Cardiovascular Research

Cardiovascular Research 2002 55(2):406-415; doi:10.1016/S0008-6363(02)00437-6
© 2002 by European Society of Cardiology

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Copyright © 2002, European Society of Cardiology

Oxidized phospholipids inhibit cyclooxygenase-2 in human macrophages via nuclear factor-B/IB- and ERK2-dependent mechanisms

Sonia Eligini^a, Marta Brambilla^a, Cristina Banfi^a, Marina Camera^a, Luigi Sironi^a, Silvia Stella Barbieri^a, Johan Auwerx^b, Elena Tremoli^a and Susanna Colli^a,*

^aE. Grossi Paoletti Center, Department of Pharmacological Sciences, University of Milan, Via Balzaretti 9, 20133 Milan, Italy
^bInstitut de Génétique et Biologie Moléculaire et Cellulaire (IGMBC), Illkirch, France

susanna.colli{at}unimi.it

^* Corresponding author. Tel.: +39-2-5835-8291; fax: +39-2-5835-8250

Received 2 January 2002; accepted 8 April 2002

	Abstract

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
Objective: Oxidized low-density lipoproteins (ox-LDL) or their components suppress macrophage inflammatory response by down-regulating cytokine synthesis, nitric oxide synthase and inducible cyclooxygenase (Cox-2). This event is crucial for the pathophysiological process leading to the formation of atherosclerotic plaque. Our present study focused on the mechanisms through which oxidized phospholipids inhibit LPS-induced Cox-2 expression in human macrophages. Methods: Macrophages were incubated with a mixture of oxidized fragmented phospholipids (ox-PAPC), present in modified LDL, and then exposed to LPS. Cox-2 was evaluated in terms of protein levels, mRNA and activity. Results: Ox-PAPC dose-dependently inhibited Cox-2 protein, mRNA and activity by preventing NF-B binding to DNA. This effect was consequent to alterations of the degradation pattern of IB. Moreover, ox-PAPC markedly prevented extracellular signal-regulated kinase (ERK2) activation, leading to Cox-2 expression, whereas activation of the transcription factor peroxisome proliferator-activated receptors (PPARs) was not influenced. Conclusion: ox-PAPC down-regulates LPS-induced Cox-2 expression in human macrophages by targeting both NF-B/IB and ERK2 pathways. An altered inflammatory response by macrophages within atheromata may contribute to the progression of atherosclerosis.

KEYWORDS Atherosclerosis; Infection/inflammation; Macrophages; Prostaglandins; Signal transduction

	1 Introduction

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
Macrophages are significant cellular participants throughout the development of atherosclerosis, which presents features of an inflammatory disease [1]. These cells are engaged to ingest modified lipoproteins, e.g., ox-LDL, through a variety of scavenger receptors and to recognize oxidized phospholipids as markers of oxidized moieties, all events that ultimately lead to the generation of the foam cells.

Once activated by inflammatory stimuli, macrophages synthesize and release eicosanoids that are potent modulators of inflammation. The rate-limiting step in their synthesis is the enzyme cyclooxygenase (Cox), present in two isoforms. Cox-1, which is constitutive and detected in most human tissues, and the inducible Cox-2, selectively expressed by a variety of cells exposed to bacterial lipopolysaccharide (LPS) and cytokines [2]. In macrophages, Cox-2 induction by LPS is characterized by ‘redundancy’ both at the extranuclear signaling level and at the level of transcriptional activation [3]. Mitogen-activated protein kinase (MAP-kinase) and protein kinase C (PKC) are involved in Cox-2 expression by LPS [4,5]. These pathways act, in a coordinate way, with several transcription factors, among which the nuclear factor -B (NF-B) [3,6]. NF-B activation is controlled by the family of inhibitors of B protein (IB) which retain the transcription factor in the cytoplasm. Treatment of macrophages with inflammatory stimuli targets the degradation of IB and allows NF-B to translocate into the nucleus [7]. NF-B activation is also regulated by members of the peroxisome proliferator-activated receptors (PPARs) family, through a mechanism termed ‘transrepression’ [8–10]. Different isoforms of PPARs have been identified so far; in particular, PPAR activation has been shown to down-regulate several inflammatory responses of monocytes and macrophages, among which is the production of inflammatory cytokines [8].

We have previously shown that ox-LDL and its phospholipid component as well, impairs Cox-2 in human macrophages, an effect mimicked by ox-PAPC, a surrogate for oxidized phospholipids obtained by autoxidation of 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine [11]. Ox-PAPC has been identified as a component of minimally oxidized LDL and its molecular structure and biological activities have been characterized [12,13].

In this study we have investigated the mechanisms through which ox-PAPC impairs Cox-2 in human macrophages.

	2 Materials and methods

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
2.1 Reagents
Cell culture medium and reagents were from BioWhittaker. Lipopolysaccharide (E. coli 0114:B4) was from Difco Labs. Benzamidine hydrochloride, leupeptin hemisulfate, soybean trypsin inhibitor, sodium orthovanadate, 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine (PAPC), bovine serum albumin (fatty acid-free and low endotoxin), clofibrate, neutral red, 3-(4,5-dimethyl-thiazol-2-yl)2,5-di-phenyltetrazolium bromide (MTT) and bisphenol A diglycidyl ether (Der Resin 332, BADGE) were from Sigma. EDTA and NaF was from Carlo Erba. Electrophoresis reagents were from Pharmacia Biotech. Wy 14,643 and 15-deoxy-^12,14-prostaglandin J₂ (15d-PGJ₂) was from Biomol, 13-oxo-octadecadienoic acid (13-oxoODE) was from Cayman Chemicals and TRIzol Reagent from Life Technologies. BRL-49653 was a kind gift by Dr. S. Trowbridge (SmithKline Beecham Pharmaceuticals).

Ox-PAPC was prepared according to Watson by transferring 1 mg PAPC in 100 µl of chloroform to a clean 16x25-mm glass test tube and evaporating the solvent under nitrogen stream [12]. The lipid residue was allowed to oxidize on exposure to air for 48 h. Caution was taken to avoid LPS contamination that was checked by the Limulus amebocyte lysate assay test (BioWhittaker). Aliquots of ox-PAPC were stored at –80 °C under nitrogen and used within 1 week.

2.2 Monocyte culture
Venous blood from healthy donors was obtained in accordance with the principles outlined in the Declaration of Helsinki. Mononuclear cells were separated by Ficoll-Paque density gradient (Pharmacia Fine Chemicals), as described [11]. Monocytes were isolated from lymphocytes by adherence to six-well plates (2 h at 37 °C, 5% CO₂). Cell preparations were >90% monocytes, as determined by nonspecific esterase staining. Macrophages were obtained by culturing monocytes for 7 days at 37 °C in a 5% CO₂ humid atmosphere in M-199 containing 2 mM glutamine, 0.5% antibiotics and 10% human heat-inactivated AB serum. They were identified by the presence of CD68Ag detected by a specific monoclonal mouse anti-human macrophage antibody (Dako). The endotoxin content of all culture materials and reagents was measured with the Limulus assay. Unless otherwise specified, macrophages were incubated with various agents in serum-free medium containing 0.2% fatty acid-free bovine serum albumin for 2 h. Medium was then removed and macrophages were exposed for further 4 h to fresh medium containing LPS (1 µg/ml). This experimental protocol was adopted in order to avoid a direct interaction between LPS and ox-PAPC, since the formation of a stable complex LPS/ox-LDL has been reported [14]. Incubation time and LPS concentration were selected on the basis of results obtained in preliminary experiments. Macrophage viability was assessed by Neutral Red and MTT assay.

2.3 Cyclooxygenase activity
Macrophages were washed with Hanks’ buffer (pH 7.4) containing 1 mg/ml BSA and incubated with the same buffer containing 10 µM sodium arachidonate. After 30 min, supernatants were harvested and the amount of TxB₂ released in the supernatant was measured by enzyme immunoassay (Cayman Chemicals), as previously described [11].

2.4 Cell extract and Western blot analysis
Whole cell lysates were prepared in RIPA buffer containing 5 mM sodium orthovanadate, 100 mM NaF, 5 µg/ml leupeptin, 5 µg/ml aprotinin, 2 mM pefabloc. Protein content was determined by the micro-bicinchoninic acid assay [15]. For cytosol extract preparation, macrophages were washed twice in ice-cold TBS, harvested in 100 µl buffer A (10 mM Hepes, pH 7.9, 1.5 mM MgCl₂, 10 mM KCl, 0.5 mM DTT) and incubated for 15 min on ice. Triton X-100 (0.4%) was added. Cytosols were separated from nuclei by centrifugation at 15 000xg for 1 min at 4 °C. SDS–PAGE was performed as previously described [11]. Membranes were subsequently incubated for 1 h at room temperature with monoclonal antibodies raised against Cox-2, mAb 29 (1/10 000) or Cox-1, mAb 10 and 11 (5 µg/ml), kindly donated by A. Habib, U 348, INSERM, Paris. Antibodies directed against the C-terminal PPAR peptide (monoclonal, 1/500), against IB (polyclonal, 1/500), and against haem-oxygenase-1 (polyclonal, 1/2000) were from Santa Cruz Biotechnology. MAP-kinase immunoblotting was performed using antibodies (1/1000) against phosphospecific p44–p42 MAP-kinase (Thr202/Tyr204, monoclonal, New England Biolabs, Celbio) and p38-kinase (pTpY^180/182, polyclonal, Biosource International), as previously described [16]. Blots were incubated with donkey anti-mouse or anti-rabbit IgG conjugated with peroxidase (Jackson ImmunoResearch) at 1/5000 for 1 h at room temperature. Monoclonal antibody directed against β-actin (Sigma) was used as internal standard for control of protein load. ECL substrates (Amersham) were used to reveal positive bands that were visualized after exposure to HyperfilmTM ECL (Amersham).

2.5 RNA extraction and reverse transcriptase-polymerase chain reaction (RT-PCR)
Total cellular RNA was extracted from macrophages with TRIzol Reagent. Cox-2 mRNA levels were detected using a coupled reverse transcription-polymerase chain reaction (RT-PCR) performed using a GeneAmp RNA PCR kit (Perkin-Elmer) according to the manufacturer's instructions. To identify Cox-2 mRNA, exact primers were synthesized by an analytical RT-PCR procedure developed for human cyclooxygenase. Cox-2 primers were 5'-TTCAAATGAGATTGTGGAAAATTGCT-3' (a 27-mer sense, oligonucleotide at position 573) and 5'-AGATCATCTCTGCCTGAGTATCTT-3' (a 24-mer antisense oligonucleotide at position 878), giving rise to a 305-bp PCR product [17]. Primers were also synthesized to amplify the cDNA encoding GAPDH, a constitutively expressed gene, as control. GAPDH primers were 5'-CCACCCATGGCAAATTCCATGGCA-3' (a 24-mer sense oligonucleotide at position 216) and 5'-TCTAGACGGCAGGTCAGGTCCACC-3' (a 24-mer antisense oligonucleotide at position 809) giving rise to a 593-bp PCR product. RNA concentration was determined spectrophotometrically. Total cellular RNA from each sample (1 µg) was reverse transcribed at 37 °C for 1 h. The same amount of the resulting cDNA was used for amplification by specific primer for human Cox-2. PCR amplification was carried out, in the presence of 0.2 µM primers, using a thermal cycler GeneAmp PCR System 2400 (Perkin-Elmer). The following conditions were used: 30 cycles with denaturation at 94 °C for 1 min; primer annealing at 55 °C for 1 min: extension at 72 °C for 1 min. The amplification of human GAPDH was used as internal standard. Aliquots (10 µl) of amplified products were resolved by electrophoresis through a 2% agarose gel. Amplified cDNA bands were detected by ethidium bromide staining.

2.6 Nuclear extracts and electrophoretic mobility shift assay
Nuclear extracts were prepared from cell suspensions using a protocol according to Dignam et al. [18], slightly modified [19]. The nuclear pellet was washed with 1 ml buffer B (20 mM Hepes, pH 7.9, 400 mM NaCl, 25% glycerol, 1.5 mM MgCl₂, 0.2 mM EDTA, 0.5 mM PMSF, and 0.5 mM DTT). Samples were vortexed for 30 min at 4 °C. Debris was removed by centrifugation at 13 000xg for 10 min. After supernatant collection, the protein content was determined and electrophoretic mobility shift assays were performed as follows: the synthetic single-stranded oligonucleotides (Eurogentec, Herstal, Belgium) containing the proximal and distal NF-B sites were annealed with the complementary primers and radiolabeled with [³²P]dCTP (Amersham). The consensus sequences for NF-B are underlined: distal NF-B site (upstream, within –455 to 428 from the transcriptional start site) 5'-GGCGGGAGA GGGGATTCCC GCGCCCCC-3'; proximal NF-B site (downstream, withinx –232 to 205 from the transcriptional start site) 5'-CAGGAGAGT GGGGACTACC CCCTCTGCT-3' [20]. Nuclear extracts (4 µg) were incubated for 20 min at room temperature with 100 000 cpm of ³²P-labeled double-strand oligonucleotides. Reactions were performed in binding buffer (18 µl) containing 60 mM KCl, 20 mM Hepes, 1 mM EDTA, 15 mM DTT, 12% glycerol and 4 µg poly(dI/dC) (Amersham). Protein–DNA complexes were separated from free DNA probe by electrophoresis through 5% nondenaturating acrylamide gels in 0.5xTBE (45 mM Tris base, 45 mM boric acid, 1 mM EDTA). Gels were dried and subjected to autoradiography. NF-B specific bands were confirmed by competition with a 100-fold molar excess of an unlabeled NF-B probe.

2.7 Transfection assays
CHO cells grown in F-12 medium were transiently transfected using a calcium phosphate precipitation method, as described by Sambrook et al. [21]. A reporter construct containing three copies of a consensus PPRE cloned upstream of the thymidine kinase-luciferase reporter (J₃-TK-LUC) was used [22]. The control vector pRSV-galactosidase (Promega) was cotransfected as a control for transfection efficiency. The cells were plated in six-well plates and transfected when 80–90% confluent, 3 h after a complete medium replacement. Cells were incubated with calcium phosphate-precipitated DNA (1 µg plasmid/well) for 5 h. After shock (2-min glycerol, 15%, v/v), fresh medium containing stimuli was added and cells were harvested 2 h later. Supernatants were removed and luciferase production was measured on cell lysates using a commercially available kit (Promega). After scraping, cells were transferred into Eppendorf tubes, vortexed, subjected to freezing and thawing and finally centrifuged at 12 000xg for 15 s. Aliquots of supernatants (20 µl) were mixed with the luciferase substrate reagent and absorbancies were determined by a fluorimeter (Lumat LB 9501, Berthold). β-Galactosidase activity was determined by standard assay (Invitrogen). Luciferase data were corrected for β-galactosidase activity and are reported as percentage activity compared with control cells.

2.8 Statistical analysis
Data are reported as means±S.E.M. Computer-assisted statistical analyses used the ANOVA program, and probability values were calculated using Fisher's protected least-significant difference test. A value of P<0.05 was considered significant.

	3 Results

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
3.1 Ox-PAPC inhibits LPS-induced Cox-2 in human macrophages
Macrophages differentiated from monocytes (7-day culture) did not express Cox-2, nor did they after incubation with ox-PAPC (5 µg/ml) for 4–24 h. In contrast, exposure to 1 µg/ml LPS for 4 h induced appreciable expression of Cox-2, recognized as a double band at 70 kDa. Ox-PAPC incubated for 2 h before LPS diminished the amount of Cox-2 in a concentration-dependent manner (Fig. 1A). The same extent of inhibition was observed when ox-PAPC was incubated longer (up to 24 h) before LPS addition (not shown). In contrast, ox-PAPC added concomitantly with, or 1 or 2 h after LPS did not affect Cox-2 (Fig. 1B). Subsequent experiments therefore were performed in macrophages exposed to ox-PAPC for 2 h prior to LPS. Ox-PAPC reduced Cox-2 activity, measured as thromboxane B₂ (TxB₂) released in the supernatant (Fig. 1C). The maximal concentration of ox-PAPC used throughout the study (10 µg/ml) has been reported to be present in 100 µg/ml ox-LDL [23].

View larger version (21K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 1 Ox-PAPC concentration-dependently inhibits Cox-2 and thromboxane B2 (TxB2) formation in LPS-stimulated macrophages. (A) Macrophages were incubated with medium or with ox-PAPC (1–10 µg/ml) for 2 h. LPS (1 µg/ml) was added and incubated for further 4 h. (B) ox-PAPC (5 µg/ml) was added to macrophage culture 2 h before, simultaneously with or after (1 or 2 h) LPS. Cox-2 and β-actin were determined in macrophage lysates by Western blot analysis. (C) Cox-2 activity was measured in macrophage supernatants as TxB2 formation, by EIA. Results are the means±S.E.M. of three independent experiments. *P<0.01 versus basal; #P<0.01 versus LPS.

 
Ox-PAPC (10 µg/ml) did not influence macrophage viability as assessed by Neutral Red (>90% macrophages were viable). A minor decrease of the cell number was observed by MTT test, which suggests a very limited, if any, toxicity.

Under these experimental conditions, the expression of Cox-1 was not influenced (not shown).

Cox-2 is characterized by the presence of a haem group. The possibility that ox-PAPC impairs Cox-2 levels through an increased expression of the haem-degrading enzyme haem oxygenase-1 (HO-1) was therefore tested. Ox-PAPC was ineffective to induce HO-1, ruling out a role of this enzyme in the observed decrease of Cox-2 levels (not shown).

Reduction of Cox-2 protein levels was consequent to attenuation of Cox-2 mRNA levels, as assessed by RT-PCR. In contrast unoxidized PAPC did not affect Cox-2 mRNA (Fig. 2).

View larger version (50K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 2 Ox-PAPC decreases Cox-2 mRNA, PAPC does not. Macrophages were incubated with medium (lanes 1 and 2) or with PAPC (5 µg/ml), either oxidized (lane 4) or unoxidized (lane 3) for 2 h. LPS (1 µg/ml) was then added for 1 h. Total RNA was extracted and Cox-2 and GAPDH mRNA levels were estimated by RT-PCR. The sequences of the primer sets and oligonucleotide probes for Cox-2 and GAPDH are given in Section 2. Results are representative of three independent experiments.

 
3.2 Ox-PAPC inhibits LPS-induced NF-B activation in human macrophages by altering the degradation of IB
Ox-LDL are known to influence the expression of several LPS-induced inflammatory genes by interfering with NF-B activation [24–26]. Nuclear extracts of macrophages exposed to LPS for 45 min formed a DNA–protein complex with the distal NF-B sequence which was prevented if a 100-fold excess of unlabeled NF-B probe has been added (Fig. 3A). No DNA–protein complex was observed when the proximal NF-B sequence was used as a probe (data not shown). This is in agreement with results published by Barrios-Rodiles et al. in the same cell type [20].

View larger version (38K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 3 Ox-PAPC inhibits LPS-induced NF-B translocation and reduces IB degradation in human macrophages. (A) EMSAs for the distal NF-B probe were performed on nuclear extracts of macrophages treated with medium alone (lanes 1 and 2) or with medium supplemented with ox-PAPC (1–5 µg/ml) (lanes 3–5). Macrophages were then exposed to LPS (1 µg/ml) for 45 min. Specificity of the binding was determined on cell extracts by competition, using a 100-fold excess of unlabeled distal probe (lane 6). Results are representative of three independent experiments. (B) Macrophages pretreated with ox-PAPC (5 µg/ml) or medium alone were stimulated with LPS for the indicated times. (C) Ox-PAPC concentration-dependently reduces IB degradation. Analysis of IB protein content was performed by Western blot on cytosolic extracts, as described in Section 2. Results are representative of three independent experiments.

 
Ox-PAPC concentration-dependently diminished the amount of the DNA–protein complex (Fig. 3A), which suggests that Cox-2 expression is impaired by the prevention of NF-B translocation.

To further explore the mechanisms involved in the effect of ox-PAPC on NF-B activation, the pattern of IB degradation was evaluated. LPS-treatment of macrophages resulted in a loss of immunoreactive IB protein, detected at 30 min from LPS addition (Fig. 3B). Macrophages, pretreated with ox-PAPC exhibited a reduced IB degradation (Fig. 3B). The effect was concentration-dependent (Fig. 3C).

3.3 Ox-PAPC inhibits LPS-induced p38 and ERK2 activation in human macrophages
NF-B activation in LPS-stimulated macrophages is under the control of MAP-kinase [3]. To define the role of MAP-kinase cascade in Cox-2 induction by LPS, human macrophages were preincubated for 30 min with 25 µM PD098059, a specific inhibitor of ERK1/2 activity and with 10 µM SB203580, which is specific for p38 kinase. Macrophages were then exposed to LPS for further 4 h. LPS-induced Cox-2 was prevented by PD098059 but not by SB203580 (not shown), which suggests a pivotal role of ERK1/2 in LPS-induced Cox-2 expression in human macrophages.

The hypothesis that ox-PAPC interferes with MAP-kinase pathway was tested by a Western blot technique using antibodies specific for the phosphorylated forms of p38 and ERK1/2. Compared to unstimulated macrophages, bands corresponding to the phosphorylated isoform of p38 kinase and ERK2, respectively, were evident in macrophages exposed to LPS (Fig. 4). The band appeared within 10–20 min, with a peak at 30–40 min and values still above the basal level by 60 min (Fig. 4A). Ox-PAPC (5 µg/ml) almost completely reduced p38 and ERK2 phosphorylation (Fig. 4A). The effect of ox-PAPC on both kinases was concentration-dependent (Fig. 4B)

View larger version (29K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 4 Ox-PAPC inhibits LPS-induced MAP-kinase activation in human macrophages. (A) Macrophages, preincubated for 2 h with ox-PAPC (5 µg/ml) or with medium alone were exposed to LPS (1 µg/ml) for indicated times. (B) Ox-PAPC concentration-dependently inhibits MAP-kinase activation. MAP-kinase immunoblotting was performed using antibodies against phosphospecific p44–p42 (ERK1/2) and p38-kinase, as described in Section 2. Results are representative of three independent experiments.

 
3.4 Effect of PPAR activators on LPS-induced Cox-2
Ox-PAPC represents a natural ligand of PPAR in human endothelial cells [27,28]. The presence of PPAR has been detected in human monocytes [29], whereas in monocytes differentiated into macrophages, the PPAR isoform predominates [29]. The presence of PPAR was detected in differentiated macrophages obtained in our culture condition (not shown). To investigate whether PPAR activation is involved in the NF-B-mediated inhibition of Cox-2, we exposed macrophages for 2 h to a series of PPAR ligands. The synthetic PPAR ligand Wy 14,643 (100 µM) and clofibrate (500 µM), did not affect Cox-2 (Fig. 5A). Similarly, BRL-49653, a synthetic high affinity PPAR ligand (Fig. 5B) did not affect Cox-2, even when incubated with macrophages for much longer time (up to 24 h) before LPS addition (not shown). The natural PPAR ligand 13-oxoODE did inhibit Cox-2 expression (Fig. 5B), but only at 50 µg/ml, a concentration twice as high as that reported to activate PPAR [30]. Another natural PPAR ligand, 15d-PGJ₂ [31,32], concentration-dependently decreased Cox-2 protein (Fig. 5C).

View larger version (21K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 5 Effect of PPAR ligands on LPS-induced expression of Cox-2 in human macrophages. Macrophages were incubated for 2 h in medium alone or in medium containing the PPAR ligands WY 14,643 (100 µM) or clofibrate (500 µM) (A) and the PPAR ligands BRL 49653 (10 µM), 13 oxo-ODE (50 µg/ml) (B) or 15d-PGJ2 (5–30 µM) (C). LPS (1 µg/ml) was then added, where indicated, for an additional 4 h. Cox-2 and β-actin proteins were determined in macrophage lysates as described in Fig. 1. This figure is representative of three independent experiments.

 
Transient transfection experiments were carried out in CHO cells, which express PPAR [33], using a reporter plasmid containing three copies of a consensus PPRE upstream of a luciferase reporter gene (J₃-TK-LUC). Ox-PAPC (5 µg/ml, 2-h incubation) did not affect luciferase activity. In contrast, BRL-49653 (10 µM), Wy 14,643 (100 µM) and 15d-PGJ₂ (10 µM), increased PPRE-luciferase activity within 2 h (Fig. 6). These data together make it unlikely an effect of ox-PAPC on Cox-2 via PPAR activation. Moreover, BADGE, an antagonist of PPAR ligands that prevents the transcriptional action of the receptor [34] did not affect the inhibition of Cox-2 exerted by ox-PAPC (data not shown).

View larger version (13K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 6 Effect of ox-PAPC and PPAR agonists on PPRE activation in transiently transfected CHO cells. Transfections were performed in 80–90% confluent cells using a reporter plasmid containing three copies of a consensus PPRE upstream of a luciferase gene. The control vector pRSV-galactosidase was cotransfected as a control for transfection efficiency. Cells were treated for 2 h with ox-PAPC (5 µg/ml), BRL-49653 (10 µM), WY 14,643 (100 µM) and 15d-PGJ2 (10 µM). Luciferase data were corrected for β-galactosidase activity and are reported as percentage activity compared with control cells. Values are the means±S.E.M. of five individual experiments.

 

	4 Discussion

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
Oxidized phospholipids are important markers for recognition of oxidized moieties by macrophages and therefore represent potential modulators of the atherogenic process. Within this context, ox-PAPC provides a useful tool to investigate mechanisms of inflammation related to atherogenesis [35,36]. This mixture of fragmented phospholipid can be indeed formed ‘in vitro’ by free-radical treatment of unsaturated phospholipids and data from ‘in vivo’ studies indicate that levels of oxidized phospholipids are increased in fatty streak lesions from cholesterol fed rabbits [13]. In addition, increased plasma levels of fragmented phosphatidylcholine in response to various forms of oxidative stress have been reported in human and rat [37].

In this study we show that ox-PAPC quickly inhibits Cox-2 expression induced by LPS in macrophages by interfering with NF-B/IB and MAP-kinase pathways. The capacity of ox-PAPC to target multiple steps involved in Cox-2 expression that act in a coordinate way to transduce signals from LPS to nucleus might explain the marked inhibitory effect observed. Inhibition of NF-B activation by ox-PAPC is consequent to alteration of IB degradation. This mechanism is identical to that described for ox-LDL in mononuclear phagocytes and considered as responsible for the inhibition of several inducible genes necessary for the inflammatory response [25].

Besides the inhibition of NF-B binding to DNA, we show, for the first time, that ox-PAPC acts also by preventing LPS-induced activation of both p38 and ERK2. This finding suggests that oxidized phospholipids may act as potential modulators of a signal transduction pathway leading to the expression of a variety of genes associated with the inflammatory response. Interestingly, from our data obtained with specific inhibitors of MAP-kinases, it emerges that only ERK1/2 is involved in Cox-2 expression induced by LPS. This finding is in accordance with a recent report which addresses a role of ERK1/2 in Cox-2 transcription through the NF-B site [3]. The observation that p38 and NF-B activation are under the control of separate pathways, at least in LPS-activated macrophages [38], may explain the lack of effect of SB203580 on Cox-2, observed in our condition. Whether activation of ERK2 results in IB phosphorylation which, in turn, inhibits NF-B translocation is an attractive issue that deserves further investigation.

A role of ox-PAPC as a regulator of gene transcription has recently been proposed [27,28]. In particular, ox-PAPC exerts, via a PPAR-mediated mechanism, a proinflammatory effect in that it stimulates the synthesis of specific monocyte adhesion molecules and monocyte activators by endothelial cells [28]. Interestingly, the same authors described an anti-inflammatory effect of PPAR-activators in LPS-stimulated endothelial cells [39]. These apparently discrepant results allowed the conclusion that PPAR ligands behave differently depending upon the presence of an inflammatory stimulus in the experimental system.

Our data rule out a role of PPAR activation in Cox-2 inhibition by ox-PAPC. Cox-2 levels are negatively affected by the natural PPAR ligands 13-oxoHODE and 15d-PGJ₂. 13-HODE, however, has been reported to act also as selective inhibitor of classical protein kinase C isoenzymes [40] and this signaling pathway is recognized as being critically involved in LPS-mediated induction of Cox-2 [5]. 15d-PGJ₂ has been recently shown to inhibit Cox-2 expression by PPAR-independent mechanisms, by affecting multiple steps in NF-B signaling pathways [41–43]. Though our study did not explore these aspects, it seems likely that such mechanisms are operative in our experimental conditions.

In conclusion, our data indicate that ox-PAPC down-regulates LPS-induced Cox-2 in macrophages by targeting both NF-B/IB and ERK2 pathways.

It can be envisaged that in the sub-endothelial microenvironment, conditions of oxidative stress/antioxidant imbalance impair the capacity of macrophages to correctly contribute to the resolution of the inflammatory response. Indeed, the capacity of macrophages within lesions to be activated is crucial for the resolution of inflammation, and consequently an impaired prostaglandin formation in foam cells, coupled with a reduction of cytokine synthesis [44,45], may represent an important contributing factor in the formation and progression of fatty streaks.

Time for primary review 27 days.

	Acknowledgments

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 
This work was supported by grants from the European Community (HIFMECH Study, contract BMH4-CT96-0272, grant to E.T.) and from the Italian Ministry of University and Scientific Research and University of Milan (Cofin, grant 9906203775/1999 to S.C. and FIRST 2000/2001, grant to S.C.).

	References

Top Abstract 1 Introduction 2 Materials and methods 3 Results 4 Discussion Acknowledgments References

 

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