This Article Abstract Full Text (PDF) Other Versions of this Article: MCB.00623-07v1 27/23/8374    most recent Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Ravaux, L. Articles by El Hadri, K. Search for Related Content PubMed PubMed Citation Articles by Ravaux, L. Articles by El Hadri, K.

 Previous Article  |  Next Article 

Molecular and Cellular Biology, December 2007, p. 8374-8387, Vol. 27, No. 23
0270-7306/07/$08.00+0     doi:10.1128/MCB.00623-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Inhibition of Interleukin-1ß-Induced Group IIA Secretory Phospholipase A2 Expression by Peroxisome Proliferator-Activated Receptors (PPARs) in Rat Vascular Smooth Muscle Cells: Cooperation between PPARß and the Proto-Oncogene BCL-6

UMR Physiologie et Physiopathologie, Université Pierre et Marie Curie, CNRS, 7 quai Saint-Bernard, 75252 Paris, France

Received 10 April 2007/ Returned for modification 14 May 2007/ Accepted 18 September 2007

	ABSTRACT

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The inflammation that occurs during atherosclerosis is characterized by the release of large amounts of group IIA secretory phospholipase A2 (sPLA2-IIA). This study was designed to define the function of the three peroxisome proliferator-activated receptors (PPARs) on sPLA2 expression in vascular smooth muscle cells (VSMCs). We found that PPAR ligands decreased sPLA2-IIA activity and inhibited mRNA accumulation under inflammatory conditions. Furthermore, interleukin-1ß-induced sPLA2-IIA promoter activity was inhibited by the three PPAR ligands and in a similar way when cells were cotransfected with PPAR, PPARß, or PPAR, plus retinoid X receptor (RXR). Our study revealed that the regulation of sPLA2-IIA gene transcription by PPAR/RXR and PPAR/RXR heterodimers requires an interaction with a PPAR response element (PPRE) of the sPLA2-IIA promoter. In contrast, PPARß operates through a PPRE-independent mechanism. In addition, we demonstrated that VSMCs expressed the transcriptional repressor BCL-6. Overexpression of BCL-6 markedly reduced sPLA2-IIA promoter activity in VSMCs, while a dominant negative form of BCL-6 abrogated sPLA2 repression by PPARß. The PPARß agonist induced a BCL-6 binding to the sPLA2 promoter in VSMCs under inflammatory conditions. The knockdown of BCL-6 by short interfering RNA abolished the inhibitory effect of the PPARß ligand on sPLA2 activity and prostaglandin E₂ release. Thus, the inhibition of sPLA2-IIA activity by PPARß agonists may provide a promising approach to impacting the initiation and progression of atherosclerosis.

	INTRODUCTION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The large family of phospholipase A2 (PLA2) enzymes hydrolyze ester bonds at the sn-2 position of glyceroacylphospholipids to produce lysophospholipids and nonesterified fatty acids such as arachidonic acid (C_20:4 [n-6]), the precursor of various proinflammatory lipid mediators, such as eicosanoids (37). To date there are four major families of PLA2. Several observations argue for the implication of type IIA secretory PLA2 (sPLA2-IIA) in the pathogenesis of various inflammatory diseases, including cancer, septic shock, rheumatoid arthritis, and atherosclerosis (38). The plasma of patients with various inflammatory diseases, particularly atherosclerosis, contains high concentrations of sPLA2-IIA (18). The synthesis of sPLA2-IIA is stimulated by inflammatory cytokines, such as interleukin-1ß (IL-1ß), IL-6, and tumor necrosis factor alpha (TNF-) (2, 20). Moreover, sPLA2-IIA is largely expressed in vascular smooth muscle cells (VSMCs) (18) in response to proinflammatory cytokines produced by infiltrated macrophages. Overproduction of human sPLA2-IIA in transgenic mice contributes to atherogenesis (19). This enzyme catalyzes the production of precursors of lipid mediators (mainly prostaglandin E₂ [PGE2]), which in turn amplify the effect of cytokines on the dedifferentiation of VSMCs (11). The enzyme was reported to have other proatherogenic properties linked to its ability to hydrolyze phospholipid monolayers of high- and low-density lipoproteins (21). Previously, our laboratory demonstrated the interplay of various transcription factors (NF-B, C/EBPß, ETS, and liver X receptor) that bind on the proximal part of the sPLA2 promoter and, more recently, has provided evidence that sPLA2-IIA could stimulate its own production in VSMCs through positive feedback (2, 3, 22). Then a better understanding of the molecular mechanisms involved in the regulation of sPLA2-IIA gene expression, in an inflammatory context, may allow the development of new inhibitors.

The peroxisome proliferator-activated receptor (PPAR) family includes three members: PPAR, PPARß, and PPAR. These ligand-activated transcription factors, belonging to the nuclear receptor superfamily, form heterodimers with the retinoid X receptor (RXR) to regulate the expression of genes involved in lipid metabolism, glucose metabolism, and inflammation (13, 25, 34). The relevance of PPAR pathways to metabolic diseases is underscored by the use of fibrates (PPAR agonists) and thiazolidinediones (PPAR agonists) to treat hyperlipidemia and hyperglycemia, respectively. Agonists of PPAR have positive effects on lipid metabolism both in animal models and in clinical practice. Indeed, agonists of PPAR, the thiazolidinediones rosiglitazone and pioglitazone, improve insulin resistance in type 2 diabetes and pioglitazone improves the dyslipidemia associated with insulin resistance. In addition to these effects, both PPAR and PPAR agonists have anti-inflammatory properties that can provide additional cardiovascular benefits (5). The role of PPARß is less well understood. In keeping with its ubiquitous expression, PPARß has been implicated in cellular proliferation and differentiation, lipid metabolism, and inflammation (4). Synthetic PPARß agonists promote cholesterol uptake (29) and efflux (41) in macrophages. In muscle, the overexpression or activation of PPARß by synthetic ligands induces lipid utilization (36, 42), while in the liver, PPARß suppresses hepatic glucose output (25). For the heart, the protective role of PPARß has been confirmed by in vitro studies showing that PPARß agonists attenuate phenylephrine-induced cardiac hypertrophy (30). PPARß has been suggested to control an inflammatory switch during atherogenesis (25). However, despite significant advancement in the search for the potential roles of PPARß in atherogenic inflammation, responses remain obscure.

Because sPLA2-IIA is thought to exert crucial proinflammatory functions in VSMCs during atherosclerosis by hydrolyzing cell membrane phospholipids into free fatty acids, the major source of eicosanoids (mainly PGE2), we investigated the effects of different PPAR activators on the regulation of the sPLA2-IIA gene expression. We demonstrate here that PPAR, -, and -ß significantly inhibited cytokine-stimulated sPLA2-IIA expression and secretion in VSMCs. While effects of PPAR and - are exerted through a PPAR response element (PPRE)-dependent mechanism, the anti-inflammatory effect of PPARß required the proto-oncogene BCL-6 in a PPRE-independent manner.

	MATERIALS AND METHODS

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Isolation and culture of VSMCs from rat aortas. VSMCs were isolated from the thoracic aortas of adult male Wistar rats as described previously (2). Cells were seeded on dishes coated with type I collagen from calf skin (Sigma) and cultured in Dulbecco's modified Eagle's medium supplemented with 10% (vol/vol) fetal calf serum (Gibco BRL, Cergy-Pontoise, France), 4 mM glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin. VSMCs were subcultured every 5 days, and experiments were performed on cells between passages 3 and 7. For the experiments, confluent cells were made quiescent by incubating them for 24 h in serum-free medium containing 0.2% fatty-acid-free bovine serum albumin before they were treated with the appropriate agents. The culture medium was then removed, and measurements of for sPLA2 activity were taken, and cells were lysed for total RNA, total proteins, or nuclear extracts preparation.

PLA2 activity. sPLA2 activity was measured using the fluorescent substrate 1-hexadecanoyl-2-(1-pyrenyldecanoyl)-sn-glycero-3-phosphoglycerol (Interchim, France) as described previously (3). Total hydrolysis of the substrate obtained by 0.1 unit of PLA2 from bee venom (Sigma) was used as a reference to calculate the PLA2 activity in the samples. Spontaneous substrate hydrolysis was assayed in fresh culture medium and subtracted from each sample value.

RNA extraction and RT-PCR analysis. Total RNA was extracted from rat VSMCs by using RNeasy kit columns (QIAGEN, Courtaboeuf, France) according to the supplier's instructions. RNA (1 µg) was reverse transcribed for 1 h at 37°C with 200 U of mouse mammary lentivirus-reverse transcriptase (RT) (Invitrogen), 100 µM random primers, and the buffer supplied by the manufacturer in a total volume of 20 µl. The reaction was terminated by heating to 95°C for 5 min. To ensure that subsequent amplification was not derived from contaminant genomic DNA, a control without mammary lentivirus-RT was included in parallel for each RNA sample. Reverse-transcribed mRNAs were amplified in a thermocycler (Hybaid Omnigene; Syngene, Ozyme, France) as described previously (3). The primer sequences used for the different tested genes are given in Table 1. PCR amplifications were performed, and PCR products were size separated by electrophoresis on 2% (wt/vol) agarose gel and visualized by ethidium bromide staining. PCR bands were quantified using the GeneGenius system (Syngene, Ozyme, France).

Western blot analysis. VSMCs were harvested and homogenized in a lysis buffer (50 mM Tris-HCl [pH 7.5], 1% Triton, 150 mM NaCl, 1 mM EDTA, 1 mM dithiothreitol, 1 mM Na₃VO₄, 0.5 mM phenylmethylsulfonyl fluoride, 50 mM NaF, 5 mM PPi, 10% glycerol) supplemented with a cocktail of antiprotease (Complete; Roche). Homogenates were centrifuged at 13,000 x g for 10 min at 4°C. The resulting supernatants were stored at –20°C until used. Proteins (20 to 40 µg/lane) were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (10% gel) and electroblotted onto al 0.45-µm-pore-size polyvinylidene difluoride membrane (Immobilon-P; Millipore). After we determined the efficiency of protein transfer and well-to-well variability with Ponceau S (Sigma-Aldrich), the membrane was incubated overnight at 4°C with a PPARß or BCL-6 rabbit polyclonal antibody (Santa Cruz Biotechnology, Inc.) at a dilution of 1:200 or 1:1,000, respectively, in 2% milk-Tris-buffered saline (TBS) with 0.1% Tween 20 (Sigma-Aldrich). The next day, the membrane was washed in TBS with 0.1% Tween before adding anti-rabbit immunoglobulin G (IgG)-horseradish peroxidase at a dilution of 1:1,000 in 5% milk-TBS with 0.1% Tween and then incubated for 1 h at room temperature. The detection of immune complex was performed using an enhanced chemiluminescence detection kit for Western blotting (Amersham) on BioMax MR Kodak film (Sigma-Aldrich).

Plasmid constructions and transfection. The [–1153; +46]sPLA2-Luc construct was obtained by PCR amplification of –1153 to +46 bp of the sPLA2 promoter. The cloning of the rat sPLA2 promoter (–488 to +46 bp) into the pGL3-basic luciferase plasmid to create [–488; +46]sPLA2-Luc has been described previously (2). The mutant PPRE-sPLA2 construct (mPPRE-sPLA2) lacks the PPAR-binding element of the sPLA2 promoter, the mutant BCL-6-sPLA2 construct (mBCL-6-sPLA2) lacks the BCL-6-binding element, and the double-mutated BCL6-PPRE-sPLA2 construct (mBCL6-mPPRE-sPLA2) lacks both PPAR and BCL-6 binding sites. The sites were replaced with the BglII restriction site by using PCR-based, site-directed mutagenesis. VSMCs were seeded, 24 h before transfection, in 24-well plates at a concentration of 20,000 cells per plate, and at 70% confluence, cells were transfected using 1.5 ml of Lipofectamine Plus (Invitrogen), 0.4 µg of reporter DNA, and 0.1 µg of pCMV-ß-galactosidase per well. For transactivation studies, 10 ng of either pcDNA3.1 PPAR, PPARß, or PPAR expression vectors and 10 ng of CMX-RXR were added. The cells were refed with Dulbecco's modified Eagle's medium containing 0.2% bovine serum albumin 3 h after we added the DNA and incubated for 24 h. Then, indicated concentrations of PPAR agonist (WY14643), PPARß agonist (L165041), or PPAR agonist (GW1929) were added and incubation continued for a further 24 h. The plasmid construct containing PPRE-3-TK-LUC (luciferase gene under control of the herpes simplex virus thymidine kinase promoter and three PPRRs was used as a control of PPAR activation. Luciferase activity was measured using a luciferase reporter assay kit, with signal detection for 12 s by a luminometer (Berthold, Pforzheim, Germany), and normalized by dividing the relative light units by ß-galactosidase activity. The degree of induction was calculated relative to the control.

EMSAs. Electrophoretic mobility-shift assays (EMSAs) were performed as described previously (2). Sequences of the oligonucleotides used are shown in Table 1. Competition assays were performed using a 100-fold molar excess of an unlabeled oligonucleotide. Samples were electrophoresed on 0.5x Tris-borate-EDTA and a 6% polyacrylamide gel. After electrophoresis, the gel was dried at 80°C and autoradiographed overnight at room temperature.

ChIP. Experiments were performed with a chromatin immunoprecipitation (ChIP) assay kit (Upstate), according to the manufacturer's procedures. Briefly, 5 x 10⁶ cells were treated with 1% formaldehyde for 10 min at 37°C. Subsequent procedures were performed on ice in the presence of protease inhibitors. Cross-linked cells were harvested, washed with phosphate-buffered saline, and lysed in sodium dodecyl sulfate lysis buffer (1% sodium dodecyl sulfate, 10 mM EDTA, 50 mM Tris-HCl, pH 8.1) for 10 min at 4°C. Chromatin was sonicated with five 10-s pulses at 30% amplitude (Sonifier, Branson Ultrasonic Corp). After centrifugation (10 min, 4°C, 14,000 x g), the supernatant was diluted 10-fold with ChIP dilution buffer (0.01% sodium dodecyl sulfate, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris-HCl, pH 8.1, 167 mM NaCl). Diluted extracts were precleared in the presence of salmon sperm DNA-protein A-agarose beads (ChIP assay kit; Upstate). One-tenth of the diluted extract was kept for a direct quantitative PCR (input). The remaining extracts were incubated for 16 h at 4°C in the presence of 1 µg of specific antibodies per milliliter, followed by 1 h of incubation with salmon sperm DNA-protein A-agarose beads. Anti-BCL6 antibodies (N-3) were purchased from Santa Cruz. Following extensive washing, bound DNA fragments were eluted with a 30-min incubation in elution buffer (1% sodium dodecyl sulfate, 0.1 M NaHCO₃). The DNA was recovered for 4 h at 65°C in elution buffer containing 200 mM NaCl and then incubated in the presence of proteinase K (20 µg/ml) for 1 h at 45°C. DNA was extracted in the presence of phenol-chloroform and chloroform-isoamyl alcohol and ethanol precipitated before being subjected to PCR.

siRNA transfection. Small interfering RNA (siRNA) duplexes designed against rat BCL-6 were siRNA BCL-6_1 (5'-AUGCAACCUUAAUCUU-3') and siRNA BCL-6_3 (5'-ACCAUACAAAUGUGACCGCUU-3'). Scrambled negative control 1 siRNA (15) was used as the control siRNA. siRNAs were transfected into cells by electroporation in an Amaxa electroporation device. One million cells were resuspended in 100 µl of Amaxa electroporation transfection solution, and 2.5 µl of siRNA (20 µM) was added. The D-33 program was used. Transfected cells were plated in two wells of a six-well plate, each containing 1 ml of complete cell culture medium. Twenty-four hours after siRNA treatment, the cells were harvested.

Statistical analysis. Analysis of variance and paired or unpaired t test were performed for statistical analysis as appropriate. Probability values less than 0.05 were considered to be statistically significant. Results are expressed as means ± standard error of the means (SEM).

	RESULTS

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
PPAR-selective synthetic ligands inhibit IL-1ß-induced sPLA2-IIA activity and mRNA accumulation in rat VSMCs. Experiments were conducted on primary cultures of VSMCs isolated from rat aortas. These cells are especially appropriate for studies of inflammatory processes, since they undergo a phenotypic change in response to proinflammatory cytokines, with an increase in the expression of adhesion proteins (VCAM-1 and MCP-1), extracellular metalloproteases, and acute-phase enzymes, such as secreted sPLA2-IIA and COX-2 (26). In the case of vascular inflammation, particularly during atherogenesis, VSMCs are activated by several proinflammatory cytokines, including IL-1ß (23). Given their antiatherogenic and anti-inflammatory properties, PPAR agonists have been proposed as promising for a therapeutic approach to treat cardiovascular diseases (7). To explore the role of PPAR activation on IL-1ß-induced sPLA2, cultured VSMCs were pretreated with PPAR agonists for 6 h prior to IL-1ß treatment (24 h). We first tested the effects of WY146043, L165041, and GW1929, agonists of PPAR, PPARß, and PPAR, respectively, and then measured the enzyme activity of sPLA2 in the cell supernatant. The three PPAR agonists caused significant decreases in IL-1ß-induced sPLA2 activity in the medium of VSMCs (Fig. 1A). In order to determine whether this effect was IL-1ß specific, we conducted sPLA2 assays on TNF--induced VSMCs. In the same way, we pretreated the cells with the PPAR agonists for 6 h prior to TNF- treatment (24 h) and then measured the enzyme activity of sPLA2 in the cell supernatant (Fig. 1B). As expected, the three PPAR agonists caused significant decreases in sPLA2 activity in the supernatant.

View larger version (27K): [in this window] [in a new window]   FIG. 1. IL-1ß-induced sPLA2 activity was repressed in response to PPAR ligands. VSMCs were preincubated for 6 h with a specific agonist of PPAR (WY14643; 100 µM), PPARß (L165041; 50 µM), or PPAR (GW1929; 1 µM) and then treated or not treated with either IL-1ß (A) or TNF-. (B) Culture mediums were collected 24 h after IL-1ß treatment, and the sPLA2 activity was measured spectrofluorimetrically. (C) Semiquantitative RT-PCR was performed on 1 µg of total RNA from each sample. Amplification was performed with sPLA2 primers and with GAPDH used as a standard gene, as shown in a representative blot. mRNA levels were quantified by the Gene Tools system. (D) VSMCs were transiently transfected with 50 ng of PPAR, PPARß, or PPAR expression vector and RXR vector in equimolar quantities. Twenty-four hours after transfection, cells were pretreated for 6 h with a specific agonist of PPAR (WY14643; 100 µM), PPARß (L165041; 50 µM), or PPAR (GW1929; 1 µM) and then treated or not treated with IL-1ß (10 ng/ml). sPLA2-IIA transcription was measured by reverse transcription/real-time PCR using the expression of the GAPDH gene as an internal standard. Results are representative of three independent experiments. *, P < 0.05 (PPAR agonist-treated versus IL-1ß-treated cells). Error bars indicate SEM.

Activators of PPAR (WY14643, LY171883, and clofibrate) have been shown to enhance the transcription of the sPLA2-IIA gene in rat mesangial cells via the PPRE-1 site (bp –909 to –888) in the rat sPLA2-IIA promoter (32). We also identified a putative PPRE binding site in the human and mouse sPLA2-IIA gene (Fig. 2). The sequence alignment of the putative sPLA2-PPRE was shown to match the PPAR-binding sequence of the known PPAR target gene for acyl-CoA oxidase. So, we performed EMSA on nuclear extracts from VSMCS to examine the binding activity of the PPARs to the rat sPLA2-PPRE. When nuclear extracts from VSMCs or CHO cells were incubated with radiolabeled PPRE as a probe, gel retardation bands were observed (Fig. 2, lanes 1 and 5). The major retarded band was obviously removed by a nonlabeled competitor with the same sequence (sPLA2-PPRE) or the consensus acyl-CoA oxidase PPRE sequence. In contrast, mutated cold probes substituted from AGGTTGTCCTCTGAACTCCACA to AGGTTGTGATCTGCGCTCCACA (the two-base mismatches are underlined) (mPPRE) did not abolish the formation of PPAR-PPRE complexes, strongly suggesting a specific binding of PPAR to the sPLA2-PPRE (Fig. 2, lanes 4 and 8). In addition, nuclear extracts from cells transfected with each of the three PPAR isotypes, along with RXR, showed the same migration pattern (lanes 9 to 11). Altogether, these results demonstrate that the putative PPRE from the rat sPLA2_-IIA gene binds PPARs in VSMCs.

View larger version (39K): [in this window] [in a new window]   FIG. 2. PPAR/RXR binds to the sPLA2 promoter in VSMCs. EMSA was performed on nuclear extracts of VSMCs incubated for 24 h with IL-1ß (10 ng/ml). Radiolabeled DNA probe corresponding to nucleotides –928 to –909 of the rat sPLA2 gene promoter (sPLA2-PPRE) was incubated with VSMC nuclear extracts with or without (–) excess of unlabeled probe corresponding to sPLA2, the consensus PPRE (cons), or mPPRE (lanes 1 to 4). A radiolabeled probe corresponding to the sPLA2-PPRE incubated with nuclear extracts of VSMCs transfected with vector expressing PPAR (lane 9), ß (lane 10), and (lane 11), or with nuclear extracts of CHO transfected with vector expressing PPAR (lane 13), -ß (lane 14), and - (lane 15) were used as a positive control for PPAR/RXR binding. Asterisks indicate nucleotides matching the consensus sequence.

View larger version (19K): [in this window] [in a new window]   FIG. 3. Effects of specific PPAR activation on the sPLA2 promoter activity in VSMCs. (A) VSMCs were transfected with the sPLA2-Luc reporter plasmid: [–1153; +46]sPLA2Luc. Twenty-four hours after transfection, cells were pretreated for 6 h with increasing amounts of specific agonist of PPAR (WY14643; 1, 100, and 500 µM), PPARß (L165041; 1, 10, and 50 µM), or PPAR (GW1929; 10, 100, 1,000 nM) and stimulated with IL-1ß (10 ng/ml) for 24 h. Cells were lysed, and luciferase activity was determined and normalized to ß-galactosidase activity. (B) VSMCs were transfected with mPPRE-sPLA2-Luc. Cells were pretreated for 6 h with WY14643 (500 µM), L165041 (50 µM), or GW1929 (1,000 nM) and stimulated with IL-1ß (10 ng/ml) for 24 h. (C and D) VSMCs were transiently transfected with the [–1153; +46]sPLA2Luc construct (C) or with mPPRE-sPLA2-Luc (D), together with 50 ng of PPAR, PPARß, or PPAR expression vector and RXR vector in equimolar quantities. Twenty-four hours after transfection, cells were pretreated for 6 h with a specific agonist of PPAR (WY14643; 100 µM), PPARß (L165041; 50 µM), or PPAR (GW1929; 1 µM) and then treated or not treated with IL-1ß (10 ng/ml). (E) VSMCs were transiently cotransfected by a plasmid construct containing the PPRE-TK-LUC. Luciferase activity was determined and normalized to ß-galactosidase activity. Results are means ± SEM (error bars) and are representative of three independent experiments. *, P < 0.05 compared with IL-1ß-treated VSMCs; **, P < 0.01; –, absence of; +, presence of.

View larger version (29K): [in this window] [in a new window]   FIG. 4. BCL-6 inhibits sPLA2 activity in rat VSMCs. VSMCs were cultured in the absence or presence of Il-1ß (10 ng/ml) for 24 h. (A) The expression of BCL-6 mRNAs and protein was assessed by RT-PCR and Western blotting analysis (Table 1). (B) VSMCs were transfected with a vector expressing a wild-type (WT) BCL-6 or mBCL-6, and BCL-6 protein expression was assessed by Western blot analysis. (C) VSMCs overexpressing BCL-6 or mBCL6 were treated or not treated with IL-1ß (10 µg/ml) for 24 h. Culture mediums were collected, and the PLA2 activity was measured spectrofluorimetrically. Luciferase activity was determined and normalized to ß-galactosidase activity. Results are means ± SEM (error bars) and are representative of three independent experiments. *, P < 0.05 (compared with IL-1ß-treated VSMCs).

View larger version (14K): [in this window] [in a new window]   FIG. 5. BCL-6 can repress IL-1ß-induced sPLA2-IIA gene transcription in VSMCs. (A) VSMCs were cotransfected with increasing amounts (0.1 to 10 ng) of pcDNA3.1 vectors expressing BCL-6. (B) VSMCs were cotransfected with luciferase reporter construct containing [–1153; +46]sPLA2Luc and increasing amounts (0.1 to 10 ng) of pcDNA3.1 vectors expressing the mBCL6. As a control, cells were transfected with pcDNA3.1 vector alone. Twenty-four hours after transfection, cells were pretreated with vehicle (dimethyl sulfoxide) or 50 µM of PPARß agonist L165041 and then incubated with IL-1ß for 24 h. The relative inductions of luciferase activity were calculated after normalization to ß-galactosidase activity. Results are means ± SEM (error bars) and are representative of three independent experiments. *, P < 0.01 (compared with IL-ß-treated cells); #, P < 0.05 (compared with cells transfected with empty vector and L165041 pretreated cells); ##, P < 0.01 (compared with cells transfected with empty vector and L165041-pretreated cells).

View larger version (67K): [in this window] [in a new window]   FIG. 6. BCL-6 binds to the sPLA2 promoter in VSMCs in vitro and in vivo. (A) EMSA was performed on nuclear extracts of VSMCs pretreated with vehicle (NT) or 50 µM of the PPARß agonist L165041 and then incubated with IL-1ß for 24 h. Radiolabeled DNA probe corresponding to nucleotides –402 to –411 of the rat sPLA2 gene promoter (BCL-6 BS) was incubated with VSMC nuclear extracts with or without (–) excess of unlabeled probe corresponding either to the sPLA2 promoter BCL-6 binding site (sPLA2) (lanes 2, 5, and 8) or to the consensus BCL-6 binding site (Cons) (lanes 3, 6, and 9). The band was removed by the addition of anti-Bcl6 antibody (Ab) (lane 10). A radiolabeled probe corresponding to the sPLA2 promoter BCL-6 binding site incubated with VSMC nuclear extracts transfected with vector expressing BCL-6 was used as a positive control for BCL-6 binding (lanes 11 to 13). Radiolabeled DNA probe corresponding to the consensus BCL-6 binding site (lanes 14 to 20) was incubated with VSMC nuclear extracts with or without excess of unlabeled probe corresponding either to sPLA2 (lanes 16 and 19) or to the consensus (lanes 17 and 20). (B) VSMCs were pretreated with vehicle (NT) or 50 µM of the PPARß agonist L165041 and then incubated with IL-1ß for 24 h. DNA-protein cross-linking was induced with formaldehyde. After fragmentation, the chromatin was precipitated with rabbit antibody against BCL-6, followed by anti-rabbit IgG. To control nonspecific precipitation, the chromatin was incubated with anti-goat IgG alone. Immunoprecipitated chromatin was amplified by PCR using primers (–488/+46) designed to amplify the rat sPLA2 promoter containing the BCL-6 binding site (–340 bp). As a positive control, lysate chromatin (Input) was also amplified. –, absence of; +, presence of.

BCL-6 targets a PPARß ligand-responsive region of the sPLA2 promoter. Next, we tested the role played by the putative BCL-6-binding site in the repression of IL-1ß-induced sPLA2 activation by the PPARß ligand. VSMCs were transiently transfected with the luciferase reporter plasmid containing either the [–1153; +46]sPLA2 promoter (Fig. 7A), mPPRE[–1153; +46]sPLA2 promoter (Fig. 7B), the mBCL-6[–1153; +46]sPLA2 promoter (Fig. 7C), or mPPRE-mBCL-6[–1153; +46] (Fig. 7D). As a first result, the PPRE mutation did not affect the basal promoter activity and preserved the IL-1ß induction. Moreover, inhibition by BCL-6 overexpression was still observed with the PPRE-deleted construct in the presence or absence of PPARß agonist (Fig. 7A and B). While PPARß repression was maintained with this construct, sPLA2 repression by PPAR and PPAR ligands was inhibited (Fig. 7B). This result is consistent with the one presented in Fig. 3 and confirms the diverse effects of PPAR and - versus PPARß. Noticeably, the double mutation of the PPAR and BCL-6 binding sites slightly and positively influenced the [–1153; +46]sPLA2 promoter induction by IL-1ß (Fig. 7D). In addition, the BCL-6 binding site mutation counteracted the inhibitory effect of BCL-6 in the presence or absence of PPARß agonist (Fig. 7C and D) as well as the repression by the PPARß ligand. In contrast, this BL-6 binding site doesn't appear as a key element of the IL-1ß-induced-sPLA2 repression by PPAR and PPAR ligands. Altogether, these results suggest that the ability of PPARß to inhibit the sPLA2-IIA promoter is dependent on the BCL-6 binding site located between positions –342 and –351 relative to the transcription initiation site.

View larger version (26K): [in this window] [in a new window]   FIG. 7. Analysis of PPRE-deleted and BCL-6-mutated constructs of the sPLA2 promoter. VSMCs were transfected with the wild-type sPLA2 promoter ([–1153; +46]sPLA2-Luc) (A), its PPRE-mutated version ([–1153; +46]sPLA2-IIA) (B), its BCL-6-mutated version (mBCL-6[–1153; +46]) (C), or its double-mutated version (mPPRE-mBCL6[–1153; +46]) (D), and cotransfected with pcDNA3.1 vectors expressing (+) or not expressing (–) BCL-6. SMCs were then treated with IL-1ß and/or PPARß ligand (10 mM) (L165041), PPAR ligand (100 µM) (WY14643), or PPAR ligand (1 µM) (GW1929) for 24 h. The relative inductions of luciferase activity were calculated after normalization to ß-galactosidase activity. Results are means ± SEM and are representative of three independent experiments. *, P < 0.05 (compared with IL-1ß-treated VSMCs).

View larger version (28K): [in this window] [in a new window]   FIG. 8. Knockdown of BCL6 abolishes PPARß ligand repression of sPLA2 activity in VSMCs. VSMCs were transfected with control siRNA or with BCL-6-specific siRNAs (BCL-6_1/BCL-6_3). RT-PCR and Western blot analysis were performed in order to verify the knockdown effects. Following transfection, cells were pretreated for 6 h with vehicle (dimethyl sulfoxide) or PPARß ligand (10 µM) before stimulation with IL-1ß (10 ng/ml) for 24 h. (B) sPLA2 activity was measured in the supernatant. (C) PGE2 release into the supernatant was quantified by enzyme-linked immunosorbent assay. sPLA2-IIA (D) and COX-2 (E) transcriptions were measured by reverse transcription/real-time PCR using GAPDH gene expression as the internal standard. The results are expressed as induction compared with the expression in cells treated with phosphate-buffered saline alone. Results are means ± SEM (error bars) and are representative of three independent experiments. *, P < 0.01 (compared with IL-1ß-treated VSMCs); #, P < 0.05 (compared with cells transfected with control siRNA and L165041-pretreated cells).

PPARß activation reduces its interaction with BCL-6. The augmentation in the DNA binding activity of BCL-6 after L165041 treatment may result from different molecular mechanisms. First, these changes may be caused by an augmentation in the expression of BCL-6. However, this possibility is unlikely because no significant changes were observed in the protein expression of either PPARß or BCL-6 after L165041 stimulation (Fig. 9A). In addition, BCL-6 may interact physically with PPARs. This association has been described for PPARß and prevents this nuclear receptor from binding to its response element and thereby inhibits its ability to repress gene transcription (25). Whether a similar mechanism affects PPARß in VSMCs is not yet known. To evaluate this possibility, we performed coimmunoprecipitation studies with isolated nuclear extracts from cotransfected VSMCs expressing PPARß and BCL-6. Immunoprecipitation was conducted in VSMCs either with antibody to BCL-6 (Fig. 9B) or with antibody to PPARß (Fig. 9C). Data shown in Fig. 9B and C demonstrate that L165041 stimulation lowered the physical interaction between BCL-6 and PPARß in transfected cells, suggesting that a decreased association between these two proteins is the mechanism through which BCL-6-repressing activity is increased after L165041 stimulation. As a control, we used siRNA to knock down BCL-6 expression. The lowered BCL-6 expression resulted in an elimination of the interaction between PPARß and BCL-6.

View larger version (46K): [in this window] [in a new window]   FIG. 9. Protein-protein interaction between BCL-6 and PPARß in primary aortic SMCs. L165041 stimulation does not affect the protein level of either BCL-6 or PPARß. Nuclear protein extracts from VSMCs stimulated with L165041 for 6 h were assayed by Western blot analysis with PPARß or BCL-6 antibodies (A). Coimmunoprecipitation was performed in VSMCs transfected or not transfected (NT) with PPARß and BCL-6 or siRNA against BCL-6 (BCL-6_3) in the absence (–) or presence (+) of PPARß ligand. Nuclear extracts from VSMCs stimulated or not stimulated with L165041 (50 µM) for 4 h were subjected to immunoprecipitation using anti-BCL-6 antibody (B) or anti-PPARß antibody (C) coupled to protein A-agarose beads. Immunoprecipitates were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis and immunoblotted with an anti-PPARß antibody (B) or an anti-BCL-6 antibody (C). IP, immunoprecipitation; IB, immunoblotting; Input, 10% of cell lysate used for IP, showing an expression level of PPARß and BCL-6.

	DISCUSSION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Inflammation is one of the major pathological responses in the cardiovascular system, and sPLA2 is known to be one of the key regulators. In VSMCs, the synthesis of this enzyme is stimulated by proinflammatory cytokines and the enzyme is responsible for the production of proinflammatory mediator promoting atherogenesis (i.e., PGE2). sPLA2-IIA is thus the culprit of an inflammation amplification mechanism (18). PPARs are currently best understood as regulators of lipid metabolism. However, PPARs are expressed in the major cell types that make up atherosclerotic lesions, including macrophages, SMCs, lymphocytes, and endothelial cells, suggesting that ligands for these receptors may act both systemically and locally to influence lesion development. In fact, the anti-inflammatory activities of PPARs have been documented extensively in vitro and in vivo (8, 23, 31). PPAR has been shown to inhibit vascular inflammation, oxidative stress, and cell growth and migration through blocking NF-B, TGF-ß/Smad, and mitogen-activated protein kinase pathways (14). Evidence has also emerged suggesting that direct vascular actions of PPAR may also play an antiatherogenic role. PPAR has been detected in both endothelium and VSMCs. It has been long known that PPAR has anti-inflammatory effects on monocytes. PPAR activation can reduce cytokine (TNF-, IL-1, and IL-6) production (23), probably by inhibiting the activities of proinflammatory transcription factors, such as NF-B, AP-1, and STAT. In addition, PPAR agonists may indirectly suppress systemic production of a proinflammatory milieu, mainly by inhibiting TNF-, plasminogen activator inhibitor-1, and IL-6 expression in adipose tissue (34). The lesser known isoform PPARß has emerged as a powerful metabolic regulator in various tissues, including fat, skeletal muscle, and heart (29). An understanding of PPARß function has been augmented through a series of preclinical studies in which PPARß activation diminishes metabolic perturbations and obesity, apparently by increasing lipid uptake and oxidation in skeletal muscle (5, 9). Recent studies reveal that PPARß activation in the liver suppresses hepatic glucose output, contributing to improved glucose homeostasis (25). Additionally, PPARß may suppress inflammation through mechanisms involving the release of anti-inflammatory factors or the stabilization of repressive complexes at inflammatory gene promoters (25).

In the present study, we found that the amounts of IL-1ß-induced sPLA2 mRNA and protein in VSMCs are inhibited by the PPAR agonists WY14643, L165041, and GW1929 (Fig. 1 and 2). Besides, transient transfection assays demonstrate that the sPLA2 promoter is a valid target for PPAR/RXR, PPARß/RXR, and PPAR/RXR heterodimer transcription factors. By EMSA, we demonstrated that those heterodimers bind to a PPRE in positions –909 to –888 of the rat sPLA2 promoter (Fig. 2). These results indicate the ability of the PPAR heterodimers to modulate sPLA2 expression at a transcriptional level. IL-1ß-induced sPLA2 inhibition by PPAR and PPAR indicates that those isoforms share common features. Unexpectedly, PPAR and PPAR isoforms need the presence of the PPRE located in the sPLA2 promoter (between –909 and –888) to be able to exert their inhibitory effects in VSMCs (Fig. 3). In accordance with this anti-inflammatory effect, PPAR agonists have been shown to suppress the NF-B-dependent induction of COX-2 and IL-6 in VSMCs (34). In contrast, PPAR appears to potentiate sPLA2-IIA expression in rat mesangial cells (32). The major experiments were repeated with CHO cells, showing that PPAR ligands repressed the transcription of the endogenous sPLA2-IIA and the use of expression vector expressing PPAR and PPAR freed us of subsidiary effects possibly caused by PPAR agonists (Fig. 5). Besides, PPAR-dependent inhibitory mechanism has also been reported through competition on a composite PPRE/AP1 site of the latter promoter (16). Altogether, these results emphasize the multiplicity and complexity of the molecular mechanisms responsible of the gene expression modulation. After activation of the PPAR/RXR heterodimer at the PPRE, the PPAR/RXR complex can recruit a great variety of nuclear receptor cofactors that modulate transcriptional activity of PPAR and RXR receptor heterodimer. Therefore, multiple mechanisms are involved in controlling the transcription of PPAR target genes in a given cell or tissue, depending on its molecular environment.

Here, we demonstrated that PPARß ligands (e.g., L165041) inhibited sPLA2 production in cultured VSMCs (Fig. 1 and 8). Transient transfection confirmed the effects observed with PPARß ligand on the endogenous sPLA2 (Fig. 3). Moreover, the use of PPARß expression vector abolished IL-1ß transcriptional activation of sPLA2 in a similar manner. In this regard, unknown endogenous PPARß ligands may be sufficient to activate the more abundant PPARß to exert their inhibition on sPLA2 expression. Unlike PPAR and - isoforms, mutations in the PPRE of the sPLA2-IIA did not significantly abolish the action of PPARß on the promoter activity. This observation prompted us to investigate the role of BCL-6 for the PPARß ligand-mediated inhibition of the sPLA2-IIA gene transcription. In cultured macrophages, overexpression of PPARß exhibits a proinflammatory effect by sequestering BCL-6, whereas the deletion of PPARß increased the availability of this inflammatory suppressor (25). A recent report indicated that a PPARß-selective ligand, L165041, inhibits phenylephrine-induced expression of an NF-B target gene, MCP-1, in cultured rat neonatal cardiomyocytes (9). However, the effects of PPARß on inflammatory responses, such as IL-1ß-induced sPLA2 production in VSMCs, remained obscure.

Because PPARß activation strongly decreases sPLA2 induction by cytokines, we examined whether VSMCs express BCL-6 mRNA and protein. VSMCs transfected with a BCL-6 expression plasmid showed a great reduction of IL-1ß sPLA2 activation. Those results indicate, for the first time, the presence of the proto-oncogene BCL-6 in rat VSMCs and moreover, its ability to inhibit a proinflammatory enzyme, sPLA-IIA. Our study adds sPLA2-IIA to the list of IL-5, IL-18, prdm1, and BCL-6 itself that are direct targets of BCL-6 (35, 40, 43). Much work has been directed toward elucidating the role of BCL-6 in physiology. Along with many other immunological defects, BCL-6^–/– mice develop a profound inflammatory disease characterized by tissue infiltration of activated eosinophils, macrophages, and T-helper type 2 (Th2) cells. Noticeably, this "Th2-type" inflammatory disease, primarily affecting the heart, results from a specific heart defect (43). Thus, the identification of direct targets is important in distinguishing the precise function of this transcription factor independently of the "downstream" changes induced in the cell.

The use of a wild-type and a mutated BCL-6 expression vector showed that the PPARß effects on IL-1ß-induced inflammation are BCL-6 dependent. This mechanism appears to require the POZ/BTB domain of BCL-6 for its inhibitory effects (1). In light of the findings that the overexpression of a wild type but not a corepressor binding mutant of BCL-6 suppressed sPLA2 promoter activation in cultured VSMCs, a plausible mechanism could be that PPARß activation in VSMCs recruits a corepressor complex through the proto-oncogene BCL-6. Coimmunoprecipitation experiments demonstrate that PPARß interacts with BCL-6 in the nucleus of VSMCs (Fig. 9). In the absence of PPARß ligand, this association prevents BCL-6 from binding to its response element (Fig. 6B) and thereby inhibits its ability to repress sPLA2 transcription. These findings are in concordance with the results reported by Lee et al. (25) on another proinflammatory gene, MCP-1. Moreover, it was recently demonstrated that the lack of BCL-6 expression in pancreatic beta cells prevented PPARß-mediated repression of inflammatory responses (24). BCL-6 can act directly or indirectly to repress chemokine expression (12). BCL-6 can also negatively regulate NF-B (10) by repressing its transcription and inhibiting its nuclear binding activity. Independently of BCL-6, PPARs may also exert anti-inflammatory effects via cross talk with NF-B. Recently, it was shown that PPARß itself may physically interact with the P65 subunit of NF-B (30). Conversely, synthetic ligand-mediated PPARß activation can inhibit NF-B activation (6). These regulatory mechanisms vary between tissues, as illustrated by the differences in PPARß-mediated repression of MCP-1 expression in liver and aorta (39). These findings implicate that PPARß might act through different mechanisms to modulate inflammation in different cell types. Although the role of PPARß in SMC proliferation has not been examined extensively, it has been reported that PPARß activation might promote VSMC proliferation (44), while other results with the selective PPARß agonist GW501516 (33) showed no effect on SMC proliferation. Indisputably, more studies will be needed to understand the tissue-specific effects of this near-ubiquitous receptor and its full potential to impact the action of metabolic syndrome and its associated disorders. The physiological relevance of the PPARß ligands in VSMCs was illuminated by measuring the impact of the ligands on the production of proinflammatory mediators. Prostanoids, including PGE2, are lipid mediators produced by sequential catalysis of COX and the respective synthase. They play a major role in the induction and/or progression of the inflammatory reaction in various models of inflammatory diseases (28). In particular, PGE2 is enhanced in atherosclerotic lesions (10). Hence, the precise regulation of sPLA2-IIA and PGE2 release strongly suggests that the synthesis of prostanoids is strongly regulated during atherogenesis. We have demonstrated here that PPARß inhibits the expression of sPLA2-IIA in VSMCs sensitized with IL-1ß, resulting in a reduction in the release of PGE2 by VSMCs. Taken together, the results of our study suggest that BCL-6-dependent PPARß regulation activity might act as a potent repressor of inflammatory events, since it appears to regulate gradual accumulation of two major inflamatory factors, sPLA2-IIA and PGE2.

Nevertheless, each active compound must be evaluated carefully in clinical studies to determine definitively whether PPARß ligand may have beneficial effects for the treatment of inflammatory vascular diseases.

	ACKNOWLEDGMENTS

 
This work was supported by the Centre National de la Recherche Scientifique and the Université Pierre et Marie Curie. L.R. was supported by doctoral fellowships from the French Ministère de l'Education Nationale, de la Recherche et de la Technologie.

We gratefully thank Ari Melnick (Department of Developmental and Molecular Biology of Albert Einstein College of Medecine) for providing the BCL-6 plasmids. We thank Walter Wahli (Center for Integrative Genomics, University of Lausanne) for providing us the PPRE-TK-Luc construct and PPAR expression vectors. The English text was edited by Paul Lazarow.

	FOOTNOTES

 
* Corresponding author. Mailing address: Université Pierre et Marie Curie (Paris 6) UMR CNRS 7079-Physiologie et Physiopathologie-Université Pierre et Marie Curie 7, quai Saint-Bernard, Bât. A, 5° étage, boîte 256, 75252 Paris Cedex 05, France. Phone: 33 1 44 27 32 05. Fax: 33 1 44 27 51 40. E-mail: michel.raymondjean{at}snv.jussieu.fr

Published ahead of print on 1 October 2007.

	REFERENCES

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
1. Ahmad, K. F., A. Melnick, S. Lax, D. Bouchard, J. Liu, C. L. Kiang, S. Mayer, S. Takahashi, J. D. Licht, and G. G. Prive. 2003. Mechanism of SMRT corepressor recruitment by the BCL6 BTB domain. Mol. Cell 12:1551-1564.[CrossRef][Medline]

2. Antonio, V., A. Brouillet, B. Janvier, C. Monne, G. Bereziat, M. Andreani, and M. Raymondjean. 2002. Transcriptional regulation of the rat type IIA phospholipase A2 gene by cAMP and interleukin-1beta in vascular smooth muscle cells: interplay of the CCAAT/enhancer binding protein (C/EBP), nuclear factor-kappaB and Ets transcription factors. Biochem. J. 368:415-424.[CrossRef][Medline]

3. Antonio, V., B. Janvier, A. Brouillet, M. Andreani, and M. Raymondjean. 2003. Oxysterol and 9-cis-retinoic acid stimulate the group IIA secretory phospholipase A2 gene in rat smooth-muscle cells. Biochem. J. 376:351-360.[CrossRef][Medline]

4. Barak, Y., D. Liao, W. He, E. S. Ong, M. C. Nelson, J. M. Olefsky, R. Boland, and R. M. Evans. 2002. Effects of peroxisome proliferator-activated receptor delta on placentation, adiposity, and colorectal cancer. Proc. Natl. Acad. Sci. USA 99:303-308.[Abstract/Free Full Text]

5. Barish, G. D., V. A. Narkar, and R. M. Evans. 2006. PPAR delta: a dagger in the heart of the metabolic syndrome. J. Clin. Investig. 116:590-597.[CrossRef][Medline]

6. Berry, E. B., J. A. Keelan, R. J. Helliwell, R. S. Gilmour, and M. D. Mitchell. 2005. Nanomolar and micromolar effects of 15-deoxy-delta 12,14-prostaglandin J2 on amnion-derived WISH epithelial cells: differential roles of peroxisome proliferator-activated receptors gamma and delta and nuclear factor kappa B. Mol. Pharmacol. 68:169-178.[Abstract/Free Full Text]

7. Brown, J. D., and J. Plutzky. 2007. Peroxisome proliferator-activated receptors as transcriptional nodal points and therapeutic targets. Circulation 115:518-533.[Abstract/Free Full Text]

8. Castrillo, A., and P. Tontonoz. 2004. PPARs in atherosclerosis: the clot thickens. J. Clin. Investig. 114:1538-1540.[CrossRef][Medline]

9. Cheng, L., G. Ding, Q. Qin, Y. Huang, W. Lewis, N. He, R. M. Evans, M. D. Schneider, F. A. Brako, Y. Xiao, Y. E. Chen, and Q. Yang. 2004. Cardiomyocyte-restricted peroxisome proliferator-activated receptor-delta deletion perturbs myocardial fatty acid oxidation and leads to cardiomyopathy. Nat. Med. 10:1245-1250.[CrossRef][Medline]

10. Cipollone, F., C. Prontera, B. Pini, M. Marini, M. Fazia, D. De Cesare, A. Iezzi, S. Ucchino, G. Boccoli, V. Saba, F. Chiarelli, F. Cuccurullo, and A. Mezzetti. 2001. Overexpression of functionally coupled cyclooxygenase-2 and prostaglandin E synthase in symptomatic atherosclerotic plaques as a basis of prostaglandin E(2)-dependent plaque instability. Circulation 104:921-927.[Abstract/Free Full Text]

11. Clément, N., M. Glorian, M. Raymondjean, M. Andreani, and I. Limon. 2006. PGE2 amplifies the effects of IL-1beta on vascular smooth muscle cell de-differentiation: a consequence of the versatility of PGE2 receptors 3 due to the emerging expression of adenylyl cyclase 8. J. Cell. Physiol. 208:495-505.[CrossRef][Medline]

12. Dent, A. L., F. H. Vasanwala, and L. M. Toney. 2002. Regulation of gene expression by the proto-oncogene BCL-6. Crit. Rev. Oncol. Hematol. 41:1-9.[Medline]

13. Devchand, P. R., H. Keller, J. M. Peters, M. Vazquez, F. J. Gonzalez, and W. Wahli. 1996. The PPARalpha-leukotriene B4 pathway to inflammation control. Nature 384:39-43.[CrossRef][Medline]

14. Diep, Q. N., R. M. Touyz, and E. L. Schiffrin. 2000. Docosahexaenoic acid, a peroxisome proliferator-activated receptor-alpha ligand, induces apoptosis in vascular smooth muscle cells by stimulation of p38 mitogen-activated protein kinase. Hypertension 36:851-855.[Abstract/Free Full Text]

15. Elbashir, S. M., J. Martinez, A. Patkaniowska, W. Lendeckel, and T. Tuschl. 2001. Functional anatomy of siRNAs for mediating efficient RNAi in Drosophila melanogaster embryo lysate. EMBO J. 20:6877-6888.[CrossRef][Medline]

16. François, M., P. Richette, L. Tsagris, M. Raymondjean, M. C. Fulchignoni-Lataud, C. Forest, J. F. Savouret, and M. T. Corvol. 2004. Peroxisome proliferator-activated receptor-gamma down-regulates chondrocyte matrix metalloproteinase-1 via a novel composite element. J. Biol. Chem. 279:28411-28418.[Abstract/Free Full Text]

17. Hajjar, D. P., and K. B. Pomerantz. 1992. Signal transduction in atherosclerosis: integration of cytokines and the eicosanoid network. FASEB J. 6:2933-2941.[Abstract]

18. Hurt-Camejo, E., S. Andersen, R. Standal, B. Rosengren, P. Sartipy, E. Stadberg, and B. Johansen. 1997. Localization of nonpancreatic secretory phospholipase A2 in normal and atherosclerotic arteries. Activity of the isolated enzyme on low-density lipoproteins. Arterioscler. Thromb. Vasc. Biol. 17:300-309.[Abstract/Free Full Text]

19. Ivandic, B., L. W. Castellani, X. P. Wang, J. H. Qiao, M. Mehrabian, M. Navab, A. M. Fogelman, D. S. Grass, M. E. Swanson, M. C. de Beer, F. de Beer, and A. J. Lusis. 1999. Role of group II secretory phospholipase A2 in atherosclerosis. 1. Increased atherogenesis and altered lipoproteins in transgenic mice expressing group IIa phospholipase A2. Arterioscler. Thromb. Vasc. Biol. 19:1284-1290.[Abstract/Free Full Text]

20. Jacques, C., G. Bereziat, L. Humbert, J. L. Olivier, M. T. Corvol, J. Masliah, and F. Berenbaum. 1997. Posttranscriptional effect of insulin-like growth factor-I on interleukin-1beta-induced type II-secreted phospholipase A2 gene expression in rabbit articular chondrocytes. J. Clin. Investig. 99:1864-1872.[Medline]

21. Jaross, W., R. Eckey, and M. Menschikowski. 2002. Biological effects of secretory phospholipase A(2) group IIA on lipoproteins and in atherogenesis. Eur. J. Clin. Investig. 32:383-393.[CrossRef][Medline]

22. Jaulmes, A., B. Janvier, M. Andreani, and M. Raymondjean. 2005. Autocrine and paracrine transcriptional regulation of type IIA secretory phospholipase A2 gene in vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol. 25:1161-1167.[Abstract/Free Full Text]

23. Jiang, C., A. T. Ting, and B. Seed. 1998. PPAR-gamma agonists inhibit production of monocyte inflammatory cytokines. Nature 391:82-86.[CrossRef][Medline]

24. Kharroubi, I., C. H. Lee, P. Hekerman, M. I. Darville, R. M. Evans, D. L. Eizirik, and M. Cnop. 2006. BCL-6: a possible missing link for anti-inflam