Molecular Human Reproduction, Vol. 7, No. 8, 731-739,
August 2001
© 2001 European Society of Human Reproduction and Embryology
Ovary and oogenesis |
The ovulatory gonadotrophin surge stimulates cyclooxygenase expression and prostaglandin production by the monkey follicle
1 Division of Reproductive Sciences, Oregon Regional Primate Research Center, 505 NW 185th Avenue, Beaverton, OR 97006 and 2 Department of Physiology and Pharmacology, Oregon Health Sciences University, 3181 SW Sam Jackson Road, Portland, OR 97201, USA
Abstract
Follicular prostaglandin concentrations increase following the gonadotrophin surge in domestic animals and rodents ~10 h before follicle rupture, suggesting a unifying role for prostaglandins in the timing of ovulation. However, little is known about prostaglandin production by the primate ovulatory follicle. In this study, adult female macaques received gonadotrophins to promote follicular development. Granulosa cells, follicular fluid, and ovaries were collected before (0 h) and 12, 24 or 36 h after administration of the ovulatory stimulus, human chorionic gonadotrophin (HCG). Cyclooxygenase (COX) isoform expression was assessed by reverse transcription–polymerase chain reaction and immunocytochemistry and follicular prostaglandin production was determined by enzyme immunoassay. COX-2 mRNA expression in granulosa cells was low at 0 h, rose 50-fold by 12 h, and remained elevated through to 36 h. COX-2 immunostaining was present in granulosa cells after, but not before, exposure to HCG. COX-1 mRNA levels did not change during the periovulatory interval, and COX-1 immunostaining of granulosa cells was not detected. Follicular fluid prostaglandin (PG) E
2
and PGF
2
concentrations were low through to 24 h but increased 100-fold at 36 h. The elevated follicular prostaglandin concentrations 4–16 h before the expected time of ovulation support the hypothesis that the time between the LH surge and increased follicular prostaglandins determines the length of the periovulatory period. Differences between the localization and timing of COX-2 expression in monkey versus non-primate follicles suggest that the pattern of COX-2 expression and activity has aspects unique to primates.
cyclooxygenase/follicle/gonadotrophin surge/ovulation/prostaglandin
Introduction
A critical role for the LH surge in initiating periovulatory events in mammals is well established. However, many of the specific gonadotrophin-stimulated processes culminating in oocyte maturation, follicle rupture, and luteinization of the follicle wall are poorly understood. Prostaglandins (PG) are putative intraovarian mediators of some of the periovulatory actions of the ovulatory gonadotrophin surge (
Murdoch et al., 1993
). The two isoforms of cyclooxygenase (also known as prostaglandin endoperoxide synthase and prostaglandin G/H synthase), COX-1 and COX-2, which catalyse the rate limiting step in the production of PG (
Vane et al., 1998
), are both present in the mammalian ovary. In rodents and domestic animals, ovarian COX-1 expression was low or non-detectable throughout the periovulatory interval (
Wong and Richards, 1991
;
Sirois, 1994
;
Sirois and Dore, 1997
). In contrast, COX-2 expression by granulosa cells and PG concentrations in follicular fluid of large preovulatory follicles rose dramatically in response to the ovulatory gonadotrophin surge (
Wong and Richards, 1991
;
Sirois, 1994
;
Sirois and Dore, 1997
). While the interval between the onset of the ovulatory gonadotrophin surge and follicle rupture varies for rodents (14 h), cows (28–30 h), and horses (36–48 h), COX-2 expression in granulosa cells and elevated PG concentrations occurs in each of these species ~10 h before follicle rupture. This observation led Sirois to propose that elevated COX-2 activity and the resulting increase in follicular fluid PG is the determining factor controlling the timing of ovulation in mammals (
Sirois and Dore, 1997
).
Prostaglandins appear to play a critical role in follicle rupture. Deletion of COX-2 gene expression in knockout mice decreases the number of ovulation sites per ovary (
Lim et al., 1997
), while mice lacking COX-1 expression have normal ovulation and fertilization (
Langenbach et al., 1995
). While administration of COX inhibitors prevents ovulation in several mammalian species including rats (
Mikuni et al., 1998
), rabbits (
Zanagnolo et al., 1996
), and sheep (
Murdoch, 1996
), limited information is available regarding PG production and action in primate follicles around the time of ovulation. PGE
2
and PGF
2
have been measured in follicular fluid obtained from women (
Jeremy et al., 1987
;
Priddy et al., 1989
), and the administration of non-selective COX inhibitors to monkeys (
Wallach et al., 1975a
) and women (
Killick and Elstein, 1987
) has been shown to block ovulation. However, a causal link between the ovulatory gonadotrophin surge and stimulation of PG production by the periovulatory follicle has not been demonstrated for primates. In these studies, we used adult female rhesus monkeys to determine if the ovulatory gonadotrophin surge stimulates COX expression and PG production by the periovulatory follicle in a temporal manner consistent with a critical role for PG in mediating follicle rupture in primates.
Materials and methods
Animals
The general care and housing of rhesus monkeys (
Macaca mulatta
) at the Oregon Regional Primate Research Center (ORPRC) were described previously (
Molskness et al., 1987
). Animal protocols and experiments were approved by the ORPRC Animal Care and Use Committee, and studies were conducted in accordance with the NIH Guide for the Care and Use of Laboratory Animals. Adult females with regular menstrual cycles were checked daily for menses, and blood samples were obtained daily from unanaesthetized monkeys by saphenous venipuncture beginning on the first day of treatment. Serum was stored at –20°C. Serum oestradiol (
Resko et al., 1975
) and progesterone (
Hess et al., 1981
) concentrations were measured by radioimmunoassay; intra- and interassay coefficients of variation did not exceed 10%. LH concentrations were determined by the mouse Leydig cell bioassay (
Ellinwood and Resko, 1980
) using monkey LH RP-1 as the standard (supplied by the NIH Hormone Distribution Program); intra- and interassay coefficients of variation for the LH bioassay did not exceed 15%. These assays were performed by the Endocrine Services Laboratory, ORPRC.
An ovarian stimulation model developed for the collection of multiple oocytes for IVF (
VandeVoort et al., 1989
) was used to obtain granulosa cells and follicular fluid (
n
= 3–5/time point
). Beginning within 3 days of initiation of menstruation, rhesus monkeys received 60 IU of recombinant human (r-h)FSH (Ares Advanced Technology, Inc., Randolph, MA, USA; days 1–6), followed by 60 IU of r-hFSH plus 60 IU r-hLH (AAT, Inc.; days 7–9) to stimulate the growth of multiple follicles. Animals also received the gonadotrophin-releasing hormone antagonist Antide (AAT, Inc.; 0.5 mg/kg body weight) daily to prevent an endogenous ovulatory LH surge. Adequate follicular development was monitored by serum oestradiol concentrations and by ultrasonography (
Wolf et al., 1996
). Follicular aspiration was performed on anesthetized animals during aseptic surgery before (0 h) and 12, 24 or 36 h after administration of 1000 IU recombinant human chorionic gonadotrophin (HCG) (AAT, Inc.; day 10). In spontaneous menstrual cycles, follicle rupture in rhesus monkeys occurs ~40 h after the ovulatory gonadotrophin surge (
Weick et al., 1973
), so the observation times span the periovulatory interval. Previous studies verified ovulation sites on ovaries and oocytes in the oviducts following this protocol (
VandeVoort et al., 1989
;
Hibbert et al., 1996
). 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 >4 mm in diameter were pooled.
Whole ovaries (
n
= 2–4/time point
) were also obtained from monkeys undergoing ovarian stimulation. Additional whole ovaries (
n
= 5
) were collected from monkeys experiencing spontaneous menstrual cycles around the expected time of the endogenous LH surge (
Weick et al., 1973
). These ovaries were obtained before the LH surge (
n
= 2
), during the LH surge (
n
= 2
), and after the LH surge (
n
= 1
) based on serum oestradiol, LH and progesterone concentrations before and at the time of organ removal. Rhesus monkey testis was obtained at necropsy.
Tissue preparation
Granulosa cells and follicular fluid were obtained from follicular aspirates as described previously (
Chaffin et al., 1999a
). Briefly, aspirates were subjected to centrifugation to pellet the granulosa cells, and the resulting supernatants (i.e. follicular fluid) were removed and stored at –80°C. A granulosa cell-enriched population was obtained by Percoll gradient centrifugation. Total RNA was obtained from granulosa cells using Trizol reagent according to the manufacturer's (Gibco-BRL, Rockville, MD, USA) instructions and was stored at –20°C. Whole ovaries and testis were fixed in 4% paraformaldehyde and embedded in paraffin.
COX-1 and COX-2 mRNA analysis
Total RNA (200 ng) was treated with DNase (Gibco-BRL) prior to reverse transcription (RT), which was performed using Molony Murine Leukemia Virus reverse transcriptase (Gibco-BRL) as previously reported (
Chaffin et al., 1999b
). Semiquantitative RT–polymerase chain reaction (PCR) based assays similar to those previously described by this laboratory (
Chaffin et al., 1999b
) were developed to assess COX-1 and COX-2 mRNA levels. Sequences for the oligonucleotides (Gibco-BRL) used for PCR are shown in Table I
. The MgCl
2
concentration, amount of cDNA included in each PCR reaction, number of PCR cycles, and primer concentrations (for which the amount of co-amplified products for the experimental gene and the internal standard gene, cyclophilin, were linear and parallel with an increasing amount of cDNA) were determined empirically for each primer set. The PCR cycle conditions were 92°C for 30 sec, 60°C for 1 min and 72°C for 1 min. The products of both primer sets were also in the exponentially increasing phase relative to the number of PCR cycles. Data are expressed as the ratio of the gene of interest to cyclophilin for each sample assayed. Sequence analysis was used to confirm the identity of PCR products. For the regions amplified in these experiments, the nucleic acid sequences of monkey COX-1 and COX-2 were 99% and 98% identical to the corresponding human sequences. Amplification using the COX-1 primers yielded two bands, one of which was determined not to be COX-1 by DNA sequencing. The identity of this PCR product is currently unknown. For this reason, only the band determined to represent COX-1 by sequence analysis was considered in the COX-1 assay; the unidentified band did not interfere with quantitative analysis of COX-1 mRNA. PCR products were separated on 2% agarose gels and photographed using Polaroid 667 film (Polaroid Corp., Cambridge, MA, USA). For each mRNA, all samples were assayed in duplicate in a single experiment. Intra-assay coefficient of variation was <15%.
|
PGE 2 and PGF 2
concentrations in follicular fluidPreliminary experiments indicated the presence of a substance in follicular fluid which interfered with the measurement of PG, so all samples were extracted before assay. Follicular fluids were acidified with HCl (final concentration 0.1 mol/l), and [ 3 H]PGE 2 or [ 3 H]PGF 2
(Dupont, Boston, MA, USA) was added to each sample to allow determination of PG recovery. Ethyl acetate (four volumes) was added to each sample, and samples were vortexed vigorously. The phases were allowed to separate at room temperature, and the organic layer was then transferred to a fresh tube and dried under nitrogen. Samples were resuspended in assay buffer (from enzyme immunoassay kit, see below), and an aliquot was subjected to scintillation counting for calculation of PG recovery, which averaged 65%. Concentrations of PGE
2
and PGF
2
in monkey follicular fluid extracts were determined by enzyme immunoassay using commercially available kits (Cayman Chemical, Ann Arbor, MI, USA), and the PG content of each sample was corrected based on PG recovery calculated for that sample. Intra- and interassay coefficients of variation averaged <10% and <20% respectively.
Immunocytochemical detection of COX-1 and COX-2
Immunocytochemical detection of COX-1 and COX-2 in ovarian tissues was performed with paraffin-embedded tissues using mouse monoclonal antibodies generated against sheep COX-1 and a synthetic peptide based on the human COX-2 sequence (Cayman Chemical) (
Sekhon et al., 1999
). Briefly, 5 µm tissue sections were deparaffinized in xylene and rehydrated through a decending alcohol series and in phosphate-buffered saline (PBS: 0.05 mol/l NaH
2
PO
4
, 0.14 mol/l NaCl, pH 7.4
). Antigen was retrieved using Antigen Retrieval Citra treatment (BioGenex Labs, San Ramon, CA, USA), and endogenous peroxidase was quenched with 2% hydrogen peroxide in methanol. After blocking with 3% non-immune serum, sections were incubated with the primary antibody at room temperature for 1 h, followed by overnight incubation at 4°C. After rinsing with PBS, sections were incubated with biotinylated secondary antibody and then with peroxide conjugated avidin solution (Vector Laboratories, Inc., Burlingame, CA, USA). All non-immune serum and antibody incubations were performed in PBS with 0.1% Triton X-100 (Sigma Chemical Co., St Louis, MO, USA). Peroxidase was visualized using nickel-DAB chromagen (Vector). The COX-1 antibody was used at a concentration of 10 µg/ml while the COX-2 antibody was used at 3 µg/ml.
Specificity of COX-2 staining was demonstrated by preabsorbing the antibody overnight at 4°C with the human COX-2 peptide used to generate this antibody at 100-fold excess (Cayman Chemical). An appropriate protein for preabsorption of the COX-1 antibody was not available. For this reason, paraffin-embedded monkey testicular tissue was used as a positive control tissue for COX-1 immunostaining. Digital images of tissue sections were obtained using a Zeiss Axioplan microscope (Carl Zeiss, New York, NY, USA) and ColorSNAP software (Photometrics, Ltd, Tucson, AZ, USA).
Data analysis
All gels were scanned, and specific bands were analysed densitometrically using NIH Image 1.40 (Research Services Branch, NIMH, Bethesda, MD, USA), which compared band size and intensity to a standard curve and calculated the optical density of each band. All data were assessed for heterogeneity of variance using Bartlett's test and log transformed when necessary. Analyses were performed using one-way analysis of variance, followed by Newman–Keuls' test when indicated. Data are presented as mean ± SEM, and significance was assumed at
P
< 0.05.
Results
Cyclooxygenase mRNA expression in periovulatory follicles
COX-2 mRNA levels in granulosa cells changed across the periovulatory interval (Figure 1
, upper panel
). COX-2 mRNA levels were low or non-detectable before the administration of the ovulatory dose of HCG (0 h). By 12 h after HCG administration, COX-2 mRNA levels rose 50-fold over levels seen at 0 h (
P
< 0.05
) and remained elevated through to 36 h. COX-1 mRNA was detected in all granulosa cell samples obtained at 0, 12 and 36 h after HCG, and the levels did not change across the periovulatory interval (Figure 1
, lower panel
). COX-1 mRNA was inconsistently detected in granulosa cells obtained 24 h after HCG; for this reason, data are not presented for this time point.
|
PGE 2 and PGF 2
concentrations in follicular fluidPGE 2 (Figure 2
, upper panel
) and PGF
2
(Figure 2
, lower panel
) concentrations in periovulatory follicular fluid samples were also determined. Concentrations of both PGs were low at 0 h and remained low through to 24 h. However, follicular fluid concentrations of both PGE
2
and PGF
2
were highest at 36 h (
P
< 0.05
), peaking at concentrations >100-fold greater than those measured at 24 h.
|
Cyclooxygenase immunostaining in periovulatory follicles
COX-2 immunostaining was detectable in monkey ovaries as a perinuclear or cytoplasmic dark precipitate (Figure 3A–I
)
. Preabsorption of the COX-2 antibody with the human COX-2 peptide used to generate this antibody (Figure 3E
) resulted in elimination of COX-2 staining when compared with an adjacent section incubated with the COX-2 antibody (Figure 3D
). Similar results were obtained when the COX-2 antibody was omitted (not shown). These data support the specificity of this antibody for detection of the monkey COX-2 protein.
|
Ovaries were obtained from monkeys undergoing ovarian stimulation before (0 h) and 12, 24 or 36 h after the administration of an ovulatory dose of HCG and processed for immunocytochemical detection of COX-2. In large follicles of ovaries obtained at 0 h (Figure 3A
;
n
= 3
) or 12 h (Figure 3B
;
n
= 2
) after HCG administration, granulosa and theca cells showed little or no COX-2 immunoreactivity. Follicles of ovaries obtained 24 h after HCG consistently showed COX-2 immunostaining in both the granulosa and theca cell layers (Figure 3C
;
n
= 4
), and strong granulosa and theca cell immunostaining was observed 36 h after HCG administration (Figure 3D
;
n
= 4
). Primary and secondary follicles as well as antral follicles <1 mm in diameter consistently lacked COX-2 immunostaining as did the ovarian surface epithelium (Figure 3F
). In ovaries from animals undergoing ovarian stimulation, antral follicles >2 mm occasionally showed COX-2 immunoreactivity in the theca, but not granulosa, cell layer (not shown).
COX-2 immunostaining was also examined in five ovaries obtained from monkeys experiencing spontaneous menstrual cycles at times which spanned the periovulatory interval. In the large (>5 mm) follicle of two ovaries obtained before the ovulatory LH surge, granulosa and theca cells showed little or no COX-2 immunoreactivity (Figure 3G
). Two ovaries were obtained during the LH surge; both the granulosa and surrounding theca cells of these large follicles showed COX-2 immunoreactivity (Figure 3H
). An additional ovary was obtained from a monkey with serum hormone concentrations and follicle morphology consistent with the period near the time of follicle rupture. Strong COX-2 immunostaining was observed in the granulosa and theca cells of this large periovulatory follicle (Figure 3I
). In these five ovaries, the primary, secondary, and antral follicles <1 mm in diameter were generally devoid of COX-2 immunostaining in both the granulosa and theca cell layers, although staining in the theca cells surrounding larger (2–3 mm) antral follicles was observed (Figure 3H
).
The specificity of immunocytochemical detection of monkey COX-1 protein was determined using monkey testicular tissue as a control. A dark precipitate representing detection of COX-1 protein was seen in the cytoplasm of epithelial cells of the rete testis, while spermatozoa and seminiferous tubule cells were consistently negative for COX-1 (Figure 3J
). No staining was observed when the primary antibody was omitted (Figure 3K
). These findings are consistent with previous reports of immunocytochemical and histochemical localization of COX-1 in the rodent reproductive tract (
Johnson and Ellis, 1977
;
Marshburn et al., 1989
) and support the use of this antibody for COX-1 detection in monkey tissues.
In monkey ovaries, COX-1 protein was not detected in the granulosa cells or stroma surrounding the follicles in any tissue section examined (Figure 3L
and data not shown
). This included primary, secondary, antral, large preovulatory, and periovulatory follicles from monkeys undergoing ovarian stimulation or experiencing spontaneous menstrual cycles. COX-1 immunoreactivity was however observed in the ovarian surface epithelium (Figure 3L
), and this immunoreactivity was not seen when the COX-1 antibody was omitted (Figure 3M
).
Discussion
This study is the first to demonstrate a temporal relationship between the ovulatory gonadotrophin surge, COX-2 expression by the periovulatory follicle, and follicular PG production in primates. Levels of COX-1 mRNA in monkey granulosa cells of large antral follicles did not change across the periovulatory interval, and COX-1 protein was not detected in the granulosa or theca cells of these same follicles. These data suggest that COX-1 is not a major source of PG in primate periovulatory follicles and are consistent with results from other mammalian species (
Wong and Richards, 1991
;
Sirois, 1994
;
Sirois and Dore, 1997
). COX-2 mRNA and protein concentrations increased in response to an ovulatory dose of gonadotrophin, and follicular fluid concentrations of PGE
2
and PGF
2
rose between 24 and 36 h after HCG administration. These data are comparable to changes in COX-2 expression and follicular fluid PG seen in other mammalian species (
Wong and Richards, 1991
;
Sirois, 1994
;
Sirois and Dore, 1997
), consistent with the hypothesis that the time between the ovulatory gonadotrophin surge and increased follicular PG production determines the length of the periovulatory interval (
Sirois and Dore, 1997
). However, the interval between initiation of COX-2 expression and the periovulatory rise in follicular fluid PG concentrations appears to be longer in primates than in other mammalian species. In addition, COX-2 was present in the theca, as well as granulosa, cells of the periovulatory follicle, in contrast with reports of an absence of COX-2 expression in the theca of domestic animals (
Liu et al., 1997
;
Sirois and Dore, 1997
). This disparity in both the temporal expression and localization of COX-2 between primates and other mammalian species suggests that important differences exist in the pathway regulating hormonal induction of COX-2 expression and modulation of PG production in periovulatory follicles.
COX-2 expression and PG production by the granulosa cells of primate periovulatory follicles appears to differ somewhat from that reported for rodents and domestic animals. In these other species, granulosa cell COX-2 mRNA and protein concentrations increase ~10 h before follicle rupture, with elevated follicular fluid PG concentrations being measured ~4 h before ovulation (
Wong and Richards, 1991
;
Sirois, 1994
;
Sirois and Dore, 1997
). A different pattern was seen in the monkey (present study). COX-2 mRNA levels increased ~28 h before the expected time of ovulation, and COX-2 protein was consistently detected in the granulosa and theca cells of periovulatory follicles 16 h before ovulation. However, PG production did not escalate until 4–16 h before the projected time of follicle rupture. Taken together, these data suggest that component(s) of the PG synthetic pathway other than COX-2 may limit PG production during these early hours of the periovulatory interval in primates. COX-2 protein was detected in the granulosa cells of follicles obtained from monkeys experiencing natural menstrual cycles after, but not before, the endogenous LH surge. These data are consistent with the pattern of COX-2 expression in ovaries from monkeys undergoing ovarian stimulation and support the use of this model in future experiments.
COX-2 protein was first detected in the theca surrounding monkey periovulatory follicles 24–36 h after HCG administration as well as after the initiation of the endogenous LH surge in natural cycles. This observation is consistent with descriptions of cyclooxygenase activity in human theca cells (
Patwardhan and Lanthier, 1981
) but is in contrast to previous reports of an absence of COX-2 protein in theca cells of domestic animals (
Liu et al., 1997
;
Sirois and Dore, 1997
). COX-2 was also present in theca cells surrounding some small antral follicles both before and after the ovulatory gonadotrophin surge, suggesting that exposure to high gonadotrophin concentrations may not be required for COX-2 expression in these cells. While it is unknown if PG of thecal origin contribute to the periovulatory rise in follicular fluid PG concentrations, theca-derived PG may play an important role in the growth, development, or rupture of the primate follicle.
COX-2 expression and PG production are regulated by several pathways in ovarian granulosa cells. In the non-primate species studied to date, ovulatory doses of gonadotrophin are required to initiate COX-2 expression in granulosa cells; lower concentrations of FSH and LH present during natural and gonadotrophin-stimulated cycles which support follicular development do not initiate COX-2 expression in these species (
Sirois, 1994
;
Liu et al., 1997
;
Sirois and Dore, 1997
). A cAMP-response element has been found in the promoter region of the rat (
Sirois et al., 1993
) and human (
Kosaka et al., 1994
)
COX-2
genes, consistent with activation of
COX-2
gene expression through the LH/HCG receptor. Recently, forskolin treatment was shown to increase COX-2 expression by bovine granulosa cells
in vitro
, demonstrating a direct relationship between elevated intracellular cAMP and initiation of COX-2 expression in these cells (
Liu et al., 2000
). Other mediators can also modulate COX-2 expression or activity in granulosa cells. Interleukin-1 (IL-1) has been seen to stimulate COX-2 expression in dispersed cells from the rat ovary (
Ando et al., 1998
) and human luteinizing granulosa cells (
Narko et al., 1997
)
in vitro
. Nitric oxide can enhance PG production by rabbit periovulatory follicles (
Yamauchi et al., 1997
), most likely through a mechanism other than regulation of COX-2 synthesis (
Goodwin et al., 1999
). Progesterone can decrease COX-2 expression by rat granulosa cells
in vitro
with equivocal effects on PG production (
Hellberg et al., 1996
;
Hedin and Eriksson, 1997
). However, no apparent effect of periovulatory progesterone depletion (
Chaffin and Stouffer, 1999
) has been observed on monkey granulosa cell COX-2 mRNA and follicular fluid PG concentrations (Duffy and Stouffer, unpublished data). While it is clear that the ovulatory gonadotrophin surge initiates granulosa cell COX-2 expression in several mammalian species, further study is needed to understand how intraovarian factors modulate COX-2 expression and activity as well as PG synthesis by the primate periovulatory follicle.
Follicular PGs may play an important role in many periovulatory processes. Administration of non-selective COX inhibitors to monkeys (
Wallach et al., 1975a
) and women (
Killick and Elstein, 1987
) has been shown to prevent ovulation; this effect could be reversed in monkeys by simultaneous administration of PGF
2
(
Wallach et al., 1975b
). A recent report of reversible infertility in women taking COX inhibitors for treatment of arthritic symptoms also supports the critical role of cyclooxygenase activity in reproductive processes (
Mendonca et al., 2000
). In rodents, non-selective or COX-2-specific PG synthesis inhibitors can also prevent ovulation when administered systemically (
Tsafriri et al., 1972
;
Mikuni et al., 1998
) or to perfused whole ovaries (
Hamada et al., 1978
;
Sogn et al., 1987
;
Mikuni et al., 1998
); ovulation can be restored by PGE
2
or PGF
2
co-administration (
Hamada et al., 1978
;
Sogn et al., 1987
). Mice lacking COX-2 have a decreased rate of ovulation (
Lim et al., 1997
) due, at least in part, to a defect in follicle rupture that can be restored with exogenous administration of PGE
2
(
Davis et al., 1999
). Further support for a key role for PGs in follicle rupture comes from studies showing that inhibition of PG production reduces the gonadotrophin- stimulated rise in proteolytic enzymes involved in follicle rupture in rats (
Reich et al., 1991
). However, normal or near-normal ovulatory efficiency has been observed in knockout mice lacking specific receptors for PGE
2
(
Hizaki et al., 1999
;
Tilley et al., 1999
) or PGF
2
(
Sugimoto et al., 1997
), so the PG(s) involved in mediating gonadotrophin-stimulated follicle rupture remain unknown. Defects in cumulus expansion have also been noted in knockout mice lacking expression of COX-2 (
Davis et al., 1999
) or the PGE receptor, EP2 (
Hizaki et al., 1999
;
Tilley et al., 1999
), suggesting an additional pathway by which PGs may reduce fertility. In monkeys (
Wallach et al., 1975a
) and women (
Killick and Elstein, 1987
) receiving non-selective COX inhibitors, normal luteal phase progesterone has been observed along with an increased incidence of luteinized unruptured follicles, suggesting that rupture, and not luteinization, was altered by PG synthesis inhibition in primates. The detection of COX-1 in the ovarian surface epithelium supports the hypothesis that prostaglandins produced by these cells may play a role in ovulation. Studies in sheep suggest that doses of indomethacin which inhibit ovulation also prevent apoptosis of ovarian surface epithelium covering the follicle apex, a process thought to be essential for follicle rupture (
Murdoch, 1996
). Collectively, these reports and current data indicate that PGs are likely mediators of follicle rupture in primates. However, identification of individual PGs and their specific roles in the regulation of various periovulatory processes awaits further study.
In summary, we have demonstrated in the rhesus monkey that the periovulatory follicle expresses COX-2 and produces PGs in response to an ovulatory dose of gonadotrophin. These data are consistent with a role for PGs in mediating the action of the LH surge to trigger follicle rupture. The ovulatory LH surge appears to be essential for COX-2 expression in the granulosa cells of the periovulatory follicle, but the regulation of the expression and activity of COX-2 and other enzymes involved in PG synthesis in the primate follicle remain poorly understood. While the data in the present study and others are suggestive, a direct link between PG action in the ovary and follicle rupture remains to be established for primates, including monkeys and women. If further studies demonstrate that locally produced PGs are required for successful ovulation in primates, this pathway may provide a useful target for the development of novel contraceptives.
Acknowledgements
The authors thank the excellent animal care and surgical staff at ORPRC for their assistance with these studies. We also appreciate the assistance of Dr Charles Chaffin in providing monkey tissues and follicular fluids for use in this study. Recombinant human gonadotrophins and Antide were generously provided by Ares Advanced Technology, Inc. This research was supported by NICHD/NIH through cooperative agreement (HD18185) as part of the Specialized Cooperative Centers Program in Reproductive Research. Dr Duffy is a Junior Investigator supported by the Andrew W.Mellon Center in Reproductive Biology, ORPRC. These studies were supported by the Medical Research Foundation of Oregon (D.M.D.), the Andrew W.Mellon Foundation (D.M.D.), and NIH grants HD20869 (R.L.S.), HD18185, and RR00163.
Notes
3 To whom correspondence should be sent. E-mail: duffyd{at}ohsu.edu 
References
Ando, M., Kol, S., Kokia, E. et al. (1998) Rat ovarian prostaglandin endoperoxide sythase-1 and -2: periovulatory expression of granulosa cell-based interleukin-1-dependent enzymes. Endocrinology, 139, 2501–2508.
Chaffin, C.L. and Stouffer, R.L. (1999) Expression of matrix metalloproteinases and their tissue inhibitor messenger ribonucleic acids in macaque periovulatory granulosa cells: time course and steroid regulation. Biol. Reprod., 61, 14–21.
Chaffin, C.L., Hess, D.L. and Stouffer, R.L. (1999a) Dynamics of periovulatory steroidogenesis in the rhesus monkey follicle after ovarian stimulation. Hum. Reprod., 14, 642–649.
Chaffin, C.L., Stouffer, R.L. and Duffy, D.M. (1999b) Gonadotropin and steroid regulation of steroid receptor and aryl hydrocarbon receptor mRNA in macaque granulosa cells during the periovulatory interval. Endocrinology, 140, 4753–4760.
Davis, B.J., Lennard, D.E., Lee, C.A. et al. (1999) Anovulation in cyclooxygenase-2-deficient mice is restored by prostaglandin E2 and interleukin-1ß. Endocrinology, 140, 2685–2695.
Duffy, D.M. and Stouffer, R.L. (1995) Progesterone receptor messenger ribonucleic acid in the primate corpus luteum during the menstrual cycle: possible regulation by progesterone. Endocrinology, 136, 1869–1876.[Abstract]
Ellinwood, W.E. and Resko, J.A. (1980) Sex differences in biologically active, immunoreactive gonadotropins in the fetal circulation of the rhesus monkey. Endocrinology, 107, 902–907.[Abstract]
Funk, C.D., Funk, L.B., Kennedy, M.E. et al. (1991) Human platelet/erythroleukemia cell prostaglandin G/H synthase: cDNA cloning, expression, and gene chromosomal assignment. FASEB J., 5, 2304–2312.[Abstract]
Goodwin, D.C., Landino, L.M. and Marnett, L.J. (1999) Effects of nitric oxide and nitric oxide-derived species on prostaglandin endoperoxide synthase and prostaglandin biosynthesis. FASEB J., 13, 1121–1136.
Hamada, Y., Wright, K.H. and Wallach, E.E. (1978) In vitro reversal of indomethacin-blocked ovulation by prostaglandin F2. Fertil. Steril., 30, 702–706.[ISI][Medline]
Hedin, L. and Eriksson, A. (1997) Prostaglandin synthesis is suppressed by progesterone in rat preovulatory follicles in vitro. Prostaglandins, 53, 91–106.[ISI][Medline]
Hellberg, P., Larson, L., Olofsson, J. et al. (1996) Regulation of the inducible form of prostaglandin endoperoxide synthase in the perfused rat ovary. Mol. Hum. Reprod., 2, 111–116.
Hess, D.L., Spies, H.G. and Hendrickx, A.G. (1981) Diurnal steroid patterns during gestation in the rhesus macaque: onset, daily variation, and the effects of dexamethasone treatment. Biol. Reprod., 24, 609–616.[Abstract]
Hibbert, M.L., Stouffer, R.L., Wolf, D.P. et al. (1996) Midcycle administration of a progesterone synthesis inhibitor prevents ovulation in primates. Proc. Natl Acad. Sci. USA, 93, 1897–1901.
Hizaki, H., Segi, E., Sugimoto, Y. et al. (1999) Abortive expansion of the cumulus and impaired fertility in mice lacking the prostaglandin E receptor subtype EP2. Proc. Natl Acad. Sci. USA, 96, 10501–10506.
Hla, T. and Neilson, K. (1992) Human cyclooxygenase-2 cDNA. Proc. Natl Acad. Sci. USA, 89, 7384–7388.
Jeremy, J.Y., Okonofua, F.E., Thomas, M. et al. (1987) Oocyte maturation and human follicular fluid prostanoids, gonadotropins, and prolactin after administration of clomiphene and pergonal. J. Clin. Endocrinol. Metab., 65, 402–406.[Abstract]
Johnson, J.M. and Ellis, L.C. (1977) The histochemical localization of prostaglandin synthetase activity in reproductive tract of the male rat. J. Reprod. Fertil., 51, 17–22.[Abstract]
Killick, S. and Elstein, M. (1987) Pharmacologic production of luteinized unruptured follicles by prostaglandin synthetase inhibitors. Fertil. Steril., 47, 773–777.[ISI][Medline]
Kosaka, T., Miyata, A., Ihara, H. et al. (1994) Characterization of the human gene (PTGS2) encoding prostaglandin-endoperoxide synthase 2. Eur. J. Biochem., 221, 889–897.[ISI][Medline]
Langenbach, R., Morham, S.G., Tiano, H.F. et al. (1995) Prostaglandin synthase 1 gene disruption in mice reduces arachidonic acid-induced inflammation and indomethacin-induced gastric ulceration. Cell, 83, 483–492.[ISI][Medline]
Lim, H., Paria, B.C., Das, S.K. et al. (1997) Multiple female reproductive failures in cyclooxygenase 2-deficient mice. Cell, 91, 197–208.[ISI][Medline]
Liu, J., Carriere, P.D., Dore, M. et al. (1997) Prostaglandin G/H synthase-2 is expressed in bovine preovulatory follicles after the endogenous surge of luteinizing hormone. Biol. Reprod., 57, 1524–1531.[Abstract]
Liu, J., Antaya, M., Boerboom, D. et al. (2000) The delayed activation of the prostaglandin G/H synthase-2 promoter in bovine granulosa cells is associated with down-regulation of truncated upstream stimulatory factor-2. J. Biol. Chem., 274, 35037–35045.
Marshburn, P.B., Clark, M.R. and Shabanowitz, R.B. (1989) Immunohistochemical localization of prostaglandin H synthase in the epididymis and vas deferens of the mouse. Biol. Reprod., 41, 491–497.[Abstract]
Mendonca, L.L., Khamashta, M.A., Nelson-Piercy, C. et al. (2000) Non-steroidal anti-inflammatory drugs as a possible cause for reversible infertility. Rheumatology (Oxford), 39, 880–882.
Mikuni, M., Pall, M., Peterson, C.M. et al. (1998) The selective prostaglandin endoperoxide synthase-2 inhibitor, NS-398, reduces prostaglandin production and ovulation in vivo and in vitro in the rat. Biol. Reprod., 59, 1077–1083.
Molskness, T.A., VandeVoort, C.A. and Stouffer, R.L. (1987) Stimulatory and inhibitory effects of prostaglandins on the gonadotropin-sensitive adenylate cyclase in the monkey corpus luteum. Prostaglandins, 34, 279–290.[ISI][Medline]
Murdoch, W.J. (1996) Differential effects of indomethacin on the sheep ovary: prostaglandin biosynthesis, intracellular calcium, apoptosis, and ovulation. Prostaglandins, 52, 497–506.[ISI][Medline]
Murdoch, W.J., Hansen, T.R. and McPherson, L.A. (1993) A review | role of eicosanoids in vertebrate ovulation. Prostaglandins, 46, 85–115.[ISI][Medline]
Narko, K., Ritvos, O. and Ristimaki, A. (1997) Induction of cyclooxygenase-2 and prostaglandin F2 receptor expression by interleukin-1ß in cultured human granulosa-luteal cells. Endocrinology, 138, 3638–3644.
Patwardhan, V.V. and Lanthier, A. (1981) Prostaglandins PGE and PGF in human ovarian follicles: endogenous contents and in vitro formation by theca and granulosa cells. Acta Endocrinol., 97, 543–550.
Priddy, A.R., Killick, S.R., Elstein, M. et al. (1989) Ovarian follicular fluid eicosanoid concentrations during the pre-ovulatory period in humans. Prostaglandins, 38, 197–202.[ISI][Medline]
Reich, R., Daphna-Iken, D., Chun, S.Y. et al. (1991) Preovulatory changes in ovarian expression of collagenases and tissue metalloproteinase inhibitor messenger ribonucleic acid: role of eicosanoids. Endocrinology, 129, 1869–1875.[Abstract]
Resko, J.A., Ploem, J.G. and Stadelman, H.L. (1975) Estrogens in fetal and maternal plasma of the rhesus monkey. Endocrinology, 97, 425–430.[Abstract]
Sekhon, HS, Jia, Y., Raab, R. et al. (1999) Prenatal nicotine increases pulmonary alpha7 nicotinic receptor expression and alters fetal lung development in monkeys. J. Clin. Invest., 103, 637–647.[ISI][Medline]
Sirois, J. (1994) Induction of prostaglandin endoperoxide synthase-2 by human chorionic gonadotropin in bovine preovulatory follicles in vivo. Endocrinology, 135, 841–848.[Abstract]
Sirois, J. and Dore, M. (1997) The late induction of prostaglandin G/H synthase-2 in equine preovulatory follicles supports its role as a determinant of the ovulatory process. Endocrinology, 138, 4427–4434.
Sirois, J., Levy, L.O., Simmons, D.L. et al. (1993) Characterization and hormonal regulation of the promotor of the rat prostaglandin endoperoxide synthase 2 gene in granulosa cells. Identification of functional and protein-binding regions. J. Biol. Chem., 268, 12199–12206.
Sogn, J.H., Curry, T.E. Jr, Brannstrom, M. et al. (1987) Inhibition of follicle-stimulating hormone-induced ovulation by indomethacin in the perfused rat ovary. Biol. Reprod., 36, 536–542.[Abstract]
Sugimoto, Y., Yamasaki, A., Segi, E. et al. (1997) Failure of parturition in mice lacking the prostaglandin F receptor. Science, 277, 681–683
Tilley, S.L., Audoly, L.P., Hicks, E.H. et al. (1999) Reproductive failure and reduced blood pressure in mice lacking the EP2 prostaglandin E2 receptor. J. Clin. Invest, 103, 1539–1545.[ISI][Medline]
Tsafriri, A., Lindner, H.R. and Lamprecht, S.A. (1972) Physiological role of prostaglandins in the induction of ovulation. Prostaglandins, 2, 1–10.[ISI][Medline]
VandeVoort, C.A., Baughman, W.L. and Stouffer, R.L. (1989) Comparison of different regimens of human gonadotropins for superovulation of rhesus monkeys: ovulatory response and subsequent luteal function. J. In Vitro Fertil. Embryo Transfer, 6, 85–91.[ISI][Medline]
Vane, J.R., Bakhle, Y.S. and Botting, R.M. (1998) Cyclooxygenases 1 and 2. Annu. Rev. Pharmacol. Toxicol., 38, 97–120.[ISI][Medline]
Wallach, E.E., de la Cruz, A., Hunt, J. et al. (1975a) The effect of indomethacin on hMG-hCG induced ovulation in the rhesus monkey. Prostaglandins, 9, 645–658.[ISI][Medline]
Wallach, E.E., Bronson, R., Hamada, Y. et al. (1975b) Effectiveness of prostaglandin F2 in restoration of hMG-hCG induced ovulation in indomethacin-treated rhesus monkeys. Prostaglandins, 10, 129–138.[ISI][Medline]
Weick, R.F., Dierschke, D.J., Karsch, F.J. et al. (1973) Periovulatory time course of circulating gonadotropic and ovarian hormones in the rhesus monkey. Endocrinology, 93, 1140–1147.[ISI][Medline]
Wolf, D.P., Alexander, M., Zelinski-Wooten, M.B. et al. (1996) Maturity and fertility of rhesus monkey oocytes collected at different intervals after an ovulatory stimulus (human chorionic gonadotropin) in in vitro fertilization cycles. Mol. Reprod. Dev., 43, 76–81.[ISI][Medline]
Wong, W.Y.L. and Richards, J.S. (1991) Evidence for two antigenically distinct molecular weight variants of prostaglandin H synthase in the rat ovary. Mol. Endocrinol., 5, 1269–1279.[Abstract]
Yamauchi, J., Miyazaki, T., Iwasaki, S. et al. (1997) Effects of nitric oxide on ovulation and ovarian steroidogenesis and prostaglandin production in the rabbit. Endocrinology, 138, 3630–3637.
Zanagnolo, V., Dharmarajan, A.M., Endo, K. et al. (1996) Effects of acetylsalicylic acid (aspirin) and naproxen sodium (naproxen) on ovulation, prostaglandin, and progesterone production in the rabbit. Fertil. Steril., 65, 1036–1043.[ISI][Medline]
Submitted on December 29, 2000; accepted on June 4, 2001.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  B. L Dozier, K. Watanabe, and D. M Duffy Two pathways for prostaglandin F2{alpha} synthesis by the primate periovulatory follicle Reproduction, July 1, 2008; 136(1): 53 - 63. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. L. Tootle and A. C. Spradling Drosophila Pxt: a cyclooxygenase-like facilitator of follicle maturation Development, March 1, 2008; 135(5): 839 - 847. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. Feuerstein, V. Cadoret, R. Dalbies-Tran, F. Guerif, R. Bidault, and D. Royere Gene expression in human cumulus cells: one approach to oocyte competence Hum. Reprod., December 1, 2007; 22(12): 3069 - 3077. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Sayasith, K. A Brown, and J. Sirois Gonadotropin-dependent regulation of bovine pituitary adenylate cyclase-activating polypeptide in ovarian follicles prior to ovulation Reproduction, February 1, 2007; 133(2): 441 - 453. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E.-M. Tsai, T.-F. Chan, Y. Chang, P.-H. Chiang, C.-Y. Chuang, C.-Y. Long, C.-Y. Chai, and J.-N. Lee Leptin Suppresses Human Chorionic Gonadotropin-Induced Cyclooxygenase-2 Expression and Prostaglandin Production in Cultured Human Granulose Luteal Cells Reproductive Sciences, December 1, 2006; 13(8): 551 - 557. [Abstract] [PDF]
|
|
|
|
 |
|
 Y. L. Pon and A. S. T. Wong Gonadotropin-Induced Apoptosis in Human Ovarian Surface Epithelial Cells Is Associated with Cyclooxygenase-2 Up-Regulation via the {beta}-Catenin/T-Cell Factor Signaling Pathway Mol. Endocrinol., December 1, 2006; 20(12): 3336 - 3350. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. Markosyan, B. L. Dozier, F. A. Lattanzio, and D. M. Duffy Primate Granulosa Cell Response via Prostaglandin E2 Receptors Increases Late in the Periovulatory Interval Biol Reprod, December 1, 2006; 75(6): 868 - 876. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. J. Bridges, C. M. Komar, and J. E. Fortune Gonadotropin-Induced Expression of Messenger Ribonucleic Acid for Cyclooxygenase-2 and Production of Prostaglandins E and F2{alpha} in Bovine Preovulatory Follicles Are Regulated by the Progesterone Receptor Endocrinology, October 1, 2006; 147(10): 4713 - 4722. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. M. Duffy, C. L. Seachord, and B. L. Dozier An Ovulatory Gonadotropin Stimulus Increases Cytosolic Phospholipase A2 Expression and Activity in Granulosa Cells of Primate Periovulatory Follicles J. Clin. Endocrinol. Metab., October 1, 2005; 90(10): 5858 - 5865. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. M. Duffy, C. L. Seachord, and B. L. Dozier Microsomal prostaglandin E synthase-1 (mPGES-1) is the primary form of PGES expressed by the primate periovulatory follicle Hum. Reprod., June 1, 2005; 20(6): 1485 - 1492. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. L. Seachord, C. A. VandeVoort, and D. M. Duffy Adipose Differentiation-Related Protein: A Gonadotropin- and Prostaglandin-Regulated Protein in Primate Periovulatory Follicles Biol Reprod, June 1, 2005; 72(6): 1305 - 1314. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. El-Nefiawy, K. Abdel-Hakim, and N. Kanayama The selective prostaglandin EP4 agonist, APS-999 Na, induces follicular growth and maturation in the rat ovary Eur. J. Endocrinol., February 1, 2005; 152(2): 315 - 323. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. M. Duffy, B. L. Dozier, and C. L. Seachord Prostaglandin Dehydrogenase and Prostaglandin Levels in Periovulatory Follicles: Implications for Control of Primate Ovulation by Prostaglandin E2 J. Clin. Endocrinol. Metab., February 1, 2005; 90(2): 1021 - 1027. [Abstract] [Full Text] [PDF]
|
|