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CARDIOVASCULAR

Selective Clearance of Macrophages in Atherosclerotic Plaques by the Protein Synthesis Inhibitor Cycloheximide

Division of Pharmacology, University of Antwerp, Wilrijk, Belgium (V.C., W.M., A.G.H., G.R.Y.D.M.); and Laboratory of Cell Biology and Histology, University of Antwerp, Wilrijk, Belgium (J.-P.T.)

Received September 14, 2006; accepted November 27, 2006.

	Abstract

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Macrophages are an essential component of unstable atherosclerotic plaques and play a pivotal role in the destabilization process. We have demonstrated previously that local delivery of the mammalian target of rapamycin (mTOR) inhibitor everolimus selectively clears macrophages in rabbit plaques. Because mTOR controls mRNA translation, inhibition of protein synthesis might induce selective macrophage cell death. We therefore investigated in the present study the effect of the protein synthesis inhibitor cycloheximide on macrophage and smooth muscle cell (SMC) viability. In vitro studies with cultured macrophages and SMCs showed that cycloheximide induced selective apoptosis of macrophages in a concentration- and time-dependent manner. Moreover, macrophages could be selectively depleted in rabbit carotid artery rings with collar-induced atherosclerotic plaques after in vitro treatment with cycloheximide. Local in vivo administration of cycloheximide via osmotic minipumps to rabbit carotid arteries with collar-induced atherosclerotic plaques significantly reduced the macrophage but not the SMC content. Cycloheximide-treated plaques showed signs of apoptosis (increased terminal deoxynucleotidyl transferase end labeling and fluorescein isothiocyanate-Val-Ala-DL-Asp(O-methyl)-fluoromethylketone labeling) that did not colocalize with SMCs. Organ chamber studies demonstrated that the functionality of SMCs and the endothelium were not influenced by cycloheximide treatment. All together, these findings demonstrate that cycloheximide decreases the macrophage load in atherosclerotic plaques by induction of apoptosis without changing SMC content or contractility.

	Materials and Methods

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Cell Culture. The murine macrophage cell line J774A.1 (American Type Culture Collection, Manassas, VA) was grown in RPMI 1640 medium (Invitrogen, Carlsbad, CA) supplemented with 10% heat-inactivated fetal bovine serum, 100 U/ml penicillin, 100 U/ml streptomycin, 50 µg/ml gentamicin, and 20 U/ml polymyxin B sulfate in a humidified 5% carbon dioxide incubator at 37°C. Murine C2C12 myoblasts (European Collection of Cell Cultures, Salisbury, UK) were grown in Dulbecco's modified Eagle's medium supplemented with 10% serum and antibiotics. Rabbit alveolar macrophages were obtained from New Zealand White rabbits fed a normal chow by bronchoalveolar lavage. In brief, rabbits were sacrificed with an overdose of sodium pentobarbital (CEVA Santé Animale, Libourne, France). The lungs were carefully removed and lavaged four times with 50 ml of phosphate-buffered saline (Invitrogen). Cells were washed twice with phosphate-buffered saline and resuspended in serum-containing RPMI 1640 medium. Smooth muscle cells were isolated from rabbit aorta by digestion with collagenase type 2 (Worthington, Lakewood, NJ) and elastase (Sigma-Aldrich, St. Louis, MO) (60–90 min, 37°C) at 300 and 5 U/ml final concentration, respectively, and cultured in serum-containing F10 Ham's medium (Invitrogen). After overnight incubation at 37°C, nonadherent cells were washed away and medium was replaced. Cells were treated with cycloheximide (Sigma-Aldrich) at different concentrations (0–30 µg/ml) and for various times (0–24 h).

To examine de novo protein synthesis, cells were treated with cycloheximide (10 µg/ml) for 1 h and pulse-labeled for 1 h at 37°C with 5 µCi of Pro-mix L-[³⁵S] in vitro cell labeling mix (GE Healthcare, Little Chalfont, Buckinghamshire, UK) in cysteine/methionine-free Dulbecco's modified Eagle's medium. After homogenization of cells in hypotonic lysis buffer (10 mM Tris, 1 mM EDTA, and 0.2% Triton X-100), labeled proteins were precipitated with 10% trichloroacetic acid, resuspended in 0.2 N NaOH, and measured by liquid scintillation counting. Evaluation of cell viability was based on the incorporation of the supravital dye neutral red by viable cells (Lowik et al., 1993).

To examine internucleosomal DNA fragmentation, cells were lysed in 0.5 ml of hypotonic lysis buffer supplemented with 250 µgof proteinase K. Lysates were incubated for 1 h at 50°C, then supplemented with 5-µl volumes of DNase-free RNase A (2 mg/ml) and incubated for an additional hour at 37°C. The samples were precipitated overnight with 0.1 volumes of 3 M sodium acetate and 1 volume of isopropanol. DNA pellets were air dried and dissolved in Tris/EDTA buffer (10 mM Tris and 1 mM EDTA, pH 7.4). After electrophoresis in 2% agarose, DNA laddering was visualized under UV light by staining the agarose with ethidium bromide.

Western Blot Analysis. Cells were lysed in an appropriate volume of Laemmli sample buffer (Bio-Rad Laboratories, Hercules, CA). Cell lysates were heat-denatured for 3 min and loaded on a 12.5% SDS-polyacrylamide gel. After gel electrophoresis, proteins were transferred to an Immobilon-P Transfer membrane (Millipore, Billerica, MA) according to standard procedures. Membranes were blocked in Tris-buffered saline containing 0.05% Tween 20 and 5% nonfat dry milk (Bio-Rad) for 1 h. After blocking, membranes were probed overnight at 4°C with primary antibodies in antibody dilution buffer (Tris-buffered saline/0.05% Tween 20 containing 1% nonfat dry milk), followed by a 1-h incubation with secondary antibody at room temperature. Antibody detection was accomplished with SuperSignal West Pico or SuperSignal West Femto Maximum Sensitivity Substrate (Pierce, Rockford, IL) using a Lumi-Imager (Roche Diagnostics, Mannheim, Germany). The following mouse primary antibodies were used: anti-caspase-3 (clone 19; BD Transduction Laboratories, Lexington, KY), anti-p53 (clone PAb 240; BD Biosciences PharMingen, San Diego, CA) and anti-CHOP (clone B-3; Santa Cruz Biotechnology, Santa Cruz, CA). Rabbit antibodies used in this study include anti-cleaved caspase-3, anti-Erk1/2, anti-phospho-Erk1/2 (Thr202/Tyr204), anti-SAPK/JNK (clone 56G8), anti-phospho-SAPK/JNK (Thr183/Tyr185), anti-p38 mitogen-activated protein kinase (MAPK), anti-phospho-p38 MAPK (Thr180/Tyr182), anti-p70 S6 kinase, anti-phospho-p70 S6 kinase (Thr389), and anti-eIF2 and anti-phospho-eIF2 (Ser51) (Cell Signaling Technology, Danvers, MA). Peroxidase-conjugated secondary antibodies were purchased from Dako Denmark A/S (Glostrup, Denmark).

In Vitro Treatment of Atherosclerotic Plaques. Male New Zealand White rabbits (2.7–3.4 kg, n = 8) were fed a diet supplemented with cholesterol (1.5%) for 14 days. After anesthesia with sodium pentobarbital (30 mg/kg IV), a nonocclusive, biologically inert, flexible silicone collar was placed around both carotid arteries and closed with silicone glue to induce atheroma-like lesions (i.e., intimal thickenings consisting of SMCs and macrophages hereafter referred to as atherosclerotic plaques) (Booth et al., 1989; Kockx et al., 1992; De Meyer et al., 1997). After another 14 days, during which the cholesterol diet was continued, the animals were sacrificed by an overdose of sodium pentobarbital. Carotid arteries were prepared free from surrounding tissues and released from the collars. Two rings were cut from each collared segment and were incubated in serum-containing Ham's F10 medium for 3 or 7 days in the presence or absence of 30 µg/ml cycloheximide. The medium was refreshed daily. After treatment, the carotid artery rings were formalin-fixed for 24 h.

In Vivo Treatment of Atherosclerotic Plaques. In male New Zealand White rabbits (3.1–3.8 kg, n = 11) with atherosclerotic plaques in the carotid arteries previously induced by positioning a silicone collar, an osmotic minipump (type 2ML1; Alzet, Cupertino, CA) was connected to each collar (Matthys et al., 1997) 14 days after initial collar placement. The pumps delivered 10 µl of solution (saline or 10 µg/ml cycloheximide) per hour locally to the carotid artery for 3 days. Thereafter, the rabbits were heparinized (150 U/kg) (LEO Pharmaceutical Products, Ballerup, Denmark) and sacrificed by an overdose of sodium pentobarbital. Three rings were cut from each collar-wrapped segment: one was formalin-fixed for 24 h, another snap-frozen in liquid nitrogen, and a third used for vascular reactivity studies.

Histological Examination. Formalin-fixed carotid artery rings were paraffin-embedded and stained with hematoxylin/eosin or Verhoef's elastin. Immunohistochemical detection was carried out using an indirect antibody conjugate technique (Kockx et al., 1992; De Meyer et al., 2000). The following primary antibodies were used: anti--SMC actin (clone 1A4; Sigma-Aldrich) and MO/RAM11 (anti-rabbit macrophages; Dako Denmark A/S) on paraffin sections; anti-CD31 (JC/70A; Dako Denmark A/S) and anti-FITC (polyclonal anti-fluorescein isothiocyanate horse radish peroxidase conjugated; Dako Denmark A/S) on frozen sections. Horseradish peroxidase-conjugated or Alexa fluor 546-conjugated secondary antibodies were obtained from Dako Denmark A/S and Invitrogen, respectively. For detection of oligonucleosomal DNA cleavage, a stringent terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) technique was used (Kockx et al., 1998). Caspase activation was detected using FITC-labeled Val-Ala-DL-Asp(O-methyl)-fluoromethylketone (FITC-VAD-fmk). The images were analyzed using a color image analysis system (Image Pro Plus 4.1; Media Cybernetics Inc., Silver Spring, MD). Fluorescence images were taken with a confocal laser scanning microscope (LSM510; Carl Zeiss Inc., Thornwood, NY) and further analyzed using the LSM510 imaging software (Carl Zeiss Inc.).

RAM11 positive areas and -SMC actin-positive areas were determined in six random regions of interest (600 x 450 µm each). Intact nuclei were counted in 12 random regions of interest (160 x 120 µm each) on hematoxylin/eosin-stained sections and expressed as the number of intact nuclei per 0.01 mm². The area of the plaque and the media was measured via planimetry on sections stained for elastin.

Vascular Reactivity Studies. A 2-mm segment was cut from each collar-wrapped carotid artery and mounted in organ chambers filled with 10 ml of physiological salt solution (118 mM NaCl, 4.7 mM KCl, 2.5 mM CaCl₂, 1.2 mM KH₂PO₄, 1.2 mM MgSO₄, 25 mM NaHCO₃, 0.025 mM CaEDTA, and 11.1 mM glucose) at 37°C, continuously aerated with 95% O₂/5% CO₂ for force measurements at 6 g loading tension as described previously (Geerts et al., 1999). Tension was measured isometrically with a Statham UC2 force transducer (Gould, Cleveland, OH) connected to a data acquisition system (Moise 3; EMKA Technologies, Paris, France). To evaluate the contractile response of the segments, KCl was used in a single concentration at the beginning of the experiment. KCl was then washed out by rinsing the organ chambers three times with physiological salt solution. Then, a concentration-response curve for serotonin was constructed. Thereafter, the organ chambers were washed three times, a concentration-response curve for phenylephrine was made, and the EC₅₀ was determined. Subsequently, phenylephrine was washed away. The segments were precontracted with phenylephrine (EC₅₀) to make a concentration-response curve for acetylcholine. After rinsing, the organ chambers the segments were precontracted again with the EC₅₀ of phenylephrine, and a concentration-response curve for spermine NONOate (Sigma-Aldrich) was constructed.

Statistical Analysis. All data are presented as mean ± S.E.M. RAM11-positive area, -SMC actin-positive area, intact nuclei, plaque, and medial area were compared using the Wilcoxon signed-rank test (in vitro experiments) or the Mann Whitney U test (in vivo experiments). Cell viability was compared among groups using one-way analysis of variance, followed by Dunnett t test when variances were equal or Dunnett T3 test when variances were unequal as assessed by the Levene's test. Vascular reactivity was evaluated by comparing the EC₅₀ (molar concentration producing half of the maximal response, reflecting the sensitivity to an agonist) and the maximal response for each agonist between control and cycloheximide-treatment with the Student's t test. All statistical analyses were carried out with the SPSS 12.0 software (SPSS, Chicago, IL). Differences were considered significant at p < 0.05.

	Results

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Macrophages but Not Smooth Muscle Cells Were Highly Susceptible to Cycloheximide Treatment. J774A.1 macrophages and SMCs isolated from rabbit aorta were treated in vitro with the protein synthesis inhibitor cycloheximide. This treatment resulted in an almost complete inhibition of de novo protein synthesis (94 ± 1 and 93 ± 1% inhibition with 10 µg/ml cycloheximide in macrophages and SMCs, respectively). Cell death was initiated in both cell types in a concentration- and time-dependent manner (Fig. 1). However, SMCs were much more resistant to cycloheximide-induced cell death compared with J774A.1 macrophages. Similar to J774A.1 cells, rabbit bronchoalveolar macrophages were highly sensitive to cycloheximide treatment and underwent rapid cell death (Fig. 1B). The type of J774A.1 macrophage cell death induced by cycloheximide was characterized by cleavage of caspase-3 and internucleosomal DNA fragmentation (Fig. 2), typical of apoptosis. Western blot analysis of MAPK signaling in J774A.1 cells and C2C12 myoblasts (as resistant to cycloheximide as rabbit SMCs; data not shown) did not reveal major differences (Fig. 3). In both cell types, the stress responsive p38 MAPK was hyperphosphorylated during cycloheximide treatment, whereas Erk1/2 and SAPK/JNK showed a transient phosphorylation status (increased phosphorylation followed by dephosphorylation). In addition, hyperphosphorylation of p70 S6 kinase and dephosphorylation of eIF2 occurred. p53 and CHOP, previously linked to cycloheximide-induced apoptosis, did not seem to be involved in the cell death process because the expression was not detectable (in case of p53) or did not change after treatment (in case of CHOP) (data not shown).

View larger version (14K): [in this window] [in a new window]   Fig. 1. Effect of cycloheximide on the viability of cultured macrophages and SMCs. A, J774A.1 macrophages () and rabbit aortic SMCs () were treated with cycloheximide (0–30 µg/ml) in serum containing medium for 24 h. B, J774A.1 macrophages (), rabbit alveolar macrophages (), and rabbit aortic SMCs () were exposed to 30 µg/ml cycloheximide for up to 24 h. Cell viability was examined by neutral red assays. Data represent the mean ± S.E.M. of three independent experiments, each performed in duplicate. *, p < 0.05; **, p < 0.01; ***, p < 0.001 versus control (analysis of variance, followed by Dunnett test).

View larger version (36K): [in this window] [in a new window]   Fig. 2. Characterization of cycloheximide-induced cell death in J774A.1 macrophages. Cells were treated with 30 µg/ml cycloheximide for 0 to 24 h. Cleavage of procaspase-3 (A) and internucleosomal DNA fragmentation (B) was analyzed using Western blotting and agarose gel electrophoresis, respectively.

View larger version (66K): [in this window] [in a new window]   Fig. 3. Western blot analysis of different phosphorylation events in J774A.1 macrophages and C2C12 myoblasts before and after cycloheximide treatment. Cells were treated with 30 µg/ml cycloheximide for 0 to 4 h.

To examine whether cycloheximide could selectively clear macrophages in atherosclerotic plaques, collared carotid artery segments from hypercholesterolemic rabbits were exposed to cycloheximide for 3 or 7 days in vitro. Thereafter, RAM11 and -SMC actin immunostains were performed to quantify the amount of macrophages and SMCs, respectively. In contrast to the -SMC positive area, the RAM11-positive area in the plaque was significantly reduced after treatment with cycloheximide (Fig. 4). As with the plaque, the -SMC positive area in the media was not affected. Cycloheximide did not affect plaque or medial area (Table 1). The number of cells in the plaque with intact nuclei progressively decreased during treatment, indicating that cycloheximide initiated apoptotic cell death. Nuclear fragmentation of medial SMCs in carotid rings treated with cycloheximide in vitro occurred only after 7 days (Table 1).

View larger version (66K): [in this window] [in a new window]   Fig. 4. In vitro effect of 30 µg/ml cycloheximide on rabbit carotid artery rings with collar-induced atherosclerotic plaques. A, immunoreactivity for RAM11 (brown = macrophages). B, immunoreactivity for -SMC actin (brown = SMCs) of carotid artery rings incubated for 7 days with serum-containing control medium or cycloheximide (CHX)-supplemented medium. P, plaque; M, media; Ad, adventitia. Scale bar, 50 µm. C, percentage RAM11-positive area in the plaque, and percentage -SMC actin-positive area in the plaque or in the media after 3 and 7 days of treatment. Data are shown as mean ± S.E.M. *, p < 0.05 versus control (Wilcoxon signed rank test).

Cycloheximide Induced Selective Macrophage Apoptosis in Vivo. Cycloheximide was locally administered in vivo by connecting an osmotic minipump to a collar around the rabbit carotid artery with previously induced atherosclerotic plaques. Pilot experiments (data not shown) demonstrated that a low concentration of cycloheximide (10 µg/ml) and a short treatment time (3 days) were essential to induce macrophage cell death in the plaque without affecting plaque or medial SMCs, which was in agreement with our in vitro findings. Under these conditions, plaques did not show significant RAM11-staining evocative of macrophage cell death compared with saline-treated plaques (Fig. 5). Initiation of apoptosis was visualized in the plaques by TUNEL, a marker for DNA fragmentation, and by FITC-VAD-fmk, a marker for caspase-activation (Fig. 6). FITC-VAD-fmk labeling was confined to plaque cells and did not colocalize with -SMC actin (Fig. 7). Moreover, -SMC actin staining in both the plaque and the media (Fig. 5), as well as CD31 staining (Fig. 8A), were similar in cycloheximide-treated versus control plaques, indicating that viability of SMCs and endothelial cells was not affected. To determine potential differences in SMC and endothelial function, vascular reactivity of cycloheximide-treated and saline-treated collared segments was compared in organ chamber studies. The contraction capacity in response to KCl, serotonin, or phenylephrine was unaltered (Fig. 9) as was the relaxation capacity in response to acetylcholine or spermine NONOate (Fig. 8, B and C). Moreover, the sensitivity to all agonists was statistically not different between control and cycloheximide-treated ring segments.

View larger version (73K): [in this window] [in a new window]   Fig. 5. In vivo effect of cycloheximide locally administered for 3 days via an osmotic minipump that was connected to a collar around atherosclerotic rabbit carotid arteries. A, immunoreactivity for RAM11 (brown = macrophages). B, immunoreactivity for -SMC actin (brown = SMCs) of carotid arteries, treated for 3 days with saline (control) or 10 µg/ml cycloheximide (CHX). P, plaque; M, media; Ad, adventitia. Scale bar, 50 µm. C, percentage RAM11-positive area in the plaque, and percentage -SMC actin-positive area in the plaque or in the media. Data are shown as mean ± S.E.M. ***, p < 0.001 versus control (Mann Whitney U test).

View larger version (56K): [in this window] [in a new window]   Fig. 6. Characterization of the type of cell death induced by cycloheximide locally administered for 3 days via an osmotic minipump that was connected to a collar around atherosclerotic rabbit carotid arteries. TUNEL labeling or FITC-VAD-fmk immunostaining is shown for saline-treated (control) or cycloheximide-treated (CHX) carotid artery rings. P, plaque; M, media; Scale bar, 50 µm.

View larger version (35K): [in this window] [in a new window]   Fig. 7. Caspase activation as a measure for apoptosis after local in vivo treatment for 3 days of atherosclerotic rabbit carotid arteries with cycloheximide (CHX) or saline (control). Apoptotic cells were visualized using FITC-VAD-fmk labeling (green), whereas smooth muscle cells were detected by immunohistochemical staining for -SMC actin (red). Scale bar, 50 µm. P, plaque; M, media.

View larger version (51K): [in this window] [in a new window]   Fig. 8. Morphology and functionality of the endothelium of atherosclerotic rabbit carotid arteries after local in vivo treatment with cycloheximide or saline via an osmotic minipump connected to a collar around the carotid arteries for 3 days. A, immunoreactivity for CD31 of saline-treated (control) or cycloheximide-treated (CHX) carotid artery rings. Scale bar, 200 µm. Responses to acetylcholine (B) and spermine-NONO-ate (C) after precontraction with phenylephrine. , cycloheximide; , saline. Data represent mean ± S.E.M.

View larger version (10K): [in this window] [in a new window]   Fig. 9. Vasoconstrictor responses to KCl, serotonin, and phenylephrine of atherosclerotic rabbit carotid arteries after local treatment for 3 days with cycloheximide () or saline () via an osmotic minipump connected to a collar around the carotid arteries. A, maximum contraction after depolarization with KCl. Contractile responses to serotonin (B) and phenylephrine (C). Data are shown as mean ± S.E.M.

	Discussion

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The major finding of this study is that the protein synthesis inhibitor cycloheximide cleared macrophages in rabbit atherosclerotic plaques by selective initiation of macrophage apoptosis. The viability and contractility of SMCs as well as the morphology and functionality of the endothelium were not affected. These results are important in terms of plaque stability because macrophages contribute greatly to SMC loss, breakdown of interstitial collagen fibers, and thinning of the fibrous cap (Galis et al., 1994; Boyle et al., 2001; Newby, 2005).

Selective clearance of macrophages in atherosclerotic plaques via macrophage-specific initiation of cell death is a novel concept in cardiovascular research, but it is hampered by specificity problems. Indeed, it is easy to trigger cell death in most vertebrate cells, including macrophages; the problem is targeting only those cells that we wish to die. This is the basis for most cancer therapies but also applies to any situation in which a particular cell population is responsible for disease. A CD11b diphtheria toxin receptor transgenic mouse has been developed in which administration of diphtheria toxin selectively depletes monocytes and macrophages (Stoneman et al., 2005). This toxin had profound effects on atherogenesis but at the same time dramatically reduced peripheral blood monocytes and tissue macrophages in a nonselective way, which makes this approach unsuitable for clinical applications. More recently, benzyloxycarbonyl-Val-Ala-DL-Asp(O-methyl)-fluoromethylketone (z-VAD-fmk), a caspase-inhibitor with broad specificity, proved to be a potent inducer of nonapoptotic cell death in J774A.1 and RAW264.7 macrophages as well as in interferon--primed primary mouse peritoneal macrophages, but not in SMCs or C2C12 myoblasts (Martinet et al., 2006). z-VAD-fmk-treated macrophages overexpressed and secreted several chemokines and cytokines, including tumor necrosis factor- (TNF). The combination of z-VAD-fmk and TNF, but not TNF alone, induced SMC necrosis, suggesting that z-VAD-fmk, despite its selective cell death-inducing capacity, might not be the preferred mechanism to clear macrophages from atherosclerotic plaques because of indirect induction of SMC death. In another study, stent-based delivery of the rapamycin analog and mTOR inhibitor everolimus selectively cleared macrophages in rabbit atherosclerotic plaques via autophagy (Verheye et al., 2007), also known as type II programmed cell death (Yoshimori, 2004), and without causing obvious adverse effects. Because mTOR controls mRNA translation, we hypothesized that inhibition of protein synthesis drives selective induction of macrophage cell death. We therefore examined in the present study the effect of the protein synthesis inhibitor cycloheximide on macrophage and SMC viability in more detail. Cycloheximide was capable of inducing macrophage cell death, but in contrast to everolimus, apoptosis and not autophagy was induced. This finding confirms earlier studies demonstrating that cycloheximide, independently of other stimuli, is capable of triggering apoptotic cell death in both cultured cells (Martin et al., 1990; Blom et al., 1999; Wu et al., 2004) and tissue (Ledda-Columbano et al., 1992; Alessenko et al., 1997; Higami et al., 2000; Ito et al., 2006). Cycloheximide is widely used in cell death research, but the detailed mechanism of cycloheximide-induced apoptosis remains unclear. Nonetheless, several possibilities have previously been considered. First, Alessenko et al. (1997) demonstrated that sublethal doses of cycloheximide caused short-term overexpression of c-myc, c-fos, and c-jun genes and long-term expression of p53. Because cycloheximide-resistant Molt-4 cells (Wu et al., 2004) do not express p53, it is tempting to speculate that cycloheximide-induced apoptosis is p53-dependent. However, expression of p53 was undetectable in the macrophages and SMC used in this study, even after stimulation with cycloheximide, and according to Higami et al. (2000), p53 seems not to be directly involved in cycloheximide-induced apoptosis. Second, it has been suggested that cycloheximide causes dysfunction of the endoplasmic reticulum because intravenous injection of cycloheximide in rats dramatically increases the mRNA levels of the endoplasmic reticulum stress genes ATF3 and CHOP (Ito et al., 2006), which have proapoptotic activity (Oyadomari and Mori, 2004). However, CHOP protein was not overexpressed in the present study after cycloheximide treatment.

Apart from the mechanism of cell death, our study raises several other interesting questions, such as: how is cycloheximide able to induce apoptosis in macrophages but not in SMCs or endothelial cells? De novo protein synthesis assays showed an equal inhibition of translation by cycloheximide in both macrophages and SMCs, which excludes differential uptake of the drug. Moreover, MAPK signaling during cycloheximide treatment was similar in both cell types. For example, we observed hyperphosphorylation of the stress-responsive p38 MAPK and SAPK/JNK as well as hyperphosphorylation of p70 S6 kinase and dephosphorylation of eIF2. The last two events are probably a final attempt of the cell to reactivate the translational machinery in response to external inhibition of protein synthesis. Because arterial macrophage-derived foam cells consume three times more oxygen than SMC (Bjornheden and Bondjers, 1987), it is conceivable that macrophages are metabolically highly active and thus more sensitive to protein synthesis inhibitors compared with SMCs. Moreover, inhibition of translation in SMCs induces a modulation toward a differentiated, quiescent, contractile phenotype (Martin et al., 2004), which renders the cells even more resistant to cell death mediated by inhibition of protein translation. Nonetheless, our data indicated that SMCs do not withstand long-term exposure to cycloheximide or treatment with high doses. Consequently, if a therapeutic strategy based on cycloheximide treatment is intended, a short-term and local intervention (e.g., drug-eluting stents with fast release) is required to clear macrophages in a plaque.

Although any reduction in macrophage numbers can improve plaque stability, it remains to be determined whether apoptosis is a suitable type of cell death to clear macrophages from atherosclerotic plaques. For many years, the apoptotic process has been considered immunologically silent. However, recent evidence suggests that apoptotic cells may also contribute to inflammatory processes via the release of pro-inflammatory cytokines and/or chemotactic factors (Schaub et al., 2000; Lauber et al., 2003). Moreover, inefficient removal of apoptotic cells may lead to induction of postapoptotic necrosis and inflammatory responses (Bjorkerud and Bjorkerud, 1996; Kockx and Knaapen, 2000; Scaffidi et al., 2002; Fink and Cookson, 2005). To prevent inflammation, cells signal their apoptotic state at an early stage to their environment, where they are recognized and engulfed by phagocytes (Savill and Fadok, 2000; Fadok et al., 2001). Hence, the consequences of macrophage apoptosis depend largely on the efficiency of apoptotic cell phagocytosis and, therefore, also on the atherosclerotic lesion state. Indeed, it has been proposed that macrophage apoptosis in early lesions has an overall beneficial effect on atherosclerosis, largely because of the ability of early plaque phagocytes to efficiently and safely clear dead macrophages (Tabas, 2005). However, in advanced atherosclerotic plaques, phagocytosis is defective (Schrijvers et al., 2005), allowing apoptotic macrophages to undergo postapoptotic necrosis, thereby increasing additional inflammation and/or thrombogenicity of the plaque. These phenomena may further promote plaque instability and increase the risk of acute atherothrombotic events (Tabas, 2005). In addition, rapid induction of massive macrophage apoptosis may not be desirable, even in early lesions, because rapid loss of macrophages would reduce scavenging of apoptotic cells. We therefore believe that selective clearance of macrophages in atherosclerotic plaques by cycloheximide treatment is only a first step toward plaque stabilization. After macrophage clearance, additional anti-inflammatory strategies may be needed to prevent secondary inflammatory responses, re-infiltration of macrophages, and short-term plaque-stabilizing effects. In conclusion, cycloheximide induces selective apoptotic cell death of macrophages within atherosclerotic plaques without a significant impact on SMCs or the endothelium.

	Acknowledgements

 
We are indebted to Hermine Fret, Jan Van Daele, Rita Van Den Bossche, Anne-Elise Van Hoydonck, and Ludo Zonnekeyn for excellent technical assistance.

	Footnotes

 
Research was supported by funding from the Fonds voor Wetenschappelijk Onderzoek (FWO) (projects G.0308.04 and G.0113.06), Special Research Fund of the University of Antwerp (Nieuwe Onderzoeksinitiatieven-Bijzonder Onderzoeksfonds) and the Bekales Foundation. Wim Martinet is a postdoctoral fellow of the Fund for Scientific Research-Flanders (Belgium, FWO).

Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.

doi:10.1124/jpet.106.113944.

ABBREVIATIONS: SMC, smooth muscle cell; mTOR, mammalian target of rapamycin; Erk, extracellular signal-regulated kinase; SAPK, stress-activated protein kinase; JNK, c-Jun NH₂-terminal kinase; MAPK, mitogen-activated protein kinase; FITC, fluorescein isothiocyanate; TUNEL, terminal deoxynucleotidyl transferase dUTP nick-end labeling; FITC-VAD-fmk, FITC-labeled Val-Ala-DL-Asp(O-methyl)-fluoromethylketone; CHOP, CCAAT/enhancer-binding protein homologous protein; eIF2, eukaryotic initiation factor 2; z-VAD-fmk, benzyloxycarbonyl-Val-Ala-DL-Asp(O-methyl)-fluoromethylketone; TNF, tumor necrosis factor-.

Address correspondence to: Dr. Guido R. Y. De Meyer, Division of Pharmacology, University of Antwerp, Universiteitsplein 1, B-2610 Wilrijk, Belgium. E-mail: guido.demeyer{at}ua.ac.be

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