Ritonavir exhibits anti-atherogenic properties on vascular smooth muscle cells
Kappert, Kaia; Caglayan, Evrena; Bäumer, Anselm Ta; Südkamp, Michaelb; Fätkenheuer, Gerdc; Rosenkranz, Stephana
From the aKlinik III für Innere Medizin, the bAbteilung für Herzchirurgie der Universität zu Köln, Joseph-Stelzmann-Str. 9, 50924 Köln, cKlinik I für Innere Medizin, Germany.
Correspondence to Dr. med. Stephan Rosenkranz, Klinik III für Innere Medizin der Universität zu Köln, Joseph-Stelzmann-Str. 9, 50924 Köln, Germany.
Tel: +49 221 478 5159; fax: +49 221 478 6490; e-mail: firstname.lastname@example.org
Received: 24 April 2003; revised: 20 August 2003; accepted: 8 September 2003.
Objectives: HIV protease inhibitors (PI) such as ritonavir have dramatically decreased HIV-related morbidity and mortality. However they exhibit significant side-effects such as hyperlipidemia, hyperglycemia with or without lipodystrophy, which may increase patients′ risk for atherosclerosis. Direct effects of PI on the vascular wall have not been investigated. Platelet-derived growth factor (PDGF) is a major contributor to atherogenesis.
Design: In the present study the effects of ritonavir on PDGF-BB-induced responses of vascular smooth muscle cells (VSMCs) were evaluated.
Methods: PDGF-induced proliferation of VSMCs was measured by BrdU-incorporation, and chemotaxis was assessed by utilizing modified Boyden chambers. Cytotoxicity and apoptosis were quantified using LDH-release- and apoptosis-kits. Immunoprecipitation and Western blot analyses were performed to evaluate βPDGF receptor (βPDGFR) expression and phosphorylation, and to monitor intracellular signaling.
Results: Pretreatment of VSMCs with ritonavir resulted in a significant concentration-dependent inhibition of PDGF-BB-induced cellular responses. At a therapeutic concentration (10 μg/ml), ritonavir significantly reduced PDGF-induced DNA synthesis and chemotaxis by 46.8 ± 5.5% and 37.2 ± 3.3%, respectively (P < 0.05 each). In addition it significantly inhibited PDGF-dependent downstream signaling, such as Erk activation. These inhibitory effects were not due to cytotoxicity or apoptosis. Instead, ritonavir inhibited the ligand-induced tyrosine phosphorylation of the βPDGFR, whereas it did not alter βPDGFR expression.
Conclusions: Ritonavir has direct effects on VSMCs at clinically relevant concentrations in vitro, as it inhibits βPDGFR activation and PDGF-dependent proliferation and migration of VSMCs. Although ritonavir may increase the risk of vascular disease by its metabolic side effects, it may exhibit anti-atherogenic properties on the cellular level.
Protease inhibitor (PI)-based highly active antiretroviral therapy (HAART) has led to an impressive improvement of morbidity and mortality in HIV-infected patients [1–3]. However, it has also been associated with metabolic alterations which may increase the risk for cardiovascular diseases . Among the effects of some PI are elevations of serum lipids, increased insulin resistance, hyperglycemia and lipodystrophy [5,6]. The extent of lipid elevations differs among the various PI. Ritonavir causes the most extensive increases of lipids , whereas other PI such as indinavir are associated with more modest elevations . Even when used as a pharmacologically boosting dose of 100 mg twice daily the lipid elevations observed with ritonavir are quite substantial .
Whether these metabolic changes cause an increase of cardiovascular diseases and thereby eventually counteract the beneficial effects of HAART is subject to an intensive yet unresolved debate. So far, population-based studies have failed to show an increase in cardiovascular diseases in HIV-infected patients , but observation times may be too short to reliably judge this issue. A recent prospective study suggests that the incidence of coronary heart disease may be increased by HAART .
Atherosclerosis is considered a chronic inflammatory disease, and lipid elevations are clearly associated with plaque formation. Several cell types including macrophages, endothelial cells (ECs), platelets and vascular smooth muscle cells (VSMCs) are involved in atherogenesis, and the migration and proliferation of VSMCs within the vascular wall are crucial events in atherosclerotic lesion formation . Platelet-derived growth factor (PDGF) is a major contributor to these processes . PDGF-ligand and -receptors (PDGFR) are significantly upregulated in atherosclerotic plaques , and inhibition of PDGFR signaling by specific antibodies or tyrosine kinase inhibitors potently inhibited atherogenesis in various models [14,15]. Recent studies demonstrated that of the various isoforms and receptor subtypes βPDGFR-mediated signals are particularly important for neointima formation following vascular injury [15,16]. The importance of PDGF in atherogenesis was recently further supported, as upregulation and activation of the βPDGFR in LDLR−/smLRP1−-mice was associated with extensive atherosclerosis, and this response was prevented by the PDGFR-specific tyrosine kinase inhibitor STI571 .
In addition to lipid elevations direct effects of PI on the cellular level have been reported, which may exert adverse as well as beneficial sequels. For example, lipodystrophy may be caused by direct effects of PI on adipocytes as shown in vitro , and subcutaneous adipocyte apoptosis occurs in lipoatrophic areas of patients in vivo. In vascular cells, ritonavir significantly decreased cell viability of human ECs at concentrations near therapeutic plasma levels . Consistent with this study, the use of HIV PI in humans is associated with endothelial dysfunction in vivo , representing an early stage of atherosclerosis. In addition to the harmful influence on endothelial integrity and function, beneficial effects of PI on the vascular wall have also been reported. In HIV-infected patients, elevated levels of endothelial markers including soluble adhesion molecules (sVCAM-1, sICAM-1) and von Willebrand factor were significantly reduced by antiretroviral therapy including PI .
The above studies provide evidence for direct actions of PI on various cell types including vascular ECs on the cellular level. Here we investigated, whether ritonavir exerts direct effects on VSMCs and affects PDGF-BB-induced cell migration and proliferation.
Cells and reagents
VSMCs were isolated from rat thoracic aorta (Wistar–Kyoto; 6–10 weeks old; Charles River Wega GmbH, Sulzfeld, Germany) by enzymatic dispersion as previously described . Cells were grown in a 5% CO2 atmosphere at 37°C in Dulbecco's modified Eagle's medium (DMEM) supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, 1% nonessential amino acids (100×) and 10% fetal calf serum (FCS). PDGF-BB was purchased from Promo Cell (Heidelberg, Germany). Antiphosphotyrosine antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, California, USA) (PY20) and Upstate Biotechnology (Lake Placid, New York, USA) (4G10). Antibodies against the βPDGFR (97A) and RasGAP (69.3) were a kind gift from Andrius Kazlauskas (Harvard Medical School, Boston, Massachusetts, USA). The phospho-Erk 1/2 (Tre 202, Tyr 204) antibody and the phospho-Akt (Ser 473) antibody were purchased from Cell Signaling (Beverly, MA, USA).
PDGF-dependent cell cycle progression was measured by a 5-bromodeoxyuridine (BrdU)-incorporation assay . Cells were synchronized overnight, followed by pre-incubation with ritonavir for either 1 or 24 h, as indicated. PDGF-BB was added to cells for 12–24 h at the indicated concentrations in the absence or presence of ritonavir. BrdU-incorporation was carried out according to the manufacturers specifications (Roche, Mannheim, Germany) with an incorporation time of 12 h.
PDGF-dependent chemotaxis was assayed using a 48-well modified Boyden chamber (NeuroProbe, Baltimore, Maryland, USA) and polyvinyl pyrrolidone-free polycarbonate filters (8 μm pores) (Poretics, Livermore, California, USA) as previously described . In brief, the lower wells of the chamber were filled with DMEM supplemented with 10 ng/ml PDGF-BB or vehicle in the presence or absence of various concentrations of ritonavir as indicated. The filters were coated with 50 mg/ml rat type I collagen (Collaborative Biomedical Products, Bedford, Massachusetts, USA) and fixed atop the bottom wells. VSMCs were trypsinized, washed, and 20 × 103 cells were placed into the top wells of the chamber. The chamber was incubated for 5 h at 37°C in a 5% CO2 atmosphere and was then disassembled. The cells on the upper surface of the filter were gently removed, and the cells on the lower surface were fixed and stained with Diff-Quick (Baxter Healthcare, Miami, Florida, USA). Chemotaxis was quantified by counting the number of cells on the lower surface of the filter in each well.
Immunoprecipitation and Western blot analysis
Quiescent VSMCs were left resting or stimulated with 1 ng/ml PDGF-BB for indicated time intervals in the presence or absence of ritonavir. The βPDGFR was immunoprecipitated as previously described , except that immune complexes were bound to protein-A sepharose. The βPDGFR immunoprecipitates representing approximately 3 × 106 cells were resolved on a 7.5% sodium dodecyl sulfate (SDS)-polyacrylamide electrophoresis gel; the proteins were transferred to Immun-Blot® PVDF-membranes (BioRad, München, Germany) and subjected to Western blot analysis, using antisera which recognize phospho-tyrosine (PY20, 4G10) or the βPDGFR (97A). To monitor the phosphorylation state of cell signaling molecules total cell lysates from resting or PDGF-stimulated cells were separated in a 12.5% SDS gel and subjected to Western blot analysis, using antisera against RasGAP, phospho-Erk 1/2, or phospho-Akt.
Cell viability and cytotoxicity assay
Cell viability was assessed by a tetrazolium compound, 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS), assay kit (Promega, Mannheim, Germany) . The assay was carried out according to the manufactures instructions. Briefly cells were synchronized by serum-deprivation and then stimulated for the indicated time periods with or without different concentrations of ritonavir. Cytotoxicity was measured by the lactate dehydrogenase (LDH) release from cells. Calculations were performed according to the following definition: Percentage cytotoxicity = 100 × [(Experimental – Culture Medium Background)/(Maximum LDH Release – Culture Medium Background)]
The apoptosis of VSMCs was analysed using a Cell Death Detection ELISAPLUS-Kit (Roche). The assay was carried out according to the manufacturers’ instructions. In brief, synchronized cells were incubated with different concentrations of ritonavir in 0%-FCS-medium for the indicated time periods. Cell-lysates were placed into a streptavidin-coated microtitre plate, followed by addition of anti-histone-biotin and anti-DNA-POD. The quantification of the amount of nucleosomes by the POD retained in the immunocomplex was determined photometrically with ABTS (2,2′-azino-bis-3-ethyl-benzthiazoline-6-sulphuric acid) as a substrate.
All data are expressed as means ± standard error of the means (SEM). Statistical analysis was evaluated by non-parametric analysis and P < 0.05 was considered significant.
Ritonavir inhibits PDGF-dependent VSMC proliferation and migration
To investigate the effects of ritonavir on PDGF-dependent cellular responses in VSMCs, we measured cell proliferation and migration after pre-treatment with ritonavir prior to PDGF stimulation. DNA synthesis was measured by BrdU incorporation assays. Stimulation of VSMCs with PDGF-BB led to a concentration-dependent increase in BrdU uptake to maximally 211.8 ± 9.6% at 20 ng/ml (P < 0.01), whereas higher concentrations did not cause a more significant induction of DNA synthesis (not shown). Therefore, the following experiments were carried out with PDGF-BB 20 ng/ml. Pre-treatment of VSMCs with ritonavir for 24 h led to a concentration-dependent decrease of PDGF-induced cell cycle progression (Fig. 1a). Whereas ritonavir 0.001–1.0 μg/ml did not significantly alter PDGF-dependent VSMC proliferation, the therapeutically relevant concentrations of 5, 10 and 20 μg/ml led to an inhibition of BrdU uptake by 43.3 ± 7.2, 46.8 ± 5.5 and 88.4 ± 15.7% (P < 0.05 each), respectively, compared to PDGF-treated control cells. This effect was also observed when ritonavir was added 1 h prior to PDGF stimulation (not shown).
PDGF-dependent migration of VSMCs was measured using a modified Boyden chemotaxis-chamber. Stimulation of VSMCs with PDGF-BB (10 ng/ml) resulted in an approximately six-fold induction of cell migration compared with non-stimulated cells. When PDGF was administered in the presence of ritonavir, PDGF-dependent chemotaxis was inhibited in a concentration-dependent manner (Fig. 1b). Whereas 1 μg/ml ritonavir did not significantly alter the migration rate after PDGF treatment (−9.9 ± 3.0%; NS), ritonavir 5 and 10 μg/ml significantly inhibited PDGF-dependent chemotaxis by 19.9 ± 3.1% (P < 0.05) and 37.2 ± 3.3% (P < 0.01), respectively, when compared to PDGF-treated cells.
Ritonavir inhibits βPDGFR-signaling
To investigate how ritonavir inhibits PDGF-induced cellular responses in VSMCs, we measured its effects on βPDGFR expression and the ligand-induced tyrosine phosphorylation of the βPDGFR. The expression levels of the βPDGFR were analysed by Western blot analyses in total cell lysates from −/+-ritonavir-pretreated VSMCs. The cells were incubated with ritonavir (10 μg/ml) for different time periods (0–24 h). No alteration of receptor levels was observed throughout the observation period (Fig. 2a).
In order to investigate the effects of ritonavir on PDGFR activation, the ligand-induced tyrosine phosphorylation of the βPDGFR was measured in the presence of various concentrations of ritonavir. To this end, VSMCs were pretreated with 1, 5 and 10 μg/ml ritonavir for 24 h and subsequently stimulated with PDGF-BB for 5 min at 37°C. The cells were then lysed, the βPDGFR was immunoprecipitated and the immunoprecipitates were subjected to Western blot analysis, using antisera against phospho-tyrosine or the βPDGFR. Treatment of VSMCs with PDGF-BB significantly increased the phospho-tyrosine content of the βPDGFR. As shown in Figure 2b, ritonavir led to a concentration-dependent reduction of the ligand-induced tyrosine phosphorylation of the βPDGFR. When normalized for the levels of βPDGFR present in the immunoprecipitates, densitometric analysis of three independent anti-phosphotyrosine blots revealed a statistically significant inhibitory effect at concentrations of 5 and 10 μg/ml (P < 0.05 each) (Fig. 2c).
We also evaluated the effects of ritonavir on PDGF-dependent downstream signaling events in VSMCs. Synchronized cells were treated with 10 μg/ml ritonavir for 24 h and stimulated with PDGF-BB (1 ng/ml) for 0, 5, 30 and 45 min prior to protein extraction. Proteins were resolved by gel electrophoresis, and subjected to Western blotting using phospho-specific antibodies recognizing MAPK (Erk 1/2) and Akt. Figure 3 demonstrates that pre-treatment with ritonavir caused a decrease of PDGF-induced MAPK activation whereas no effect on Akt phosphorylation could be found.
Ritonavir does not alter VSMC viability or apoptosis at relevant concentrations
To rule out that the inhibitory effects of ritonavir on βPDGFR signaling and PDGF-dependent cellular effects in VSMCs were due to cytotoxicity, we performed cell viability and apoptosis assays. The effect of ritonavir on VSMC viability was determined by an MTS assay, as described earlier . First, VSMCs were treated with 20 μg/ml ritonavir for different time-periods. At 24, 48 and 72 h, the cell viability rates were 87.8 ± 3.7, 60.0 ± 7.2 and 40.0 ± 28.0%, respectively, showing a clear, time-dependent decline at all analysed time points compared with untreated control cells (all P < 0.05, Fig. 4a). At time points less than 24 h (4, 8 and 12 h), no significant decrease in cell viability could be observed (not shown). After we established these kinetics of cellular injury due to ritonavir treatment, we performed dose-dependency experiments, in order to determine the concentrations of ritonavir that may cause declining VSMC viability. Since cell viability was first decreased at ritonavir (20 μg/ml) after 24 h, the following experiments were performed at this time point. Cells were treated with different concentrations of ritonavir (1, 10, 20 and 100 μg/ml) for 24 h (Fig. 4b). A significant decrease of VSMC viability could be demonstrated in cells treated with 20 and 100 μg/ml ritonavir (−13.2 ± 3.7 and −21.0 ± 5.8%, respectively, P < 0.05 each), but not in cells treated with therapeutically relevant concentrations of 1 and 10 μg/ml.
To test whether ritonavir causes VSMC apoptosis, cell death detection enzyme-linked immunosorbent assays were performed. VSMCs were incubated in the presence or absence of ritonavir (1, 10, 20 and 100 μg/ml) for 24 h (Fig. 4c). Treatment with ritonavir at 20 and 100 μg/ml caused an increase of apoptotic cells to 201.8 ± 3.3 and 216.5 ± 5.6%, respectively (P < 0.05 each) compared with control cells. In contrast, lower concentrations of ritonavir (1 and 10 μg/ml) did not induce VSMC apoptosis, as shown in Figure 4c.
These data indicate that ritonavir at 10 μg/ml and lower has no effect on VSMC viability and apoptosis. However, treatment with higher concentrations (20 and 100 μg/ml) causes significant cytotoxic ad proapoptotic cellular responses.
The present study demonstrates for the first time that ritonavir has significant impact on VSMC functions at clinically relevant concentrations. In the presence of 10 μg/ml ritonavir PDGF-dependent VSMC migration and cell cycle progression were significantly attenuated. The inhibitory effects correlated with impaired βPDGFR-activation, without altering receptor-levels. These potentially beneficial effects on VSMCs were observed at concentrations, which did not cause cell death.
This is the first study demonstrating a direct anti-proliferative effect of an HIV PI on VSMCs. More specifically, we show that ritonavir inhibits the proliferation and migration of VSMCs in response to PDGF. Numerous studies have demonstrated that signal relay by PDGFRs plays a critical role in all phases of atherogenesis (reviewed in ). PDGF exerts its biological effects via activation of two subtypes of receptor tyrosine kinases, termed α and βPDGFR [27,28]. Since βPDGFR-mediated signals are particularly important for neointima formation following vascular injury , we focused on this receptor subtype. Once activated, the βPDGFR specifically promotes signal relay cascades, leading to downstream events such as MAP kinase activation, which subsequently mediate PDGF-dependent cellular responses .
Here we show, that ritonavir inhibits the ligand-induced tyrosine phosphorylation (activation) of the βPDGFR at clinically relevant concentrations. As a consequence, ritonavir attenuates PDGF-dependent downstream events such as Erk phosphorylation and cellular responses including VSMC proliferation and migration. As these responses are critical events during the formation of atherosclerotic plaques, ritonavir exerts direct effects on VSMCs in vitro which may protect from atherogenesis on the cellular level. These effects are of potential benefit against atherogenesis, as selective inhibition of the βPDGFR was shown to significantly reduce the formation of atherosclerotic plaques in various models [17,29–32]. Inhibition of PDGFR signaling by monoclonal antibodies or specific tyrosine kinase inhibitors potently attenuates the atherogenic and/or restenotic process [17,30,31]. Functional blockade of the βPDGFR by injection of a monoclonal antibody into apolipoprotein E-deficient mice led to a significant reduction of aortic atherosclerotic lesion size and intimal VSMCs . Injection of a mouse/human chimeric anti-βPDGFR antibody also led to an approximately 50% decrease in intimal area and intima-to-media ratio in non-human primates . These studies collectively indicate that PDGF plays a key role in the development of intimal lesions at sites of vascular injury and that PDGFR inhibition is sufficient to suppress atherogenesis in vivo.
The clinical use of ritonavir is associated with significant metabolic disorders, which represent well-characterized risk factors for atherosclerotic vascular disease [33–35]. However, the clinical consequences of these side effects are not quite clear. In some studies, they were associated with an increased risk for cardiovascular events [11,20], whereas recent studies did not show an increased cardiovascular risk in HAART-treated patients [10,36]. Mercie and co-workers recently analysed the carotid intima–media thickness, a surrogate marker of atherosclerosis, in HIV-infected patients in a multivariate analysis. Although lipodystrophy and HAART were primarily associated with an increased intima–media thickness, the effect disappeared after adjustment for other cardiovascular risk factors . Furthermore, a retrospective analysis of > 35 000 patients who received HAART demonstrated that there is no increased risk for cardiovascular or cerebrovascular events and related mortality in these patients . This may be explained by relatively short observation times, or by the protective effects of ritonavir on the cellular level that may counteract the atherogenic profile in patients treated with HAART. The present study reveals that ritonavir – at clinically relevant concentrations – attenuates PDGF-induced proliferation and migration of cultured VSMCs, which are critical events in atherogenesis. Clearly, these observations were only obtained in vitro, and thus cannot be extrapolated to the in vivo situation in humans. Nevertheless, our findings offer a molecular explanation for the fact, that the increased individual cardiovascular risk profile in HAART-treated patients does not necessarily lead to an increase of ischemic events.
The serum concentrations of ritonavir in HIV-infected patients depend on inter-individual differences and drug interactions, particularly with other antiretroviral drugs . The maximal plasma concentration of fully dosed ritonavir (600 mg twice daily) is 11.2 μg/ml . We studied cellular responses of ritonavir at concentrations between 1 and 100 μg/ml. Consistent with others , we found that the type of cellular response depends on the concentration used, and that the ‘therapeutic range’ of ritonavir is rather limited. Inhibition of PDGF-dependent cell cycle progression and chemotaxis of VSMCs in vitro was obtained at clinically relevant concentrations. To ensure that these inhibitory effects are not linked to cytotoxic or pro-apoptotic events, we performed cell viability and apoptosis assays. These revealed that relevant concentrations (1, 5 and 10 μg/ml) did not attenuate cell viability, whereas higher concentrations (≥ 20 μg/ml) led to cytotoxic and apoptotic responses of VSMCs. This is consistent with the finding, that ritonavir reduced cell viability (> 11 μg/ml) and induced apoptosis (> 22 μg/ml) of cultured human ECs . Therefore, the effects of ritonavir on βPDGFR activation and PDGF-induced responses in VSMCs were investigated only at low concentrations up to 10 μg/ml. It is unlikely that the inhibitory effects of ritonavir on VSMC function at these concentrations are due to cytotoxicity. Furthermore, pre-incubation of ritonavir for 1 h prior to PDGF-BB was sufficient of altering DNA-synthesis, indicating that the inhibitory effect was due to direct attenuation of receptor activation rather than alterations of gene expression or cytotoxic effects.
In summary, our findings offer a novel, molecular explanation for the fact that HAART-treated patients may not be at increased risk for vascular complications despite severe pro-atherogenic metabolic changes. We do not know whether ritonavir also affects signaling by other growth factors, as we have only investigated PDGF-mediated events. Future studies should focus on the comparison of different PI and evaluate substance-specific or class-specific effects on signal relay by various growth factors. Also, the currently used antiretroviral combinations in HAART may be analysed in order to monitor for possible additive effects. Clinical and experimental studies will show, whether substance-specific differences among PI in terms of altering vascular cell functions in vitro as well as in vivo exist.
We thank Manuela Uebel for excellent technical assistance.
Sponsorship: Ritonavir was provided by Abbott (Wiesbaden, Germany). This work was supported by the Köln Fortune Program, Faculty of Medicine, University of Cologne (84/2002 to K.K.) and by the Deutsche Forschungsgemeinschaft (Ro 1306/2-1 to S.R.).
1. Palella FJ, Jr., Delaney KM, Moorman AC, Loveless MO, Fuhrer J, Satten GA, et al. Declining morbidity and mortality among patients with advanced human immunodeficiency virus infection. HIV Outpatient Study Investigators. N Engl J Med 1998, 338:853–860.
2. Hammer SM, Squires KE, Hughes MD, Grimes JM, Demeter LM, Currier JS, et al. A controlled trial of two nucleoside analogues plus indinavir in persons with human immunodeficiency virus infection and CD4 cell counts of 200 per cubic millimeter or less. AIDS Clinical Trials Group 320 Study Team. N Engl J Med 1997, 337:725–733.
3. Cameron DW, Heath-Chiozzi M, Danner S, Cohen C, Kravcik S, Maurath C, et al. Randomised placebo-controlled trial of ritonavir in advanced HIV-1 disease. The Advanced HIV Disease Ritonavir Study Group. Lancet 1998, 351:543–549.
4. Behrens G, Schmidt H, Meyer D, Stoll M, Schmidt RE. Vascular complications associated with use of HIV protease inhibitors. Lancet 1998, 351:1958.
5. Carr A, Samaras K, Burton S, Law M, Freund J, Chisholm DJ, et al. A syndrome of peripheral lipodystrophy, hyperlipidaemia and insulin resistance in patients receiving HIV protease inhibitors. AIDS 1998, 12:F51–58.
6. Walli R, Herfort O, Michl GM, et al. Treatment with protease inhibitors associated with peripheral insulin resistance and impaired oral glucose tolerance in HIV-1-infected patients. AIDS 1998, 12:F167–173.
7. Sullivan AK, Nelson MR. Marked hyperlipidaemia on ritonavir therapy. AIDS 1997, 11:938–939.
8. Manfredi R, Chiodo F. Disorders of lipid metabolism in patients with HIV disease treated with antiretroviral agents: frequency, relationship with administered drugs, and role of hypolipidaemic therapy with bezafibrate. J Infect 2001, 42:181–188.
9. Arnaiz JA, Mallolas J, Podzamczer D, Gerstoft J, Lundgren JD, Cahn P, et al. Continued indinavir versus switching to indinavir/ritonavir in HIV-infected patients with suppressed viral load. AIDS 2003, 17:831–840.
10. Bozzette SA, Ake CF, Tam HK, Chang SW, Louis TA. Cardiovascular and cerebrovascular events in patients treated for human immunodeficiency virus infection. N Engl J Med 2003, 348:702–710.
11. Friis-Moller N, Weber R, Reiss P, Thiebaut R, Kirk O, d'Arminio Monforte A, et al. Cardiovascular disease risk factors in HIV patients–association with antiretroviral therapy. Results from the DAD study. AIDS 2003, 17:1179–1193.
12. Ross R. The pathogenesis of atherosclerosis: a perspective for the 1990s. Nature 1993, 362:801–809.
13. Raines EW, Ross R. Multiple growth factors are associated with lesions of atherosclerosis: specificity or redundancy? Bioessays 1996, 18:271–282.
14. Fishbein I, Waltenberger J, Banai S, Rabinovich L, Chorny M, Levitzki A, et al. Local delivery of platelet-derived growth factor receptor-specific tyrphostin inhibits neointimal formation in rats. Arterioscler Thromb Vasc Biol 2000, 20:667–676.
15. Leppanen O, Janjic N, Carlsson MA, Pietras K, Levin M, Vargeese C, et al. Intimal hyperplasia recurs after removal of PDGF-AB and -BB inhibition in the rat carotid artery injury model. Arterioscler Thromb Vasc Biol 2000, 20:E89–95.
16. Klinghoffer RA, Mueting-Nelsen PF, Faerman A, Shani M, Soriano P. The two PDGF receptors maintain conserved signaling in vivo despite divergent embryological functions. Mol Cell 2001, 7:343–354.
17. Boucher P, Gotthardt M, Li WP, Anderson RG, Herz J. LRP: role in vascular wall integrity and protection from atherosclerosis. Science 2003, 300:329–332.
18. Lenhard JM, Furfine ES, Jain RG, Ittoop O, Orband-Miller LA, Blanchard SG, et al. HIV protease inhibitors block adipogenesis and increase lipolysis in vitro. Antiviral Res 2000, 47:121–129.
19. Zhong DS, Lu XH, Conklin BS, Lin PH, Lumsolen AB, Yao Q, et al. HIV protease inhibitor ritonavir induces cytotoxicity of human endothelial cells. Arterioscler Thromb Vasc Biol 2002, 22:1560–1566.
20. Stein JH, Klein MA, Bellehumeur JL, McBride PE, Wiebe DA, Otvos JD, et al. Use of human immunodeficiency virus-1 protease inhibitors is associated with atherogenic lipoprotein changes and endothelial dysfunction. Circulation 2001, 104:257–262.
21. Wolf K, Tsakiris DA, Weber R, Erb P, Battegay M. Antiretroviral therapy reduces markers of endothelial and coagulation activation in patients infected with human immunodeficiency virus type 1. J Infect Dis 2002, 185:456–462.
22. Chamley-Campbell J, Campbell GR, Ross R. The smooth muscle cell in culture. Physiol Rev 1979, 59:1–61.
23. Rosenkranz S, Knirel D, Dietrich H, Flesch M, Erdmann E, Bohm M. Inhibition of the PDGF receptor by red wine flavonoids provides a molecular explanation for the ‘‘French paradox’'. FASEB J 2002, 16:1958–1960.
24. Rosenkranz S, DeMali KA, Gelderloos JA, Bazenet C, Kazlauskas A. Identification of the receptor-associated signaling enzymes that are required for platelet-derived growth factor-AA-dependent chemotaxis and DNA synthesis. J Biol Chem 1999, 274:28335–28343.
25. Baxter RM, Secrist JP, Vaillancourt RR, Kazlauskas A. Full activation of the platelet-derived growth factor beta-receptor kinase involves multiple events. J Biol Chem 1998, 273:17050–17055.
26. Ross R. Atherosclerosis is an inflammatory disease. Am Heart J 1999, 138:S419–420.
27. Heldin CH, Westermark B. Mechanism of action and in vivo role of platelet-derived growth factor. Physiol Rev 1999, 79: 1283–1316.
28. Rosenkranz S, Kazlauskas A. Evidence for distinct signaling properties and biological responses induced by the PDGF receptor alpha and beta subtypes. Growth Factors 1999, 16:201–216.
29. Hart CE, Kraiss LW, Vergel S, Gilbertson D, Kenagy R, Kirkman T, et al. PDGFbeta receptor blockade inhibits intimal hyperplasia in the baboon. Circulation 1999, 99:564–569.
30. Davies MG, Owens EL, Mason DP, Lea H, Tran PK, Vergel S, et al. Effect of platelet-derived growth factor receptor-alpha and -beta blockade on flow-induced neointimal formation in endothelialized baboon vascular grafts. Circ Res 2000, 86: 779–786.
31. Bilder G, Wentz T, Leadley R, Amin D, Byan L, O'Conner B, et al. Restenosis following angioplasty in the swine coronary artery is inhibited by an orally active PDGF-receptor tyrosine kinase inhibitor, RPR101511A. Circulation 1999, 99:3292–3299.
32. Sano H, Sudo T, Yokode M, Murayama T, Kataoka H, Takakura N, et al. Functional blockade of platelet-derived growth factor receptor-beta but not of receptor-alpha prevents vascular smooth muscle cell accumulation in fibrous cap lesions in apolipoprotein E-deficient mice. Circulation 2001, 103: 2955–2960.
33. Carr A, Cooper DA. Images in clinical medicine. Lipodystrophy associated with an HIV-protease inhibitor. N Engl J Med 1998, 339:1296.
34. Periard D, Telenti A, Sudre P, Cheseaux JJ, Halfan P, Reymond MJ, et al. Atherogenic dyslipidemia in HIV-infected individuals treated with protease inhibitors. The Swiss HIV Cohort Study. Circulation 1999, 100:700–705.
35. Mulligan K, Grunfeld C, Tai VW, Algren H, Pang M, Chernoff DN, et al. Hyperlipidemia and insulin resistance are induced by protease inhibitors independent of changes in body composition in patients with HIV infection. J Acquir Immune Defic Syndr 2000, 23:35–43.
36. Mercie P, Thiebaut R, Lavignolle V, Pellegrin JL, Yvorra-Vives MC, Morlat P, et al. Evaluation of cardiovascular risk factors in HIV-1 infected patients using carotid intima-media thickness measurement. Ann Med 2002, 34:55–63.
37. Hsu A, Granneman GR, Bertz RJ. Ritonavir. Clinical pharmacokinetics and interactions with other anti-HIV agents. Clin Pharmacokinet 1998, 35:275–291.
38. Lea AP, Faulds D. Ritonavir. Drugs 1996, 52:541–546; discussion 547–548.
This article has been cited 6 time(s).
American Journal of Physiology-Heart and Circulatory Physiology17 beta-estradiol attenuates PDGF signaling in vascular smooth muscle cells at the postreceptor levelAmerican Journal of Physiology-Heart and Circulatory Physiology
Biochemical and Biophysical Research CommunicationsRitonavir does not inhibit calpain in vitroBiochemical and Biophysical Research Communications
Experimental Biology and MedicineRitonavir Increases CD36, ABCA1 and CYP27 Expression in THP-1 MacrophagesExperimental Biology and Medicine
Medecine Et Maladies Infectieuses
Tolerance of boosted antiproteases
Medecine Et Maladies Infectieuses, 34():
JAIDS Journal of Acquired Immune Deficiency SyndromesMolecular Mechanisms of HIV Protease Inhibitor-Induced Endothelial DysfunctionJAIDS Journal of Acquired Immune Deficiency Syndromes
HIV protease inhibitor; ritonavir; atherosclerosis; vascular smooth muscle cells; migration; proliferation; platelet-derived growth factor
© 2004 Lippincott Williams & Wilkins, Inc.
Highlight selected keywords in the article text.