Department of Internal Medicine, IDIBAPS, Hospital Clinic, Faculty of Medicine, University of Barcelona, Spain
Received 15 February 1999; in revised form 16 August 1999; accepted 23 August 1999
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ABSTRACT |
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INTRODUCTION |
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MATERIALS AND METHODS |
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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, 20x-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 (ß-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 (20500 mM; 902300 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.
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RESULTS |
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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.
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DISCUSSION |
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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 (90460 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.
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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IDIBAPS, Institut d'Investigacions Biomèdiques August Pi i Sunyer.
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