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BOR - Papers in Press, published online ahead of print September 28, 2005.
Biol Reprod 2005, 10.1095/biolreprod.105.043729
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BIOLOGY OF REPRODUCTION 74, 153–160 (2006)
DOI: 10.1095/biolreprod.105.043729
© 2006 by the Society for the Study of Reproduction, Inc.

Research Article

Troglitazone Regulates Peroxisome Proliferator-Activated Receptors and Inducible Nitric Oxide Synthase in Murine Ovarian Macrophages1

Cadence E Minge, Natalie K. Ryan, Kylie H. Van Der Hoek, Rebecca L. Robker, and Robert J. Norman 2

Research Centre for Reproductive Health, Department of Obstetrics and Gynaecology, University of Adelaide, Queen Elizabeth Hospital, Woodville, South Australia 5011, Australia

ABSTRACT

Peroxisome proliferator-activated receptor-gamma (PPARG) and PPAR-alpha (PPARA) control metabolic processes in many cell types and act as anti-inflammatory regulators in macrophages. PPAR-activating ligands include thiazolidinediones (TZDs), such as troglitazone, once frequently used to treat insulin resistance as well as symptoms of polycystic ovary syndrome (PCOS). Since macrophages within the ovary mediate optimal follicle development, TZD actions to improve PCOS symptoms are likely to be partly mediated through these specifically localized immune cells. In mouse ovary, PPARG protein was expressed in granulosa cells and in isolated cells localized to theca, stroma, and corpora lutea, consistent with EMR1+ macrophages. Isolation of immune cells (EMR1+ or H2+) showed that Pparg and Ppara were expressed in ovarian macrophages at much higher levels than in peritoneal macrophages. Ovulatory human chorionic gonadotropin downregulated expression of Pparg and Ppara in EMR1+ ovarian macrophages, but no hormonal responsiveness was observed in H2+ cells. Downstream anti-inflammatory effects of PPARG activation were analyzed by in vitro treatment of isolated macrophages with troglitazone. Interleukin-1 beta (Il1b) expression was not altered, and tumor necrosis factor-alpha (Tnf) expression was affected in peritoneal macrophages only. In ovarian macrophages, inducible nitric oxide synthase (Nos2), an important proinflammatory enzyme that regulates ovulation, was significantly reduced by troglitazone treatment, an effect that was restricted to cells from the preovulatory ovary. Thus, expression of PPARs within ovarian macrophages is hormonally regulated, reflecting the changing roles of these cells during the ovulatory cycle. Additionally, ovarian macrophages respond directly to troglitazone to downregulate expression of proinflammatory Nos2, providing mechanistic information about ovarian effects of TZD treatment.

cytokines, immunology, ovary

INTRODUCTION

Troglitazone is a member of a family of drugs called thiazolidinediones (TZDs), which also includes rosiglitazone (Avandia, GlaxoSmithKline) and pioglitazone (Actos, Takeda/Eli Lilly) [1, 2]. Thiazolidinediones have been demonstrated to be effective therapies for polycystic ovary syndrome (PCOS) [3, 4], an endocrine disorder characterized by elevated plasma androgen levels, chronic oligo- or amenorrhea, and arrested ovarian follicle growth and associated with hyperlipidemia, hyperinsulinemia, obesity, and diabetes mellitus [5]. Little is understood of the mechanisms by which these drugs might directly affect ovarian function in PCOS to restore regular menstrual cyclicity.

TZDs exert many of their effects through a mechanism that involves activation of the gamma isoform of the peroxisome proliferator-activated receptor (PPARG), a nuclear receptor. TZD-induced activation of PPARG alters the transcription of several genes that are considered key regulators of energy homeostatsis [6], insulin sensitivity [2, 7], and inflammation [8, 9]. Another important PPAR subtype, PPARA, controls fatty acid beta-oxidation pathways in the liver and in immune cells [10, 11].

While some biological responses to TZD are clearly mediated by improvements to insulin sensitivity of central and peripheral glucose-monitoring tissues, there are likely direct TZD targets and functions within the ovary. PPARA and PPARG mRNA and protein are both clearly present within the ovary [12–16], and activators of PPAR have been shown to influence steroid production by cultured granulosa, luteal, and theca cells [17–20]. Wu et al. [21] have previously reported that troglitazone treatment can reverse the expression imbalance between insulin receptor substrates 1 and 2 (IRS1 and IRS2) in human PCOS granulosa cells, although TZD effects on other ovarian cells, such as immunoregulatory macrophages, have not yet been examined.

In nonreproductive tissues, PPARG has been shown to have a variety of actions within resident macrophage immune cells, such as regulating cytokine mRNA expression to modulate overall inflammatory responses [8]. TZD activation of PPARG has been shown to affect monocyte/macrophage function by downregulating transcription of proinflammatory genes [22–26], such as inducible nitric oxide synthase (NOS2), gelatinase B, interleukins (IL) 1 beta (IL1B), IL6, IL12, and tumor necrosis factor alpha (TNF) [9, 23, 27].

Many of these same inflammatory mediators are required in the ovary for optimal follicle growth and ovulation, which is likened to an inflammatory-type response essential for follicle rupture but which must be resolved during formation of the corpus luteum [28]. In particular, TNF is an important factor in follicular development and atresia and luteal formation, maintenance, and regression [29]; IL1B promotes prostaglandin biosynthesis, plasminogen activator production, and collagenase activation; and nitric oxide is an important regulator of vasodilation and leukocyte chemotaxis within the ovary [30].

Our hypothesis is that, as in other tissues, macrophages within the ovary express PPARs and that PPARG in particular presents as a direct target for TZD action in these cells. Nuclear activation of PPARG in these cells would contribute to the regulation of inflammatory events during ovulation. PPARG protein was identified within the murine ovary, and messenger RNA from purified macrophages was isolated from the ovaries of gonadotropin-primed mice and examined for Ppara and Pparg expression at key time points of the ovulatory cycle. PPARG-regulated proinflammatory cytokine production within these cells following in vitro treatment with the TZD troglitazone provides insight into the activated function of PPARG and also implicates ovarian macrophages as direct targets of TZD therapy.

MATERIALS AND METHODS

Animals and Ovulation Induction

Sexually immature SV129 female mice (23–26 days of age) were used throughout this study. The animals were bred and housed at The Queen Elizabeth Hospital animal house at 24°C on a 14L:10D illumination cycle with water and pelleted food available ad libitum. The ethics committees of both the Queen Elizabeth Hospital and the University of Adelaide approved all experiments, and the animals were handled in accordance with the Australian Code of Practices for the Care and Use of Animals for Scientific Purposes. Mice were injected i.p. with 5 IU eCG (Intervet, Broxmeer, The Netherlands) in 0.1 ml of PBS (Dulbecco phosphate-buffered saline; Invitrogen Corporation, Auckland, NZ) with 0.1% BSA (wt/vol) (fraction V; Sigma Aldrich Chemical Co., St. Louis, MO) and 48 h later were either killed (preovulatory) or injected i.p. with 5 IU hCG (Pregnyl; N.V. Organon, Oss, The Netherlands) in 0.1 ml of saline and killed at 6 h (preovulatory) or 15, 24, or 48 h (postovulatory) after hCG. For Western blots and immunohistochemical studies, mature, naturally cycling mice housed in the same conditions were used.

Western Blots

Ovary, adipose tissue, and peritoneal macrophages (isolated from peritoneal lavage fluid) were homogenized in a 6 M urea and 0.1% Triton buffer containing protease inhibitors EDTA, benzamidine, leupeptin, and Sigma {alpha}-inhibitor cocktail (Sigma Aldrich) and then centrifuged, and the supernatant was collected. Total cellular protein was determined by Bradford assay (Bio-Rad), and 30 µg of protein from each extract were separated on 12% reducing SDS-polyacrylamide gels and transferred to polyvinylidene difluoride membrane (Immobilon-P; Millipore Corp., Bedford, MA). Prestained protein size markers from Bio-Rad were included in all Western blots. Nonspecific antibody binding was blocked by 1 h of room-temperature incubation of membranes with Tris-buffered saline and Tween 20 (TBST; 10 mM Tris [pH 7.5], 150 mM NaCl, and 0.05% Tween 20) containing 3% skim milk (wt/vol). Blots were then incubated at room temperature for 1 h with anti-PPARG-primary antibody 1:1000 (#2492; Cell Signaling Technology, Beverly, MA) in 3% milk, followed by 4 x 5-min washes in TBST and then incubated for 1 h with horseradish peroxidase-linked anti-rabbit IgG 1:10000 (#7074; Cell Signaling Technology), then washed 4 x 5 min with TBST. Enhanced chemiluminescence detection was performed using ECL Western Blotting Detection Reagents from Amersham Biosciences.

Immunohistochemistry

The distribution of PPARG and EMR1 protein expression in the murine ovary was determined by an avidin-biotin amplified peroxidase method. Ovarian tissue was collected from mature female mice and postdissection was fixed immediately in 4% paraformaldehyde (wt/vol) (BDH Laboratory Supplies, Poole, UK) at 4°C for 6 h, followed by overnight incubation in 18% sucrose (wt/vol) (Sigma Aldrich) at 4°C. Tissues were then embedded in optimal cutting temperature compound and snap-frozen in liquid nitrogen. Six-micrometer serial tissue sections were cut using a –20°C cryostat (Leica Instruments GmbH), defrosted at room temperature for 30 min, washed in 96% ethanol (vol/vol), and then washed in PBS (3 x 5 min). Endogenous peroxidase activity was blocked with 0.3% hydrogen peroxide (vol/vol) for 10 min and washed with PBS. The primary antibody, anti-PPARG (#2492; Cell Signaling Technology), at 1:50 in PBS/1% BSA/10% goat serum (vol/wt/vol) or anti-EMR1 (MCAP407; Serotec, Oxford, UK) at 1:500 in PBS/1% BSA/10% rabbit serum (vol/wt/vol), was then incubated for 2 h at room temperature in a humid chamber. Following PBS washes, antibody labeling was detected by incubation with either biotinylated goat anti-rabbit (Vector Laboratories Inc., Burlingame, CA) for PPARG or biotinylated rabbit anti-rat (DAKO Corporation, Carpinteria, CA) for EMR1, at 1:500 dilution in blocking buffer as described previously, for 2 h at room temperature. Following PBS washes, sections were incubated with avidin-linked horseradish peroxidase (DAKO) at 1:500 in PBS for 40 min at room temperature. The enzyme was visualized with Sigma-Fast DAB tablets (Sigma Aldrich), diluted in water, and applied for 10 min at room temperature. Negative controls in which primary antibody was replaced with PBS/1% BSA/10% serum were performed for both secondary antibodies. The sections were then dehydrated and mounted in DPX mountant (BDH Laboratory Supplies, Poole, UK).

Macrophage Isolation

Groups of 9–10 hormonally primed female SV129 mice were killed at selected pre- and postovulation time points as described earlier. All reagents were purchased from Sigma unless otherwise indicated. Resting peritoneal macrophages were obtained by peritoneal lavage, performed on four animals with 3 ml cold HBSS/EDTA/AZ (Hanks balanced salt solution [modified]/0.35 mg/ml sodium bicarbonate/0.625µl/ml gentamycin [DBL]/0.01% sodium azide [wt/vol]/5 mM EDTA). The addition of sodium azide to wash media at this subcytotoxic concentration inhibits macrophage capping and internalization of labeling antibodies bound to membrane antigens [31]. Recovered ovaries were dissected free of fat and connective tissue and examined for evidence of appropriate maturational stage before weighing. Tissues were then digested in {alpha}-MEM (MultiCel buffered with 2.98 mg/ml Hepes acid salt and 3.25 mg/ml Hepes sodium salt, with added 0.22 mg/ml sodium bicarbonate, 0.31 mg/ml calcium chloride dihydrate, and 0.624µl/ml gentamycin) containing collagenase (1 mg/20 mg tissue) and DNase (25µl/20 mg tissue). Tissues were incubated with gentle rocking for 45 min at 25°C, with additional agitation every 15 min. The ovarian digest was filtered through a 70-µm cell strainer (BD Falcon, Bedford, MA), and all tissue remnants were recovered and incubated for a further 20 min at 4°C in 3 ml HBSS/EDTA/AZ. Filtered cells and collected peritoneal lavage fluid were then centrifuged (200 x g at 4°C for 10 min). The tissue remnants were agitated by pipette and filtered once more without washing before being used to resuspend the ovarian pellet. The peritoneal pellet was also resuspended in 3 ml cold HBSS/EDTA/AZ. Both samples were underlayed with 3 ml fetal calf serum (FCS) and centrifuged (300 x g at 4°C for 10 min). Pellets were resuspended in 1 ml HBSS/10% FCS. Next anti-H2 and anti-EMR1 (both hybridoma supernatant) antibodies were used to specifically isolate the tissue macrophages. These hybridomas were obtained from the American Type Culture Collection (Rockville, MD) and are specific for EMR1 (also known as F4/80, a macrophage antigen of unknown function) and H2 (MHC class II) and have been used before [31]. H2 positivity defines mature macrophages and dendritic cells [32], while EMR1 positivity defines recently recruited as well as mature macrophages [33, 34]. In each sample, 500 µl of antibody were incubated with 500 µl of cells with gentle rocking for 1 h at 4°C with 500 µl of either anti-H2 or anti-EMR1 antibody in glass blood collection tubes (BD Vacutainer, silicon-coated interior). The FCS wash was repeated, and the pellets were resuspended in 1 ml HBSS/Gent. Labeled cells were next incubated on antibody panning plates for 2 h at 4°C with very gentle rocking. Panning plates were prepared by coating 3 cm Sarstedt microbiology-grade Petri dishes with 12.5µl anti-rat IgG (heavy and light chain, EMD Biosciences Inc., San Diego, CA) in PBS for 24 h at 4°C, followed by a PBS wash, and blocking with 10% FCS in PBS (vol/vol) at 4°C for 20 min. Unbound cells were washed off with 10 ml HBSS/10% FCS (vol/vol), and bound cells were lysed with 400 µl Tri Reagent (Sigma Aldrich). Cellular contents were scraped from the Petri dish and stored at –80°C until mRNA isolation. Maximal cell numbers and optimal purity of the cell population isolated using this method has been confirmed previously by establishing viability, positivity for leukocyte common antigen (CD45), phagocytic ability, negative progesterone secretion, and follicle-stimulating hormone receptor mRNA expression corresponding to less than 1% potential contamination [35].

Effect of Troglitazone on Isolated Macrophage Function

The effect of the TZD troglitazone on immune cells resident in the ovary and peritoneum was limited to cells positive for H2 because of the acquisition of a significantly greater number of cells with use of this antibody. All cells were isolated as described previously. Following removal of unbound cells from the antibody panning plate, bound macrophages were incubated for 24 h at 37°C in 5% CO2 in air with 800 µl Macrophage Serum Free Media (Gibco, Auckland, NZ) with or without 5µM troglitazone (Biomol, Plymouth Meeting, PA). This concentration of troglitazone was selected after experiments to determine maximal dosage without cytotoxic effects (data not shown). Cells were then lysed with Tri Reagent as described previously.

mRNA Extraction

Total cellular RNA was extracted using a modified Tri Reagent protocol, where 20 µg glycogen (Roche Diagnostics GmbH, Mannheim, Germany) were added to each sample before overnight precipitation at –20°C. To eliminate potential contamination by genomic DNA, each sample was DNase treated (Promega, Madison, WI) according to the manufacturer's instructions, dissolved in 20 µl PCR-grade water, and assayed using a Ribogreen RNA quantification kit (Molecular Probes, Eugene, OR) according to the manufacturer's instructions.

Reverse Transcription Real-Time PCR

Forty nanograms of mRNA were reversed transcribed using random primers (Roche) and a SuperScript II RNase H– Reverse Transcriptase (Invitrogen, Carlsbad, CA), RNaseOUT modified preamplification system for first-strand cDNA synthesis according to the manufacturer's protocol. For each reverse transcription, a control was performed in which all incubations and buffers were identical but no Superscript RT enzyme was added, verifying the absence of contaminating genomic DNA in PCR reactions. Complementary DNA templates were then subjected to fluorometric semiquantitative real-time PCR. For analysis of Il1b, Nos2, and Tnf mRNA, primers were designed using Primer Express software and synthesized by GeneWorks (Thebarton, SA, Australia):

Hprt1—F 5'-CTTCCTCCTCAGACCGCTTTT-3'

Hprt1—R 5'-AACCTGGTTCATCATCGCTAAC-3'

Il1b—F 5'-TGAAGTTGACGGACCCCAAA-3'

Il1b—R 5'-TGATGTGCTGCTGCGAGATT-3'

Tnf—F 5'-CCAGGCGGTGCCTATGTC-3'

Tnf—R 5'-GGCCATTTGGGAACTTCT-3'

Nos2—F 5'-CATCAGGTCGGCCATCACT-3'

Nos2—R 5'-CGTACCGGATGAGCTGTGAA-3'

Amplified cDNA was measured using SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA). For Ppara and Pparg mRNA analysis, TaqMan probes were purchased as an assay-on-demand kit from Applied Biosystems, containing sequence-specific forward and reverse primers and the fluorogenic TaqMan MGB probe (FAM dye-labeled; Applied Biosystems). Both Ppara and Pparg target sequences were amplified from regions within the ligand-binding domain, in a location that crossed exon 6 and 7 boundaries. For every 9 µl of diluted cDNA and 1 µl of primer and probe mix, 10 µl TaqMan Universal PCR Master Mix (containing AmpliTaq Gold DNA Polymerase and AmpErase UNG; Applied Biosystems) were added to the reaction well, which was run using the ABI PRISM 5700 Sequence Detection System (Applied Biosystems). Hypoxanthine-guanine-phosphoribosyltransferase (Hprt1) primers or an assay-on-demand kit was used as an internal control for every sample.

Data Analysis

For genes of interest, mRNA content was calculated for each sample relative to the housekeeping gene Hprt1. Hprt1 mRNA has previously been shown to be constitutively expressed in alveolar lung macrophages [36] and unaffected by the treatments administered within these experiments, as the critical threshold (CT) value per microgram of RNA did not vary statistically across treatment groups. Before collation and analysis of data, samples with a CT value >36 were considered negative and therefore not detectable. All real-time RT-PCR data were analyzed using the equation {Delta}{Delta}CT, where {Delta}CT is the difference between the gene of interest and the housekeeping gene. Data were subsequently normalized to become the fold change compared to a designated time point or experimental condition (e.g., 48 h post-eCG). To evaluate differences between groups, data were subjected to one-way ANOVA with Tukey post hoc analysis. At specific time points, macrophage populations were compared with a t-test. In all cases, differences were considered significant at P < 0.05. All statistical evaluation was performed using the software packages GraphPad InStat version 2.04a (GraphPad Software Inc., San Diego, CA) and SigmaStat for Windows version 2.03 (Jandel Corp., San Ramon, CA).

RESULTS

OvarianPPARG Expression and Localization with EMR1 in the Ovary

Western blots of 30-µg protein extracts from ovarian, peritoneal macrophage, and adipose tissue samples with anti-PPARG antibody revealed a similar level of expression in the ovary and adipose tissue (Fig. 1). The sample of resting, unstimulated peritoneal macrophages did not contain any detectable PPARG. This experiment also confirmed the specificity of this anti-PPARG antibody for ovarian immunohistochemical analysis.


Figure 1
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  		FIG. 1. Peroxisome proliferator-activated receptor (PPARG) protein in the murine ovary. Ovarian, peritoneal macrophage, and adipose protein extracts (30 µg) were analyzed by Western blot using anti-PPARG antibody. PPARG was detected in the ovarian and adipose tissue samples but not in resting peritoneal macrophages

Immunohistochemical localization of PPARG protein in mouse ovarian tissue showed intense staining within the nuclei of granulosa cells of both small and large follicles as well as moderate staining within discrete cells localized to thecal, stromal, and luteal regions (Fig. 2, a, c, and e). These results are consistent with reports localizing Pparg mRNA within the rat ovary [19, 37]. The macrophage marker protein EMR1 antigen was detected in a distribution typical of ovarian macrophages, within the thecal cell layers and corpora lutea (Fig. 2, b, d, and f). There were several areas of colocalization within the thecal, stromal, and luteal regions, suggesting coexpression of genes for PPARG and EMR1 in a subset of ovarian cells.


Figure 2
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  		FIG. 2. Peroxisome proliferator-activated receptor (PPARG) and EMR1 localization within the murine ovary. Serial ovarian sections were stained for PPARG (a, c, and e) or the macrophage marker EMR1 (b, d, and f) and visualized with DAB. The strongest PPARG staining is found in granulosa cells of all developing follicles. Within the thecal layer of small follicles (a and b) are isolated cells that express PPARG colocalized with cells expressing EMR1. This is also observed in the theca of larger and atretic follicles (c and d) and within corpora lutea (e and f). Original magnification x40

Pparg Expression by Ovarian Macrophages

Cells were isolated from ovaries based on their positivity for either H2 or EMR1 (markers frequently used to characterize macrophage identity [31, 33]) at two time points pre- and three time points postovulation. Real time-PCR analysis showed that macrophages within the ovary express Pparg throughout follicular development (Fig. 3a), and although gonadotropin stimulation does not modify Pparg expression in H2+ cells, EMR1+ macrophage displayed significantly reduced Pparg expression 6 h post-LH-stimulated ovulation (P < 0.02). Restricted expression within the EMR1+ cells returns to preovulatory levels as corpora lutea development progresses. Preovulation, no significant difference in receptor expression was observed between cell H2+ and EMR1+ types. Immediately following ovulation, H2+ ovarian cells expressed more Pparg than EMR1+ cells (P < 0.05) because of the apparent decrease in receptor expression observed in ovarian EMR1+ cells postovulation. At every stage of the induced cycle there was significantly more Pparg found within both H2+ and EMR1+ ovarian macrophages than in the corresponding macrophage population of the peritoneum (P < 0.001) (Fig. 3b). In isolated peritoneal macrophages, expression of Pparg was low, and no measurable response to hormonal stimulation was observed within these cells across the induced cycle.


Figure 3
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  		FIG. 3. Pparg expression in isolated macrophages. All data are normalized to Hprt1 and are expressed as the mean fold change ± SEM from the control group ovarian H2+ at time point eCG, which is given the value 1. In macrophages isolated from the ovary (a) Pparg expression is not hormonally regulated in H2+ cells (black bars) but is significantly downregulated in EMR1+ cells (white bars) 15 h post-hCG (* P = 0.018), with measurements returning to preovulatory levels following this. Peritoneal macrophages (b) do not respond to gonadotropic stimulation. At each time point there is significantly more Pparg mRNA within macrophages from the ovary than the peritoneum (P < 0.001) (n = 4 repeated experiments with 9–10 mice per hormonal treatment)

Ppara Expression by Ovarian Macrophages

Real time-PCR analysis also revealed that ovarian macrophages express detectable quantities of Ppara (Fig. 4a). Across the induced cycle, H2+ cells maintained relatively stable Ppara expression, while EMR1+ macrophages responded to hCG injection by rapidly downregulating transcription within 6 h (P < 0.05) and maintaining a reduced level of expression throughout the next 48 h. Preovulation, EMR1+ ovarian macrophages contained significantly more Ppara mRNA than H2+ cells, but levels were similar following the postovulatory downregulation observed in the EMR1+ cells. This contrasts with observations of Pparg expression, where significant differences between the macrophage subpopulations did not emerge until after ovulation. Messenger RNA levels of Ppara from peritoneal macrophages positive for both macrophage markers was very low throughout the cycle and could not be detected in several samples (Fig 4b).


Figure 4
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  		FIG. 4. Ppara expression in isolated macrophages. All data are normalized to Hprt1 and are expressed as the mean fold change ± SEM from the control group ovarian H2+ at time point eCG, which is given the value 1. In macrophages isolated from the ovary (a) Ppara expression is not hormonally regulated in H2+ cells (black bars) but is significantly downregulated in EMR1+ cells (white bars) immediately after hCG administration (* P < 0.05). In peritoneal macrophages (b) Ppara expression is low across the induced cycle. At 6 h (EMR1+), 24 h (EMR1+), and 48 h post-hCG (H2+ and EMR1+), mRNA expression is not detectable (ND) (n = 4 repeated experiments with 9–10 mice per hormonal treatment)

Effect of Troglitazone onPparg Expression

To investigate the effect of PPARG activation on mRNA expression, H2+ cells were isolated as described previously from preovulatory mice (48 h post-eCG) or postovulatory mice (48 h post-hCG) and treated for a further 24 h in vitro with 5 µM troglitazone. Pparg was analyzed in the macrophages from both the ovary and the peritoneal cavity and compared to non-troglitazone-treated controls (Fig. 5). Untreated ovarian macrophages demonstrated significant differences in Pparg mRNA content, unlike previous experimental results, which showed equal expression at eCG and 48 h post-hCG time points, indicating a potential influence of culture conditions on macrophage phenotype. Preovulatory ovarian macrophages did not display any change in Pparg mRNA in response to troglitazone. Treated cells isolated from the postovulatory murine ovary, however, demonstrated a 55% decrease in Pparg compared to the same population of cells unexposed to troglitazone. Pre- and postovulation expression of the Pparg transcript within H2+ peritoneal cells was minimal to undetectable and was not measurably influenced by the addition of troglitazone.


Figure 5
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  		FIG. 5. Troglitazone regulation of macrophage Pparg expression. Macrophages are isolated from ovaries (a) and peritoneal cavity (b) of mice pre- (48 h post-eCG) and postovulation (48 h post-hCG). Cells were cultured 24 h either with 5 µM troglitazone (black bars) or without (white bars). Results are expressed as the mean fold change ± SEM. The control group with the designated value 1 is the preovulation nontreated ovarian group. Graph bars with the same letter are not significantly different (all significant differences P < 0.05) (n = 4 repeated experiments with 9–10 mice per hormonal treatment)

Effect of Troglitazone on Inflammatory Mediator mRNA Expression

Expression of mRNA for the inflammatory mediators TNF, IL1B, and NOS2 was assessed in ovarian and peritoneal H2+ macrophages following 24-h culture with 5 µM troglitazone and compared to nontreated controls (Fig. 6). While ovarian macrophages displayed an upregulation of Tnf expression following ovulation, troglitazone did not mediate any response from these cells. In contrast, Tnf production within peritoneal macrophages was significantly reduced, by nearly half, following exposure to 5 µM troglitazone. Ovarian Il1b expression was not significantly affected by troglitazone, and similarly no effect was seen in comparative populations of peritoneal macrophages. Troglitazone treatment did, however, lead to a significant downregulation of Nos2 production before ovulation, effectively reducing expression by 80% into the range of lower postovulatory levels (P = 0.03). The pattern of preovulatory downregulation of Nos2 expression in peritoneal cells mirrored the results found in ovarian macrophages but did not reach significance.


Figure 6
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  		FIG. 6. Troglitazone regulation of macrophage gene expression. Macrophages are isolated from ovaries (a) and peritoneal cavity (b) of mice pre- (48 h post-eCG) and postovulation (48 h post-hCG). Cells were cultured 24 h either with 5 µM troglitazone (black bars) or without (white bars). Expression of inflammatory mediators Tnf (i), Il1b (ii), and Nos2 (iii) was analyzed. Results are expressed as the mean fold change ± SEM. The control group with the designated value 1 is the preovulation nontreated group. Graph bars with the same letter are not significantly different (all significant differences P < 0.05) (n = 4 repeated experiments with 9–10 mice per hormonal treatment)

DISCUSSION

In the ovary, tissue-bound macrophages are especially important in the cyclic reproductive events because they secrete a number of inflammatory and immunomodulating cytokines and enzymes and perform phagocytotic duties associated with tissue remodeling [38, 39]. There are pathological examples where immune dysfunction might contribute to ovarian disease, including premature ovarian failure, PCOS, and endometriosis [39, 40]. Treatment of these disorders might involve administration of PPARG-activating TZDs, including troglitazone (which used to be frequently prescribed), that affect insulin sensitivity, glucose utilization [2, 7], and immune cell activity [8, 9]. PPARG has been shown to have a variety of actions within certain populations of resident macrophages to regulate cytokine mRNA expression to consequently modulate overall inflammatory responses.

Our results reveal that within the ovary, macrophage Pparg levels are carefully maintained across the ovarian cycle, with a transient suppression coinciding with the brief window of time when the structural reorganization of ovulation necessitates the production of inflammatory cytokines. Interestingly the ovulatory downregulation of Pparg occurs specifically in EMR1+ macrophages, while H2+ cells appear resistant to such gonadotropin-stimulated effects, displaying no hormonal regulation of either Pparg or Ppara genes. H2+ cells are considered to be dendritic cells or specialized macrophages capable of antigen presentation. These results suggest specific differences in the functions of macrophages expressing EMR1 and/or H2 and support observations in the mouse uterus [31] of variations in lineage development and important functional differences between these macrophage populations. Pparg gene expression was downregulated by treatment with troglitazone, but only in ovarian macrophages collected after ovulation. The minimal level of peritoneal macrophage Pparg expression was not affected. Troglitazone-induced downregulation of Pparg has been previously described in vitro in differentiated 3T3-L1 adipocytes [41]. However, in hepatocytes treated in vitro [42] or in vivo [43], hepatic Pparg mRNA and protein increased. Such results, perhaps indicating tissue-specific troglitazone effects, are here shown to be also dependent on other microenvironmental factors, such as follicle maturation state. Interestingly, we found that although the levels of Pparg in peritoneal macrophages were undetectable, troglitazone was still able to influence the production of Tnf in these cells when isolated from animals before ovulation. Similarly, in ovarian macrophages, troglitazone exerted a greater effect on Nos2 expression when preovulatory Pparg levels were relatively reduced. This may suggest PPARG-independent actions or pathways that are interesting in light of other experiments showing PPARG-independent anti-inflammatory effects of TZDs [44] as well as the maintenance of TZD-mediated effects in PPARG-deficient macrophages [26, 45]. In addition, more recent studies suggest that the natural PPARG ligand 15-deoxy-{Delta}12,14 prostaglandin J2 (15d-PGJ2) and some TZDs may operate independently of PPARG through direct induction of other anti-inflammatory mediators, such as suppressor of cytokine signaling 1 [46], to thereby inhibit immune responses. The relatively high range of troglitazone concentration employed in this experiment would potentially allow such nonspecific effects to occur.

High expression and activity of PPARG is a mediator of a type 2 immune response that results in anti-inflammatory immune activation. Thus our results here are consistent with the hypothesis emerging from several sources of evidence in our laboratory that ovarian macrophages produce a protective anti-inflammatory response during tissue remodeling at ovulation [35]. In normal fertile cycles the induction of cyclooxygenase 2 in response to the LH surge results in synthesis of prostaglandins [47], including 15d-PGJ2 [48], that may interact endogenously with PPARG in macrophages surrounding follicles. Macrophages respond with a downregulation of type 1 inflammatory compounds such as NOS2 as well as a transient downregulation of the PPAR receptor as we demonstrate here using exogenous troglitazone in vitro. If future appropriate in vivo experimentation reveals the same cellular responses, the administration of troglitazone in PCOS patients can be concluded to have a direct effect in ovarian macrophages and may mediate in part the therapeutic effects.

This study is the first to localize PPARG protein in the murine ovary and to also show that macrophages from these ovaries express both Ppara and Pparg at levels dramatically higher than macrophages residing in nonreproductive tissues. A physiological role for PPARG is suggested to be important within the ovary, where normal fertility is dependent on the appropriate induction as well as resolution of macrophage-derived inflammatory signals. Pparg levels are also responsive to hormonal stimulation across the ovarian cycle, and we have shown that ovarian macrophages respond to troglitazone in vitro by downregulating an important inflammatory mediator (Nos2) in these cells. Such new information about TZDs contributes to our understanding of PCOS treatment, as these drugs that have traditionally been thought to target discrete nonreproductive tissues to primarily improve insulin sensitivity are here shown to also influence additional aspects of ovarian physiology.

ACKNOWLEDGMENTS

We thank Dr. Darryl Russell for technical assistance and critical reading of this manuscript.

FOOTNOTES

1 Supported by the National Health and Medical Research Council of Australia. Back

2 Correspondence: Robert J. Norman, Research Centre for Reproductive Health, University of Adelaide, Queen Elizabeth Hospital, First Floor, Maternity Building, 28 Woodville Rd., Woodville SA 5011, Australia. FAX: 61 8 8222 7521; robert.norman{at}adelaide.edu.au Back

Received: 10 May 2005.

First decision: 30 May 2005.

Accepted: 27 September 2005.

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