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CARDIOVASCULAR
and 
Ligands Differentially Affect Smooth Muscle Cell Proliferation and Migration
Institute of Cardiovascular Sciences, St. Boniface General Hospital Research Centre, Winnipeg, Manitoba, Canada (P.Z., B.W., M.F., N.Y., K.M.); and Departments of Human Nutritional Sciences (M.F., N.Y., C.G.T.) and Physiology (P.Z.), University of Manitoba, Winnipeg, Manitoba, Canada
Received September 26, 2005; accepted January 9, 2006.
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and 
are expressed in smooth muscle cells (SMCs). This study was designed to compare the effects of PPAR and PPAR on SMC proliferation and migration and to determine how they operate. Treatment of SMCs from porcine coronary artery revealed that mitogen-stimulated DNA synthesis was blocked by the PPAR ligand 4-chloro-6-(2,3-xylidino)-2-pyrimidinylthioacetic acid (WY14,643) and 15-deoxy-
12,14 prostaglandin J2 (15d-PGJ2) (a putative PPAR agonist) but not by the PPAR agonist rosiglitazone or the PPAR
/
ligand 2-methyl-4-((4-methyl-2-(4-trifluoromethylphenyl)-1,3-thiazol-5-yl)-methylsulfanyl)phenoxy acetic acid (GW501516). Inhibition of DNA synthesis by clofibrate and 2-(4-(2-(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2-methylproprionic acid (GW7647) confirmed that SMC proliferation is affected by PPAR. This conclusion was supported by the fact that WY14,643 also inhibited the proliferation of H4IIE hepatoma cells (expressing only PPAR) but not A10 SMCs (expressing only PPAR1). In contrast, the effective inhibition of all cell types with 15d-PGJ2 indicated that this compound probably operates via a PPAR-independent mechanism. Interestingly, rosiglitazone did not inhibit DNA synthesis of either H4IIE or A10 cells, suggesting that the activation of PPAR does not influence cell proliferation. Phosphorylation of cyclin-dependent kinase 2 and expression of proliferating cell nuclear antigen were inhibited by WY14,643 but not by rosiglitazone or 15d-PGJ2, indicating that PPAR prevents progression into S phase. Although rosiglitazone did not block SMC proliferation, it (like WY14,643) reduced neointimal hyperplasia in vitro. This observation can be rationalized by the fact that both WY14,643 and rosiglitazone inhibit SMC migration, probably through matrix metalloproteinase 9. Our study therefore shows that selective interference with mediators of cell cycle progression and cell migration via activation of PPARs may prevent growth-related vascular diseases such as restenosis and atherosclerosis.
). To date, three distinct PPARs, designated PPAR (NR1C1), PPAR (NR1C3), and PPAR/ (NR1C2), have been identified. It is now apparent from numerous studies that the PPARs can indirectly influence vascular disease (Lee et al., 2003). More intriguing, however, are two reports that suggest these proteins may affect vascular smooth muscle cells (SMCs) directly (Hu et al., 2002; Zahradka et al., 2003).
Although PPAR levels are much lower in vasculature tissue compared with hepatic and adipose tissues, each of the three PPAR isoforms has been detected in vascular SMCs (Marx et al., 2004). PPAR agonists have been shown to suppress the nuclear factor 
B-dependent induction of cyclooxygenase 2 and interleukin-6 in vascular SMCs (Staels et al., 1998). Because these observations suggest PPAR participates in the inflammatory response of SMCs to cytokines in addition to lipid and lipoprotein metabolism, there is speculation that PPAR agonists may be able to suppress progression of atherosclerotic lesions (Marx et al., 2004). In agreement with this prospect, PPAR agonists have been shown to inhibit the activation of nuclear factor B in SMCs and consequently decrease both chemokine secretion and matrix metalloproteinase expression (Chinetti et al., 2001). In addition, PPAR agonists can decrease both SMC migration and proliferation (Miwa et al., 2000; Gouni-Berthold et al., 2001). PPAR has therefore been associated with both atherogenesis and the response to injury. Although PPAR/ is expressed in SMCs, its function has yet to be established. Oliver et al. (2001), however, have reported PPAR/ may influence reverse cholesterol transport through its ability to regulate ABCA1 transporter expression.
There is support for the concept that PPARs are important factors in atherogenesis, and this view is substantiated by the results of a limited clinical study that showed that troglitazone, a PPAR agonist, reduced the thickness of the carotid artery after administration to patients with type 2 diabetes (Minamikawa et al., 1998). Furthermore, the inhibition of SMC proliferation and migration by PPAR agonists has led to speculation that these compounds may prevent intimal hyperplasia after revascularization (Bishop-Bailey et al., 2002). Similar data are now accumulating for PPAR, because the PPAR agonist WY14,643 inhibits mitogen-induced DNA synthesis (Hu et al., 2002; Zahradka et al., 2003). In contrast, expression of PPAR/ may promote SMC proliferation (Zhang et al., 2002).
In an earlier study, we observed that both WY14,643 (a PPAR agonist) and 15d-PGJ2 (a putative PPAR agonist) inhibited DNA synthesis after mitogen stimulation of human SMCs (Zahradka et al., 2003). These results led to the conclusion that these compounds might prove effective in preventing neointimal proliferation after vascular injury. However, a mechanism to explain the growth-inhibitory actions of PPAR agonists was lacking. We therefore examined the effect of PPAR agonists on several mediators of cell cycle progression in SMCs. Both retinoblastoma (Rb) protein and cdk2 were studied, based on evidence that Rb phosphorylation by cdk2 is required for the release of the E2F transcription factor (Andres, 2004) and subsequent expression of genes coding for proliferating cell nuclear antigen (PCNA) and cyclins E and A (Stevens and La Thangue, 2003). In addition, cyclin D levels and IKK (I B kinase) phosphorylation were monitored, because both have been shown essential for SMC proliferation (Zahradka et al., 2002; Andres, 2004). In the current investigation, we present data that confirm the antiproliferative activity of PPAR agonists and identify a target for their behavior. Furthermore, we demonstrate 15d-PGJ2 does not operate via PPAR. Finally, we establish that both PPAR and PPAR agonists block neointimal hyperplasia but that they operate via different mechanisms.
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Materials and Methods |
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). PCA-SMCs were propagated in Dulbecco's minimal essential medium (DMEM; Invitrogen, Burlington, ON, Canada) containing 20% FBS (Invitrogen). When 75% confluent, the growth medium was replaced with DMEM supplemented with 5 µg/ml transferrin, 1 nM selenium, 20 mM ascorbate, and 10 nM insulin for 5 days. A10 SMCs and H4IIE hepatoma cells were cultured as described previously (Saward and Zahradka, 1996; Yau et al., 1998). Quiescence was achieved by placing A10 and H4IIE cells into serum-free media for 72 h, with A10 cells receiving the same supplement as the PCA-SMCs. Quiescent cells were used for all growth assays, and PPAR agonists were added 60 min before mitogen stimulation. Specific agents used in these studies included WY14,643 (Cayman; Ann Arbor, MI), rosiglitazone [5-((4-(2-(methyl-2-pyridinylamino)ethoxy)phenyl)methyl)-2,4-thiazolidinedione; generously provided by SmithKline Beecham, Harlow, Essex, UK], 15d-PGJ2 (Cayman), GW501516 (Calbiochem, San Diego, CA), GW7647 (Calbiochem), and clofibrate [2-(4-chlorophenoxy)-2-methylpropanoic acid ethyl ester; Calbiochem].
DNA Synthesis. Triplicate sets of quiescent cells, prepared in 24-well dishes, were treated with mitogen ± PPAR agonist in the presence of 1 µCi [3H]thymidine (PerkinElmer Life and Analytical Sciences, Boston, MA). The labeling period used for each cell type was established previously (Saward and Zahradka, 1996; Yau et al., 1998). Incorporation of radiolabel into DNA was monitored by trichloroacetic acid precipitation.
Migration Assay. Migration of PCA-SMCs through polycarbonate filters with 5-µm pores was measured with a Boyden chamber (48-well unit) as described previously (Yau et al., 2003). Serum-free DMEM containing 0.1 µg/ml PDGF was placed in the lower compartment, whereas inhibitors were added to the upper compartment. After 48 h in a standard CO2 incubator, the cells on the underside of the membrane were visualized with Giemsa stain and quantified.
Reverse Transcription-Polymerase Chain Reaction Amplification. Total RNA was isolated from cells in six-well culture dishes or frozen white adipose tissue with TRIzol (Invitrogen). The RNA was resuspended in RNase-free water, and concentration was determined by spectrophotometric absorbance at 260 nm. Reverse transcription of 1 µg of RNA was conducted (after removal of possible genomic DNA contamination with DNase I) according to the protocol (62°C annealing temperature) recommended for the Access RT-PCR System (Promega, Madison, WI). The number of amplification cycles was empirically determined for each primer pair to identify the logarithmic phase. The specific forward and reverse oligodeoxynucleotide primers used were: GAPDH (s) 5'-CGCTGTGAACGGATTTGGCCGTAT-3', GAPDH (as) 5'-AGCCTTCTCCATGGTGGTGAAGAC-3'; rat PPAR (s) 5'-AAGACGCTTGTGGCCAAGAT-3', rat PPAR (as) 5'-ATGTCGCAGAATGGCTTCCT-3'; porcine PPAR (s) 5'-CGTGGCACTGAACATCGAAT-3', porcine PPAR (as) 5'-CGGTCTCGGCATCTTCTAGG-3'; rat PPAR1 (s) 5'-ACAAGACTACCCTTTACTGAAATTACC-3', rat PPAR1 (as) 5'-GTCTTCATAGTGTGGAGCAGAAATGCT-3'; porcine PPAR1 (s) 5'-CAGATTTGGTGGAAGCCAACT-3' porcine PPAR1 (as) 5'-CGTTTAAGGAAACAACCTTCCTG-3'; rat PPAR2 (s) 5'-TACAGCAAATCTCTGTTTTATGCTGTT-3', rat PPAR2 (as) 5'-GTCTTCATAGTGTGGAGCAGAAATGCT-3'; and porcine PPAR2 (s) 5'-GTTCCATGCTGTTATGGGTGAA-3', porcine PPAR2 (as) 5'-GCATCGCTTTCTGGGTCAAT-3' (Zahradka et al., 1995; Marx et al., 1998; Tanaka et al., 1999; Benson et al., 2000). Amplification products were analyzed by electrophoresis in 2% agarose gels, and the intensity of the Vistra Green-stained bands was quantified with a Storm 850 Imaging System and ImageQuant software (Amersham Biosciences Inc., Baie d'Urfé, QC, Canada). Control reactions (minus RNA, minus RT, and minus primers) were used to demonstrate the specificity of the PCR reaction.
Western Blot Analysis. Western blotting of cellular proteins (10 µg) separated by SDS-polyacrylamide gel electrophoresis in a 7.5% gel and transferred to a polyvinylidene difluoride membrane (Roche Diagnostics, Laval, QC, Canada) was conducted as described previously (Yau et al., 2003). Membranes were probed with antibodies to Thr-160 phospho-cdk2 (Cell Signaling Technology Inc., Beverly, MA), Ser-780 phospho-Rb (Cell Signaling), PCNA (Dako, Glostrup, Denmark), Ser-180/Ser-181 phospho-IKK/ (Cell Signaling), cyclin A (NeoMarkers, Fremont, CA), cyclin D (Upstate Biotechnology, Lake Placid, NY), cyclin E (Santa Cruz Biotechnology, Inc., Santa Cruz, CA), -tubulin (Sigma-Aldrich, St. Louis, MO), and horseradish peroxidase-conjugated secondary antibody (1:10,000 diluted) was detected using the ECL chemiluminescent system (Amersham).
Gelatin Zymography. The procedure was conducted with media samples recovered from organ culture after a 48-h incubation period as described previously (Zahradka et al., 2004).
Coronary Artery Organ Culture. Segments of porcine coronary artery, injured by inflation of an angioplasty catheter (3.5 x 20 mm) for 1 min, were cultured in 24-well dishes containing 20% FBS in DMEM as described previously (Wilson et al., 1999). Media, including treatments, were changed every 2nd day. Vessels harvested from culture were embedded, sectioned, and stained in Lee's methylene blue. Digital images were captured with a DAGE-MTI CCD camera (DAGE-MTI, Michigan City, IN) and analyzed with StainPoint software (Lynx Graphics Ltd., Winnipeg, MB, Canada; http://www.lynxgl.com) to quantify the neointimal index: intimal area/medial area.
Data Measurement and Statistical Analysis. Radiotracer assay data and densitometric scans of Western blots were quantified and plotted as means ± S.E.M. of individual experiments (n = 3). Morphometry was performed with eight replicates per treatment. Treatment means were compared using one-way analysis of variance, whereas all other data were analyzed with the unpaired Student's t test. Statistical significance was set at P < 0.05.
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agonist, or 15d-PGJ2, a putative PPAR agonist, significantly reduced the growth-stimulatory actions of PDGF, in accordance with data published for human vascular SMCs (Zahradka et al., 2003). In contrast, neither rosiglitazone nor GW501516 were capable of inhibiting DNA synthesis. Although these results were not surprising for a PPAR/ agonist such as GW501516, they were unexpected for the PPAR agonist rosiglitazone, given the potency demonstrated by 15d-PGJ2. We therefore explored in more detail the relationship between PPAR activation and cell proliferation.
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Activation of PPAR Blocks SMC Proliferation. The presence of PPAR in human and rodent SMCs (Diep et al., 2000; Zahradka et al., 2003) suggests that porcine SMCs should be responsive to PPAR agonists, and the results shown in Fig. 1 support this premise. Because WY14,643 has been described as a specific PPAR agonist (Lee et al., 1995), we examined its ability to block cell proliferation over a range of concentrations and with two distinct mitogens. Quiescent PCA-SMCs were stimulated with either PDGF-BB (0.1 µg/ml) or FBS (2%, v/v). Both mitogens significantly increased DNA synthesis, with FBS being considerably more potent than PDGF (2798 ± 660 versus 1072 ± 76%, respectively). In both cases, however, the addition of WY14,643 produced a concentration-dependent reduction in thymidine incorporation (Fig. 2, A and B). Interestingly, the effectiveness of WY14,643 varied with the two mitogens, because 250 µM was able to reduce DNA synthesis to near basal levels for PDGF-treated cells (with 100 µM producing a significant reduction), whereas a concentration of 500 µM was required for FBS-stimulated PCA-SMCs. This difference in efficacy was clearly evident upon comparison of the respective EC50 values for PDGF-(120 µM) and FBS (275 µM)-stimulated cells.
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and SMC proliferation, we tested two structurally distinct PPAR agonists. Like WY14,643, both clofibrate and GW7647 produced a concentration-dependent reduction in thymidine incorporation (EC50 values 245 and 3.2 µM, respectively) when added to PCA-SMCs stimulated with PDGF (Fig. 2, C and D). These data support the view that activation of PPAR can block SMC proliferation. Effect of PPAR Agonists on A10 SMCs and H4IIE Hepatomas. To determine whether inhibition of cell proliferation was a general property of PPAR agonists, we extended our study to include A10 SMCs and H4IIE hepatoma cells. Each cell type was stimulated with a mitogen (A10 cells with PDGF, H4IIE cells with insulin) that significantly increased thymidine incorporation by approximately 8- and 2-fold, respectively (Fig. 3). The addition of WY14,643 had no inhibitory effect on DNA synthesis after mitogen stimulation of A10 SMCs (Fig. 3A), but 250 µM WY14,643 significantly inhibited H4IIE cell proliferation (Fig. 3B). In contrast, 5 µM 15d-PGJ2 blocked the proliferation of both A10 SMCs and H4IIE hepatomas (Fig. 3, C and D). Interestingly, as seen with the PCA-SMCs (Fig. 1), the addition of 10 µM rosiglitazone had no effect on the proliferation of either A10 or H4IIE cell type (Fig. 3, E and F).
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, PPAR1, and PPAR2. It was observed that PPAR was present in PCA-SMCs (Fig. 4A) and H4IIE hepatomas (Fig. 4B) but not in A10 SMCs (Fig. 4C). In contrast, neither PPAR1 nor PPAR2 was expressed in H4IIE cells, although PPAR1 (but not PPAR2) was detected in both PCA and A10 SMCs (Fig. 4). RNA isolated from rat white adipose tissue was used as a positive control for PPAR2 (Fig. 4D). These results indicated there is a correlation between PPAR expression and inhibition of cell proliferation by WY14,643; however, the divergent actions of 15d-PGJ2 and rosiglitazone could not be traced to differences in PPAR expression.
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PPAR Activation Interferes with Cell Cycle Progression. A possible link between the putative PPAR agonist docosahexaenoic acid and cell cycle progression via cdk2 has been reported (Terano et al., 1999). We therefore examined the effect of PPAR agonists on the cell cycle mediators in quiescent PCA-SMCs stimulated with PDGF. It was observed that cdk2 phosphorylation was prevented by the addition of WY14,643 (Fig. 5A), whereas neither rosiglitazone nor 15d-PGJ2 inhibited this event. Likewise, the increase in PCNA elicited by PDGF was blocked by WY14,643 but not rosiglitazone or 15d-PGJ2 (Fig. 5B). In contrast, these compounds had no effect on PDGF-dependent phosphorylation of either Rb (Fig. 5C) or IKK (Fig. 5D). Likewise, PDGF-stimulated expression of cyclins D and A was unaffected by the PPAR agonists (Fig. 5E); however, cyclin E levels were reduced upon treatment with PGJ2 but not WY14,643 or rosiglitazone (Fig. 5F).
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). SMCs make a significant contribution to neointimal lesion formation after their transformation to the synthetic phenotype, which allows proliferation and migration. Based on the data presented above, WY14,643 would be expected to prevent neointimal hyperplasia after injury. In addition, because it has previously been reported that PPAR agonists can prevent neointimal hyperplasia (Law et al., 1996), rosiglitazone would be expected to perform similarly. We therefore used an organ culture model of balloon angioplasty (Wilson et al., 1999) to evaluate the actions of these compounds. Segments of porcine coronary arteries injured by balloon inflation were cultured for 14 days in the presence or absence of either 250 µM WY14,643 or 10 µM rosiglitazone. Control vessel segments that had not been injured were cultured in parallel. Morphometric analysis showed that the lesion of balloon-injured vessels was 3.8-fold larger than the control (Fig. 6). In the presence of WY14,643, the neointima was only 1.6-fold larger than control, a decline of 76%. Likewise, rosiglitazone decreased the neointimal area by 65%. These results confirm that both WY14,643 and rosiglitazone can inhibit neointimal formation after vascular injury.
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and PPAR Agonists Inhibit SMC Migration. Injury-induced neointimal proliferation requires both SMC proliferation and migration. Because our data suggest that rosiglitazone does not operate by inhibiting cell proliferation (Fig. 1), we investigated its ability to interfere with migration. Furthermore, because migration is dependent upon activation of specific cell cycle mediators, including cdk2 (Andres, 2004), we concurrently explored the possibility that WY14,643 could also impede SMC migration. PDGF-induced migration of PCA-SMCs was therefore quantified 48 h after seeding into a Boyden chamber. In the absence of a PPAR agonist, PDGF stimulated cell migration to the lower chamber of the apparatus; however, inclusion of WY14,643, 15d-PGJ2, or rosiglitazone significantly inhibited migration under these conditions (Fig. 7). In contrast, 10 µM GW501516 was ineffective. These results establish that agonists of PPAR and PPAR are potent inhibitors of SMC migration, but it is unlikely that PPAR/ has a role in this process.
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To investigate the possible mechanism by which migration is hindered, we examined the effect of the PPAR agonists on MMP2 and MMP9 production in organ culture. Injured vessels were placed into culture for 48 h in the presence of various agonists and release of MMP2 and MMP9 into the medium was measured by gelatin zymography (Zahradka et al., 2004). The levels of both latent and active MMP2 were unchanged relative to control except in the presence WY14,643, whereas MMP9 levels were reduced by both WY14,643 and rosiglitazone (Fig. 7, B and C). Interestingly, although 15d-PGJ2 was a potent inhibitor of migration, it did not affect MMP production. These results confirm that rosiglitazone and 15d-PGJ2 operate via distinct mechanisms and that 15d-PGJ2 probably does not function through PPAR.
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Discussion |
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with WY14,643 inhibits the proliferation of SMCs and extends these observations to cell migration. These results are supported by our finding that the PPAR ligands clofibrate, which is clinically used to reduce serum triglyceride levels, and GW7647 also prevent DNA synthesis by SMCs. We further demonstrated that the antiproliferative effects of these PPAR agonists are restricted to cells expressing PPAR, presumably by inhibiting cdk2 phosphorylation and PCNA expression. Although the PPAR agonist rosiglitazone was unable to block the proliferative response of SMCs to mitogens, it did inhibit cell migration, which is sufficient to prevent neointimal hyperplasia after vascular injury. Finally, our data support the view that the antiproliferative effect of 15d-PGJ2 is probably mediated via cyclin E but independent of PPAR. These results provide some clarification concerning the distinct physiological actions of PPAR and PPAR on vascular tissues.
In this study, we have established that activation of PPAR inhibits DNA synthesis, cell migration, and neointimal hyperplasia (Figs. 2, 6, and 7) using three distinct PPAR agonists for this purpose. This approach was necessitated by the fact that WY14,643 has been reported to both function as a highly selective PPAR agonist (Jiang et al., 1998; Seimandi et al., 2005) and to activate PPAR (Lehmann et al., 1997) when used at 100 µM. However, the similar response seen with the different PPAR ligands, as well as the ineffectiveness of rosiglitazone, supports the conclusion that was reached. Nonetheless, we did note that mitogenic potency apparently influences the degree of inhibition obtained with WY14,643, because a higher concentration was required to block the actions of FBS than PDGF. This observation may explain the variability in WY14,643 potency described in different publications.
Although there are few examples of a role for PPAR in SMC proliferation, Terano et al. (1999) reported the putative PPAR agonist docosahexaenoic acid blocks the phosphorylation of cdk2, a key event in the progression from G1 to S phase. We have observed that WY14,643 also inhibits cdk2 phosphorylation (Fig. 5A), which suggests this process may be the primary target for PPAR. cdk2 activation mediates transactivation of genes coding for proteins involved in DNA synthesis, including PCNA, which is responsible for recruiting DNA polymerase to the prereplication complex (Waga and Stillman, 1998). Because WY14,643 also suppressed PDGF-dependent induction of PCNA (Fig. 5B), inhibition of DNA synthesis by WY14,643 probably results from the absence of PCNA. Evidence of a relationship between cdk2 and PCNA (Sever-Chroneos et al., 2001; Starkel et al., 2005) supports this argument. Furthermore, the inability to phosphorylate cdk2 in the presence of WY14,643 suggests that this compound may operate through a cdk-activating kinase, a mechanism previously identified for mevastatin (Ukomadu and Dutta, 2003). The latter possibility would also explain why WY14,643 had no effect on cyclin expression (Fig. 5, E and F).
Recognition that PPARs are expressed in vascular SMCs (Marx et al., 2004) led to studies that examined whether PPAR agonists could positively affect vascular function directly, independent of their ability to decrease serum lipid levels. Benson et al. (2000) showed that activation of PPAR with troglitazone inhibited both SMC growth and migration. Similar data have been obtained for 15d-PGJ2 and other agonists belonging to the thiazolidinedione class (Law et al., 2000), thus providing a plausible mechanism to explain how troglitazone treatment prevented neointimal hyperplasia after balloon angioplasty of the rat carotid artery (Law et al., 1996). Based on these findings, it has been generally accepted that PPAR can modulate the proliferation of SMCs. On the other hand, the results of our study with rosiglitazone and 15d-PGJ2 suggest that PPAR may not influence SMC proliferation. This conclusion is based on the fact that H4IIE cells (chosen for comparative purposes because PPAR is strongly expressed in the liver) do not express PPAR1 (Fig. 3) yet still are sensitive to the actions of 15d-PGJ2. Therefore, the antiproliferative actions of 15d-PGJ2 cannot be mediated by PPAR, an argument that has been made by other investigators (Jozkowicz et al., 2001; Lennon et al., 2002). At the same time, none of the cell types we used exhibited sensitivity to rosiglitazone, a ligand that shows a high degree of specificity for PPAR (Seimandi et al., 2005). Therefore, it is unlikely that PPAR activation inhibits cell proliferation. Although this finding is not consistent with the conclusions reached in other studies, it should be recognized that many of the formative studies of PPAR function used 15d-PGJ2 and other agonists with uncertain specificity. On the other hand, it must be noted that the cells used for this investigation did not express detectable amounts of PPAR2 (Fig. 4), whereas it was strongly expressed in white adipose tissue, where the bulk of PPAR action has been reported. It is therefore possible that PPAR2 could be responsible for modulating cell proliferation in these tissues.
Although the role of PPAR/ in SMC proliferation has not been extensively examined, Zhang et al. (2002) have reported that PPAR/ activation promotes SMC proliferation. Our results with the selective PPAR/ agonist GW501516 (Seimandi et al., 2005), however, have shown that PPAR/ has no effect on SMC proliferation. Furthermore, GW501516 was unable to prevent PDGF-mediated SMC migration. These results clearly establish that PPAR activation is not a factor in the vascular response to injury.
Because neointimal hyperplasia requires both cell proliferation and migration (Newby and Zaltsman, 2000), and inhibition of either process is sufficient to prevent neointimal hyperplasia (Zahradka et al., 2004), the reduction in intimal lesion formation produced by WY14,643 (Fig. 6) was not surprising. On the other hand, the selective PPAR agonist rosiglitazone had no effect on DNA synthesis. This result was unexpected considering the general view that PPAR can modulate SMC proliferation (Marx et al., 2004). Nevertheless, the ability to prevent cell migration (Fig. 7A) could explain why rosiglitazone is an effective inhibitor of neointimal hyperplasia, regardless of its effect on proliferation, as we previously observed with the MMP inhibitor GM6001 (Zahradka et al., 2004). Interestingly, rosiglitazone and WY14,643 both reduced MMP9 production by vessels that had been subjected to injury by balloon inflation, as previously reported by Shu et al. (2000) for monocytic cells, yet only WY14,643 affected MMP2 (Fig. 7, B and C). These data support the argument that PPAR agonists impede neointimal hyperplasia by blocking cell migration and suggest that PPAR and PPAR probably operate via distinct pathways. In addition, the fact that 15d-PGJ2 had no effect on MMP levels, in conjunction with its unique action on cyclin E (Fig. 5), is strong evidence that this compound does not influence SMC proliferation and migration via PPAR.
The availability of cell lines that differentially express PPAR and PPAR has made it possible to distinguish the contribution of these isoforms to SMC proliferation. A10 SMCs may be a particularly interesting system for examining the details of PPAR function without the need to isolate cells from PPAR-null animals. The inhibitory effect of PPAR agonists on SMC proliferation sheds new light on the possible role of this receptor in vascular disease processes. Furthermore, we have identified a target, cdk2, which may be used to investigate the mechanism by which PPAR modulates both SMC proliferation and migration.
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Footnotes |
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ABBREVIATIONS: PPAR, peroxisome proliferator-activated receptor; SMC, smooth muscle cell; WY14,643, 4-chloro-6-(2,3-xylidino)-2-pyrimidinylthioacetic acid; 15d-PGJ2, 15-deoxy-12,14 prostaglandin J2; GW501516, 2-methyl-4-((4-methyl-2-(4-trifluoromethylphenyl)-1,3-thiazol-5-yl)-methylsulfanyl)phenoxy acetic acid; GW7647, 2-(4-(2-(1-cyclohexanebutyl-3-cyclohexylureido)ethyl)phenylthio)-2-methylproprionic acid; Rb, retinoblastoma; cdk2, cyclin-dependent kinase 2; PCNA, proliferating cell nuclear antigen; IKK, IB kinase; PCA-SMC, porcine coronary artery smooth muscle cell; DMEM, Dulbecco's minimal essential medium; FBS, fetal bovine serum; PDGF, platelet-derived growth factor; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; s, sense; as, antisense; MMP, matrix metalloproteinase; GM6001, N-[2(R)-2-(hydroxamido carbonylmethyl)-4-methylpentanoyl]-L-tryptophane methylamide.
Address correspondence to: Peter Zahradka, Institute of Cardiovascular Sciences, St. Boniface Research Centre, 351 Tache Avenue, Winnipeg, MB, Canada R2H 2A6. E-mail: peterz{at}sbrc.ca
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