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The Journal of Immunology, 2002, 169: 2619-2626.
Copyright © 2002 by The American Association of Immunologists

Activation of Peroxisome Proliferator-Activated Receptor {gamma} by Nitric Oxide in Monocytes/Macrophages Down-Regulates p47phox and Attenuates the Respiratory Burst1

Andreas von Knethen and Bernhard Brüne2

Institute of Cell Biology, University of Kaiserslautern, Kaiserslautern, Germany


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
NO appears as an important determinant in auto and paracrine macrophage function. We hypothesized that NO switches monocyte/macrophage function from a pro- to an anti-inflammatory phenotype by activating anti-inflammatory properties of the peroxisome proliferator-activated receptor (PPAR){gamma}. NO-releasing compounds (100 µM S-nitrosoglutathione or 50 µM spermine-NONOate) as well as inducible NO synthase induction provoked activation of PPAR{gamma}. This was proven by EMSAs, with the notion that supershift analysis pointed to the involvement of PPAR{gamma}. PCR analysis ruled out induction of PPAR{gamma} mRNA as a result of NO supplementation. Reporter assays, with a construct containing a triple PPAR response element in front of a thymidine kinase minimal promoter driving the luciferase gene, were positive in response to NO delivery. DNA binding capacity as well as the transactivating capability of PPAR{gamma} were attenuated by addition of the antioxidant N-acetyl-cysteine or in the presence of the NO scavenger 2-phenyl-4,4,5,6-tetramethyl-imidazoline-1-oxyl 3-oxide. Having established that NO but not lipophilic cyclic GMP analogs activated PPAR{gamma}, we verified potential anti-inflammatory consequences. The oxidative burst of macrophages, evoked by phorbol ester, was attenuated in association with NO-elicited PPAR{gamma} activation. A cause-effect relationship was demonstrated when PPAR response element decoy oligonucleotides, supplied in front of NO delivery, allowed to regain an oxidative response. PPAR{gamma}-mediated down-regulation of p47 phagocyte oxidase, a component of the NAD(P)H oxidase system, was identified as one molecular mechanism causing inhibition of superoxide radical formation. We conclude that NO participates in controlling the pro- vs anti-inflammatory phenotype of macrophages by modulating PPAR{gamma}.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Inflammation provokes the generation of multiple mediators, including NO. It has been shown that monocytes/macrophages are prominent producers of NO, with the implication that NO is a key component of the antimicrobial and tumoricidal immune response (1). NO not only serves as a cytotoxic molecule to produce cell demise along an apoptotic or necrotic pathway but also gained attention as a regulator of immune function (2, 3). Part of these activities is attributed to the ability to regulate protein expression. Although in many cases the primary targets for NO are uncertain, a number of signal transducers, i.e., transcription factors, are known to be positively or negatively regulated by NO. Examples comprise the redox-sensitive factors NF-{kappa}B or AP-1 (4, 5, 6). As a consequence, the expression of cytokines, apoptotic proteins, or immediate early genes such as cyclooxygenase-2 are modulated, thus stressing the function of NO as a messenger molecule (7, 8).

The anti-inflammatory properties of the nuclear hormone receptor family known as peroxisome proliferator-activated receptors (PPARs)3 were recently established (9). Three subtypes, PPAR{alpha}, PPAR{beta} (also known as PPAR{delta}), and PPAR{gamma} have been described so far, originally found in association with obesity, diabetes, and atherosclerosis (10). All of them act as ligand-dependent transcription factors which, upon heterodimerization with the 9-cis retinoic acid receptor (RXR), bind to the PPAR response element (PPRE) to modulate target gene expression (11). PPAR{gamma} has been shown to be activated by natural agonists such as 15-deoxy-{Delta}12,14-PGJ2 (15d-PGJ2) or synthetic antidiabetic thiazolidinediones, i.e., ciglitazone, with the outcome to reduce proinflammatory cytokine as well as reactive nitrogen species production in monocytes/macrophages (12, 13). In addition, activation may be achieved by oxidized low-density lipoprotein via the CD36 scavenger receptor (14, 15), which has been linked to the development of atherosclerosis. Experiments performed with cells derived from murine embryonic stem cells that were homozygous for a null mutation in the PPAR{gamma} gene questioned the anti-inflammatory properties of PPAR{gamma} (16) but verified its role in CD36 expression (17).

Considering the properties of NO as a messenger molecule during innate and adaptive immunity, we sought to correlate NO actions to an anti-inflammatory macrophage phenotype. Therefore, we asked whether NO may activate PPAR{gamma}. We demonstrate activation of PPAR{gamma} in macrophages by chemically distinct NO donors as well as endogenously synthesized NO by inducible NO synthase (iNOS) induction and link an active PPAR{gamma} with an attenuated oxidative burst. Inhibition of superoxide radical formation was due to PPAR{gamma}-mediated down-regulation of p47 phagocyte oxidase (p47phox), one important component of the NAD(P)H oxidase enzyme complex. We conclude that anti-inflammatory properties of NO may be transmitted by gene regulation affected by PPAR{gamma} activation.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials

PMA, N-acetyl-cysteine (NAC), 2-phenyl-4,4,5,6-tetramethyl-imidazoline-1-oxyl 3-oxide (PTIO), LPS, and 8-bromo-cGMP were purchased from Sigma (Deisenhofen, Germany). Spermine-NONOate (spermine-NO) was obtained from Bio-Trend (Cologne, Germany). Hydroethidine (HE) was from Molecular Probes (Leiden, The Netherlands). The polyclonal anti-PPAR{gamma}2 Ab and L-NG-nitroarginine methyl ester (L-NAME) were from Alexis (Grünberg, Germany). The polyclonal anti-PPAR{alpha} and anti-c-Jun Abs were obtained from Santa Cruz Biotechnology (Heidelberg, Germany). Culture supplements and FCS were ordered from Biochrom (Berlin, Germany). 15d-PGJ2 and ciglitazone were bought from Biomol (Hamburg, Germany). Murine rIFN-{gamma} was from Roche Diagnostics (Mannheim, Germany). The luciferase assay kit was obtained from Promega (Mannheim, Germany) and the {beta}-galactosidase detection kit was from Tropix (Mannheim, Germany). Oligonucleotides were ordered from Eurogentec (Seraing, Belgium). All other chemicals were of the highest grade of purity and commercially available.

GSNO synthesis

S-nitrosoglutathione (GSNO) was synthesized and characterized as previously described (18).

Cell culture

The mouse monocyte/macrophage cell line RAW 264.7 and the premonocytic human cell lines U937 and THP1 were maintained in RPMI 1640 supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% heat-inactivated FCS (complete RPMI). All experiments were performed using complete RPMI.

Flow cytometry of oxygen radical production (HE assay)

Cells were cultured under nonadherent conditions. Following a prestimulation regime, 5 x 105 cells were incubated for 30 min with 1 µM PMA. Thereafter, 3 µM HE was added and incubations went on for 30 min. Cells were harvested, washed with PBS, and resuspended in 200 µl PBS. Flow cytometry analysis was performed using a Coulter Epics XL flow cytometer (Beckman Coulter, Krefeld, Germany) and HE was measured through a 630-nm long pass filter (FL3). Data from 10,000 cells were collected to reach significance.

Nuclear protein extraction

Preparation of crude nuclear extract was basically as described (19). Briefly, following cell activation for the times indicated, 4 x 106 cells were washed in 1 ml of ice-cold PBS, centrifuged at 1,000 x g for 5 min, resuspended in 400 µl ice-cold hypotonic buffer (10 mM HEPES/KOH, 2 mM MgCl2, 0.1 mM EDTA, 10 mM KCl, 1 mM DTT, 0.5 mM PMSF, pH 7.9), left on ice for 10 min, vortexed, and centrifuged at 15,000 x g for 30 s. Sedimented nuclei were resuspended in 50 µl ice-cold saline buffer (50 mM HEPES/KOH, 50 mM KCl, 300 mM NaCl, 0.1 mM EDTA, 10% glycerol, 1 mM DTT, 0.5 mM PMSF, pH 7.9), left on ice for 20 min, vortexed, and centrifuged at 15,000 x g for 5 min at 4°C. Aliquots of the supernatant, containing nuclear proteins, were frozen in liquid nitrogen and stored at -70°C. Protein was determined using a Bio-Rad II kit (Bio-Rad, Hercules, CA).

EMSAs

An established EMSA method, with slight modifications, was used (20). Nuclear protein (20 µg) was incubated for 20 min at room temperature with 20 µg BSA, 2 µg poly(dI-dC) (from Amersham Biosciences, Freiburg, Germany), 2 µl buffer D (20 mM HEPES/KOH, 20% glycerol, 100 mM KCl, 0.5 mM EDTA, 0.25% Nonidet P-40, 2 mM DTT, 0.5 mM PMSF, pH 7.9), 4 µl buffer F (20% Ficoll-400, 100 mM HEPES/KOH, 300 mM KCl, 10 mM DTT, 0.5 mM PMSF, pH 7.9), and 20,000 cpm of a 32P-labeled oligonucleotide in a final volume of 20 µl. Supershift Abs (2 µg) were added as indicated. DNA-protein complexes were resolved at 180 V for 4 h in a taurine-buffered, native 6% polyacrylamide gel (4% for supershifts), dried, and visualized (autoradiography using a Fuji x-ray film). Oligonucleotide probes were labeled by a filling reaction using the Klenow fragment (Roche Diagnostics). A total of 1 pmol oligonucleotide was labeled with 50 µCi of [{alpha}-32P]dCTP (3,000 Ci/mmol; Amersham, Braunschweig, Germany) and cold nucleotides (dATP, dTTP, dGTP; Life Technologies, Eggenstein, Germany), purified on a CHROMA SPIN-10 column (Clontech Laboratories, Heidelberg, Germany), and stored at -20°C until use. Oligonucleotides with the consensus PPRE site (bold letters) were used (21): 5'-GGTAAAGGTCAAAGGTCAAT-3' and 3'-ATTTCCAGTTTCCAGTTAGCCG-5'.

PPRE reporter gene assay

The plasmid J3TK pGL3, containing three copies of the human apoAII gene promoter PPRE-containing J site cloned upstream of the thymidine kinase promoter in the pGL3 luciferase expression vector, was kindly provided by B. Staels (Institut National de la Santé et de la Recherche Médicale, Unité 325, Institute Pasteur, Lille, France) (22). Macrophages were transiently transfected using the DEAE-dextran method as previously described (23). Cell selection was unnecessary because the expression of a macrophage-unrelated protein was analyzed. Briefly, 1 day before transfection, cells were seeded in suspension at a density of 1 x 106 cells/ml. A total of 1 x 107 cells were harvested, washed twice with PBS, and incubated for 3 h at 37°C in 1 ml RPMI 1640 supplemented with 50 mM Tris-HCl (pH 7.3), 400 µg DEAE-dextran, 20 µg luciferase reporter construct (J3TK pGL3), and 5 µg CMV-{beta}-galactosidase plasmid as an internal control. To discard the DNA/DEAE-dextran mixture, cells were washed twice with PBS, seeded at a density of 1 x 106 cells/ml, and cultured for 24 h. Afterward cells were stimulated for 15 h with NO donors or 15d-PGJ2. Cell extracts were assayed for luciferase and {beta}-galactosidase activity. For calculations, luciferase activity was normalized for {beta}-galactosidase by using the following formula: luciferase activity/{beta}-gal activity.

RNA extraction and semiquantitative RT-PCR

RNA was extracted using peqGOLD RNAPure (Peqlab, Erlangen, Germany) according to the distributor’s manual. Reverse transcription reactions and PCR for murine and human PPAR{gamma} and GAPDH were performed using the Advantage RT-for-PCR kit and the Advantage 2 Polymerase Mix (Clontech Laboratories). The sequences of the primers were as follows: murine PPAR{gamma} 480–1228(480–1228) (24), TA = 63°C: 5'->3', ATGGCCATTGAGTGCCGAGTCTG; 3'->5', GGCTTTTGAGGAACTCCCTGGTCA. Murine GAPDH 47–1048(47–1048) (25), TA = 63°C: 5'->3', ATGGTGAAGGTCGGTGTGAACGG; 3'->5', TTACTCCTTGGAGGCCATGTAGGGC.

Annealing temperatures were calculated using the primer design program Oligo (MBI, St. Leon-Rot, Germany). The number of amplification cycles (25 for GAPDH and 30 for PPAR{gamma}) was necessary to achieve exponential amplification where product formation is proportional to starting DNA. Products were run on a 1% agarose gel and were ethidium bromide stained. Controls of isolated RNA omitting reverse transcription were used during PCR to guarantee genomic DNA-free RNA preparations (data not shown).

Decoy approach

Cells were exposed to an oligonucleotide containing a PPRE consensus site as specified for the EMSA. Cells were seeded at a density of 1 x 106 cells/well into six-well plates. Oligonucleotides (3 µM) were added 24 h before cell stimulation. Cell stimulation was performed as indicated. Oligonucleotide sequences were identical to those used for EMSA. To guarantee oligonucleotide stability within the cells, oligonucleotides containing a phosphorothioate backbone were applied. For control reasons oligonucleotides with a mutated PPRE site were used (mutated sites in bold letters): 5'-GGTAAAGAACAAAGAACAAT-3' and 3'-ATTTCTTGTTTCTTGTTAGCCG-5'.

Griess assay

To this end, nitrite, a stable end product of NO metabolism, was determined in the supernatant of RAW 264.7 cells. Cells were seeded in six-well plates at a density of 1 x 106 cells/well. After incubations for 24 h, cells were exposed to 100 µM GSNO, 3 µM ciglitazone, and 1 µM 15-dPGJ2, or remained as controls. After 2 h medium was changed and cells were directly stimulated with 1 µg/ml LPS in combination with 10 U/ml IFN-{gamma} for 15 h. The amount of nitrite was determined spectrophotometrically by the Griess assay (Promega, Heidelberg, Germany) according to the manufacturer’s instructions. Nitrite concentrations in the supernatants were calculated by comparison with standard concentrations of NaNO2 dissolved in culture medium. Unless otherwise stated, the reported values are the mean (±SD) of three separate experiments, each performed in triplicate.

Northern blot

Total RNA was extracted using peqGOLD RNAPure (Peqlab) according to the distributor’s manual. Twenty micrograms of total RNA was used for Northern blotting. The probes for p22phox and p47phox were generated using the human GenBank sequences. The used primer sequences were as follows: Human p22phox 31–605(31–605) (26), TA = 63°C: 5'->3', GGGGCAGATCGAGTGGGCC; 3'->5', CGTCGGTCACCGGGATGGG. Human p47phox 131–871(131–871) (27), TA = 63°C: 5'->3', TCTACCGGCGCTTCACCGAGA; 3'->5', CGTCTTGCCCCGACTTTTGCA.

Subsequently, cDNA was cloned into a pDrive cloning vector (Qiagen, Hilden, Germany). After EcoRI (Roche Diagnostics) restriction cleavage and gel extraction, probes were radiolabeled with [{alpha}-32P]dCTP using the Rediprime II random prime labeling system (Amersham). Blots were also probed with a 28S RNA probe to assess equal loading.

Statistical analysis

Each experiment was performed at least three times and statistical analysis was performed using the two-tailed Student t test. Otherwise representative data are shown.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Activation of PPAR{gamma} by NO-releasing compounds and iNOS induction

In a first set of experiments we evaluated activation of PPAR{gamma} by gel shift analysis. Experimentally, monocytes/macrophages were dose-dependently exposed to chemically distinct NO donors. As shown in Fig. 1GoA, GSNO stimulated binding of PPAR{gamma} to radioactively labeled oligonucleotides in U937 cells. Activation was pronounced at 100 µM, somewhat lower at 200 µM, and virtually absent at doses of 50 µM as well as concentrations at or above 500 µM GSNO. Controls show minor or no PPAR{gamma} activation. To exclude any cell type artifact we repeated these studies in THP1 cells (Fig. 1GoB). In THP1 monocytes GSNO, at a concentration of 100 µM, evoked PPAR{gamma} activation, with lower or higher doses being inactive.



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  		FIGURE 1. Activation of PPAR{gamma} in response to NO-releasing compounds. Activation of PPAR{gamma} was analyzed by EMSA as described in Materials and Methods. A, U937 cells were dose-dependently stimulated with GSNO (50–1000 µM) for 1 h. B, THP1 cells were stimulated with increasing concentrations of GSNO (50–1000 µM) for 1 h. For controls cell stimulation was omitted. C, Supershift analysis of the active PPAR complex was performed as described in Materials and Methods. RAW 264.7 macrophages were stimulated with 100 µM GSNO or 50 µM spermine-NO for 1 h. For supershift analysis a PPAR{gamma}2 Ab (lanes 2 and 6), a PPAR{alpha} Ab (lanes 3 and 7), or a c-Jun Ab (lanes 4 and 8) was included. Incubations without supershift Abs are shown in lanes 1 and 5. D, RAW 264.7 cells were stimulated for 15 h with a combination of 10 µg/ml LPS and 100 U/ml IFN-{gamma} with or without 1 mM L-NAME. For controls cell stimulation was omitted. Data are representative of three similar experiments.

 
In extending studies we demonstrated that a chemically distinct NO donor such as spermine-NO activated PPAR{gamma} as well, with the notion that 50 µM was the most effective concentration (data not shown). Similar results were obtained using the murine macrophage-like cell line RAW 264.7 (data not shown). It appeared that activation of PPAR{gamma} was achieved at low concentrations of NO donors that elicited neither necrosis nor an apoptotic phenotype.

We went on to verify the identity of the PPAR-oligonucleotide complex by supershift analysis (Fig. 1GoC). This was achieved by adding PPAR{gamma} vs PPAR{alpha} Abs to the binding assay followed by EMSA analysis. Specificity was further verified using an unrelated anti-c-Jun Ab in parallel. While the PPAR{gamma} antiserum shifted the protein-oligonucleotide complex to a higher m.w., the PPAR{alpha} Ab did not. The c-Jun Ab was ineffective as well. Conclusively, the {gamma}-isoform of PPARs is activated in response to NO.

The biological significance shown for NO donor-mediated effects was verified in experiments using RAW 264.7 macrophages (Fig. 1GoD). After LPS/IFN-{gamma} treatment, an established iNOS-inducing regimen in these cells, activation of PPAR{gamma} was clearly visible by EMSA analysis (Fig. 1GoD, lane 3). Interestingly, inhibition of iNOS by L-NAME markedly reduced activation of PPAR{gamma} (Fig. 1GoD, lane 2), thus pointing to the involvement of endogenously generated NO in PPAR{gamma} activation.

In the following set of experiments we looked for the time-dependent activation of PPAR{gamma} by exposing U937 cells to GSNO or spermine-NO (Fig. 2GoA) and RAW 264.7 macrophages to spermine-NO (Fig. 2GoB). With 100 µM GSNO we elicited an immediate onset of the PPAR{gamma} response after 30 min. The strongest activation was achieved after 1–2 h, with a decreased responsiveness after 4 h.



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  		FIGURE 2. Time-dependent activation of PPAR{gamma} by NO donors. Activation of PPAR{gamma} was analyzed by EMSA as described in Materials and Methods. A, U937 cells were stimulated with 100 µM GSNO or 50 µM spermine-NO for 30 min up to 4 h. B, RAW 264.7 macrophages were exposed to 50 µM spermine-NO for 30 min up to 4 h. C, U937 cells were stimulated with 100 µM GSNO or 3 µM ciglitazone for 1 h alone or in combination. For controls cell stimulation was omitted. Data are representative of three similar experiments.

 
Spermine-NO evoked PPAR{gamma} activation after 30 min and promoted the strongest response after 1 h, with a declining responsiveness toward the control level at 2 and 4 h as shown for U937 (Fig. 2GoA) and RAW 264.7 cells (Fig. 2GoB). To confirm specific PPAR{gamma} activation we conducted costimulation experiments with the PPAR{gamma} agonists ciglitazone and 15d-PGJ2 in combination with the NO donor GSNO. As shown for U937 cells in Fig. 2GoC, ciglitazone, used at a concentration of 3 µM, itself led to PPAR{gamma} activation. This was significantly increased with the simultaneous addition of 100 µM GSNO. Similar results were obtained using 15d-PGJ2 in combination with GSNO (data not shown).

A well-established signal transduction pathway for NO is activation of the soluble guanylyl cyclase with the formation and action of cGMP (28, 29).

To analyze for the potential involvement of cGMP in conferring activation of PPAR{gamma} in our system, we exposed THP1 as well as RAW 264.7 cells to the lipophilic cyclic GMP analog 8-bromo-cGMP (1 mM) for 30 min up to 4 h (Fig. 3Go). Unlike NO, 8-bromo-cGMP did not reproduce PPAR{gamma} activation, thus pointing to a cGMP-independent NO signaling mechanism.



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  		FIGURE 3. Lipophilic cGMP analogs do not achieve PPAR{gamma} activation. A, THP1 cells were stimulated with 50 µM spermine-NO for 1 h or 1 mM 8-bromo-cGMP for 30 min up to 4 h. B, RAW 264.7 macrophages were exposed to 100 µM GSNO for 1 h or 1 mM 8-bromo-cGMP for 30 min up to 4 h. Activation of PPAR{gamma} was followed by EMSA as described in Materials and Methods. Data are representative of three similar experiments.

 
In continuation of the study we asked whether NO promoted PPAR{gamma} transactivation, besides enhancing DNA binding. We used the luciferase reporter assay to demonstrate transcriptional activation of PPAR{gamma} in response to NO donors and 15d-PGJ2, which represents a commonly accepted control activator (Fig. 4Go). At most effective concentrations GSNO and spermine-NO evoked a 2-fold activation in RAW 264.7 cells, while 15d-PGJ2 elicited an ~4-fold increase of luciferase activity. The response toward NO and 15d-PGJ2 was determined 15 h after cell stimulation.



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  		FIGURE 4. NO-evoked PPAR{gamma} reporter gene activation. RAW 264.7 macrophages were cotransfected with the J3TK-plasmid and a plasmid encoding {beta}-galactosidase. Luciferase and {beta}-galactosidase expression were analyzed after both activities had been normalized as described in Materials and Methods. Cell stimulation was for 15 h with 100 µM GSNO, 50 µM spermine-NO, 1 µM 15d-PGJ2, or vehicle (control). Data are means ± SD of three individual experiments (*, p <= 0.05 vs control).

 
We concluded that NO stimulated binding of PPAR{gamma} to the DNA and concomitantly caused transactivation of a PPAR-responsive reporter system.

Specificity of the NO response

Considering the possibility that PPAR{gamma} is transcriptionally regulated by NO we analyzed the mRNA level by RT-PCR in RAW 264.7 macrophages (Fig. 5Go). Neither GSNO (100 µM) nor spermine-NO (50 µM), both supplied up to 4 h, revealed any variations in the mRNA content of PPAR{gamma} after normalization with the GAPDH standard.



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  		FIGURE 5. mRNA expression of PPAR{gamma} in response to NO donors. Expression of PPAR{gamma} mRNA was analyzed by RT-PCR. RAW 264.7 cells were stimulated with 100 µM GSNO or 50 µM spermine-NO for the times indicated. Semiquantitative RT-PCR was performed as described in Materials and Methods. Data are representative of three similar experiments.

 
Excluding a transcriptional regulation of PPAR{gamma} in response to NO delivery, we assumed a direct activation mechanism to be more likely. Along that line we asked whether antioxidants such as NAC may interfere with PPAR{gamma} activation. This was tested by preincubating RAW 264.7 cells for 1 h with 1 mM NAC before the addition of 100 µM GSNO or 50 µM spermine-NO, supplied for 1 h (Fig. 6GoA). The efficacy of NAC was demonstrated in gel shift assays, when activation of PPAR{gamma} by GSNO and spermine-NO was largely attenuated in the presence of the antioxidant. Moreover, NAC suppressed the transactivation capacity of PPAR{gamma} (Fig. 6GoB). For these experiments 1 mM NAC was supplied for 1 h in front of NO donors, followed by determination of luciferase activity 15 h later.



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  		FIGURE 6. NAC and PTIO attenuate PPAR{gamma} activation in response to NO. A, RAW 264.7 macrophages were incubated for 1 h with 1 mM NAC before the addition of 100 µM GSNO or 50 µM spermine-NO. EMSA was performed after 1 h as described in Materials and Methods. For controls cell stimulation was omitted. Data are representative of three similar experiments. B, RAW 264.7 macrophages were cotransfected with the J3TK plasmid and a plasmid encoding {beta}-galactosidase. Luciferase and {beta}-galactosidase expression were analyzed after both activities had been normalized as described in Materials and Methods. As indicated, cells were prestimulated for 1 h with 1 mM NAC or 100 µM PTIO, or remained as controls, before the addition of 100 µM GSNO, 50 µM spermine-NO, or vehicle (control) for 15 h. Data are means ± SD of three individual experiments (*, p <= 0.05 vs untreated sample; **, p <= 0.05 vs GSNO-treated sample; ***, p <= 0.05 vs spermine-NO-treated sample).

 
The 2-fold activation, normally seen with NO donors, was completely absent with NAC being present, implying that oxidative or nitrosative modifications are involved in PPAR{gamma} activation. It was our further intention to pinpoint NO as the molecule setting into motion the activation cascade that culminated in PPAR{gamma} activation. Therefore, we used the NO scavenger PTIO (Fig. 6GoB). The NO scavenger, preincubated at a dose of 100 µM for 1 h, completely attenuated PPAR{gamma}-evoked luciferase activity in response to GSNO and spermine-NO. Control determinations showed no interference of PTIO or NAC on the basal PPAR{gamma} response.

NO attenuated the oxidative burst in macrophages via activation of PPAR{gamma}

Decoy oligonucleotides can be used to scavenge and thereby to inactivate transcription factors (30, 31). Using this experimental approach we provided evidence that NO donors attenuated reactive oxygen species (ROS) formation via PPAR{gamma} activation (Fig. 7Go). As shown in Fig. 7GoA, oxidation of HE was elicited in response to 1 µM PMA in RAW 264.7 macrophages. This was measured by flow cytometry and is shown by the rightward shift of the HE signal (Fig. 7GoA). Prestimulation with 100 µM GSNO (Fig. 7GoB) or 50 µM spermine-NO (Fig. 7GoC) for 15 h eradicated ROS formation because NO donors attenuated the HE shift in response to PMA.



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  		FIGURE 7. Oxygen radical production in macrophages under the influence of NO and PPAR{gamma}. ROS production was analyzed in RAW 264.7 macrophages by flow cytometry using 3 µM HE as the redox-sensitive dye as described in Materials and Methods. Cell stimulation was achieved with 1 µM PMA and data are representative of three or more similar experiments. Manipulations are as follows: A, controls (gray) vs PMA stimulation (black); B, PMA stimulation (black) vs PMA with 100 µM GSNO being preincubated for 15 h (gray); C, PMA stimulation (black) vs PMA with 50 µM spermine-NO being preincubated for 15 h (gray); D, PMA response (black) vs a treatment with PPRE decoy oligonucleotides, 100 µM GSNO, and PMA as described in the text (gray); E, PMA treatment (black) vs PMA with 100 µM PTIO and 100 µM GSNO being preincubated for 16 and 15 h (gray); F, PMA response (black) vs PMA with 1 mM 8-bromo-cGMP being preincubated for 15 h (gray).

 
The presence of PPRE decoy oligonucleotides attenuated the down-modulatory behavior of GSNO and allowed to regain HE oxidation after PMA addition (Fig. 7GoD). The soft shoulder seen in the trace from PPRE decoy oligonucleotides/GSNO/PMA-treated cells may refer to a population of macrophages that have not been reached by oligonucleotides, therefore still attenuating ROS formation. The specificity for NO in diminishing the oxidative burst was further demonstrated when coincubations of PTIO, GSNO, and PMA allowed ROS generation (Fig. 7GoE). In line with the observation that cGMP does not activate PPAR{gamma} (Fig. 3Go), prestimulation with 1 mM 8-bromo-cGMP turned out to be ineffective in inhibiting ROS formation following PMA addition (Fig. 7GoF).

We conclude that NO, delivered by the breakdown of GSNO or spermine-NO, activates PPAR{gamma} by a cGMP-independent mechanism that in turn attenuates the proinflammatory signal of the macrophage oxidative burst.

NO-mediated inhibition of superoxide radical production is due to PPAR{gamma}-dependent down-regulation of p47phox

Modulation of gene expression in response to PPAR{gamma} activation is known and associated with, e.g., inhibition of iNOS expression (13). To verify our system we used RAW 264.7 macrophages, known to express iNOS in response to LPS/IFN-{gamma} treatment. To establish that NO-activated PPAR{gamma} may attenuate iNOS expression, we treated cells for 2 h with 100 µM GSNO, changed medium, and stimulated macrophages with 1 µg/ml LPS in combination with 10 U/ml IFN-{gamma} for 15 h. Thereafter nitrite was determined by the Griess assay as the metabolic end-product of iNOS-generated NO. Prestimulation of the cells with GSNO attenuated nitrite production by 50% compared with cells treated with LPS/IFN-{gamma} only (27 ± 9 vs 14 ± 7 µM nitrite). To make the contribution of PPAR{gamma} more likely, PPAR{gamma} agonists such as 15d-PGJ2 or ciglitazone were used. Prestimulation for 2 h with either 15d-PGJ2 or ciglitazone decreased LPS/IFN-{gamma}-mediated NO production by ~70% (27 ± 9 vs 9 ± 5 µM nitrite for 15d-PGJ2 or 11 ± 7 µM nitrite for ciglitazone).

We now focused on components of the NAD(P)H oxidase system, known to be involved in the superoxide radical formation, to identify the molecular mechanism responsible for attenuating the oxidative burst. We analyzed the expression pattern of different members of this multifactor complex on RNA levels by Northern blotting. Using this experimental system we demonstrate as depicted for U937 cells that NO-mediated PPAR{gamma} activation leads to p47phox down-regulation. As shown in Fig. 8Go, upper panel, transcription of p47phox was clearly reduced in response to 50 and 100 µM GSNO. To verify the role of PPAR{gamma} activation concerning this effect, we applied 15d-PGJ2 and ciglitazone, two known specific PPAR{gamma} agonists, to the cells with a similar outcome. Expression of p47phox was significantly diminished. p22phox, another component of the NAD(P)H oxidase system, was not altered by NO exposure or by PPAR{gamma} agonist addition compared with untreated controls (Fig. 8Go, lower panel). To assess equal loading, filters were hybridized with a 28S RNA probe (data not shown).



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  		FIGURE 8. Down-regulation of p47phox in response to NO-dependent PPAR{gamma} activation. Regulation of p22phox and p47phox expression was analyzed on total RNA level by Northern blotting as described in Materials and Methods. Cells were treated with 50 or 100 µM GSNO, 1 µM 15d-PGJ2, or 3 µM ciglitazone for 24 h or remained as controls. The blots were hybridized first with a probe specific for p47phox (upper panel) and second with one specific for p22phox (lower panel). Data are representative of three similar experiments.

 
We assume that NO-dependent PPAR{gamma} activation leads to attenuation of at least p47phox expression, one necessary component of the NAD(P)H oxidase, responsible for oxidative burst generation.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this work we presented evidence that NO-releasing compounds activated the anti-inflammatory properties of the transcription factor PPAR{gamma} and concomitantly attenuated the oxidative burst in monocytes/macrophages, likely due to down-regulated p47phox expression. Activation of PPAR{gamma} was confirmed by gel shift analysis, reporter gene assays, and a decoy oligonucleotide approach. The involvement of NO was substantiated by using the NO scavenger PTIO and the antioxidant NAC with the notion that lipophilic cGMP analogs were unable to reduce superoxide formation. Attenuation of p47phox expression was shown by Northern blotting. To verify the biological significance of these results obtained with NO donors, RAW 264.7 macrophages were stimulated with LPS/IFN-{gamma} to endogenously produce NO as a result of iNOS induction. As shown by gel shift analysis, PPAR{gamma} was activated in response to endogenously synthesized NO, because blocking iNOS significantly attenuated PPAR{gamma} activation. We conclude that iNOS-generated NO suffices in provoking DNA binding of PPAR{gamma}. Taking the lipid-soluble nature of NO into account, it appears attractive to hypothesize that cells in the direct neighborhood to NO-producing cells are affected with the outcome of an impaired proinflammatory signaling cascade in those target cells.

Reactive oxygen- and nitrogen-derived species are implicated as effector molecules in the immune system, serving major functions during immunological host defense, mainly as a result of macrophage and neutrophil activation (32). However, reactive oxygen and nitrogen species operate as modulators of signal transducing pathways as well, thus characterizing them as intra- and intercellular messenger molecules (33). In line, redox-controlled transcription factors, oxidative susceptible thiol groups, or redox-sensitive phosphorylation events allow to channel the action of reactive species into established intracellular communication systems (2, 34).

The observation that NO-provoked PPAR{gamma}-activation is unrelated to cGMP signaling appears in line with several recent reports on direct NO/target interactions. For example, activation of c-Ha-Ras (p21) via nitrosylation of cysteine 118 has recently been shown by using electrospray ionization mass spectrometry (34), up-regulation of transcription factors such as NF-{kappa}B has been noted in a NO-dependent manner in a murine model of hemorrhagic shock (6), and protein tyrosine phosphorylation is achieved by NO. Moreover, the transcription factor HIF1{alpha} turned out to be NO responsive (35), and NO is needed to activate tyrosine kinase 2 and to tyrosine phosphorylate STAT4 (36). In this work we show, by using chemically distinct NO donors, that PPAR{gamma} is activated by NO. This is substantiated by using a NO scavenger to attenuate PPAR{gamma} activation. The use of NAC to abrogate the NO signal may aim toward its NO scavenging ability or its antioxidant properties. In any case, the interference of NAC appears rational. Although our studies excluded cGMP to activate PPAR{gamma}, molecular details on NO action remain unknown so far. In this respect, extending examinations are needed to address various possibilities, such as a direct interaction of NO with PPAR{gamma}, phosphorylation events, or NO-evoked generation of a PPAR{gamma} activator. Of note, DNA binding of PPAR{gamma} appeared fast and was noticed 1–2 h after the addition of NO. The rapid response points to activation of a preformed transcription factor rather than a mechanism involving enhanced PPAR{gamma} expression. This assumption is strengthened by our observation that mRNA of PPAR{gamma} remained unchanged in response to NO.

Activation of PPAR{gamma} by NO is contrasted by inhibition seen at higher concentrations of NO donors. An attenuated DNA binding ability of PPAR{gamma} at higher doses of GSNO appears in some agreement with studies of Kröncke et al. (37). They observed reduced DNA binding activities of zinc finger transcription factors, among them the RXR, at high doses of NO donors (>=0.5 mM). Taking into consideration that PPAR{gamma} heterodimerizes with RXR before DNA binding, one may envision an attenuated promoter binding activity of PPAR{gamma} at elevated NO concentrations. The importance of RXR in transcription factor complex activation/inhibition will be an important issue of future experiments. As seen in our study, at concentrations >200 µM GSNO we lost the ability of NO to stimulate DNA binding or to cause transactivation of PPAR{gamma} (data not shown).

Activation of PPAR{gamma} by NO results in desensitization of macrophages with the consequence of an attenuated oxidative burst. A cause-effect relationship is established by the successful use of a decoy oligonucleotide approach that allowed to regain an oxidative signal despite the presence of NO. Decoy oligonucleotide approaches have been shown to block transcriptional activity with high efficacy by scavenging active transcription factors (31). PPAR{gamma} is known for its anti-inflammatory properties and is reported to exert a negative effect on proinflammatory cytokine and/or iNOS expression in macrophages, such as RAW cells (12, 13). The identified anti-inflammatory property of PPAR{gamma} is in contrast to the work of Chawla et al. (16), who showed in macrophages derived from murine embryonic stem cells that were homozygous for a null mutation in the PPAR{gamma} gene, that inhibitory effects on cytokine production and inflammation may be receptor independent. Concerning this data we assume a concentration-dependent, PPAR{gamma}-independent effect, which might not occur using lower doses of the applied PPAR{gamma}-specific and unspecific agonists.

On a molecular basis PPAR{gamma} has recently been shown to antagonize coactivators such as the CREB-binding protein from interacting with its cognate target gene, thereby attenuating up-regulation of, e.g., iNOS (13), or blocking TNF-{alpha} formation (12). Having these reports in mind, our decoy approach may imply either that transactivation of PPAR{gamma} with concomitant formation of a PPAR{gamma}-responsive gene product interferes with the oxidative burst or that active PPAR{gamma} complexes and thereby removes another factor that presumably is required to express constituents of the NAD(P)H oxidase. In any case, PPAR{gamma} decoy oligonucleotides will interfere with gene expression with the limitation that unknown transcription factors/coactivators, distinct from PPAR{gamma}, may be targeted by this approach. Taking advantage of published data that iNOS expression is inhibited by PPAR{gamma} activation (13), we provided evidence that our system is in line with published results. Prestimulation for 2 h with GSNO inhibits LPS/IFN-{gamma}-mediated nitrite production by ~50%. However, complete inhibition was not achieved and control experiments performed with specific PPAR{gamma} agonists revealed ~70% inhibition. This may depend on the presence of IFN-{gamma}, which antagonizes PPAR{gamma}-provoked inhibitory mechanisms (38). Having demonstrated that PPAR{gamma} targets established systems, e.g., NO formation, we provided further evidence on PPAR{gamma} in attenuating ROS formation. Results came from Northern blotting experiments, showing inhibition of p47phox expression in response to NO-releasing compounds as well as PPAR{gamma}-specific agonists; we assume that attenuated p47phox expression as one component of the NAD(P)H oxidase system accounts for reduced ROS formation (39). These observations are in some analogy with the results of Inoue et al. (40), demonstrating inhibition of p22phox expression in response to PPAR{gamma} agonists in primary endothelial cells. Regulation of NAD(P)H oxidase components has been described in human cultured monocytes, where decreased gp91phox, p22phox, and p47phox expression in response to diminished binding of the transcription factor PU.1 to the corresponding promoter sites reduced the ability to produce microbicidal oxidants (41). Endogenous production of NO seems insufficient to down-regulate p47phox (data not shown). Rather, up-regulation of p47phox upon LPS/IFN-{gamma} stimulation was shown (42). PPAR{gamma}-mediated effects are counter-regulated by IFN-{gamma} (38). Further studies will need to analyze the involvement of p47phox promoter regions in conferring this opposing effect of LPS/IFN-{gamma} vs NO.

Inhibition of the oxidative burst by NO has been known for some time. In neutrophils NO blocked O2- formation with some indication that assembly of the NAD(P)H oxidase is affected (43). Along that line, NO attenuated the oxidative burst in murine microglia as well (44). In analogy to our studies, this was unrelated to cGMP signaling or a simple scavenging of O2-, thus implying activation of PPAR{gamma} as a rational explanation for these observations.

Macrophages are key players during the innate immune response. Immunological activation of macrophages is achieved by cytokines and bacterial components (45). Cell activation results in the release of various proinflammatory cytokines and reactive nitrogen as well as oxygen species (46). The excessive release of these mediators results in the development of whole body inflammation, which is closely related to the clinical symptoms of sepsis or septic shock (47). During sepsis the early hyperinflammatory phase is counterbalanced by an anti-inflammatory response, characterized by monocyte deactivation (48). It appears attractive to hypothesize whether the formation of NO in macrophages not only represents an early cytotoxic signal but also shifts the balance toward an anti-inflammatory response via activation of PPAR{gamma}, which would be in line with recent reports describing NO as an inhibitor of proinflammatory cytokine expression (7, 49, 50). In further studies it will be essential to compare the cytokine profile of activated macrophages to that of a deactivated macrophage phenotype, to study the impact of NO on this cytokine balance, and to elucidate the role of PPAR{gamma} during this activation-deactivation transition.


   Acknowledgments
 
We thank Sabine Häckel for expert technical assistance.


   Footnotes
 
1 This work was supported by grants of the Deutsche Forschungsgemeinschaft (SFB 263 TP B10, Br 999, and Deutsche Krebshilfe). Back

2 Address correspondence and reprint requests to Dr. Bernhard Brüne, Institute of Cell Biology, University of Kaiserslautern, Erwin-Schroedinger-Strasse, 67663 Kaiserslautern, Germany. E-mail address: bruene{at}rhrk.uni-kl.de Back

3 Abbreviations used in this paper: PPAR, peroxisome proliferator-activated receptor; p47phox, p47 phagocyte oxidase; GSNO, S-nitrosoglutathione; spermine-NO, spermine-NONOate; PPRE, PPAR response element; NAC, N-acetyl-cysteine; PTIO, 2-phenyl-4,4,5,6-tetramethyl-imidazoline-1-oxyl 3-oxide; L-NAME, L-NG-nitroarginine methyl ester; 15d-PGJ2, 15-deoxy-{Delta}12,14-PGJ2; HE, hydroethidine; RXR, retinoic acid receptor; iNOS, inducible NO synthase; ROS, reactive oxygen species. Back

Received for publication February 1, 2002. Accepted for publication June 28, 2002.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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