody (2.0 µg/ml) was a rabbit polyclonal generated against a synthetic human PGDH peptide (Cayman Chemical, Ann Arbor, MI); an antirabbit IgG-horseradish peroxidase conjugate secondary antibody (Amersham, Piscataway, NJ) was used at a dilution of 1:20,000. Bands were detected by chemiluminescence (ECL, Amersham). Blots were then stripped of primary and secondary antibodies following instructions provided by the membrane manufacturer, and Western blotting was performed on the stripped membranes using a mouse antitubulin primary antibody (1:1000 dilution, Sigma-Aldrich, St. Louis, MO) and an antimouse IgG-horseradish peroxidase conjugate secondary antibody (Amersham). Molecular size of bands representing PGDH and tubulin proteins were determined in comparison with prestained standards (Bio-Rad). Films were scanned and analyzed densitometrically using SigmaGel software (Jandel Scientific, San Rafael, CA). Tubulin levels in granulosa cell lysates were not different between treatment groups. All data are expressed as a ratio of PGDH/tubulin content for each granulosa cell sample.
Immunohistochemical detection of PGDH
Immunohistochemical detection of PGDH in ovarian tissues was performed with 5-µm sections of paraffin-embedded tissues as previously described (11) using the anti-PGDH antibody described above for Western blotting (4 µg/ml) and a biotinylated bovine antirabbit IgG secondary antibody and peroxide conjugated avidin solution (Vector Laboratories, Burlingame, CA); peroxidase activity was visualized with Nova Red Chromagen (Vector Laboratories). In some experiments, the primary antibody was preabsorbed with the peptide used to generate the antibody (0.5 µg peptide/1.0 µg primary antibody) for 1 h at room temperature before incubation with tissue sections.
PGE2 and PGE2 metabolite (PGEM) concentrations in follicular fluid
Follicular fluid obtained from aspirates was acidified and extracted with ethyl acetate before assay as previously described (11). [3H]PGE2 was added to each sample before extraction to correct for procedural losses; the mass of [3H]PGE2 added was <0.1% of the total PGE2 content of each follicular fluid sample. Samples were resuspended in assay buffer (see below), and an aliquot was subjected to scintillation counting to calculate PG recovery, which averaged 84%. Concentrations of PGE2 and PGEM in monkey follicular fluid were determined by enzyme immunoassay (EIA, Cayman Chemical), and the PGE2 or PGEM content of each sample was corrected based on PGE2 recovery calculated for each sample. The intra- and interassay coefficients of variation for the PGE2 EIAs were 14.6 and 13.4%, respectively. The intra- and interassay coefficients of variation for the PGEM EIA were 7.0 and 15.8%, respectively.
Granulosa cell culture
Granulosa cells obtained from monkeys undergoing controlled ovarian stimulation before the administration of an ovulatory dose of gonadotropin (0 h) were plated on tissue culture plates coated with fibronectin and maintained at 37 C in 5% CO2 in serum-free DMEM-Ham’s F12 medium containing insulin (2 µg/ml), transferrin (5 µg/ml), selenium (0.25 nmol), aprotinin (25 mg/ml), and human low-density lipoprotein (25 µg/ml) as previously described (7). Beginning at the time of plating, cultures of 100,000 cells/well were treated with hCG (100 ng/ml, Serono Reproductive Biology Institute), progesterone (10 nM, Sigma-Aldrich), or no treatment (control) for up to 48 h. In additional experiments, granulosa cells were treated with indomethacin (100 nM, Sigma-Aldrich) to prevent endogenous PGE2 production or indomethacin with PGE2 (1 µg/ml, Cayman Chemical) for up to 48 h. Cells were lysed in situ with Trizol reagent, and total RNA was prepared with the addition of glycogen (10 µg) to improve recovery; RT-PCR was then performed as described above.
Data analysis
All data were assessed for heterogeneity of variance using Bartlett’s test and log transformed when Bartlett’s test yielded a significance of <0.05; data presented in Figs. 4
and 5 were log transformed before further analysis. PGDH mRNA levels in granulosa cells obtained before and after hCG administration in vivo, follicular fluid PGE2, and follicular fluid PGEM levels were compared using one-way ANOVA, followed by Newman-Keuls test. Tubulin levels as determined by Western blotting were assessed by the Kruskal-Wallis test. A robust statistical test could not be identified for analysis of the PGDH protein data presented in Fig. 2; the large SD at 24 h after hCG prevented parametric analysis, whereas the number of samples per time point limited the power of nonparametric analysis. Therefore, no statistical analysis of these data is presented. Analysis of PGDH mRNA in cultured granulosa cells was performed using one-way ANOVA with one repeated measure (blocked for individual animal) at each time point examined, followed by Newman-Keuls test. Data in Figs. 1, 4, and 5 are presented as mean ± SEM, and significance was assumed at P < 0.05. Individual PGDH/tubulin ratios are shown in Fig. 2, with the mean for each group indicated by the bar.
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Results |
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To determine whether PGDH was expressed by granulosa cells of the monkey periovulatory follicle in response to the ovulatory gonadotropin surge, granulosa cells were obtained at specific times which span the 40-h periovulatory interval in primates (22), and total granulosa cell RNA was assessed for PGDH mRNA levels (Fig. 1
). PGDH mRNA levels were low at 0 h hCG, rose 10-fold to peak at 12 h hCG, and were low again at 24 to 36 h after hCG administration (P < 0.05).
Granulosa cell PGDH protein content was assessed by semiquantitative Western blotting. PGDH protein levels were high 0–12 h after hCG administration, variable at 24 h after hCG, and nondetectable in all granulosa cell samples assayed by 36 h after hCG (Fig. 2). A longer exposure time for the Western blot did indicate the presence of PGDH in all samples, but the densitometric values for some samples (n = 2 at 24 h and n = 4 at 36 h) were below the linear range of the standard curve used to quantify PGDH levels.
Immunohistochemistry indicated that PGDH expression within monkey periovulatory follicles was restricted to the granulosa cells (Fig. 3). PGDH was localized to the cytoplasm of granulosa cells as well as luminal epithelial cells of the monkey seminal vesicle, previously reported to express PGDH (27). PGDH was detected in granulosa, but not theca, cells of periovulatory follicles. Moderate PGDH immunostaining was observed in granulosa cells of ovaries obtained 0, 12, and 24 h after hCG administration; lower levels of PGDH immunostaining were observed 36 h after hCG.
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In a previous report by this laboratory, expression of COX-2 mRNA was shown to be elevated 12 h after hCG administration in monkey granulosa cells (11), suggesting that PGE2 synthesis may be initiated early in the periovulatory interval. However, monkey follicular fluid PGE2 levels remained low until 36 h after hCG administration, which is 4 h before the expected time of follicle rupture (11). Therefore, we hypothesized that PGE2 levels during the periovulatory interval may be modulated by metabolism of PGE2 via the activity of PGDH present in granulosa cells; a high rate of PGE2 metabolism may continue through 24 h after hCG administration, after which PGDH protein levels decline (Figs. 2 and 3). To test this hypothesis, follicular fluid levels of PGEM and PGE2 were determined (Fig. 4). PGEM levels were low at 0 h hCG, elevated 12 h after hCG, peaked 24 h after hCG, and remained high 36 h after hCG administration (P < 0.05); PGEM levels measured at 24 h were 56-fold greater than those measured at 0 h. PGE2 levels were low 0–12 h after hCG, rose 10-fold over 0-h levels by 24 h after hCG, and increased 100-fold over 0-h levels by 36 h after hCG administration (P < 0.05).
PGDH regulation by gonadotropin, progesterone, and PGE2
The studies described above demonstrate that the ovulatory gonadotropin surge initially increases granulosa cell PGDH expression in vivo but that this increase is not sustained throughout the periovulatory interval. The ovulatory gonadotropin surge also increases follicular progesterone production in vivo (24), and progesterone has been implicated as an important regulator of PGDH expression in nonovarian tissues (28, 29). In addition, PGE2 levels in follicular fluid increase during the periovulatory interval (11) (Fig. 4), and binding of PGE2 to its EP2 receptor increases intracellular cAMP (30), which has been shown to regulate PGDH expression (29). Binding of hCG to the LH/hCG receptor on granulosa cells also increases intracellular cAMP as well as other second messengers (31). To determine whether gonadotropin or progesterone acts directly at granulosa cells to regulate PGDH mRNA levels, monkey granulosa cells obtained from large periovulatory follicles before administration of hCG (0 h) were cultured in the presence of an ovulatory concentration of hCG (100 ng/ml) or progesterone (10 nM, Fig. 5). After 24 h of culture, hCG stimulated PGDH mRNA levels above controls (P < 0.05); progesterone treatment did not alter PGDH levels. After 48 h of culture, treatment with either hCG or progesterone decreased PGDH expression when compared with control levels (P < 0.05). To determine whether PGE2 regulates PGDH expression by granulosa cells, cells were exposed to indomethacin (100 nM) to prevent endogenous PGE2 production in the absence or presence of PGE2 (1 µg/ml); PGE2 did not alter granulosa cell PGDH expression after 24 or 48 h in vitro when compared with cultures treated with indomethacin only (n = 4–5/group, not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The pattern of accumulation of PGE2 and PGEM in follicular fluid after hCG administration indicates that PGE2 is both produced and metabolized by the follicle during the periovulatory interval. Although both granulosa and theca cells of primate follicles express COX-2 (11) and produce PGE2 in vitro (7, 32, 33), detection of PGDH only in granulosa cells indicates that conversion of PGE2 into PGEM occurs solely in the granulosa cells. Low levels of both PGE2 and PGEM in follicular fluid before hCG administration likely reflect a low rate of PGE2 synthesis. Increased follicular fluid PGEM, but not PGE2, 12 h after hCG suggests that PGE2 is synthesized but is converted to PGEM via the activity of PGDH at this time. Higher follicular fluid PGE2 levels at 24 h hCG is likely due to increased PGE2 synthesis; increased PGEM levels indicate that PGE2 metabolism via PGDH also continues. The large variation among PGDH levels at 24 h after hCG may reflect a period of transition from high PGDH to low PGDH protein expression. At 36 h hCG, just before ovulation, PGE2 levels continue to rise while PGEM levels plateau; this may be due to increased PGE2 synthesis but a decreased rate of PGE2 conversion to PGEM, consistent with the low levels of PGDH protein as detected by immunohistochemistry and PGDH below the sensitivity of the semiquantitative Western blotting technique used for these studies. It is unclear why PGEM levels are maintained in follicular fluid when PGDH protein levels are greatly reduced 36 h after hCG administration. Remaining low levels of PGDH activity may be sufficient to convert PGE2 into PGEM at the same rate that PGEM is removed from follicular fluid by further metabolism. Taken together, these data are consistent with a role for PGDH to reduce follicular fluid PGE2 levels 12–24 h after hCG administration, after PGE2 synthesis has been initiated within the primate periovulatory follicle.
PGDH is the primary enzyme responsible for the metabolism of PGE2. PGDH catalyzes the conversion of PGE2 into 15-keto PGE2, a biologically inactive PG (16). The 
13–15-ketoprostaglandin reductase then converts 15-keto PGE2 into an unstable PGE2 derivative, which undergoes nonenzymatic degradation into the group of PGE2-derived metabolites measured as PGEM in our assay (34, 35). Further degradation yields molecules that can be derived from several bioactive PGs. Measurement of products specific to PGE2 metabolism likely underestimates the amount of PGE2 degraded within the follicle but does provide a relative measure of PGE2 metabolism and, therefore, PGDH activity. Conversion of PGEMs into metabolites not unique to PGE2 may explain why PGEM levels are lower than PGE2 levels in follicular fluid at all times examined. PGDH expression and activity has been shown to be hormonally regulated in the ovary (36 ; present study) and other reproductive tract tissues (18, 28), and studies using tissues such as kidney, lung, and uterus suggest that PGDH activity is often the hormonally or developmentally regulated step in PG metabolism (37, 38). The ability to directly address PGDH activity in vivo is limited because most known PGDH inhibitors either inhibit COX activity (and, therefore, PGE2 synthesis) or are ligands for PPAR
, known to be present in follicular granulosa cells (39, 40). In contrast to PGDH, 13–15-ketoprostaglandin reductase activity is not thought to be a key regulated step in PG metabolism (16). PGE2 can also be converted to PGF2
via the activity of the PGE 9-ketoreductase (41). PGE 9-ketoreductase is present in the ovaries of rabbits and sheep (42, 43), but PGE 9-ketoreductase expression in the primate follicle has not been addressed. Although PGDH is likely the primary enzyme responsible for removal of PGE2 from the follicular fluid pool, the ability of other enzymes to metabolize PGE2 must also be considered.
Gonadotropin is likely the major regulator of follicular PGDH expression during the first half of the periovulatory interval. The ovulatory gonadotropin surge increased PGDH mRNA levels in granulosa cells in vivo, and elevated granulosa cell PGDH protein levels were maintained until midway through the periovulatory interval. hCG also increased PGDH mRNA levels in granulosa cells after 24 h in vitro, confirming a direct effect of gonadotropin to increase granulosa cell PGDH expression. This ability of an ovulatory dose of gonadotropin to increase granulosa cell PGDH mRNA levels and maintain elevated PGDH protein levels is consistent with the ability of hCG to increase intracellular cAMP through interaction with the LH/CG receptor located on granulosa cells (44) and the presence of a cAMP response element in the promoter region of the human PGDH gene (29). However, exposure to PGE2, also thought to raise granulosa cell cAMP via the EP2 receptor (30), did not alter PGDH mRNA levels in vitro, and it is possible that hCG may regulate PGDH gene expression through a cAMP-independent mechanism. Taken together, these data suggest that the ovulatory surge of gonadotropin, perhaps acting via elevated granulosa cell cAMP levels, rapidly increases PGDH expression. Even though PGE2 synthesis via COX-2 may be initiated early in the periovulatory interval, PGDH activity could maintain low intrafollicular PGE2 concentrations and lengthen the time between the ovulatory gonadotropin surge and the accumulation of PGE2 within the follicle.
Decreased PGDH expression late in the periovulatory interval may be due to direct action of either gonadotropin or gonadotropin-stimulated progesterone on granulosa cells. Although exposure to an ovulatory concentration of gonadotropin increased granulosa cell PGDH mRNA levels at the earliest time points examined (12 h in vivo and 24 h in vitro), PGDH expression was not maintained and decreased in vivo by 36 h and in vitro after 48 h of exposure to gonadotropin. After 36 h of hCG exposure in vivo, PGDH protein levels in granulosa cells were very low, as evidenced by weak immunohistochemical staining and levels below the limits of detection by Western blotting. This biphasic action of gonadotropin to increase and then decrease PGDH expression may reflect the ability of the LH/CG receptor to stimulate multiple intracellular signaling molecules including cAMP and phospholipase C (31). Although cAMP can increase the activity of the human PGDH promoter (29), human trophoblasts respond to increased intracellular cAMP with decreased PGDH expression (45), suggesting that the ability of cAMP to regulate transcription of the PGDH gene is modulated by multiple factors. In addition, gonadotropin initiates the process of luteinization within granulosa cells both in vivo and in vitro, and changes associated with luteinization may contribute to the decline in PGDH expression observed after 36–48 h of gonadotropin exposure in vivo and in vitro, respectively. For example, gonadotropin stimulation increases granulosa cell progesterone production (24); elevated intrafollicular progesterone is essential for follicle rupture and successful ovulation (23, 46). In the present study, exposure of granulosa cells to progesterone decreased PGDH mRNA levels in vitro, consistent with the low PGDH mRNA levels measured in granulosa cells obtained 36 h after hCG administration in vivo. Previous studies demonstrated that hCG treatment of monkey granulosa cells in vitro stimulates progesterone production (7), and studies in nonovarian transfected cells demonstrated that progesterone stimulates the activity of the human PGDH promoter, even though the classical progesterone response element could not be identified within the promoter region (29). Interestingly, PGDH expression in placental cells is also enhanced in response to progesterone exposure (47). Additional studies using progesterone synthesis inhibitors and/or progesterone receptor antagonists in vivo and in vitro during gonadotropin exposure will be required to rigorously test the hypothesis that progesterone receptor-mediated progesterone action regulates PGDH expression in the granulosa cells of primate periovulatory follicles.
The studies presented here represent the first report of dynamic PGDH expression within the follicle throughout the periovulatory interval and suggest a role for PG metabolism in the regulation of bioactive PG levels within the primate periovulatory follicle. PGs were once thought to be produced in or near the tissues where they act, then enter the circulation for inactivation at the lung, which possesses very high PGDH activity (17). Therefore, tissue levels of PGs were thought to be regulated primarily by the rate of PGE2 synthesis, especially by the activity of COX-1 or COX-2. However, regulation of follicular PG concentrations may occur at multiple levels; availability of adequate arachidonic acid as substrate for PG synthesis, regulation of enzymes involved in PG production, and metabolism of PGs produced may play coordinated roles in the control of tissue PGE2 levels. The presence of PGDH and the timing of PGDH expression within PG-producing and -responsive tissues supports our hypothesis that PGDH plays a key regulatory role in the control of PG levels within the periovulatory follicle. Almost 2 decades ago, PGDH activity was identified in the corpora lutea of rabbits and was thought to be involved in the regulation of PGF2-induced luteolysis (36); the studies presented here extend these findings to suggest a key role for PGDH to regulate local concentrations of bioactive PGs in the periovulatory follicle as well. Therefore, the ovulatory gonadotropin surge may coordinate the expression of enzymes involved in both PG synthesis and PG metabolism within the primate periovulatory follicle to regulate a critical reproductive function, ovulation.
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Acknowledgments |
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Footnotes |
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Abbreviations: COX, Cyclooxygenase; EIA, enzyme immunoassay; PG, prostaglandin; PGDH, PG dehydrogenase; PGEM, PGE2 metabolite.
This work was supported by National Institutes of Health Grant HD38972 (to D.M.D.).
Received June 28, 2004.
Accepted October 27, 2004.
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13 reductase with specific assay methods. Prostaglandins Leukot Essent Fatty Acids 38:25–30[CrossRef][Medline]
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