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Simvastatin induces impairment in skeletal muscle while heart is protected
BBRC Biochemical and Biophysical Research Communications 338 (2005) 1426–1434 www.elsevier.com/locate/ybbrc
Simvastatin induces impairment in skeletal muscle while heart is protected Pascal Sirvent a,b, Sylvain Bordenave b, Marianne Vermaelen b, Belle Roels c, Guy Vassort a, Jacques Mercier b, Eric Raynaud b, Alain Lacampagne a,* a
c
INSERM U637 CHU A. de Villeneuve, 34295 Montpellier, France b EA 701, Universite´ Montpellier I, 34000 Montpellier, France UPRES EA 3759 Faculty of Sport Sciences, 700 avenue Pic St Loup, 34090 Montpellier, France Received 17 October 2005 Available online 26 October 2005
Abstract 3-Hydroxy-3-methylglutaryl-coenzyme A reductase inhibitors (statins) are widely used to reduce plasma cholesterol concentration. However, statins are also known to induce various forms of muscular toxicity. We have previously shown that acute application of simvastatin on human skeletal muscle samples induced a cascade of cellular events originating from mitochondria and resulting in a global alteration of Ca2+ homeostasis. The present study was designed to further define the origin of the mitochondria impairment and to understand the apparent lack of deleterious effect on the heart. Using fluorescence imaging analysis and oxygraphy on human and rat skinned skeletal muscle samples, we show that the simvastatin-induced mitochondria impairment results from inhibition of the complex I of respiratory chain. Similar simvastatin-induced mitochondria impairment and alteration of Ca2+ homeostasis occur in permeabilized but not in intact ventricular rat cardiomyocytes. In intact rat skeletal muscle fibers from the flexor digitorum brevis muscle, the simvastatin-induced alteration of Ca2+ homeostasis is abolished when monocarboxylate transporter (MCT4) is inhibited. The impairment of complex I by simvastatin might be the primary step of its cellular deleterious effects leading to muscle fiber death. This mechanism is seen specifically in skeletal muscles. This specificity should be in part attributed to a preferential uptake of statins by MCT4 that is not expressed in cardiomyocytes. 2005 Elsevier Inc. All rights reserved. Keywords: Human skeletal muscle; Rat cardiomyocytes; Calcium; Mitochondria; Myotoxicity; Hypercholesterolemia; Satins; Monocarboxylate transporter
Hypercholesterolemia is a well-known risk factor for coronary heart disease. Among various drugs developed to reduce plasma cholesterol concentration, 3-hydroxy-3methylglutaryl-coenzyme A (HMG-CoA) reductase inhibitors (statins) are likely to be the most promising drug [1,2]. Statins decrease cholesterol content by selective inhibition of HMG-CoA reductase, rate-limiting step in the mevalonate synthesis pathway [3]. Clinical trials have demonstrated a reduction in cardiovascular-related morbidity and mortality in treated patients with or without coronary artery disease [4–6]. In addition to the benefits of lowering cholesterol, it has been shown that statins present choles*
Corresponding author. Fax: +33 467 415 242. E-mail address: [email protected] (A. Lacampagne).
0006-291X/$ - see front matter 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2005.10.108
terol-independent effects allowing for improved endothelial function, enhanced stability of atherosclerotic plaques, decreased inflammation and oxidative stress, and inhibited thrombogenic response in the vascular wall [7–10]. Adverse reactions involving skeletal muscles (myalgias, cramps, weakness, and exercise intolerance) are the most frequent, occurring in 1–7% of statin-treated patients [11]. These adverse effects can occur with or without increase in creatine kinase (CK) level. Some rare cases of fatal rhabdomyolysis have also been reported, with comedication or overdose [11,12]. If these adverse effects do not call into question efficiency and global safety of statin treatments, they can disturb treated patients in their everyday life. Taking into account the great number of statintreated patients, which would increase again in the next
P. Sirvent et al. / Biochemical and Biophysical Research Communications 338 (2005) 1426–1434
few years, comprehension of mechanisms of statin-induced myotoxicity seems to be determinant in order to prevent this adverse effect. We have recently shown that acute applications of simvastatin on human skeletal muscle samples trigger a large release of Ca2+ from the SR, resulting from early alterations of mitochondrial function [13]. The present study was designed to further define the origin of the mitochondrial impairment as well as to understand the apparent lack of deleterious effect on heart. Here, we demonstrate on human and rat muscle samples that: (1) the simvastatin-induced Ca2+ release does not affect cardiac cells and is specific of skeletal muscle whatever muscle phenotype, (2) the simvastatin-induced Ca2+ homeostasis impairment is predominantly dependent on statin uptake through monocarboxylate-transporter (MCT4), (3) a specific alteration of complex I of the respiratory chain appears to be the initial step of a cascade of deleterious cellular effects leading to Ca2+ homeostasis alteration. Materials and methods Human skeletal muscle samples. Muscle samples are prepared as previously described [13]. Fifteen healthy voluntary men aged between 25 and 45 were recruited in order to take part in the study. They did not take any medication known to potentially interfere with the protocol. The study was approved by the Local Ethics Committee and conformed to the Declaration of Helsinki regarding the use of human subject and informed consent was obtained from all the subjects after explanation of the nature of the study and potential risks. Muscle biopsies were performed in the inferior third level of the vastus lateralis muscle under local anesthesia with a Bergstro¨m needle. For imaging experiments, biopsies (40 mg) were immediately placed in the internal-like medium (in mol/L: K-glutamate 140, Hepes 10, phosphocreatine 20, Na2ATP 5, MgCl2 4.53, EGTA 1, CaCl2 0.29, and malate 2, pH 7.0). For in situ mitochondrial respiration studies, biopsies were placed in an ice-cold relaxing solution (in mM: EGTA–calcium buffer 10 (free Ca2+ concentration: 100 nM), imidazole 20, KH2PO4 3, MgCl2 1, taurine 20, DTT 0.5, MgATP 5, and phosphocreatine 15, pH 7.1). Rat muscle and heart samples. Male Wistar rats weighting 300 g (Janvier; Le Genest-St-Isle, France) were used. This investigation conforms to the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 85-23, Revised 1996) and European directives (96/ 609/EEC) were used. EDL and soleus muscle were dissected and kept in an external medium containing (in mM): NaCl 145, KCl 4, CaCl2 1.8, MgSO4 1, Hepes 10, and glucose 10, pH 7.4. For imaging experiments, muscles were then set in the dissecting chamber in a solution containing (in mM): K-glutamate 140, Hepes 10, MgCl2 10, and EGTA 0.1, pH 7.0. For in situ mitochondrial respiration studies, samples were placed in the same ice-cold relaxing solution as for human biopsies. Flexor digitorum brevis (FDB) muscles were dissected and enzymatically dissociated with 3 mg/ml collagenase (type 1; Sigma) dissolved in an external medium (see above) at 37 C for 1 h. Bundles of fibers were then transferred to an external medium without collagenase, and fibers were separated mechanically by gentle trituration. Intact single fibers were plated on extracellular matrix (Sigma)-coated coverslips attached across a 12-mm hole in the bottom of 35-mm petri dishes. Cardiac ventricular cells were enzymatically isolated as described earlier [14]. Briefly, rats were anesthetized by intraperitoneal injection of pentobarbital (200 mg/100 g). The heart was perfused retrogradely with a Ca2+-free Hanks-Hepes-buffered solution (in mM: NaCl 117, KCl 5.7, NaHCO3 4.4, KH2PO4 1.5, MgCl2 1.7, Hepes 21, glucose 11, and taurine 20, pH 7.2, adjusted with NaOH and bubbled with 100% O2) for 5 min at 37 C and then with an
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enzyme-containing solution (same as above with 1.0 mg/ml collagenase type IV (Worthington, Freehold, NJ, USA) and 20 lM Ca2+) for 25– 30 min. The ventricles were then gently dissociated with forceps in the same medium without enzyme. The cells were then filtered and further washed several times in a solution containing 0.3 mM Ca2+. Finally, myocytes were kept in a medium containing 1 mM Ca2+ and 0.5% bovine serum albumin (BSA). Cardiomyocytes were permeabilized as described by others [15]. Cardiomyocytes were placed in an internal medium (containing in mM: 120 potassium aspartate, 3 MgATP, 2 EGTA, 0.11 CaCl2 (free [Ca2+] = 100 nM), 10 phosphocreatine, and 5% dextran (40,000); pH 7.2.) with saponin (0.01% for 45–60 s). Experiments were performed with 50 lM Fluo-3 potassium salt (TefLabs, Austin, TX, USA) in the internal medium. Measurements of cytoplasmic Ca2+. Bundles of 2–5 fibers were manually dissected under a dissecting microscope (Stemi 200 Zeiss, France). Fibers were mounted in an experimental chamber containing an internal medium and permeabilized by adding saponin 0.01%, 30 s at room temperature. Fibers were slightly stretched to a sarcomere length around 2.8–3.2 lm to reduce movement artifacts. Fibers were then bathed in an internal medium containing the fluorescent Ca2+ indicator Fluo-3 (pentapotassium salt; TefLabs, Austin, TX, USA; 50 lM). FDB fibers and isolated cardiomyocytes bathed in an external medium, were loaded during 30 min at room temperature with Fluo-3 AM (5 lM, TefLabs, Austin, TX, USA). Variations in intracellular Ca2+ were measured by time-series (1 image/5 s) of x–y confocal fluorescent images (Zeiss LSM 510 Meta, 63· objective, NA = 1.2, H2O immersion). Fluo-3 was excited with an argon/krypton laser at 488 nm, and emitted fluorescence was recorded at 525 nm. Mean fluorescence was calculated in regions of interest (see Fig. 1, white box) and reported as a function of time. Measurements of mitochondrial Ca2+. To assess mitochondrial Ca2+, fibers were incubated for 30 min at room temperature with Rhod-2 AM (5 lM) in the internal medium. Rhod-2 is taken up preferentially into polarized mitochondria. After washes (3 times 5 min in internal medium) to remove cytosolic Rhod-2, variations in mitochondrial Ca2+ were recorded by time-series (1 image/5 s) of x–y confocal images. Fluorescence intensity was recorded at 570 nm, with an excitation at 543 nm provided by a helium/neon laser. Measurements of mitochondrial membrane potential. Mitochondrial membranes were labeled with the rhodamine-based fluorescent cationic lipophilic tetramethylrhodamine ethyl ester (TMRE, 100 nM), which distributes across the inner mitochondrial membrane in accordance with the Nernst equation. After washes (3 times 5 min in internal medium), variations in mitochondrial membrane potential were recorded by timeseries (1 image/5 s) of x–y confocal images. Fluorescence intensity was recorded above 560 nm, with an excitation at 543 nm provided by a helium/neon laser. Mean fluorescence was calculated in small regions of interest corresponding to mitochondrial staining and reported as a function of time. Measurements of mitochondrial respiration. Respiratory parameters of the total mitochondrial population were studied in situ on fresh skeletal muscle fibers as previously described [16]. Bundles of muscle fibers were manually isolated and saponin-skinned (50 lg/ml saponin for 30 min at 4 C). Respiration rates were determined at 27 C with a Clark electrode (Strathkelvin, Glasgow, Scotland) in an oxygraphic cell containing respiration solution (same composition as the relaxing solution, except that MgATP and phosphocreatine were replaced by 3 mM phosphate and 2 mM fatty acid-free bovine serum albumin). Respiration rates were expressed in micromoles of O2 per minute per gram of dry weight. Basal oxygen consumption without ADP (V0), maximal respiration rate (Vmax) with a saturating concentration of ADP and acceptor control ratio (ACR), and the ratio between Vmax and V0, which represents the level of coupling between oxidation and phosphorylation, have been measured. At the end of each measurement, cytochrome c test used to investigate the outer mitochondrial membrane integrity [17]. Respiration rates were recorded in the presence of glutamate (5 mM)/malate (2 mM) or pyruvate (10 mM)/malate (2 mM) or palmitoyl-carnitine (40 lM)/malate (2 mM). Basal oxygen consumption without ADP (V0) was first
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Fig. 1. Simvastatin triggers mitochondrial Ca2+ release. Fibers were loaded with Rhod-2 AM to monitor mitochondrial [Ca2+]. (A) Acute applications of simvastatin (50 lM) resulted in a dose-dependent decrease in fluorescence indicating a mitochondrial Ca2+ efflux. (B) In fibers pretreated with both cyclosporin A and clonazepam, simvastatin did not induce Ca2+ release. (C) Dose-dependent effects of simvastatin on mitochondrial Rhod-2 fluorescence. (*p < 0.05 compare to control values.)
recorded, and Vmax was determined in the presence of 2 mM ADP. Increasing concentrations of simvastatin diluted were then added. For calculation of the ACR, V0 was recorded with 50 lM final concentration of simvastatin; 2 mM ADP was then added for maximal activation of respiration. The effects of simvastatin on respiratory chain complexes were investigated at maximal respiration rates (ADP 2 mM) and for 50 lM of simvastatin. The specific substrates and inhibitors used to study the effects of simvastatin on the respiratory chain complexes are summarized in Fig. 5. Drugs and chemical reagents. All drugs used were purchased from Sigma, except for Fluo-3 and Rhod-2 (Teflab) and TMRE (Molecular Probes). Simvastatin was a generous gift from Merck laboratories. The provided lactone form of simvastatin was converted to the acidic pharmacologically active form as described by others [18]. Since simvastatin is diluted in ethanol, a series of control experiments containing the same amount of solvent was performed to exclude a potential effect of solvent. Statistical analysis. Statistical comparisons were done by a variance analysis (ANOVA). Data are expressed as means ± SEM and difference was considered significant with p < 0.05.
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Bundles of human muscle fibers were loaded with Rhod2 AM to monitor mitochondrial Ca2+ (Fig. 1). Acute application of simvastatin (3–100 lM) resulted in a dosedependent release of Ca2+ by mitochondria (Figs. 1A and C) that reached 59 ± 5% (n = 7) depletion at maximum (10 lM). Efflux of Ca2+ from mitochondria can be carried out either by the permeability transient pore (PTP) or the Na+–Ca2+ exchanger (NCE), respectively, blocked by cyclosporine A and clonazepam. Pre-treatment of fibers with 0.2 lM clonazepam or 0.5 lM cyclosporine A did not block the statin-induced Ca2+ release by mitochondria.
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Fig. 2. Simvastatin induces a fall in mitochondrial membrane potential. (A) Acute application of simvastatin (50 lM) on permeabilized muscle fibers loaded with TMRE resulted in a dose-dependent decrease in fluorescence compared to control conditions. (B) Pre-treatment of fibers with cyclosporin A and clonazepam did not prevent mitochondrial membrane potential fall. (C) Dose-dependent effects of simvastatin on mitochondrial TMRE fluorescence. (*p < 0.05 compare to control values.)
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deshydrogenase complex, Krebs cycle, and respiratory chain. With both substrates, maximal ADP-stimulated respiration was inhibited by 20% after applying 50 lM simvastatin (Figs. 4A–D). We used FCCP to uncouple oxidative phosphorylation, using glutamate/malate as substrate (Figs. 5C and D). In this uncoupled condition, simvastatin induced a stronger inhibition of respiration rate (38%). This difference in percentage of inhibition is explained by the difference in maximal rate of respiration known to be slower in coupled than in uncoupled oxidative phosphorylation. These observations on uncoupled respiratory chain render the hypothesis of an uncoupling effect of simvastatin very unlikely. Taken together, these data strongly suggest an effect of simvastatin upstream to the ATP synthase, i.e., one or more of the complexes I–IV of the respiratory chain of the mitochondria.
However, pre-treatment with both drugs simultaneously completely abolished this phenomenon (Figs. 1B and C). Changes in mitochondrial membrane potential were then assessed on bundles of human muscle fibers loaded with TMRE (Fig. 2). Acute applications of simvastatin (1–100 lM) resulted in rapid depolarizations (47 ± 1% at maximum, n = 6) of mitochondrial membrane that were dose-dependent (Figs. 2A and C). Pre-treatment with both 0.2 lM clonazepam and 0.5 lM cyclosporine A did not prevent this phenomenon (Figs. 2B and C). This result implies that the fall in mitochondrial membrane potential is independent of NCE and PTP opening. It might result from an upstream step such as an alteration of the respiratory chain electron flux, which allows the formation of the electrochemical gradient through the inner membrane. Simvastatin alters mitochondrial respiration
Simvastatin affects mainly complex I of respiratory chain Mitochondrial respiration was assessed on bundles of saponin-skinned human skeletal muscle fibers. Acute application of simvastatin resulted in a dose-dependent inhibition of maximal ADP-stimulated rate of oxygen consumption (Figs. 3A and B). This effect was statistically significant at 10 lM, and independent of muscle phenotype since simvastatin similarly altered maximal ADP-stimulated respiration rate on both soleus and EDL muscle fibers (Figs. 3C and D). We then evaluated mitochondrial respiration with different substrates to determine the origin of simvastatin-induced impairment. We used palmitoyl-carnitine/malate as substrate to assess lipid pathway, testing b-oxidation, Krebs cycle, and respiratory chain. For glucidic pathway, we used pyruvate/malate as substrate, testing pyruvate A
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We used specific substrates and inhibitors to sequentially evaluate the effect of simvastatin on the different complexes of respiratory chain (Figs. 5A and B). Glutamate/ malate as substrates allow to activate complexes I, III, and IV of respiratory chain. The use of succinate and rotenone would restrict the activity of the respiratory chain to complexes II, III, and IV. Finally, the use of TMPD (N,N,N 0 ,N 0 -tetramethyl-p-phenylenediamine) and ascorbate should test specifically the activity of complex IV. Acute applications of simvastatin induced a strong inhibition of maximal ADP-stimulated respiration rate with glutamate/malate as substrate. No significant inhibition could be seen using succinate as substrate, and only a very weak inhibition with TMPD and ascorbate (Fig. 5C). Complex C
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Fig. 3. Simvastatin inhibits mitochondrial respiration rate. (A) Mitochondrial respiration was estimated with acute applications of various concentrations of simvastatin on permeabilized bundles of human skeletal muscle fibers using pyruvate/malate as substrate. (B) Dose-dependent decrease in the maximal ADP-stimulated respiration rate induced by simvastatin. (C) Mitochondrial respiration recorded on permeabilized rat Soleus (left) and EDL (right) muscle fibers. (D) In both muscle types, acute applications of 10 lM simvastatin resulted in a decrease of maximal ADP-stimulated respiration rate to the same extent. (n = 8; *p < 0.05 compare to Vmax.)
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Fig. 4. Simvastatin alters glucidic and lipidic mitochondrial metabolic pathway. (A,B) Mitochondrial respiration was recorded on permeabilized human skeletal muscle fibers in the presence of palmitoyl-carnitine or pyruvate as main substrate. (C) Acute applications of simvastatin (50 lM) resulted in a 20% decrease in maximal ADP-stimulated respiration rate with both substrates. FCCP has been used to decouple mitochondrial respiratory chain. Acute application of simvastatin (50 lM) on decoupled mitochondria induced a strong decrease in respiration rate. (D) Changes in the rate of O2 consumption ( VO2), in the presence of simvastatin relative to the rates measured in respective control conditions. (n = 8; *p < 0.05 compare to control values.)
IV is not a limiting step in the respiratory chain. In normal conditions, when global respiration is activated, it functions at 50% of its maximal activity reached with TMPD/ascorbate, accounting for the discrepancy of observations between succinate/rotenone and TMPD/ascorbate. Indeed, the slight decrease observed in succinate/rotenone is likely to be only due to the inhibition of complex IV that is only partially activated in this condition. These results demonstrate that simvastatin effect is mainly due to an inhibition of complex I of respiratory chain. Effects of simvastatin on cytosolic and mitochondrial Ca2+: does simvastatin alter Ca2+ homeostasis in cardiomyocytes? Secondary to mitochondrial impairment, the acute application of simvastatin on skinned human skeletal muscle fibers resulted in a delayed large increase in [Ca2+]i (250 ± 25%, n = 36, Fig. 6A) as previously observed [13]. Vastus lateralis is known to be composed of both oxidative (slow twitch) and glycolytic (fast twitch) muscle fibers. In order to assess the effects of simvastatin on muscle fibers with different phenotypes, we used permeabilized cut myofibers manually dissected from rat EDL (fast) and soleus (slow) muscles. Acute application of simvastatin resulted in a delayed large transient Ca2+ release in both situations similarly (Figs. 6B and C). Acute application of simvastatin was also tested on rat ventricular cardiomyocytes to verify whether the adverse effect of simvastatin was generalized to all types of striated muscles. When applied on intact cardiac cells, simvastatin used at 200 lM did not trigger a large increase in intracellular Ca2+ similar to that seen in skeletal muscle,
even after more than 1 h of exposure (Fig. 7A). This implies that simvastatin does not easily diffuse into the cell or that the cascade of intracellular events triggered by simvastatin in skeletal muscle does not occur in cardiomyocytes. To answer this point, acute applications of simvastatin were performed on saponin-skinned cardiomyocytes (Fig. 7B). In these conditions, acute application of 200 lM simvastatin resulted in a fast increase in resting level of Ca2+ followed after 7–8 min by a larger increase in cytoplasmic [Ca2+] to some extent comparable to that observed on permeabilized skeletal muscle fibers. In some experiments, saponin-skinned cardiomyocytes were also loaded with TMRE to assess mitochondrial membrane potential. Acute application of simvastatin triggered a depolarization of mitochondrial membrane correlated with the first increase in [Ca2+]i, suggesting that this increase in resting Ca2+, as previously demonstrated in skeletal muscle [13], resulted from a mitochondrial Ca2+ efflux (not shown). This phenomenon was absent in intact cardiomyocytes. Human muscle biopsies do not allow us to obtain intact adult muscle fibers. To evaluate the effects of acute applications of simvastatin on intact skeletal muscle cells, we used intact rat flexor digitorum brevis (FDB) fibers dissociated enzymatically. In contrast with cardiomyocytes, simvastatin induced in intact muscle fibers a large increase in [Ca2+]i although with a prolonged delay and preceded by a slow increase in basal [Ca2+]i (Fig. 7C). Monocarboxylate transporter (MCT) and particularly the type 4 (MCT4) has been shown to mediate uptake of statins in rat mesanglial cells [19] and bovine kidney NBL-1 cells [19]. Interestingly, while MCT4 is largely expressed in skeletal muscle, it is
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Fig. 5. Simvastatin-induced mitochondrial respiratory chain impairment. (A) Schematic representation of complexes of mitochondrial respiratory chain. Different substrates and inhibitors have been used to isolate and study independently the different complexes as indicated. (B) Complexes I + III + IV were analyzed with glutamate/malate as substrate, and inhibited by rotenone. Complexes II + III + IV were analyzed with succinate as substrate. TMPD/ascorbate have been used to analyze complex IV of respiratory chain. (C) Acute applications of simvastatin resulted in a strong inhibition of respiration with glutamate/malate as substrate, and a low inhibition of respiration with TMPD/ascorbate as substrate (n = 8). *p < 0.05 compare to control values.
absent in cardiomyocyte [20]. We then hypothesized that the striated muscle specificity of statins might be due to statin transport through MCT4. Thus, experiments were conducted on intact rat FDB fibers pre-incubated with 100 lM p-chloromercurbenzene sulfonate (PCMBS), a potent inhibitor of MCT4 [21]. In this condition, the release of calcium triggered by simvastatin was totally abolished (Fig. 7D).
Statins are widely prescribed and demonstrated to be highly efficient to prevent cardiovascular diseases [6,22]. However, they are also known to induce skeletal muscle abnormalities ranging from myalgia to myopathy with an incidence of 1–7% [1,23]. If myopathy is not recognized and treatment maintained, necrosis of muscle and rhabdomyolysis may occur with an incidence of 0.04–0.2% [24,25]. We recently reported that acute application of simvastatin on human skeletal muscle fibers induced a mitochondrialdependent impairment of Ca2+ homeostasis [13]. This might contribute to the deleterious effects of statins such as calpaı¨n activation and apoptosis as recently reported [26]. In this study, we further investigated the cellular effect of simvastatin and observed that this adverse effect: (1) is specific of skeletal muscle and independent of muscle phenotype, (2) is linked to MCT4 present in skeletal muscle while absent in cardiac tissue, and (3) results primarily from an alteration of the mitochondrial respiratory chain mainly focused on an inhibition of complex I (i.e., NADH dehydrogenase). As previously reported [13], simvastatin triggers Ca2+ efflux by both NCE and PTP responsible for a SR-Ca2+ overload subsequently followed by a large SR-dependent Ca2+ release. Mitochondrial calcium efflux is associated with depolarization of the inner membrane of the mitochondria and increased NADH autofluorescence, suggesting alteration of mitochondrial energetic function by simvastatin. The present study demonstrates that Ca2+ efflux through PTP and NCE is a consequence of mitochondrial membrane potential fall observed after acute application of simvastatin. Indeed, inner membrane depolarization occurs even in the absence of Ca2+ efflux through PTP and NCE. Since proton flux through respiratory chain is the precursor of the inner mitochondrial membrane potential, the origin of simvastatin deleterious effects could be linked to an altered mitochondrial respiratory chain and mitochondrial energetic function. Oxymetric measurements of mitochondrial respiration clearly demonstrate that simvastatin alters mitochondrial energetic function at the origin of the inner membrane potential fall independently of the metabolic profile. In order to further determine the origin of simvastatininduced impairment of mitochondrial respiration, we assess simvastatin effects using different substrates and inhibitors of respiratory chain complexes. Using palmitoyl-carnitine/malate, or pyruvate/malate as substrates, we demonstrated that simvastatin potentially affects in the same range lipidic and glucidic energetic pathways. Thus, simvastatin affects commune downstream targets of these pathways, namely Krebs cycle or more probably respiratory chain. We used FCCP to uncouple oxidative phosphorylation and dissociate complex V from the others. Although we cannot totally exclude a direct effect of simvastatin on complex V of respiratory chain (ATP synthase), the strong inhibition of respiration rate in these conditions
P. Sirvent et al. / Biochemical and Biophysical Research Communications 338 (2005) 1426–1434
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Fig. 6. Simvastatin-induced alteration in Ca homeostasis. x–y fluorescence images recorded in bundles of skinned human skeletal muscle fibers (A), in skinned rat EDL (B), and soleus (C) loaded with Fluo-3 (50 lM). In each image, average fluorescence is measured in a region of interest and reported as a function of time. Acute application of simvastatin resulted in a delayed (10 min) increase in [Ca2+]i independently of muscle phenotype.
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Fig. 7. Effect of simvastatin on ventricular cardiomyocytes. x–y fluorescence images recorded in intact (A) and saponin-skinned (B) rat ventricular myocytes and in intact rat FDB enzymatically dissociated fibers in the absence (C) or in the presence on 100 lM MCT4 inhibitor PCMBS (D). Intact fibers were loaded with Fluo-3 AM (5 lM in the external medium for 30 min) and permeabilized cells were incubated with 50 lM Fluo-3 pentapotassium salt in the internal medium. In each image, fluorescence is measured in a region of interest and reported as a function of time. Simvastatin was used at the concentration of 200 lM. Intact cardiomyocytes exhibited spontaneous Ca2+ transients that were independent of the presence of simvastatin.
supports the contention that a major target of simvastatin is focused on the four first complexes of the respiratory chain. As a matter of fact, our data indicated that complex I is the main complex affected by simvastatin. The larger simvastatin effect on respiration rate occurs when using glutamate/malate as substrates No significant effects could be observed with succinate and rotenone, strongly supporting a direct effect of simvastatin on complex I, while complexes II seem to be unaffected. This is also in agreement with the simvastatin-induced increase in NADH auto fluo-
rescence observed previously [13]. Moreover, using TMPD and ascorbate as substrate, we could notice a weak inhibition by simvastatin of complex IV. Mazat et al. [27] have shown that limited inhibition of complex IV as observed here did not affect global respiration rate. Based on these observations they concluded that complex IV is not a limiting step of respiratory chain. Considering the low amplitude of the simvastatin-induced inhibition of this complex, we assume that this effect would not participate predominantly to the global inhibition of respiratory chain induced
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by simvastatin. The mechanism by which simvastatin inhibits complex I remains to be elucidated. However, one can postulate that the interaction of simvastatin with this complex might block the transfer of electrons within the complex or between complex I and II. Skeletal and heart muscles shear a lot of homologies in regard to their metabolism and regulation of Ca2+ homeostasis and contractile function. Nevertheless, deleterious effects of statins were to this date only reported in skeletal muscles. As a matter of fact, it has been recently reported that administration (40 mg/kg/day) of pravastatin for 3 weeks in mice induced marked ultrastructure alterations of mitochondria in skeletal muscle while the ultrastructure of the heart and liver of the treated animals appeared normal [28]. In the present work, we further sought to test this skeletal muscle specificity of statinsÕ deleterious effects. Therefore, acute applications of simvastatin on both intact and permeabilized isolated rat cardiomyocytes have been performed. While simvastatin was unable to alter Ca2+ homeostasis in intact cardiomyocytes, one could observe on permeabilized cells a large release of Ca2+ preceded by a slight elevation in cytosolic Ca2+. As in skeletal muscle fibers [13], this first phase was correlated with a mitochondrial membrane depolarization. A similar phenomenon has already been observed in heart cells with DCP, an inhibitor of mitochondrial respiration [29]. Acute application of DCP on heart cells induced a mitochondrial membrane potential fall, a release of Ca2+ by mitochondria, and a delayed large increase in [Ca2+]i comparable with the effects observed in our study. Thus, simvastatin presents a similar potential to alter both skeletal muscle and heart cellsÕ internal machinery. It has been shown that simvastatin diffuses in kidney cells through the monocarboxylate transporter MCT4. This isoform is abundant in skeletal muscle but absent in heart tissue [30]. Although other transporters such as organic anion transporter have been proposed for potential statin uptake [31], their relative contribution compared to MCTs is not clear. Inhibition of MCT4 abolished simvastatin-induced alteration in Ca2+ homeostasis in intact skeletal muscles, suggesting a predominant participation of MCT4 in statin uptake. This could account for the difference observed in intact skeletal muscle and heart cells with acute applications of simvastatin. We cannot however totally exclude that simvastatin diffuses into cardiomyocytes during chronic treatment via other pathways. Functional differences between heart and skeletal muscles might also account for this specificity. Among them, one can notice the predominant role of sodium/calcium exchanger (NCX) in heart cells to control Ca2+ homeostasis. While skeletal muscle has a high capacity to reuptake its intracellular Ca2+ to the SR, cardiomyocytes tend to avoid cytoplasmic and sarcoplasmic reticulum Ca2+-overload by extruding Ca2+ through NCX [32]. In conclusion, we demonstrate here that a specific alteration of complex I of respiratory chain represents the early step of a cascade of cellular deleterious mechanisms induced by simvastatin. This effect can be seen on both
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slow and fast type skeletal muscles, but not on intact heart cells. Cardiomyocytes appear to be protected by their lack of MCT4 which mediates statin uptake in skeletal muscle. The effects described here in vitro could easily account for the adverse effects observed in vivo in statin-treated patients. It would be important to further elucidate the myotoxicity induced by statins and interesting to confirm this demonstration with muscle biopsies from statin-treated patients and suffering from myalgia. Acknowledgments This study was supported by grants from INSERM, the Association Franc¸aise contre les Myopathies (AFM), the Fondation pour la Recherche Me´dicale (FRM), and the Agence Franc¸aise de Se´curite´ Sanitaire des Produits de Sante´ (AFSSAPS). References [1] D.J. Maron, S. Fazio, M.F. Linton, Current perspectives on statins, Circulation 101 (2000) 207–213. [2] C.J. Vaughan, A.M. Gotto Jr., Update on statins: 2003, Circulation 110 (2004) 886–892. [3] J.A. Tobert, G.D. Bell, J. Birtwell, I. James, W.R. Kukovetz, J.S. Pryor, A. Buntinx, I.B. Holmes, Y.S. Chao, J.A. Bolognese, Cholesterol-lowering effect of mevinolin, an inhibitor of 3-hydroxy-3methylglutaryl-coenzyme a reductase, in healthy volunteers, J. Clin. Invest. 69 (1982) 913–919. [4] Design and baseline results of the Scandinavian Simvastatin Survival Study of patients with stable angina and/or previous myocardial infarction, Am. J. Cardiol. 71 (1993) 393–400. [5] F.M. Sacks, M.A. Pfeffer, L.A. Moye, J.L. Rouleau, J.D. Rutherford, T.G. Cole, L. Brown, J.W. Warnica, J.M. Arnold, C.C. Wun, B.R. Davis, E. Braunwald, The effect of pravastatin on coronary events after myocardial infarction in patients with average cholesterol levels. Cholesterol and Recurrent Events Trial investigators, N. Engl. J. Med. 335 (1996) 1001–1009. [6] Prevention of cardiovascular events and death with pravastatin in patients with coronary heart disease and a broad range of initial cholesterol levels. The Long-Term Intervention with Pravastatin in Ischaemic Disease (LIPID) Study Group, N. Engl. J. Med. 339 (1998) 1349–1357. [7] M. Takemoto, K. Node, H. Nakagami, Y. Liao, M. Grimm, Y. Takemoto, M. Kitakaze, J.K. Liao, Statins as antioxidant therapy for preventing cardiac myocyte hypertrophy, J. Clin. Invest. 108 (2001) 1429–1437. [8] J.K. Liao, Isoprenoids as mediators of the biological effects of statins, J. Clin. Invest. 110 (2002) 285–288. [9] J. Davignon, Beneficial cardiovascular pleiotropic effects of statins, Circulation 109 (2004) III39–III43. [10] J.P. Halcox, J.E. Deanfield, Beyond the laboratory: clinical implications for statin pleiotropy, Circulation 109 (2004) II42–II48. [11] M. Ucar, T. Mjorndal, R. Dahlqvist, HMG-CoA reductase inhibitors and myotoxicity, Drug Safety 22 (2000) 441–457. [12] S. Bellosta, R. Paoletti, A. Corsini, Safety of statins: focus on clinical pharmacokinetics and drug interactions, Circulation 109 (2004) III50–III57. [13] P. Sirvent, J. Mercier, G. Vassort, A. Lacampagne, Simvastatin triggers mitochondria-induced Ca2+ signaling alteration in skeletal muscle, Biochem. Biophys. Res. Commun. 329 (2005) 1067–1075. [14] O. Cazorla, A. Lacampagne, J. Fauconnier, G. Vassort, SR33805, a Ca2+ antagonist with length-dependent Ca2+-sensitizing properties in cardiac myocytes, Br. J. Pharmacol. 139 (2003) 99–108.
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[15] V. Lukyanenko, S. Gyorke, Ca2+ sparks and Ca2+ waves in saponinpermeabilized rat ventricular myocytes, J. Physiol. 521 (Pt 3) (1999) 575–585. [16] C. Thomas, P. Sirvent, S. Perrey, E. Raynaud, J. Mercier, Relationships between maximal muscle oxidative capacity and blood lactate removal after supramaximal exercise and fatigue indexes in humans, J. Appl. Physiol. 97 (2004) 2132–2138. [17] V.A. Saks, V.I. Veksler, A.V. Kuznetsov, L. Kay, P. Sikk, T. Tiivel, L. Tranqui, J. Olivares, K. Winkler, F. Wiedemann, W.S. Kunz, Permeabilized cell and skinned fiber techniques in studies of mitochondrial function in vivo, Mol. Cell. Biochem. 184 (1998) 81– 100. [18] M. Jakobisiak, S. Bruno, J.S. Skierski, Z. Darzynkiewicz, Cell cyclespecific effects of lovastatin, Proc. Natl. Acad. Sci. USA 88 (1991) 3628–3632. [19] K. Nagasawa, K. Nagai, Y. Sumitani, Y. Moriya, Y. Muraki, K. Takara, N. Ohnishi, T. Yokoyama, S. Fujimoto, Monocarboxylate transporter mediates uptake of lovastatin acid in rat cultured mesangial cells, J. Pharm. Sci. 91 (2002) 2605–2613. [20] A.P. Halestrap, N.T. Price, The proton-linked monocarboxylate transporter (MCT) family: structure, function and regulation, Biochem. J. 343 (Pt. 2) (1999) 281–299. [21] J.E. Manning Fox, D. Meredith, A.P. Halestrap, Characterisation of human monocarboxylate transporter 4 substantiates its role in lactic acid efflux from skeletal muscle, J. Physiol. 529 (Pt. 2) (2000) 285–293. [22] J.R. Downs, M. Clearfield, S. Weis, E. Whitney, D.R. Shapiro, P.A. Beere, A. Langendorfer, E.A. Stein, W. Kruyer, A.M. Gotto Jr., Primary prevention of acute coronary events with lovastatin in men and women with average cholesterol levels: results of AFCAPS/ TexCAPS. Air Force/Texas Coronary Atherosclerosis Prevention Study, Jama 279 (1998) 1615–1622.
[23] W.R. Garnett, Interactions with hydroxymethylglutaryl-coenzyme A reductase inhibitors, Am. J. Health Syst. Pharm. 52 (1995) 1639–1645. [24] Y. Berland, H. Vacher Coponat, C. Durand, M. Baz, R. Laugier, J.L. Musso, Rhabdomyolysis with simvastatin use, Nephron 57 (1991) 365–366. [25] M.A. Omar, J.P. Wilson, T.S. Cox, Rhabdomyolysis and HMG-CoA reductase inhibitors, Ann. Pharmacother. 35 (2001) 1096–1107. [26] J. Sacher, L. Weigl, M. Werner, C. Szegedi, M. Hohenegger, Delineation of myotoxicity induced by 3-hydroxy-3-methylglutaryl CoA reductase inhibitors in human skeletal muscle cells, J. Pharmacol. Exp. Ther. 314 (2005) 1032–1041. [27] J.P. Mazat, T. Letellier, F. Bedes, M. Malgat, B. Korzeniewski, L.S. Jouaville, R. Morkuniene, Metabolic control analysis and threshold effect in oxidative phosphorylation: implications for mitochondrial pathologies, Mol. Cell. Biochem. 174 (1997) 143–148. [28] M. Bergman, H. Salman, M. Djaldetti, S. Alexandrova, I. Punsky, H. Bessler, Ultrastructure of mouse striated muscle fibers following pravastatin administration, J. Muscle Res. Cell Motil. 24 (2003) 417– 420. [29] D. Hudman, R.D. Rainbow, C.L. Lawrence, N.B. Standen, The origin of calcium overload in rat cardiac myocytes following metabolic inhibition with 2,4-dinitrophenol, J. Mol. Cell. Cardiol. 34 (2002) 859–871. [30] A. Bonen, Lactate transporters (MCT proteins) in heart and skeletal muscles, Med. Sci. Sports Exerc. 32 (2000) 778–789. [31] M. Takeda, R. Noshiro, M.L. Onozato, A. Tojo, H. Hasannejad, X.L. Huang, S. Narikawa, H. Endou, Evidence for a role of human organic anion transporters in the muscular side effects of HMG-CoA reductase inhibitors, Eur. J. Pharmacol. 483 (2004) 133–138. [32] H. Lullmann, A. Ziegler, Calcium, cell membrane, and excitation– contraction coupling, J. Cardiovasc. Pharmacol. 10 (Suppl. 1) (1987) S2–S8.
Simvastatin induces impairment in skeletal muscle while heart is protected Pascal Sirvent a,b, Sylvain Bordenave b, Marianne Vermaelen b, Belle Roels c, Guy Vassort a, Jacques Mercier b, Eric Raynaud b, Alain Lacampagne a,* a
c
INSERM U637 CHU A. de Villeneuve, 34295 Montpellier, France b EA 701, Universite´ Montpellier I, 34000 Montpellier, France UPRES EA 3759 Faculty of Sport Sciences, 700 avenue Pic St Loup, 34090 Montpellier, France Received 17 October 2005 Available online 26 October 2005
Abstract 3-Hydroxy-3-methylglutaryl-coenzyme A reductase inhibitors (statins) are widely used to reduce plasma cholesterol concentration. However, statins are also known to induce various forms of muscular toxicity. We have previously shown that acute application of simvastatin on human skeletal muscle samples induced a cascade of cellular events originating from mitochondria and resulting in a global alteration of Ca2+ homeostasis. The present study was designed to further define the origin of the mitochondria impairment and to understand the apparent lack of deleterious effect on the heart. Using fluorescence imaging analysis and oxygraphy on human and rat skinned skeletal muscle samples, we show that the simvastatin-induced mitochondria impairment results from inhibition of the complex I of respiratory chain. Similar simvastatin-induced mitochondria impairment and alteration of Ca2+ homeostasis occur in permeabilized but not in intact ventricular rat cardiomyocytes. In intact rat skeletal muscle fibers from the flexor digitorum brevis muscle, the simvastatin-induced alteration of Ca2+ homeostasis is abolished when monocarboxylate transporter (MCT4) is inhibited. The impairment of complex I by simvastatin might be the primary step of its cellular deleterious effects leading to muscle fiber death. This mechanism is seen specifically in skeletal muscles. This specificity should be in part attributed to a preferential uptake of statins by MCT4 that is not expressed in cardiomyocytes. 2005 Elsevier Inc. All rights reserved. Keywords: Human skeletal muscle; Rat cardiomyocytes; Calcium; Mitochondria; Myotoxicity; Hypercholesterolemia; Satins; Monocarboxylate transporter
Hypercholesterolemia is a well-known risk factor for coronary heart disease. Among various drugs developed to reduce plasma cholesterol concentration, 3-hydroxy-3methylglutaryl-coenzyme A (HMG-CoA) reductase inhibitors (statins) are likely to be the most promising drug [1,2]. Statins decrease cholesterol content by selective inhibition of HMG-CoA reductase, rate-limiting step in the mevalonate synthesis pathway [3]. Clinical trials have demonstrated a reduction in cardiovascular-related morbidity and mortality in treated patients with or without coronary artery disease [4–6]. In addition to the benefits of lowering cholesterol, it has been shown that statins present choles*
Corresponding author. Fax: +33 467 415 242. E-mail address: [email protected] (A. Lacampagne).
0006-291X/$ - see front matter 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2005.10.108
terol-independent effects allowing for improved endothelial function, enhanced stability of atherosclerotic plaques, decreased inflammation and oxidative stress, and inhibited thrombogenic response in the vascular wall [7–10]. Adverse reactions involving skeletal muscles (myalgias, cramps, weakness, and exercise intolerance) are the most frequent, occurring in 1–7% of statin-treated patients [11]. These adverse effects can occur with or without increase in creatine kinase (CK) level. Some rare cases of fatal rhabdomyolysis have also been reported, with comedication or overdose [11,12]. If these adverse effects do not call into question efficiency and global safety of statin treatments, they can disturb treated patients in their everyday life. Taking into account the great number of statintreated patients, which would increase again in the next
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few years, comprehension of mechanisms of statin-induced myotoxicity seems to be determinant in order to prevent this adverse effect. We have recently shown that acute applications of simvastatin on human skeletal muscle samples trigger a large release of Ca2+ from the SR, resulting from early alterations of mitochondrial function [13]. The present study was designed to further define the origin of the mitochondrial impairment as well as to understand the apparent lack of deleterious effect on heart. Here, we demonstrate on human and rat muscle samples that: (1) the simvastatin-induced Ca2+ release does not affect cardiac cells and is specific of skeletal muscle whatever muscle phenotype, (2) the simvastatin-induced Ca2+ homeostasis impairment is predominantly dependent on statin uptake through monocarboxylate-transporter (MCT4), (3) a specific alteration of complex I of the respiratory chain appears to be the initial step of a cascade of deleterious cellular effects leading to Ca2+ homeostasis alteration. Materials and methods Human skeletal muscle samples. Muscle samples are prepared as previously described [13]. Fifteen healthy voluntary men aged between 25 and 45 were recruited in order to take part in the study. They did not take any medication known to potentially interfere with the protocol. The study was approved by the Local Ethics Committee and conformed to the Declaration of Helsinki regarding the use of human subject and informed consent was obtained from all the subjects after explanation of the nature of the study and potential risks. Muscle biopsies were performed in the inferior third level of the vastus lateralis muscle under local anesthesia with a Bergstro¨m needle. For imaging experiments, biopsies (40 mg) were immediately placed in the internal-like medium (in mol/L: K-glutamate 140, Hepes 10, phosphocreatine 20, Na2ATP 5, MgCl2 4.53, EGTA 1, CaCl2 0.29, and malate 2, pH 7.0). For in situ mitochondrial respiration studies, biopsies were placed in an ice-cold relaxing solution (in mM: EGTA–calcium buffer 10 (free Ca2+ concentration: 100 nM), imidazole 20, KH2PO4 3, MgCl2 1, taurine 20, DTT 0.5, MgATP 5, and phosphocreatine 15, pH 7.1). Rat muscle and heart samples. Male Wistar rats weighting 300 g (Janvier; Le Genest-St-Isle, France) were used. This investigation conforms to the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 85-23, Revised 1996) and European directives (96/ 609/EEC) were used. EDL and soleus muscle were dissected and kept in an external medium containing (in mM): NaCl 145, KCl 4, CaCl2 1.8, MgSO4 1, Hepes 10, and glucose 10, pH 7.4. For imaging experiments, muscles were then set in the dissecting chamber in a solution containing (in mM): K-glutamate 140, Hepes 10, MgCl2 10, and EGTA 0.1, pH 7.0. For in situ mitochondrial respiration studies, samples were placed in the same ice-cold relaxing solution as for human biopsies. Flexor digitorum brevis (FDB) muscles were dissected and enzymatically dissociated with 3 mg/ml collagenase (type 1; Sigma) dissolved in an external medium (see above) at 37 C for 1 h. Bundles of fibers were then transferred to an external medium without collagenase, and fibers were separated mechanically by gentle trituration. Intact single fibers were plated on extracellular matrix (Sigma)-coated coverslips attached across a 12-mm hole in the bottom of 35-mm petri dishes. Cardiac ventricular cells were enzymatically isolated as described earlier [14]. Briefly, rats were anesthetized by intraperitoneal injection of pentobarbital (200 mg/100 g). The heart was perfused retrogradely with a Ca2+-free Hanks-Hepes-buffered solution (in mM: NaCl 117, KCl 5.7, NaHCO3 4.4, KH2PO4 1.5, MgCl2 1.7, Hepes 21, glucose 11, and taurine 20, pH 7.2, adjusted with NaOH and bubbled with 100% O2) for 5 min at 37 C and then with an
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enzyme-containing solution (same as above with 1.0 mg/ml collagenase type IV (Worthington, Freehold, NJ, USA) and 20 lM Ca2+) for 25– 30 min. The ventricles were then gently dissociated with forceps in the same medium without enzyme. The cells were then filtered and further washed several times in a solution containing 0.3 mM Ca2+. Finally, myocytes were kept in a medium containing 1 mM Ca2+ and 0.5% bovine serum albumin (BSA). Cardiomyocytes were permeabilized as described by others [15]. Cardiomyocytes were placed in an internal medium (containing in mM: 120 potassium aspartate, 3 MgATP, 2 EGTA, 0.11 CaCl2 (free [Ca2+] = 100 nM), 10 phosphocreatine, and 5% dextran (40,000); pH 7.2.) with saponin (0.01% for 45–60 s). Experiments were performed with 50 lM Fluo-3 potassium salt (TefLabs, Austin, TX, USA) in the internal medium. Measurements of cytoplasmic Ca2+. Bundles of 2–5 fibers were manually dissected under a dissecting microscope (Stemi 200 Zeiss, France). Fibers were mounted in an experimental chamber containing an internal medium and permeabilized by adding saponin 0.01%, 30 s at room temperature. Fibers were slightly stretched to a sarcomere length around 2.8–3.2 lm to reduce movement artifacts. Fibers were then bathed in an internal medium containing the fluorescent Ca2+ indicator Fluo-3 (pentapotassium salt; TefLabs, Austin, TX, USA; 50 lM). FDB fibers and isolated cardiomyocytes bathed in an external medium, were loaded during 30 min at room temperature with Fluo-3 AM (5 lM, TefLabs, Austin, TX, USA). Variations in intracellular Ca2+ were measured by time-series (1 image/5 s) of x–y confocal fluorescent images (Zeiss LSM 510 Meta, 63· objective, NA = 1.2, H2O immersion). Fluo-3 was excited with an argon/krypton laser at 488 nm, and emitted fluorescence was recorded at 525 nm. Mean fluorescence was calculated in regions of interest (see Fig. 1, white box) and reported as a function of time. Measurements of mitochondrial Ca2+. To assess mitochondrial Ca2+, fibers were incubated for 30 min at room temperature with Rhod-2 AM (5 lM) in the internal medium. Rhod-2 is taken up preferentially into polarized mitochondria. After washes (3 times 5 min in internal medium) to remove cytosolic Rhod-2, variations in mitochondrial Ca2+ were recorded by time-series (1 image/5 s) of x–y confocal images. Fluorescence intensity was recorded at 570 nm, with an excitation at 543 nm provided by a helium/neon laser. Measurements of mitochondrial membrane potential. Mitochondrial membranes were labeled with the rhodamine-based fluorescent cationic lipophilic tetramethylrhodamine ethyl ester (TMRE, 100 nM), which distributes across the inner mitochondrial membrane in accordance with the Nernst equation. After washes (3 times 5 min in internal medium), variations in mitochondrial membrane potential were recorded by timeseries (1 image/5 s) of x–y confocal images. Fluorescence intensity was recorded above 560 nm, with an excitation at 543 nm provided by a helium/neon laser. Mean fluorescence was calculated in small regions of interest corresponding to mitochondrial staining and reported as a function of time. Measurements of mitochondrial respiration. Respiratory parameters of the total mitochondrial population were studied in situ on fresh skeletal muscle fibers as previously described [16]. Bundles of muscle fibers were manually isolated and saponin-skinned (50 lg/ml saponin for 30 min at 4 C). Respiration rates were determined at 27 C with a Clark electrode (Strathkelvin, Glasgow, Scotland) in an oxygraphic cell containing respiration solution (same composition as the relaxing solution, except that MgATP and phosphocreatine were replaced by 3 mM phosphate and 2 mM fatty acid-free bovine serum albumin). Respiration rates were expressed in micromoles of O2 per minute per gram of dry weight. Basal oxygen consumption without ADP (V0), maximal respiration rate (Vmax) with a saturating concentration of ADP and acceptor control ratio (ACR), and the ratio between Vmax and V0, which represents the level of coupling between oxidation and phosphorylation, have been measured. At the end of each measurement, cytochrome c test used to investigate the outer mitochondrial membrane integrity [17]. Respiration rates were recorded in the presence of glutamate (5 mM)/malate (2 mM) or pyruvate (10 mM)/malate (2 mM) or palmitoyl-carnitine (40 lM)/malate (2 mM). Basal oxygen consumption without ADP (V0) was first
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Fig. 1. Simvastatin triggers mitochondrial Ca2+ release. Fibers were loaded with Rhod-2 AM to monitor mitochondrial [Ca2+]. (A) Acute applications of simvastatin (50 lM) resulted in a dose-dependent decrease in fluorescence indicating a mitochondrial Ca2+ efflux. (B) In fibers pretreated with both cyclosporin A and clonazepam, simvastatin did not induce Ca2+ release. (C) Dose-dependent effects of simvastatin on mitochondrial Rhod-2 fluorescence. (*p < 0.05 compare to control values.)
recorded, and Vmax was determined in the presence of 2 mM ADP. Increasing concentrations of simvastatin diluted were then added. For calculation of the ACR, V0 was recorded with 50 lM final concentration of simvastatin; 2 mM ADP was then added for maximal activation of respiration. The effects of simvastatin on respiratory chain complexes were investigated at maximal respiration rates (ADP 2 mM) and for 50 lM of simvastatin. The specific substrates and inhibitors used to study the effects of simvastatin on the respiratory chain complexes are summarized in Fig. 5. Drugs and chemical reagents. All drugs used were purchased from Sigma, except for Fluo-3 and Rhod-2 (Teflab) and TMRE (Molecular Probes). Simvastatin was a generous gift from Merck laboratories. The provided lactone form of simvastatin was converted to the acidic pharmacologically active form as described by others [18]. Since simvastatin is diluted in ethanol, a series of control experiments containing the same amount of solvent was performed to exclude a potential effect of solvent. Statistical analysis. Statistical comparisons were done by a variance analysis (ANOVA). Data are expressed as means ± SEM and difference was considered significant with p < 0.05.
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Bundles of human muscle fibers were loaded with Rhod2 AM to monitor mitochondrial Ca2+ (Fig. 1). Acute application of simvastatin (3–100 lM) resulted in a dosedependent release of Ca2+ by mitochondria (Figs. 1A and C) that reached 59 ± 5% (n = 7) depletion at maximum (10 lM). Efflux of Ca2+ from mitochondria can be carried out either by the permeability transient pore (PTP) or the Na+–Ca2+ exchanger (NCE), respectively, blocked by cyclosporine A and clonazepam. Pre-treatment of fibers with 0.2 lM clonazepam or 0.5 lM cyclosporine A did not block the statin-induced Ca2+ release by mitochondria.
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Fig. 2. Simvastatin induces a fall in mitochondrial membrane potential. (A) Acute application of simvastatin (50 lM) on permeabilized muscle fibers loaded with TMRE resulted in a dose-dependent decrease in fluorescence compared to control conditions. (B) Pre-treatment of fibers with cyclosporin A and clonazepam did not prevent mitochondrial membrane potential fall. (C) Dose-dependent effects of simvastatin on mitochondrial TMRE fluorescence. (*p < 0.05 compare to control values.)
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deshydrogenase complex, Krebs cycle, and respiratory chain. With both substrates, maximal ADP-stimulated respiration was inhibited by 20% after applying 50 lM simvastatin (Figs. 4A–D). We used FCCP to uncouple oxidative phosphorylation, using glutamate/malate as substrate (Figs. 5C and D). In this uncoupled condition, simvastatin induced a stronger inhibition of respiration rate (38%). This difference in percentage of inhibition is explained by the difference in maximal rate of respiration known to be slower in coupled than in uncoupled oxidative phosphorylation. These observations on uncoupled respiratory chain render the hypothesis of an uncoupling effect of simvastatin very unlikely. Taken together, these data strongly suggest an effect of simvastatin upstream to the ATP synthase, i.e., one or more of the complexes I–IV of the respiratory chain of the mitochondria.
However, pre-treatment with both drugs simultaneously completely abolished this phenomenon (Figs. 1B and C). Changes in mitochondrial membrane potential were then assessed on bundles of human muscle fibers loaded with TMRE (Fig. 2). Acute applications of simvastatin (1–100 lM) resulted in rapid depolarizations (47 ± 1% at maximum, n = 6) of mitochondrial membrane that were dose-dependent (Figs. 2A and C). Pre-treatment with both 0.2 lM clonazepam and 0.5 lM cyclosporine A did not prevent this phenomenon (Figs. 2B and C). This result implies that the fall in mitochondrial membrane potential is independent of NCE and PTP opening. It might result from an upstream step such as an alteration of the respiratory chain electron flux, which allows the formation of the electrochemical gradient through the inner membrane. Simvastatin alters mitochondrial respiration
Simvastatin affects mainly complex I of respiratory chain Mitochondrial respiration was assessed on bundles of saponin-skinned human skeletal muscle fibers. Acute application of simvastatin resulted in a dose-dependent inhibition of maximal ADP-stimulated rate of oxygen consumption (Figs. 3A and B). This effect was statistically significant at 10 lM, and independent of muscle phenotype since simvastatin similarly altered maximal ADP-stimulated respiration rate on both soleus and EDL muscle fibers (Figs. 3C and D). We then evaluated mitochondrial respiration with different substrates to determine the origin of simvastatin-induced impairment. We used palmitoyl-carnitine/malate as substrate to assess lipid pathway, testing b-oxidation, Krebs cycle, and respiratory chain. For glucidic pathway, we used pyruvate/malate as substrate, testing pyruvate A
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We used specific substrates and inhibitors to sequentially evaluate the effect of simvastatin on the different complexes of respiratory chain (Figs. 5A and B). Glutamate/ malate as substrates allow to activate complexes I, III, and IV of respiratory chain. The use of succinate and rotenone would restrict the activity of the respiratory chain to complexes II, III, and IV. Finally, the use of TMPD (N,N,N 0 ,N 0 -tetramethyl-p-phenylenediamine) and ascorbate should test specifically the activity of complex IV. Acute applications of simvastatin induced a strong inhibition of maximal ADP-stimulated respiration rate with glutamate/malate as substrate. No significant inhibition could be seen using succinate as substrate, and only a very weak inhibition with TMPD and ascorbate (Fig. 5C). Complex C
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Fig. 3. Simvastatin inhibits mitochondrial respiration rate. (A) Mitochondrial respiration was estimated with acute applications of various concentrations of simvastatin on permeabilized bundles of human skeletal muscle fibers using pyruvate/malate as substrate. (B) Dose-dependent decrease in the maximal ADP-stimulated respiration rate induced by simvastatin. (C) Mitochondrial respiration recorded on permeabilized rat Soleus (left) and EDL (right) muscle fibers. (D) In both muscle types, acute applications of 10 lM simvastatin resulted in a decrease of maximal ADP-stimulated respiration rate to the same extent. (n = 8; *p < 0.05 compare to Vmax.)
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Fig. 4. Simvastatin alters glucidic and lipidic mitochondrial metabolic pathway. (A,B) Mitochondrial respiration was recorded on permeabilized human skeletal muscle fibers in the presence of palmitoyl-carnitine or pyruvate as main substrate. (C) Acute applications of simvastatin (50 lM) resulted in a 20% decrease in maximal ADP-stimulated respiration rate with both substrates. FCCP has been used to decouple mitochondrial respiratory chain. Acute application of simvastatin (50 lM) on decoupled mitochondria induced a strong decrease in respiration rate. (D) Changes in the rate of O2 consumption ( VO2), in the presence of simvastatin relative to the rates measured in respective control conditions. (n = 8; *p < 0.05 compare to control values.)
IV is not a limiting step in the respiratory chain. In normal conditions, when global respiration is activated, it functions at 50% of its maximal activity reached with TMPD/ascorbate, accounting for the discrepancy of observations between succinate/rotenone and TMPD/ascorbate. Indeed, the slight decrease observed in succinate/rotenone is likely to be only due to the inhibition of complex IV that is only partially activated in this condition. These results demonstrate that simvastatin effect is mainly due to an inhibition of complex I of respiratory chain. Effects of simvastatin on cytosolic and mitochondrial Ca2+: does simvastatin alter Ca2+ homeostasis in cardiomyocytes? Secondary to mitochondrial impairment, the acute application of simvastatin on skinned human skeletal muscle fibers resulted in a delayed large increase in [Ca2+]i (250 ± 25%, n = 36, Fig. 6A) as previously observed [13]. Vastus lateralis is known to be composed of both oxidative (slow twitch) and glycolytic (fast twitch) muscle fibers. In order to assess the effects of simvastatin on muscle fibers with different phenotypes, we used permeabilized cut myofibers manually dissected from rat EDL (fast) and soleus (slow) muscles. Acute application of simvastatin resulted in a delayed large transient Ca2+ release in both situations similarly (Figs. 6B and C). Acute application of simvastatin was also tested on rat ventricular cardiomyocytes to verify whether the adverse effect of simvastatin was generalized to all types of striated muscles. When applied on intact cardiac cells, simvastatin used at 200 lM did not trigger a large increase in intracellular Ca2+ similar to that seen in skeletal muscle,
even after more than 1 h of exposure (Fig. 7A). This implies that simvastatin does not easily diffuse into the cell or that the cascade of intracellular events triggered by simvastatin in skeletal muscle does not occur in cardiomyocytes. To answer this point, acute applications of simvastatin were performed on saponin-skinned cardiomyocytes (Fig. 7B). In these conditions, acute application of 200 lM simvastatin resulted in a fast increase in resting level of Ca2+ followed after 7–8 min by a larger increase in cytoplasmic [Ca2+] to some extent comparable to that observed on permeabilized skeletal muscle fibers. In some experiments, saponin-skinned cardiomyocytes were also loaded with TMRE to assess mitochondrial membrane potential. Acute application of simvastatin triggered a depolarization of mitochondrial membrane correlated with the first increase in [Ca2+]i, suggesting that this increase in resting Ca2+, as previously demonstrated in skeletal muscle [13], resulted from a mitochondrial Ca2+ efflux (not shown). This phenomenon was absent in intact cardiomyocytes. Human muscle biopsies do not allow us to obtain intact adult muscle fibers. To evaluate the effects of acute applications of simvastatin on intact skeletal muscle cells, we used intact rat flexor digitorum brevis (FDB) fibers dissociated enzymatically. In contrast with cardiomyocytes, simvastatin induced in intact muscle fibers a large increase in [Ca2+]i although with a prolonged delay and preceded by a slow increase in basal [Ca2+]i (Fig. 7C). Monocarboxylate transporter (MCT) and particularly the type 4 (MCT4) has been shown to mediate uptake of statins in rat mesanglial cells [19] and bovine kidney NBL-1 cells [19]. Interestingly, while MCT4 is largely expressed in skeletal muscle, it is
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Fig. 5. Simvastatin-induced mitochondrial respiratory chain impairment. (A) Schematic representation of complexes of mitochondrial respiratory chain. Different substrates and inhibitors have been used to isolate and study independently the different complexes as indicated. (B) Complexes I + III + IV were analyzed with glutamate/malate as substrate, and inhibited by rotenone. Complexes II + III + IV were analyzed with succinate as substrate. TMPD/ascorbate have been used to analyze complex IV of respiratory chain. (C) Acute applications of simvastatin resulted in a strong inhibition of respiration with glutamate/malate as substrate, and a low inhibition of respiration with TMPD/ascorbate as substrate (n = 8). *p < 0.05 compare to control values.
absent in cardiomyocyte [20]. We then hypothesized that the striated muscle specificity of statins might be due to statin transport through MCT4. Thus, experiments were conducted on intact rat FDB fibers pre-incubated with 100 lM p-chloromercurbenzene sulfonate (PCMBS), a potent inhibitor of MCT4 [21]. In this condition, the release of calcium triggered by simvastatin was totally abolished (Fig. 7D).
Statins are widely prescribed and demonstrated to be highly efficient to prevent cardiovascular diseases [6,22]. However, they are also known to induce skeletal muscle abnormalities ranging from myalgia to myopathy with an incidence of 1–7% [1,23]. If myopathy is not recognized and treatment maintained, necrosis of muscle and rhabdomyolysis may occur with an incidence of 0.04–0.2% [24,25]. We recently reported that acute application of simvastatin on human skeletal muscle fibers induced a mitochondrialdependent impairment of Ca2+ homeostasis [13]. This might contribute to the deleterious effects of statins such as calpaı¨n activation and apoptosis as recently reported [26]. In this study, we further investigated the cellular effect of simvastatin and observed that this adverse effect: (1) is specific of skeletal muscle and independent of muscle phenotype, (2) is linked to MCT4 present in skeletal muscle while absent in cardiac tissue, and (3) results primarily from an alteration of the mitochondrial respiratory chain mainly focused on an inhibition of complex I (i.e., NADH dehydrogenase). As previously reported [13], simvastatin triggers Ca2+ efflux by both NCE and PTP responsible for a SR-Ca2+ overload subsequently followed by a large SR-dependent Ca2+ release. Mitochondrial calcium efflux is associated with depolarization of the inner membrane of the mitochondria and increased NADH autofluorescence, suggesting alteration of mitochondrial energetic function by simvastatin. The present study demonstrates that Ca2+ efflux through PTP and NCE is a consequence of mitochondrial membrane potential fall observed after acute application of simvastatin. Indeed, inner membrane depolarization occurs even in the absence of Ca2+ efflux through PTP and NCE. Since proton flux through respiratory chain is the precursor of the inner mitochondrial membrane potential, the origin of simvastatin deleterious effects could be linked to an altered mitochondrial respiratory chain and mitochondrial energetic function. Oxymetric measurements of mitochondrial respiration clearly demonstrate that simvastatin alters mitochondrial energetic function at the origin of the inner membrane potential fall independently of the metabolic profile. In order to further determine the origin of simvastatininduced impairment of mitochondrial respiration, we assess simvastatin effects using different substrates and inhibitors of respiratory chain complexes. Using palmitoyl-carnitine/malate, or pyruvate/malate as substrates, we demonstrated that simvastatin potentially affects in the same range lipidic and glucidic energetic pathways. Thus, simvastatin affects commune downstream targets of these pathways, namely Krebs cycle or more probably respiratory chain. We used FCCP to uncouple oxidative phosphorylation and dissociate complex V from the others. Although we cannot totally exclude a direct effect of simvastatin on complex V of respiratory chain (ATP synthase), the strong inhibition of respiration rate in these conditions
P. Sirvent et al. / Biochemical and Biophysical Research Communications 338 (2005) 1426–1434
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Fig. 6. Simvastatin-induced alteration in Ca homeostasis. x–y fluorescence images recorded in bundles of skinned human skeletal muscle fibers (A), in skinned rat EDL (B), and soleus (C) loaded with Fluo-3 (50 lM). In each image, average fluorescence is measured in a region of interest and reported as a function of time. Acute application of simvastatin resulted in a delayed (10 min) increase in [Ca2+]i independently of muscle phenotype.
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Fig. 7. Effect of simvastatin on ventricular cardiomyocytes. x–y fluorescence images recorded in intact (A) and saponin-skinned (B) rat ventricular myocytes and in intact rat FDB enzymatically dissociated fibers in the absence (C) or in the presence on 100 lM MCT4 inhibitor PCMBS (D). Intact fibers were loaded with Fluo-3 AM (5 lM in the external medium for 30 min) and permeabilized cells were incubated with 50 lM Fluo-3 pentapotassium salt in the internal medium. In each image, fluorescence is measured in a region of interest and reported as a function of time. Simvastatin was used at the concentration of 200 lM. Intact cardiomyocytes exhibited spontaneous Ca2+ transients that were independent of the presence of simvastatin.
supports the contention that a major target of simvastatin is focused on the four first complexes of the respiratory chain. As a matter of fact, our data indicated that complex I is the main complex affected by simvastatin. The larger simvastatin effect on respiration rate occurs when using glutamate/malate as substrates No significant effects could be observed with succinate and rotenone, strongly supporting a direct effect of simvastatin on complex I, while complexes II seem to be unaffected. This is also in agreement with the simvastatin-induced increase in NADH auto fluo-
rescence observed previously [13]. Moreover, using TMPD and ascorbate as substrate, we could notice a weak inhibition by simvastatin of complex IV. Mazat et al. [27] have shown that limited inhibition of complex IV as observed here did not affect global respiration rate. Based on these observations they concluded that complex IV is not a limiting step of respiratory chain. Considering the low amplitude of the simvastatin-induced inhibition of this complex, we assume that this effect would not participate predominantly to the global inhibition of respiratory chain induced
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by simvastatin. The mechanism by which simvastatin inhibits complex I remains to be elucidated. However, one can postulate that the interaction of simvastatin with this complex might block the transfer of electrons within the complex or between complex I and II. Skeletal and heart muscles shear a lot of homologies in regard to their metabolism and regulation of Ca2+ homeostasis and contractile function. Nevertheless, deleterious effects of statins were to this date only reported in skeletal muscles. As a matter of fact, it has been recently reported that administration (40 mg/kg/day) of pravastatin for 3 weeks in mice induced marked ultrastructure alterations of mitochondria in skeletal muscle while the ultrastructure of the heart and liver of the treated animals appeared normal [28]. In the present work, we further sought to test this skeletal muscle specificity of statinsÕ deleterious effects. Therefore, acute applications of simvastatin on both intact and permeabilized isolated rat cardiomyocytes have been performed. While simvastatin was unable to alter Ca2+ homeostasis in intact cardiomyocytes, one could observe on permeabilized cells a large release of Ca2+ preceded by a slight elevation in cytosolic Ca2+. As in skeletal muscle fibers [13], this first phase was correlated with a mitochondrial membrane depolarization. A similar phenomenon has already been observed in heart cells with DCP, an inhibitor of mitochondrial respiration [29]. Acute application of DCP on heart cells induced a mitochondrial membrane potential fall, a release of Ca2+ by mitochondria, and a delayed large increase in [Ca2+]i comparable with the effects observed in our study. Thus, simvastatin presents a similar potential to alter both skeletal muscle and heart cellsÕ internal machinery. It has been shown that simvastatin diffuses in kidney cells through the monocarboxylate transporter MCT4. This isoform is abundant in skeletal muscle but absent in heart tissue [30]. Although other transporters such as organic anion transporter have been proposed for potential statin uptake [31], their relative contribution compared to MCTs is not clear. Inhibition of MCT4 abolished simvastatin-induced alteration in Ca2+ homeostasis in intact skeletal muscles, suggesting a predominant participation of MCT4 in statin uptake. This could account for the difference observed in intact skeletal muscle and heart cells with acute applications of simvastatin. We cannot however totally exclude that simvastatin diffuses into cardiomyocytes during chronic treatment via other pathways. Functional differences between heart and skeletal muscles might also account for this specificity. Among them, one can notice the predominant role of sodium/calcium exchanger (NCX) in heart cells to control Ca2+ homeostasis. While skeletal muscle has a high capacity to reuptake its intracellular Ca2+ to the SR, cardiomyocytes tend to avoid cytoplasmic and sarcoplasmic reticulum Ca2+-overload by extruding Ca2+ through NCX [32]. In conclusion, we demonstrate here that a specific alteration of complex I of respiratory chain represents the early step of a cascade of cellular deleterious mechanisms induced by simvastatin. This effect can be seen on both
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slow and fast type skeletal muscles, but not on intact heart cells. Cardiomyocytes appear to be protected by their lack of MCT4 which mediates statin uptake in skeletal muscle. The effects described here in vitro could easily account for the adverse effects observed in vivo in statin-treated patients. It would be important to further elucidate the myotoxicity induced by statins and interesting to confirm this demonstration with muscle biopsies from statin-treated patients and suffering from myalgia. Acknowledgments This study was supported by grants from INSERM, the Association Franc¸aise contre les Myopathies (AFM), the Fondation pour la Recherche Me´dicale (FRM), and the Agence Franc¸aise de Se´curite´ Sanitaire des Produits de Sante´ (AFSSAPS). References [1] D.J. Maron, S. Fazio, M.F. Linton, Current perspectives on statins, Circulation 101 (2000) 207–213. [2] C.J. Vaughan, A.M. Gotto Jr., Update on statins: 2003, Circulation 110 (2004) 886–892. [3] J.A. Tobert, G.D. Bell, J. Birtwell, I. James, W.R. Kukovetz, J.S. Pryor, A. Buntinx, I.B. Holmes, Y.S. Chao, J.A. Bolognese, Cholesterol-lowering effect of mevinolin, an inhibitor of 3-hydroxy-3methylglutaryl-coenzyme a reductase, in healthy volunteers, J. Clin. Invest. 69 (1982) 913–919. [4] Design and baseline results of the Scandinavian Simvastatin Survival Study of patients with stable angina and/or previous myocardial infarction, Am. J. Cardiol. 71 (1993) 393–400. [5] F.M. Sacks, M.A. Pfeffer, L.A. Moye, J.L. Rouleau, J.D. Rutherford, T.G. Cole, L. Brown, J.W. Warnica, J.M. Arnold, C.C. Wun, B.R. Davis, E. Braunwald, The effect of pravastatin on coronary events after myocardial infarction in patients with average cholesterol levels. Cholesterol and Recurrent Events Trial investigators, N. Engl. J. Med. 335 (1996) 1001–1009. [6] Prevention of cardiovascular events and death with pravastatin in patients with coronary heart disease and a broad range of initial cholesterol levels. The Long-Term Intervention with Pravastatin in Ischaemic Disease (LIPID) Study Group, N. Engl. J. Med. 339 (1998) 1349–1357. [7] M. Takemoto, K. Node, H. Nakagami, Y. Liao, M. Grimm, Y. Takemoto, M. Kitakaze, J.K. Liao, Statins as antioxidant therapy for preventing cardiac myocyte hypertrophy, J. Clin. Invest. 108 (2001) 1429–1437. [8] J.K. Liao, Isoprenoids as mediators of the biological effects of statins, J. Clin. Invest. 110 (2002) 285–288. [9] J. Davignon, Beneficial cardiovascular pleiotropic effects of statins, Circulation 109 (2004) III39–III43. [10] J.P. Halcox, J.E. Deanfield, Beyond the laboratory: clinical implications for statin pleiotropy, Circulation 109 (2004) II42–II48. [11] M. Ucar, T. Mjorndal, R. Dahlqvist, HMG-CoA reductase inhibitors and myotoxicity, Drug Safety 22 (2000) 441–457. [12] S. Bellosta, R. Paoletti, A. Corsini, Safety of statins: focus on clinical pharmacokinetics and drug interactions, Circulation 109 (2004) III50–III57. [13] P. Sirvent, J. Mercier, G. Vassort, A. Lacampagne, Simvastatin triggers mitochondria-induced Ca2+ signaling alteration in skeletal muscle, Biochem. Biophys. Res. Commun. 329 (2005) 1067–1075. [14] O. Cazorla, A. Lacampagne, J. Fauconnier, G. Vassort, SR33805, a Ca2+ antagonist with length-dependent Ca2+-sensitizing properties in cardiac myocytes, Br. J. Pharmacol. 139 (2003) 99–108.
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