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Department of Obstetrics and Gynecology (C.M.K., T.E.C.), Chandler Medical Center, University of Kentucky, Lexington, Kentucky 40536-0298; and Institut de Biologie Animale (O.B., W.W.), Université de Lausanne, CH-1050 Lausanne, Switzerland
Address all correspondence and requests for reprints to Carolyn M. Komar, Department of Obstetrics and Gynecology, Chandler Medical Center, 800 Rose Street, Room MS 331, University of Kentucky, Lexington, Kentucky 40536-0298. E-mail: ckomar{at}uky.edu
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, 
, and 
) were localized
by in situ hybridization. Changes in the levels of mRNA
for the PPARs were determined by ribonuclease protection
assays.
PPAR mRNA was localized primarily to granulosa cells, and levels of
expression did not change during follicular development. Four hours
post-hCG, levels of mRNA for PPAR decreased (P
< 0.05) but not uniformly in all follicles. At 24 h post-hCG,
levels of PPAR mRNA were reduced 64%, but some follicles maintained
high expression. In contrast, mRNAs for PPAR and were located
primarily in theca and stroma, and their levels did not change during
the intervals studied. To investigate the physiologic significance of
PPAR in the ovary, granulosa cells from PMSG-primed rats were
cultured for 48 h with prostaglandin J2
(PGJ2) and ciglitazone, PPAR activators. Both compounds
increased progesterone and E2 secretion (P <
0.05).
These data suggest that PPAR is involved in follicular development,
has a negative influence on the luteinization of granulosa cells,
and/or regulates the periovulatory shift in steroid production. The
more general and steady expression of PPARs and indicate that
they may play a role in basal ovarian function.
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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(NR1C1),
PPAR [ß, NUC-1, fatty acid-activated receptor (FAAR),
NR1C2], and PPAR (NR1C3). Each PPAR is transcribed from an
individual gene. The gene encoding PPAR gives rise to different
species of mRNA resulting from alternate splicing and the use of
different promoters (2, 3, 4). PPARs are activated by various factors such as fibrates, herbicides, industrial plasticizers, insulin, fatty acids including arachidonic acid and its eicosanoid metabolites, nonsteroidal antiinflammatory drugs, and thiazolidinediones (insulin-sensitizing drugs; see Ref. 1 for a review). The activity of PPARs can also be modulated by phosphorylation (1). PPARs heterodimerize with 9, cis-retinoic acid receptors and bind to PPAR response elements (PPREs) present in the promoter region of target genes, thereby regulating transcription.
Since their initial discovery, PPARs have been found to be involved in
a variety of cellular functions, some which could directly influence
ovarian physiology. PPARs can bind to estrogen response elements
(5, 6), inactively occupying the estrogen response element
and preventing access to the ER (5). PPARs also regulate
the activity and expression of aromatase, an enzyme involved in the
biosynthesis of estrogen. In human breast adipose tissue and
granulosa-lutein cells, aromatase activity is inhibited by the
activation of PPAR (7, 8). The activation of PPAR
has also been shown to inhibit progesterone production by cultured
porcine (9) and human (10) granulosa cells.
This effect of PPAR on progesterone production may result from its
ability to decrease the activity of 3ß-hydroxysteroid dehydrogenase
(9).
PPARs are capable of modulating the expression and activity of proteolytic enzymes which could affect tissue remodeling and angiogenesis that occurs during ovarian follicular development, ovulation, and luteal formation. For example, PPARs regulate gelatinase B (11, 12, 13) and plasminogen activator (14), proteases known to be stimulated during follicular growth and ovulation. The promoter for another protease, stromelysin, contains a PPRE suggesting that its expression is regulated by PPARs (15).
Other factors that have been shown to be involved in ovarian
function, such as endothelin-1 (16), nitric oxide synthase
(17, 18, 19), and cyclooxygenase-2 (COX-2; 20, 21), are also regulated by PPARs. PPAR decreases the
secretion of endothelin-1 from endothelial cells (22) and
inhibits the expression of nitric oxide synthase in macrophages
(12) and vascular smooth muscle cells (23). A
PPRE has been identified upstream of the COX-2 transcriptional start
site (24), and activation of PPARs modulates the
expression of COX-2 (24, 25, 26). Taken together, these
findings indicate that there are a number of ways PPARs could regulate
ovarian function.
All three PPAR isotypes have been identified in the rat ovary
(27), and mRNA for PPAR has been found in the cow
(2, 28) and human ovary (29). However, it is
not completely understood where the PPARs are localized and produced in
the ovary, how expression of the PPARs changes during the ovarian
cycle, how they may be regulated, or their role in ovarian function.
The studies described herein were designed to investigate the
localization and expression of PPARs in the ovary and their potential
to regulate ovarian steroidogenesis.
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Materials and Methods |
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12, 14-PGJ2
from Cayman Chemical (Ann Arbor, MI). RNase A was obtained from Amresco
Inc. (Solon, OH). TRIZOL was supplied by Life Technologies, Inc. (Rockville, MD). Optimal cutting temperature (OCT)
embedding compound came from VWR Scientific (Atlanta, GA).
Animals
All animal procedures were approved by the University of
Kentucky Animal Care and Use Committee. Female, Sprague Dawley rats
(Harlan Sprague Dawley, Inc., Indianapolis, IN) were
injected with 10 IU PMSG sc on d 23 of age to initiate follicular
development. At 0 (no PMSG), 6, 12, 24, and 48 h post-PMSG,
ovaries were collected from a subset of these animals (n =
3–4/time point) to examine the expression of PPARs during follicular
growth. The remaining animals received 10 IU hCG sc 48 h post-PMSG
to stimulate ovulation and luteal development. Animals were killed 0
(no hCG), 4, 8, 12, and 24 h after treatment with hCG (n =
3/time point) and their ovaries collected. Ovaries were frozen in
liquid nitrogen and stored at -70 C until RNA isolation, or placed in
OCT and stored at -70 C until sectioned for in situ
hybridization.
Tissues were collected from a second group of animals during follicular
development and the periovulatory period to localize mRNA for PPAR
and apoptotic cells in adjacent tissue sections. Female Sprague Dawley
rats were injected with 10 IU PMSG sc on d 23 of age. Ovaries were
collected from a subset of these animals 0 (no PMSG) and 24 h
post-PMSG (n = 2/time point). The remaining animals received 10 IU
of hCG 48 h post-PMSG. Animals in this latter group were killed 0
(no hCG), 4, and 24 h after treatment with hCG (n = 3/time
point) and their ovaries collected. All ovaries were placed in OCT and
snap frozen. Tissues were stored at -70 C until sectioned for in
situ hybridization and the detection of apoptotic cells.
To investigate the role of PPAR in steroidogenesis in
vitro, granulosa cells were collected from PMSG-primed immature
rats. On d 23 of age, female rats received 10 IU PMSG sc; 48 h
later, granulosa cells were collected as described previously
(30).
In situ hybridization
Ovaries collected from the first group of animals during
follicular development and the periovulatory period were sectioned at 8
µm and mounted on ProbeOn Plus slides (Fisher Scientific, Pittsburgh, PA). Tissue sections were fixed in 4%
paraformaldehyde, washed twice in PBS (pH 7.6), followed by two washes
in 0.75% glycine. Slides were rinsed twice in PBS and then washed in
triethanolamine buffer with 0.25% acetic anhydride. Following another
PBS wash, sections were dehydrated via passage through a series of
alcohol washes.
Sense and antisense riboprobes for PPAR, , and were
synthesized using a MAXISCRIPT kit (Ambion, Inc. Austin,
TX) and [-33P]UTP (10 µmCi/ml; NEN Life Science Products, Boston, MA). Tissues were hybridized with
radiolabeled probe (1 x 106 cpm) in 50 µl
hybridization buffer (100 µg/ml salmon sperm DNA, 250 µg/ml total
yeast RNA, 250 µg/ml yeast transfer RNA, 20 mM Tris, 1
mM EDTA, 300 mM NaCl, 50% formamide, 10%
dextran sulfate, and 1x Denhart’s) at 60 C for 15–18 h.
Nonspecifically bound RNA transcripts were removed by washing tissue
sections in 2x standard saline citrate (SSC) and treating them with
RNase A (0.1 mg/ml) for 30 min at 45 C. Tissue sections were then
washed in 0.2x SSC, followed by a 1 h wash in 0.1x SSC at 55 C.
After air drying, slides were dipped in Kodak NTB2
emulsion and exposed at 4 C for 4–6 (PPAR) or 6–8 (PPARs and
) wk. Slides were brought to room temperature, developed, and
counterstained with hematoxylin.
Ovarian tissue collected from the second group of animals to determine
whether expression of PPAR mRNA was correlated with the presence of
apoptotic cells was serially sectioned (10 µm) and placed on ProbeOn
Plus slides. Tissues were processed as described above with the
following modification. mRNA corresponding to PPAR was localized
using 35S-labeled riboprobes (10 µmCi/ml;
ICN Biomedicals, Inc., Irvine, CA).
RNase protection assay
Total RNA was isolated from ovaries collected during follicular
development and the periovulatory period using TRIZOL reagent and
quantified by spectrophotometry. Plasmids containing rat cDNAs for
PPARs , , and , and mouse cDNA for ribosomal protein L32 (the
latter 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 Ambion, Inc.’s MAXISCRIPT kit and
[-32P]UTP (10 mCi/ml; NEN Life Science Products).
RNase protection assays were carried out as described previously (31, 32) using RNA isolated from 3–4 animals/time point (each sample assayed once). Briefly, samples of total RNA (1–6 µg) were hybridized for 15–18 h at 50 C with excess radiolabeled antisense riboprobe. Loading variation between samples was standardized by including L32 riboprobe in all hybridization reactions. Protected RNA fragments were analyzed by electrophoresis through a 5% acrylamide/8 M urea gel. Quantification of band intensity in the gels was determined using a phosphor-imager (Molecular Dynamics, Inc., Sunnyvale, CA). The band intensity of mRNA for each PPAR isotype was normalized to the corresponding band for L32 per sample.
Detection of apoptotic cells
Apoptotic cells were identified in serial sections of frozen
ovarian tissue (10 µm) using ApoAlert (CLONTECH Laboratories, Inc., Palo Alto, CA). The manufacturer’s instructions were
followed with a few modifications. Tissue sections were allowed to come
to room temperature before proceeding with fixation and were not
treated with proteinase K. The tissue sections were not stained with
propidium iodide, but rather mounted with Vectashield mounting medium
(Vector Laboratories, Inc., Burlingame, CA) containing
propidium iodide. Three slides/animal with three tissue sections/slide
were processed to detect apoptotic cells.
Cell culture
Granulosa cells were pooled and 1 x
106 cells cultured per ml of DMEM-Ham’s F-12
containing 1% BSA, 0.01% pyruvic acid, 0.22% bicarbonate, and ITS
(insulin, transferrin, selenium) at 37 C in an atmosphere of 95%
O2:5% CO2. Depending on
the number of cells collected, cultured cells were treated in duplicate
or triplicate as follows (n = 5 independent experiments): control,
ciglitazone (25 and 50 µM), or PGJ2
(10 and 25 µM). Treatments were added to the cells at the
time of plating. Forty-eight hours after the initiation of culture,
media were collected to measure the concentrations of E2 and
progesterone by RIA. Cell viability was assessed at the end of culture
by extracting and analyzing total cellular RNA in formaldehyde/agarose
gel.
RIA
Concentrations of E2 and progesterone were determined in culture
media by using Coat-A-Count tubes (Diagnostic Products
Co., Los Angeles, CA), which are direct, solid phase
125I RIA kits. These kits are designed to
determine concentrations of steroids in serum, and our laboratory has
verified their ability to measure levels of steroids in conditioned
culture media (data not shown). Assay sensitivity is 0.03 ng/ml for
progesterone and 50 pg/ml for E2. The inter and intraassay coefficients
of variation were 7.9% and 5.9% for progesterone, and 4.8% and 3.6%
for E2, respectively.
Statistical analysis
Levels of mRNA for the PPAR isotypes in ovarian tissue collected
during follicular development and the periovulatory period were
analyzed by one-way ANOVA. Posthoc comparisons were made using Tukey’s
HSD. Concentrations of progesterone and E2 in conditioned culture media
were subjected to square root transformation before being analyzed by
one-way ANOVA. Posthoc comparisons were made using the
Student-Newman-Kuels test. A P < 0.05 was considered
significant.
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Results |
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and 2,
mRNA for PPAR is limited primarily to granulosa cells throughout
follicular development and the periovulatory period. A few follicles in
some of the animals contained theca extema that labeled weakly for
PPAR mRNA, but no consistent pattern was observed (date not shown).
During follicular development (Fig. 1), granulosa cells in the majority
of follicles expressed high levels of mRNA for PPAR at all time
points studied before and after PMSG administration. However, the
granulosa cells in some follicles were less intensely labeled than
others.
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in
granulosa cells decreased after the administration of hCG (Fig. 2
). The
expression pattern at 0 h (48 h post-PMSG, no hCG) was very
similar to the pattern seen during follicular development, with
granulosa cells in most follicles labeled intensely for PPAR mRNA
(Fig. 2, a and g). By 4 h post-hCG, granulosa cells in a number of
large follicles lost expression of mRNA for PPAR. Interestingly, the
expression of mRNA for PPAR did not decrease uniformly in all
follicles; rather some follicles lost expression, whereas others
maintained high expression (Fig. 2, b and h). At 24 h post-hCG,
PPAR expression in luteinizing granulosa cells was undetectable, but
labeling remained high in follicles (Fig. 2, c and i).
In contrast to the expression of mRNA for PPAR, mRNA for both
PPAR and was expressed primarily in theca and stroma tissues.
The expression patterns of PPAR and did not change during
follicular development or the periovulatory period; therefore, only the
representative time point of 48 h post-PMSG is shown (Fig. 3 and 4, respectively). As can be
seen in Fig. 3, the expression of mRNA
for PPAR was low, not much higher than background. Labeling of mRNA
for PPAR, although seen throughout the tissue, was more intense in
theca and interstitial tissues than in granulosa cells (Fig. 4).
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and remained steady
throughout follicular development and the
periovulatory period (Figs. 5 and 6, respectively). The level of mRNA for
PPAR also remained steady during follicular development (Fig. 5).
However, by 4 h post-hCG, there was a significant decrease in the
levels of mRNA for PPAR (P < 0.05; Fig. 6). The
level of mRNA for PPAR remained low and at 24 h post-hCG levels
were reduced by 64% compared with those at 0 h (P <0.01).
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mRNA than others during follicular development, and the finding
that some ovarian structures maintained expression for PPAR mRNA
24 h post-hCG, we hypothesized that the expression of mRNA for
PPAR may be related to follicular health. To test this idea, ovarian
tissue was collected from a second group of animals during follicular
development and the periovulatory period. mRNA for PPAR and the
presence of apoptotic cells were identified in serial tissue
sections.
As seen in tissue collected from the first group (Fig. 1), the majority
of follicles in tissue collected from animals in the second group 0 and
24 h post-PMSG contained granulosa cells expressing mRNA for
PPAR (data not shown). However, few follicles contained apoptotic
cells in ovaries from the second group of animals, and there was no
correlation between the labeling intensity of mRNA for PPAR and the
presence of apoptotic cells (data not shown). The same trend was
observed during the periovulatory period. Tissue collected 4 h
post-hCG are presented in Fig. 7 and
represent the findings in periovulatory tissues. There was no
relationship between the relative level of PPAR mRNA expression and
the presence or absence of apoptotic cells.
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expression after the administration
of hCG led us to hypothesize that this PPAR isotype may be involved in
the periovulatory shift in steroid production from E2 to progesterone.
To begin testing this hypothesis, granulosa cells were cultured in the
presence and absence of PGJ2 (10 and 25
µM; an endogenous activator of PPAR; 33)
and ciglitazone (25 and 50 µM; a member of the
thiazolidinedione drug family that specifically activates PPAR).
The addition of PGJ2 to cultured rat granulosa
cells resulted in a dose-dependent increase in progesterone secretion
(Fig. 8A). There was a 3-fold increase in
basal progesterone secretion when cells were treated with 10
µM PGJ2. In the presence of 25
µM PGJ2, progesterone secretion
increased 14-fold (P < 0.05). However, only the high
dose of ciglitazone (50 µM) resulted in a
significant, 3-fold increase in progesterone secretion
(P < 0.05). Basal E2 secretion was stimulated by the
high dose of each PPAR agonist (Fig. 8B). Treatment with 25
µM PGJ2 and both 25 and
50 µM ciglitazone resulted in a significant
30% increase in E2 secretion (P < 0.05).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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has been reported to be expressed in ovarian tissue from humans
(29) and cattle (2, 28), and all three PPAR
isotypes have been identified in ovarian tissue taken from an adult rat
(27). The data presented here extend those initial
findings by characterizing the expression of each PPAR throughout
follicular development and the periovulatory period in the rat. Our
findings show that the expression of PPAR is restricted primarily to
granulosa cells of developing follicles, whereas mRNAs for PPAR and
were localized to the theca and stroma. The high expression of mRNA
for PPAR both before and after the administration of PMSG suggests
that this isotype may play a role in follicular development. Of
particular interest was the observation that levels of mRNA for PPAR
decreased in response to treatment with hCG, which mimics the LH surge.
This finding indicates that PPAR may be inhibitory to the transition
of granulosa cells into luteal cells. In contrast to the expression of
PPAR, the steady expression of mRNAs for PPAR and throughout
follicular development and the periovulatory period, coupled with their
more general distribution throughout ovarian tissue, suggests that
these two PPARs may be important factors in maintaining basal ovarian
function.
Although the granulosa cells in the majority of follicles were labeled
for PPAR mRNA during follicular development, some follicles were
more intensely labeled than others. A difference in follicular PPAR
labeling was also seen during the periovulatory period. The decrease in
granulosa cell PPAR expression in response to gonadotropin treatment
did not occur uniformly in all follicles; rather, some follicles lost
expression whereas others maintained high levels of expression, even
24 h post-hCG. This finding led us to hypothesize that PPAR
expression may relate to follicular health. However, the lack of
association between the expression of mRNA for PPAR and the presence
of apoptotic cells in follicles during follicular development and the
periovulatory period indicate that PPAR expression is not correlated
with cellular health. Another possible explanation for the difference
in PPAR expression between granulosa cells in different follicles is
that the follicles with lower expression are not as far along the
developmental pathway as others in their cohort. This latter hypothesis
is currently under investigation.
In our culture system, activation of PPAR in rat granulosa cells
resulted in an increase in both progesterone and E2 secretion. These
findings differ from previous reports of PPAR inhibiting
steroidogenesis in cultured human and porcine granulosa cells (7, 9, 10). In those studies, the activation of PPAR inhibited
progesterone (9, 10) and E2 production (7).
The apparent dichotomy between our findings with rat ovarian cells and
those in human and pig ovarian cells could result from the stage of
cellular differentiation and/or species variation. Another possible
explanation may be the fact that in the current study the steroid
content of conditioned media was measured 48 h after the
initiation of culture. Because the granulosa cells were collected at a
time when they express high levels of mRNA for PPAR (Fig. 2; 48
h post-PMSG), the activation of PPAR in these cells in
vitro could initially have had a stimulatory effect on E2
secretion. Because granulosa cells spontaneously luteinize when placed
in culture, treatment with PPAR agonists could have augmented
progesterone production late in the culture period. Data to support
this hypothesis comes from a study by Löhrke, et al.
(1998). Mid-cycle bovine luteal cells cultured in the presence of
PGJ2 or ciglitazone increased progesterone
production (28). In addition, putative inhibition of
PPAR expression by treating bovine luteal cells with
aurintricarboxylic acid, resulted in decreased progesterone secretion
(28).
There are a number of studies demonstrating that
troglitazone, a member of the thiazolidinedione drug
family that is capable of activating PPAR, is an effective treatment
for some women with polycystic ovary syndrome (PCOS). PCOS is a leading
cause of infertility in premenopausal women and is characterized by
hyperandrogenism, anovulation, and frequently insulin resistance.
Treatment of these women with troglitazone reduced
androgen levels, improves hyperinsulinemia, and in some women restored
ovulation (35, 36, 37, 38). These findings suggest that PPAR
activity can influence thecal androgen production. Such an activity of
PPAR would be interesting because this factor has already been shown
to influence granulosa cell steroid production, and we have localized
mRNA for PPAR primarily to granulosa cells. Whether our data reflect
a species difference between the rat and human, or a difference in
expression between gonadotropin-treated and naturally cycling animals
is the focus of current and future studies. Results from such studies
will yield important information concerning not only how PPARs may
influence ovarian function, but also how drugs in clinical use
(thiazolidinediones and fibrates) impact ovarian physiology.
In summary, the data presented here show that PPAR is expressed in
the ovary, primarily in the granulosa cells of developing follicles.
Following the LH surge, levels of mRNA for PPAR decline, indicating
that this factor may play a role in follicular development, and/or be
inhibitory to the transition of granulosa cells into luteal cells. We
have also shown that activation of PPAR in cultured granulosa cells
can stimulate the secretion of both progesterone and E2. Further
investigation of this PPAR isotype in ovarian function will lead to a
better understanding of follicular growth and differentiation,
steroidogenesis, as well as the regulation of ovarian gene expression
by the gonadotropin surge.
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Acknowledgments |
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Footnotes |
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Abbreviations: COX-2, cyclooxygenase-2; FAAR, fatty acid-activated receptor; hCG, human CG; NR1C1–NR1C, a family of nuclear hormone receptors belonging to the steroid receptor superfamily; PGJ2, prostaglandin J2; PPRE, PPAR response elements; PCOS, polycystic ovary syndrome.
Received March 12, 2001.
Accepted for publication June 14, 2001.
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References |
|---|
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1 and 2: genetic
mapping and differential expression of the two isoforms. Biochem
Biophys Res Commun 239:857–861[CrossRef][Medline]
3 mRNA: a distinct PPAR subtype
transcribed from an independent promoter. FEBS Lett 438:55–60[CrossRef][Medline]
(mPPAR) gene: alternative promoter use and different splicing yield
two mPPAR isoforms. Proc Natl Acad Sci USA 92:7921–7925 (12,14)-prostaglandin J2 (PGJ2) a peroxisome
proliferator activating receptor (PPAR ) ligand on human ovarian
steroidogenesis. Program of the 81st Annual
Meeting of The Endocrine Society, San Diego, CA, P3-247 (Abstract)
activators inhibit gene
expression and migration in human vascular smooth muscle cells. Circul
Res 83:1097–1103 is a negative
regulator of macrophage activation. Science 391:79–82
. Am J Pathol 153:17–23 ligands are potent
inhibitors of angiogenesis in vitro and in vivo.
J Biol Chem 274:9116–9121 in corpus luteum function. Rev
Reprod 5:1–5[Abstract]
on vascular
endothelial function. Biochem Biophys Res Commun 254:757–763[CrossRef][Medline]
ligands inhibit nitric
oxide synthesis in vascular smooth muscle cells. Hypertension 35:1232–1236 but not by
PPAR activators. Nature 393:790–793[CrossRef][Medline]
, -ß and - in the adult
rat. Endocrinology 137:354–366[Abstract]
in lutein cells. J Endocrinol 159:429–439[Abstract]
is activated by inducers
of adipogenesis, including thiazolidinedione drugs. Eur J Biochem 239:1–7[Medline]
-hydroxylase, in bovine preovulatory follicles. Biol Reprod 64:1797–180512,
14-prostaglandin J2 is a ligand for
the adipocyte determination factor PPAR. Cell 83:803–812[CrossRef][Medline]
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K. A. Brown, D. Boerboom, N. Bouchard, M. Dore, J. G. Lussier, and J. Sirois Human Chorionic Gonadotropin-Dependent Regulation of 17{beta}-Hydroxysteroid Dehydrogenase Type 4 in Preovulatory Follicles and Its Potential Role in Follicular Luteinization Endocrinology, April 1, 2004; 145(4): 1906 - 1915. [Abstract] [Full Text] [PDF]
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P. Froment, S. Fabre, J. Dupont, C. Pisselet, D. Chesneau, B. Staels, and P. Monget Expression and Functional Role of Peroxisome Proliferator-Activated Receptor-{gamma} in Ovarian Folliculogenesis in the Sheep Biol Reprod, November 1, 2003; 69(5): 1665 - 1674. [Abstract] [Full Text] [PDF]
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C. M. Komar and T. E. Curry Jr Inverse Relationship Between the Expression of Messenger Ribonucleic Acid for Peroxisome Proliferator-Activated Receptor {gamma} and P450 Side Chain Cleavage in the Rat Ovary Biol Reprod, August 1, 2003; 69(2): 549 - 555. [Abstract] [Full Text] [PDF]
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 K. Toda, T. Okada, C. Miyaura, and T. Saibara Fenofibrate, a ligand for PPAR{alpha}, inhibits aromatase cytochrome P450 expression in the ovary of mouse J. Lipid Res., February 1, 2003; 44(2): 265 - 270. [Abstract] [Full Text] [PDF]
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in tissue
sections from an ovary collected 48 h post-PMSG. a, darkfield
image (magnification x40) of a tissue section hybridized with an
antisense riboprobe as described in Materials and
Methods; b, corresponding brightfield image; c, darkfield image
of a tissue section hybridized with a sense riboprobe (magnification
x40).
