Activation of the innate and adaptive immune system contributes to the progression of cardiovascular disease (CVD) in the general population,1,2 and inflammation and immune activation are associated with mortality, including deaths related to CVD, in patients infected with the HIV.3,4 Immune activation may mediate HIV disease progression, vascular disease, diabetes,3,5–8 and an increased risk of both venous and arterial thrombosis9–20 in HIV-infected subjects. Lymphocyte activation, as measured by CD38 and HLA-DR expression on CD4+ and CD8+ T cells, is predictive of disease course in untreated HIV infection21 and of CD4+ T-cell reconstitution after initiation of antiretroviral therapy (ART).22 We have reported a direct relationship between the proportion of activated CD8+ T cells and mean common carotid artery intima-media thickness (IMT) in HIV disease, and an increased proportion of activated CD8+ T cells in HIV-infected patients with coronary plaque (IMT > 1.5 cm) compared with these proportions in patients without plaque.23 In HIV-infected women, T-cell activation was also associated with subclinical atherosclerosis,24,25 providing further evidence for a relationship between T-cell activation in CVD risk in chronic HIV disease.
Several strategies to reduce chronic immune activation in treated HIV disease are underway, including this trial: Stopping Atherosclerosis and Treating Unhealthy bone with RosuvastatiN in HIV (SATURN-HIV). Statins, or 3-hydroxy-3-methylglutaryl coenzyme A (HMG-CoA) reductase inhibitors, have anti-inflammatory effects,26,27 and here, HIV-infected subjects receiving successful ART, and who had normal low-density lipoprotein-cholesterol (LDL-C) levels, but elevated levels of immune activation, were randomized to receive rosuvastatin (10 mg daily) or placebo. We reported that 24 weeks of rosuvastatin treatment resulted in significant reductions in markers of monocyte subset activation28 and in vascular inflammation (lipoprotein-associated phospholipase A2, Lp-PLA2).29 Rosuvastatin had no effect on systemic inflammation or T-cell activation28; results in discord with the findings of a small study where 8-week administration of high-dose atorvastatin (80 mg) reduced modestly the proportion of HLA-DR expressing CD8+ T cells30 in patients not receiving ART. We hypothesized that rosuvastatin therapy may take longer to reduce T-cell activation in treated subjects than in ART-naive subjects, because of the lower levels of T-cell activation generally reported in treated vs untreated HIV disease. Here, we present the results of a prespecified secondary analysis aimed at assessing the effects of statin administration on markers of immune activation and inflammation at 48 weeks.
SATURN-HIV is a randomized, double-blind placebo-controlled study designed to measure the effect of rosuvastatin on markers of cardiovascular risk, skeletal health, and immune activation in HIV disease and is registered on clinicaltrials.gov (identifier: NCT01218802). The study was approved by the Institutional Review Board of University Hospitals Case Medical Center (Cleveland, OH), and all subjects signed a written consent before enrollment. Randomization was conducted by the Case investigational pharmacist at 1:1 to active rosuvastatin 10 mg daily vs matching placebo. Randomization was stratified by protease inhibitor use. Study drugs (active and placebo) were provided by AstraZeneca.
All subjects were aged 18 years or older, without known coronary disease or diabetes, and on stable ART for at least 3 months and cumulative ART duration of at least 6 months, with HIV-1 RNA <1000 copies per milliliter and fasting LDL-C ≤130 mg/dL and fasting triglycerides ≤500 mg/dL. Subjects were excluded if they had a history of myocardial infarction, were pregnant or lactating, or had another active or controlled inflammatory condition. Additional entry criteria included evidence of either heightened T-cell activation, identified as the proportion of CD8+ T cells that expressed CD38+HLA-DR+ ≥19%, or levels of high sensitivity C-reactive protein (CRP) ≥2 mg/L. The T-cell activation cutoff (>19%) was determined based on our previous data linking this level of immune activation to immunologic failure in treated HIV infection.31 The CRP cutoff was selected based on criteria established in the JUPITER statin trial.27
At screening visit, self-reported demographics and medical history were obtained, along with a targeted physical examination including height and weight measurements. Blood was drawn after a 12-hour fast for lipoproteins and high sensitivity CRP. Percent CD8+ T-cell activation was determined as described below. If enrollment criteria were met, subjects returned within 30 days for entry evaluations. At entry, a fasting blood draw was obtained for a lipid profile and for measurement of soluble markers of immune activation. HIV-1 RNA levels and CD4+ cell counts were obtained as part of routine care.
Whole-blood samples were collected into EDTA-containing tubes. Peripheral blood mononuclear cell (PBMC) samples were separated by centrifugation with Ficoll-Hypaque and were cryopreserved until analyzed in batch. Plasma was isolated by centrifugation for 10 minutes at 400g and was frozen at −80°C until thawed once and analyzed in batch.
Fresh whole-blood (300-μL) samples were incubated for 15 minutes on ice with FACS Lyse buffer (BD Biosciences, San Diego, CA) and were then washed in buffer (phosphate-buffered saline with 1% bovine serum albumin and 0.1% sodium azide). Cells were then stained for 30 minutes in the dark, on ice, and then washed in buffer and fixed in 1% paraformaldehyde. Monocyte subsets were identified by size, granularity, and by expression of CD14 and CD16.32 Isotype gating was used to identify positive expression of surface markers. Cell surface molecule expression was monitored by staining cells with fluorochrome-labeled antibodies: anti-Tissue Factor (TF) (fluorescein isothiocyanate; American Diagnostica, Stamford, CT), anti-CD14 (Pacific Blue), anti-CD16 (phycoerythrin, PE; both from BD Pharmingen, San Diego, CA).
The proportion of activated T cells required to determine eligibility was measured by analyses of freshly collected whole-blood samples, processed as above. T cells were identified by size and granularity and by positive expression of CD3 and CD4 or CD8. T-cell activation was measured using anti-CD38 (PE), anti-HLA-DR (fluorescein isothiocyanate), anti-CD3 (Peridinin Chlorophyll Protein Complex) anti-CD8 (allophycocyanin-cy7), anti-CD4 (allophycocyanin; all from BD Biosciences).
Assessment of T-cell activation for the baseline/entry, 24-, and 48-week time points was performed by comparing expression of surface markers on cryopreserved PBMC samples from each patient. Samples were thawed and analyzed in batch, to maintain uniformity of cytometer settings among time points for each patient. In addition to the T-cell markers described above, analysis of frozen PBMC samples also included a stain for cell viability (Live/Dead Violet, Pacific Blue) and an additional activation marker PD-1 (PE-Cy7; BD Pharmingen).
Monocytes were analyzed in real time, on fresh whole-blood samples, as above using a Miltenyi MACS Quant flow cytometer (Miltenyi Biotec, Bergisch Gladbach, Germany). MACS Quantify software (version 2.21031.1; Miltenyi Biotec) was used to analyze the data. Initial assessment of T-cell activation for screening/enrollment was performed using an LSR II flow cytometer (Becton-Dickinson, San Jose, CA) and FACSDiva software version 6.1.1 (BD Biosciences). Longitudinal T-lymphocyte activation was analyzed on the MACS Quant.
Plasma samples were collected at baseline, 24 and 48 weeks. Levels of soluble CD14 and CD163 were measured using Quantikine enzyme-linked immunosorbent assay (ELISA) kits (R&D Systems, Minneapolis, MN). Levels of the proinflammatory cytokine IL-6 and soluble receptors of tumor necrosis factor alpha (sTNF-RI and sTNF-RII), interferon gamma–inducible protein 10 (IP-10 or CXCL10), and the cellular adhesion molecules soluble vascular cell adhesion molecule 1 (sVCAM-1) and soluble intercellular adhesion molecule 1 (sICAM-1), were determined by quantitative sandwich ELISAs (R&D Systems). The D-dimer level was determined by immunoturbidimetric assay on a STA-R Coagulation Analyzer (Diagnostica-Stago, Parsippany, NJ). The fibrinogen level was determined by particle-enhanced immunonephelometric assay on a BNII nephelometer (Siemens, Malvern, PA). All inter-assay coefficients of variance were <12%, except for very low D-dimer values (<0.20 μg/mL), for which the inter-assay coefficient of variance was 21%. Intra-assay coefficients of variance were not calculated for all but 1 marker because of the small number of samples in each run; for IP-10, the inter-assay coefficient of variance was 3.3%. Plasma Lp-PLA2 concentrations were measured with an ELISA (PLAC Test; diaDexus, South San Francisco, CA). Inter- and intra-assay coefficients of variance were <5% and <9%, respectively. Calculated medians of the measurements were used in the analysis.
This was a prespecified preplanned analysis to assess changes from baseline to 48 weeks in plasma inflammatory and coagulation indices and markers of lymphocyte and monocyte activation.
All analyses were initially performed using intent-to-treat principles based on randomized treatment assignment that used all available data. Modifications to randomized treatment and missing values were ignored. As-treated analyses did not differ from intent-to-treat analyses; therefore, only the former data are presented.
Demographics, clinical characteristics, fasting metabolic parameters, and inflammatory and coagulation markers are described by the study group. Continuous measures are described by medians and interquartile ranges, and nominal variables are described with frequencies and percentages.
Nominal variables were compared using χ2 analysis or Fisher exact test. Continuous measures were tested for normality. For between-group comparisons (at baseline and changes from baseline to 48 weeks), normally distributed variables were compared using t tests, and nonnormally distributed variables were compared using Wilcoxon rank sum tests. For within-group changes from baseline to 48 weeks, normally distributed variables were compared with a paired t test and nonnormally distributed variables were compared with Wilcoxon signed rank test. The nonparametric Spearman correlation was used to estimate correlations among changes of markers of inflammation and immune activation at week 48. Nominal P values are presented throughout, with no correction for multiple comparisons.
All analyses were done using SAS 9.3 (SAS Institute Inc., Cary, NC).
Two hundred and two patients were screened for enrollment; 55 patients were screen failures resulting in an enrollment of 147 patients between March 2011 and August 2012 in the SATURN-HIV study (N = 72 in the rosuvastatin arm, N = 75 in the placebo arm, reported previously,28 and in Figure S1, Supplemental Digital Content, http://links.lww.com/QAI/A608). Demographic information and baseline characteristics are displayed in Table 1; indices were similar between groups. Overall, the median age of the patients was 47 years, 78% were male and 70% were African American, 29% were Caucasian, and 1% was Hispanic. Baseline immune activation markers were also similar between groups, except for the proportion of CD14DimCD16+ monocytes that expressed TF (statin arm = 21.8% and placebo arm = 18.9%, P = 0.047, Table 1); this difference was controlled for in the week 48 analysis. There was no difference in the number of subjects from each group on antihypertensive medication (statin: n = 20; placebo: n = 18, P = 0.60). Seven subjects had active hepatitis B (statin: n = 3; placebo: n = 4, P = 0.74), and 12 subjects had active hepatitis C (statin: n = 5; placebo: n = 7, P = 0.60).
Subject Disposition and Safety Data
Nineteen subjects (6 statin; 13 placebo) withdrew before the week 48 analysis: 16 were lost to follow-up. Two subjects withdrew due to grade 2 myalgias with normal CPK levels; both were on placebo. One subject (also on placebo) was diagnosed with diabetes and dropped out of the study. One additional subject in the statin group stopped treatment at week 5 because of hospitalization for hydration to treat grade 3 myalgias without rhabdomyolysis or renal compromise, but continued to be followed on study, off study drug, and myalgia resolved soon after study drug was discontinued. Three subjects (all on placebo) changed ART regimens between baseline and 48 weeks: one was started on abacavir in place of didanosine, one switched Truvada to Epzicom, and another stopped lamivudine/zidovudine and started emtricitabine/tenofovir and maraviroc. Overall, 14 subjects (7 on statin and 7 on placebo) had HIV-1 RNA >50 copies per milliliter at week 48. The patient flowchart is provided as Figure S1 (see Supplemental Digital Content, http://links.lww.com/QAI/A608). HIV-1 RNA levels, CD4+ T cell counts, CD4/CD8 cell ratios, or their 48-week changes did not differ between the groups at 48 weeks.
Changes in Fasting Lipid levels
Lipid profiles in the rosuvastatin group changed by week 48, with a significant decrease in LDL levels (−23.4%, P < 0.0001) and a nonsignificant increase in high-density lipoprotein (relative increase from baseline = 0.7%, P = 0.76) and triglyceride (5.5%, P = 0.47). Patients receiving placebo did not have significant changes in these indices (high-density lipoprotein = 3.7%, P = 0.33; LDL = 7.5%, P = 0.053; and triglycerides = 4.2%, P = 0.51). The difference between groups was significant only for the changes in LDL-C (P < 0.0001 between groups).
Changes in Markers of Immune Activation
There were several significant differences between treatment arms when comparing levels of cellular and plasma markers of immune activation at week 48 (Table 2), extending our findings from the 24-week timepoint.28,29 Rosuvastatin administration reduced levels of sCD14 by 10.4%, relative to baseline levels, whereas subjects receiving placebo had a 0.5% increase in sCD14 levels (P = 0.006, Fig. 1A). Subjects receiving rosuvastatin also had a greater reduction in the proportion of TF+ CD14DimCD16+ (patrolling) monocytes compared with the reduction seen in subjects receiving placebo (−41.6% vs −18.8%, P = 0.005, Fig. 1B). Among the patients receiving rosuvastatin, there were reductions from baseline levels in the proportions of TF expressing cells among all monocyte subsets (TF+ CD14+CD16−, −52.1%, P = 0.0002; TF+ CD14+CD16+, −33.7%, P < 0.0001; TF+ CD14DimCD16+, −41.6%, P < 0.0001, Table 2). Proportions of all monocytes that expressed TF tended to decrease more in patients receiving a statin (−43.2%) compared with the decrease measured in patients receiving placebo (−27.2%, P = 0.087). Among patients in the active group, we did not see a significant relationship between changes in relative LDL levels and changes in the proportions of monocyte subsets that expressed TF or the soluble markers of monocyte activation, sCD14 and sCD163 (Table 3).
Plasma levels of vascular inflammation (Lp-PLA2) were reduced significantly from baseline in the statin arm (−12.2%) compared with the placebo arm (−1.7%, P = 0.0007, Fig. 1C). There was a modest relationship between relative changes in LDL and Lp-PLA2 (r = 0.25, P = 0.05) among patients receiving rosuvastatin. There were no differences in the relative changes of several markers between the groups at week 48; however, many markers were reduced relative to baseline within the statin group, including levels of sCD163 (−12.3%, P = 0.001), sTNF-RI (−0.7%, P = 0.03), and fibrinogen (−6.0%, P = 0.014). Also among the active group, the relative changes in VCAM-1, TNF-RII, and fibrinogen were all related (Table 3), and none of these changes were related to changes in LDL.
Plasma levels of interferon gamma–induced protein 10 (IP-10) were reduced significantly, relative to baseline levels, in patients receiving rosuvastatin (−27.5%), compared with levels in patients receiving placebo (−8.2%, P = 0.03, Fig. 1D). Relative changes in IP-10/CXLC10 levels among the statin group were related to relative changes in plasma D-dimer (r = 0.37, P = 0.012), VCAM-1 (r = 0.61, P < 0.0001), fibrinogen (r = 0.45, P = 0.002), and sCD163 (r = 0.34, P = 0.022) levels (Table 3).
We also assessed markers of T-cell activation; proportions of both CD4+ and CD8+ T cells that express CD38 and HLA-DR were reduced significantly by statin treatment (−38.1% and −44.8%, respectively) compared with placebo (−7.8%, P = 0.0009, and −27.4%, P = 0.0035, Figs. 2B, C). Proportions of cells that express CD38, HLA-DR, and the T-cell exhaustion marker PD-1 are also reduced by statin treatment compared with placebo, among CD4+ (−42.5% vs −24.1%, P = 0.0019) and CD8+ T cells (−45.5% vs −26.3%, P = 0.0016, Table 2). Relative changes in activation marker expression among CD4+ and CD8+ T cells in the active group were not related to changes in LDL or changes in viremia but were directly related to changes in both IP-10 and VCAM-1 levels (Table 3).
The SATURN-HIV study is the first double-blind, randomized placebo-controlled clinical trial of a statin as an immunomodulatory therapy in HIV-infected subjects on suppressive ART. Overall, changes from baseline in monocyte and vascular activation markers at week 24 of treatment were similar at week 48.28,29 Rosuvastatin treatment, compared with placebo, reduced significantly plasma levels of sCD14, a marker that independently predicts morbidity in HIV disease,6 by week 24, and this decline plateaued between weeks 24 and 48. Statin treatment also decreased levels of sCD163, but this decrease was not significantly different from the one measured in the placebo arm; further studies that aimed at defining the potentially divergent mechanisms behind sCD14 and sCD163 induction may prove valuable. Rosuvastatin treatment reduced the proportion of circulating TF+ patrolling (CD14DimCD16+) monocytes by week 24, and this decline continued through 48 weeks. Expression of TF can initiate the extrinsic clotting pathway,33 and we have shown previously that TF levels on monocytes are related to plasma levels of D-dimer34; potentially linking monocyte activation to coagulation in HIV-infected subjects. Statin treatment reduced the proportion of TF+ monocytes in all subsets, but this reduction was not significantly different from reductions seen in placebo recipients among the traditional (CD14+CD16−) and inflammatory (CD14+ CD16+) subsets. The procoagulant phenotype of monocyte subsets we report in HIV disease mirrors that of uninfected patients who have recently experienced an acute coronary event,32 and proportions of CD16+ monocytes in HIV-infected and uninfected subjects have been linked to morbidity,35,36 linking monocyte activation to CVD progression in HIV disease.
Rosuvastatin treatment, compared with placebo treatment, also reduced significantly levels of Lp-PLA2, at week 24, and this decline persisted through week 48. Lp-PLA2 is a specific marker of vascular inflammation, and several studies have shown that increases in Lp-PLA2 predict cardiovascular events in the general population.37–39 By reducing both the procoagulant phenotype of monocytes and inflammation of the vascular endothelium, which may act as a substrate for clot formation, one could speculate that rosuvastatin treatment may reduce coagulation risk in HIV disease.
Plasma levels of IP-10/CXCL10 were reduced from baseline by 48 weeks in the statin group compared with the placebo group; a result that was not seen at 24 weeks. Increased IP-10 levels have been associated with rapid disease progression in acute HIV infection40 and with immunologic treatment failure after ART initiation.41
Markers of T-cell activation were reduced by 48 weeks of rosuvastatin treatment; this is a new result, as 24 weeks of statin treatment had little effect on these indices. Expression of CD38 and HLA-DR on CD8+ T cells is directly related to disease course in untreated HIV disease,21 to plasma levels of LPS in HIV-infected patients,32,42 to proportions of CD16+ monocytes,32 and inversely to CD4+ T-cell reconstitution after initiation of ART.22 Our previous work reported a direct relationship between the proportion of activated CD8+ T cells and mean common carotid artery IMT in HIV disease, and an increased proportion of activated CD8+ T cells among HIV-infected patients with coronary plaque (IMT > 1.5 cm) compared with those without plaque.23 Some T cells are capable of homing to vascular endothelium,43,44 and up to 50% of lymphocytes in human carotid artery plaques are activated CD8+ T cells.45 Ganesan et al30 reported a modest decrease (−3%) in the proportion of CD8+CD38+HLA-DR+ cells after ART-naive patients received high-dose (80 mg) atorvastatin treatment. Here, for the first time, we show that rosuvastatin treatment has beneficial effects on T-cell activation (both CD4+ and CD8+) in subjects on ART. Another notable finding in our study is that rosuvastatin also resulted in significant reductions in the proportions of T cells that express PD-1, a marker reflective of cellular exhaustion and impaired proliferative capacity after T-cell receptor engagement.46 By reducing PD-1 expression, statin treatment may enhance the functionality of T cells in HIV-infected patients.
Forty-eight weeks of statin treatment reduced cellular markers of T-cell activation, monocyte activation, and circulating markers of immune activation, many of which have been linked to disease progression and cardiovascular risk in HIV disease. The decrease in levels of these markers was independent of changes in LDL-C levels, suggesting an anti-inflammatory effect of statins that is not related to an effect on lipid levels. Although we cannot report the exact mechanism by which rosuvastatin treatment is reducing T-cell, monocyte, and endothelial cell activation, one could speculate that rosuvastatin may be reducing the inflammatory response to microbial translocation by reducing Toll-like receptor 4 levels47 or by increasing expression of Kruppel-like factor 2, which can be modulated by statin treatment and has anti-inflammatory effects in several cell types, including T cells,48 endothelial cells,49 and myeloid cells.50 The temporal relationship between the reduction in “innate immune activation” at 24 weeks (sCD14, monocyte activation) and the reduction in “adaptive immune activation” (T-cell activation) at 48 weeks must also be investigated further. Mechanistic studies assessing the effects of rosuvastatin treatment on regulators of innate immune signaling and inflammation are planned; perhaps early, sustained alterations in innate activation provide beneficial effects to the immune system that enable T-cell activation markers to decrease (CD38, HLA-DR) and T-cell responsiveness to improve (lower PD-1 expression).
The SATURN-HIV study has generated several novel observations, but it has limitations. The study population was very specific: HIV-positive subjects with stable ART and low or undetectable plasma HIV-1 RNA levels, normal LDL-C, and heightened baseline immune activation. These patients are also mostly African American men; these factors may potentially limit the generalizability of this result to the HIV-infected population as a whole. By reducing vascular inflammation, the procoagulant phenotype of monocytes, and markers of T-cell activation, we would predict that statin treatment would reduce thrombotic risk, but this is planned for longer studies with clinical events as study end points. SATURN-HIV does provide though several potential directions for further mechanistic studies and provides rationale for a longer term clinical end point study, exploring the effects of statin treatment on morbidity and mortality in chronic treated HIV disease.
1. Hansson GK. Inflammation
, atherosclerosis, and coronary artery disease. N Engl J Med. 2005;352:1685–1695.
2. Hansson GK, Hermansson A. The immune system in atherosclerosis. Nat Immunol. 2011;12:204–212.
3. Kuller LH, Tracy R, Belloso W, et al.. Inflammatory and coagulation biomarkers and mortality in patients with HIV infection. PLoS Med. 2008;5:e203.
4. Emery S, Neuhaus JA, Phillips AN, et al.. Major clinical outcomes in antiretroviral therapy (ART)-naive participants and in those not receiving ART at baseline in the SMART study. J Infect Dis. 2008;197:1133–1144.
5. Kalayjian RC, Machekano RN, Rizk N, et al.. Pretreatment levels of soluble cellular receptors and interleukin-6 are associated with HIV disease progression in subjects treated with highly active antiretroviral therapy. J Infec Dis. 2010;201:1796–1805.
6. Sandler NG, Wand H, Roque A, et al.. Plasma levels of soluble CD14 independently predict mortality in HIV infection. J Infec Dis. 2011;203:780–790.
7. Brown TT, Tassiopoulos K, Bosch RJ, et al.. Association between systemic inflammation
and incident diabetes in HIV-infected patients after initiation of antiretroviral therapy. Diabetes Care. 2010;33:2244–2249.
8. Ross AC, Rizk N, O'Riordan MA, et al.. Relationship between inflammatory markers, endothelial activation markers, and carotid intima-media thickness in HIV-infected patients receiving antiretroviral therapy. Clin Infect Dis. 2009;49:1119–1127.
9. Tabib A, Leroux C, Mornex JF, et al.. Accelerated coronary atherosclerosis and arteriosclerosis in young human-immunodeficiency-virus-positive patients. Coron Artery Dis. 2000;11:41–46.
10. Matta F, Yaekoub AY, Stein PD. Human immunodeficiency virus infection and risk of venous thromboembolism. Am J Med Sci. 2008;336:402–406.
11. Hsue PY, Lo JC, Franklin A, et al.. Progression of atherosclerosis as assessed by carotid intima-media thickness in patients with HIV infection. Circulation. 2004;109:1603–1608.
12. Hsue PY, Hunt PW, Sinclair E, et al.. Increased carotid intima-media thickness in HIV patients is associated with increased cytomegalovirus-specific T-cell responses. AIDS. 2006;20:2275–2283.
13. Hsue PY, Giri K, Erickson S, et al.. Clinical features of acute coronary syndromes in patients with human immunodeficiency virus infection. Circulation. 2004;109:316–319.
14. Friis-Moller N, Weber R, Reiss P, et al.. Cardiovascular disease risk factors in HIV patients–association with antiretroviral therapy. Results from the DAD study. AIDS. 2003;17:1179–1193.
15. Friis-Moller N, Reiss P, Sabin CA, et al.. Class of antiretroviral drugs and the risk of myocardial infarction. N Engl J Med. 2007;356:1723–1735.
16. Mauri L, Hsieh WH, Massaro JM, et al.. Stent thrombosis in randomized clinical trials of drug-eluting stents. N Engl J Med. 2007;356:1020–1029.
17. Crum-Cianflone NF, Weekes J, Bavaro M. Thromboses among HIV-infected patients during the highly active antiretroviral therapy era. AIDS Patient Care STDS. 2008;22:771–778.
18. Fultz SL, McGinnis KA, Skanderson M, et al.. Association of venous thromboembolism with human immunodeficiency virus and mortality in veterans. Am J Med. 2004;116:420–423.
19. Klein SK, Slim EJ, de Kruif MD, et al.. Is chronic HIV infection associated with venous thrombotic disease? A systematic review. Neth J Med. 2005;63:129–136.
20. Triant VA, Lee H, Hadigan C, et al.. Increased acute myocardial infarction rates and cardiovascular risk factors among patients with human immunodeficiency virus disease. J Clin Endocrinol Metab. 2007;92:2506–2512.
21. Giorgi JV, Hultin LE, McKeating JA, et al.. Shorter survival in advanced human immunodeficiency virus type 1 infection is more closely associated with T lymphocyte activation than with plasma virus burden or virus chemokine coreceptor usage. J Infect Dis. 1999;179:859–870.
22. Hunt PW, Martin JN, Sinclair E, et al.. T cell activation is associated with lower CD4+ T cell gains in human immunodeficiency virus-infected patients with sustained viral suppression during antiretroviral therapy. J Infect Dis. 2003;187:1534–1543.
23. Longenecker CT, Funderburg NT, Jiang Y, et al.. Markers of inflammation
and CD8 T-cell activation, but not monocyte activation, are associated with subclinical carotid artery disease in HIV-infected individuals. HIV Med. 2013;14:385–390.
24. Kaplan RC, Sinclair E, Landay AL, et al.. T cell activation and senescence predict subclinical carotid artery disease in HIV-infected women. J Infect Dis. 2011;203:452–463.
25. Kaplan RC, Sinclair E, Landay AL, et al.. T cell activation predicts carotid artery stiffness among HIV-infected women. Atherosclerosis. 2011;217:207–213.
26. Jain MK, Ridker PM. Anti-inflammatory effects of statins: clinical evidence and basic mechanisms. Nat Rev Drug Discov. 2005;4:977–987.
27. Ridker PM, Danielson E, Fonseca FA, et al.. Rosuvastatin
to prevent vascular events in men and women with elevated C-reactive protein. N Engl J Med. 2008;359:2195–2207.
28. Funderburg NT, Jiang Y, Debanne SM, et al.. Rosuvastatin
treatment reduces markers of monocyte activation in HIV-infected subjects on antiretroviral therapy. Clin Infect Dis. 2014;58:588–595.
29. Eckard AR, Jiang Y, Debanne SM, et al.. Effect of 24 weeks of statin therapy on systemic and vascular inflammation
in HIV-infected subjects receiving antiretroviral therapy. J Infect Dis. 2014;209:1156–1164.
30. Ganesan A, Crum-Cianflone N, Higgins J, et al.. High dose atorvastatin decreases cellular markers of immune activation without affecting HIV-1
RNA levels: results of a double-blind randomized placebo controlled clinical trial. J Infect Dis. 2011;203:756–764.
31. Lederman MM, Calabrese L, Funderburg NT, et al.. Immunologic failure despite suppressive antiretroviral therapy is related to activation and turnover of memory CD4 cells. J Infect Dis. 2011;204:1217–1226.
32. Funderburg NT, Zidar DA, Shive C, et al.. Shared monocyte subset phenotypes in HIV-1
infection and in uninfected subjects with acute coronary syndromes. Blood. 2012;120:4599–4608.
33. Mackman N. Role of tissue factor
in hemostasis and thrombosis. Blood Cells Mol Dis. 2006;36:104–107.
34. Funderburg NT, Mayne E, Sieg SF, et al.. Increased tissue factor
expression on circulating monocytes
in chronic HIV infection: relationship to in vivo coagulation and immune activation. Blood. 2010;115:161–167.
35. Baker JV, Hullsiek KH, Singh A, et al.. Immunologic predictors of coronary artery calcium progression in a contemporary HIV cohort. AIDS. 2014;28:831–840.
36. Rogacev KS, Cremers B, Zawada AM, et al.. CD14++CD16+ monocytes
independently predict cardiovascular events: a cohort study of 951 patients referred for elective coronary angiography. J Am Coll Cardiol. 2012;60:1512–1520.
37. Vittos O, Toana B, Vittos A, et al.. Lipoprotein-associated phospholipase A2 (Lp-PLA2): a review of its role and significance as a cardiovascular biomarker. Biomarkers. 2012;17:289–302.
38. Brilakis ES, Khera A, Saeed B, et al.. Association of lipoprotein-associated phospholipase A2 mass and activity with coronary and aortic atherosclerosis: findings from the Dallas Heart Study. Clin Chem. 2008;54:1975–1981.
39. Brilakis ES, McConnell JP, Lennon RJ, et al.. Association of lipoprotein-associated phospholipase A2 levels with coronary artery disease risk factors, angiographic coronary artery disease, and major adverse events at follow-up. Eur Heart J. 2005;26:137–144.
40. Jiao Y, Zhang T, Wang R, et al.. Plasma IP-10 is associated with rapid disease progression in early HIV-1
infection. Viral Immunology. 2012;25(4):333–337.
41. Stylianou E, Aukrust P, Bendtzen K, Muller F, Froland SS. Interferons and interferon (IFN)-inducible protein 10 during highly active anti-retroviral therapy (HAART)-possible immunosuppressive role of IFN-alpha in HIV infection. Clin Exp Immunol. 2000;119(3):479–485.
42. Brenchley JM, Price DA, Schacker TW, et al.. Microbial translocation is a cause of systemic immune activation in chronic HIV infection. Nat Med. 2006;12:1365–1371.
43. Dimayuga PC, Chyu KY, Kirzner J, et al.. Enhanced neointima formation following arterial injury in immune deficient Rag-1-/- mice is attenuated by adoptive transfer of CD8 T cells. PLoS One. 2011;6:e20214.
44. Quan L, Jian Z, Ping Z, et al.. Proteinase-activated receptor-1 mediates allogeneic CD8(+) T cell-induced apoptosis of vascular endothelial cells. Med Oncol. 2009;26:379–385.
45. Grivel JC, Ivanova O, Pinegina N, et al.. Activation of T lymphocytes
in atherosclerotic plaques. Arterioscler Thromb Vasc Biol. 2011;31:2929–2937.
46. Petrovas C, Price DA, Mattapallil J, et al.. SIV-specific CD8+ T cells express high levels of PD1 and cytokines but have impaired proliferative capacity in acute and chronic SIVmac251 infection. Blood. 2007;110:928–936.
47. Methe H, Kim JO, Kofler S, et al.. Statins decrease Toll-like receptor 4 expression and downstream signaling in human CD14+ monocytes
. Arterioscler Thromb Vasc Biol. 2005;25:1439–1445.
48. Bu DX, Tarrio M, Grabie N, et al.. Statin-induced Kruppel-like factor 2 expression in human and mouse T cells reduces inflammatory and pathogenic responses. J Clin Invest. 2010;120:1961–1970.
49. Sen-Banerjee S, Mir S, Lin Z, et al.. Kruppel-like factor 2 as a novel mediator of statin effects in endothelial cells. Circulation. 2005;112:720–726.
50. Tuomisto TT, Lumivuori H, Kansanen E, et al.. Simvastatin has an anti-inflammatory effect on macrophages via upregulation of an atheroprotective transcription factor, Kruppel-like factor 2. Cardiovasc Res. 2008;78:175–184.