Interferon Regulatory Factor-1 Mediates PPAR-Induced Apoptosis in Vascular Smooth Muscle Cells
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《动脉硬化血栓血管生物学》
From the Cardiovascular Research Institute (Y.L., F.L.M., H.X., J.Z., M.F., Y.E.C.), Morehouse School of Medicine, Atlanta, GA and the Cardiovascular Research Institute (X.Z., Y.E.C.), Peking University Health Science Center, Beijing 100083, China.
Correspondence to Yuqing E. Chen, Cardiovascular Research Institute, Morehouse School of Medicine, 720 Westview Drive SW, Atlanta, GA 30310. E-mail echen@msm.edu
Abstract
Objective— Peroxisome proliferator-activated receptor (PPAR) possesses general beneficial effects on the cardiovascular system, such as inhibition of vascular lesion formation and atherosclerosis. However, molecular mechanisms for these effects are yet to be fully defined. The aim of this study is to elucidate whether interferon regulatory factor-1 (IRF-1), a transcriptional factor with anti-proliferative and pro-apoptotic properties, mediates PPAR-induced apoptosis in vascular smooth muscle cells (VSMCs).
Methods and Results— Using Northern and Western blot analyses, we documented that PPAR ligands, including ciglitazone, troglitazone, and GW7845, significantly increased IRF-1 expression in VSMCs; however, the PPAR ligand (Wy14643) and PPAR ligand (GW0742) did not affect its expression. PPAR-induced IRF-1 expression was abrogated by pretreatment with the PPAR antagonist GW9662. In contrast, adenoviral expression of PPAR in VSMCs dramatically increased IRF-1 level. Furthermore, PPAR activation increased IRF-1 promoter activity but did not affect IRF-1 mRNA stability. Finally, reducing IRF-1 expression by antisense technology attenuated PPAR-induced VSMC apoptosis through decreasing cyclin-dependent kinase inhibitor p21cip1 and caspase-3 activity.
Conclusion— Our data demonstrate that IRF-1 is a novel PPAR target gene and mediates PPAR-induced VSMC apoptosis.
Key Words: peroxisome proliferator-activated receptor ? interferon regulatory factor-1 ? vascular smooth muscle cells ? gene expression ? apoptosis ? atherosclerosis
Introduction
Peroxisome proliferator-activated receptors (PPARs) including , , and /? are ligand-activated nuclear transcriptional factors. PPARs form heterodimers with retinoid X receptors (RXR), which are also members of nuclear factors, to bind to PPAR responsive element (PPRE), which either activates or represses its target genes.1 PPAR is most abundantly expressed in fat cells and has been linked to adipocyte differentiation and insulin sensitivity. The anti-diabetic "insulin-sensitizer" drug class, thiazolidinediones (TZDs), are pharmacological PPAR ligands.2 In addition, the endogenous ligands appear to be polyunsaturated fatty acids and eicosanoids, such as 15-deoxy--12, 14-prostaglandin J2 (15d-PGJ2).3
Mounting data suggest that PPAR is an important determinant of vascular structure and function.4 It is expressed in endothelial cells,5 vascular smooth muscle cells (VSMCs),6–8 monocytes/macrophages,9 and T lymphocytes.10 Recent studies documented that PPAR ligands inhibit atherosclerotic lesions11–13 and neointimal formation.14,15 TZDs have a pro-apoptotic action16–20 in addition to anti-proliferative,21–23 anti-inflammatory, 7,24 and anti-fibrotic25 effects in VSMCs. However, the molecular mechanisms for these effects in VSMCs are not quite fully defined, because the PPAR target genes in VSMCs are not well characterized. We recently performed cDNA microarrays to profile the changes in expression of thousands of mRNA in human aortic smooth muscle cells (HASMCs). More than 50 genes are either upregulated or downregulated by at least 2-fold after treatment with the TZDs.25 Interestingly, interferon regulatory factor-1 (IRF-1), a transcription factor, was one of the few upregulated genes by TZD treatment. This led us to hypothesize that TZDs increase IRF-1 gene expression via PPAR activation.
IRF-1 was initially identified as a regulator of interferon (IFN)-/? genes.26 It is well known that IRF-1 is able to promote apoptosis in many cell types,27–30 in addition to its pivotal role in anti-proliferation and differentiation. Intriguingly, Horiuchi et al reported that IRF-1 could upregulate interleukin-1?-converting enzyme expression and then induce apoptosis in VSMCs.31 Therefore, we hypothesized that PPAR activation might promote VSMC apoptosis by increasing IRF-1 expression.
It is important to note that a recent series of reports have documented that in the vascular system, TZDs can induce apoptosis in many cell types, including macrophages,32 lymphocytes,33 endothelial cells, 34,35 and VSMCs.16–20 However, whether TZD-induced apoptosis is a PPAR-dependent or independent effect is poorly understood. In the present study, we documented that TZDs could upregulate IRF-1 expression in VSMCs in a time- and dose-dependent manner. Using adenoviral-mediated PPAR overexpression and a PPAR antagonist, we revealed that IRF-1 is a novel PPAR target gene in VSMCs. In addition, reducing IRF-1 expression by antisense technology attenuated PPAR-induced VSMC apoptosis. Thus, our data suggest that the PPAR-regulated IRF-1 pathway is a newfound mechanism for VSMC survival.
Methods
Materials
Ciglitazone, troglitazone, and Wy14,643 were purchased from BioMol Research Laboratories (Plymouth Meeting, PA). GW7845, GW0742, and GW9662 were obtained from GlaxoSmithKline (Research Triangle Park, NC).3 TNF- and actinomycin D were purchased from Sigma. Hoechst 33342 and propidium iodide were purchased from Molecular Probes. Rabbit anti-human-IRF-1 (sc-497), rabbit anti-rat-IRF-1 (sc-640), rabbit anti-p21cip1 (sc-397), and anti-?-actin polyclonal antibody (sc-1616) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PPAR1 expression plasmid, pcDNA3.1-PPAR, was constructed in our laboratory as described previously.25 The pCMV-GFP plasmid was purchased from Invitrogen.
Cloning of Human IRF-1 Gene Promoter
Using the human IRF-1 mRNA sequence (GenBank Accession Number NM_002198) to blast the human genome database, we obtained the human IRF-1 gene sequence, including its promoter sequence (GenBank Accession Number NT_03477204). Based on this sequence information, we designed two primers to amplify a 2-kb human IRF-1 promoter by polymerase chain reaction. This human IRF-1 promoter (nt -1915 to nt +85) was cloned into a luciferase reporter plasmid (PGL3 basic; Promega, Madison, WI). The resultant plasmid was designated as pIRF-1 to 2kbLuc.
Adenovirus Recombination
The preparation and infection of the adenovirus containing PPAR or the control GFP were performed as described previously.36
Cell Culture
HASMCs were purchased from BioWhittaker (San Diego, CA) and cultured in smooth muscle cell growth medium-2 containing 5% FBS, 2 ng/mL human basic fibroblast growth factor, 0.5 ng/mL human epidermal growth factor, 50 μg/mL gentamicin, 50 ng/mL amphotericin B, and 5 μg/mL bovine insulin. For all experiments, passages 5 to 7 cells were grown to 80% to 90% confluence and made quiescent by Opti-MEM I (Invitrogen) for 24 hours. GW9662 was added to cells 1 hour before each PPAR agonist treatment. Actinomycin D was added 2 hours after ciglitazone treatment. Rat aortic vascular smooth muscle cells (RASMCs) were prepared from the thoracic aorta of 8- to 10-week-old male Sprague-Dawley rats using the explant technique. Relatively pure (>98%) RASMCs were confirmed by smooth muscle -actin staining (data not shown). The cells were cultured in DMEM/F12 (Invitrogen) containing 10% fetal bovine serum (FBS), 100 U/mL penicillin, 100 mg/mL streptomycin, and 1 mmol/L L-glutamine. Passages 8 to 10 cells were grown to 70% to 80% confluence and made quiescent by Opti-MEM I for 24 hours.
Northern Blot Analysis
Using RNeasy kit (Qiagen), 20 μg of total RNA was isolated from each condition and then subjected to electrophoresis through 1% formaldehyde-agarose gels. After transfer to nylon membranes (Perkin–Elmer Life Sciences), the RNA was cross-linked to the membrane by a UV cross-linker (Bio-Rad). 32P-Labeled cDNA probes were generated using the random primer labeling system (Life Technologies). Blots were pre-hybridized, hybridized, and washed once with 2x SSC at 65°C and once with 0.1x SSC, 1.0% SDS (wt/vol) at 65°C over the course of 15 minutes. A GAPDH cDNA probe was used to normalize for loadings.
Western Blot Analysis
Cells were washed two times by cold PBS and then lysed in solubilization buffer (20 mmol/L Tris·HCl, pH 7.5; 150 mmol/L NaCl; 1 mmol/L EDTA; 1 mmol/L EGTA; 1% Triton X-100; 2.5 mmol/L sodium pyrophosphate; 1 mmol/L sodium vanadate; 10 μg/mL each of aprotinin and leupeptin; 2 mmol/L phenylmethylsulfonyl fluoride). Also, 50 μg of total cell lysate was subjected to SDS-polyacrylamide gel electrophoresis and electrotransferred to PVDF membrane (Bio-Rad). After blocking in 20 mmol/L Tris·HCl, pH 7.6, containing 150 mmol/L NaCl, 0.1% Tween 20, and 5% (wt/vol) non-fat dry milk, blots were incubated with the specific first antibodies overnight at 4°C. The blots were then incubated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology). The immunoactivity was visualized through enhanced chemiluminescence detection system (Amersham Pharmacia Biotech) according to the manufacturer’s instructions. The lane loading differences were normalized using actin antibody.
Transient Transfection and Luciferase Assay
CV-1 cells (the African green monkey kidney fibroblast cell line from ATCC, Cat no.: CCL-70) were grown to 80% to 85% confluence in DMEM/F12 supplemented with 10% FBS and were transiently transfected in 24-well plates by using LipofectAMINE 2000 (Invitrogen) with reporter and expression plasmids as described. Green fluorescence protein (GFP) expression plasmid was co-transfected as the control for transfection efficiency. The total amount of transfected DNA was kept constant by using a corresponding empty vector DNA. Twenty-four hours after transfection, cells were cultured for 24 hours in Opti-MEM I and incubated for 24 hours in the same medium containing the appropriate reagents for the experiments. Each transfection was performed in triplicate at least 3 times. Reporter luciferase assay kit (Promega) was used to measure the luciferase activity in the cells according to the manufacturer’s instruction with a luminometer (Victor II, Perkin–Elmer). The luciferase activity was normalized by GFP.
Oligonucleotides and Transfection of RASMCs
Rat IRF-1 morpholino oligo antisense (rIRF-1-antisense), GAATGATGCCCGAGATGC (-111/-94),31 and its invert (rIRF-1-invert), CGTAGAGCCCGTAGTAAG, were purchased from Gene Tools (Philomath, OR). The oligonucleotides were transfected into RASMCs according to the manufacturer’s protocol.
Apoptosis Assay and Morphological Examination
Approximately, 2x105 RASMCs were seeded into a 12-well plate after transfection with rat IRF-1 invert and antisense oligonucleotides; the cells were cultured in growth medium for another 48 hours and then made quiescent using serum-free DMEM/F12 for 48 hours. The quiescent cells were treated with 5 μmol/L of ciglitazone for 24 hours or 72 hours and were then stained with Hoechst 33342 for apoptotic nuclear morphology.37,38 Caspase-3 activation was monitored by fluorometric assay kit (R&D Systems, Minneapolis, MN).
Statistical Analysis
Each experiment was repeated a minimum of 3 times. Statistical analyses were performed by analysis of variance and unpaired two-tailed Student’s t test. Data are presented as means ± SE. The value P<0.05 allows the data to be considered significant.
Results
Activation of PPAR Increases IRF-1 Expression in VSMCs
To characterize the role of PPAR in VSMCs, we globally profiled the gene expression in HASMCs in response to PPAR activators as we previously described.25 IRF-1, which is a well-characterized mediator in promoting VSMC apoptosis,31 was one of the few upregulated genes by TZD treatment (2.5-fold). This led us to postulate that IRF-1 might be a PPAR target gene in VSMCs.
We first examined whether PPAR activators regulate IRF-1 expression in HASMCs. Cells were treated with different PPAR activators, including ciglitazone (10 μmol/L), GW7845 (1 μmol/L), and troglitazone (10 μmol/L) for 2 hours in Northern blot analyses and for 24 hours in Western blot analyses. We documented that all PPAR activators tested significantly increased IRF-1 expression at both mRNA and protein levels, whereas the PPAR ligand Wy14643 (100 μmol/L) had no effect on IRF-1 expression (Figures 1 A and 1B). TNF- (10 ng/mL) was used as the positive control in this study. In addition, we found that PPAR-specific ligand GW0742 (0.2 μmol/L) did not change the expression of IRF-1 in HASMCs (data not shown).
Figure 1. PPAR activation increases IRF-1 expression in HASMCs. A and B, HASMCs were made quiescent by Opti-MEM I (Invitrogen) for 24 hours and then treated with the vehicle (0.1% DMSO, lane 1), ciglitazone (10 μmol/L, lane 2), GW7845 (1 μmol/L, lane 3), troglitazone (10 μmol/L, lane 4), Wy14,643 (100 μmol/L, lane 5), and TNF- (10 ng/mL, lane 6) for 2 hours for Northern blotting analyses (A) and 24 hours for Western blotting analyses (B). C, Representative Northern blot showing ciglitazone-induced IRF-1 expression in a time-course manner. D, Representative Northern blot showing ciglitazone-induced IRF-1 expression in a dose-dependent manner. The relative values normalized by GAPDH or ?-actin, and the control was assigned as an arbitrary value of 1 (n=3, *P<0.05 and **P<0.01 vs vehicle control).
IRF-1 is a phosphoprotein, and its dephosphorylated form has reduced DNA-binding activity in vitro.39 Our results show that PPAR activation increased both phosphorylated and dephosphorylated forms of IRF-1 protein in HASMCs (Figure 1B).
Similar results were obtained using RASMCs (data not shown).
Time Course and Dose-Dependent Effects of PPAR Activation on IRF-1 Gene Expression in VSMCs
HASMCs were treated with 10 μmol/L of ciglitazone for 0, 0.25, 0.5, 1, 2, 6, 12, and 24 hours. Afterwards, the levels of IRF-1 mRNA in the cells were determined by Northern blot analyses. As shown in Figure 1C, the levels of IRF-1 mRNAs induced by ciglitazone were dramatically increased at 1 hour, reached the maximal level at 2 hours, but decreased to some extent at 6 and 12 hours. The mRNA levels increased again at 24 hours. These levels are consistent with previous reports40,41 in which a biphasic expression of IRF-1 mRNA was also documented in VSMCs.
In addition, increasing the ciglitazone dose (1, 5, 10 μmol/L) upregulated IRF-1 mRNA in a dose-dependent manner after treatment for 2 hours (Figure 1D). Similarly, we found that other PPAR ligands, including GW7845 and troglitazone, induced IRF-1 expression in a time- and dose-dependent manner (data not shown). Taken together, our data reveal that PPAR activation upregulates IRF-1 gene expression in VSMCs.
Upregualtion of IRF-1 Expression by TZD Is Mediated by PPAR
Recent reports suggest that PPAR agonists in the TZD class have various PPAR-independent effects in many cell types.42–44 To determine if the upregulation of IRF-1 expression in the present study is mediated by PPAR in HASMCs, the cells were pretreated with or without GW9662, a PPAR-specific antagonist, at 1 μmol/L for 1 hour before the addition of 10 μmol/L of ciglitazone. As shown in Figure 2A, inhibition of PPAR by GW9662 abolished the ciglitazone-induced IRF-1 expression. In addition, increased IRF-1 expression by other PPAR ligands such as GW7845 (1 μmol/L), a high-affinity and non-TZD PPAR agonist, was also abrogated by GW9662 (Figure 2B)
Figure 2. PPAR antagonist abrogates PPAR-induced IRF-1 expression in HASMCs. Cells were pretreated with or without GW9662 (1 μmol/L) for 1 hour before the addition of 10 μmol/L of ciglitazone (A) or 1 μmol/L of GW7845 (B) for 2-hour stimulation. Total RNA was isolated from the treated cells for Northern blotting analyses. The relative IRF-1 mRNA levels were normalized by GAPDH, and the control was assigned as an arbitrary value of 1 (n=3).
To further determine that PPAR directly upregulates IRF-1 expression in HASMCs, the cells were infected with PPAR adenovirus at 2 or 5 plaque forming units (pfu)/cell for 2 hours. The GFP adenovirus was used as the control. The infected cells were cultured in growth medium for 24 hours before they were switched to Opti-MEM I for another 24 hours. The quiescent cells were then treated with ciglitazone (10 μmol/L) or the vehicle (0.1% of DMSO) for 2 hours. As shown in Figure 3, adenoviral expression of PPAR significantly upregulated IRF-1 mRNA level in a dose-dependent manner. After treatment with ciglitazone for PPAR adenovirus-infected HASMCs, IRF-1 mRNA levels increased dramatically compared with the treatment with DMSO. Taken together, these data suggest that TZD-induced IRF-1 expression in VSMCs is mediated by PPAR.
Figure 3. Overexpression of PPAR dramatically increased IRF-1 expression in HASMCs. Cells were infected with PPAR adenovirus (2 or 5 pfu/cell) or Ad-GFP (5 pfu/cell) for 2 hours and cultured in growth medium for 24 hours before they were switched to Opti-MEM I for another 24 hours. The quiescent cells were then treated with ciglitazone (10 μmol/L) or the vehicle (0.1% of DMSO) for 2 hours. Northern blot analyses were performed to analyze IRF-1 mRNA levels. The relative IRF-1 mRNA levels were normalized by GAPDH, and the control was assigned as an arbitrary value of 1 (n=3, **P<0.01 vs Ad-GFP without ciglitazone stimulation).
Upregulation of IRF-1 mRNA Expression by PPAR Activation Does Not Affect the Stability of IRF-1 mRNA
To evaluate whether IRF-1 mRNA stability contributes to the upregulation of IRF-1 mRNA expression by PPAR activation, we examined the half-life of IRF-1 mRNA in HASMCs (Figure I, available online at http://atvb.ahajournals.org). Northern blot analysis was performed with the addition of actinomycin D (5 μg/mL) after 2 hours of ciglitazone (10 μmol/L) or vehicle stimulation. Consistent with the previous report,45 we documented that the half-life of IRF-1 mRNA was 30 minutes. In addition, there was no significant difference between ciglitazone-treated and DMSO-treated cells. These data imply that PPAR-induced IRF-1 expression in VSMCs occurs mainly at the transcriptional level.
PPAR Activation Increases the IRF-1 Promoter Activity
To further examine the mechanisms by which PPAR activation upregulates IRF-1 expression, we investigated the regulation of the IRF-1 promoter in CV-1, a widely used cell line for testing PPAR-mediated transcription. Without transfecting the PPAR expression plasmid, treatment of CV-1 cells with ciglitazone (5 μmol/L) had no effect on IRF-1 promoter activation. When PPAR expression plasmid (100 ng/well) was transfected into these cells, ciglitazone strongly increased IRF-1 promoter activation (Figure II, available online at http://atvb.ahajournals.org). In addition, similar results were obtained using other PPAR agonists, including troglitazone and GW7845 (data not shown). Therefore, our results indicate that PPAR activation increases IRF-1 expression at the transcriptional level in a PPAR-dependent mechanism.
IRF-1 Mediates the VSMC Apoptosis Induced by PPAR Activation
Although it has been documented that PPAR activation can induce VSMC apoptosis,16–20 the molecular mechanisms behind the process are poorly understood. Identification of IRF-1 as a novel PPAR target gene in the present study led us to postulate that IRF-1 might mediate the VSMC apoptosis induced by PPAR activation. To address this hypothesis, we used a highly characterized antisense method to reduce the IRF-1 expression in VSMCs. The IRF-1 antisense oligonucleotide and its control were synthesized as previously described.31 The successful transfection into RASMCs was documented by Western blot analyses. As shown in Figure 4A, the IRF-1 antisense significantly reduced basal and ciglitazone-induced (5 μmol/L) IRF-1 expression in VSMCs. In addition, the p21cip1, a well-characterized IRF-1 target gene, was also significantly reduced in the IRF-1 antisense-treated VSMCs. Intriguingly, we found that RASMCs transfected with IRF-1 antisense exhibited a marked reduction in apoptosis induced by PPAR activation using the nuclear chromatin morphology analyses (Figure 4B), the apoptotic cells exhibiting the brightly fluorescent, condensed, and coalesced chromatin staining patterns. In addition, similar results were obtained by FACS analyses (data not shown). Consistent with the apoptotic morphology analysis, we further documented that the reduction in IRF-1 expression in RASMCs significantly decreased caspase 3 activity induced by ciglitazone stimulation (Figure 4C). Hence, our data demonstrate that IRF-1 mediates PPAR-induced VSMC apoptosis.
Figure 4. IRF-1 mediates VSMC apoptosis induced by ciglitazone. A, RASMCs were successfully transfected with IRF-1 antisense or IRF-1-invert oligonucleotides. The expression levels of IRF-1 and its target gene, p21cip1, were documented by Western blot analyses. The 72-hour serum-freed cells were treated with ciglitazone (5 μmol/L) or vehicle (DMSO) for 24 hours. The values were normalized by ?-actin, and the control was assigned as an arbitrary value of 1. B, The 72-hour treated RASMCs, to be analyzed by nuclear chromatin morphology, were stained with Hoechst 33342 and assessed for the characteristic condensed, coalesced, chromatin pattern of apoptotic cells. C, The samples for Western blot analyses in A were used for caspase-3 activity assay. The relative values were expressed as means ± SD (n=3), and the control (IRF-1-invert with DMSO treatment) was assigned as an arbitrary value of 1. Three independent experiments showed similar results.
Discussion
PPAR is highly expressed in human46 and murine atherosclerotic lesions,47 as well as in the neointimal smooth muscle cells of rat arteries after balloon angioplasty injury.11–13 In this study, we documented that IRF-1 is a novel PPAR target gene in VSMCs, and suggested that the upregulation of IRF-1 by PPAR activation might be one of molecular mechanisms for PPAR-inhibiting neointimal formation and atherosclerotic lesions.
Recent studies suggest that PPAR is expressed in the vasculature, and increasing evidence indicates an important role of PPAR activation in the cardiovascular system. When activated by TZDs, the anti-diabetic "insulin-sensitizer" drugs, PPAR inhibits the development of atherosclerotic lesions11–13 and neointimal formation in vivo.14,15 The proposed anti-proliferative, 21–23 anti-inflammatory,7,24 and anti-fibrotic25 effects help to explain the protective roles of PPAR in vasculature.4 In addition, it has been recently documented that PPAR activated by TZDs induced VSMC apoptosis;16–20 however, the underlying molecular mechanisms are unclear.
It has been well documented that IRF-1 regulates the transcription of an array of target genes that are involved in apoptosis, such as interleukin-1? converting enzyme,27 iNOS,48,49 angiotensin II type 2 receptor,50 caspase 1 and caspase 7, and p21Cip1.30 Intriguingly, Horiuchi et al studied the effect of serum withdrawal on RASMCs and observed the upregulation of IRF-1.31 Using antisense treatment, they demonstrated that IRF-1 inhibition attenuated apoptotic changes. In addition, Wessely et al recently reported that IRF-1 is highly expressed in human vascular lesions and exhibits growth inhibition in coronary artery smooth muscle cells and functions as a negative mediator that inhibits neointimal formation in the mouse carotid ligation model.51 Thus, IRF-1 is a crucial mediator in VSMC apoptosis and plays a critical role in vascular lesion formation. Indeed, we have identified for the first time that IRF-1 mediates TZD-induced VSMC apoptosis by PPAR. Interestingly, IRF-2, which is another isoform of IRF family, has been documented to countervail IRF-1-induced apoptosis in many cell types, including VSMCs.31,50 In the present study, we did not find that TZD stimulation had any effect on IRF-2 expression in VSMCs (data not shown).
The cyclin-dependent kinase inhibitor p21cip1 is an IRF-1 target gene in VSMCs.51 Several reports suggested that p21cip1 could function as negative regulator of VSMC growth.37,52–54 In this study, we documented that PPAR activation by TZDs increased p21cip1 expression (Figure 4), suggesting that the molecular link of PPARIRF-1p21cip1 pathway leads to the activation of caspase 3 and apoptosis in VSMCs. However, a recent report documented that TZD could inhibit PDGF-stimulated p21cip1 expression by protein kinase C (PKC) and may subsequently block survival signaling and promote apoptosis.55 It should be pointed out that TZD treatment could induce VSMC apoptosis by upregulation51 or downregulation55 of p21cip1. Although LaBaer et al suggested that the cell concentration of p21cip1 may be one explanation of why p21cip1 can function as either a positive or a negative regulator of growth,56 we believe there could be a more complicated explanation thus requiring further studies.
TZDs could increase the expression of p53 and Gadd45, which are two apoptotic genes, and then induce VSMC apoptosis.17 However, whether the increased expression of Gadd45 was directly regulated by PPAR remains unknown. It was noted that TZD-induced Gadd45 expression required at least 12 hours of treatment in VSMCs, suggesting that PPAR indirectly regulated Gadd45. Indeed, a recent article demonstrated that the transcriptional factor Oct-1 was responsible for PPAR-induced Gadd45 expression, suggesting that the PPAROct-1Gadd45 pathway significantly contributes to TZD-induced apoptosis.20 Our data demonstrated that the PPARIRF-1p21cip1 pathway regulates VSMC apoptosis. Interestingly, we retrieved the human GADD45 gene sequence, including its promoter, from GenBank (Accession Number AC046149) and found one consensus IRF-1 binding site (caaaactGAAAat) located between nt-1667 and nt-1655 within its 3-kb promoter by computer analyses. Thus, IRF-1 may regulate GADD45 gene expression. Further studies are required to explore the interactions and relative contributions among these pathways.
Although it was reported that docosahexaenoic acid, a low-affinity ligand of PPAR, could also induce VSMC apoptosis,57 the very potent PPAR agonist Wy14,643 and several known natural PPAR agonists were not able to induce the morphological changes of apoptosis in RASMCs, nor did they increase caspase-3 activity.58 In this report, we demonstrated that PPAR activation does not change IRF-1 expression. We can thus conclude that activation of PPAR is most likely not responsible for VSMC apoptosis caused by docosahexaenoic acid.57
We cloned the 2-kb human IRF-1 promoter and documented that PPAR activation increased the IRF-1 promoter activity. In addition, PPAR activation did not affect IRF-1 mRNA stability. Taken together, our data suggest that PPAR-induced IRF-1 expression in VSMCs occurs at the transcriptional level. However, computer analysis of this 2-kb human IRF-1 promoter did not revealed any putative PPAR response element. Currently, we are performing systematic deletion mapping of this IRF-1 promoter to characterize the PPAR response sequence.
In summary, our data suggest that IRF-1 is a novel PPAR target gene in VSMCs and that PPAR-induced VSMC apoptosis by the PPARIRF-1p21cip1 pathway provides a new insight into understanding the roles of PPAR in vasculature. The characterization of these PPAR signaling pathways in the vasculature may have important implications for the treatment of vascular diseases.
Acknowledgments
We thank Dr Timothy M. Willson at GlaxoSmithKline for the PPAR ligands. This work was partially supported by NIH grants HL068878, HL03676, S06GM08248, and G12-RR03034. Y. L. (grant no. 0225323B) and M. F. (grant no. 0225214B) are supported by American Heart Association Southeast Affiliate Fellowships.
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Correspondence to Yuqing E. Chen, Cardiovascular Research Institute, Morehouse School of Medicine, 720 Westview Drive SW, Atlanta, GA 30310. E-mail echen@msm.edu
Abstract
Objective— Peroxisome proliferator-activated receptor (PPAR) possesses general beneficial effects on the cardiovascular system, such as inhibition of vascular lesion formation and atherosclerosis. However, molecular mechanisms for these effects are yet to be fully defined. The aim of this study is to elucidate whether interferon regulatory factor-1 (IRF-1), a transcriptional factor with anti-proliferative and pro-apoptotic properties, mediates PPAR-induced apoptosis in vascular smooth muscle cells (VSMCs).
Methods and Results— Using Northern and Western blot analyses, we documented that PPAR ligands, including ciglitazone, troglitazone, and GW7845, significantly increased IRF-1 expression in VSMCs; however, the PPAR ligand (Wy14643) and PPAR ligand (GW0742) did not affect its expression. PPAR-induced IRF-1 expression was abrogated by pretreatment with the PPAR antagonist GW9662. In contrast, adenoviral expression of PPAR in VSMCs dramatically increased IRF-1 level. Furthermore, PPAR activation increased IRF-1 promoter activity but did not affect IRF-1 mRNA stability. Finally, reducing IRF-1 expression by antisense technology attenuated PPAR-induced VSMC apoptosis through decreasing cyclin-dependent kinase inhibitor p21cip1 and caspase-3 activity.
Conclusion— Our data demonstrate that IRF-1 is a novel PPAR target gene and mediates PPAR-induced VSMC apoptosis.
Key Words: peroxisome proliferator-activated receptor ? interferon regulatory factor-1 ? vascular smooth muscle cells ? gene expression ? apoptosis ? atherosclerosis
Introduction
Peroxisome proliferator-activated receptors (PPARs) including , , and /? are ligand-activated nuclear transcriptional factors. PPARs form heterodimers with retinoid X receptors (RXR), which are also members of nuclear factors, to bind to PPAR responsive element (PPRE), which either activates or represses its target genes.1 PPAR is most abundantly expressed in fat cells and has been linked to adipocyte differentiation and insulin sensitivity. The anti-diabetic "insulin-sensitizer" drug class, thiazolidinediones (TZDs), are pharmacological PPAR ligands.2 In addition, the endogenous ligands appear to be polyunsaturated fatty acids and eicosanoids, such as 15-deoxy--12, 14-prostaglandin J2 (15d-PGJ2).3
Mounting data suggest that PPAR is an important determinant of vascular structure and function.4 It is expressed in endothelial cells,5 vascular smooth muscle cells (VSMCs),6–8 monocytes/macrophages,9 and T lymphocytes.10 Recent studies documented that PPAR ligands inhibit atherosclerotic lesions11–13 and neointimal formation.14,15 TZDs have a pro-apoptotic action16–20 in addition to anti-proliferative,21–23 anti-inflammatory, 7,24 and anti-fibrotic25 effects in VSMCs. However, the molecular mechanisms for these effects in VSMCs are not quite fully defined, because the PPAR target genes in VSMCs are not well characterized. We recently performed cDNA microarrays to profile the changes in expression of thousands of mRNA in human aortic smooth muscle cells (HASMCs). More than 50 genes are either upregulated or downregulated by at least 2-fold after treatment with the TZDs.25 Interestingly, interferon regulatory factor-1 (IRF-1), a transcription factor, was one of the few upregulated genes by TZD treatment. This led us to hypothesize that TZDs increase IRF-1 gene expression via PPAR activation.
IRF-1 was initially identified as a regulator of interferon (IFN)-/? genes.26 It is well known that IRF-1 is able to promote apoptosis in many cell types,27–30 in addition to its pivotal role in anti-proliferation and differentiation. Intriguingly, Horiuchi et al reported that IRF-1 could upregulate interleukin-1?-converting enzyme expression and then induce apoptosis in VSMCs.31 Therefore, we hypothesized that PPAR activation might promote VSMC apoptosis by increasing IRF-1 expression.
It is important to note that a recent series of reports have documented that in the vascular system, TZDs can induce apoptosis in many cell types, including macrophages,32 lymphocytes,33 endothelial cells, 34,35 and VSMCs.16–20 However, whether TZD-induced apoptosis is a PPAR-dependent or independent effect is poorly understood. In the present study, we documented that TZDs could upregulate IRF-1 expression in VSMCs in a time- and dose-dependent manner. Using adenoviral-mediated PPAR overexpression and a PPAR antagonist, we revealed that IRF-1 is a novel PPAR target gene in VSMCs. In addition, reducing IRF-1 expression by antisense technology attenuated PPAR-induced VSMC apoptosis. Thus, our data suggest that the PPAR-regulated IRF-1 pathway is a newfound mechanism for VSMC survival.
Methods
Materials
Ciglitazone, troglitazone, and Wy14,643 were purchased from BioMol Research Laboratories (Plymouth Meeting, PA). GW7845, GW0742, and GW9662 were obtained from GlaxoSmithKline (Research Triangle Park, NC).3 TNF- and actinomycin D were purchased from Sigma. Hoechst 33342 and propidium iodide were purchased from Molecular Probes. Rabbit anti-human-IRF-1 (sc-497), rabbit anti-rat-IRF-1 (sc-640), rabbit anti-p21cip1 (sc-397), and anti-?-actin polyclonal antibody (sc-1616) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PPAR1 expression plasmid, pcDNA3.1-PPAR, was constructed in our laboratory as described previously.25 The pCMV-GFP plasmid was purchased from Invitrogen.
Cloning of Human IRF-1 Gene Promoter
Using the human IRF-1 mRNA sequence (GenBank Accession Number NM_002198) to blast the human genome database, we obtained the human IRF-1 gene sequence, including its promoter sequence (GenBank Accession Number NT_03477204). Based on this sequence information, we designed two primers to amplify a 2-kb human IRF-1 promoter by polymerase chain reaction. This human IRF-1 promoter (nt -1915 to nt +85) was cloned into a luciferase reporter plasmid (PGL3 basic; Promega, Madison, WI). The resultant plasmid was designated as pIRF-1 to 2kbLuc.
Adenovirus Recombination
The preparation and infection of the adenovirus containing PPAR or the control GFP were performed as described previously.36
Cell Culture
HASMCs were purchased from BioWhittaker (San Diego, CA) and cultured in smooth muscle cell growth medium-2 containing 5% FBS, 2 ng/mL human basic fibroblast growth factor, 0.5 ng/mL human epidermal growth factor, 50 μg/mL gentamicin, 50 ng/mL amphotericin B, and 5 μg/mL bovine insulin. For all experiments, passages 5 to 7 cells were grown to 80% to 90% confluence and made quiescent by Opti-MEM I (Invitrogen) for 24 hours. GW9662 was added to cells 1 hour before each PPAR agonist treatment. Actinomycin D was added 2 hours after ciglitazone treatment. Rat aortic vascular smooth muscle cells (RASMCs) were prepared from the thoracic aorta of 8- to 10-week-old male Sprague-Dawley rats using the explant technique. Relatively pure (>98%) RASMCs were confirmed by smooth muscle -actin staining (data not shown). The cells were cultured in DMEM/F12 (Invitrogen) containing 10% fetal bovine serum (FBS), 100 U/mL penicillin, 100 mg/mL streptomycin, and 1 mmol/L L-glutamine. Passages 8 to 10 cells were grown to 70% to 80% confluence and made quiescent by Opti-MEM I for 24 hours.
Northern Blot Analysis
Using RNeasy kit (Qiagen), 20 μg of total RNA was isolated from each condition and then subjected to electrophoresis through 1% formaldehyde-agarose gels. After transfer to nylon membranes (Perkin–Elmer Life Sciences), the RNA was cross-linked to the membrane by a UV cross-linker (Bio-Rad). 32P-Labeled cDNA probes were generated using the random primer labeling system (Life Technologies). Blots were pre-hybridized, hybridized, and washed once with 2x SSC at 65°C and once with 0.1x SSC, 1.0% SDS (wt/vol) at 65°C over the course of 15 minutes. A GAPDH cDNA probe was used to normalize for loadings.
Western Blot Analysis
Cells were washed two times by cold PBS and then lysed in solubilization buffer (20 mmol/L Tris·HCl, pH 7.5; 150 mmol/L NaCl; 1 mmol/L EDTA; 1 mmol/L EGTA; 1% Triton X-100; 2.5 mmol/L sodium pyrophosphate; 1 mmol/L sodium vanadate; 10 μg/mL each of aprotinin and leupeptin; 2 mmol/L phenylmethylsulfonyl fluoride). Also, 50 μg of total cell lysate was subjected to SDS-polyacrylamide gel electrophoresis and electrotransferred to PVDF membrane (Bio-Rad). After blocking in 20 mmol/L Tris·HCl, pH 7.6, containing 150 mmol/L NaCl, 0.1% Tween 20, and 5% (wt/vol) non-fat dry milk, blots were incubated with the specific first antibodies overnight at 4°C. The blots were then incubated with horseradish peroxidase-conjugated secondary antibody (Santa Cruz Biotechnology). The immunoactivity was visualized through enhanced chemiluminescence detection system (Amersham Pharmacia Biotech) according to the manufacturer’s instructions. The lane loading differences were normalized using actin antibody.
Transient Transfection and Luciferase Assay
CV-1 cells (the African green monkey kidney fibroblast cell line from ATCC, Cat no.: CCL-70) were grown to 80% to 85% confluence in DMEM/F12 supplemented with 10% FBS and were transiently transfected in 24-well plates by using LipofectAMINE 2000 (Invitrogen) with reporter and expression plasmids as described. Green fluorescence protein (GFP) expression plasmid was co-transfected as the control for transfection efficiency. The total amount of transfected DNA was kept constant by using a corresponding empty vector DNA. Twenty-four hours after transfection, cells were cultured for 24 hours in Opti-MEM I and incubated for 24 hours in the same medium containing the appropriate reagents for the experiments. Each transfection was performed in triplicate at least 3 times. Reporter luciferase assay kit (Promega) was used to measure the luciferase activity in the cells according to the manufacturer’s instruction with a luminometer (Victor II, Perkin–Elmer). The luciferase activity was normalized by GFP.
Oligonucleotides and Transfection of RASMCs
Rat IRF-1 morpholino oligo antisense (rIRF-1-antisense), GAATGATGCCCGAGATGC (-111/-94),31 and its invert (rIRF-1-invert), CGTAGAGCCCGTAGTAAG, were purchased from Gene Tools (Philomath, OR). The oligonucleotides were transfected into RASMCs according to the manufacturer’s protocol.
Apoptosis Assay and Morphological Examination
Approximately, 2x105 RASMCs were seeded into a 12-well plate after transfection with rat IRF-1 invert and antisense oligonucleotides; the cells were cultured in growth medium for another 48 hours and then made quiescent using serum-free DMEM/F12 for 48 hours. The quiescent cells were treated with 5 μmol/L of ciglitazone for 24 hours or 72 hours and were then stained with Hoechst 33342 for apoptotic nuclear morphology.37,38 Caspase-3 activation was monitored by fluorometric assay kit (R&D Systems, Minneapolis, MN).
Statistical Analysis
Each experiment was repeated a minimum of 3 times. Statistical analyses were performed by analysis of variance and unpaired two-tailed Student’s t test. Data are presented as means ± SE. The value P<0.05 allows the data to be considered significant.
Results
Activation of PPAR Increases IRF-1 Expression in VSMCs
To characterize the role of PPAR in VSMCs, we globally profiled the gene expression in HASMCs in response to PPAR activators as we previously described.25 IRF-1, which is a well-characterized mediator in promoting VSMC apoptosis,31 was one of the few upregulated genes by TZD treatment (2.5-fold). This led us to postulate that IRF-1 might be a PPAR target gene in VSMCs.
We first examined whether PPAR activators regulate IRF-1 expression in HASMCs. Cells were treated with different PPAR activators, including ciglitazone (10 μmol/L), GW7845 (1 μmol/L), and troglitazone (10 μmol/L) for 2 hours in Northern blot analyses and for 24 hours in Western blot analyses. We documented that all PPAR activators tested significantly increased IRF-1 expression at both mRNA and protein levels, whereas the PPAR ligand Wy14643 (100 μmol/L) had no effect on IRF-1 expression (Figures 1 A and 1B). TNF- (10 ng/mL) was used as the positive control in this study. In addition, we found that PPAR-specific ligand GW0742 (0.2 μmol/L) did not change the expression of IRF-1 in HASMCs (data not shown).
Figure 1. PPAR activation increases IRF-1 expression in HASMCs. A and B, HASMCs were made quiescent by Opti-MEM I (Invitrogen) for 24 hours and then treated with the vehicle (0.1% DMSO, lane 1), ciglitazone (10 μmol/L, lane 2), GW7845 (1 μmol/L, lane 3), troglitazone (10 μmol/L, lane 4), Wy14,643 (100 μmol/L, lane 5), and TNF- (10 ng/mL, lane 6) for 2 hours for Northern blotting analyses (A) and 24 hours for Western blotting analyses (B). C, Representative Northern blot showing ciglitazone-induced IRF-1 expression in a time-course manner. D, Representative Northern blot showing ciglitazone-induced IRF-1 expression in a dose-dependent manner. The relative values normalized by GAPDH or ?-actin, and the control was assigned as an arbitrary value of 1 (n=3, *P<0.05 and **P<0.01 vs vehicle control).
IRF-1 is a phosphoprotein, and its dephosphorylated form has reduced DNA-binding activity in vitro.39 Our results show that PPAR activation increased both phosphorylated and dephosphorylated forms of IRF-1 protein in HASMCs (Figure 1B).
Similar results were obtained using RASMCs (data not shown).
Time Course and Dose-Dependent Effects of PPAR Activation on IRF-1 Gene Expression in VSMCs
HASMCs were treated with 10 μmol/L of ciglitazone for 0, 0.25, 0.5, 1, 2, 6, 12, and 24 hours. Afterwards, the levels of IRF-1 mRNA in the cells were determined by Northern blot analyses. As shown in Figure 1C, the levels of IRF-1 mRNAs induced by ciglitazone were dramatically increased at 1 hour, reached the maximal level at 2 hours, but decreased to some extent at 6 and 12 hours. The mRNA levels increased again at 24 hours. These levels are consistent with previous reports40,41 in which a biphasic expression of IRF-1 mRNA was also documented in VSMCs.
In addition, increasing the ciglitazone dose (1, 5, 10 μmol/L) upregulated IRF-1 mRNA in a dose-dependent manner after treatment for 2 hours (Figure 1D). Similarly, we found that other PPAR ligands, including GW7845 and troglitazone, induced IRF-1 expression in a time- and dose-dependent manner (data not shown). Taken together, our data reveal that PPAR activation upregulates IRF-1 gene expression in VSMCs.
Upregualtion of IRF-1 Expression by TZD Is Mediated by PPAR
Recent reports suggest that PPAR agonists in the TZD class have various PPAR-independent effects in many cell types.42–44 To determine if the upregulation of IRF-1 expression in the present study is mediated by PPAR in HASMCs, the cells were pretreated with or without GW9662, a PPAR-specific antagonist, at 1 μmol/L for 1 hour before the addition of 10 μmol/L of ciglitazone. As shown in Figure 2A, inhibition of PPAR by GW9662 abolished the ciglitazone-induced IRF-1 expression. In addition, increased IRF-1 expression by other PPAR ligands such as GW7845 (1 μmol/L), a high-affinity and non-TZD PPAR agonist, was also abrogated by GW9662 (Figure 2B)
Figure 2. PPAR antagonist abrogates PPAR-induced IRF-1 expression in HASMCs. Cells were pretreated with or without GW9662 (1 μmol/L) for 1 hour before the addition of 10 μmol/L of ciglitazone (A) or 1 μmol/L of GW7845 (B) for 2-hour stimulation. Total RNA was isolated from the treated cells for Northern blotting analyses. The relative IRF-1 mRNA levels were normalized by GAPDH, and the control was assigned as an arbitrary value of 1 (n=3).
To further determine that PPAR directly upregulates IRF-1 expression in HASMCs, the cells were infected with PPAR adenovirus at 2 or 5 plaque forming units (pfu)/cell for 2 hours. The GFP adenovirus was used as the control. The infected cells were cultured in growth medium for 24 hours before they were switched to Opti-MEM I for another 24 hours. The quiescent cells were then treated with ciglitazone (10 μmol/L) or the vehicle (0.1% of DMSO) for 2 hours. As shown in Figure 3, adenoviral expression of PPAR significantly upregulated IRF-1 mRNA level in a dose-dependent manner. After treatment with ciglitazone for PPAR adenovirus-infected HASMCs, IRF-1 mRNA levels increased dramatically compared with the treatment with DMSO. Taken together, these data suggest that TZD-induced IRF-1 expression in VSMCs is mediated by PPAR.
Figure 3. Overexpression of PPAR dramatically increased IRF-1 expression in HASMCs. Cells were infected with PPAR adenovirus (2 or 5 pfu/cell) or Ad-GFP (5 pfu/cell) for 2 hours and cultured in growth medium for 24 hours before they were switched to Opti-MEM I for another 24 hours. The quiescent cells were then treated with ciglitazone (10 μmol/L) or the vehicle (0.1% of DMSO) for 2 hours. Northern blot analyses were performed to analyze IRF-1 mRNA levels. The relative IRF-1 mRNA levels were normalized by GAPDH, and the control was assigned as an arbitrary value of 1 (n=3, **P<0.01 vs Ad-GFP without ciglitazone stimulation).
Upregulation of IRF-1 mRNA Expression by PPAR Activation Does Not Affect the Stability of IRF-1 mRNA
To evaluate whether IRF-1 mRNA stability contributes to the upregulation of IRF-1 mRNA expression by PPAR activation, we examined the half-life of IRF-1 mRNA in HASMCs (Figure I, available online at http://atvb.ahajournals.org). Northern blot analysis was performed with the addition of actinomycin D (5 μg/mL) after 2 hours of ciglitazone (10 μmol/L) or vehicle stimulation. Consistent with the previous report,45 we documented that the half-life of IRF-1 mRNA was 30 minutes. In addition, there was no significant difference between ciglitazone-treated and DMSO-treated cells. These data imply that PPAR-induced IRF-1 expression in VSMCs occurs mainly at the transcriptional level.
PPAR Activation Increases the IRF-1 Promoter Activity
To further examine the mechanisms by which PPAR activation upregulates IRF-1 expression, we investigated the regulation of the IRF-1 promoter in CV-1, a widely used cell line for testing PPAR-mediated transcription. Without transfecting the PPAR expression plasmid, treatment of CV-1 cells with ciglitazone (5 μmol/L) had no effect on IRF-1 promoter activation. When PPAR expression plasmid (100 ng/well) was transfected into these cells, ciglitazone strongly increased IRF-1 promoter activation (Figure II, available online at http://atvb.ahajournals.org). In addition, similar results were obtained using other PPAR agonists, including troglitazone and GW7845 (data not shown). Therefore, our results indicate that PPAR activation increases IRF-1 expression at the transcriptional level in a PPAR-dependent mechanism.
IRF-1 Mediates the VSMC Apoptosis Induced by PPAR Activation
Although it has been documented that PPAR activation can induce VSMC apoptosis,16–20 the molecular mechanisms behind the process are poorly understood. Identification of IRF-1 as a novel PPAR target gene in the present study led us to postulate that IRF-1 might mediate the VSMC apoptosis induced by PPAR activation. To address this hypothesis, we used a highly characterized antisense method to reduce the IRF-1 expression in VSMCs. The IRF-1 antisense oligonucleotide and its control were synthesized as previously described.31 The successful transfection into RASMCs was documented by Western blot analyses. As shown in Figure 4A, the IRF-1 antisense significantly reduced basal and ciglitazone-induced (5 μmol/L) IRF-1 expression in VSMCs. In addition, the p21cip1, a well-characterized IRF-1 target gene, was also significantly reduced in the IRF-1 antisense-treated VSMCs. Intriguingly, we found that RASMCs transfected with IRF-1 antisense exhibited a marked reduction in apoptosis induced by PPAR activation using the nuclear chromatin morphology analyses (Figure 4B), the apoptotic cells exhibiting the brightly fluorescent, condensed, and coalesced chromatin staining patterns. In addition, similar results were obtained by FACS analyses (data not shown). Consistent with the apoptotic morphology analysis, we further documented that the reduction in IRF-1 expression in RASMCs significantly decreased caspase 3 activity induced by ciglitazone stimulation (Figure 4C). Hence, our data demonstrate that IRF-1 mediates PPAR-induced VSMC apoptosis.
Figure 4. IRF-1 mediates VSMC apoptosis induced by ciglitazone. A, RASMCs were successfully transfected with IRF-1 antisense or IRF-1-invert oligonucleotides. The expression levels of IRF-1 and its target gene, p21cip1, were documented by Western blot analyses. The 72-hour serum-freed cells were treated with ciglitazone (5 μmol/L) or vehicle (DMSO) for 24 hours. The values were normalized by ?-actin, and the control was assigned as an arbitrary value of 1. B, The 72-hour treated RASMCs, to be analyzed by nuclear chromatin morphology, were stained with Hoechst 33342 and assessed for the characteristic condensed, coalesced, chromatin pattern of apoptotic cells. C, The samples for Western blot analyses in A were used for caspase-3 activity assay. The relative values were expressed as means ± SD (n=3), and the control (IRF-1-invert with DMSO treatment) was assigned as an arbitrary value of 1. Three independent experiments showed similar results.
Discussion
PPAR is highly expressed in human46 and murine atherosclerotic lesions,47 as well as in the neointimal smooth muscle cells of rat arteries after balloon angioplasty injury.11–13 In this study, we documented that IRF-1 is a novel PPAR target gene in VSMCs, and suggested that the upregulation of IRF-1 by PPAR activation might be one of molecular mechanisms for PPAR-inhibiting neointimal formation and atherosclerotic lesions.
Recent studies suggest that PPAR is expressed in the vasculature, and increasing evidence indicates an important role of PPAR activation in the cardiovascular system. When activated by TZDs, the anti-diabetic "insulin-sensitizer" drugs, PPAR inhibits the development of atherosclerotic lesions11–13 and neointimal formation in vivo.14,15 The proposed anti-proliferative, 21–23 anti-inflammatory,7,24 and anti-fibrotic25 effects help to explain the protective roles of PPAR in vasculature.4 In addition, it has been recently documented that PPAR activated by TZDs induced VSMC apoptosis;16–20 however, the underlying molecular mechanisms are unclear.
It has been well documented that IRF-1 regulates the transcription of an array of target genes that are involved in apoptosis, such as interleukin-1? converting enzyme,27 iNOS,48,49 angiotensin II type 2 receptor,50 caspase 1 and caspase 7, and p21Cip1.30 Intriguingly, Horiuchi et al studied the effect of serum withdrawal on RASMCs and observed the upregulation of IRF-1.31 Using antisense treatment, they demonstrated that IRF-1 inhibition attenuated apoptotic changes. In addition, Wessely et al recently reported that IRF-1 is highly expressed in human vascular lesions and exhibits growth inhibition in coronary artery smooth muscle cells and functions as a negative mediator that inhibits neointimal formation in the mouse carotid ligation model.51 Thus, IRF-1 is a crucial mediator in VSMC apoptosis and plays a critical role in vascular lesion formation. Indeed, we have identified for the first time that IRF-1 mediates TZD-induced VSMC apoptosis by PPAR. Interestingly, IRF-2, which is another isoform of IRF family, has been documented to countervail IRF-1-induced apoptosis in many cell types, including VSMCs.31,50 In the present study, we did not find that TZD stimulation had any effect on IRF-2 expression in VSMCs (data not shown).
The cyclin-dependent kinase inhibitor p21cip1 is an IRF-1 target gene in VSMCs.51 Several reports suggested that p21cip1 could function as negative regulator of VSMC growth.37,52–54 In this study, we documented that PPAR activation by TZDs increased p21cip1 expression (Figure 4), suggesting that the molecular link of PPARIRF-1p21cip1 pathway leads to the activation of caspase 3 and apoptosis in VSMCs. However, a recent report documented that TZD could inhibit PDGF-stimulated p21cip1 expression by protein kinase C (PKC) and may subsequently block survival signaling and promote apoptosis.55 It should be pointed out that TZD treatment could induce VSMC apoptosis by upregulation51 or downregulation55 of p21cip1. Although LaBaer et al suggested that the cell concentration of p21cip1 may be one explanation of why p21cip1 can function as either a positive or a negative regulator of growth,56 we believe there could be a more complicated explanation thus requiring further studies.
TZDs could increase the expression of p53 and Gadd45, which are two apoptotic genes, and then induce VSMC apoptosis.17 However, whether the increased expression of Gadd45 was directly regulated by PPAR remains unknown. It was noted that TZD-induced Gadd45 expression required at least 12 hours of treatment in VSMCs, suggesting that PPAR indirectly regulated Gadd45. Indeed, a recent article demonstrated that the transcriptional factor Oct-1 was responsible for PPAR-induced Gadd45 expression, suggesting that the PPAROct-1Gadd45 pathway significantly contributes to TZD-induced apoptosis.20 Our data demonstrated that the PPARIRF-1p21cip1 pathway regulates VSMC apoptosis. Interestingly, we retrieved the human GADD45 gene sequence, including its promoter, from GenBank (Accession Number AC046149) and found one consensus IRF-1 binding site (caaaactGAAAat) located between nt-1667 and nt-1655 within its 3-kb promoter by computer analyses. Thus, IRF-1 may regulate GADD45 gene expression. Further studies are required to explore the interactions and relative contributions among these pathways.
Although it was reported that docosahexaenoic acid, a low-affinity ligand of PPAR, could also induce VSMC apoptosis,57 the very potent PPAR agonist Wy14,643 and several known natural PPAR agonists were not able to induce the morphological changes of apoptosis in RASMCs, nor did they increase caspase-3 activity.58 In this report, we demonstrated that PPAR activation does not change IRF-1 expression. We can thus conclude that activation of PPAR is most likely not responsible for VSMC apoptosis caused by docosahexaenoic acid.57
We cloned the 2-kb human IRF-1 promoter and documented that PPAR activation increased the IRF-1 promoter activity. In addition, PPAR activation did not affect IRF-1 mRNA stability. Taken together, our data suggest that PPAR-induced IRF-1 expression in VSMCs occurs at the transcriptional level. However, computer analysis of this 2-kb human IRF-1 promoter did not revealed any putative PPAR response element. Currently, we are performing systematic deletion mapping of this IRF-1 promoter to characterize the PPAR response sequence.
In summary, our data suggest that IRF-1 is a novel PPAR target gene in VSMCs and that PPAR-induced VSMC apoptosis by the PPARIRF-1p21cip1 pathway provides a new insight into understanding the roles of PPAR in vasculature. The characterization of these PPAR signaling pathways in the vasculature may have important implications for the treatment of vascular diseases.
Acknowledgments
We thank Dr Timothy M. Willson at GlaxoSmithKline for the PPAR ligands. This work was partially supported by NIH grants HL068878, HL03676, S06GM08248, and G12-RR03034. Y. L. (grant no. 0225323B) and M. F. (grant no. 0225214B) are supported by American Heart Association Southeast Affiliate Fellowships.
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