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Regular Article |
a Department of Obstetrics and Gynecology, Chandler Medical Center, University of Kentucky, Lexington, Kentucky 40536-0293
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 
, and 
) in luteal tissue, ovaries were collected from gonadotropin-treated, immature rats on Days 1, 4, 8, and 14 of pseudopregnancy and from adult, cycling animals on each day of the estrous cycle. Ovaries were processed for in situ hybridization or RNA isolation for analysis by RNase protection assay. The expression of PPAR mRNA was abundant in granulosa cells of developing follicles during both pseudopregnancy and the estrous cycle and was low to undetectable in CL from pseudopregnant rats. However, luteal tissue in cycling animals, especially CL remaining from previous cycles, had high levels of PPAR mRNA. The PPAR mRNA was localized mainly in the theca and stroma, and PPAR mRNA was expressed throughout the ovary. Levels of mRNA for PPAR decreased between Days 1 and 4 of pseudopregnancy, and PPAR mRNA levels were lower on the day of estrus compared to pro- and metestrus (P < 0.05). The PPAR mRNA levels remained steady throughout the estrous cycle and pseudopregnancy. These data illustrate a difference in the luteal expression of mRNA for PPAR between the adult, cycling rat and the immature, gonadotropin-treated rat. This differential pattern of expression may be related to the difference in timing of the preovulatory prolactin surge, because the gonadotropin-primed animals would not experience a prolactin surge coincident with the LH surge, as occurs in adult, cycling animals. Additionally, the expression pattern of PPAR mRNA indicates that it may be involved in cellular functions involved with maintaining basal ovarian function, whereas PPAR may play a role in lipid metabolism in the theca and stroma.
corpus luteum, follicle, granulosa cells, ovary, theca cells
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The PPAR superfamily is comprised of three members: , , and . Each PPAR isotype is transcribed from an individual gene. The PPARs are capable of influencing gene transcription by heterodimerizing with the 9-cis, retinoic acid receptor (RXR) and binding to PPAR response elements (PPREs) in the promoter region of target genes. Some of the genes that have been shown to be regulated by the PPARs are involved in tissue remodeling, angiogenesis, and prostaglandin production [5–8].
Tissue remodeling and angiogenesis are required for restructuring of the postovulatory follicle into the CL. The PPARs could affect these processes in numerous ways. For example, PPARs have been shown to regulate the activity and expression of two proteases, gelatinase B [7, 9–11] and urokinase plasminogen activator (uPA) [5]. Another protease, stromelysin, has a PPRE in its promoter region [6], suggesting that expression of this protease is also regulated by PPARs. These proteases have been identified in ovarian and/or luteal tissue from various species (for review, see [12, 13]), indicating that PPARs could regulate protease activity and influence luteal formation and/or regression. In addition, activation of PPAR has been shown to inhibit angiogenesis both in vitro and in vivo [5]. This inhibition of blood vessel development by PPAR occurs, in part, by increasing the level of mRNA for plasminogen-activator inhibitor-1, a potent regulator of uPA [5, 14], and by decreasing the expression of mRNA for vascular endothelial growth factor receptors [5].
Prostaglandins are another group of important players in both luteal development and regression. Interestingly, PPARs are activated by arachidonic acid and its eicosanoid metabolites (for review, see [15]). In turn, PPARs can regulate prostaglandin synthesis by modulating the expression of cyclooxygenase-2 (COX-2) [8, 16, 17]. A PPRE has been identified in the COX-2 promoter that allows direct regulation of COX-2 expression by the PPARs [8]. The ability of PPARs to regulate COX-2 expression and the fact that eicosanoids can activate PPARs suggest that a feedback system may exist between the synthesis of prostaglandins and PPAR activity.
Progesterone produced by the CL is essential for the maintenance of pregnancy, as mentioned above. Agonists for PPAR have been shown to influence progesterone production by cultured rat [18] and porcine [19] granulosa cells, human granulosa-lutein cells [20], and bovine luteal cells [21]. Löhrke et al. [21] identified PPARs in bovine luteal tissue at both the mRNA and protein level. Although the amount of protein for PPAR in bovine luteal cells declined between the early and midluteal phase of the estrous cycle [21, 22], activation of PPAR in cultured midcycle bovine luteal cells stimulated progesterone secretion [21]. These data suggest that PPAR may influence luteal cell function by regulating progesterone production.
In a previous study, we showed that mRNAs corresponding to the PPAR isotypes are expressed in ovarian tissue from gonadotropin-primed, immature rats during follicular development and the periovulatory period [18]. Because PPARs are able to influence events involved with luteal development and steroidogenesis, the present study examined the expression of mRNA for the PPARs , , and in ovarian tissue from pseudopregnant rats to determine if the PPARs are present in a temporal and spatial pattern associated with the formation, function, and/or regression of the CL. We also investigated the expression of mRNA for the PPARs in ovarian tissue from cycling rats to compare the gonadotropin-driven, immature animal model to the naturally cycling adult.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All animal procedures were approved by the University of Kentucky Animal Care and Use Committee. Immature and mature, virgin Sprague-Dawley rats were kept on a 14L:10D photoperiod, with lights-on at 0400 h. To examine the cellular expression pattern of mRNA for the PPAR isotypes in luteal tissue, ovaries were collected on various days during pseudopregnancy to correspond to formation (Day 1), function (Days 4 and 8), and regression (Day 14) of the CL. On Day 23 of age, 10 IU of eCG (generously provided by Dr. A.F. Parlow, National Hormone and Peptide Program, National Institute of Diabetes, Digestive, and Kidney Diseases, Torrance, CA) were administered to the immature animals s.c., and 48 h post-eCG, animals received 10 IU of hCG (Sigma, St. Louis, MO) s.c. to trigger ovulation and luteal development. Animals were killed 1, 4, 8, and 14 days after treatment with hCG (n = 3 per time point). Ovaries were collected, snap-frozen, and stored at -70°C until RNA isolation or placed in optimal cutting temperature compound (VWR Scientific, Atlanta, GA) and stored at -70°C until sectioned for in situ hybridization.
The expression patterns of mRNAs corresponding to the PPAR isotypes were also examined in ovarian tissue from cycling animals. This allowed comparison of the immature, gonadotropin-treated model with the naturally cycling animal as well as investigation of PPAR expression in luteal tissue of various ages. Adult animals (aged 50 days) were monitored for reproductive cyclicity by daily examination of vaginal cytology. Only those rats demonstrating at least two normal, 4-day estrous cycles were utilized. Ovaries were removed from animals under ether anesthesia between 1000 and 1100 h on the morning of the proestrous, estrous, metestrous, and diestrous (n = 3 animals/day) stage of the cycle. All ovaries were processed as described above.
In Situ Hybridization
Ovaries collected from animals during pseudopregnancy and the estrous cycle were sectioned (thickness, 8 µm) and mounted on ProbeOn Plus slides (Fisher Scientific, Pittsburgh, PA). Tissue sections (6–8 sections/animal/antisense probe, 3–4 sections/animal/sense probe) were processed for in situ hybridization as described previously [18], with the modification that slides were hybridized at 55°C. Sense and antisense riboprobes for the PPAR isotypes were synthesized using rat cDNAs (generously provided by Dr. Walter Wahli, Université de Lausanne, Lausanne, Switzerland), Ambion's MAXISCRIPT kit (Ambion, Inc., Austin, TX), and [-35S]uridine-5'-trisphosphate (10 µCi/ml; ICN Biomedicals, Inc., Irving, CA).
RNase Protection Assay
Total RNA was isolated from individual cycling and pseudopregnant rat ovaries using Trizol (Gibco BRL, Rockville, MD) and quantified by spectrophotometry. Rat cDNAs for the PPARs , , and and mouse cDNA for ribosomal protein L32 (kindly provided by Dr. O.-K. Park-Sarge, University of Kentucky, Lexington, KY) were linearized with the appropriate restriction enzymes. Antisense riboprobes were transcribed using the MAXISCRIPT kit and [-32P]uridine-5'-trisphosphate (10 Ci/ml; New England Nuclear, Boston, MA).
The RNase protection assays were carried out as described previously [23, 24]. Briefly, samples of total RNA were hybridized for 15–18 h at 50°C with excess radiolabeled antisense riboprobe (n = 3 individual samples/time point). Loading variation between samples was standardized by including L32 riboprobe in all hybridization reactions. Quantification of band intensity in the gels was determined using a PhosphorImager (Molecular Dynamics, Sunnyvale, CA). The band intensity of mRNAs corresponding to the PPAR isotypes were normalized to the corresponding band for L32 per sample.
Statistical Analysis
Levels of mRNA for the PPARs determined by RNase protection assay in ovarian tissue collected during pseudopregnancy and the estrous cycle were analyzed by one-way ANOVA. Post-hoc comparisons were made using the Tukey honestly significant difference test. Differences were considered to be significant at P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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was found primarily in the theca and stroma, with low levels of expression in the CL (Fig. 1a). PPAR mRNA was expressed throughout the ovary, with labeling mainly seen in the theca, stroma, and luteal tissue (Fig. 1d). Messenger RNA for PPAR was localized primarily to granulosa cells of developing follicles. Expression of PPAR mRNA in luteal tissue was low to undetectable (Fig. 1g).
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To determine whether the levels of mRNA for the PPAR isotypes changed during pseudopregnancy, RNase protection assays were used. As shown in Figure 2, the levels of mRNA for PPARs and remained steady throughout pseudopregnancy. In contrast, the levels of mRNA for PPAR decreased between Days 1 and 4 (P < 0.05) and then remained steady for the rest of pseudopregnancy.
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The investigation of mRNAs for the PPAR isotypes in ovarian tissue from naturally cycling animals showed that the cellular expression pattern of mRNA for PPAR was different from the pattern seen in ovarian tissue from pseudopregnant animals (Fig. 3). In both animal models, PPAR mRNA was abundant in granulosa cells of developing follicles. However, the CL of the pseudopregnant animals contained little to no labeling for PPAR mRNA, but mRNA for PPAR was present in luteal tissue from cycling animals.
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In the cycling animal, morphological differences enable one to differentiate between luteal tissue from previous estrous cycles and that formed during the current estrous cycle (Fig. 4b). As described by Simpson et al. [25], newly forming CL are comprised mainly of large luteal cells characterized by large, darkly staining nuclei, whereas CL from previous cycles contain more stromal cells and connective tissue components. The relative expression of PPAR mRNA appeared to be low in the newly forming CL and higher in luteal tissue from previous cycles (Fig. 4a). In addition, mRNA for PPAR was identified in the theca externa of developing follicles in ovarian tissue from cycling rats (Figs. 3 and 4c).
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The pattern of cellular expression for PPAR mRNA in ovarian tissue from cycling animals did not differ from what was observed during pseudopregnancy. Messenger RNA for PPAR was seen throughout the ovary, primarily in theca, stroma, and luteal tissues (Fig. 5a). The expression of mRNA for PPAR was also very similar to that seen during pseudopregnancy, with low to undetectable levels of expression in the CL on proestrus, estrus, and diestrus (Fig. 5d). On metestrus, one animal had newly forming CL that labeled for PPAR mRNA, whereas ovaries from the other two animals collected on this day of the cycle had low to undetectable levels of mRNA in all luteal tissue (Fig. 6).
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RNase protection assays were used to measure the levels of mRNA corresponding to the PPAR isotypes in cycling rat ovaries. Levels of mRNA for PPARs and did not change during the estrous cycle (Fig. 7). However, the levels of mRNA for PPAR were lower on the day of estrus compared to levels on the days of proestrus and metestrus (P < 0.05). On diestrus, PPAR mRNA levels were not different from those on the other days of the cycle.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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in luteal tissue differs between naturally cycling and gonadotropin-primed rats. The expression of PPAR mRNA was low to undetectable in luteal tissue from immature, pseudopregnant animals. However, the expression of mRNA for PPAR was relatively high in luteal tissue from cycling animals, particularly in luteal tissue from previous cycles. These data suggest that a difference in the endocrine environment exists between naturally cycling and gonadotropin-treated rats, resulting in the observed differential pattern of PPAR mRNA expression. One possibility is that the expression of mRNA for PPAR in luteal tissue is regulated by prolactin, because the immature, gonadotropin-treated animals would not have a prolactin surge coincident with the LH surge, as occurs in the adult, cycling animal [26]. The hypothesis that prolactin regulates the expression of PPAR mRNA in luteal tissue is currently under investigation.
Previous work has shown a relationship between PPAR and steroidogenic enzymes in ovarian cells [27]. Activation of PPAR inhibited the activity and expression of aromatase in cultured human granulosa-lutein cells [27]. It is possible that PPAR may also be correlated to the expression of steroidogenic enzymes in luteal tissue. For example, expression of PPAR may be related to the expression of 20-hydroxysteroid dehydrogenase (20-HSD), an enzyme responsible for converting progesterone into the inactive metabolite 20-dihydroprogesterone. This enzyme has been hypothesized to play a role in the decrease in progesterone seen during luteolysis in mice and rats. In a study conducted by Yoshida et al. [28], adult, cycling animals were rendered pseudopregnant by cervical stimulation. Examination of luteal tissue collected from these animals demonstrated that the expression of mRNA for 20-HSD was low in newly forming CL on Days 6 and 10 and increased on Days 16 and 18 of pseudopregnancy. In the present study, expression of mRNA for PPAR in cycling animals was low in newly forming CL, whereas expression of mRNA for PPAR was relatively high in luteal tissue from previous cycles. These findings suggest that PPAR may play a role in luteal regression or in the change in progesterone production as CL age in adult, cycling rats.
Further evidence supporting a role for PPAR in modifying progesterone production comes from the expression of PPAR mRNA in granulosa cells versus luteal tissue. The high expression of PPAR mRNA in granulosa cells of developing follicles reported in the present study and previously [18] correlates with follicular estradiol production [29–31]. The reduction of mRNA for PPAR after the LH surge [18] and the low level of expression in newly forming CL exhibits an inverse relationship to the increase in progesterone production as CL develop. Progesterone production decreases as the CL ages, coincident with an increase in the expression of mRNA for PPAR seen in CL from cycling rats. Although activation of PPAR in cultured bovine luteal cells resulted in increased progesterone secretion [21], species differences and/or differences in the cellular response related to the stage of luteal differentiation may exist. Current experiments in our laboratory are examining the ability of PPAR to influence steroidogenesis in luteal tissue.
The localization and pattern of expression of mRNA for PPAR did not change during pseudopregnancy. However, the levels of mRNA for PPAR in ovarian tissue from pseudopregnant rats decreased between Days 1 and 4. This decrease most likely reflects a reduction in the proportion of ovarian RNA from follicular tissue as the CL develop, not a true reduction in the amount of mRNA for PPAR. In other words, as the CL develop, the amount of total ovarian RNA derived from follicular tissue, which contains PPAR mRNA, is diluted.
Messenger RNA corresponding to PPAR was located mainly in the theca and stroma of ovaries from both pseudopregnant and cycling animals. These findings agree with those of previous work illustrating the expression of PPAR mRNA in the theca and stroma of ovarian tissue collected from immature, gonadotropin-primed animals during follicular development and the periovulatory period [18]. The finding that the levels of mRNA for PPAR were lower on the day of estrus compared to the days of proestrus and metestrus may reflect the loss of theca and stroma as the follicular tissue remaining after ovulation differentiates into luteal tissue. Because PPAR has been shown to be an important player in lipid metabolism [15, 32], this factor may be involved in the metabolic processes and steroidogenesis occurring in these ovarian tissue compartments.
The expression of mRNA for PPAR throughout ovarian tissue observed in the current study, and previously [18], is in agreement with its ubiquitous distribution in other tissues [33, 34]. Although assigning specific functions to this PPAR isotype has been difficult, the use of selective ligands and knock-out technologies has shown that PPAR is involved in reverse cholesterol transport [35], development, nerve myelination, and lipid metabolism [36]. Because it is expressed throughout ovarian tissue at a constant level during the ovarian cycle, PPAR may be a factor involved in basal ovarian function.
PPARs have been shown to play a role in angiogenesis [5]. Besides regulating blood vessel development, PPARs could also influence the ovarian vasculature by their ability to regulate endothelin-1 (ET-1) and nitric oxide synthase. Endothelin-1 is a potent vasoconstrictor, and recent studies have shown that it is also an important player in ovarian function [37, 38]. Nitric oxide synthase synthesizes nitric oxide, a vasodilator, from arginine. Nitric oxide is involved not only in luteolysis [39] but also in ovarian cyclicity [40], ovulation [40–42], oocyte maturation [41], and follicular development [43, 44]. Activation of PPAR decreases the secretion of ET-1 from endothelial cells [45] and inhibits the expression of nitric oxide synthase in macrophages [10] and vascular smooth muscle cells [46]. Hence, PPARs could affect the ovarian vasculature by regulating angiogenesis as well as impacting ET-1 and nitric oxide production.
In conclusion, the ability of PPARs to influence tissue remodeling, angiogenesis, lipid metabolism, and steroidogenesis, coupled with the current findings that these factors are expressed in various ovarian tissue compartments, indicate that PPARs may be important players in follicular and luteal function. The difference in the pattern of expression of mRNA for PPAR between the naturally cycling adult and the immature, gonadotropin-treated animal suggests that expression of the PPARs may be influenced by the hormonal environment. Future studies are needed to gain a better understanding of how the PPARs are regulated in the ovary and to identify the cellular processes in which these factors are involved during the ovarian cycle.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Supported by grants NIH:HD23195 (T.E.C.) and NCRR P20 15592 (T.E.C., C.M.K.). 
2 Correspondence and current address: Carolyn Komar, Department of Animal Science, 2356F Kildee Hall, Iowa State University, Ames, IA 50011-3150. FAX: 515 294 4471; ckomar{at}iastate.edu
Accepted: December 13, 2001.
Received: October 1, 2001.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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ligands are potent inhibitors of angiogenesis in vitro and in vivo. J Biol Chem 1999; 274:9116-9121 or reduces secretion of matrix metalloproteinase 9 but not interleukin 8 from human monocytic THP-1 cells. Biochem Biophys Res Commun 2000; 267:345-349[CrossRef][Medline]. Am J Pathol 1998; 153:17-23 is a negative regulator of macrophage activation. Science 1998; 391:79-82 activation in human endothelial cells increases plasminogen activator inhibitor type-1 expression. Arterioscler Thromb Vasc Biol 1999; 19:546-551 but not by PPAR activators. Nature 1998; 393:790-793[CrossRef][Medline] (12,14)-prostaglandin J2 (PGJ2) a peroxisome proliferator activating receptor (PPAR ) ligand on human ovarian steroidogenesis. Endocrinology 1999; (suppl 1):491 (abstract P3-247) in lutein cells. J Endocrinol 1998; 159:429-439[Abstract] protein in bovine large lutein cells. J Reprod Fertil 2000; 118:153-161[Abstract]-hydroxylase, in bovine preovulatory follicles. Biol Reprod 2001; 64:1797-1805 activators regulate genes governing lipoprotein metabolism, vascular inflammation and atherosclerosis. Curr Opin Lipidol 1999; 10:245-257[CrossRef][Medline], -ß and - in the adult rat. Endocrinology 1996; 137:354-366[Abstract]). Mol Cell Biol 2000; 20:5119-5128 in corpus luteum function. Rev Reprod 2000; 5:1-5[Abstract] on vascular endothelial function. Biochem Biophys Res Commun 1999; 254:757-763[CrossRef][Medline] ligands inhibit nitric oxide synthesis in vascular smooth muscle cells. Hypertension 2000; 35:1232-1236This article has been cited by other articles:
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