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CD36 is a membrane glycoprotein expressed on platelets, monocytes, macrophages, and several other cell types that was recently demonstrated to be involved in platelet activation in response to oxidized phospholipids, including oxidized LDL. Although the role of CD36 in other vascular cells has not been well defined, previous studies have demonstrated that cd36-knockout (cd36–/–) mice have prolonged thrombosis times after vascular injury, which can be protective in the state of hyperlipidemia. Here, we found significantly less ROS in the vessel walls of cd36–/– mice compared with WT after chemically induced arterial injury, suggesting that CD36 may contribute to ROS generation in the VSMCs themselves. Gene expression analysis revealed that the antioxidant enzymes peroxiredoxin-2 (Prdx2) and heme oxygenase-1 were upregulated in cd36–/– VSMCs. Molecular dissection of the pathway in isolated mouse VSMCs revealed CD36 ligand-dependent induction of Fyn phosphorylation, with subsequent phosphorylation and degradation of the redox-sensitive transcription factor Nrf2. Chromatin immunoprecipitation experiments further showed that Nrf2 directly occupied the Prdx2 promoter. The importance of this pathway was evidenced by increased ROS generation in prdx2–/– mice and decreased thrombosis times in both prdx2–/– and nrf2–/– mice after vascular injury. These data suggest that CD36-mediated downregulation of antioxidant systems in VSMCs may contribute to its prothrombotic, proinflammatory, and atherogenic effects.
CD36, a class B scavenger receptor, is an 88-kDa membrane glycoprotein expressed on platelets, monocytes, macrophages, and several other cell types (1). It is a multifunctional receptor with independent capacity to bind 3 major classes of ligands: modified phospholipids, long-chain fatty acids, and proteins containing thrombospondin structural homology domains. Although CD36 was first identified on platelets, its role in platelet function remained undefined until recently, when our group showed that CD36 promotes platelet activation in response to oxidized phospholipids, including oxidized LDL (oxLDL) (2). We found that genetic deletion of CD36 in mice protected them from the prothrombotic state associated with hyperlipidemia (2) and oxidant stress and also significantly prolonged thrombotic occlusion times after vascular injury induced by ferric chloride (FeCl3) (3). The mechanisms underlying this phenotype remain incompletely understood, but the antithrombotic effects of CD36 deficiency were dependent on the dose of FeCl3 used to induce injury (3) and platelet transfusion studies revealed that they were in part platelet dependent. Furthermore, CD36-mediated signaling through specific Src family kinases and MAPKs was shown to promote platelet activation after vascular injury (4). The endogenous CD36 ligands generated during FeCl3-induced injury include endothelial cell–derived microparticles (EMPs), which we showed bind specifically to platelet CD36 and enhance platelet reactivity (3).
These studies, while showing a critical role for platelet CD36 in thrombosis, do not rule out a role for CD36-mediated signaling in other vascular cells in promoting thrombosis. Although CD36 is not expressed on large-vessel endothelial cells (5, 6), several groups have reported that VSMCs in culture express low levels of CD36 (7–12) and that expression can be upregulated by PPARγ (9). No function for CD36 has been identified, however, on VSMCs. Since FeCl3 and other injuries induce thrombosis via generation of oxidant damage and since CD36 has been reported to promote ROS formation in macrophages in vitro and in a murine cerebral ischemia model in vivo (13, 14), we hypothesized that CD36 on VSMCs could contribute to vascular injury by generating further oxidant stress and that part of the “thrombo-protection” seen in cd36–/– mice is due to diminished ROS production in the vessel wall.
In this manuscript, we show that CD36 is abundantly expressed by murine arterial smooth muscle cells in vivo and in vitro and absence of CD36 attenuated ROS production in response to FeCl3-induced vascular injury in a mouse carotid artery thrombosis model. Using proteomic and genetic approaches, we found that CD36 regulates expression of the antioxidant enzymes peroxiredoxin-2 (Prdx2) and heme oxygenase-1 (HO-1) in VSMCs, and defined a CD36-dependent pathway mediated by the redox-sensitive transcription factor, NF-E2–related factor-2 (Nrf2), responsible for this regulation. Targeting CD36 may enhance activity of Nrf2 and thereby provide an effective strategy to restore endogenous antioxidant defenses in cardiovascular diseases.
To determine whether CD36 contributes to vascular oxidant stress after FeCl3-induced carotid injury, we measured ROS formation in the vessel wall by direct injection of a fluorescent superoxide indicator, dihydroethidium (DHE), also known as hydroethidine, which forms 2-hydroethidium (EOH) after reacting with superoxide anion (15). In WT animals, we observed a rapid increase in vessel fluorescence that was dose and time dependent, peaking at approximately 7-fold that of baseline with 12.5% FeCl3 (data not shown). Fluorescence intensity was significantly decreased in the vessel wall of cd36–/– mice compared with WT (Figure (Figure1A).1A). Laser confocal microscopy examination of cross sections of dissected thrombus-containing vessels demonstrated that the fluorescence was mainly within the vessel wall, not the cells within the thrombi (Figure (Figure1B),1B), suggesting that the origin of the ROS was vascular cells. The level of ROS in the vessel walls of cd36–/– mice was similar to that seen in WT mice treated with edaravone (3-methyl-1 phenyl-2-pyrazoline-5-1), a potent free radical scavenger (Figure (Figure1C).1C). Interestingly, edaravone administration significantly prolonged occlusion times in WT mice after FeCl3-induced injury (Figure (Figure1D),1D), phenocopying untreated cd36–/– mice. Edaravone also induced a small but marked increase in the vessel occlusion time in cd36–/– mice injured with the high dose of FeCl3 (not shown).
These data suggest that CD36-mediated enhancement of oxidant stress contributes to thrombosis. Since microparticles (MPs) can be generated from endothelial cells by oxidant stress and function as CD36-dependent platelet-activating ligands, we measured circulating MP levels after FeCl3 injury by immunofluorescence microscopy and dot immunoblot assay. There was no difference in circulating endothelial-derived or total MP levels between WT and cd36–/– mice after sham operation (data not shown). Circulating levels of EMPs, however, increased in both WT and cd36–/– mice after carotid injury with 7.5% FeCl3, but as seen in the dot blot assay (Figure (Figure1E),1E), levels were significantly lower in cd36–/– mice. Immunofluorescence analysis of the MPs showed fewer total MPs and EMPs in the cd36–/– mice. Injury with 12.5% FeCl3 further increased circulating MPs, but no difference was seen between groups (data not shown). These results are consistent with our prior studies (3) showing decreased incorporation of endothelial antigens in clots induced in cd36–/– animals associated with protection from thrombosis after injury with 7.5% but not 12.5% FeCl3.
To determine potential mechanisms underlying the antioxidative effect of CD36 deletion in the vessel wall, we performed 2D difference gel electrophoresis comparing carotid artery segments from WT and cd36–/– mice. Proteins that were differentially expressed were then identified by mass spectrometry. One of the proteins with the greatest difference in expression (~2-fold) was the phase II antioxidant enzyme Prdx2, a member of a ubiquitous family of antioxidant enzymes known to detoxify ROS and reactive nitrogen oxide species (16, 17). To confirm differential Prdx2 expression, the contralateral noninjured carotid arteries from 18 experimental mice in which the left carotids were used for thrombosis assays were combined into 2 pools and analyzed by immunoblotting assay. As shown in Figure Figure2A,2A, Prdx2 levels were 3- to 4-fold higher in the cd36–/– mice than in the WT. Expression levels of glutathione peroxidase-1, catalase, and superoxide dismutase-1 did not differ between cd36–/– and WT carotid arteries (not shown). Immunohistochemical stains of the thrombosed carotid artery segments revealed that Prdx2 was mainly localized in the smooth muscle layer and that the staining intensity was markedly higher in the cd36–/– mice (Figure (Figure2B).2B). Prdx2 expression was also examined in other cells and tissues, including bone marrow, macrophages, platelets, and heart. Levels were markedly higher in hearts from cd36–/– mice and modestly increased in bone marrow–derived cells (Figure (Figure2,2, C–F), suggesting that CD36 may regulate Prdx2 levels in multiple cell types.
Immunohistochemical staining demonstrated that CD36 is abundantly expressed in the aortae (Figure (Figure3A)3A) and carotid arteries (not shown) of WT C57BL/6 mice. To explore mechanisms by which CD36 influences Prdx2 expression in the vessel wall, VSMCs were cultured from explants of thoracic aortic arteries of cd36–/– and WT mice (18) These cells showed characteristic staining for smooth muscle actin by immunocytofluorescence microscopy (not shown), and RT-PCR assays confirmed CD36 mRNA expression in WT but not in cd36–/– cells (Figure (Figure3B).3B). To demonstrate that the CD36 expressed by cultured VSMCs is functionally active, we showed that WT but not cd36–/– VSMCs bound and internalized 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate–labeled (DiI-labeled) NO2LDL, a form of oxLDL that is a highly specific CD36 ligand (not shown).
A cis-acting antioxidant responsive element (ARE) regulates transcription of many genes encoding detoxification enzymes and antioxidant proteins that function in cellular defense systems (19). The transcription factor Nrf2 is the major regulator of ARE-driven gene expression, but whether Nrf2 regulates Prdx2 transcription is not known. By analyzing the nucleotide sequences 10 kb upstream of the Prdx2 ATG translational start site, we identified a core ARE (cARE) site (TGACTCTGCA) at position –8518 to –8508, a so-called extended ARE (eARE) site (TGAGGTAGCA) at –7836 to –7826, and 2 “eARE-like” sites located between the core and extended sites (20), suggesting that Nrf2 may regulate Prdx2 expression (Supplemental Figure 1; supplemental material available online with this article; doi: 10.1172/JCI42823DS1). We therefore performed ChIP assays to determine whether Nrf2 binds to the putative ARE sites in the Prdx2 promoter using the cultured VSMCs. As shown in Figure Figure4A,4A, an antibody to Nrf2 precipitated the cARE and eARE sites from both WT and cd36–/– VSMCs, suggesting that Nrf2 may regulate Prdx2 expression. To determine whether Nrf2 binding to the Prdx2 promoter sites has functional consequences, an 869-bp region from the Prdx2 promoter that includes the cARE, eARE, and eARE-like sites (P6; Figure Figure4B),4B), as well as truncated constructs (P1–P5: Figure Figure4B),4B), was cloned into a luciferase reporter vector, pGL2-Basic, and transfected into WT VSMCs using Lipofectamine LTX and Plus Reagent. Luciferase reporter activities examined 48 hours later showed (Figure (Figure4C)4C) that constructs containing the eARE site but not the cARE site induced significant luciferase activity, indicating that the eARE alone is sufficient to induce transcription. Luciferase activity was increased by another 2-fold with P6, indicating that the cARE site may contribute to transcriptional activity, but the absence of luciferase activity with P1 suggests that cARE alone is not sufficient to induce transcription. The “eARE-like” sites had no activity, either alone (P2) or on combination with eARE (P5 vs. P3) or with cARE (P4 vs. P1). Mutations in the eARE site (Figure (Figure4B)4B) at sequences previously shown to inhibit activity in other systems (21) significantly diminished luciferase reporter activity (M1, M2, and M3; Figure Figure4C),4C), further demonstrating that a functional eARE site plays a key role in Nrf2-mediated Prdx2 expression.
We next sought to determine whether Nrf2 could induce Prdx2 expression in vascular cells. A plasmid expression vector encoding mouse Nrf2 was used to generate a pool of Nrf2-overexpressing mouse VSMCs, and immunoblots of lysates from these cells demonstrated dramatic upregulation of Prdx2 expression compared with that of cells transfected with a control plasmid, empty pCDNA3 (Figure (Figure4D). 4D).
Nrf2 is normally sequestered in the cytoplasm by a binding partner, Keap1, but activation signals including electrophiles and oxidant stress disrupt the Nrf2/Keap1 complex, allowing Nrf2 to translocate to the nucleus where it binds ARE sequences and induces transcription of specific genes. To determine whether CD36 signaling regulates Nrf2 and subsequently Prdx2 expression in the vessel wall, we first demonstrated by real-time PCR that prdx2 mRNA levels were significantly higher in cd36–/– VSMCs than in WT cells (Figure (Figure5A).5A). Immunoblot assays showed that Prdx2 and Nrf2 protein expression were also significantly higher in cd36–/– VSMCs compared with WT (Figure (Figure5B).5B). Importantly, cellular fractionation studies showed that Nrf2 accumulated in the nuclei of cd36–/– cells to a far greater extent than in WT cells (Figure (Figure5C).5C). Cytosolic Nrf2 levels were also slightly higher in cd36–/– VSMCs (Figure (Figure5D).5D). In these experiments, the efficacy of cell fractionation was documented using anti-TATA binding protein and anti–α-tubulin antibodies to detect nuclear and cytoplasmic proteins, respectively. To further demonstrate that deletion of cd36 induced Prdx2 overexpression via Nrf2 signaling, Nrf2 expression was knocked down in cd36–/– VSMCs using shRNA, and Prdx2 expression was shown to be significantly decreased (Figure (Figure5E).5E). These data suggest that CD36-mediated signaling may inhibit Nrf2 nuclear translocation and/or promote Nrf2 nuclear export and degradation.
A common theme in CD36 signal transduction in platelets, mononuclear phagocytes, and endothelial cells is activation of specific Src family kinases, including Fyn and Lyn (1, 22). Studies by Jain and Jaiswal in hepatoma cells demonstrated that Fyn-induced phosphorylation of Nrf2 at tyrosine 568 led to nuclear export and degradation (23). We thus hypothesized that CD36-dependent activation of Fyn in VSMCs would lead to Nrf2 degradation and hence decreased Prdx2 expression. To explore this hypothesis, we first examined whether inhibition of Src family kinase activity affected Nrf2 nuclear accumulation in VSMCs. As shown by immunofluorescence in Figure Figure6A6A (PP2), a broadly active Src family kinase inhibitor significantly increased nuclear Nrf2 in WT VSMCs, but caused less of an increase in cd36–/– cells, suggesting that Src kinases may play some role in Nrf2 nuclear export or degradation. The fact that cells treated with PP3, a nonactive control analog of PP2, showed significantly lower nuclear Nrf2 in WT VSMCs than in cd36–/– cells is consistent with the Western blot assay of nuclear/cytosol fractionation showed in Figure Figure5C.5C. Treatment of WT VSMCs with 1-(Palmitoyl)-2-(5-keto-6-octene-dioyl)phosphatidylcholine (KodiA-PC) (24), a specific CD36 ligand, dramatically increased Fyn phosphorylation (Figure (Figure6B),6B), suggesting functional CD36/Fyn signaling in VSMCs. Interestingly, tyrosine-phosphorylated Nrf2 also increased in the same manner (Figure (Figure6B).6B). These data strongly suggest that CD36 controls Nrf2 activity by regulating Fyn activity. In further support of this conclusion, we found that VSMCs cultured from thoracic aortae of fyn–/– mice markedly increased Nrf2 and Prdx2 expression compared with WT cells (Figure (Figure6C). 6C).
To determine whether the increase in Nrf2 activity associated with deletion of CD36 protects vascular cells from oxidative stress, VSMCs were serum starved for 24 hours and then reexposed to 10% serum in the presence or absence of 250 μM H2O2 for timed periods up to 6 hours. Figure Figure7A7A shows that H2O2 induced dramatically more HO-1 expression in cd36–/– cells compared with WT, with induction seen as early as 3 hours and increasing further at 6 hours. Lower (50 μM) or higher (1 mM) concentrations of H2O2 produced similar effects (not shown). These data suggest that CD36 expression inhibits the antioxidant system in response to oxidant stimulation.
To further demonstrate the prooxidant effect of CD36, VSMCs were treated with different concentration of FeCl3 and cellular ROS generation was quantified by HPLC detection of EOH (25, 26). FeCl3 increased superoxide generation in a dose-dependent manner, as has been previously reported (27). The level of EOH was significantly lower in FeCl3-treated cd36–/– cells compared with WT VSMCs and was no different in treated cd36–/– cells than untreated cells (Figure (Figure7B).7B). To show whether Prdx2 overexpression in WT VSMCs phenocopies cd36–/– cells, human Prdx2 cDNA was transfected into WT VSMCs and a cell pool was generated by G418 selection. Prdx2 overexpression significantly decreased FeCl3-induced superoxide anion generation when compared with WT VSMCs, almost to the level seen in cd36–/– (Figure (Figure7B).7B). In contrast, shRNA-based gene knockdown of nrf2 in cd36–/– cells significantly increased FeCl3-induced superoxide formation when compared with cd36–/– cells. These data show that both loss of cd36 and overexpression of Prdx2 provoke a strong oxidant effect and cd36–/– induced antioxidant effect via upregulation of Nrf2 and Prdx2.
To show that Prdx2 expression in the vessel wall may induce a protective effect in vivo, we measured ROS generation after FeCl3-induced carotid injury in prdx2–/– mice. As predicted, FeCl3 treatment induced more ROS generation in the vessel walls of prdx2–/– mice than in prdx2 WT mice (Figure (Figure7C).7C). Importantly, arterial thrombi formed more quickly and were larger in both prdx2- and nrf2–/– mice compared with WT, resulting in a significantly shorter time to occlusion (Figure (Figure7D). 7D).
Obstructive and thrombotic cardiovascular diseases are multifactorial and involve complex interplay among lifestyle (diet, smoking, exercise, ethanol consumption) and fixed (genotype, age, menopausal status, sex) causative factors (28). An initiating step is often endothelial injury induced by cellular oxidative stress precipitated by the production of damaging ROS by many cell types including VSMCs and monocytes/macrophages (28). Endothelial injury can promote thrombosis by multiple mechanisms, including generation of MPs. ROS also contribute to injury by oxidizing lipoproteins such as LDL within the vessel wall, facilitating uptake of these particles by macrophages. Under conditions in which lipid uptake cannot be balanced by natural efflux pathways, lipids accumulate in the cells, leading to formation of foam cells, a hallmark of early atherosclerotic lesions (1, 29, 30). Foam cells are also a source of inflammatory cytokines and further ROS production, contributing to lesion progression. CD36 plays several important roles in these processes (31–33). On macrophages and platelets, it is a critical receptor for oxLDL and MPs (1, 3) and its engagement by these ligands activates the cells, contributing to the thromboinflammatory state (1, 3, 4, 34). CD36-mediated oxLDL uptake by macrophages is essential for foam cell formation in vitro and in vivo and CD36 signaling in macrophages alters cytoskeletal dynamics, inhibiting migration and thus preventing egress of the cells from atheroinflammatory lesions (35).
To this point, evidence linking CD36 to human atherothrombotic disease includes reports that CD36 genetic polymorphisms are associated with myocardial infarction (36, 37) and that hyperlipidemia is associated with CD36-dependent platelet hyperreactivity (2). Numerous studies of mouse model systems have validated the importance of CD36 in vascular disease. cd36–/– mice show diminished atherosclerosis in both apoe- and ldlr–/– backgrounds (33, 38), demonstrate decreased ischemia/reperfusion injury after cerebral infarction (13), and are partially protected from pathological thrombosis in response to vascular injury (3). In this manuscript, we demonstrate what we believe is a novel mechanism by which CD36 participates in the vascular injury response and also for the first time, to our knowledge, demonstrate a biological function for CD36 on vascular smooth muscle cells. We found that CD36 mediates VSMC ROS generation in vitro and in vivo by downregulating expression of the redox-sensitive transcription factor Nrf2, which in turn leads to loss of expression of phase II antioxidant enzymes, including Prdx2 and HO-1.
Figure Figure88 outlines a model for this CD36-regulated pathway. Oxidative vascular injury, such as that induced by hyperlipidemia and inflammation in apoe–/– mouse atherosclerosis models or by topical FeCl3 application in commonly used murine thrombosis models, leads to dissociation of the transcription factor Nrf2 from its cytoplasmic partner, Keap1/INrf2. Nrf2 then enters the nucleus, where it binds to specific ARE elements in key antioxidant genes, including HO-1 and prdx2. Expression of these genes then attenuates oxidant stress. Oxidative stress, however, also leads to generation of specific CD36 ligands, such as MPs and oxidized phospholipids. Engagement of these ligands by CD36 recruits and activates Fyn kinase, which in turn phosphorylates Nrf2. Phosphorylated Nrf2 is transported out of the nucleus and degraded. CD36 in VSMCs thus sensitizes the cells to oxidant stress, contributing to atherogenesis and thrombosis. Our observation that prdx2–/– mice as well as nrf2–/– mice showed a prothrombotic phenotype after FeCl3-induced injury highlights the importance of this endogenous antioxidant system in vivo. Interestingly, we observed increased expression of Nrf2 and Prdx2 in VSMCs from fyn–/– mice, phenocopying the cd36 nulls and strongly supporting the model that CD36-mediated Fyn activation is a critical component of the pathway. The data presented in this paper also show enhanced expression of the antioxidant pathway in cd36–/– mice under basal conditions. This raises the possibility that some CD36-mediated effects are ligand independent, although we cannot rule out the presence of endogenous CD36 ligands in these studies.
Attenuation of endogenous antioxidant systems is probably not the sole mechanism by which CD36 promotes ROS generation. Studies from our lab and others strongly suggest that CD36 signaling in macrophages directly induces ROS formation; blockade of CD36 or inhibition of NADPH oxidase (Nox) decreased oxLDL-induced ROS formation (35, 39). The intracellular pathway from CD36 to Nox has not been delineated, but recent studies from our lab and others show that CD36 may activate Vav family guanine nucleotide exchange factors. These are known to influence activation of the small molecular weight G proteins associated with Nox.
Interestingly, several studies have reported that Nrf2 regulates CD36 expression in macrophages and preadipocytes in response to oxLDL stimulation (40–43), suggesting that oxidative stress might “feed forward” through CD36 to decrease natural antioxidant responses and promote further injury. A recent paper by Sussan et al., however, surprisingly showed that loss of Nrf2 was protective against development of atherosclerosis (44), perhaps related to decrease in macrophage CD36 expression. This observation is discordant with many reports supporting the hypothesis that prooxidant conditions promote atherosclerosis while antioxidant conditions protect. Indeed, a different group clearly demonstrated that macrophage Nrf2 expression did not promote atherosclerosis in hyperlipidemic ldlr–/– mice and that Nrf2 was atheroprotective in early stages of atherogenesis (45). Together, these data suggest that the cellular context of Nrf2 expression, its downstream activities, and the regulation of its activity are complex and that further studies are necessary to reveal the mechanistic connections among Nrf2, CD36, and vascular disease.
In summary, we have shown that deletion of CD36-attenuated ROS production in response to FeCl3 induced vascular injury in a mouse carotid artery thrombosis model and defined a CD36-dependent signaling pathway in VSMCs mediated by the redox-sensitive transcription factor Nrf2, which downregulates endogenous antioxidant defenses. Transfection of Nrf2 or its downstream targets (such as HO-1) has been shown to inhibit ROS generation in vitro and in vivo and to ameliorate atherosclerosis and obstructive vascular diseases (46–48). These results are consistent with our model and suggest that targeting CD36 could enhance the Nrf2 pathway and provide a novel strategy to restore natural antioxidant defenses in cardiovascular diseases.
Mice of various genetic backgrounds were anesthetized and subjected to left common carotid artery injury as previously described (3, 4) by topical application of a 1 × 2 mm piece of filter paper saturated with 7.5% or 12.5% FeCl3 for 1 minute. Clot formation was observed in real time by fluorescence microscopy, and time to complete cessation of blood flow was determined through visual inspection using video image capture as previously described (3, 4). The end points of occlusion were set as (a) cessation of blood flow for more than 30 seconds and (b) no occlusion seen over the entire 30-minute time of observation (time was recorded as 30 minutes). In some studies, hydroethidine (10 μg/g in 150 μl saline) with or without edaravone (1 μg/g) was injected into the right jugular vein 5 minutes prior to carotid injury to detect ROS formation in the vessel wall. Fluorescence was recorded by intravital microscopy with a fixed exposure time of 2 ms throughout the observation period.
All procedures and manipulations of animals were approved by institutional animal care and use committees (IACUCs) at Cleveland Clinic in accordance with the United States Public Health Service Policy on the Humane Care and Use of Animals and the NIH Guide for the care and use of laboratory animals (8th edition. Revised 2010).
After undergoing carotid artery thrombosis assays, mice were sacrificed and the thrombosed left carotid arteries were harvested, fixed in 4% formaldehyde overnight, and embedded in paraffin. Six-micrometer sections were cut, and immunohistochemical staining was performed for prdx2 expression. For frozen sections, the vessels were embedded in OCT and snap frozen in liquid nitrogen. For VSMC staining, cells were seeded in 4-well chamber glass slide (Lab-Tek), fixed in cold acetone (–20°C) for 10 minutes, washed with PBS, and permeabilized with 0.1% Triton X-100 in PBS for 5 minutes at room temperature. Slides were then incubated with primary antibodies followed by Alexa Fluor 488– or 594–conjugated secondary antibodies and then analyzed by laser confocal fluorescence microscopy using a Leica TCS-SP2 AOBS upright microscope. To quantify Nrf2 nuclear translocation, the fluorescence density of 15 randomly selected nuclei from 3 fields were analyzed by ImageJ software (NIH).
Heparin (100 USP in 100 μl saline) was injected into the right jugular vein prior to arterial injury to prevent thrombosis. After 1 minute application of FeCl3, whole blood was collected 5 minutes later by cardiac puncture. 300 μl was diluted in 700 μl PBS, then centrifuged at 4300 g for 5 minutes to remove cells and large cell fragments. The supernatants were centrifuged at 100,000 g for 120 minutes at 4°C to pellet MPs. Pelleted MPs were resuspended in 50 μl HEPES-Tyrode’s buffer and stored at –80°C (3). MP solutions (5 μl) were applied to siliconized glass slides and stained with FITC-conjugated anti–annexin V and anti-CD105 antibodies. Images were analyzed with laser confocal microscopy. MPs were also analyzed by a dot immunoblot assay using 10 μl of the MP solution and the same antibodies. Dot density was analyzed with NIH ImageJ.
The uninjured right carotid arteries of 18 WT and 18 cd36–/– mice that had been subjected to FeCl3-induced thrombosis in the contralateral carotid were harvested and pooled into 2 groups of 9; then proteins were extracted using the mirVana PARIS kit (Ambion). 100 μg from each extract was treated with the ReadyPrep 2-D Cleanup Kit (Bio-Rad). Proteins (50 μg) from WT mice were labeled with Cy3 (Invitrogen), those from cd36–/– mice with Cy5, and an internal control mixture from both mice (25 μg each) with Cy2. CyDye–labeled samples were mixed together, diluted to 450 μl with DeStreak Rehydration Solution (Amersham), and loaded onto a 24-cm IPG strip (pH 3-10 NL) gel for isoelectric focusing (IEF) using the Amersham IPGphor IEF system. The IPG strips were then reduced and alkylated and placed onto 24-cm Jule Pre-Cast Gels (12.5% SDS-PAGE) overlaid with 0.5% agarose in Laemmli buffer. Gels were run using an Ettan DALT 12 electrophoresis system. The 2D gels were scanned with the Typhoon imaging system and analyzed with DeCyder 6 software (GE). Gels were stained with Coomassie blue, and spots with more than 1.5-fold difference comparing WT and cd36 nulls identified by DeCyder 6 analysis were picked and subjected to mass spectrometry analysis with LCQ or LTQ ion trap systems (Thermo Fisher).
Proteins (40 μg) from carotid extracts or cultured VSMCs were electrophoresed in 7.5% to 12.5% SDS-PAGE gradient gels and transferred onto PVDF membranes. After blockade with 5% nonfat milk, membranes were incubated with antibodies and developed with Amersham ECL Plus Western Blotting Detection Reagents. Membranes were stripped and reblotted with other antibodies. An anti-actin antibody was used as a loading control. The density of each band was analyzed with ATTO Densitography.
Mouse VSMC cultures were established from thoracic aorta explants of 8-week-old male mice as previously described (18, 49). Cells were identified as VSMCs by immunocytochemistry using a monoclonal antibody to smooth muscle α-actin (A2547; Sigma-Aldrich). Cell passages 4 to 12 were used in this study. After treatment, cells were washed with cold PBS and lysed in 20 mM MOPS, pH 7.0; 2 mM EGTA; 5 mM EDTA; 30 mM sodium fluoride; 60 mM β-glycerophosphate, pH 7.2; 20 mM sodium pyrophosphate; 1 mM sodium orthovanadate; 1% Triton X-100 containing proteinase inhibitor cocktail (Roche). BioVision Nuclear/Cytosol Fractionation Kit (BioVision Research Products) was used for extraction of cytosol and nuclear proteins based on the manufacturer’s protocol.
LDL was isolated and purified from normal volunteers and was oxidized using a myeloperoxidase, glucose oxidase, nitrate system as previously described to generate NO2LDL (50). NO2LDL was labeled with the fluorescent probe Dil (Molecular Probes) as previously described (51). WT and cd36–/– VSMCs were seeded in 4-chamber slides at 103 cells/well, cultured for overnight, and then exposed to Dil-NO2 LDL (10 μg/ml) for 60 minutes, fixed in 10% formalin, and analyzed by laser confocal microscopy.
VSMCs were seeded at 2 × 106/10-cm plate, cultured overnight, and then synchronized for 24 hours prior to stimulation with DMEM containing 10% FCS for 3 hours. ChIP was performed using the EZchip Kit (Millipore) following the manufacturer’s protocol. Anti-Nrf2 antibody (sc-722; Santa Cruz Biotechnology Inc.) and normal rabbit IgG were used to immunoprecipitate DNA/protein complexes. Two pairs of primers were designed to amplify the Prdx2 promoter region containing the cARE (forward: GACAGGTGACAGGTGCTTTCAGAT; reverse: TCAGCATCCTCATAGCACCTTAGA) and eARE (forward: CCTTCCTGATGCTGCGACCCTTTA; reverse: CCCCAGGATAGCAGCGAGCATTCA).
An 869-bp region from the Prdx2 promoter that includes the cARE, eARE, and eARE-like sites as well as truncated constructs from the same region shown in Supplemental Figure 1 and Figure Figure4B4B was amplified with specific primers. For mutation studies, the eARE site (TGAGGTAGCA) was divided into 3 parts as shown in Figure Figure4B4B and the nucleotides in each part were changed to thymine in the reverse primers. All of the forward primers were linked with agctttaCTCGAG at the 5′ end containing an XhoI recognition site, and reverse primers were linked with actgcaataagctt at the 3′ end containing a HindIII recognition site. PCR products were cloned into a luciferase reporter vector, pGL2-Basic (Promega), and the plasmids then transformed into One Shot Top 10 competent cell (Invitrogen) and cultured and purified with QIAprep Spin Miniprep Kit (QIAGEN). 1 μg of each plasmid was transfected into WT VSMCs (cultured in 6-well) using Lipofectamine LTX and Plus Reagent (Invitrogen). Luciferase reporter activities were examined 48 hours later using an assay kit from Promega.
Total RNA was extracted from VSMCs using RNeasy Mini Kit (QIAGEN). 1 μg of the total RNA was treated with DNase I, and cDNA was constructed using AMV First Strand cDNA Synthesis Kit for RT-PCR (Roche). Real-time PCR was performed using SYBR Green PCR Master Mix (Applied Biosystems) with an iCycler iQ Real-Time PCR detection system (52).
The pCDNA3/mNrf2 plasmid vector (53), empty pCDNA3 vector, and pCR3.1/hPrdx2 plasmid (54) were transfected into WT VSMCs using the Lipofectamine LTX and Plus Reagent, and cells were selected with G418 antibiotic. A pool of Nrf2- or Prdx2-overexpressing cells as well as pcDNA empty vector control cells was established and expressions of Nrf2 as well as Prdx2 were examined by Western blot. In other experiments, mouse Nrf2 shRNA plasmid (Santa Cruz Biotechnology Inc.) was transfected into cd36–/– cells as above and a cell pool was selected with puromycin (7.5 μg/ml final concentration). Prdx2 expression after nrf2 gene knockdown in cd36–/– cells was examined by Western blot assay.
2 × 106 VSMCs were seeded on 10-cm plates and cultured for 24 hours. The cells were then treated with FeCl3 in a final concentration of 0.05% to 0.2% in culture media for 30 minutes at 37°C (27) and then washed with cold PBS 2 times and incubated in 3 ml Hanks’ buffer composed of 1.3 mM CaCl2, 0.8 mM MgSO4, 5.4 mM KCl, 0.4 mM KH2PO4, 4.3 mM NaHCO3, 137 mM NaCl, 0.3 mM Na2HPO4, and 5.6 mM glucose, pH 7.4, containing diethylenetriamine pentaacetic acid (DTPA) (100 μM) and a final DHE concentration of 50 μM for an additional 30 minutes at 37°C. Cells were washed twice with cold PBS, harvested in acetonitrile (1.5 ml/plate), sonicated (10 s, 1 cycle at 8 w), and centrifuged at 12,000 g for 10 minutes at 4°C (25). Supernatants were lyophilized to dryness and pellets maintained at –20°C in the dark until analyzed. Samples were resuspended in 150 μl PBS/DTPA (100 μM) and 100 μl aliquots injected into a 250 mm × 4.6 mm, 5 μm Kromasil C18 column on an HPLC system (Agilent 1100; Agilent Technologies). Solutions A (pure acetonitrile) and B (water/10% acetonitrile/0.1% trifluoroacetic acid) were used as a mobile phase at a flow rate of 0.5 ml/min (25). DHE was monitored by ultraviolet absorption at 245 nm. EOH and ethidium were monitored by fluorescence detection with excitation 510 nm and emission 595 nm. Quantification was performed by comparison of integrated peak areas between the obtained and standard solutions under identical chromatographic conditions. EOH standard was prepared as previously described (15).
One-way ANOVA (Bonferroni/Dunn) was used to determine the differences among groups. Unpaired t test was used to determine differences between groups. Data were represented as mean ± SEM, and P < 0.05 was considered significant. Each in vitro experiment was repeated as least 3 times with different passage cells.
This project was supported by NIH grants HL81011 and HL092747 (to R.L. Silverstein) and HL66109 (to S.P. Reddy). We thank Patricia DiBello in the Cleveland Clinic Department of Cell Biology for her assistance in HPLC analysis.
Conflict of interest: The authors have declared that no conflict of interest exists.
Citation for this article: J Clin Invest. 2010;120(11):3996–4006. doi:10.1172/JCI42823.