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Department of Microbiology & Immunology (H.Z., P.B.H.), Division of Gastroenterology, Department of Internal Medicine and McGuire Veterans Affairs Medical Center (W.M.P.), Department of Physiology (V.L.), and Division of Pulmonary and Critical Care Medicine, Department of Internal Medicine (R.N.), Virginia Commonwealth University, Richmond, Virginia
Received for publication March 16, 2005.
Accepted for publication June 23, 2005.
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Abstract |
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). Most patients on HAART develop hyperlipidemia, lipodystrophy, and insulin resistance (Carr et al., 1998; Koster et al., 2003; Beregszaszi et al., 2003; Bongiovanni et al., 2004). Moreover, HAART significantly increases the risk of premature atherosclerosis and myocardial infarction (Holmberg et al., 2004). Recent studies suggest that PIs disrupt cellular lipid homeostasis by increasing the levels of transcriptionally active sterol regulatory element-binding proteins (SREBPs) (Riddle et al., 2001; Williams et al., 2004), endoplasmic reticulum (ER) membrane-bound transcription factors that when proteolytically activated increase the expression of dozens of genes involved in lipid metabolism (Horton et al., 2002). However, the cellular/molecular mechanisms underlying the HIV PI-associated metabolic abnormalities remain elusive and maybe multifactorial.
The unfolded protein response (UPR) is an intracellular signaling pathway, which uses unique regulatory mechanisms to cope with the accumulation of unfolded or misfolded proteins in the ER and plays an important role in regulating cell growth, differentiation, and apoptosis (Oyadomari and Mori, 2004; Zhang and Kaufman, 2004). Several UPR components have been identified in mammalian cells, which include three transducers [ER transmembrane kinase/endoribonuclease IRE1, doubled-stranded RNA-activated protein kinase-like ER kinase, and activating transcription factor 6 (ATF-6)] and one master regulator [an ER chaperone protein (BiP/GRP78; Zhang and Kaufman, 2004)]. Under various physiological and pathological conditions, the protein folding in the ER is impaired, which causes ER stress. Many ER stress inducers may activate the UPR: accumulation of misfolded proteins, inhibition of N-linked glycosylation, depletion of ER calcium stores, glucose deprivation, redox status, and accumulation of free cholesterol in the ER (Feng et al., 2003; Rutkowski and Kaufman, 2004). Although an elicited response aids the cell in surviving stress, prolonged activation of the UPR ultimately leads to programmed cell death.
Macrophages are the most prominent cell type present in atherosclerotic lesions and play essential roles in all phases of atherosclerosis (Tabas, 2004). It has been demonstrated recently that accumulation of excess free cholesterol in the ER can activate the UPR and induce apoptosis in macrophages (Feng et al., 2003). In the present study, we examined the effects of different HIV PIs on lipid metabolism and the UPR activation in macrophages. The results show that HIV PIs increased levels of transcriptionally active SREBPs, induced accumulation of intracellular free cholesterol, decreased ER calcium stores, activated the UPR, and significantly increased apoptosis in macrophages. These results provide novel insights into the cellular mechanisms whereby HIV PIs induce lipid dysregulation and accelerate atherosclerosis.
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Materials and Methods |
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Cell Culture and HIV PI Treatment. Mouse J774A.1 macrophages were maintained in DMEM supplemented with 10% FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin at 37°C with 5% CO2. Cells from passages six to nine were used in these studies. Ritonavir and atazanavir were dissolved in ethanol. Indinavir was dissolved in H2O. HIV PIs were directly added to culture medium (final concentration, 5 to 50 µM) and incubated for 0.5 to 24 h.
Isolation of Mouse Peritoneal Macrophages. Adult male C57BL/6 mice were injected intraperitoneally with 0.5 ml of phosphate-buffered saline (PBS) containing 40 µg of concanavalin A. The macrophages were harvested 72 h after injection by peritoneal lavage. The harvested cells were cultured in DMEM containing 10% fetal bovine serum (FBS) and 20% L-cell-conditioned medium (Stanley et al., 1976; Koster et al., 2003). The medium was replaced every 24 h until the macrophages were confluent.
Filipin Staining of Free Cholesterol. Mouse J774A.1 macrophages were plated on 22 x 22-mm glass cover slips in six-well plates. The medium was replaced after 24 h, and cells were treated with control vehicle, HIV PIs (5-50 µM), or thapsigargin (100 nM) for 24 h. The cells were fixed with 3.7% formaldehyde in PBS for 10 min and permeabilized with 0.1% Triton X-100 in PBS for 3 min at 4°C. After washing with PBS, the cells were stained with filipin (50 µg/ml) dissolved in PBS containing 0.5% BSA for 30 min at 37°C followed by extensive washing with PBS. The coverslips were mounted onto glass slides using antifade mounting solution. The filipin-free cholesterol complexes were visualized by fluorescence microscopy using Olympus epifluorescence microscope with excitation at 380/40 nm and emission at 485/35 nm.
Measurement of Intracellular Free Cholesterol and Cholesterol Ester. Mouse J774A.1 macrophages were plated on 100-mm plates. The medium was replaced after 24 h, and cells were treated with control vehicle, HIV PIs (5-25 µM), or thapsigargin (100 nM) for 24 h. The cells were collected and washed with PBS. The intracellular total and free cholesterol were measured by using Wako Cholesterol E and Free Cholesterol assay kits (Wako Bioproducts). The amount of cholesterol ester was calculated by subtracting free cholesterol from total cholesterol.
Western Blot Analysis. The nuclear extract was isolated from cells as described previously (Williams et al., 2004). The protein concentration was determined using Bio-Rad protein assay reagent. The nuclear extract (15 µg of protein) was resolved on 10% Bis-Tris or 7% Tris-Acetate NuPAGE Novex gels and transferred to Nitrocellulose membranes. Immunoblots were blocked overnight at 4°C with 5% nonfat milk in Tris-buffered saline and incubated with antibodies to CHOP, XBP-1, ATF-4, SREBP-1, SREBP-2, or lamin B. Immunoreactive bands were detected using horseradish peroxidase-conjugated secondary antibody and the Western Lightning Chemiluminescence reagent plus. The density of immunoblot was analyzed using Image J computer software (http://rsb.info.nih.gov/ij/).
Cholesterol Esterification Assay. Mouse J774A.1 macrophages were plated in DMEM containing 10% FBS for 24 h. Media were replaced with DMEM containing 10% lipoprotein-deficient bovine calf serum. Cells were treated with ritonavir (15 or 25 µM) or control vehicle for 24 h in the presence (20 µg/ml) or absence of LDL and pulsed with [3H]oleate (2 Ci/ml) for 2 h. The lipids were extracted with hexane/isopropyl alcohol [3:2 (v/v)] as described previously (Williams et al., 2004). A recovery standard (30 µg of cholesteryl oleate, 30 µg of triolein, and 0.0005 µCi of [14C]cholesteryl oleate) was added. The extracted samples were dried under a nitrogen gas atmosphere. The lipids were separated by thin-layer chromatography using heptane/ethyl ether/acetic acid [90:30:1 (v/v/v)] and visualized by iodine. The [3H]cholesteryl oleate was quantified by liquid scintillation spectrometry.
Analysis of Apoptosis by Annexin V and Propidium Iodine Staining. Mouse J774A.1 macrophages were treated with various concentrations of ritonavir for 24 h and stained with Annexin V and propidium iodide using BD ApoAlert Annexin V kit according to the protocol recommended by the manufacturer. Annexin V/propidium iodide-stained cells were visualized under confocal fluorescence microscope with a 40x oil immersion objective using a dual filter set for FITC and rhodamine. Cells stained with Annexin V and propidium iodide were further analyzed by two-color flow cytometry to quantify the apoptotic cells. Annexin V and propidium iodide emissions were detected in the FL1 and FL2 channels of a Cytomics FC 500 flow cytometer (Beckman Coulter, Fullerton, CA). At least 10,000 cells were analyzed in each experiment.
Measurement of Caspase-12 Activity. Mouse J774A.1 macrophages were treated with various concentrations of ritonavir or thapsigargin (100 nM) for 24 h. The total cell lysate was prepared. The caspase-12 activity was measured by using Caspase-12 fluorometric assay kit according to the manufacturer instruction (BioVision).
Assay of Endoplasmic Reticulum Calcium Pools. Mouse J774A.1 macrophages were grown on 22 x 40-mm coverslips and treated with vehicle or ritonavir for 24 h. The cells were loaded with 4 µM Fura-2 AM and 0.3% Plurinic F-127 in HBSS at 37°C for 60 min and incubated in HBSS for additional 30 min, then mounted on the stage of an Axioskop 2 plus upright fluorescence microscope (Carl Zeiss GmbH, Jena, Germany) equipped with a 40x objective. After washing with HBSS without Ca2+ and Mg2+ for three times, the cells were stimulated with 100 nM thapsigargin. Fluorescence images (510-nm emission after alternate 340- and 380-nm excitation) before and after addition of thapsigargin were collected at 15-s intervals through a cooled charge-coupled device camera (TILL Photonics LLC, Martinsreid, Germany) which is attached to an image intensifier (VS4-1845; Videoscope, Washington, DC), an epifluorescent light source (Polychrome IV; TILL Photonics), a 515 nm dichroic beam splitter, and a 535 nm emission filter (20 nm band pass; Omega Optical, Brattleboro, VT). The 340:380 ratios of individual cells in these images were analyzed using TILLvisION version 3.1 imaging software.
RNA Isolation and Real-time Quantitative PCR. Total cellular RNA was isolated from mouse J774A.1 macrophages after treatment with ritonavir (15 µM), thapsigargin (100 nM), or vehicle control for 24 h, using an Ambion RNAqueous kit. Total RNA (10 µg) was used for first-strand cDNA synthesis using High-Capacity cDNA archive kit (Applied Biosystems). The mRNA levels of ABCA1, ABCG1, CD36, and LDLR were quantified using the specific gene expression assay kits for mouse ABCA1, ABCG1, CD36, and LDLR on an ABI PRISM 7700 Sequence Detection System. The mRNA values for each gene were normalized to internal control 
-actin mRNA. The ratio of normalized mean value for each treatment groups to vehicle control was calculated.
Oil Red O Staining. Mouse J774A.1 macrophages were plated on 22 x 22-mm glass coverslips in six-well plates. The medium was replaced after 24 h. Cells were loaded with Ac-LDL (50 µg/ml) or vehicle control for 24 h then treated with ritonavir (15 µM), thapsigargin (100 nM), or vehicle control for 24 h. Cells were fixed with 3.7% formaldehyde in PBS for 30 min followed by two washes with PBS. The cells were stained with 0.2% Oil Red O in 60% 2-propanol for 10 min and washed with three times with PBS. The images were taken with the use of a microscope (Olympus, Tokyo, Japan) equipped with image recorder under 40x lenses.
Statistical Methods. Student's t test was employed to analyze the differences between sets of data. Statistics were performed using Prism 4 (GraphPad, San Diego, CA).
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). Treatment of mouse macrophages with ritonavir (5-25 µM) significantly activated caspase-12. At a concentration of 15 µM, caspase-12 activity was increased by 1.9-fold (Fig. 4C).
Depletion of ER Calcium Stores by HIV PIs. The ER is a principal site of the synthesis for protein, sterols, cholesterol, and other lipids. Maintenance of ER calcium homeostasis is essential for many cellular functions. Perturbation of ER calcium homeostasis is expected to induce the ER stress. Thapsigargin (a sarcoplasmic/ER calcium-ATPase inhibitor) depletes the ER calcium stores and activates the UPR in many different cells (Yamaguchi and Wang, 2004). To investigate the possible mechanisms of the HIV PI-induced ER stress and activation of the UPR, we assessed the effect of ritonavir on ER calcium stores in mouse J774A.1 macrophages with the fluorescent calcium indicator fura-2/AM. After treatment with different concentrations of ritonavir for 24 h, cells were loaded with fura-2/AM and switched to calcium-free medium. ER calcium release induced by 100 nM thapsigargin was recorded by fluorescence microscopy. Treatment of mouse J774A.1 macrophage with ritonavir decreased the ER calcium content in a dose-dependent manner (Fig. 5). Cells treated with 25 µM ritonavir remarkably reduced the response to thapsigargin, indicating that ER calcium stores were depleted. Atazanavir and indinavir had similar effects on ER calcium stores (data not shown).
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; Hui, 2003; Fontas et al., 2004). Our previous studies showed that indinavir increased the levels of activated SREBP-1 and SREBP-2 in rat hepatocytes (Williams et al., 2004). To examine the effect of HIV PIs on the expression of SREBPs in mouse J774A.1 macrophages, cells were treated with HIV PIs (ritonavir, indinavir, and atazanavir) at therapeutic concentration (15 µM) for various times. The levels of transcriptionally active mature SREBP-1 and SREBP-2 were significantly increased as shown in Fig. 6. Treatment of these cells with ritonavir (15 µM) for 6 h, increased the levels of mature SREBP-1 and SREBP-2 by 273% and 82% (p < 0.05), respectively. This HIV PI also increased the level of the precursor form of SREBP-1 (data not shown).
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Effects of Ritonavir on Free Cholesterol Accumulation and Esterification in Macrophages. Maintenance of the cellular cholesterol homeostasis is crucial to various cellular functions. To further define whether HIV PI-induced activation of SREBPs is associated with a corresponding increase in intracellular free cholesterol, mouse macrophages were treated with different concentrations of ritonavir (5-50 µM) for 24 h, and the free cholesterol was detected with filipin staining. As shown in Fig. 7A, ritonavir modestly induced the accumulation of free cholesterol in intracellular membranes of mouse macrophages. Treatment with 15 to 25 µM ritonavir increased intracellular free cholesterol content (Fig. 7B). However, treatment with thapsigargin (100 nM) for 24 h did not cause the accumulation of free cholesterol in macrophage (Fig. 7, A and B). These results indicate that activation of UPR is not the direct cause of the HIV PI-induced free cholesterol accumulation in mouse macrophages. We were surprised to find that even though intracellular cholesterol levels increased, there was a significant decrease in endogenous cholesterol esterification (Fig. 7C). In contrast, exogenous cholesterol esterification was not affected (Fig. 7D). The amount of total esterified cholesterol was not affected by ritonavir (Fig. 7B).
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Effects of Ritonavir on the Expression of ABCA1, ABCG1, CD36, and LDLR in Macrophages. To determine whether HIV PIs regulate the genes related to cholesterol uptake and efflux, such as LDLR, CD36, ABCA1, and ABCG1, we used quantitative PCR to measure the mRNA level of these receptors and lipid transporters in control and ritonavir-treated cells. Mouse J774A.1 cells were treated with ritonavir (15 µM), thapsigargin (100 nM), or control vehicle for 24 h, and total RNA was isolated. The mRNA levels of ABCA1, ABCG1, LDLR, and CD 36 were quantified using specific gene expression assay kits. The results (Table 1) show that ritonavir had no effect on mRNA levels of ABCA1 and ABCG1, but significantly increased CD36 and LDLR mRNA levels. However, thapsigargin, a positive control of ER stress inducer, significantly decreased CD36 mRNA level and had little effect on ABCA1, ABCG1, and LDLR mRNA levels.
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Ritonavir Activated the UPR and Induced Apoptosis in Foam Cells. Lipid-laden macrophages (foam cells) are found in all stages of atherosclerosis. Unstable human atherosclerotic plaques constitute numerous foam cells. Apoptosis of foam cells is extremely detrimental. To further examine whether HIV PIs can also induce ER stress, activate the UPR, and increase apoptosis in foam cells, mouse J774A.1 cells were loaded with Ac-LDL (50 µg/ml) or vehicle control for 24 h and then treated with ritonavir (15 µM), thapsigargin (100 nM), or vehicle for 24 h. The intracellular lipids were stained with oil red O. The expression of CHOP, ATF-4, and XBP-1 were detected by Western blot; the apoptotic cells were detected by Annexin V-FITC/propidium iodide staining with confocal fluorescence microscopy. As shown in Fig. 8, A and B, ritonavir increased the lipid accumulation not only in Ac-LDL-loaded cells but also in normal macrophages, suggesting that ritonavir may promote the foam cell formation. In Ac-LDL-induced foam cells, ritonavir modestly increased the accumulation of the intracellular free cholesterol, but significantly increased cholesterol ester accumulation (Fig. 8C). Treatment of foam cells with ritonavir (0-50 µM) induced dose-dependent apoptosis. As shown in Fig. 9, Ac-LDL-induced foam cells were more sensitive to ritonavir compared with normal cholesterol loaded macrophages. Likewise, ritonavir activated the UPR in foam cells (Fig. 10).
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), promote atherosclerosis (Sklar and Masur, 2003), and increase the risk of myocardial infarction (Friis-Moller et al., 2003) in patients with HIV infection. Studies using rodent macrophages (Dressman et al., 2003) and hepatocytes (Liang et al., 2001; Riddle et al., 2001) have shown that HIV PIs dysregulate cellular lipid metabolism. However, the exact mechanism(s) altering lipid metabolism is unclear and may be multifactorial. Our current studies show that HIV PIs (ritonavir, indinavir, and atazanavir) activate the UPR and induce apoptosis in both normal cholesterol loaded macrophages and in Ac-LDL-loaded macrophages. We were surprised to find that HIV PIs markedly increased the mature forms of nuclear SREBP-1 and SREBP-2 (Fig. 6) and increased cholesterol in intracellular membranes (Fig. 7). HIV PI-induced disruption of lipid homeostasis and enhanced programmed cell death occurred in macrophages at concentrations (5-15 µM) that are within serum levels reported for patients infected with HIV who are taking these medications (Flexner, 1998).
In mammalian cells, the ER is involved in the proper folding and post-translational modification of proteins. The ER also maintains very low membrane cholesterol content (Feng et al., 2003) and is very sensitive to alterations in cellular homeostatic mechanisms. A number of cellular stress conditions can alter the normal rate of folding and processing of newly synthesized proteins (Pahl, 1999). The ER has evolved cell signaling pathways to respond to the accumulation of unfolded or misfolded proteins, collectively referred to as the UPR (reviewed in Zhang and Kaufman, 2004). Activation of the UPR allows mammalian cells to increase protein folding and degradation pathways and differentially inhibits most protein synthesis. However, prolonged activation of the UPR can lead to apoptosis.
Macrophages are the major cell type found in atherosclerotic lesions, and inappropriate and/or excessive apoptosis of these cells is believed to play a key role in both the initiation and the progression of atherosclerosis. Compounds that activate the UPR in vivo seem to accelerate cardiovascular disease. For example, patients with hyperhomocystinemia accumulate high serum levels of homocysteine because of genetic defects in genes involved in homocysteine metabolism (Outinen et al., 1999). Hyperhomocystinemia is an independent risk factor for atherosclerosis and cardiovascular disease. Homocysteine has been shown to be an activator of the UPR in hepatocytes and vascular endothelial cells (Werstuck et al., 2001). Activation of the UPR by homocysteine is associated with dysregulation of lipid metabolism, increased apoptosis, and accelerated atherosclerosis. We hypothesize that HIV PIs accelerate atherosclerosis and cardiovascular disease as a result of their ability to disrupt normal lipid metabolism and to activate the UPR.
It is currently unclear how HIV protease inhibitors activate the UPR; a number of cellular stress signals, such as depletion of ER calcium stores, increased cholesterol in ER membranes, deprivation of glucose (Oyadomari and Mori, 2004), or inhibition of proteasome activity (Parker et al., 2005) can activate this system. Recent studies have shown that overloading of free cholesterol in the ER causes depletion of ER calcium stores (by inhibiting the sarcoplasmic-endoplasmic reticulum calcium ATPase-2b) (Li et al., 2004), activates the UPR, and induces apoptosis in macrophages (Feng et al., 2003). In the present study, we discovered that HIV PIs induced accumulation of free cholesterol and lipid in intracellular membranes of mouse macrophages and depleted ER calcium stores (as shown in Figs. 5; 7, A and B; and 8). However, there was no direct effect of HIV PIs on ER calcium release (data not shown). Therefore, one possible mechanism by which HIV PIs activate the UPR in macrophages is the accumulation of intracellular free cholesterol followed by the depletion of ER calcium stores.
Macrophages have acquired multiple mechanisms to prevent the accumulation of excess free cholesterol, including an increase in cholesterol esterification, induction of free cholesterol efflux, and down-regulation of cholesterol biosynthesis (Zhang and Kaufman, 2003). HIV PIs may promote foam cell formation and atherosclerosis by modulation of CD36 (Dressman et al., 2003) and the LDLR (Tran et al., 2003) and by increasing cholesterol biosynthesis. In the present study, we demonstrated that ritonavir significantly up-regulates CD36 and LDLR mRNA expression but has no effect on ABCA1 and ABCG1 mRNA levels. It has been demonstrated that overloading of free cholesterol in macrophages decreases ABCA1-mediated cholesterol efflux by accelerating the degradation of ABCA1 protein (Feng and Tabas, 2002). Thus, HIV PI-induced accumulation of intracellular free cholesterol and lipid may indirectly impair the normal function of ABCA1 protein. It has also been shown that disruption of intracellular cholesterol transport may adversely affect cholesterol efflux and cholesterol esterification (Tabas, 2002).
It has been reported that depletion of glucose causes the accumulation of unfolded proteins, induces ER stress, and activates the UPR (Pahl, 1999). It has been demonstrated that indinavir selectively and reversibly inhibits the glucose transporter isoform GluT4 at therapeutic concentrations (Murata et al., 2002; Koster et al., 2003) and impairs SREBP-1 intranuclear localization (Caron et al., 2001). These observations suggest another possible mechanism by which HIV PIs might induce ER stress and activate the UPR. However, whether HIV PIs disrupt glucose homeostasis in macrophages needs further investigation.
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) suggest that inhibition of proteasome activity and differential glucose transport by HIV PIs are proximal events eliciting the UPR, which can regulate lipogenic pathways in hepatocytes or adipocytes. Several studies have shown that ritonavir is a reversible inhibitor of proteasome and inhibits the chymotrypsin-like activity of the proteasome while enhancing the trypsin-like activity at high concentration (50-100 µM) (Lagathu et al., 2004; Laurent et al., 2004). However, a block in proteasomal housekeeping functions in intact cells by ritonavir requires 50 to 100 µM concentrations of the drug (Andre et al., 1998). In the present studies, the UPR is activated by ritonavir at concentrations (5-15 µM) at which essential functions of the proteasome are not yet blocked. In vitro studies have also shown that different HIV PIs differ in their effects on proteasome activity. Indinavir and nelfinavir have no effect on proteasome activity even at high concentration (100 µM) (Andre et al., 1998). However, in our studies, we observed that indinavir induced the UPR activation and disrupted lipid metabolism at clinically relevant concentration (15 µM) in macrophages. Nelfinavir is also able to induce the UPR activation at even lower concentrations (1-10 µM) (unpublished data). Therefore, the HIV PI-induced activation of the UPR may not be due to the dysfunction of the proteasome in macrophages. In summary, the current study provides novel insights into how HIV PIs induce ER stress, activate the UPR, and induce apoptosis in macrophages (Fig. 11). The effects of HIV PIs on accumulation of intracellular free cholesterol and lipids and depletion of ER calcium stores represent a potential mechanism by which HIV PIs induce atherosclerosis and cardiovascular disease in patients infected with HIV who are undergoing HARRT. Different HIV PIs have different effects in different cell types. We cannot rule out the involvement of other target proteins in the HIV PI-induced activation of the UPR and apoptosis in macrophages. A better understanding of the cellular and molecular mechanisms of HIV PI-induced metabolic abnormality may provide useful information for the development of new drugs and therapeutic strategies.
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Acknowledgements |
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Footnotes |
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ABBREVIATIONS: HIV, human immunodeficiency virus; PI, protease inhibitor; HAART, highly active antiretroviral therapy; SREBP, sterol regulatory element-binding protein; ER, endoplasmic reticulum; UPR, unfolded protein response; AM, acetoxymethyl ester; LDLR, low-density lipoprotein receptor; LDL, low-density lipoprotein; Ac-LDL, acetylated low-density lipoprotein; FBS, fetal bovine serum; PBS, phosphate-buffered saline; CHOP, C/EBP homologous protein; DMEM, Dulbecco's modified Eagle's medium; FITC, fluorescein isothiocyanate; HBSS, HEPES-buffered saline solution; XBP-1, X-box-binding protein 1; ATF, activating transcription factor.
Address correspondence to: Dr. Phillip B. Hylemon, Department of Microbiology and Immunology, Virginia Commonwealth University, P.O. Box 980678, Richmond, VA 23298-0678. E-mail: hylemon{at}hsc.vcu.edu
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