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CELLULAR AND MOLECULAR
Unità di Farmacologia, Dipartimento Farmacobiologico, Facoltà di Farmacia, Università di Bari, Bari, Italy
Received December 7, 2006; accepted February 6, 2007.
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Abstract |
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). The different statins currently available (simvastatin, atorvastatin, lovastatin, pravastatin, fluvastatin, rosuvastatin) are generally well tolerated in patients. However, side effects may arise in skeletal muscle, ranging from transient increase in creatine kinase, muscle pain, and cramps to myositis and potentially life-threatening rhabdomyolysis (Bellosta et al., 2004; Rosenson, 2004). Despite the numerous studies describing myopathy in animals and humans, the molecular mechanism of statin-induced myotoxicity has not been completely elucidated. A variety of hypotheses have been formulated to explain such toxicity, including impairment of glycoprotein synthesis in the muscle membrane as well as reduction of ubiquinone concentration in mitochondria, causing severe deficits in energy metabolism (Evans and Rees, 2002).
Interestingly, it was also proposed that an alteration of structures involved in Ca2+ homeostasis could play a pivotal role in producing myocyte injury. After 2 to 3 months of chronic treatment of rats with simvastatin, the voltage threshold for contraction (mechanical threshold, MT), a calcium-sensitive index of excitation-contraction coupling, was shifted toward more negative potentials in fast-twitch muscle fibers (Pierno et al., 1999), an effect that is compatible with an increase of resting cytosolic calcium concentration ([Ca2+]i). Moreover, in vitro studies showed a statin-induced elevation of [Ca2+]i and suggested a possible interference of the drug with intracellular Ca2+ stores (Nakahara et al., 1994; Inoue et al., 2003; Sirvent et al., 2005), which in turn might be responsible of cell damages via activation of Ca2+-dependent proteolytic enzymes (Sacher et al., 2005). It is noteworthy that all statin-induced effects on skeletal muscle are strictly dependent on their lipophilicity, showing highly hydrophilic pravastatin, and no muscle toxicity, even at high doses (Nakahara et al., 1994; Pierno et al., 1995, 1999). Recently, a multidisciplinary study aimed to the identification of the cellular and tissue targets of a chronic treatment with newer powerful and lipophilic statins, such as fluvastatin and atorvastatin, on rat skeletal muscle was conducted (Pierno et al., 2006). Both statins resulted more potently than simvastatin in producing skeletal muscle injury. Indeed, although no histological changes were observed on muscle fibers, both statin treatments produced a significant negative shift of the potentials of MT at which fibers contracted. In addition, statins altered muscle excitability by reducing resting chloride conductance (gCl), the electrical parameter sustained by the voltage-gated chloride channel CLC-1 critical for the maintenance of membrane stability. Because CLC-1 is negatively regulated by the Ca2+-dependent protein kinase C (De Luca et al., 1998; Rosenbohm et al., 1999), the observed reduction of gCl supports the hypothesis that statin administration could effectively interfere with calcium-handling mechanism.
Based on the above findings, we performed a 2-month in vivo administration with fluvastatin and atorvastatin to rats and evaluated ex vivo the effect of drug treatment on resting [Ca2+]i by fura-2 cytofluorimetry on tendon to tendon intact fibers of extensor digitorum longus (EDL) muscle. In parallel, to further elucidate the statin-induced Ca2+ release mechanism, we characterized the statin cellular effects after acute in vitro application.
Our data showed that both statins caused a sustained increase in cytosolic Ca2+ levels by interfering with intracellular stores, such as mitochondria and sarcoplasmic reticulum. Considering the crucial role of resting calcium ions in skeletal muscle function and maintenance, our findings contribute to a better understanding of the mechanism responsible for cramps, myalgia, and other muscular side effects induced by this widely used class of hypolipidemic drugs.
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Materials and Methods |
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). The doses of drugs were chosen based on data present in the literature regarding human and rodents, as reported previously (Pierno et al., 2006). Fluvastatin and atorvastatin dissolved in CMC (0.5%) suspension were administered orally via an esophageal cannula once a day for 2 months. For each rat, the weight-related dose was formulated so that the maximal volume of drug-containing suspension was 1 ml. Because the results obtained from the two control groups (untreated and CMC-treated rats) were similar, we have combined and showed data as unique control.
In Vivo Study: Determination of the Forelimb Muscle Strength. The forelimb muscle force of control and treated rats was evaluated before and every week until the end of the treatment by means of a grip strength meter (Columbus Instruments, Columbus, OH). Five rats per group (five rats treated with 5 mg/kg fluvastatin, five rats treated with 20 mg/kg fluvastatin, five rats treated with 5 mg/kg atorvastatin, five rats treated with 10 mg/kg atorvastatin, and five control rats) were analyzed. For this measure, the rats were allowed to grasp a triangular ring connected to a force transducer and then gently pulled away until the grip was broken. The transducer saved the force value at this point as a measure of the maximal resistance; the animal can develop with its forelimbs (De Luca et al., 2003). Five measures (one to five) were taken from each animal within 3 min, and each value was normalized with respect to the weight of the animal. We calculated the difference between the fifth measure and the first measure in each animal as a muscle fatigability index. The mean values (±S.E.M.) of this difference of control and of treated rats were then analyzed for significance (by ANOVA test).
Dissection of Native Muscle Fibers. For ex vivo and in vitro experiments, EDL muscles were removed from the animal under deep urethane anesthesia (1.2 g/kg body weight). Soon after the surgery, the rats that were still anesthetized were euthanized by anesthetic overdose. EDL muscles of the different groups of treated rats and of control rats were pinned in a dissecting dish containing 95% O2/5% CO2-gassed normal physiological solution at room temperature (22°C) for further dissection. Small bundles of 10 to 15 fibers arranged in a single layer were dissected lengthwise, tendon to tendon, with the use of microscissors, as described elsewhere (Fraysse et al., 2003).
Fura-2 Fluorescence Measurements in Intact Muscle Fibers. Calcium measurements were performed using the membranepermeant Ca2+ indicator fura-2 acetoxymethyl ester (Invitrogen, Pero, Milano, Italy). Loading of muscle fibers was performed for 2 h at 25°C in normal physiological solution containing 5 µM fura-2 acetoxymethyl ester mixed to 0.05% (v/v) Pluronic F-127 (Invitrogen). After loading, muscle fibers were washed with normal physiological solution and mounted in a modified RC-27NE experimental chamber (Warner Instrument Inc., Hamden, CT) on the stage of an inverted Eclipse TE300 microscope (Nikon, Tokyo, Japan) with a 40x Plan-Fluor objective (Nikon). The mean sarcomere length was set to 2.5 to 2.7 µm. Fluorescence measurements were made using a QuantiCell 900 integrated imaging system (Visitech International Ltd, Sunderland, UK) as described previously (Fraysse et al., 2003, 2004).
During experiments, pairs of background subtracted images of fura-2 fluorescence (510 nm) after excitation at 340 and 380 nm were acquired, and ratiometric images (340/380 nm) were calculated for each muscle fiber of the preparation using QC2000 software. Subsequently fluorescence ratio values were converted to the resting cytosolic calcium [Ca2+] (in nanomolars) after a calibration procedure using the equation: [Ca2+]i = (R–Rmin)/(Rmax–R) x KD x 
, where R is the ratio of fluorescence excited at 340 nm to that excited at 380 nm; KD is affinity constant of fura-2 for calcium, which was taken as 145 nM (Molecular Probes); and is a parameter according to Grynkiewicz et al. (1985) and was determined experimentally in situ in ionomycin-permeabilized muscle fibers as described previously (Fraysse et al., 2003). Rmin and Rmax were determined in muscle fibers incubated in Ca2+-free normal physiological solution containing 10 mM EGTA and in normal physiological solution, respectively.
Determination of Sarcolemmal Permeability to Divalent Cations. The manganese quench technique was used to estimate the sarcolemmal permeability to divalent cations. Mn2+ enters via the same routes as Ca2+ but accumulates inside the cell. As Mn2+ quenches the fluorescence of fura-2, the reduction of the intensity of fura-2 fluorescence can be used as an indicator of the time integral of Mn2+ influx (Parekh and Penner, 1997). Muscle preparations were perfused for 2 min with normal physiological solution containing 0.5 mM Mn2+ as a surrogate of CaCl2 (quenching solution). To measure the Mn2+ influx through the store-operated Ca2+ channels, a proper protocol for SOCE activation was used (Kurebayashi and Ogawa, 2001; Zhao et al., 2005, 2006). Particularly, the fibers were perfused with 10 µM thapsigargin and 40 mM caffeine for 15 min in zero Ca2+ extracellular solution to induce sarcoplasmic reticulum (SR) Ca2+ depletion; successively, 0.5 mM Mn2+ was added to extracellular solution. During the quenching protocol, the fluorescence of fura-2 excited at 360 nm was acquired at 1 Hz.
MT for Contraction. Soon after the removal from the rat, EDL muscles tied at the end of each tendon were placed on a glass rod located in a 25-ml muscle bath chamber maintained at 30°C and continuously perfused with 95% O2/5% CO2-gassed normal physiological solution. The MT for contraction was determined using a two-microelectrode point voltage-clamp method in the presence of 3 µM tetrodotoxin, as described previously (Pierno et al., 2006). Depolarizing current pulses of increasing duration (5–500 ms) were given repetitively at a rate of 0.3 Hz, whereas the impaled fibers were viewed continuously with a stereomicroscope (100x magnification). The command voltage was increased until contraction was just visible and the threshold membrane potential at this point was read from a digital sample-and-hold voltmeter. The mean threshold membrane potential (millivolt) ± S.E.M. (n fibers) was plotted as a function of the pulse duration t (millisecond), and the relationship was fit using the equation V(t) = [H–R x exp(–t/
R)]/[1–exp(–t/R)], where H is the holding potential (–90 mV), R (millivolt) is the rheobase voltage, and R (millisecond) is the time constant to reach R. The MT values were expressed as the fitted R parameter along with the mean ±S.E. that was determined from the variance-covariance matrix in the nonlinear least-squares fitting algorithm.
Solution and Chemical Compounds. The normal physiological solution contained 148 mM NaCl, 4.5 mM KCl, 2.5 mM CaCl2,1mM MgCl2, 0.44 mM NaH2PO4, 12 mM NaHCO3, and 5.5 mM glucose. The pH of all solutions was adjusted to 7.3 to 7.4 by bubbling them with 95% O2/5% CO2. The calcium free-solution had the same composition of the normal physiological solution, with the exception that CaCl2 was omitted and 10 mM EGTA was added. The quenching solution had the same composition, with the exception that 0.5 mM MnCl2 was substituted for CaCl2. All chemicals cited above and ionomycin, caffeine, ruthenium red, cyclosporine A, and thapsigargin were purchased from Sigma (St. Louis, MO).
Commercially available fluvastatin (Lescol, fluvastatin sodium, A.I.C. number 029163021; Novartis Novartis, Basel, Switzerland) and atorvastatin (Torvast, atorvastatin calcium, A.I.C. number 033007042; Pfizer Ireland Pharmaceuticals, County Cork, Ireland) in CMC (0.5%) suspension were used for in vivo chronic treatment to rats. For in vitro experiments, fluvastatin sodium salt purchased from Calbiochem was used.
Statistical Analysis. The ANOVA followed by Bonferroni's t test was used to evaluate statistical differences between treated and control groups. For the in vitro studies, significance levels were calculated using Student's unpaired t test.
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Results |
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; Dulhunty, 1992), we first investigated the effect of the in vivo statin chronic treatment on the gross morphologic fibers aspect and on the excitation-calcium release coupling. To this purpose, we examined skeletal muscle fibers samples by visible light microscopy and exposed them to a depolarizing high potassium solution. As illustrated in Fig. 2 (top), there was no light microscopic changes in EDL muscle fibers of rats treated with 5 mg kg–1 day–1 fluvastatin and atorvastatin at both dosages compared with control rats. In contrast, fluvastatin-related muscle of rats treated at the dose of 20 mg kg–1 day–1 showed a compromised muscle fiber integrity characterized by a remarkable collapse and disarray of sarcomeres. This structural change was accompanied by the alteration of the voltage-dependent Ca2+ release. Indeed, the amplitude of the Ca2+ transient induced by 100 mM K+-depolarizing solution showed by this group of treated animals was reduced by approximately 55% with respect to the control rats (Fig. 2, bottom).
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Measurements of resting [Ca2+]i. In line with previous studies (Fraysse et al., 2003, 2006), EDL fast-twitch skeletal muscle fibers of adult control rats showed a value of resting [Ca2+]i of 26.9 ± 0.8 nM. Both in vivo fluvastatin and atorvastatin administration induced a significant increase of [Ca2+]i in all of the treated groups, although the effect was more important for animals treated with 20 mg kg–1 day–1 fluvastatin (Fig. 3). Indeed, the resting [Ca2+]i was 31 ± 0.8 and 43 ± 2.3 nM in rats treated with 5 mg kg–1 day–1 fluvastatin and 20 mg kg–1 day–1 fluvastatin, respectively. In the case of atorvastatin, the maximal effect was already observed with the lower dose, reaching a 20% increase of [Ca2+]i.
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). Because the maximal amplitude value of caffeine-induced calcium transient in rat EDL intact muscle fibers was obtained by application of 40 mM caffeine (Fraysse et al., 2003), we determined the caffeine responsiveness of the different animal groups at this dose.
Compared with control rats, a slight amplitude reduction of the calcium transient induced by 40 mM caffeine was observed after treatment with 5 mg kg–1 day–1 fluvastatin and atorvastatin at both doses. The lowering of the responsiveness to caffeine was more marked in 20 mg kg–1 day–1 fluvastatin-treated rats, showing a 80% reduction of the calcium transient amplitude (Fig. 4B).
Effect of in Vitro Application of Fluvastatin on Calcium Homeostasis of Skeletal Muscle Fibers. To gain insight into the cellular mechanism of statin-induced calcium handling alteration, we evaluated the drug effect after in vitro acute application on rat skeletal muscle fibers. Based on the results obtained with the in vivo statins administration, we focused our attention on fluvastatin that produced the most evident alteration of the calcium homeostasis in treated rat muscle fibers.
Mechanical threshold and resting [Ca2+]i. We previously demonstrated that muscle fibers from fluvastatin-treated rats needed less depolarization to contract showing a rheobase voltage (R) shift of 3 to 4 mV toward more negative potentials compared with control rats (Pierno et al., 2006). Accordingly, the in vitro application of fluvastatin on EDL muscle produced a negative shift of the voltage threshold for contraction. Particularly, the effect was observed already after application of 50 µM fluvastatin that shifted R of approximately 2 mV toward a more negative potential with respect to the control value (data not shown) and became statistically significant at higher concentration. Indeed, as shown in Fig. 5, 200 µM fluvastatin produced a marked shift of strength-duration curve at each pulse duration. The R value obtained by the fit of the experimental points were –66.0 ± 0.15 and –69.5 ± 0.25 mV in the absence and presence of 200 µM fluvastatin, respectively.
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200 µM. For the dosage below 200 µM, fluvastatin effect was characterized by only a slow-rising phase responsible for a slight increase of [Ca2+]i. Therefore, in further investigations, we routinely applied fluvastatin at 200 µM.
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Mn2+ quench rate. In agreement with the reduction of the cation membrane permeability observed in 20 mg kg–1 day–1 fluvastatin-treated animals, in vitro application of fluvastatin reduced fluorescence quench. Particularly, the mean quench rate was 3.9 ± 0.15 and 2.89 ± 0.16% min–1 before and after application of 200 µM fluvastatin, respectively.
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; Zhao et al., 2005, 2006). Thus, a reduction of SOCE activity could account for the reduced caffeine or potassium-induced calcium transient observed in muscle fibers after in vivo treatment with 20 mg kg–1 day–1 fluvastatin.
To confirm this hypothesis, we evaluated the effect of fluvastatin on Mn2+ permeability after SOCE activation. To this end, SR Ca2+ stores of muscle fibers were depleted by treatment with thapsigargin and caffeine (Kurebayashi and Ogawa, 2001; Zhao et al., 2006; Liantonio et al., 2007).
As shown in Fig. 7A, activation of SOCE induced by SR Ca2+ depletion led to a steep Mn2+ quenching of fura-2 fluorescence. Interestingly, we found that the preincubation of muscle fibers with 200 µM fluvastatin significantly reduced the rate of Mn2+ entry following depletion of SR Ca2+ store (Fig. 7A). Indeed, the mean quench rate resulted in 9.1 ± 0.15 (n = 15) and 5.1 ± 0.12% min–1 (n = 13) in nontreated and fluvastatin-treated muscle fibers, respectively.
Investigation of the source of calcium release. Removal of external Ca2+ in the bath solution did not abolish the fluvastatin-induced [Ca2+]i rise, producing an increase of 108 ± 22 nM, a value not significantly different from that obtained in the presence of extracellular calcium (Fig. 7B). Together with the lack of Mn2+ permeability enhancement after fluvastatin in vitro application (see above), this result strongly suggested that fluvastatin induced a [Ca2+]i increase probably by favoring a calcium release from intracellular stores. Thus, we attempted to identify the relevant intracellular source. We investigated the possible involvement of SR using ruthenium red, an inhibitor of the SR Ca2+ release channels (RyR). Pretreatment with ruthenium red, at a concentration (5 µM) that completely prevented the 40 mM caffeine effect (Liantonio et al., 2007), significantly reduced the fluvastatin-induced [Ca2+]i increase, although a [Ca2+]i increase of 22.5 ± 1.2 nM was still observed. (Figs. 6A and 7B). These data indicated that fluvastatin mobilized Ca2+ mainly from SR but that another intracellular calcium store was involved. It is well known that mitochondria play a role in skeletal muscle [Ca2+]i (Saris and Carafoli, 2005). To assess a possible mitochondrial origin of the fluvastatin-induced [Ca2+]i increase, we evaluated the effect of such statin in the presence of CsA, an inhibitor of the mitochondrial permeability transition pore (PTP) (Crompton et al., 1999; Liantonio et al., 2007).
In agreement with previous study (Liantonio et al., 2007), the application of 2 µM CsA did not alter the baseline of [Ca2+]i (data not shown). Successive addition of 200 µM fluvastatin failed to produce any [Ca2+]i increase (Fig. 7B). Thus, the inhibition of the mitochondrial PTP completely abolished the fluvastatin-induced [Ca2+]i increase.
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Discussion |
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), and an alteration of calcium homeostasis frequently results in muscle damage. Accordingly, elevated resting [Ca2+]i levels were found in pathophysiological conditions of skeletal muscle, such as ageing (Navarro et al., 2001; Fraysse et al., 2006) or muscular dystrophy (Fraysse et al., 2004; De Luca et al., 2003). In the present study, we investigated the potential relationship between statin myotoxicity and calcium homeostasis by assessing the effects of a chronic treatment with fluvastatin and atorvastatin on fura-2-loaded rat intact skeletal muscle fibers.
Either 5 mg kg–1 day–1 fluvastatin or atorvastatin at both doses produced a significant increase in [Ca2+]i, with a concomitant decrease of the caffeine responsiveness and no changes in sarcolemmal calcium permeability. These findings clearly indicated a drug capability of altering calcium homeostasis by interfering with intracellular targets involved with calcium handling mechanisms. It is noteworthy that 20 mg kg–1 day–1 fluvastatin-treated animals, in addition to a marked increase of [Ca2+]i, showed a completely compromised sarcomere organization and a significant decrease of the depolarization-induced intracellular calcium peak. It might be possible that, at this high drug dosage together with the disruption of calcium homeostasis, a series of other cellular mechanisms may take place, all events accounting for the described detrimental effects. An interesting hypothesis could be that fluvastatin administration would produce an alteration of the T-tubule membrane composition. Indeed, the T-tubule membrane system of striated muscle cells was highly enriched in cholesterol (Rosemblatt et al., 1981), and an alteration of the amount of this component caused a reduction of the depolarization-induced muscle tension (Launikonis and Stephenson, 2001). In support of this hypothesis, it was recently reported that skeletal muscle biopsies from statin-treated patients are characterized by a breakdown of the T-tubular system and by subsarcolemmal ruptures (Draeger et al., 2006).
The degree of calcium homeostasis impairment induced by the statin administration well correlated with the in vivo muscle function performance alteration. Indeed, rats treated with 20 mg kg–1 day–1 fluvastatin showed a reduced force production after prolonged activity, thus indicating a fatigue-related decrease in muscle function of this animal group with respect to control rats.
To define the statin cellular mechanism of action, leading to calcium homeostasis alteration, an in vitro study was conducted. It is noteworthy that micromolar acute application of fluvastatin caused changes of [Ca2+]i, resembling the effect obtained after the in vivo administration. Indeed, fluvastatin was capable of producing a shift of MT for contraction toward more negative potentials and an increase of resting [Ca2+]i. Furthermore, in vitro fluvastatin application reduced the cation membrane permeability at rest, an effect that resulted particularly markedly when SOCE was activated. These results allowed us to explain some of the statin-induced effects observed after in vivo treatment. Indeed, it was recently demonstrated that, in skeletal muscle, SOCE is essential for the maintenance of Ca2+ homeostasis by ensuring the refilling of intracellular calcium storage (Kurebayashi and Ogawa, 2001; Zhao et al., 2005, 2006). The direct statin SOCE inhibition would lead to a chronic depletion of the intracellular stores, thus justifying the drastic reduction of the potassium- and caffeine-induced Ca2+ transients, characterizing the muscle fibers of 20 mg kg–1 day–1 fluvastatin-treated animals. The in vivo and in vitro characterization of the type of the voltage-insensitive permeable calcium current, effectively involved in the SOCE-related effect induced by statins, will be very useful to define the role and the relevance of such phenomenon in mediating the statin-induced myotoxicity.
Accordingly, to quench measurements performed on both in vivo and in vitro statin-treated fibers, the amplitude of the fluvastatin-induced increase of [Ca2+]i did not vary after withdrawal of extracellular calcium, thus strongly indicating that the drug effect on resting [Ca2+]i is not due to an increase of the sarcolemmal cationic permeability but rather to an internal Ca2+ store depletion.
By using appropriate tools, we determined the sequence of the statin-induced Ca2+ release mechanism. Mitochondria appeared as the cellular structure responsible for the earlier event, leading to a sustained [Ca2+]i elevation. Indeed, the partial and the complete lack of effect of fluvastatin observed in the presence of ruthenium red and CsA, respectively, led us to propose that a mitochondrial Ca2+ efflux represents the early event, which in turn caused the subsequent large SR Ca2+ release. It is noteworthy that our findings represent the first direct evidence of such mechanism on intact skeletal muscle fibers where the e-c coupling system functionality is ensured and are in agreement with previous studies performed on cultured myoblasts as well as on skinned fibers (Nakahara et al., 1994; Inoue et al., 2003; Sirvent et al., 2005).
To support direct activity of fluvastatin on mitochondria, it was reported that various lipophilic statins produced alterations in mitochondrial function associated with dissipation of the electric potential across the inner mitochondrial membrane (Kaufmann et al., 2006; Velho et al., 2006), an event that could account for the opening of PTP observed in our investigations. Furthermore, tight structural association between the SR and mitochondria has been described in many cell types (Rizzuto et al., 1998), and recently, a close functional connection between the two cellular structures has been reported in adult fast- and slow-twitch skeletal muscle (Shkryl and Shirokova, 2006). The statin-induced mitochondrial Ca2+ efflux, albeit of small entity, might be highly localized near SR microdomains in close contact with mitochondria, sufficiently resulting in activating Ca2+-induced Ca2+ release from SR. Nevertheless, we cannot exclude the hypothesis that the Ca2+ movement triggered by the opening of PTP might activate a successive adequate cellular signal, which in turn could cause a direct stimulation of calcium efflux from SR.
It is noteworthy that the in vitro reported mitochondrial calcium efflux could allow us also to explain the alteration of caffeine responsiveness observed after in vivo administration. Indeed, because the RyR channel activity was depressed through direct Ca2+-dependent regulation (Fill and Copello, 2002), the reduced caffeine responsiveness, particularly marked for 20 mg kg–1 day–1 fluvastatin-treated animals, could probably be due to an alteration of these SR calcium release channels, other than SOCE inhibition, consequent to the persistent mitochondria-derived Ca2+ stimulation. Finally, we did not exclude that a reduced electrochemical driving force due to the increased cytosolic calcium amount could account for the limited caffeine effect.
All our data support the finding that statins induced an alteration in Ca2+ homeostasis in skeletal muscle fibers. The effects on Ca2+ signaling elicited by statins on vascular smooth muscle (de Sotomayor et al., 2005) and on endothelial cells (O'Meara and Kinsella, 2004; Heinke et al., 2004) explain some of the beneficial cardiovascular effects of HMG-CoA reductase inhibitors that are not related to their lipid-lowering properties. In contrast, such an effect on skeletal muscle could cause various cell damages strongly contributing to the pathogenesis of myopathy induced by statins. The most probable way that elevation in free cytosolic Ca2+ acts to disrupt excitation-contraction coupling in skeletal muscle is via stimulation of certain Ca2+-activated enzymes, such as calpains, phospholipase A2, or protein kinase C. In this regard, it will be highly interesting to investigate whether the statin-induced increase in resting [Ca2+]i may contribute to the decrease of gCl (Pierno et al., 1995, 2006) through a calcium-dependent protein kinase C-dependent CLC-1 inhibition. This may allow a better understanding of the cross-talk mechanism between the two main early targets, cytosolic calcium level and macroscopic gCl, most probably contributing to statin-induced myotoxicity (Pierno et al., 2006).
In conclusion, herein we give the first evidence that an increase of [Ca2+]i in intact tendon-to-tendon rat skeletal muscle fibers can be obtained upon in vivo treatment with fluvastatin or atorvastatin. This effect was attributable to a Ca2+ release from SR and mitochondria. Considering the crucial role of resting calcium ions in skeletal muscle function and maintenance, our findings considerably contribute to the understanding of the myopathic mechanism induced by this widely used class of hypolipidemic drugs.
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Acknowledgements |
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: HMG-CoA, 3-hydroxy-methyl-glutaryl coenzyme A; EDL, extensor digitorum longus; [Ca2+]i, intracellular calcium concentration; MT, mechanical threshold; gCl, resting chloride conductance; SR, sarcoplasmic reticulum; RyR, ryanodine receptor; PTP, permeability transition pore; RR, ruthenium red; CsA, cyclosporine A; SOCE, store-operated calcium entry; ANOVA, analysis of variance.
Address correspondence to: Prof. Diana Conte Camerino, Section of Pharmacology, Dept. of Pharmacobiology, Faculty of Pharmacy, University of Bari, Via Orabona, 4, I-70125 Bari, Italy. E-mail: conte{at}farmbiol.uniba.it
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