OUP user menu


Montserrat Cofán, Josep M. Nicolás, Joaquim Fernández-Solà, Jordi Robert, Esther Tobías, Emilio Sacanella, Ramón Estruch, Alvaro Urbano-Márquez
DOI: http://dx.doi.org/10.1093/alcalc/35.2.134 134-138 First published online: 1 March 2000


Alcohol misuse frequently leads to muscle weakness, which may also occur in the setting of acute and chronic alcoholic myopathies. At the cellular level, ethanol has been found to interfere with signalling mechanisms in cardiac myocytes, skeletal myotubes, and smooth muscle cells. In this study, we focused on the effects of ethanol on the intracellular calcium ([Ca2+]i) transients responsible for excitation–contraction (EC) coupling in isolated mouse skeletal fibres loaded with the fluorescent Ca2+ indicator fura-2. Following electrical stimulation, ethanol caused a significant reversible dose-dependent reduction in [Ca2+]i transient amplitude, already significant at 100 mM ethanol (P = 0.03), without modifying resting [Ca2+]i. Evaluating the potential loci for the effects of ethanol, we indirectly measured sarcolemmal Ca2+ entry by monitoring Mn2+-quenching of intracellular fura-2 via the nitrendipine-sensitive Ca2+ channels during electrical pacing. Ethanol at doses of 20 mM and greater caused a dose-dependent reduction in the rate of fura-2 quenching (all at P < 0.05). Moreover, the intracellular pool of Ca2+ releasable by caffeine was found to be reduced at a minimum of 300 mM ethanol (P = 0.05). We conclude that ethanol reduces the [Ca2+]i transients underlying EC coupling in single mouse skeletal muscle fibres. This acute effect of ethanol was primarily due to an inhibitory effect of ethanol on sarcolemmal Ca2+ influx via voltage-operated Ca2+-channels and, to a lesser extent, to a reduction in the Ca2+ sarcoplasmic reticulum loading state. This inhibitory effect of ethanol may be implicated in the development of muscle weakness with alcohol consumption.


The ergolytic effect of ethanol has long been recognized. Although acute and chronic myopathies are the most usual reasons for muscle weakness in alcoholics, acute intake of ethanol may lead to muscle weakness, without structural muscle damage (Urbano-Márquez et al., 1989; Fernández-Solà et al., 1996). Eichner (1993) observed that, on drinking alcohol, athletes exhibited a decrease in their muscle strength and contraction velocity, which was reversible after ethanol suppression. It is well known that calcium-dependent excitation– contraction (EC) coupling is the main mechanism responsible for skeletal muscle strength. Previous studies in other muscle specimens, such as rat cardiomyocytes (Thomas et al., 1989, 1996; Danziger et al., 1991; Nicolás et al., 1996), cultured myotubes (Cofán et al., 1995; Nicolás et al., 1998), and smooth muscle cells (Zhang et al., 1992), have demonstrated that acute exposure to ethanol interferes with EC coupling in a dose-dependent manner, mainly through a reduction in [Ca2+]i transients. However, no studies have evaluated the effect of ethanol on EC coupling in mammalian mature skeletal muscle fibres. Using fluorescence imaging techniques, we have measured the acute effects of this toxin on the pathways involved in EC coupling, a potential cellular mechanism for ethanol-induced muscle weakness. The results showed that ethanol depressed the [Ca2+]i transients underlying EC coupling in skeletal muscle fibres due, at least in part, to an inhibitory effect of ethanol on voltage-operated Ca2+-channel activity and a reduction in the calcium content at the sarcoplasmic reticulum.


Isolation of muscle fibres and loading

Intact fibres from the flexor digitorum brevis and interosseal muscles of CD-1 mice were isolated by a classical enzymatic dissociation process. Mice were killed by cervical dislocation, muscles were removed and incubated at 37°C for 90 min in Tyrode solution (140 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, pH 7.4 with NaOH) containing collagenase (2 mg/ml) (Sigma, St Louis, MI, USA). They were then rinsed with collagenase-free Tyrode solution and stored up to 6 h at 4°C until used. Intact single fibres were separated from the muscle mass by gentle trituration with the tip of a disposable pipette. Isolated fibres were loaded with 4 μM fura-2 acetoxymethyl ester (Molecular Probes, Eugene, OR, USA) in incubation buffer [121 mM NaCl, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 1.25 mM CaCl2, 10 mM glucose, 5 mM NaHCO3, 10 mM HEPES, 0.01% bovine serum albumin (BSA), pH 7.4 with NaOH] for 25 min at 37°C, with gentle shaking.

Measurement of [Ca2+]i transients and contraction

After loading with fura-2, isolated fibres were washed with incubation buffer and placed on a laminin-coated coverslip in an open flow chamber (2 ml volume) mounted on the stage of a Diaphot-300 inverted epifluorescence microscope (Nikon, Tokyo, Japan). The stage, 20×-glycerine immersion objective (Nikon) and chamber were maintained at 17°C. The chamber was equipped with platinum electrodes to allow electrical field stimulation of the fibres within the field of view, which was applied at 0.25 Hz (5 ms duration) (A385, WPI, Sarasota, FL, USA). The electrical stimulator and excitation filter changer were synchronized with the imaging computer by a prepulse from an interval generator (A300, WPI). The cells were perfused continuously at a flow rate of 7 ml/min with modified incubation buffer containing 2 mM CaCl2.

The imaging system and techniques for obtaining digital images of cellular fluorescence at high-time resolution have been described in detail previously (O'Rourke et al., 1990; Nicolás et al., 1996). In brief, an electrically cooled charge-coupled device (CCD) camera (CH250, Photometrics, Tucson, AZ, USA) was used in a mode that allowed a consecutive series of images to be stored in the photon wells of the CCD chip. To achieve this, the CCD was masked, and the image of the cell was focused on one end of the detector. The image was shifted periodically along the detector, so that a series of subimages was accumulated on the CCD, digitized and stored as a single image in the memory of the imaging computer (MacIntosh 840AV, Apple Inc., Cupertino, CA, USA). To evaluate the contractile cycle of the fibres, series of 8- and 16-ms time-resolved fluorescence images were created by averaging four and two contractions, respectively. Fluorescence images were collected alternately at excitation wavelengths of 340 and 380 nm (10 nm bandwidth filters) to excite the Ca2+-bound and Ca2+-free forms of this ratiometric dye, respectively. The emission wavelength was 510 nm (120 nm bandwidth filter). In order to minimize photobleaching, a computer-controlled shutter was used to limit the exposure of the cells to excitation light. The individual contractions of most fibres were reproducible and image pairs co-registered accurately. [Ca2+]i values were calculated from the 340 to 380 nm ratios, as described previously (Grynkiewicz et al., 1985).

At the end of each experiment, fibres were exposed to butanedione monoxime (20 mM) to prevent fibre shrinkage, and then to digitonin (30 μg/ml) and ethylene glycol-bis (β-aminoethyl ether) N,N,N,N-tetraacetic acid (EGTA) (25 mM) for 10 min (Sigma). This treatment releases the intracellular fura-2 leaving the residual fluorescence at each wavelength due to fibre autofluorescence and any compartmentalized dye. The residual fluorescence was measured over the same region of the fibre as the Ca2+-dependent fluorescence. To ensure that a steady state was achieved, [Ca2+]i was obtained under control conditions and after 10 min incubation with ethanol (20–500 mM; 90–2300 mg/dl), and after a 10-min washout period. Studies with fluorescent dyes indicated a complete chamber buffer exchange within 1 min. Fibres were selected for study based on their overall physical appearance (elongated rods with well-defined sarcomere structure and no blebs), quiescence in the absence of stimuli, and their ability to contract in response to electrical stimulation. Because fibre size exceeded the field of study, only one edge of the fibre was focused and the contractile response was not analysed.

Measurement of sarcolemmal Ca2+channel activity using Mn2+-influx

A potential locus at which ethanol may interact with EC coupling in skeletal muscle cells is at the level of sarcolemmal ion channels. Muscle action potential and contraction are dependent on Ca2+ currents, and it is generally accepted that hormones or drugs that modulate the inward calcium current amplitude affect muscle contraction. To investigate the possible role of ethanol on voltage-dependent Ca2+ entry, we evaluated the effects of ethanol on sarcolemmal Mn2+-influx via the nitrendipine-sensitive Ca2+ channels by measuring the rate of fura-2 quenching by Mn2+ entry when electrical stimulation was applied. Isolated fibres preloaded with fura-2/AM (7 μM) were perfused at a flow rate of 7 ml/min with a modified incubation buffer (130 mM NaCl, 4.7 mM KCl, 0.1 mM KH2PO4, 1.2 mM MgSO4, 0.1 mM CaCl2, 10 mM glucose, and 10 mM HEPES, pH 7.4 with NaOH) at room temperature (22°C). After a perfusion period of 10 min in which electrical stimulation at 0.25 Hz (5 ms duration) was applied and a field of contractile fibres was selected, the system was switched to a modified incubation buffer which contained 0.2 mM MnCl2, and the electrical pacing was stopped. Fluorescence images were collected every 2 s using 360 nm excitation and 510 nm emission filters. After 100 s, during which the basal quench rate was determined, electrical stimulation (1 Hz, 100 ms duration) was initiated in order to activate Mn2+ entry via voltage-sensitive Ca2+ channels. The electrically stimulated Mn2+ quench rate was completely inhibited by nitrendipine (10 μM) (LC Laboratories, Woburn, MA, USA), indicating that it reflects the activity of L-type Ca2+ channels.

Measurements of caffeine releasable Ca2+pool

To evaluate the sarcoplasmic reticulum (SR) Ca2+ loading state, caffeine (15 mM), dissolved in modified incubation buffer without CaCl2 or BSA, was perfused directly onto one fibre for 10 s using a puffer micropipette (6-μm tip diameter) Narishige IM-300 (Tokyo, Japan) at a pressure of 6 psi. Fibres were perfused continuously at a flow rate of 7 ml/min with incubation buffer containing 2 mM CaCl2 at room temperature (22°C). Because the SR Ca2+ content is modified by the stimulation frequency, fibres were stimulated electrically (0.25 Hz, 5 ms), and caffeine was puffed onto the fibres in phase with the ongoing electrical pulse at the time point of the next stimulation pulse. Pairs of images were collected every 2 s, and fluorescence values at each time point were expressed as a ratio to the initial resting fibre fluorescence. This ratio was calibrated in terms of [Ca2+]i using the initial [Ca2+]i value obtained from the 340 to 380 nm fluorescence measurements, as described previously (Renard et al., 1994). Individual fibres acted as their own controls, with the response to caffeine being compared before and after treatment with ethanol.

Statistical analysis

Standard statistical methods from the SPSS Statistical Analysis System V6.1 (SPSS, Chicago, USA) were used. Paired and unpaired Student's t-tests were used to analyse differences. Correlation studies were obtained by Pearson's correlation coefficient and regression analysis. All variables are expressed as means ± SEM, and a level of probability (P) lower than 0.05 was judged as being significant.


Effect of ethanol on [Ca2+]itransients

We evaluated the influence of ethanol on electrically triggered Ca2+ transients in single skeletal fibres that showed repetitive Ca2+ mobilization in response to the electrical field stimulation. In control cells (n = 60), [Ca2+]i increased from a mean resting value of 93 ± 4 nM to a peak of 1121 ± 46 nM approximately 16 ms after the initiation of the electrical stimulus, returning to basal values after 112 to 128 ms of the depolarization. Ca2+ mobilization was inhibited by 90% in fibres pretreated with 10 μM nitrendipine, indicating that the electrically triggered [Ca2+]i transients were dependent on the activity of nitrendipine-sensitive voltage-operated Ca2+ channels. The amplitude of [Ca2+]i transients was significantly reduced under exposure to ethanol at doses of 100 mM and greater, ranging from 11 ± 3% at 100 mM to 22 ± 10 at 500 mM (P < 0.05), whereas the resting [Ca2+]i was unchanged (Table 1). The reductions in [Ca2+]i transients were totally reversible after a 10-min washout period. Low ethanol concentrations (20 and 40 mM) did not exert a significant depressant effect on [Ca2+]i transients. The reduction caused in the [Ca2+]i transients correlated significantly with the dose of ethanol used (r = –0.48, P < 0.001).

View this table:
Table 1.

Effect of ethanol on [Ca2+]i transients (peak over resting values) in isolated mouse skeletal fibres

[Ca2+]i transients (nM)
Values are in mM and are expressed as means ± SEM; n, number of fibres evaluated; *P < 0.05 and **P < 0.01 compared to basal conditions, respectively; †P < 0.05 compared to ethanol exposure.
Control (zero ethanol) (n = 8)954 ± 52963 ± 62971 ± 78
Ethanol 20 mM (n = 10)893 ± 134885 ± 144887 ± 125
Ethanol 40 mM (n = 10)1164 ± 1411140 ± 1341205 ± 182
Ethanol 100 mM (n = 10)1269 ± 2311045 ± 224*1347 ± 255†
Ethanol 300 mM (n = 12)891 ± 140741 ± 95*831 ± 86†
Ethanol 500 mM (n = 10)1011 ± 150631 ± 75**895 ± 120†

Effect of ethanol on the sarcolemmal Ca2+channel activity

The effect of ethanol on the rate and extent of cytosolic fura-2 quenching by extracellular Mn2+ during electrical stimulation was measured in isolated muscle fibres. Fluorescence (F360) data, collected every 2 s for 5 min, showed a very small decline of 3% in the Ca2+-independent fluorescence, due to photobleaching. When 0.2 mM MnCl2 was added to the buffer and electrical stimulation was not applied, F360 decreased by 10 ± 1% over the same period, a magnitude similar to that observed in fibres pretreated with nitrendipine (10 μM) and stimulated electrically. This showed that the basal quench was primarily independent of voltage-operated Ca2+ channels. By contrast, fibres exposed to a voltage-sensitive Ca2+ channel agonist Bay-K 8644 (1 μM) (LC Laboratories, Woburn, MA, USA) exhibited a threefold increase in the electrically induced Mn2+-quench of fura-2, compared with control fibres (P = 0.01).

Ethanol caused a dose-dependent reduction in the rate of Mn2+ quench (r = –0.57, P < 0.001), achieving an inhibition of 62 ± 11% at 500 mM ethanol (Fig. 1). Moreover, the total amount of fura-2 quenched by Mn2+ during the 100 s under electrical stimulation was also reduced, achieving significance at low ethanol concentrations of 20 mM (21 ± 5%, P < 0.001). These data suggest that ethanol may interfere with sarcolemmal Ca2+ channel activity, diminishing Ca2+ entry into the cells.

Fig. 1.

Nitrendipine-sensitive voltage-operated calcium channel activity. Channel activity was evaluated indirectly by measuring the percentage of cellular fura-2 quenched by Mn2+ entry (F360). The increase in the rate of quench caused by the electrical stimulation was compared in control and ethanol-treated fibres. Ethanol caused a dose-dependent reduction in the rate of quench (r = –0.57, P < 0.001), with a significant decrease at ethanol doses of 20 mM and greater. *P < 0.05; **P < 0.001.

Sarcoplasmic reticulum Ca2+loading state

The magnitude of [Ca2+]i transient also depends on the amount of Ca2+ released from the SR. Because the ethanol-induced decline in [Ca2+]i transient could be due to a reduction in the availability of SR Ca2+, we examined the SR Ca2+ content by measuring the caffeine-induced increase in [Ca2+]i. As the increase of [Ca2+]i evoked by caffeine could cause sarcolemmal depolarization and inward currents that might contribute to the [Ca2+]i signal (Renard et al., 1994), caffeine (15 mM) was applied in modified incubation buffer without Ca2+ and BSA. When caffeine was puffed in phase with the ongoing electrical pulse, more than 95% of fibres responded to caffeine, with similar Ca2+ mobilization at both the initial and a second exposure to caffeine 10 min later (753 ± 119 nM and 799 ± 156 nM, respectively). Ethanol at concentrations of up to 100 mM did not affect the caffeine-induced increase of [Ca2+]i (Table 2). Only ethanol at 300 mM and above consistently decreased the magnitude of the caffeine-releasable Ca2+ pool, with a mean decrease of 35 ± 9% of control values for 300 mM ethanol (P = 0.004) (Table 2).

View this table:
Table 2.

Effect of ethanol on caffeine-induced sarcoplasmic reticulum calcium release (peak over resting values) in isolated mouse skeletal muscle fibres

Ca release (nM)
Values are in mM and are expressed as means ± SEM; n, number of fibres evaluated; *P < 0.05 and **P < 0.01 compared to basal conditions respectively.
Control (zero ethanol) (n = 14)753 ± 119799 ± 196
Ethanol 20 mM (n = 8)722 ± 127827 ± 246
Ethanol 40 mM (n = 9)932 ± 260716 ± 266
Ethanol 100 mM (n = 10)744 ± 260780 ± 185
Ethanol 300 mM (n = 10)790 ± 136559 ± 127*
Ethanol 500 mM (n = 8)762 ± 109465 ± 127**


The present study supports the hypothesis that ethanol interferes with EC coupling by decreasing the availability of Ca2+ needed to generate proper [Ca2+]i transients. Therefore, it is feasible that muscle weakness observed in very drunk patients as well as in the setting of acute and chronic alcoholic myopathies (Urbano-Márquez et al., 1989), may to some extent be mediated by the deleterious effect of ethanol on EC coupling. Acute ethanol may cause muscle weakness in the absence of histological evidence of myopathy (Fernández-Solà et al., 1996). In this sense, it is well known that the ergolytic effect of ethanol in athletes (Eichner, 1993) is more evident in middle distance running than in sprinting (McNaughton and Preece, 1986). Although acute exercise may lead to enzyme leakage as a result of damage to muscle fibres, Clarkson and Reichsman (1990) reported that ethanol did not cause any significantly greater increase in the plasma levels of muscle enzymes during exertion.

No recent study has evaluated the effect of ethanol on EC coupling in muscle fibres from mammalian origin. Preliminary studies with frog muscle preparations showed that alcohol reduced the contractile response and modified [Ca2+]i (Pape and Baylor, 1985; Oz and Frank, 1995). In stimulated mammalian whole-muscle specimens, ethanol exerted a marked decrease in the contractility at concentrations as low as 21.7 mM (Taylor et al., 1992; Pagala et al., 1995). However, the effect of ethanol on EC coupling at the level of mature skeletal muscle fibres of mammalian origin has not been studied. For this reason, we isolated mouse muscle fibres and loaded them with a fluorescent indicator, in order to assess the Ca2+-dependent steps of EC coupling. Ca2+ mobilization values obtained and the kinetics of the [Ca2+]i transients under control conditions were comparable to those reported in the literature when using similar models (Vergara et al., 1991; Imbert et al., 1995).

Similarly to other striated muscle specimens, we observed that ethanol acutely reversibly decreased [Ca2+]i transients in a dose-dependent manner (Thomas et al., 1989; Danziger et al., 1991; Nicolás et al., 1996), an effect that is mediated primarily by an inhibitory effect of ethanol on sarcolemmal voltage-operated Ca2+ channel activity (Takeda et al., 1984; Habuchi et al., 1995) and by reducing the amount of releasable Ca2+ available for EC coupling (Onishi et al., 1984; Thomas et al., 1996). Similar cellular effects have been described with a number of anaesthetic agents (Kress, 1995). By interfering with EC coupling in skeletal muscle cells, ethanol can exert deleterious consequences in muscle strength. We have already made the same observations when evaluating human cultured myotubes (Nicolás et al., 1998). Nevertheless, the inhibitory effects of ethanol on [Ca2+]i transients were somewhat lower than those documented in cardiomyocytes and cultured myotubes. While in the latter cells, 100 mM ethanol caused a decrease of about 25% in the magnitude of the [Ca2+]i transients (Nicolás et al., 1996, 1998), in the present study, mouse skeletal fibre [Ca2+]i transients were inhibited by 11%. This, we believe, is in accordance with the lower clinical consequences of ethanol misuse in skeletal muscle compared to heart muscle.

The maximum concentrations of ethanol used in this study were higher than those generally determined in alcoholics. Nevertheless, significant effects on Ca2+ channel activity and [Ca2+]i transients were observed with 20 to 100 mM (90–460 mg/dl) ethanol, which covers the range that can be encountered during binge drinking, and blood-ethanol concentrations as high as 300 mM have been encountered (Berlid and Hasselbalch, 1981; Johnson et al., 1982; O'Neill et al., 1984). Although the pharmacological effects at higher ethanol concentrations do not necessarily reflect all of the key changes in skeletal muscle physiology that occur at lower levels of ethanol, the data presented are consistent with the view that impairment of EC coupling may contribute to muscle weakness in alcoholism.


This work was supported with grants from Fondo de Investigaciones Sanitarias FIS 98/0330 and FIS 99/0115.


  • * Author to whom correspondence and reprint requests should be addressed at: Department of Internal Medicine, Hospital Clinic, Villarroel 170, 08036 Barcelona, Spain.

  • IDIBAPS, Institut d'Investigacions Biomèdiques August Pi i Sunyer.


View Abstract