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CELLULAR AND MOLECULAR

Delineation of Myotoxicity Induced by 3-Hydroxy-3-methylglutaryl CoA Reductase Inhibitors in Human Skeletal Muscle Cells

Center of Biomolecular Medicine and Pharmacology (J.S., M.W., C.S., M.H.), Department of Anaesthesiology and Intensive Care Medicine (L.W.), Medical University of Vienna, Vienna, Austria

Received March 21, 2005; accepted May 19, 2005.

	Abstract

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The 3-hydroxy-3-methyl-glutaryl-CoA reductase inhibitors (statins) are widely used and well tolerated cholesterol-lowering drugs. In rare cases, side effects occur in skeletal muscle, including myositis or even rhabdomyolysis. However, the molecular mechanisms are not well understood that lead to these muscle-specific side effects. Here, we show that statins cause apoptosis in differentiated human skeletal muscle cells. The prototypical representative of statins, simvastatin, triggered sustained intracellular Ca²⁺ transients, leading to calpain activation. Intracellular chelation of Ca²⁺ completely abrogated cell death. Moreover, ryanodine also completely prevented the simvastatin-induced calpain activation. Nevertheless, an activation of the ryanodine receptor by simvastatin could not be observed. Downstream of the calpain activation simvastatin led to a translocation of Bax to mitochondria in a caspase 8-independent manner. Consecutive activation of caspase 9 and 3 execute apoptotic cell death that was in part reversed by the coadministration of mevalonic acid. Conversely, the simvastatin-induced activation of calpain was not prevented by mevalonic acid. These data delineate the signaling cascade that leads to muscle injury caused by statins. Our observations also have implications for improving the safety of this important medication and explain to some extent why physical exercise aggravates skeletal muscle side effects.

Usually, statins are well tolerated (S4 group, 1994, 2000; Sacks et al., 1997; Bellosta et al., 2004). However, side effects may arise in skeletal muscle. These may range from transient increases in creatine kinase (CK), muscle pain, and cramps to myositis and potentially life-threatening rhabdomyolysis (Farmer and Torre-Amione, 2000; Bellosta et al., 2004; Rosenson, 2004). Physical exercise predisposes patients who take statins to develop muscle specific side effects (Thompson et al., 1997; Sinzinger and O'Grady, 2004). This is also true for some forms of comedication (Bellosta et al., 2004). The concomitant administration of inhibitors of cytochrome oxidase P450 3A4 (e.g., macrolide antibiotics, cyclosporin A, or azole antimycotics) puts patients at a higher risk (Farmer and Torre-Amione, 2000; Bellosta et al., 2004; Rosenson, 2004). The molecular mechanisms have not yet been identified that underlie statin-induced lysis of skeletal muscle (Thompson et al., 1997; Sinzinger and O'Grady, 2004). It is specifically not clear why skeletal muscle is so susceptible to toxicity of statins (Rosenson, 2004). Cardiac muscle, for example, is not adversely affected by statins.

To gain mechanistic insight, in the present work we have delineated the signaling pathways that are activated by statins in differentiated primary human skeletal muscle cells. Our experiments show that statins cause sustained increases in cytosolic Ca²⁺ levels, which are essentially linked to the development of apoptosis in differentiated human skeletal muscle cells.

	Materials and Methods

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Cell Culture. All experiments were carried out with differentiated primary human skeletal muscle cells obtained from skeletal muscle biopsies of healthy individuals who underwent diagnostic testings. The leftover material (100–400 mg) was used to isolate satellite cells. The procedure was approved by the local ethic committee of the Medical University of Vienna (Dr. Weigl; Department of Anaesthesiology, General Hospital, Vienna, Austria). Satellite cells were kept in growth medium (Ham's F-12, 15% fetal calf serum, 50 ng/500 ml epidermal growth factor, 10 mg/500 ml insulin, 200 µg/500 ml dexomethasone, 250 mg/500 ml fetuin and bovine serum albumin, 78 g/500 ml glucose, 200 mM L-glutamine, 5000 units/ml penicillin G, 5000 µg/ml streptomycin, and 250 µg/500 ml amphotericine B), and differentiation was initiated by switching to differentiating medium (Dulbecco's modified Eagle's medium supplemented with 5% horse serum, and 4 mM L-glutamine, 100 ng/ml insulin, and 0.1 µg/ml gentamicin) as described previously (Weigl et al., 2000).

Caspase Assays. Differentiated human skeletal muscle cells were maintained in the absence and presence of statins and inhibitors at concentrations and for incubation times indicated in the respective figures. Thereafter, the cells were washed with phosphate-buffered saline and lysed with ice-cold caspase-lysis buffer (25 mM HEPES, pH 7.4, 5 mM EDTA, 1 mM EGTA, 5 mM MgCl₂, and 5 mM dithiothreitol) supplemented with protease inhibitors (1.4 µg/ml aprotinin, 10 µg/ml leupeptin, and 100 µM pefablock). Microsomal fractions and cytosol were separated by centrifugation at 45,000g at 4°C for 20 min. The pellet was resuspended in caspase lysis buffer, and samples were stored at –80°C. All steps starting with cell lysis were carried out on ice. Aliquots of the supernatant (10–50 µg) were incubated in reaction buffer (25 mM HEPES, pH 7.4, 6.6% sucrose, 1.4 CHAPS, and 5 mM dithiothreitol) and 50 µM 7-amino-4-trifluoro-methylcoumarin (AFC)-conjugated substrate at 37°C for 90 min in the dark. The caspase-specific substrates for caspase were 3 Ac-Asp-Glu-Val-Asp-AFC (Ac-DEVD-AFC); for caspase 8, Ac-Leu-Glu-Thr-Asp-AFC (Ac-LETD-AFC); and for caspase 9, Ac-Leu-Glu-His-Asp-AFC (Ac-LEHD-AFC). The cleavage of the caspase substrates was measured at an excitation wavelength of 405 nm and an emission wavelength of 535 nm by a fluorescence plate reader (Wallac 1420 multilabel counter VICTOR-2; PerkinElmer Life and Analytical Sciences, Wellesley, MA). As a negative control, the AFC-conjugated substrates were diluted in lysis buffer and reaction buffer in the absence of protein.

Calpain Assay. Calpain activity was measured with a calpain kit obtained from Calbiochem (San Diego, CA). Purified calpain was used as a positive control. As a negative control, the reaction was carried out in the presence of calpain inhibitor I (N-acetyl-Leu-Leu-Nle-CHO), calpain inhibitor II (N-acetyl-Leu-Leu-Met-CHO), calpain inhibitor IV (Z-Leu-Leu-Tyr-CH₂F), or in the absence of protein. After incubation for 90 min at 37°C in the dark, cleavage of the fluorescent calpain substrate Ac-Leu-Leu-Tyr-AFC (Ac-LLY-AFC) was monitored with a fluorescence plate reader as already described for the caspase assays (i.e., excitation at 405 nm and emission at 535 nm).

Immunohistochemistry and Western Blot Analysis. The pellets and supernatants prepared from differentiated human skeletal muscle cells were used for Western blot analysis. Protein samples (5–15 µg) were diluted in sample buffer, heated for 2 min at 95°C, and resolved on SDS-polyacrylamide gels. The proteins were transferred to nitrocellulose membranes (pore size, 0.45 µm; Schleicher & Schuell, Dassel, Germany), and immunodetection was carried out with a polyclonal rabbit IgG antibody against Bax (N-20; diluted 1:250; Santa Cruz Biotechnology, Inc., Santa Cruz, CA), a monoclonal mouse antibody #17 against a synthetic peptide homologous to amino acids 313 to 330 of lamin-associated polypeptide 2 (LAP2) (1:500; a kind gift from Dr. Roland Foisner, Department of Biochemistry and Molecular Cell Biology, Vienna Biocenter, Veinna, Austria) and a monoclonal mouse IgG antibody against actin (AC-40) from Sigma Chemical Co. (St. Louis, MO) (1:500) as primary antibodies. Secondary antibodies (diluted 1:5000) conjugated to horseradish peroxidase allowed detection by an enhanced chemiluminescence detection system from Pierce Chemical (Rockford, IL). The intensity of the bands of interest was quantified with the Scion image software (www.scioncorp.com) and compared with the corresponding loading control (actin band). For immunohistochemistry, differentiated human skeletal muscle cells were fixed with 4% paraformaldehyde for 30 min. The cells were washed with 50 mM NH₄Cl and subsequently permeabilized for 5 min with a solution containing 0.1% Triton X-100 and 0.1% citrate in phosphate-buffered saline. The cells were blocked with 2% bovine serum albumin and exposed to Alexa 488-conjungated Mito-Tracker to visualize mitochondria. All the antibodies were diluted 1:200 and incubated for 1 h at 37°C. Immunostaining of cells was visualized with a goat anti-rabbit Cy3-conjugated (diluted 1:500; Amersham Biosciences, Inc., Vienna, Austria) or goat anti-mouse Alexa 488-conjugated secondary antibody (diluted 1:500; Molecular Probes, Leiden, The Netherlands). As a negative control, cells were treated under identical conditions without applying a first antibody. Images were collected using a confocal microscope from Carl Zeiss (Jena, Germany) equipped with an argon laser system (LSM 410). The digitized pictures were stored and analyzed off-line using MetaMorph software (Universal Imaging Corporation, Downingtown, PA).

Ca²⁺ Release Experiments. Differentiated human skeletal muscle cells were loaded with 10 µM fura-2/AM (Molecular Probes) in the presence of 0.025% Pluronic acid for 45 min and prepared for fluorescence photometry in suspension by collecting the cells in Tyrode's solution to achieve a concentration of 2.5 to 5 x 10⁶ cells/ml (Weigl et al., 2003). The fluorescence (excitation at 340 nm and 380 nm and the corresponding emission at 510 nm; 5-nm slits) was continuously recorded with a fluorescence photometer (F-4500; Hitachi, Tokyo, Japan). The ratio of the signals obtained at 340 and 380 nm was used to visualize a proportional value for the intracellular Ca²⁺ concentration and was done off-line.

High-Affinity [³H]Ryanodine Binding. Heavy sarcoplasmic reticulum membranes were prepared from rabbit hind leg and back skeletal muscle as described previously (Klinger et al., 1999). For high-affinity [³H]ryanodine binding, heavy sarcoplasmic reticulum membranes (50 µg) were incubated in 100 µl of binding buffer containing 40 mM HEPES·NaOH, pH 7.4, 200 mM KCl, 15 mM NaCl, 0.5 mM EGTA, 0.45 mM CaCl₂, 20 nM [³H]ryanodine, 1 µM aprotinin, 1 µM leupeptin, 100 µM pefablock, and the drug concentrations indicated in the figure legends. The samples were incubated for 120 min at 30°C. Nonspecific binding was determined in the presence of 100 µM ryanodine. The reaction was terminated by filtration over glass fiber filters (presoaked in 1% polyethylenimine) using a Skatron vacuum filtration device. The filters were rinsed with 10 ml of ice-cold 10 mM Tris-HCl, pH 7.4, and 500 mM NaCl, and the remaining radioactivity on the filters was determined by liquid scintillation counting.

Terminal Deoxynucleotidyl Transferase dUTP Nick-End Labeling (TUNEL) Assay. Apoptotic DNA fragmentation was visualized with an in situ cell death detection kit (Roche Diagnostics, Penzberg, Germany). As a negative control, mock incubations with only label solution containing fluorescent-deoxyuridine triphosphate nucleotides were carried out to measure background staining. The fluorescein labels incorporated in DNA were detected immediately or mounted and analyzed within 48 h. Fluorescence was visualized using a confocal argon laser microscope (LSM410; Carl Zeiss) with an excitation at 488 nm and an emission at 515 nm.

Miscellaneous Procedures. All experiments were carried out in triplicate, and each experiment was repeated at least two times. All data are presented as mean ± S.D., if not otherwise stated. The data of concentration-response curves were fitted by nonlinear least-squares regression to the Hill equation. Statistical significance (p < 0.05) was determined with Student's t test and for multiple comparison with ANOVA and post hoc Scheffé's test. The protein concentration was determined with the Bio-Rad Coomassie Blue kit (Bio Rad, Munich, Germany) or the bicinchoninic acid assay (Micro-BCA; Pierce Chemical) using bovine serum albumin as a protein standard.

	Results

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Simvastatin-Induced Apoptosis. Differentiated primary human skeletal muscle cells underwent apoptosis when exposed to simvastatin. Simvastatin (Fig. 1A), and to a similar extent lovastatin (data not shown), reduced the number of viable cells in a time- and concentration-dependent manner. The reduction in cell number was prevented in part by coadministration of mevalonic acid (Fig. 1B). Even a mevalonic acid concentration of 10 mM could not overcome the simvastatin-induced cell death. This indicates that a mechanism independent of the mevalonate pathway contributes to cell toxicity (Fig. 1B). This conclusion is based on the following observations.

View larger version (45K): [in this window] [in a new window]   Fig. 1. Simvastatin-induced apoptosis in differentiated human skeletal muscle cells. A, differentiated human skeletal muscle cells were treated with 1 (open triangle), 3 (filled circle), 30 (filled square), and 100 µM (open diamond) simvastatin or the carryover of 1% DMSO (open circle) for 24 h. Four phase contrast pictures were collected from each time point and sample treatment. The number of cells in four areas of equal size was counted in each picture, and the mean of the cell numbers was plotted against time. B, differentiated human skeletal muscle cells were treated for 24 h in the absence (CTL) and presence of 30 µM simvastatin (30S) or a combination of 30 µM simvastatin and 0.1, 1, or 10 mM mevalonic acid (30S+m.a.) or mevalonic acid alone at 0.1, 1, or 10 mM (0.1m.a., 1m.a., and 10m.a.). Trypan blue-stained cells were counted in four phase contrast pictures of equal size and subtracted from the total number of cells. The columns represent the mean ± S.D. C, TUNEL assay was carried out with differentiated human skeletal muscle cells that had been incubated in the absence (CTL) and presence of 30 µM simvastatin (30 µM S) for 24 h. Pictures were generated at a magnification of 10x for CTL and 20x for simvastatin (SIM) by phase contrast and confocal fluorescence microscopy. D, differentiated human muscle cells were incubated in the absence (CTL) and presence of 1 µM (1S) or 30 µM simvastatin (30S) for 24 h. LAP2 and actin were visualized in the cytosolic (10 µg; SUPERNATANT) and membrane (20 µg; PELLET) fractions. The intensity of the protein bands was quantified using the Scion Image software. The columns represent the mean ± S.E.M. Statistical significance for multiple comparison was calculated with ANOVA and post hoc Scheffé's test.

To address the mechanism by which simvastatin induces apoptosis, we analyzed the kinetics of caspase activation, by using selective fluorigenic substrates (Fig. 2). Caspase 3 is an effector caspase (Ferri and Kroemer, 2001). After 8 h of simvastatin exposure, caspase 3 activation was detectable (Fig. 2A) and sustained up to 30 h. Caspase 3 activation coincided with the sharp rise in caspase 9 activity (Fig. 2B), but the peak activation in caspase 9 activity occurred clearly before that of caspase 3. To confirm the conjecture that statins trigger apoptosis only via a caspase 9/caspase 3 cascade, we also measured caspase 8 activity. Within 36 h of simvastatin exposure, caspase 8 activity remained undetectable (Fig. 2C). Tumor necrosis factor- is known to trigger caspase 8 activity in myoblasts and was therefore used as a positive control (Stewart et al., 2004).

View larger version (45K): [in this window] [in a new window]   Fig. 2. Caspase and calpain activation by simvastatin in differentiated skeletal muscle cells. Differentiated human skeletal muscle cells were harvested at the indicated time points, and the cytosolic fraction (50 µg) was incubated with the 50 µM of the substrate for caspase 3 (A), caspase 9 (B), or caspase 8 (C) in caspase reaction buffer for 90 min at 37°C in the dark. The skeletal muscle cells were incubated in the absence (filled circle) and presence of 10 µM simvastatin (open triangles) or 10 µM simvastatin plus 1 mM mevalonic acid (filled square). After 36 h, cells treated with 0.1% DMSO (open diamond) were also prepared to control for the solvent carryover in the simvastatin sample. The filled triangle (B and C) shows the reaction buffer plus caspase substrate in the absence of cellular lysate. Caspase 8 activity was triggered with 10 ng/ml tumor necrosis factor- (filled diamond) (C). The data represent mean values ± S.E.M. obtained from two independent experiments in triplicates that were repeated twice. Differentiated human skeletal muscle cells were also exposed to increasing concentrations (1, 10, and 30 µM) of simvastatin (SIM) and lovastatin (LOV) for 24 h and compared with control (CTL). Caspase 3 (D), caspase 9 (E), and calpain (F) activity were measured in the cytosolic fraction (50 µg) as described above. The columns represent mean values ± S.E.M. obtained from three to 12 independent experiments carried out in duplicates. Statistical significance for multiple comparison was calculated with ANOVA and post hoc Scheffé's test (*, p < 0.05; **, p < 0.01; ***, p < 0.005; ****, p < 0.0001).

Simvastatin Triggers the Mitochondrial Pathway of Apoptosis. The role of calpain in apoptosis is a matter of debate; calpain was proposed to be both upstream and down-stream of the caspase cascade (for review, see Goll et al., 2003). We therefore focused first on the activation of the mitochondrial pathway of apoptosis. Simvastatin caused activation that was in magnitude comparable with the effect of staurosporine (Fig. 3B, right-hand column). If the proapoptotic action of simvastatin depends on the mitochondrial pathway, its effect ought to be abolished by an inhibitor of the mitochondrial transition pore. We verified this prediction by using bongkrekic acid, an inhibitor of the mitochondrial adenine transporter, which is an essential component of the mitochondrial transition pore (Gross et al., 1999). The addition of bongkrekic acid suppressed the activation of caspase 3 (Fig. 3A) and 9 (Fig. 3B) by 90 and 80%, respectively. In a similar manner, the simvastatin-induced activation of caspase 3 and 9 was abrogated by the simultaneous application of a cell-permeable pan-caspase inhibitor.

View larger version (37K): [in this window] [in a new window]   Fig. 3. Simvastatin-induced mitochondrial pathway of apoptosis. Differentiated human skeletal muscle cells were incubated for 24 h in the absence (CTL) or presence of 30 µM simvastatin (30 S); 4 µM bongkrekic acid (BA); 100 µM pan-caspase-inhibitor N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethyl ketone, Z-VAD-FMK (pCi); 1 mM mevalonic acid (m.a.); 30 µM simvastatin plus 4 µM bongkrekic acid (30S+BA); 30 µM simvastatin plus 100 µM Z-VAD-FMK (30S+pCi); and 30 µM simvastatin plus 1 mM mevalonic acid (30S+m.a.). Thereafter, the cytosolic fractions were assayed for caspase 3 (A) and caspase 9 (B) activity. The columns represent mean values ± S.E.M. obtained from two independent experiments carried out in duplicates, which were repeated with similar results. C, differentiated human skeletal muscle cells were incubated in the absence (CTL) and presence of 30 µM simvastatin (30S) or a combination of 30 µM simvastatin plus 1 mM mevalonic acid (30S+m.a.) for 24 h to perform Western blot analysis. The cytosolic (8 µg; SUPERNATANT) and membrane (12 µg; PELLET) fraction was analyzed for Bax and actin as a loading control. The intensity of the protein-bands was quantified and plotted as relative intensity (mean ± S.E.M.). D, similarly treated cells were subjected to immunohistochemistry at a magnification of 63x, using a confocal laser scanning microscope. The mitochondria were stained with Mito-Tracker (1:200; excitation 488 nm; emission 515), and Bax was visualized with a specific antibody. Statistical significance for multiple comparison was calculated with ANOVA and post hoc Scheffé's test.

Simvastatin-Triggered Ca²⁺ Release Activates Calpain. The proteolytic activity of calpain is activated by a long-lasting increase in the intracellular Ca²⁺ concentration (Goll et al., 2003). Therefore, we monitored the intracellular Ca²⁺ concentration in mass suspension. Under these conditions simvastatin triggered a sustained rise in the intracellular Ca²⁺ concentration (Fig. 4A). This amplitude of the Ca²⁺ response was comparable with that in the presence of extracellular EGTA, indicating that an intracellular Ca²⁺ release is triggered (Fig. 4B, open triangles). The simvastatin-triggered Ca²⁺ release did not return to basal resting Ca²⁺ concentrations but remained elevated. The amplitude of the Ca²⁺ transient was not attributable to an effect of the solvent, because dimethyl sulfoxide (DMSO) alone had no effect below 0.3%. Nevertheless, between 0.3 and 1.0% DMSO, a slight increase in the Ca²⁺ concentration was observed, which was subtracted from the Ca²⁺ release amplitudes at the respective simvastatin concentrations (Fig. 4B).

View larger version (34K): [in this window] [in a new window]   Fig. 4. Simvastatin-triggered Ca2+ release activates calpain. A, differentiated human skeletal muscle cells (2.5–5 x 105 cells) were resuspended in Tyrode's solution and loaded with 10 µM fura-2/AM. The simvastatin (SIM)-induced Ca2+ release is shown in a representative experiment. B, increasing concentrations of simvastatin were added to cell suspensions in Tyrode's solution in the absence (filled circles) and presence of 4 mM EGTA (open triangles), and the peak amplitude of the Ca2+ release is given. The data points represent the mean ± S.E.M. of two to six experiments. C, differentiated human skeletal muscle cells were incubated for 24 h in the absence (CTL) or presence of 30 µM simvastatin (30S), 4 µM bongkrekic acid (BA), 1 mM mevalonic acid (m.a.), or the combination of 30 µM simvastatin plus 4 µM bongkrekic acid (BA+S) and the combination of 30 µM simvastatin plus 1 mM mevalonic acid (m.a.+30S). The cytosolic fraction of the cells (20 µg) was used for calpain activity measurements. D, similar to C, cells were alternatively treated with 10 µM calpain inhibitor I (Ci), 30 µM simvastatin plus 10 µM calpain inhibitor I (30S+Ci), or 500 nM staurosporine plus 10 µM calpain inhibitor I (500St+Ci). Calpain activity was measured in the cytosolic fraction of the cell lysate (50 µg). The columns represent the mean ± S.E.M. obtained from three independent experiments in duplicates. Statistical significance for multiple comparison was calculated with ANOVA and post hoc Scheffé's test.

Chelation of the intracellular Ca²⁺ concentration with BAPTA-AM was indeed sufficient to prevent a reduction of cell number in the presence of simvastatin (Fig. 5A). In line with this finding, the simvastatin-induced activation of caspase 3, caspase 9, and calpain was abrogated by the co-administration of BAPTA-AM (Fig. 5, B–D). Interestingly, coadministration of ryanodine was also capable of preventing activation of the apoptotic-signaling cascade. However, it is unlikely that simvastatin directly activates the ryanodine receptor. The protective action of ryanodine is rather related to a block of the Ca²⁺-induced Ca²⁺ release that amplifies the simvastatin induced Ca²⁺ transient. We measured high-affinity [³H]ryanodine binding, which is proportional to the opening of the ryanodine receptor and that is sensitive to ryanodine receptor channel openers (Hohenegger et al., 1996; Klinger et al., 1999). We did not detect any significant alteration in [³H]ryanodine binding by simvastatin or lovastatin at concentrations up to 300 µM (Fig. 6). Conversely, caffeine, a ryanodine receptor agonist, gave a significant stimulation of [³H]ryanodine binding. The activation of caspase 3 and caspase 9 by simvastatin was also prevented by the calpain inhibitor I (Fig. 5, B and C) or by a combination of calpain inhibitor I and II (data not shown). Together, these data show that the simvastatin-evoked elevation of intracellular Ca²⁺ levels suffices to activate calpain, which is upstream of the mitochondrial pathway of apoptosis.

View larger version (32K): [in this window] [in a new window]   Fig. 5. Role of intracellular Ca2+ on simvastatin-induced apoptosis. A, differentiated human skeletal muscle cells were treated in the absence (CTL) and presence of 30 µM simvastatin (30S) or a combination of 30 µM simvastatin plus 1 µM BAPTA-AM (30S+BAPTA) for 24 h, and the number of cells was counted as described in Fig. 1. The columns show the mean ± S.E.M. normalized to control. Caspase 3 (B), caspase 9 (C), and calpain activities (D) were measured with cytosolic fractions from differentiated human skeletal muscle cells, incubated in the absence (CTL) and presence of 30 µM simvastatin (30S), a combination of 30 µM simvastatin and 1 µM BAPTA-AM (BAPTA+S), 30 µM ryanodine (Ry), a combination of 30 µM simvastatin and 30 µM ryanodine (Ry+S), 10 µM calpain inhibitor I (Ci), and a combination of 30 µM simvastatin and 10 µM calpain inhibitor I (Ci+S) for 24 h. The experiments were carried out in duplicates and repeated twice; data represent mean ± S.E.M. Statistical significance for multiple comparison was calculated with ANOVA and post hoc Scheffé's test.

View larger version (23K): [in this window] [in a new window]   Fig. 6. High-affinity [3H]ryanodine binding is not stimulated by statins. Basal high-affinity [3H]ryanodine binding (CTL) to sarcoplasmic reticulum membranes from rabbit skeletal muscle was stimulated with 10 mM caffeine (Caffeine) or increasing concentrations of simvastatin (open triangle), lovastatin (filled circle), or the corresponding carry over of the solvent DMSO (filled square). Error bars indicate the standard deviation of three experiments carried out in duplicates. Statistical significance was determined with Student's t test.

	Discussion

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Statins are well tolerated, cholesterol-lowering drugs. Because of the high prevalence of hypercholesterinemia and because of their demonstrated efficacy, statins became one of the most widely prescribed classes of medications. However, in rare cases, myopathy, myositis, or rhabdomyolysis occurs during therapy (Bellosta et al., 2004; Rosenson, 2004). The incidence of skeletal muscle side effects is significantly increased by elevated dose regimes or by the concomitant administration of drugs that block CYP3A4 and thereby the degradation of some statins. These circumstances have led to the withdrawal of cerivastatin from the international market in 2001 (Griffin, 2001). The incidence of myopathies may be also facilitated by preexisting conditions such as diabetes mellitus, hypothyroidism, surgery, trauma, or excessive alcohol intake (Bellosta et al., 2004; Rosenson, 2004). A genetic linkage of individuals susceptible for statin induced myopathies has not been investigated. Interestingly, physical exercise is also an aggravating factor for the development of muscle pain, myositis, and CK elevation (Sinzinger and O'Grady, 2004). However, CK elevation is an unreliable parameter for the development and severity of a myopathy (Dujovne, 2002; Phillips et al., 2002; Rosenson, 2004). Patients have been described who generate symptoms of severe muscle pain and restriction in mobility despite normal serum CK levels (Phillips et al., 2002; Bellosta et al., 2004).

Here, we present a signaling cascade in differentiated human skeletal muscle cells that is triggered by simvastatin-induced Ca²⁺ transients and is capable of explaining the phenomena described above. This evidence is based on the following observations. 1) Simvastatin and lovastatin induced apoptosis in human skeletal muscle cells in a concentration- and time-dependent manner (Figs. 1 and 2). Whereas a high simvastatin concentration of 10 µM triggered the mitochondrial pathway within 24 h, similar effects were seen with lower concentrations only after 36 or 48 h (data not shown). 2) The statin-induced reduction in cell number is in part abrogated by mevalonic acid. The fact that only 30 to 50% of the reduction in viable cells was due to inhibition of HMG-CoA reductase inhibition is indicative of an additional, mevalonate-independent mechanism underlying simvastatin induced cell death. 3) The application of simvastatin to fura-2-loaded skeletal muscle cells initiated a long-lasting Ca²⁺ transient that led to activation of calpain. 4) Abrogation of the intracellular Ca²⁺ elevation was sufficient to prevent activation of calpain and caspases 9 and 3. Moreover, the chelation of intracellular Ca²⁺ completely rescued skeletal muscle cells from simvastatin-induced cell death. A rough estimate of the EC₅₀ value for simvastatin and lovastatin triggered caspase activity is in the range of 1 to 5 µM (Fig. 2, D and C). An earlier observation in neonatal rat skeletal myocytes shows that cell viability, monitored by mitochondrial dehydrogenase activity, is inhibited half-maximally at 21.2 µM simvastatin and 20.2 µM lovastatin, whereas the IC₅₀ for simvastatin and lovastatin to inhibit the HMG-CoA reductase was between 0.09 and 0.12 µM (Masters et al., 1995). Although we have not directly determined the reduction of the endogenous cholesterol synthesis in differentiated human skeletal muscle cells, mevalonic acid was capable of reversing activation of caspase 3 and 9. Interestingly, mevalonic acid prevented the mitochondrial insertion of Bax but not its redistribution into the cytosolic compartment (Fig. 3C). This may be indicative for Bax targeting to another membrane system. Recent experimental evidence accumulates that Bax may insert into the endoplasmic reticulum, resulting in depletion of this Ca²⁺ store with consecutive caspase activation (Zong et al., 2003; Oakes et al., 2005).

Statin-induced apoptosis via the activation of calpain is clearly in line with previous observations that the Ca²⁺-dependent protease calpain may trigger apoptosis (Goll et al., 2003). In the case of differentiated human skeletal muscle cells, calpain is clearly upstream of the mitochondrial pathway. A cell-permeable calpain inhibitor was able to overcome the activation of calpain and the mitochondrial pathway of apoptosis (Fig. 5). The chelation of intracellular Ca²⁺ was also sufficient to prevent the simvastatin-induced calpain activation as well as activation of the caspases 9 and 3. Strikingly, such a treatment kept the number of viable cells identical to that of the control (Fig. 5). Conversely, coadministration of mevalonic acid was not capable of abrogating simvastatin-induced reduction in cell numbers (Fig. 1B) and calpain activation. A possible target of such a mevalonate-independent trigger mechanism of apoptosis is via the Ca²⁺ release channel of the sarcoplasmic reticulum, the ryanodine receptor. This interpretation is strengthened by the finding that in the absence of extracellular Ca²⁺, the amplitude of the simvastatin induced Ca²⁺ release is comparable with that in the presence of extracellular Ca²⁺ (Fig. 4B). Moreover, the plant alkaloid ryanodine could block the simvastatin-induced activation of calpain. Nevertheless, a direct activation of the ryanodine receptor by simvastatin is unlikely for the following reasons. 1) High-affinity [³H]ryanodine binding to sarcoplasmic reticulum fractions revealed no activation of the ryanodine receptor by simvastatin (Fig. 6). Possibly, simvastatin may bind to a cytoplasmic target protein that then may activate the ryanodine receptor. In such a case, an activation in the [³H]ryanodine binding would not be seen because we used sarcoplasmic reticulum membranes. 2) It also has to be mentioned that due to limited human cell material, the above-described binding studies have been done with rabbit skeletal muscle. However, it cannot be excluded that species differences may account for the absence of a statin effect on the [³H]ryanodine binding. 3) Elevation of the cytoplasmic Ca²⁺ concentration could be triggered by a statin-induced inhibition of the Ca²⁺ ATPase of the sarcoplasmic reticulum. Such a scenario, however, does not explain the protective feature of ryanodine to prevent calpain and caspase activation by simvastatin (Fig. 5). It is therefore clear that intensive effort has to be undertaken to address the exact target of simvastatin induced Ca²⁺ release. Nakahara and colleagues investigated in L6 rat myoblasts simvastatin induced Ca²⁺ transients. A target for the described Ca²⁺ fluxes has not been elucidated. Besides an intracellular Ca²⁺ release mechanism, they also described a Ca²⁺ influx, which together may contribute to cellular damage (Nakahara et al., 1994). In human skeletal muscle cells, a Ca²⁺ influx plays only a minor role, because we readily can detect a simvastatin-induced Ca²⁺ release in the presence of extracellular EGTA (Fig. 4B). The amplitudes of the Ca²⁺ release in EGTA-treated cells were similar to that obtained in the presence of extracellular Ca²⁺.

Statins increase the incidence of myopathy and aggravate their clinical signs in the presence of physical exercise (Thompson et al., 1997; Sinzinger and O'Grady, 2004). Repetitive skeletal muscle stimulation elevates the intracellular Ca²⁺ concentration. In the presence of statins, this physiological regulation of the skeletal muscle may facilitate muscular side effects, even in well trained individuals (Sinzinger and O'Grady, 2004). Under pathological conditions, in 11 patients with exertional rhabdomyolysis, the intracellular Ca²⁺ concentration in skeletal muscle biopsies was directly measured and found to be elevated up to 1.2 µM compared with 0.12 µM in control individuals (Lopez et al., 1995). Importantly, the clinical symptoms of rhabdomyolysis in these individuals were reversed with dantrolen, an inhibitor of the ryanodine receptor. Our observation, that ryanodine was capable to prevent calpain and caspase activation, corroborates the conjecture that simvastatin-induced Ca²⁺ transients may be amplified by a Ca²⁺-induced Ca²⁺ release mechanism via the ryanodine receptor (Fig. 4). Obviously, this mechanism was sufficient to keep the intracellular Ca²⁺ concentration at elevated levels to support the proteolytic activity of calpain (Fig. 5). In human Duchenne muscular dystrophy, elevated intracellular Ca²⁺ concentrations have been found to be responsible for increased proteolytic calpain activity (Alderton and Steinhardt, 2000). Calpain was shown to cleave caspases but mainly into inactive fragments. Thus, calpain thereby exerts an antiapoptotic action (Chua et al., 2000; Goll et al., 2003). Conversely, in a cerebral hypoxiaischemia model Ca²⁺-mediated activation of calpain triggered caspase 3-induced apoptosis (Blomgren et al., 2001). Similar proapoptotic mechanisms were found in other cell systems, initiated by a calpain activation (for review, see Goll et al., 2003).

The therapeutic administration of 40 mg of simvastatin per day has been summarized to give peak plasma levels of 10 to 34 ng/ml (25–80 nM) (Bellosta et al., 2004). Due to CYP3A4 inhibition, these plasma levels may increase by a factor of 10 to 15 and may reach at maximum 1.2 µM simvastatin. Interestingly, in a phase I dose finding study lovastatin was administrated to give plasma concentrations of up to 3.9 µM to treat cancer patients (Thibault et al., 1996). The authors report that myopathy was the dose-limiting factor. Interestingly, the reduction of ubiquinone was transient and not dose-dependent, but substitution with ubiquinone reduced and rescued the individuals from muscular side effects. The elevated statin concentrations described here are close to the concentration range where we observed half-maximum caspase and calpain activation. Moreover, caspase 3, caspase 9, and calpain were already activated by 1 µM simvastatin significantly; thus, underlining the clinical relevance of this study (Fig. 2, D–F).

	Acknowledgements

 
We thank Drs. Michael Freissmuth and Laura Malaga-Dieguez for helpful discussion, Dr. Roland Foisner for the generous gift of the LAP2 antibody, and A. Karel for technical assistance.

	Footnotes

 
This work was supported by Austrian Science Foundation Grant FWF P15973 [SwissProt] and Herzfelder'sche Familienstiftung (to M.H.).

doi:10.1124/jpet.105.086462.

ABBREVIATIONS: HMG, 3-hydroxy-3-methylglutaryl coenzyme A reductase; CK, creatine kinase; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]propanesulfonate; AFC, 7-amino-4-trifluoro-methylcoumarin; LAP2, lamin-associated polypeptide 2, heavy sarcoplasmic reticulum; TUNEL, terminal deoxynucleotidyl transferase dUTP nick-end labeling; ANOVA, analysis of variance; DMSO, dimethyl sulfoxide; BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-acetoxymethyl ester.

Address correspondence to: Dr. Martin Hohenegger, Center of Biomolecular Medicine and Pharmacology, Medical University of Vienna, Waehringerstrasse 13A, 1090 Vienna, Austria. E-mail: martin.hohenegger{at}meduniwien.ac.at

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