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INFLAMMATION, CYTOKINES, AND TEMPERATURE REGULATION
Department of Physiology, University of Aarhus, DK-8000 Århus C, Denmark
Submitted 4 April 2002 ; accepted in final form 5 March 2003
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Ca2+-ionophore; skeletal; extensor digitorum longus; lactate dehydrogenase
Ca2+ ions have been shown to activate intracellular phospholipases (20) and proteases (3, 4). These enzymes in turn degrade cellular lipids, proteins, and membranes and eventually lead to sarcolemmal leakage and loss of intracellular components to the surrounding water phase (21). Moreover, the loss of cellular integrity (by allowing direct diffusion of Ca2+ ions from the extracellular phase into the cytoplasm) might start a vicious cycle with self-increasing degradation of the muscle cells (for review, see Ref. 17).
In 1983, Jones et al. (21) found that electrical stimulation (100 Hz for 0.5 s every 5 s) led to a release of lactate dehydrogenase (LDH) from isolated mouse soleus and extensor digitorum longus (EDL). This was later confirmed (16) in rat EDL during long-term low-frequency stimulation (1 Hz continuously). Furthermore, Jones et al. (22) showed that increasing the permeability of the membrane to Ca2+ by using either the Ca2+ ionophore A23187 [GenBank] or a detergent likewise led to a release of LDH from mouse soleus.
In this article, we tested the hypothesis that an increased influx of Ca2+ is a causative factor in inducing loss of muscle cell integrity as evidenced by the loss of large intracellular molecules (LDH) as well as the additional uptake of Ca2+. Moreover, we wanted to examine whether this increase in Ca2+ uptake persists after cessation of stimulation. To do this, we applied three different experimental approaches: 1) prolonged low-frequency stimulation, 2) electroporation, and 3) exposure of the muscle to the Ca2+ ionophore A23187 [GenBank] . In each case, the uptake of Ca2+ was quantified and related to the release of LDH.
Long-term stimulation at low frequency (1 Hz for 120 or 240 min) was chosen because it resembles the average spontaneous activity (0.3–2.8 Hz) measured in vivo in rat EDL (19) and at the same time allows comparison with the results of our previous studies (15, 16). Electroporation provided a method of acutely inducing pores in the sarcolemma (7) that allowed entry of Ca2+, and A23187 [GenBank] provided a more specific method that allowed Ca2+ to gain access to the cytoplasm without the use of electrical pulses.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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18
mm; diameter, 2 mm) to allow a more ready access to oxygen and substrate
during incubation. Diffusion was further improved by stirring, which was
achieved by constant bubbling with a 95% O2-5% CO2
mixture. Muscle preparation and incubation. The animals were killed by cervical dislocation followed by decapitation, and intact EDL muscles were dissected out as previously described (8). The standard incubation medium was a Krebs-Ringer bicarbonate buffer (pH 7.4) that contained (in mM) 120.2 NaCl, 25.1 NaHCO3, 4.7 KCl, 1.2 KH2PO4, 1.2 MgSO4, 1.3 CaCl2, and 5 or 10 D-glucose. Most incubations took place at 30°C in a volume of 2–5 ml. In a few experiments, the temperature was changed to 20 or 37°C. After preparation, the muscles were mounted on electrodes at resting length except in the force-measurement experiments, where it was possible to adjust the length to the optimal length for twitch-force development (on average, 108% of resting length). All muscles were equilibrated for a minimum of 30 min in the standard medium before further incubation. Control experiments that measured tetanic force (1.5-s pulse trains at 90 Hz) showed that EDL muscles allowed to rest for 8 h underwent no significant decrease in tetanic force. However, when stimulated every 5 min with 1.5-s, 90-Hz pulse trains, tetanic force gradually decreased by 44% over 8 h.
Determination of LDH. Muscle cell integrity was monitored by measuring the release of LDH into the incubation medium. This was done essentially as previously described (16). After excision, the muscles were mounted at resting length on the same muscle holders that were used for electrical stimulation. The muscles were then washed 3 x 30 min at 30°C in standard Krebs-Ringer bicarbonate buffer and 1 x 30 min in a buffer similar to that used in the subsequent experiment. During the experiments, the muscles were moved to new tubes every 30 min. Buffer samples were taken from the tubes immediately after removal of the muscles. BSA was added to the buffer samples at a final concentration of 0.1%, and a 250-µl sample was mixed with 2.65 ml of phosphate buffer (0.1 M K2HPO4 titrated with KH2PO4 to pH 7.0) that contained NADH (0.3 mM) and pyruvate (0.8 mM). The absorbance of NADH was monitored at 340 nm for 4 min (Lambda 20, Perkin-Elmer) at 30°C. In all experiments, the minor spontaneous release of LDH in resting control muscles mounted at resting length was monitored.
45Ca uptake. Uptake of 45Ca was determined as earlier described (9, 15). After equilibration and in some cases stimulation in unlabeled buffer, the muscles were incubated in buffer that contained 45Ca (0.1–0.5 µCi/ml) for periods that varied between 15 and 240 min. Depending on the experiment, the concentration of cold Ca2+ in the buffer was between 0.1 and 5.0 mM. In some experiments of shorter duration (15–60 min), incubation was followed by washout for 4 x 30 min at 0°C in Ca2+-free Krebs-Ringer bicarbonate buffer that contained 0.5 mM EGTA to remove extracellular 45Ca.
Finally, the muscles were blotted, weighed, and soaked overnight in 2–3 ml of 0.3 M trichloroacetic acid (TCA). The next day, 45Ca activity of the TCA extract was determined by liquid scintillation counting (Tri-Carb 2100 TR, Packard), and 45Ca uptake was determined on the basis of the activity of 45Ca in the incubation medium. When washout was performed, a previously determined correction factor (1.6) for the loss of cellular 45Ca that occurred during the 4 x 30-min washout at 0°C was applied when the uptake was calculated (15). Where no washout was applied, the values were corrected for the content of 45Ca in the extracellular space. The extracellular space determined by using [14C]sucrose was found to be 23% in EDL.
Ca2+ content. After the experimental protocol, the muscles were blotted and weighed, and the tendons were carefully cut off. The muscles that weighed 20–25 mg were soaked overnight in 3 ml of 0.3 M TCA. Ca2+ content was determined by atomic absorption spectrophotometry (Philips PU 9200, Pye Unicam, Cambridge, UK) by using 1.5 ml of the TCA extract after addition of KCl to a final concentration of 2.4 mM. The muscle extracts were measured against a blank and standards (12.5, 25, and 50 µM CaCl2) that contained the same amount of TCA and KCl (12). Where no washout was applied, the values were corrected for the Ca2+ content in the extracellular space.
Electroporation. Electroporation was performed by using an ECM 830 square-wave electroporator (BTX, San Diego, CA). The muscles were taken down from the holders and placed between two aluminum electrodes in a small cuvette that contained 1.0 ml of Krebs-Ringer bicarbonate buffer with the same extracellular Ca2+ concentration ([Ca2+]o) as in the experiment. The cuvette was then placed with the aluminum electrodes adjacent to the metal contacts in the safety stand, and the appropriate square pulses were delivered. The shape of the pulse was checked on an oscilloscope. All muscles received three pulses at a frequency of 1 Hz, a duration of 200 µs, and an amplitude of 313 or 500 V/cm. Control muscles were taken off the holders and placed in the cuvette but were not electroporated. The muscles were then remounted on the holders at resting length.
Electrical stimulation. An experimental setup that allowed the simultaneous direct stimulation of 12 muscles was used. Each muscle was mounted between two platinum electrodes on either side of the central part of the muscle at resting length so as to allow isometric contractions. Single pulses with an amplitude of 10 V and a duration of 1 ms were used in all experiments except during indirect stimulation, where the pulse duration was reduced to 0.02 ms. Frequency, amplitude, and pulse duration were checked with an oscilloscope (HAMAG HM 207).
Force measurements during electroporation experiments. Force was measured essentially as described earlier (10). The muscles were mounted between two platinum electrodes on either side of the central part of the muscle at optimal length for twitches, and force was checked by using two short tetanic contractions at 30 and 90 Hz. The muscles were then allowed to rest. Twenty minutes before electroporation, force was checked using a short tetanic contraction (90 Hz for 0.5 s). After a 20-min rest, the muscles were taken off the transducers and placed in the cuvette for electroporation. After the muscles received the electroporation pulses, they were remounted and stretched to the same baseline tension as before electroporation. In the subsequent 70–90 min, force was checked at chosen intervals. No change in baseline tension was observed during that period.
Determination of sucrose space. Extracellular volume was determined by using [14C]sucrose as a marker. Muscles were equilibrated for 90 min in Krebs-Ringer bicarbonate buffer that contained sucrose (1 mM) and [14C]sucrose (0.5 µCi/ml). After incubation, the muscles were blotted, weighed, and soaked overnight in 2 ml of 0.3 M TCA. The next day, [14C]sucrose activity in the TCA extract was determined by liquid scintillation counting (Tri-Carb 2100 TR), and the uptake of [14C]sucrose in the muscles was calculated by comparison with the activity of [14C]sucrose in the incubation medium. In one experiment, muscles were dried to constant weight before being soaked in TCA to determine total water content.
The effects of electroporation were examined by using two experimental procedures: 1) muscles were incubated in [14C]sucrose for 30 min before electroporation to allow the isotope to diffuse into the core of the muscles and for 60 min after the electroporation; and 2) in other experiments, this incubation was followed by a wash of 4 x 15 min in ice-cold Na+-free Tris-sucrose buffer to remove the major part of the extracellular [14C]sucrose. This allowed us to assess the amount of [14C]sucrose that was trapped inside the cells during the transient opening and resealing of the plasma membrane.
Chemicals and isotopes. All chemicals were of analytical grade. A23187 [GenBank] , TTX, and tubocurarine were purchased from Sigma Chemical (St. Louis, MO); NADH and pyruvate were from Boehringer Mannheim (Germany). 45Ca (1.31 Ci/mmol) and [14C]sucrose (0.6 Ci/mmol) were from Amersham International (Aylesbury, Bucks, UK).
Statistics. Results are given as mean values ± SE. The statistical significance of any difference between groups was ascertained by using a t-test for unpaired observations. All comparisons made were between separate groups of muscles.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Experiments that use the extracellular marker [14C]sucrose showed that 120 min of stimulation at 1 Hz gave rise to an increase in the [14C]sucrose space from 23.5 ± 0.7 to 38.3 ± 1.3% (P < 0.001; n = 9 vs. 9) measured in a 90-min period after stimulation. This increase (14.8%) could not be accounted for by augmented total muscle water content, because this only increased 2% (n = 6 pairs of muscles), but it probably represents increased access of [14C]sucrose to the intracellular water space of some fibers due to membrane damage. Another indication of cell membrane leaks is the total Na+ content of the muscle. Because extracellular Na+ is 145.3 mM, the increase in [14C]sucrose space of 14.8% would correspond to an increase in Na+ content of 21.5 µmol/g of wet wt. Flame photometric measurements showed that Na+ content increased by 23.6 µmol/g of wet wt (from 42.3 to 65.9 µmol/g of wet wt).
After stimulation at 1 Hz for 120 min at 1.3 mM Ca2+, resting 45Ca uptake was increased even long after cessation of stimulation. After 60 min of recovery, resting 45Ca uptake was still increased 3.6-fold (to 0.23 ± 0.01 µmol·g of wet wt-1·15 min-1), and after 120 min rest, the increase was 2.3-fold (to 0.17 ± 0.02 µmol·g of wet wt-1·15 min-1).
As shown in Table 1, the poststimulatory increase in resting 45Ca uptake was not significantly affected by the addition of TTX. Thus in contrast to the uptake during stimulation (15, 16), the poststimulatory increase in 45Ca uptake was not due to influx through the voltage-gated Na+ channels but rather reflects nonspecific leakage through the plasma membrane.
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Also total Ca2+ content was found to increase during recovery. Immediately after stimulation for 120 min at 1 Hz with 1.3 mM Ca2+, the Ca2+ content of the muscles was 1.50 ± 0.07 µmol/g of wet wt, and after a subsequent 120 min of recovery, the Ca2+ content had increased to 2.34 ± 0.33 µmol/g of wet wt (P < 0.001; n = 6).
To examine whether this poststimulatory uptake of Ca2+ was important for the maintenance of integrity during recovery, the release of LDH was measured under the same experimental conditions. Buffer samples for determination of LDH release were taken before the onset of electrical stimulation, during the final 30 min of the 120-min stimulation, and after a subsequent 90–120 min of recovery. As can be seen from Fig. 3, there is no increase in LDH release during the final 30 min of stimulation. However, in the interval from 90 to 120 min after the cessation of stimulation, an eightfold increase in LDH release was found that reached a value of 3.1 ± 0.4 U·g of wet wt-1·30 min-1. When [Ca2+]o was reduced to 0.3 mM, the LDH release in the final 30 min of the recovery period was significantly lower (1.5 ± 0.2 U·g of wet wt-1·30 min-1; P < 0.001). Increasing Ca2+ to 5.0 mM, however, caused no further increase in LDH release in the final 30 min of the recovery period (3.1 ± 0.3 U·g of wet wt-1·30 min-1; data not shown).
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To test the hypothesis that the increase in LDH release was due to Ca2+-activated enzymatic processes that degraded the sarcolemma, we investigated whether release of LDH showed temperature dependence. This was done in a separate series of experiments where EDL muscles were stimulated continuously at 1 Hz for 240 min at 20, 30, or 37°C in the standard buffer that contained 1.3 mM Ca2+. As shown in Fig. 4A, the Ca2+ content of the resting muscles did not differ significantly between 20 and 37°C. However, the Ca2+ content of the stimulated muscles increased with temperature (by 37% from 20 to 30°C; P < 0.005; and by 33% from 30 to 37°C; P < 0.05). As shown in Fig. 4B, stimulation-induced LDH release showed a much more marked temperature dependence. At 30°C, LDH release from the stimulated muscles was fivefold larger than at 20°C (P < 0.001), and at 37°C, the release was increased 2.5-fold compared with the release at 30°C (P < 0.001). In contrast, the resting muscles showed no significant increase in LDH release with increasing temperature.
Effects of electroporation. The long-term stimulation experiments leave uncertainties as to whether the observed leakages could be due to impaired energy metabolism or mechanical strain. It was of interest, therefore, to characterize the effects of acutely induced cell membrane leakage. When a large electric field is applied across the muscle within a short interval of time, the integrity of the sarcolemma breaks down and pores are created in the membrane. These pores are transient but allow the passage of otherwise nonpermeant ions and molecules such as extracellular markers (14). This allows for a rapid increase in Ca2+ uptake without previous long-term stimulation and contractile activity.
Measurements of force after electroporation showed that the loss of force and the ability to recover could be graded by application of different voltages of varying durations. As shown in Fig. 5, the application of three 200-µs pulses of 500 V/cm induced a complete and lasting loss of force. From a series of preliminary force-recovery experiments, we chose to use an electroporation paradigm of three pulses of 200-µs duration at a voltage of 313 V/cm. This resulted in an almost complete loss of force in the first 10 min after electroporation. Subsequently, the force underwent a slow recovery and reached 85% of the control level after 50 min.
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The degree of permeabilization of the muscle cells was monitored by measuring the [14C]sucrose uptake after electroporation. As shown in Table 2, electroporation induced a small but significant increase in the [14C]sucrose uptake. Because a large part of the [14C]sucrose taken up resides in the extracellular space, an attempt was made to reduce this component by washing the muscles after the incubation in [14C]sucrose. As shown in Table 2, even after a washout of 4 x 15 min, the amount of [14C]sucrose retained was considerably (threefold) larger in the muscles that were exposed to electroporation.
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As shown in Table 3, in the standard Krebs-Ringer bicarbonate buffer, electroporation increased 45Ca uptake four- to fivefold when measured during the subsequent 30 or 60 min (first and second columns). When the Ca2+ concentration of the buffer was increased to 5.0 mM Ca2+, electroporation increased 45Ca uptake almost eightfold (second and third columns).
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As shown in Fig. 6, electroporation induced an early and marked increase in LDH release that was evident within the first 15 min. In the subsequent period, LDH release decreased but remained significantly and markedly higher (8- to 12-fold) than the controls throughout the period of measurement (P < 0.01). The release of LDH clearly depended on the Ca2+ concentration in the buffer. Thus at 5.0 mM Ca2+, the initial increase in LDH release was three times larger than that measured at 1.3 mM Ca2+ (P < 0.001), and during the following 150 min, the LDH release at 5.0 mM Ca2+ was significantly larger than at 1.3 mM Ca2+ (P < 0.005). Control experiments where muscles were electroporated in a Krebs-Ringer buffer that contained 0.1 mM Ca2+ or no Ca2+ with the addition of 0.5 mM EGTA showed a very large LDH release that was probably due to instability of the membrane under the (nearly) Ca2+-free conditions (data not shown).
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In view of the marked effect of electroporation on 45Ca uptake, it could be envisaged that the observed uptake of 45Ca during direct electrical stimulation could be due to electroporation. We have previously shown that direct electrical stimulation at 40 Hz increased the rate of 45Ca uptake 34-fold in EDL muscle (16). To test whether this could be due to electroporation of the membrane, we investigated whether indirect stimulation via the nerve would result in similar increases in the rate of 45Ca uptake. Isolated EDL muscles were stimulated at 40 Hz for 30 s by using pulses of 0.02-ms duration so as to achieve indirect stimulation via the nerve. The rate of 45Ca uptake in the resting muscles was 4.5 ± 0.4 nmol·g of wet wt-1·min-1, whereas the rate of 45Ca uptake during stimulation was 66.8 ± 16.6 nmol·g of wet wt-1·min-1 (P < 0.01; n = 5). Thus indirect stimulation induced a 15-fold increase in the rate of 45Ca uptake. Addition of tubocurarine completely blocked the excitation-induced increase in 45Ca uptake.
Effects of A23187 [GenBank] . To identify a more specific effect of Ca2+ on LDH release, the effects of the Ca2+ ionophore A23187 [GenBank] were examined. Figure 7 shows the time course of the effects of A23187 [GenBank] (2 x 10-5 M) on 45Ca uptake and Ca2+ content in EDL muscles. A significant increase in 45Ca uptake could be detected within the first 15 min of incubation. After 120 min of incubation, the muscles treated with A23187 [GenBank] had taken up 45Ca corresponding to 2.7 µmol Ca2+/g of wet wt, which represented an increase of 1.9 µmol/g of wet wt compared with the untreated controls. In accordance with this, the increase in total Ca2+ content at 120 min was measured to 1.8 µmol/g of wet wt. A similar close correspondence between 45Ca uptake and increase in total Ca2+ content was observed in all other incubation periods.
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As shown in Fig. 8A, in experiments performed under identical conditions, it was found that A23187 [GenBank] caused a marked increase in the release of LDH, which was clearly detectable and highly significant (P < 0.005) already over the first 15 min. The LDH release showed a steep increase in the first 60 min that was followed by a somewhat slower increase a release as high as 13.3 U·g of wet wt-1·30 min-1 in the interval 150–180 min of incubation. When [Ca2+]o was decreased to 0.1 mM, the LDH release was markedly reduced (to 8.3 U·g of wet wt-1·30 min-1 at 180 min). At 0 mM Ca2+, the release was further reduced, and finally when the buffer contained 0 mM Ca2+ and 0.5 mM EGTA, no effect of A23187 [GenBank] could be detected.
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As another indicator of cell damage induced by A23187 [GenBank] and Ca2+, Na+ content was measured. As shown in Fig. 8B, the increase in LDH release that was induced by A23187 [GenBank] was clearly dependent on the Ca2+ content of the buffer and associated with an increase in Na+ content. Because A23187 [GenBank] transports only divalent cations, this increase in Na+ content could not be attributed to a direct action of A23187 [GenBank] ; rather, it must have been due to an increased permeability of the cellular membrane that allowed Na+ from the extracellular compartment to enter the interior of the muscle cells.
To investigate whether LDH release depended on the amount of Ca2+ taken up, experiments were performed that measured 45Ca uptake and LDH release as a result of incubation with A23187 [GenBank] at several different concentrations of [Ca2+]o. As shown in Fig. 9, LDH release increased in proportion to the stimulation of 45Ca uptake. The relation between the data on Ca2+ uptake and LDH release indicates that at a [Ca2+]o of 0.6 mM, a saturation plateau was reached where further increase in [Ca2+]o caused no additional membrane damage.
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Control experiments showed that A23187 [GenBank] (2 x 10-5 M) caused no significant increase in resting tension at a [Ca2+]o of either 1.3 or 5.0 mM.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Our results confirm and extend those obtained by Jones et al. (21, 22) that linked intracellular enzyme release after stimulation with extracellular Ca2+ and support the suggestion that entry of Ca2+ during stimulation initiates the subsequent release of LDH (16). Moreover, their observations that LDH release is further augmented when the muscles are stimulated under hypoxic conditions and that this is prevented by the removal of extracellular Ca2+ (22) are well in line with the Ca2+ hypothesis. Hypoxia has in many instances been found to increase intracellular Ca2+ concentration, and recently it was shown in myotubes that this was due to an increased influx of Ca2+ through sarcolemmal Na+ channels (24). It could be envisaged, therefore, that the excitation-induced increase in 45Ca uptake that we observed in our experiments was due to a shortage of oxygen that arose from the increased energy demand during contractions. However, preliminary experiments with isolated EDL muscles showed that during 30 min of exposure to anoxic buffer, resting 45Ca uptake was unchanged, and electrical stimulation (30 s at 40 Hz) still induced a 15-fold increase in 45Ca uptake. This indicates that the marked excitation-induced stimulation of 45Ca uptake is not caused by hypoxia, which expectedly would be much more pronounced at 40 Hz than at 1 Hz. Moreover, if the effect of excitation on 45Ca uptake was solely the result of hypoxia, it would be expected to disappear in muscles incubated for 30 min in the absence of oxygen.
Long-term electrical stimulation. As shown in Fig. 1, during long-term stimulation at 1 Hz, there is a highly significant correlation between 45Ca uptake and LDH release, which indicates loss of the integrity of the sarcolemma. The present observation that after 120 min of electrical stimulation at 1 Hz the muscles showed a marked increase in the resting uptake of extracellular Ca2+ suggests that prolonged stimulation elicits a more lasting leakage of the sarcolemma. This was evident for at least 120 min poststimulation. The increased uptake of Ca2+ could be a prerequisite for more extensive damage. In line with this, the eightfold increase in LDH release observed from 90 to 120 min after the cessation of stimulation was dependent on [Ca2+]o.
Interestingly enough, the previously observed release of LDH after 240 min of stimulation at 1 Hz (16) was not higher than that observed after 120 min of stimulation at 1 Hz and 120 min of rest. Also, the Ca2+ content of the muscles after 240 min of stimulation was not higher than after 120 min of stimulation and 120 min of rest. This indicates that the damage is initiated during the first 120 min of stimulation, and in the following 120 min of rest, the damage to the cellular membrane progresses and finally allows even the large LDH molecule to leak out. Another important implication of this is that although stimulation is stopped, further entry of Ca2+ may occur and possibly exacerbate the damage process.
Mechanical damage has been proposed as a likely primary mechanism for the observed increase in Ca2+ uptake and LDH release elicited by contractile activity (1, 2). Mechanical damage may be important during eccentric contractions. However, our present and previous observations on isotonic and isometric contractions indicate that mechanical damage is not a likely explanation for the increased uptake of Ca2+ during stimulation, and that LDH release seems to be due to the increased uptake of Ca2+ and not primarily to mechanical damage. We have the following arguments: 1) LDH release correlates with the stimulation-induced increase in 45Ca uptake (see Fig. 1); 2) the excitation-induced early increase in 45Ca uptake in EDL is rapid in onset (30 s of stimulation at 40 Hz), large (up to 34-fold), and does not lead to any detectable increase in LDH release over the following 120 min (16); 3) uptake of Ca2+ is considerably larger in muscles that undergo isotonic contractions at zero load (where mechanical strain on the muscle is very low) compared with muscles that contract isometrically (15); and 4) although force production during long-term stimulation (240 min at 1 Hz) was similar in EDL and soleus, 45Ca uptake and LDH release were 5- and 10-fold larger, respectively, in EDL than in soleus (15, 16).
The large effect of electroporation on 45Ca uptake raises the question whether direct electrical stimulation might induce electroporation and account for the observed uptake of 45Ca. We have previously reported a 34-fold increase in the rate of 45Ca uptake in EDL muscles stimulated at 40 Hz (16). However, we also showed that this was suppressed by TTX. In the present article, we show that indirect stimulation via the nerve also results in a marked (15-fold) increase in the rate of 45Ca uptake, which is completely suppressed by tubocurarine. Taken together, these observations make it unlikely that the observed stimulation-induced uptake of 45Ca is due to electroporation.
When measured during 240 min of chronic low-frequency stimulation, the release of LDH was strongly dependent on temperature. At 20°C, the stimulation-induced increase was 0.38 U·g of wet wt-1·30 min-1, whereas at 37°C, the increase was 6.29 U·g of wet wt-1·30 min-1, which corresponds to a 17-fold increase. In contrast, compared with the LDH release, the uptake of Ca2+ was only slightly affected by temperature. Thus increasing the temperature from 30 to 37°C only resulted in a 33% increase in the Ca2+ content of the stimulated muscles. These results suggest that the LDH release at higher temperatures is not solely the result of a higher uptake of Ca2+, but that the loss of cellular integrity depends on an enzymatic process with a rather high temperature coefficient. Recently, Warren et al. (31) reported a 33-fold increase in LDH release after eccentric exercise of mouse EDL when the temperature was increased from 25 to 37°C.
The strong dependency on temperature may be very important for the cell damage that develops during intense exercise, where the deep-core muscle temperature may reach values as high as 41°C (18). From our data, it can be calculated that if the temperature in a muscle increases from 34°C at rest to 41°C during intense exercise (29), the risk of developing cell leakage will increase 2.5-fold.
Electroporation. The effect of acute cell membrane damage was studied by using electroporation. Electroporation has been shown to induce the formation of numerous small pores (20–120 nm in diameter) in the sarcolemma (7). This is likely to cause a general leakage of ions and lead to depolarization and loss of excitability. Indeed, we find that electroporation leads to a prompt and general loss of contractility with a slow recovery to 85% of the control level within 50 min. This is in accordance with the observed membrane-resealing time for isolated muscle cells exposed to electroporation (6). The almost complete loss of muscle force indicates that virtually all fibers are affected by the electroporation.
In intact mouse skeletal muscle in vivo, electroporation was shown to increase the uptake of the extracellular marker CrEDTA (14). We find that electroporation leads to an increased uptake of [14C]sucrose, which is a strong indication of a loss of plasma-membrane integrity. This loss is transient, as can be seen from the significant increase in the amount of [14C]sucrose that was retained after washout at 0°C (see Table 2). Moreover, this is evidence that the extracellular marker penetrated into the muscle fibers and after the resealing of the pores was trapped in the cytoplasm.
Electroporation led to a marked increase in 45Ca uptake. Again the uptake was dependent on [Ca2+]o. In buffer that contained 1.3 mM Ca2+, 45Ca uptake (measured over 60 min) increased almost fourfold with electroporation, and at 5.0 mM Ca2+, the increase was eightfold. In line with this, electroporation also led to a large release of LDH. This was most pronounced within the first 15 min but stayed significantly elevated throughout the 180-min period of observation. LDH release was larger from the muscles electroporated at 5.0 mM Ca2+ compared to the release from the muscles electroporated at 1.3 mM Ca2+. The large LDH response within the first 15 min after electroporation can be attributed to the large influx of Ca2+ that occurred via the electroporation pores. However, a certain leakage of LDH through the pores that directly arises from the electroporation cannot be excluded, although the dependency of the release on [Ca2+]o suggests that Ca2+-activated processes comprise the major component. What is perhaps even more interesting is the release of LDH observed from the period 60 min after electroporation and onward. Here, force had recovered to 85% of the control level, and we therefore expected that most of the pores created by electroporation would be resealed. This delayed LDH release must have been due to Ca2+-dependent damage to the sarcolemnma. Thus, by eliciting a rapid uptake of Ca2+, electroporation activates a lasting release of LDH.
A few experiments were conducted at (nearly) Ca2+-free conditions to separate the direct effects of electroporation from the Ca2+-mediated effects on LDH release. Unfortunately, however, electroporation at (nearly) Ca2+-free conditions resulted in very large releases of LDH. Also at (nearly) Ca2+-free conditions, the control muscles not undergoing electroporation showed a large LDH release after being positioned in a cuvette and remounted on the muscle holder. Muscle cells are very apt at repairing the membrane (26); thus damage caused by handling or by Ca2+-activated mechanisms is likely to be repaired. However, Ca2+ is needed for this repair mechanism to function properly. Thus we assume that at the low Ca2+ concentrations tested, the muscle cells were not able to repair the damage to the sarcolemma that occurred as a result of handling and/or electroporation and thus resulted in large increases in LDH release despite no influx of Ca2+. On the other hand, it is also clear that to achieve a prolonged increase in LDH release, as observed in many of our experiments, a continuous degradation of the membrane must occur.
A23187. To identify the role of Ca2+ in eliciting membrane leakage, we characterized the time course of the effects of the Ca2+ ionophore A23187 [GenBank] , which is a rather specific agent that favors the entry of Ca2+ into the muscle cells without causing muscle contraction. In muscles not undergoing any contractions and therefore not likely to suffer mechanical damage, A23187 [GenBank] produced a progressive loss of LDH, which was detectable within the first 15 min of incubation and correlates to the rate of 45Ca uptake. Moreover, the observation that A23187 [GenBank] produced no LDH release in the absence of Ca2+ indicates that A23187 [GenBank] as such causes no leakage of the plasma membrane. These experiments indicate that Ca2+ alone is enough to elicit skeletal muscle damage. In A23187 [GenBank] -treated muscles, LDH release was elicited at Ca2+-uptake levels much smaller than those observed during electrical stimulation (see Figs. 1 and 9). We have no clear explanation for this difference.
Perspectives
The present observations confirm and extend previous studies (17, 20, 21) in documenting that Ca2+ gaining access to the cytoplasm from the extracellular phase elicits loss of cellular integrity. The new aspect is that this process seems to be self-increasing, which possibly leads to more extensive muscle damage. This may explain why the damage process sometimes continues for several hours after cessation of muscle work (13, 23, 28). The observations are important for the understanding, prevention, and treatment of loss of muscle integrity that arise during work or sports events. Moreover, the widespread muscle cell damage that is elicited by electrical shocks (25) or other conditions where the permeability of the membrane is increased (e.g., sepsis, rhabdomyolysis, or traumatic cell damage) may be worsened by subsequent entry of Ca2+ into the muscle cells and a following additional activation of Ca2+-sensitive proteases and lipases.
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ACKNOWLEDGMENTS |
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This work was supported by grants from the Danish Medical Research Council Grant (J. N. 9802488), the Danish Biomembrane Center, Aarhus Universitets Forskningsfond, and Kulturministeriets Udvalg for Idrætsforskning.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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