Transcription Factor CHF1/Hey2 Regulates Neointimal Formation In Vivo and Vascular Smooth Muscle Proliferation and Migration In Vitro
Objective— To determine the role of the cardiovascular-restricted, hairy-related bHLH transcription factor, CHF1/Hey2, in the biological response to vascular injury.
Methods and Results— We investigated the response of CHF1/Hey2-deficient mice to vascular injury in vivo and the response of primary cultured vascular smooth muscle cells (VSMCs) from these mice to growth factors in vitro. Neointima formation after arterial wire injury is decreased in knockout (KO) compared with wild-type (WT) mice (0.025±0.011 mm2 in WT [n=13]) versus 0.016±0.008 mm2 in KO (n=12; P<0.05) and is accompanied by reduced cellular proliferation. CHF1/Hey2-deficient VSMCs proliferate slowly compared with WT VSMCs and also show decreased migration in response to platelet-derived growth factor (PDGF) (62.6±10.3 CPF versus 37.2±13.5 CPF; P<0.01) and heparin-binding epidermal growth factor-like growth factor (HB-EGF) (27.4±7.7 CPF versus 6.4±3.7 CPF, P<0.05). Furthermore, lamellipodia formation and membrane ruffling induced by these chemoattractants are diminished in KO VSMCs, which is correlated with decreased activation of the small GTPase Rac1. Although total Rac1 protein was not changed in KO VSMCs, the level of the Rac guanine exchange factor (GEF), Sos1, was decreased.
Conclusions— CHF1/Hey2 is an important regulator of vascular smooth muscle cell (VSMC) accumulation during vascular remodeling and responsiveness to growth factors in vitro.
The vascular smooth muscle cell (VSMC) is a major component of the arterial wall and plays a critical role in the development of occlusive vascular lesions.1 In normal vessels, VSMCs are quiescent, differentiated, contractile, and function to maintain vascular tone and blood pressure. In response to injury, VSMCs undergo a phenotypic transition whereby they proliferate, migrate from the medial layer to the intima, secrete matrix metalloproteinases, decrease their expression of contractile proteins, and increase their expression of extracellular matrix molecules. The critical steps that regulate this phenotypic transition are not completely understood.
Previously, we and other groups cloned the bHLH protein CHF1/Hey2.2–7 CHF1/Hey2 (also known as Hesr-2, HRT2, HERP1, and gridlock) is one of only a few tissue-restricted bHLH transcription factors, along with relatives CHF2/Hey1 and CHF3/HeyL, expressed in VSMCs. Studies of the zebrafish homologue of CHF1/Hey2, gridlock, suggested a role in development of the aorta6 and in patterning of the arterial and venous systems.8 Other studies have suggested that CHF1/Hey2 gene expression decreases after vascular injury and growth factor stimulation, and that immortalized smooth muscle cell lines overexpressing the related gene CHF2/Hey1 (also known as HRT1 and Hesr-1) show enhanced smooth muscle cell growth and protection against apoptosis.9,10 CHF2/Hey1 has also been suggested to play a role in vascular endothelial cell differentiation.11 Despite these interesting findings about CHF1/Hey2 and its relative CHF2/Hey1, the function of CHF1/Hey2 in the adult vasculature has not been demonstrated in vivo by loss of function studies.
We and others have reported that CHF1/Hey2-deficient mice have ventricular septal defects and cardiomyopathy.12–14 In all cases, no anatomic vascular defect was reported. In this study, we evaluated these mice for a functional vascular phenotype. Our results show that neointima formation after arterial wire injury was markedly diminished in knockout (KO) mice when compared with wild-type (WT) mice and that CHF1/Hey2-deficient VSMCs show decreased proliferation and ability to migrate in response to platelet-derived growth factor (PDGF) and heparin-binding epidermal growth factor-like growth factor (HB-EGF). Furthermore, this defect is associated with decreased lamellipodia formation, membrane ruffling, decreased activation of Rac1, and decreased expression of the Rac GEF Sos1. These lines of evidence suggest that CHF1/Hey2 is an important regulator of VSMC responses to arterial injury via regulation of growth factor responsiveness.
Generation of CHF1/Hey2-deficient mice has been described previously.14 Although the majority of these mice die perinatally from complications of ventricular septal defects and a primary cardiomyopathy, a small percentage of mice on a mixed genetic background (129/SvJ and C57BL/6) survive to adulthood with no ventricular septal defects or cardiomyopathy. The mice used in this study had no evidence of cardiomyopathy at the time of injury or the time of harvest. WT littermates were used as controls in all experiments.
Femoral Artery Injury, Immunohistochemistry, and Morphometry
Mouse femoral artery injury and tissue harvesting was performed essentially as described.15 Sections were selected from the vessel at 0.5 mm, 1.0 mm, and 1.5 mm from the site of wire entry and stained with a proliferating cell nuclear antigen (PCNA) staining kit (Zymed), followed by counterstaining with hematoxylin. To assess the intimal and/or medial proliferating cell population, PCNA-positive cells were counted and the PCNA labeling index, the ratio of PCNA positive cells to hematoxylin positive cells, was calculated. For quantitative morphometric analysis, sections were stained with combined Masson–elastic stain. The intimal and medial areas were measured using National Institutes of Health Image software and the average intima, media, and intima-to-media ratio were calculated. All measurements were made by blinded observers.
Culture of Aortic Smooth Muscle Cells, RNA Preparation, and Reverse-Transcription Polymerase Chain Reaction
Three different preparations of WT and KO VSMCs were isolated from the aortas of 8- to 10-week-old matched littermate mice of mixed genetic background (129/SvJ × C57BL/6; 4 to 8 mice for WT and KO VSMCs). The aortas were excised, washed in phosphate-buffered saline, and incubated in DMEM containing 1 mg/mL of Collagenase type II (Worthington Biochemical Corp) for 10 to 15 minutes. Then, under microscopic guidance, the adventitia was removed with fine forceps, the vessels were incised longitudinally, and the endothelial cells were gently scraped off. The vessels were then transferred to culture dishes containing DMEM with 1 mg/mL of Collagenase type I (Worthington Biochemical Corp) and 0.125 mg/mL of Elastase type III (Sigma), minced with scissors, incubated in 37°C, 5% CO2 humidified incubator, and the cells were mechanically dispersed by pipetting vigorously every 10 minutes for 40 to 60 minutes until >90% of the cells were dispersed under the microscope. The cells were centrifuged at 1600 rpm for 5 minutes, then resuspended in 3 mL DMEM with 20% fetal calf serum (FCS), 2% penicillin–streptomycin, and cultured in plates or flasks. Cultured cells were verified to be VSMCs by immunostaining with antismooth muscle α-actin antibody (Sigma) and antismooth muscle calponin antibody (Sigma), followed by counterstaining with Hoechst 33258 (Sigma). RNA isolation and reverse-transcription polymerase chain reaction (RT-PCR) was performed essentially as described.14 Primer sequences for RT-PCR of Rac GEFs and GAPDH are available on request.
Assessment of Cell Growth and Migration
VSMCs were used between passages 3 and 16 in these experiments. Cells were initially plated in triplicate for each time point at a density of 10 000 cells per well of a 12-well tray (surface area 3.8 cm2). Plating efficiency was nearly identical for both types of cells (data not shown). Cells were cultured in DMEM, 20% FCS, and harvested at 3 and 6 days. Cell number was determined with a Z1 particle counter according to the manufacturer’s instructions (Beckman Coulter). Six independent experiments were performed with 2 different paired isolates. Statistical analysis was performed with ANOVA.
The ability of cultured VSMCs to migrate in response to growth factors was assessed by a modified Boyden chamber assay,16 with minor modifications. Briefly, cells were serum-starved 48 to 72 hours, placed in the upper chambers (20 000 cells) of 24-well transwell dishes (8-μm pore size; Corning Science Products, Acton, Mass) in triplicate, and the bottom chambers were filled with DMEM, 0.4% FCS, or DMEM, 0.4% FCS plus 5 ng/mL PDGF-BB (Sigma, St. Louis, Mo) or 10 ng/mL HB-EGF (Calbiochem, San Diego, Calif). The transwell plates were placed in 37°C, 5% CO2 incubator for 4 hours. Attachment was nearly identical for both WT and KO cells (data not shown). The membranes were fixed and stained with PROTOCOL HEMA3 staining kit (Fisher Diagnostics) according to the manufacturer’s instructions. Cells on the lower surface of the membrane were counted manually in 3 consecutive 200× fields on 3 different membranes and averaged. We performed 9 independent experiments from 3 paired isolates. Statistical significance was determined by a Student t test.
Assessment of Lamellipodia and Membrane Ruffling in VSMCs After Growth Factor Stimulation
Chemoattractant-induced lamellipodia formation and membrane ruffling are commonly regarded as essential for cell migration. To assess the effect of CHF1/Hey2 on these cytoskeletal alterations, WT and KO VSMCs were serum-starved for 48 hours, incubated for 30 minutes with 5 ng/mL of PDGF-BB or 10 ng/mL of HB-EGF, and stained with ALEXA fluor 568-labeled phalloidin according to the manufacturer’s instructions (Molecular Probes).
Assessment of Rac1 Activation
Measurement of GTP-bound Rac1 using a coprecipitation method with PAK-1 PBD agarose (Upstate Cell Signaling Solutions, Lake Placid, NY) was performed according to the manufacturer’s instructions with minor modifications. Briefly, after 48 to 72 hours of serum starvation, cells were stimulated with 5 ng/mL of PDGF or 50 ng/mL of HB-EGF for 0, 2, or 5 minutes. Then, cells were lysed with magnesium-containing lysis buffer (25 mmol/L HEPES, pH 7.5, 150 mmol/L NaCl, 1% Triton-X, 10% glycerol, 25 mmol/L NaF, 10 mmol/L MgCl2, 1 mmol/L EDTA, 1 mmol/L Na3VO4, 10 μg/mL leupeptin, and 10 μg/mL aprotinin), and PAK-1 PBD agarose was added to the cell lysate immediately. After incubation for 30 to 60 minutes at 4°C, agarose beads were collected, washed 3 times, resuspended with Laemmli sample buffer, and boiled for 5 minutes. After centrifuging the sample, supernatant and control lysate were analyzed by Western blotting using anti-Rac1 antibody (Upstate Cell Signaling Solutions).
Decreased Neointimal Formation After Wire Injury in KO Mice
In response to wire injury, neointimal formation occurs with proliferation and migration of VSMCs from the medial layer of the artery. Before wire injury, there was no difference in femoral arterial size, and no intimal thickening was present in either population (data not shown). In WT mice, intimal thickening was evident 28 days after injury (0.025±0.011 mm2, N=12), whereas in KO mice, intimal thickening was reduced by 36% (0.016±0.008 mm2, N=13; P<0.05) (Figure 1A to 1E). In KO mice, the intima/media ratio was reduced by 28% at 28 days (2.1±1.1 versus 1.3±0.7, P<0.05; Figure 1F). Our results clearly show that absence of CHF1/Hey2 results in decreased intimal thickening after wire injury.
Decreased Cell Proliferation After Injury in KO Mice
Vascular injury is well-known to induce VSMC proliferation and migration from the medial layer of the arterial wall to the intima. At 5 days after wire injury, we observed a significant difference in PCNA labeling index both in intima (WT 68.4±12.7% versus KO 23.7±13.4%, N=5 for WT, N=4 for KO; P<0.002) (Figure 2A) and in the medial layer (WT 55.7±11.2% versus KO 30.9±9.4%; P<0.009) (Figure 2B). These results indicate that the VSMC proliferative response to wire injury in vivo is blunted in CHF1/Hey2-deficient mice.
Proliferation of Cultured VSMCs Is Decreased in Cells Lacking CHF1/Hey2
VSMCs from WT and KO mice were morphologically identical and were positive for smooth muscle α-actin and smooth muscle calponin (Figure 3A and data not shown). We compared the proliferation of WT and KO VSMCs by standard growth curves. The KO cells grew slowly in comparison with WT cells, demonstrating a statistically significant difference in proliferation in the presence of fetal calf serum at 6 days after plating (Figure 3B).
Chemotactic Response to PDGF and HB-EGF Is Decreased in KO VSMCs
In a modified Boyden chamber assay, CHF1/Hey2-deficient VSMCs had a significantly decreased migratory response to both PDGF (62.6±10.3/CPF versus 37.2±13.5/CPF; P<0.01) and HB-EGF (27.4±7.7/CPF versus 6.4±3.7/CPF; P<0.05) (Figure 4). Because PDGF and HB-EGF are considered to be important for VSMC migration in vivo after vascular injury, it is likely that the decreased responsiveness to PDGF and HB-EGF seen in vitro is in part responsible for decreased neointimal formation in CHF1/Hey2 KO mice.
Cytoskeletal Reorganization and Activation of Rac1 After Growth Factor Stimulation Are Decreased in VSMCs Lacking CHF1/Hey2
Cell motility induced by chemoattractants such as PDGF and HB-EGF requires lamellipodia formation and membrane ruffling, a process that is often dependent on the small GTPase Rac1.17,18 We observed decreased membrane ruffling and formation of lamellipodia in response to PDGF and HB-EGF in KO VSMCs (Figure 5). To understand the underlying mechanism, we examined the effect of growth factor signaling on Rac1 activation. As shown in Figure 6A, the increase in GTP-bound Rac1 seen after growth stimulation is blunted in KO VSMCs, although total Rac1 is not altered. To identify potential mechanisms, we measured the expression of several Rac GEFs and adapter molecules (Tiam1, Tiam2, Sos1, Vav2, Abi1, and Eps8) by RT-PCR and found that Sos1 is specifically decreased in the aortic smooth muscle layer and cultured VSMC, whereas others are not affected (Figure 6B and data not shown). This decreased expression of Sos1 is specific to VSMCs and is not observed in liver cells, which is consistent with the expression pattern of CHF1/Hey2. These findings suggest that the intracellular mechanism by which CHF1/Hey2 regulates responsiveness to growth factors involves Rac1 and the Rac GEF Sos1.
Although it is commonly accepted that phenotypically modulated VSMCs contribute to the development of vascular lesions, little is known regarding the transcriptional mechanisms that control VSMC migration and proliferation. In the present study, we investigated the role of the transcription factor CHF1/Hey2 in the vascular response to wire injury in mice. Our results clearly show that absence of CHF1/Hey2 results in decreased formation of the neointima after wire injury, most likely through decreased VSMC proliferation and migration as a result of decreased responsiveness to growth factors and decreased activation of Rac1. Rac signaling has been shown to be essential for both cell proliferation19 and cell motility.20
The response of the arterial wall to injury has been studied extensively, and many growth factors have been implicated in formation of neointima. Proliferation and migration of smooth muscle cells are hypothesized to be essential for neointimal formation in atherosclerosis and after mechanical arterial injury (ie, angioplasty) through common mechanisms.1 PDGF, released by adherent and aggregating platelets at the site of injury, is considered to play a primary role in vascular smooth muscle proliferation and migration in vivo.21 HB-EGF is localized in the smooth muscle cells and macrophages of human atherosclerotic plaques,22 and expression of HB-EGF in neointimal cells is induced by balloon injury in rat carotid arteries.23 In the present study, CHF1/Hey2 KO mice showed decreased neointimal formation after vascular injury, and cultured CHF1/Hey2 KO VSMCs showed decreased responsiveness to both PDGF and HB-EGF. Our findings link CHF1/Hey2 transcriptional regulation to growth factor signaling, and they are the first to our knowledge to show that CHF1/Hey2 deficiency causes a functional phenotype in VSMCs. These findings suggest a broader role for CHF1/Hey2 in the biological response to vascular injury and link a developmentally regulated transcription factor with an adult phenotype.
Cell growth and motility induced by growth factors such as PDGF and HB-EGF require binding of these growth factors to their receptor tyrosine kinases, followed by reorganization of the cytoskeleton and cell shape changes.24 We initially assessed the activation of proximal signaling pathways and found no significant differences in expression of PDGF or EGF receptors, activation of the MAP kinases ERK1/2, p38, or activation of the survival kinase AKT (Figure I, available online at http://atvb.ahajournals.org and data not shown). Membrane ruffling and formation of lamellipodia after PDGF or HB-EGF treatment, however, were decreased in KO VSMCs when compared with WT VSMCs (Figure 5). In addition, Rac1 activation and Sos1 expression were decreased (Figure 6). Sos1 has been shown to function as a Ras GEF and Rac GEF downstream of receptor tyrosine kinases and is essential for normal embryonic development.25,26 Decreased expression of Sos1 would be expected to decrease the conversion of inactive GDP-bound Rac1 to active GTP-bound Rac1. It is likely that CHF1/Hey2 functions in vascular wall remodeling by regulating the expression of Sos1 or other signaling molecules critical for growth factor responsiveness and cell motility. Because CHF1/Hey2 is a tissue-restricted transcriptional repressor, loss of Sos1 expression in knockout VSMCs is likely to be an indirect effect and is not seen in KO liver cells (Figure 6B).
Other possible mechanisms that have been reported to play a role in vascular remodeling after injury include regulation of cell cycle genes and regulation of apoptosis. The related gene, CHF2/Hey1 (also known as HRT1) has been suggested to promote VSMC growth and survival through suppression of the cyclin kinase inhibitor p21 and by increased expression of the anti-apoptotic kinase AKT, based on analysis of stable transformants of the immortalized smooth muscle cell line A7r5.10 Our work is consistent with a role in facilitating smooth muscle proliferation; however, we do not observe any alterations in the protein expression of the cell cycle-related molecules cyclin A, cyclin D1, cyclin E, Cdk2, Cdk4, p21, and p27 in synchronized populations of WT and KO cells during serum starvation or after stimulation with growth factors (Figures II and III, available online at http://atvb.ahajournals.org and data not shown). Similarly, although we can easily measure apoptosis after vascular injury, we do not see significantly increased apoptosis in the knockout vessels when compared with WT (data not shown). We have also assessed our KO VSMCs for a compensatory increase in the related protein CHF2/Hey1, but we see no change in expression (data not shown). These findings indicate that the predominant effect of CHF1/Hey2 loss of function in our system results in VSMC proliferation and migration defects, which may be mediated by alteration in Rac1 activation, although other mechanisms cannot be ruled out.
One potential limitation of our study is that the mixed genetic background of the mice used may introduce significant variation in lesion size27 and also in the responses of individually derived smooth muscle cell lines. To control for variation in the in vivo model, we have used littermate controls and appropriate sample sizes, quantitative morphometry, and statistical methods to verify that the changes seen are not caused by genetic variation. Similarly, we have taken great care in choosing littermate controls for VSMC preparations and have made each preparation from pools of carefully matched individuals. We have also made multiple preparations of WT and KO cells and have diligently repeated all the experiments with these different preparations. We have also performed careful statistical analysis of our findings to derive our conclusion that CHF1/Hey2 regulates VSMC proliferation and migration in response to growth factors. Nevertheless, genetic background effects cannot be completely ruled out. Ideally, we would perform our experiments on pure C57BL/6 background KO mice; however, these mice do not survive to weaning and thus cannot be used for adult vascular studies (Y. Sakata and M.T. Chin, unpublished data, 2004).
Another potential limitation to our study is the possibility of survival bias introduced by selection. The mice used in this study have survived 2 different selection processes. Approximately two-thirds of the KO mice die from ventricular septal defects in the perinatal period, and the majority of the survivors later die from cardiomyopathy.14 The mice used in this study have survived both selections and therefore may have compensatory mechanisms that confound our analysis. We are currently generating C57BL/6 background conditional KO mice and hope to address these issues of genetic background and selection bias in the future. Despite these caveats, our results demonstrating a significant effect of gene deletion on vascular injury and vascular smooth muscle function raise the exciting possibility that therapy based on CHF1/Hey2-dependent pathways may eventually be useful in the treatment of occlusive vascular disease.
An additional issue is that CHF1/Hey2 (a.k.a. HRT2) expression has been reported to initially decrease in rat carotid arteries after injury and to also decrease in rat aortic smooth muscle cells after PDGF stimulation.9 Although these findings are consistent with the hypothesis that CHF1/Hey2 prevents cell growth, which is at variance with our current findings, the authors of this study were careful not to make this conclusion and clearly recognize the limitations of expression studies without genetic confirmation. We observe a similar decrease in CHF/Hey2 expression in our cultured mouse aortic smooth muscle cells after PDGF treatment (data not shown). Decreased expression after growth factor stimulation in conjunction with genetic loss of function studies presented in this article suggest that the decrease in expression likely reflects the activation of a negative feedback loop, whereby growth signaling pathways are turned off after the initial stimulus.
Our previous work has shown that CHF1/Hey2 can regulate vascular endothelial growth factor expression2 and that CHF1/Hey2 is critical for normal ventricular septation during embryonic development, as well as normal myocardial function.12–14 Others have reported that CHF1/Hey2 is responsive to Notch signaling,7,9,28 and that Notch3 has an important role in the biology of the vessel wall.9,29–31 Our current findings add further genetic evidence through loss of function studies that CHF1/Hey2 is an important transcriptional regulator of signal-responsiveness in target cardiovascular cells during tissue remodeling in vivo and in vitro.
We thank Jianxin Sun, Koh Kawasaki, and Yoshiyuki Rikitake for advice on smooth muscle cell migration assays. We thank James K. Liao and William Boisvert for critical reading of the manuscript. This work was supported by a grant from the National Institutes of Health (HL67141) and an American Heart Association National grant-in-aid to M.T.C., and in part by National Institutes of Health grants HL57506 and HL73852 to D.I.S.
- Received May 5, 2004.
- Accepted August 23, 2004.
Chin MT, Maemura K, Fukumoto S, Jain MK, Layne MD, Watanabe M, Hsieh C-M, Lee M-E. Cardiovascular basic helix loop helix factor 1, a novel transcriptional repressor expressed preferentially in the developing and adult cardiovascular system. J Biol Chem. 2000; 275: 6381–6387.
Nakagawa O, Nakagawa M, Richardson JA, Olson EN, Srivastava D. HRT1, HRT2, and HRT3: a new subclass of bHLH transcription factors marking specific cardiac, somatic, and pharyngeal arch segments. Devel Biol (Orlando). 1999; 216: 72–84.
Zhong TP, Rosenberg M, Mohideen M-APK, Weinstein B, Fishman MC. gridlock, an HLH gene required for assembly of the aorta in zebrafish. Science. 2000; 287: 1820–1824.
Iso T, Sartorelli V, Chung G, Shichinohe T, Kedes L, Hamamori Y. HERP, a new primary target of Notch regulated by ligand binding. Mol Cell Biol. 2001; 21: 6071–6079.
Wang W, Campos AH, Prince CZ, Mou Y, Pollman MJ. Coordinate Notch3-hairy-related transcription factor pathway regulation in response to arterial injury. Mediator role of platelet-derived growth factor and ERK. J Biol Chem. 2002; 277: 23165–23171.
Henderson AM, Wang SJ, Taylor AC, Aitkenhead M, Hughes CC. The basic helix-loop-helix transcription factor HESR1 regulates endothelial cell tube formation. J Biol Chem. 2001; 276: 6169–6176.
Sakata Y, Kamei CN, Nakagami H, Bronson R, Liao JK, Chin MT. Ventricular septal defect and cardiomyopathy in mice lacking the transcription factor CHF1/Hey2. Proc Natl Acad Sci U S A. 2002; 99: 16197–16202.
Roque M, Fallon JT, Badimon JJ, Zhang WX, Taubman MB, Reis ED. Mouse model of femoral artery denudation injury associated with the rapid accumulation of adhesion molecules on the luminal surface and recruitment of neutrophils. Arterioscler Thromb Vasc Biol. 2000; 20: 335–342.
Bornfeldt KE, Raines EW, Nakano T, Graves LM, Krebs EG, Ross R. Insulin-like growth factor and platelet-derived growth factor-BB induce directed migration of human arterial smooth muscle cells via signalling pathways that are distinct from those of proliferation. J Clin Invest. 1994; 93: 1266–1274.
Moore KA, Sethi R, Doanes AM, Johnson TM, Pracyk JB, Kirby M, Irani K, Goldschmidt-Clermont PJ, Finkel T. Rac1 is required for cell proliferation and G2/M progression. Biochem J. 1997; 326 (Pt 1): 17–20.
Anand-Apte B, Zetter BR, Viswanathan A, Qiu RG, Chen J, Ruggieri R, Symons M. Platelet-derived growth factor and fibronectin-stimulated migration are differentially regulated by the Rac and extracellular signal-regulated kinase pathways. J Biol Chem. 1997; 272: 30688–30692.
Heldin CH, Westermark B. Mechanism of action and in vivo role of platelet-derived growth factor. Physiol Rev. 1999; 79: 1283–1316.
Miyagawa J, Higashiyama S, Kawata S, Inui Y, Tamura S, Yamamoto K, Nishida M, Nakamura T, Yamashita S, Matsuzawa Y, et al. Localization of heparin-binding EGF-like growth factor in the smooth muscle cells and macrophages of human atherosclerotic plaques. J Clin Invest. 1995; 95: 404–411.
Igura T, Kawata S, Miyagawa J, Inui Y, Tamura S, Fukuda K, Isozaki K, Yamamori K, Taniguchi N, Higashiyama S, Matsuzawa Y. Expression of heparin-binding epidermal growth factor-like growth factor in neointimal cells induced by balloon injury in rat carotid arteries. Arterioscler Thromb Vasc Biol. 1996; 16: 1524–1531.
Innocenti M, Tenca P, Frittoli E, Faretta M, Tocchetti A, Di Fiore PP, Scita G. Mechanisms through which Sos-1 coordinates the activation of Ras and Rac. J Cell Biol. 2002; 156: 125–136.
Wang DZ, Hammond VE, Abud HE, Bertoncello I, McAvoy JW, Bowtell DD. Mutation in Sos1 dominantly enhances a weak allele of the EGFR, demonstrating a requirement for Sos1 in EGFR signaling and development. Genes Dev. 1997; 11: 309–320.
Dansky HM, Charlton SA, Sikes JL, Heath SC, Simantov R, Levin LF, Shu P, Moore KJ, Breslow JL, Smith JD. Genetic background determines the extent of atherosclerosis in ApoE-deficient mice. Arterioscler Thromb Vasc Biol. 1999; 19: 1960–1968.
Nakagawa O, McFadden DG, Nakagawa M, Yanagisawa H, Hu TH, Srivastava D, Olson EN. Members of the HRT family of basic helix-loop-helix proteins act as transcriptional repressors downstream of Notch signaling. Proc Natl Acad Sci U S A. 2000; 97: 13655–13660.
Joutel A, Corpechot C, Ducros A, Vahedi K, Chabriat H, Mouton P, Alamowitch S, Domenga V, Cecillion M, Marechal E, Maciazek J, Vayssiere C, Cruaud C, Cabanis EA, Ruchoux MM, Weissenbach J, Bach JF, Bousser MG, Tournier-Lasserve E. Notch3 mutations in CADASIL, a hereditary adult-onset condition causing stroke and dementia.[comment]. Nature. 1996; 383: 707–710.
Campos AH, Wang W, Pollman MJ, Gibbons GH. Determinants of Notch-3 receptor expression and signaling in vascular smooth muscle cells: implications in cell-cycle regulation. Circ Res. 2002; 91: 999–1006.
Wang W, Prince CZ, Mou Y, Pollman MJ. Notch3 signaling in vascular smooth muscle cells induces c-FLIP expression via ERK/MAPK activation. Resistance to Fas ligand-induced apoptosis. J Biol Chem. 2002; 277: 21723–21729.