1Division of Pulmonary and Critical Care Medicine, Department of Medicine, University of Michigan Medical School, Ann Arbor 48109-0360; and 2Department of Pathology, Wayne State University School of Medicine, Detroit, Michigan 48201
Submitted 27 June 2003 ; accepted in final form 7 November 2003
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ABSTRACT |
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15-deoxy-12,14-prostaglandin J2; rosiglitazone; peroxisome proliferator-activated receptor-
2
Peroxisome proliferator-activated receptors (PPARs) are ligand-activated transcription factors belonging to the nuclear hormone receptor family. Three PPAR isoforms (,
, and
), each with a specific pattern of expression, have been identified. PPAR-
plays a critical role in adipocyte differentiation and glucose homeostasis (18, 26). Ligands for PPAR-
include a variety of compounds such as hydroxyeicosatetraenoic acids, hydroxyoctadecanoic acids, 15-deoxy-
12,14-prostaglandin J2 (15d-PGJ2), and the antidiabetic thiazolidinediones (9, 16, 20). In addition to expression in adipocytes, PPAR-
expression has been found in immune cells. In immune cells, treatment with PPAR-
ligands results, for the most part, in downregulation of inflammatory responses (10, 11, 15, 24).
PPAR- has been characterized in other monocyte/macrophage populations; however, no prior studies have assessed the expression, regulation, or functional significance of PPAR-
in AMs. Further investigation of the role of PPAR-
in AMs may provide important insights into pulmonary host defense mechanisms. Therefore, we characterized PPAR-
expression in AMs and investigated the effects of selective PPAR-
ligands on the regulation of immune responses by these cells.
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MATERIALS AND METHODS |
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Animals. C57BL/6 wild-type mice (Jackson Laboratory) were housed in specific pathogen-free conditions within the animal care facility at the University of Michigan until the day of death. All experimental procedures were approved by the University of Michigan's University Committee on Use and Care of Animals. The experiments were performed in adherence to the National Institutes of Health guidelines on the use of laboratory animals.
Bronchoalveolar lavage and AM isolation. Mice were euthanized by asphyxia in a high-CO2 environment. The trachea was then exposed and intubated using a 1.7-mm-OD polyethylene catheter. Bronchoalveolar lavage (BAL) was performed by instillation of Dulbecco's PBS containing 5 mM EDTA in 1-ml aliquots. PBS was instilled at 10 ml per mouse; 9 ml of lavage fluid were retrieved. Lavaged cells from each group of animals were pooled. Lavaged cells consisted of >95% AMs for each of the groups examined (data not shown). Cells were then washed, seeded onto tissue culture-treated plates (Costar, Cambridge, MA), and allowed to adhere for 1 h, and nonadherent cells were washed away.
Peritoneal lavage and peritoneal macrophage isolation. Mice were euthanized by asphyxia in a high-CO2 environment. Peritoneal lavage was then performed by instillation of Dulbecco's PBS containing 5 mM EDTA in 1-ml aliquots into the peritoneal cavity. PBS was instilled at 5 ml per mouse; 4 ml of lavage fluid were retrieved. Lavaged cells from each group of animals were pooled. Cells were then washed, seeded onto tissue culture-treated plates, and allowed to adhere for 1 h, and nonadherent cells were washed away.
Macrophage culture conditions. AMs were washed and resuspended in RPMI 1640 (Life Technologies). For culture supernatants and isolation of RNA, cells were plated at 5 x 105 and 1 x 106 cells/ml, respectively, into six-well tissue culture plates. The cells were pretreated with PPAR- ligands for 4 h and then incubated with LPS (100 ng/ml) for 16 h in RPMI 1640 supplemented with 100 U/ml penicillin and 100 µg/ml streptomycin (Life Technologies) and 10% FCS (Life Technologies) for ELISA and for 6 h for mRNA analysis at 37°C under an atmosphere of 5% CO2. AM culture supernatants were harvested and stored at -80°C until analyzed by ELISA. In separate experiments, adherent peritoneal macrophages (PMs) and AMs in RPMI 1640 were washed and treated with TRIzol reagent (Life Technologies) for RNA isolation and stored at -80°C until analyzed by RT-PCR.
Real-time quantitative PCR (TaqMan). Gene-specific primers and probes were designed using Primer Express software (Perkin-Elmer/PE Applied Biosystems, Foster City, CA). The sequences were as follows: 5'-CCTGACGGGTCTCGGTTG-3' (forward) and 5'-TGTCCTGAATATCAGTGGTTCACC-3' (reverse) for PPAR-1, 5'-GCTTCTTTCAAATCTTGTCTGTCACACAGTCCTG-3' for PPAR-
1 probe, 5'-TGGGTGAAACTCTGGGAGATTC-3' (forward) and 5'-AATTTCTTGTGAAGTGCTCATAGGC-3' (reverse) for PPAR-
2, and 5'-CCTGTTGACCCAGAGCATGGTGCC-3' for PPAR-
2 probe. Oligonucleotide primers and TaqMan probe for 18S rRNA internal control were purchased from Perkin-Elmer/PE Applied Biosystems. The real-time quantitative RT-PCR was performed essentially following the manufacturer's protocol. Briefly, the reaction mixture contained 5.5 mM MgCl2, 500 µM dNTP, 2.5 µM random hexamers, 200 nM FAM probe, and forward and reverse primers at 600 nM in a final volume of 25 µl and was analyzed in an ABI PRISM 7700 sequence detection system. Relative quantitation of PPAR-
mRNA levels was plotted as fold change compared with day 0 of 3T3-L1 cells. 18S rRNA was used for normalization. TaqMan RT reactions were performed in triplicates, and the experiments were repeated independently at least three times.
Western blot analysis and antibodies. Western blot analysis was performed using previously described methods (32). Briefly, crude protein extracts were obtained by lysing 2 x 106 cells in a buffer [50 mM Tris·HCl (pH 7.6), 1% Nonidet P-40, 2 mM EDTA, 0.5% sodium deoxycholate, 150 mM NaCl, 1 mM sodium orthovanadate, 2 mM EGTA, 4 mM sodium p-nitrophenyl phosphate, and 100 mM sodium fluoride] with protease inhibitors (0.5% leupeptin, 0.5% aprotinin, and 0.2% phenylmethylsulfonyl fluoride). Samples containing 20 µg of total protein were electrophoresed on SDS-polyacrylamide gels and transferred to nitrocellulose membranes by electroblotting. Membranes were probed with antibodies as indicated and then with horseradish peroxidase-conjugated mouse or rabbit secondary antibodies (Pierce, Rockford, IL) and West Pico chemiluminescence detection reagents (Pierce). A murine monoclonal antibody against PPAR- (E8, Santa Cruz Biotechnology, Santa Cruz, CA) or a rabbit polyclonal antibody against inducible NO synthase (iNOS; Santa Cruz Biotechnology) was used at 1:200 for 1 h at room temperature. Mouse monoclonal antibody against
-actin was obtained from Sigma Chemical. In each experiment, blots were stripped and probed for
-actin to confirm equal loading.
Immunofluorescence staining. AMs plated on glass coverslips were initially rinsed with Dulbecco's modified Eagle's medium (GIBCO, Grand Island, NY) for 30 s at ambient temperature and then fixed in 4% formaldehyde for 5 min. Cells were then washed three times in PBS before permeabilization and after each subsequent step. Permeabilization was performed in buffer consisting of 1.0% Triton in 50 mM PIPES (pH 7.0), 90 mM HEPES (pH 7.0), 0.5 mM MgCl2, 0.5 mM EGTA, and 75 mM KCl for 5 min at room temperature. Coverslips were sequentially incubated with mouse monoclonal antibody against PPAR- (Santa Cruz Biotechnology) and FITC-labeled anti-mouse antibody (Jackson ImmunoResearch Laboratories, West Grove, PA), each for 60 min at room temperature. Cells were then visualized and photographed using a Zeiss fluorescence microscope.
Transient reporter assays. AMs were plated in 12-well plates at a density of 2 x 106 cells/well. The cells were transfected in the presence of 10% FCS at 2 µg/well with the PPAR--responsive reporter plasmid FATP3X-tk-luciferase, which contains three copies of the mouse FATP PPRE upstream of a firefly luciferase gene and thymidine kinase promoter, using Effectene (according to the manufacturer's instructions). The Renilla luciferase plasmid pRL-SV40 (Promega, Madison, WI; 0.1 µg /well) was also transfected as an internal control for monitoring transfection efficiency and for normalizing the firefly luciferase activity. After 24 h of transfection, cells were washed and then treated with PPAR-
ligands in the presence of serum. After 24 h, protein extracts generated from harvested cells were assessed for luciferase and control vector using a dual-luciferase assay system (Promega).
Measurement of H2O2 release. A peroxidase-based fluorometric assay was used to measure extracellular release of H2O2, as previously described (32). Briefly, AMs were plated in RPMI 1640 supplemented with 10% FCS at a density of 1 x 106 cells/well in six-well dishes. Cells were treated with 10 µM rosiglitazone or DMSO (vehicle control) for 16 h. Cells were washed with Dulbecco's PBS, pH 7.4, and then incubated with a reaction mixture containing 100 µM homovanillic acid, 5 U/ml horseradish peroxidase (type IV), and 1 mM HEPES in Hanks' balanced salt solution without phenol red. PMA (1 µM) was immediately added to this reaction mixture. The reaction mixture was collected after 1 h of incubation, pH was adjusted to 10.0 with 0.1 M glycine-NaOH buffer, and fluorescence was measured at excitation and emission wavelengths of 321 and 421 nm, respectively.
RT-PCR amplification of AM mRNA. Total cellular RNA from AMs was isolated from 1 ml of TRIzol reagent (GIBCO) following the TRIzol protocol. Expression of mRNA was determined by RT-PCR using an RT-PCR system kit (Access, Promega) following the manufacturer's protocol. The following primer pairs were used for specific mRNA amplification: 5'-CCT-GTA-GCC-CAC-GTC-GTA-GC-3' (forward) and 5'-TTG-ACC-TCA-GCG-CTG-AGT-TG-3' (reverse) for TNF-, 5'-TGA-GCT-GCG-CTG-TCA-GTG-CCT-3' (forward) and 5'-AGA-AGC-CAG-CGT-TCA-CCA-GGA-3' (reverse) for KC, 5'-TGC-CTG-AAG-ACC-CTG-CCA-AGG-3' (forward) and 5'-GGT-AGC-CTT-GCC-TTT-GTT-CAG-3' (reverse) for MIP-2, 5'-CTA-TGC-TGC-CTG-CTC-TTA-3' (forward) and 5'-ATG-GCC-TTG-TAG-ACA-CCT-3' (reverse) for IL-10, 5'-ATG-TTG-TAG-AGG-TGG-ACT-3' (forward) and 5'-GGA-CTG-CTA-CTG-CTC-TTG-AT-3' (reverse) for IL-12 p40, and 5'-ATG-GAT-GAC-GAT-ATC-GCT-C-3' (forward) and 5'-GAT-TCC-ATA-CCC-AGG-AAG-G-3' (reverse) for
-actin. After amplification, the cDNA products were separated on a 2% agarose gel containing 0.3 mg/ml (0.003%) of ethidium bromide, and bands were visualized and photographed using UV transillumination.
Murine cytokine ELISAs. Murine TNF, IL-12, KC, and MIP-2 were quantitated using a modification of a double-ligand method, as previously described (38). Briefly, flat-bottomed 96-well microtiter plates were coated with 50 µl/well of rabbit antibody against the various cytokines for 16 h at 4°C and then washed. The plates were rinsed four times with wash buffer, diluted cell-free supernatants (50 µl) in duplicate were added, and the samples were incubated for 1 h at 37°C. The plates were washed four times, biotinylated rabbit antibodies against the specific cytokines (50 µl/well) were added, and the plates were incubated for 30 min at 37°C. The plates were washed four times, streptavidin-peroxidase conjugate (Bio-Rad Laboratories, Richmond, CA) was added, and the plates were incubated for 30 min at 37°C. The plates were washed again four times, and chromogen substrate (Bio-Rad Laboratories) was added. The plates were incubated at room temperature to the desired extinction, and the reaction was terminated with 3 M H2SO4 solution (50 µl/well). Plates were read at 490 nm in an ELISA reader.
Statistical analysis. Values are means ± SE. Statistical significance was determined using a two-tailed Student's t-test. All calculations were performed on the Prism 3.0 statistical program (Graphpad Software, San Diego, CA). P < 0.05 was considered significant.
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RESULTS |
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To assess murine AM PPAR- protein expression, we analyzed protein extracts by Western blot analysis in AMs obtained from C57BL/6 mice. Resident PMs were also isolated from these mice by peritoneal lavage. Significant constitutive expression of PPAR-
protein was detected in AMs; expression in PMs was minimal (Fig. 2A). To determine cellular localization of PPAR-
, immunofluorescence staining of freshly prepared murine AMs was performed. Figure 2B demonstrates that PPAR-
is predominantly localized in the nucleus. Western blot analysis of nuclear and cytosolic fractions confirmed PPAR-
nuclear compartmentalization, with almost undetectable expression in the cytosolic fraction (data not shown).
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IL-4 is a cytokine that has been shown to inhibit proinflammatory cytokine expression and NO production from macrophages (2, 31). Treatment of murine AMs with IL-4 (10 ng/ml) resulted in induction of PPAR- protein expression (Fig. 3).
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Constitutive PPAR- in AMs is functionally active. We assessed whether PPAR-
expressed by AMs could be activated. AMs were transfected with a PPAR-dependent promoter gene construct, FATP3x-tk-luciferase, containing three direct repeats of the fatty acid transport protein PPAR response element subcloned upstream of the thymidine kinase promoter. Both 15d-PGJ2 (5 µM) and rosiglitazone (10 µM) activated transcription of the FATP3x-tk-luciferase reporter gene (2.2- and 3.7-fold, respectively; Fig. 4).
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Effect of PPAR- ligands on H2O2 production and iNOS expression. To determine whether activation of PPAR-
inhibited AM oxidative burst, we examined the effects of PPAR-
ligands on the extracellular release of H2O2 from AMs. Murine AMs were treated with 10 µM rosiglitazone for 16 h and then stimulated with 1 µM PMA and assayed for H2O2 production using a peroxidase-based fluorometric assay. Rosiglitazone decreased PMA-induced H2O2 release from AMs (Fig. 5). In addition, we assessed the effects of rosiglitazone on LPS + IFN-
-induced iNOS expression. LPS (100 ng/ml) and IFN-
(75 U/ml) significantly induced iNOS expression in AMs, an effect that is inhibited by treatment with 10 µM rosiglitazone for 16 h (Fig. 6).
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Effect of PPAR- ligands on murine AM cytokine production. To assess the effects of PPAR-
ligands on proinflammatory cytokine production, murine AMs were treated with selective PPAR-
agonists and then stimulated with LPS. AMs were pretreated for 4 h with rosiglitazone, troglitazone, ciglitazone, or 15d-PGJ2 at various concentrations and then with LPS (100 ng/ml) for 16 h. Levels of the proinflammatory cytokines IL-12 and TNF-
and the chemokines MIP-2 and KC were then measured. All ligands significantly inhibited IL-12 production by AMs, with less effect on TNF-
production (Fig. 7). Effects on KC and MIP-2 production were not statistically significant. 15d-PGJ2 produced the most significant inhibition of TNF-
production by AMs. Likewise, IL-12 (p40) mRNA levels were most significantly inhibited by rosiglitazone treatment, with less effect on TNF-
and no significant effect on KC and MIP-2 mRNA expression. No expression of IL-10 mRNA was noted in the LPS- or LPS + rosiglitazone-treated cells. PPAR-
ligands were used at doses that were nontoxic and did not affect cell viability as measured by 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium assay (data not shown). DMSO, used as a vehicle for PPAR-
ligands, did not affect cell viability or modulate cytokine production at the concentrations used in this study (data not shown).
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DISCUSSION |
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Our results are unique, inasmuch as no prior studies have characterized the significance of PPAR- ligands in modulation of AM function. In addition, in contrast to freshly isolated PMs, we demonstrated that AMs constitutively express high PPAR-
levels. The significance of this difference is unclear, but macrophages isolated from different anatomic sites differ functionally and phenotypically. AMs are known to vary from other macrophage populations, and from PMs in particular (6). Considerable differences in cell surface receptor expression, antigen-presenting capacity, eicosanoid production, and phagocytosis between AMs and other macrophage populations have been observed (4, 5, 12, 19). The reason for constitutively high expression of PPAR-
by AMs may be the surrounding milieu and environmental location of these cells. It is vital that host responses in the lung be tightly regulated. Unchecked inflammatory responses by AMs can lead to injury of surrounding tissue (30). PPAR-
may thus provide an important immunosuppressive function in lung AMs. Likewise, IL-4 has been shown to inhibit proinflammatory cytokine expression, NO production, and CD14 cell surface expression from macrophages. Moreover, this cytokine can induce the expression of other anti-inflammatory mediators, including IL-1 receptor antagonist and TNF-soluble receptors from monocytes (2, 14, 29, 31). IL-4 has also been shown to increase 12/15-lipoxygenase synthesis in macrophages, leading to the generation of endogenous PPAR-
ligands (13). As in other studies, IL-4 increased PPAR-
expression in AMs (13, 39).
Analysis of the specific PPAR- isoforms expressed by AMs revealed preferential expression of PPAR-
2 relative to PPAR-
1 in these cells. PPAR-
2 is expressed mostly in fat tissue, whereas PPAR-
1 is found at low levels in other tissues (28). The functional and biological significance of the differential expression of PPAR-
isoforms is still not completely known. PPAR-
2 has been shown to be critically important in adipogenesis, in contrast to PPAR-
1 (22). The expression of the PPAR-
variants has not been well characterized in monocytes/macrophages, although PPAR-
1 appears to be more highly expressed in murine PMs (13). The significance of high expression of PPAR-
2 requires further investigation. In addition to being highly expressed, we have demonstrated that PPAR-
in AMs is functionally active and localized in the nuclear compartment.
AMs generate superoxide, which dismutates to H2O2 by assembly and activation of the multicomponent NADPH oxidase. This respiratory burst enzyme is primarily responsible for bactericidal killing as part of the innate host response (3). We observed inhibition of PMA-stimulated oxidative burst in AMs by rosiglitazone. Recent data suggest that endogenously generated NO activates PPAR- and results in PPAR-
-mediated downregulation of p47 phagocyte oxidase, a subunit of NADPH oxidase, in RAW 264.7 and U937 cells (37). This suggests a potential mechanism for the attenuation of oxidative burst activity in AMs. Our results also demonstrated reduction of iNOS expression in these cells with rosiglitazone treatment. We and others have demonstrated that NO is a critical regulator of pulmonary host defenses (25, 35). PPAR-
may play an important role in this regulation.
IL-12 is a pivotal cytokine for the development of Th1 cells and the initiation of cell-mediated immune responses to several pathogens (34). Overall, the various PPAR- ligands were not potent inhibitors of LPS-induced cytokine production, with the exception of effects on IL-12 and TNF-
production by 15d-PGJ2. 15d-PGJ2 was the only ligand that demonstrated inhibition of multiple cytokines and suggests, in light of other studies, activation of PPAR-
-independent pathways (8, 36). Our results are consistent with data examining the role of PPAR-
ligands in regulation of monocyte/macropage cytokine production, although some interesting differences exist (1, 15, 33). Similar to the study by Thieringer et al. (33), we observed only modest inhibitory effects on proinflammatory cytokine production by PPAR-
ligands. However, they did not examine ligand effects on production of IL-12, the cytokine found to be most significantly inhibited in our study. Although Alleva et al. (1) demonstrated potent inhibition of LPS-induced IL-12 production with PPAR-
agonists, they used activated macrophages (thioglycolate-elicited PMs), in contrast to our noninflammatory AMs. In addition, they also found significant inhibition of TNF-
with ligands other than 15d-PGJ2. The levels of LPS-induced IL-12 production were also significantly higher in thioglycolate-elicited PMs than AMs, although the degree of IL-12 inhibition by PPAR-
ligands was similar (1). Effects of PPAR-
ligands on PMA-induced cytokine production by AMs were not analyzed, as in the study by Jiang et al. (15), secondary to the finding that PMA stimulation of AMs did not induce cytokine production (as measured by ELISA, data not shown). Similar to other studies, we found that ciglitazone was less potent at inhibiting cytokine production than other PPAR-
ligands tested (1, 33). No effects of any of the ligands on inhibition of chemokine production were seen. Because of concerns of the potential apoptotic/toxic effects of PPAR-
agonists, we carefully monitored the effects of these ligands on cell viability. In all studies, PPAR-
ligands were used at doses that did not affect viability. Our results indicate that higher doses of ciglitazone (>3 µM), troglitazone (>3 µM), rosiglitazone (>16 µM), and 15d-PGJ2 (>7 µM) were needed to affect viability (data not shown). Finally, similar to the results obtained by Alleva et al., we did not find induction in expression of the anti-inflammatory cytokine IL-10 with LPS + rosiglitazone over LPS alone, suggesting that IL-10 likely is not involved in PPAR-
-mediated AM deactivation.
In summary, we have characterized the expression and functional roles of PPAR- in AM deactivation. These cells play a vital role in pulmonary host defense and inflammatory processes (17). Further investigation into the mechanisms by which PPAR-
regulates AM function will improve our understanding of the role of PPAR-
in immune-mediated lung diseases.
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ACKNOWLEDGMENTS |
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This work was supported in part by National Heart, Lung, and Blood Institute Grants K08 HL-070068 (R. C. Reddy) and HL-58200, HL-57243, and P50 HL-60289 (T. J. Standiford). R. C. Reddy holds a Research Award from the American Lung Association.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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