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¹ Department of Zoology, La Trobe University Bundoora Campus, Melbourne, Victoria 3086, Australia

	Abstract

Top Abstract Introduction Methods Results Discussion References

 
This study investigated the effects of elevated, physiological levels of intracellular free [Ca²⁺] on depolarization-induced force responses, and on passive and active force production by the contractile apparatus in mechanically skinned fibres of toad iliofibularis muscle. Excitation–contraction (EC) coupling was retained after skinning and force responses could be elicited by depolarization of the transverse-tubular (T-) system. Raising the cytoplasmic [Ca²⁺] to 1 µM or above for 3 min caused an irreversible reduction in the depolarization-induced force response by interrupting the coupling between the voltage sensors in the T-system and the Ca²⁺ release channels in the sarcoplasmic reticulum. This uncoupling showed a steep [Ca²⁺] dependency, with 50% uncoupling at 1.9 µM Ca²⁺. The uncoupling occurring with 2 µM Ca²⁺ was largely prevented by the calpain inhibitor leupeptin (1 mM). Raising the cytoplasmic [Ca²⁺] above 1 µM also caused an irreversible decline in passive force production in stretched skinned fibres in a manner graded by [Ca²⁺], though at a much slower relative rate than loss of coupling. The progressive loss of passive force could be rapidly stopped by lowering [Ca²⁺] to 10 nM, and was almost completely inhibited by 1 mM leupeptin but not by 10 µM calpastatin. Muscle homogenates preactivated by Ca²⁺ exposure also evidently contained a diffusible factor that caused damage to passive force production in a Ca²⁺-dependent manner. Western blotting showed that: (a) calpain-3 was present in the skinned fibres and was activated by the Ca²⁺exposure, and (b) the Ca²⁺ exposure in stretched skinned fibres resulted in proteolysis of titin. We conclude that the disruption of EC coupling occurring at elevated levels of [Ca²⁺] is likely to be caused at least in part by Ca²⁺-activated proteases, most likely by calpain-3, though a role of calpain-1 is not excluded.

(Received 23 December 2004; accepted after revision 2 March 2005; first published online 3 March 2005)
Corresponding author E. Verburg: Department of Zoology, La Trobe University, Bundoora, Victoria 3086, Australia. Email: e.verburg{at}latrobe.edu.au

	Introduction

Top Abstract Introduction Methods Results Discussion References

 
Force development in skeletal muscle is triggered by a rise in intracellular [Ca²⁺] (Melzer et al. 1995; Gordon et al. 2000). Depolarization of the transverse-tubular (T-) system activates specialized voltage sensors, which in turn open the Ca²⁺ release channels in the apposing sarcoplasmic reticulum (SR) at the triad junction, leading to an increase in intracellular [Ca²⁺] and consequent force development by the contractile apparatus. Sustained high levels of intracellular Ca²⁺, however, can damage muscle function. We have previously shown that raising the free [Ca²⁺] in the cytoplasmic space to 23 and 100 µM for 10 s completely abolished depolarization-induced Ca²⁺ release in skinned fibres from the iliofibularis of the toad, with half-maximal effect occurring with 10 µM Ca²⁺ (Lamb et al. 1995). The reduction in depolarization-induced Ca²⁺ release was irreversible over the time scale of the experiment (>30 min), and was due to interruption of the coupling between the voltage sensors and the Ca²⁺ release channels. A comparable Ca²⁺-induced reduction in Ca²⁺ release has also been demonstrated in intact fibres (Chin & Allen, 1996; Chin et al. 1997). As suggested by Lamb et al. (1995), it seems likely that this Ca²⁺-dependent disruption of excitation–contraction (EC) coupling is the major cause of ‘low-frequency fatigue’ (Chin & Allen, 1996; Chin et al. 1997), a phenomenon in which repeated vigorous activation of a muscle fibre leads to a long-lasting decrease in force production that is most evident when the muscle is tested with low-frequency stimulation (Edwards et al. 1977; Allen et al. 1995).

The mechanistic basis of this Ca²⁺-dependent uncoupling is unknown. One reason for this is that the precise Ca²⁺ dependence and time dependence of the uncoupling are currently not well defined, which makes it difficult to quantitatively relate the uncoupling to particular biochemical reactions. Our previous work (Lamb et al. 1995) primarily examined the effect of relatively brief (10 s) exposures to quite high [Ca²⁺] (10 µM); these levels are similar to or substantially greater than current estimates of the average cytoplasmic [Ca²⁺] reached during a tetanus (1–20 µM; Konishi et al. 1991; Chin & Allen, 1996; Baylor & Hollingworth, 2003). Studies on intact muscle fibres (e.g. Chin & Allen, 1996) do not give a precise indication of the Ca²⁺ dependence and time dependence of the uncoupling because the intracellular [Ca²⁺] is not under close experimental control, and almost certainly varies considerably between different cytoplasmic regions during stimulation, being much higher near the sites of Ca²⁺ release. The present study, using mechanically skinned fibres with functional EC coupling, was able to avoid these problems and apply a known [Ca²⁺] within the physiological range throughout the intracellular space, and thereby accurately characterize the Ca²⁺ dependence and time dependence of the uncoupling effect.

Previous work has ruled out some mechanisms as being involved in the uncoupling, such as oxidation–reduction and phosphorylation–dephosphorylation events (Lamb et al. 1995). The present study specifically investigates whether Ca²⁺-activated neutral proteases (calpains) could be responsible for the uncoupling. We show here that calpain-3 is retained in the skinned fibres and is activated by the Ca²⁺ exposure. We also use the loss of passive force production to demonstrate that the Ca²⁺ treatment activates diffusible Ca²⁺-dependent proteases in the fibres that are capable of proteolysing titin. Titin, also known as -connectin, is a large elastic protein that keeps the thick filament centred in the sarcomere by anchoring it to the Z-line. In the skinned fibres preparation used here, where there is no extracellular matrix, titin should be predominantly responsible for the passive force produced when a fibre is stretched (Horowits et al. 1986; Neagoe et al. 2003; Tskhovrebova & Trinick, 2003). Furthermore, calpains readily proteolyse titin (Goll et al. 2003; Taveau et al. 2003; Kramerova et al. 2004). Given the known properties of the three most abundant muscle calpains, the observed concentration- and time-dependent effects of Ca²⁺ and actions of various inhibitors indicate that the proteolysis of titin, and likewise the EC uncoupling, are probably mediated at least in part by calpain-3.

	Methods

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Skinned fibre preparation

All procedures were done in accordance with guidelines of the La Trobe University Animal Ethics Committee. Cane toads (Bufo marinus) were killed by double pithing, and the iliofibularis muscles were removed. The muscles were pinned at resting length under paraffin oil and kept on ice. As previously described (Lamb & Stephenson, 1990), muscle fibres were mechanically skinned, mounted on a force transducer (AME875; SensoNor, Norway), stretched to 1.2 times resting length and transferred to a bath containing 2 ml of the appropriate potassium-based solution, according to the type of experiment involved (see below). Force responses were amplified, and recorded on a chart recorder or on a computer with Powerlab hardware (400 series; ADInstruments, Sydney, Australia) and Chart software (version 4.2). All experiments were carried out at room temperature (24 ± 2°C).

Solutions

All chemicals were obtained from Sigma (St Louis, MO, USA), unless stated otherwise. All solutions had a pH of 7.10 ± 0.01 and an osmolality of 295 ± 5 mosmol kg^–1, and 1 mM free Mg²⁺, unless stated otherwise. Free [Ca²⁺] at 0.1 µM was verified with a Ca²⁺-sensitive electrode (Orion, Cambridge, MA, USA). Leupeptin 1 mM, hemi-sulphate (ICN Biomedials, Aurora, OH, USA) calpastatin (10 µM, human recombinant domain I; Calbiochem, San Diego, CA, USA) and calpeptin (30 µM, made from 60 mM stock in DMSO) were added to the solutions as required, with the fibre always pre-equilibrated in the agent for at least 30 s.

Depolarization-induced Ca²⁺-release experiments.  The ‘K-HDTA’ solution used to polarize the T-system contained (mM): K⁺, 126; Na⁺, 37; hexamethylene-diamine-tetraacetic acid (HDTA^2–; Fluka, Buchs, Switzerland), 50; total ATP, 8; total magnesium, 8.6; creatine phosphate (CrP), 10; total EGTA, 0.05; Hepes, 90; N₃^–, 1 or 0; pH 7.10 and pCa 7.0 (0.1 µM free Ca²⁺). The ‘Na-HDTA’ solution was similar but with all K⁺ replaced with Na⁺. The ‘load solution’ was the K-HDTA solution with 1 mM total EGTA at pCa 6.7 (0.2 µM free Ca²⁺). The ‘low [Mg²⁺] solution’ was similar to the K-HDTA solution, but with a free [Mg²⁺] of only 50 µM (2.1 mM total Mg²⁺) (see Lamb & Stephenson, 1994).

Contractile apparatus experiments.  These solutions had strong Ca²⁺ buffering (see Stephenson & Williams, 1981). The ‘relaxing solution’ was similar to K-HDTA solution with all HDTA replaced with 50 mM EGTA (pCa > 9), and the ‘maximum-activation solution’ had 49.5 mM CaEGTA and 0.5 mM free EGTA to give a free [Ca²⁺]20 µM (pCa 4.7). Solutions of intermediate [Ca²⁺] were made by appropriate mixture of these two solutions. Ca²⁺-sensitivity was examined using a sequence of solutions of progressively higher [Ca²⁺], limiting the estimate of maximum activation to a 5 s exposure to 4 µM (pCa 5.4) until after the 40 µM Ca²⁺-rigor treatment. The force–[Ca²⁺] data in each case were fitted with a Hill curve.

Elevated [Ca²⁺] and rigor treatment.  The ‘control-rigor solution’ was similar to K-HDTA solution, but with all ATP and CrP replaced iso-osmotically with additional HDTA (68 mM total), the total Mg²⁺ reduced to 1.5 mM to keep the free [Mg²⁺] at 1 mM, and 0.25 mM free EGTA (pCa 8). This solution was used to initially induce rigor, as well as being the control treatment and the wash solution after Ca²⁺ treatment. The ‘Ca²⁺-rigor solutions’ were similar to the control-rigor solution, but with Ca²⁺ added to set free [Ca²⁺] in the range of 1–100 µM.

EC coupling and its disruption by elevated [Ca²⁺]

The response of the skinned fibre to depolarization was measured at 1 min intervals by transferring the fibre to Na-HDTA solution, and after each force response was completed (2–3 s), the fibre was returned to the K-HDTA to repolarize the T-system. The fibre was then washed twice in the zero-ATP control-rigor solution (pCa 8) and moved for 3 min to an identical solution (control treatment) or to another rigor solution with elevated [Ca²⁺] (test treatment), before finally being washed for 30 s in control rigor solution (pCa 8) again and returned to K-HDTA solution (e.g. Fig. 1). Depolarization-induced responses were then measured both before and after loading the SR with additional Ca²⁺ by 15–20 s in the loading solution. At the end of each experiment, the skinned fibre was exposed to the low [Mg²⁺] solution to verify that the SR was indeed adequately loaded with Ca²⁺ (see Fig. 1), and then maximum Ca²⁺-activated force was determined.

View larger version (17K): [in this window] [in a new window]   Figure 1.  Exposure to physiological levels of intracellular [Ca2+] disrupts excitation–contraction (EC) coupling Depolarization-induced force responses were elicited in a skinned fibre by substituting all K+ in the solution with Na+ (Depol.). A, when the rigor treatment included a 3 min exposure to 2 µM free Ca2+, it caused an irreversible reduction in the depolarization-induced force response by more than 50% even after additional sarcoplasmic reticulum (SR) loading. B, in another fibre, a 3 min exposure to 3.5 µM free Ca2+ completely abolished the depolarization-induced response. Lowering the free [Mg2+] to 50 µM (low Mg2+), which stimulates the SR Ca2+ release channels directly, indicated that the SR was loaded above the endogenous level and that the Ca2+ release channels were still functional. Maximum Ca2+-activated force was ascertained in a heavily Ca2+-buffered solution (Max, pCa 4.7). Moving the fibre between solutions caused small force artefacts. The time scale was 1 s during the depolarizations and low [Mg2+], and 30 s elsewhere.

Each skinned fibre segment was mounted on the force transducer at its resting length; this length was determined by microscopic observation, and by adjusting the length to just below that producing any passive force. Whilst in relaxing solution, the fibre was stretched over 5 s to 2.0 times the resting length, and passive force production was recorded as in Fig. 3. The fibre segment was then subjected to one of a number of possible treatments. In some cases the fibre was kept in its stretched state and transferred to K-HDTA solution (at pCa 8) for 1 min and then to the control-rigor solution, with or without subsequent Ca²⁺ exposure (e.g. Fig. 3B and A, respectively); in such cases exposure to the rigor solution produced little or no increase in force production. In other cases, fibre length was reduced back to 1.2 times resting length for at least 2 min before exposing the fibre segment to the Ca²⁺-rigor solution (e.g. Fig. 3C). After the required treatment, each fibre was returned to its resting length for 2 min, and then again stretched in relaxing solution to 2.0 times its original resting length, and passive force production was measured. Other fibres were subjected to a number of treatments during a single stretch episode (e.g. Fig. 5). Fibres pretreated with Triton X-100 were given the usual initial 2 min equilibration period, followed by 5 min in relaxing solution with 2% (v/v) Triton X-100, and a further 5 min washing in relaxing solution before being stretched like other skinned fibres.

View larger version (11K): [in this window] [in a new window]   Figure 3.  Elevated [Ca2+] treatment during stretch damages passive force development A, passive force production in a fibre stretched to 2.0 times resting length (in relaxing solution), and then exposed to the control rigor solution (pCa 8). After the fibre was returned to its resting length for 2 min, stretching it again to 2.0 times resting length gave passive force similar to that upon the first stretch. B, when another fibre was treated similarly, except exposed for 3 min to 40 µM Ca2+ during the rigor period, force declined markedly during the Ca2+ exposure, and there was subsequently a great reduction in the amount of passive force produced at 2.0 times resting length. C, in another fibre in which the length was reduced to 1.2 times of resting length before applying the rigor and Ca2+ treatment, passive force production (at 2.0 times resting length) was virtually identical before and after the treatment.

View larger version (12K): [in this window] [in a new window]   Figure 5.  Elevated [Ca2+] damages passive force development A, complete protocol of the passive force experiment in which a skinned fibre was stretched to 2.0 times slack length and then treated. In this example the fibre was stretched to final length in two steps, producing the two peaks in passive force at the beginning. Transferring the fibre to the control-rigor solution (Rig.) initially caused a small increase in force, but then force subsequently declined with approximately its original time course. The fibre was given various treatments, and then rigor was reversed by returning to the control K-HDTA solution (see Methods) with ATP (Post), and the fibre was returned to close to resting length. B, expanded view of period in rigor solutions showing the off-on-off-on effect of elevated [Ca2+] on the rate of decay of passive force. Step reductions in force often occurred when moving the fibre between solutions, most likely because of stretch-induced breaking of the rigor bridges and possibly also titin filaments. C, expanded view of the rigor period in another fibre, showing effect of 1 mM leupeptin (leu). The first two periods are with leupeptin present, in which the rate of decay of passive force in 40 µM Ca2+ is unchanged compared with the preceding control rigor period. The Ca2+ effect could be switched on again by washing out the leupeptin and re-exposing the fibre to 40 µM Ca2+.

The iliofibularis muscle alone, or with adjoining thigh muscles, was homogenized in Ca²⁺-rigor solution containing 40 µM free Ca²⁺ (0.25 g tissue per 1 ml), to which 5 mM total CaCl₂ was added immediately after homogenization. After allowing 60 s for initiation of any Ca²⁺-activated processes, the free [Ca²⁺] was rapidly lowered to pCa 6.4 by adding EGTA to a final concentration of 20–25 mM, and then this ‘preactivated’ low-[Ca²⁺] homogenate was immediately used to treat a skinned fibre. This was compared with control treatments using homogenates prepared without raising the [Ca²⁺] or with 1 mM leupeptin added.

Measurement of calpain-3 activation in muscle homogenates

Muscle homogenates were prepared as described above, with an additional incubation for 10 min in 4% SDS. Samples were mixed 1:2 with SDS loading buffer (0.125 M Tris-HCl, 10% glycerol, 4% SDS, 4 M urea, 10% mercaptoethanol and 0.001% bromophenol blue, pH 6.8), heated at 95°C for 4 min, and stored at –20°C until analysed. Homogenates of rat skeletal (mixed) muscle were prepared in a similar way to the tool homogenate. Denatured protein was separated on an 8% SDS-PAGE gel (100 V for 15 min, 160 V for 60 min) and transferred to nitrocellulose (15–25 V for 30–90 min). Membranes were exposed overnight to mouse anticalpain-3 (monoclonal 12A2; Novocastra, Newcastle, UK) diluted 1 in 50 in blocking buffer (5% skim-milk powder in phosphate-buffered saline with 0.025% Tween (PBST)), followed by goat antimouse HRP (Bio-Rad, Hercules, CA, USA) secondary antibody, diluted 1 in 5000 in blocking buffer for 45–60 min. Bands were visualized using Opti-4 CN substrate (Bio-Rad). Densitometry of scanned images (Hewlett Packard Scanjet 5200C) was analysed using Quantity One software (Bio-Rad).

Measurement of calpain-3 activation in single fibres

Fibres were mechanically skinned under paraffin oil and placed in a 10 µl drop of control-rigor solution under oil. Once three fibres had been collected, they were transferred to a 10 µl drop of either a fresh control-rigor solution or a rigor solution containing 2, 8 or 40 µM Ca²⁺ for 1 or 3 min. The fibres were then placed in a fresh aliquot of control-rigor solution that was collected and mixed 1:1 with SDS loading buffer, heated to 95°C for 4 min, and stored at –20°C until analysed. Western blots were run as described for muscle homogenates; however, an amplified detection method using streptavidin was required due to the smaller amount of protein being separated. This involved using 1% BSA in PBST as the blocking buffer, with a number of additional washes being required following exposure to the goat antimouse HRP antibody (1 x 10 min, Bio-Rad Amplification Reagent; 4 x 5 min 20% DMSO in PBST; 2 x 5 min PBST and 1 x 30 min streptavidin HRP diluted 1 in 5000 in 1% BSA in PBST). The remaining steps were as described for the muscle homogenates.

Extraction of titin from muscle and single fibres

To examine titin in muscle, freshly excised toad iliofibularis and rat soleus muscle was chopped into fine pieces, and titin extraction buffer (8.7% SDS, 0.1 M Tris-Cl, pH 8.8, 5 mM EGTA, 50 mM DTT) was added at a ratio of 1:4 (w/v). Samples were heated at 65°C for 5 min, and incubated at room temperature for 90–120 min before being spun at 13 000 g for 20 min. The supernatant was mixed 1:1 with 50% glycerol in PBS with bromophenol blue and stored at –20°C until analysed. To obtain degraded titin, chopped muscle was placed in titin extraction buffer as above, but was heated to 30–35°C for 30 min and left at room temperature for a further 90 min before the 65°C heating step. The muscle was then treated as described. Toad single fibres were also analysed for titin. Following the measurements of passive force (see earlier), each fibre was added to 5 µl of titin extraction buffer and treated in the same way as the whole muscle. Fibres were analysed for titin within 1 week of being collected.

Measurement of titin in muscle and single fibres

Low-percentage acrylamide gels (Anderson et al. 2002; Minajeva et al. 2002) with silver staining were used to detect titin in rat and toad muscle homogenates and toad single fibres. Fifteen-well, 1.5 mm thick SDS-PAGE gels were prepared, with final concentrations as follows: 2.8% acrylamide/bis-acrylamide, 37.5:1 (Bio-Rad), 0.1% SDS, 0.375 M Tris-HCl, pH 8.6, 0.075% ammonium persulphate, 0.06% TEMED, and poured using a mini-protean 3 electrophoresis apparatus (Bio-Rad). Electrophoresis was performed at room temperature (10 mA for 30 min, 20 mA for 3 h and 30 mA for approximately 90 min). In all cases electrophoresis was continued at the final current for 30 min once the dye front had left the bottom of the gel. Gels were silver-stained using GelCode SilverSNAP Stain Kit II (Pierce, IL, USA). In brief, this procedure required a number of washes as follows: 2 x 5 min double-distilled water (ddH₂O); 2 x 15 min 30% ethanol/10% acetic acid; 2 x 5 min 10% ethanol; 2 x 5 min ddH₂O; 1 min 0.2% SilverSNAP sensitiser; 2 x 1 min ddH₂O; 25 min 2% SilverSNAP stain; 2 x 20 s ddH₂O; 4–10 min 2% SilverSNAP developer. Development was stopped with two washes (1 and 10 min) of 5% acetic acid, and the gels were then placed in ddH₂O before being scanned (Hewlett Packard Scanjet 5200C). To determine the ratio of full-length titin to degraded titin (T1 and T2, respectively), densitometry of the bands was quantified using Quantity One software (Bio-Rad).

	Results

Top Abstract Introduction Methods Results Discussion References

 
Disruption of EC coupling at moderately elevated levels of intracellular [Ca²⁺]

In order to apply Ca²⁺ at a set concentration throughout the skinned fibre, all ATP was removed from the bathing solution so as to prevent Ca²⁺ uptake by the SR pump. The skinned fibre developed rigor in this solution (pCa 8), but this was readily reversible and in itself had little effect on subsequent responses to depolarization (response after control-rigor treatment: 94 ± 2% of pretreatment response, n= 10). Exposing a fibre for 3 min to elevated [Ca²⁺] at concentrations as low as 1 µM caused irreversible disruption of EC coupling. The extent of this uncoupling depended on the free [Ca²⁺] (Figs 1 and 2). Following exposure to 2 µM Ca²⁺, depolarization-induced force response was significantly reduced to approximately half of the pretreatment controls (P < 0.001, n= 6) (Figs 1A and 2), with force almost fully abolished by 3.5 µM Ca²⁺ (pCa 5.4, P < 0.001, n= 3) (Figs 1B and 2). A Hill curve fitted to the mean data (Fig. 2) indicated that 50% uncoupling occurred with a 3 min exposure to 1.9 µM Ca²⁺ (pCa 5.72). These data demonstrate that the EC uncoupling occurs at physiological intracellular Ca²⁺ levels.

View larger version (11K): [in this window] [in a new window]   Figure 2.  Ca2+ dependence of EC uncoupling Mean size (±S.E.M.) of depolarization-induced force response after a 3 min exposure at the indicated [Ca2+] (in pCa units), expressed as a percentage of the response before treatment. Ca2+ was applied during rigor as in Fig. 1, and n is the number of fibres. The data were fitted with a Hill curve with 50% inhibition at pCa 5.72 (i.e. 1.9 µM Ca2+) and a Hill coefficient of 3.5.

Effects of Ca²⁺ exposure on passive force production

The effect of raised [Ca²⁺] on passive force production in stretched fibres was used as an assay of whether proteases present in the fibres were activated by the Ca²⁺ treatment. Stretching the skinned fibre segments to 2.0 times resting length resulted in large passive force (Fig. 3). As seen previously in frog fibres (Higuchi, 1992), passive force did not stay constant upon stretching but instead declined over time, initially quite rapidly over the first 30 s, and then at a progressively slower rate (e.g. see start of Fig. 3A). When a fibre was kept stretched and then exposed to control-rigor solutions (pCa 8) for 4.5 min (e.g. Fig. 3A), it had no noticeable effect on the time course of the force and the peak of the passive force produced when subsequently shortening and re-stretching the fibre to same final length (i.e. 2.0 times original resting length) was very similar to that when first applying the stretch (mean 94 ± 3%, n= 10; Fig. 4). In contrast, when the [Ca²⁺] in the rigor solution was 8 or 40 µM (e.g. Fig. 3B), force declined substantially during the exposure, and the peak of the passive force upon re-stretching the fibre was greatly reduced (to 77 ± 3 and 31 ± 4% of initial value for 8 and 40 µM Ca²⁺, respectively; Fig. 4). This decline in passive force was irreversible; the peak of the passive force remained virtually unchanged at the low level when the same stretch was applied after the fibre had been kept at resting length for a further 10 min (not shown). If 1 mM leupeptin was present in the 40 µM Ca²⁺-rigor solution, the passive force only decreased by 15 ± 5% of the initial level (Fig. 4), which was significantly less reduction than with 40 µM Ca²⁺ exposure in the absence of leupeptin (P < 0.05) and not significantly different from the contol-rigor case.

View larger version (15K): [in this window] [in a new window]   Figure 4.  Summary of effects of treatment on passive force Mean (±S.E.M.) of peak of passive force following indicated treatment, expressed as a percentage of that before the treatment. Experiments were performed as shown in Fig. 3, with a 3 min exposure to the [Ca2+] indicated. The number of fibres is shown for each mean. Each fibre was stretched to 2.0 times resting length during the exposure except in case indicated (far right). leu, 1 mM leupeptin. *Significantly less than 100%. **P < 0.01 and ***P < 0.001, mean is significantly less than for the control-rigor case.

The peak of the passive force reached when stretching a fibre to 2.0 resting length was typically slightly larger than the maximum active force expected for the respective fibre cross-sectional area. In general, maximum Ca²⁺-activated force was not assessed in the above passive force experiments, both to avoid additional exposure to elevated [Ca²⁺], and also because in every case where a fibre was exposed to the 40 µM Ca²⁺-rigor solution for more than 1 min whilst stretched, it invariably broke before reaching maximum Ca²⁺-activated force. Such breakage was extremely rare in all other circumstance (i.e. if the fibre was not stretched during the Ca²⁺ exposure, or if leupeptin was present with the Ca²⁺, or if the [Ca²⁺] was never raised).

Rate of decline of force in stretched fibres in rigor solutions

The rate of decline of force in stretched fibres exposed to rigor and Ca²⁺-rigor solutions was used as a sensitive assay of changes in passive force. This avoided the problems involved in shortening and re-stretching a fibre to a given precise length, as well as those due to changes in fibre properties occurring over time. In these experiments, the fibre length was kept constant at 2.0 times resting length, and the rate of force decline in various conditions was compared with that in control-rigor solution (pCa 8) measured immediately before and afterwards (see Fig. 5A). The rate of decline of force during each rigor solution condition was expressed as a percentage of the force just prior to initially inducing rigor (percentage force per minute) and also expressed relative to that measured during the control-rigor periods in the same fibre.

When a stretched fibre was exposed to a rigor solution with elevated [Ca²⁺], the force always declined much more rapidly than during the control-rigor period (e.g. Fig. 5B). The rate of decrease in passive force showed a clear [Ca²⁺] dependence, with a significant effect being apparent even at 1.2 µM free [Ca²⁺] (Fig. 6), where the rate of decline increased twofold compared with control level (P < 0.05, paired difference with control-rigor period, n= 6).

View larger version (16K): [in this window] [in a new window]   Figure 6.  Effect of various treatments on the rate of decline of passive force A, mean (±S.E.M.) rate of decline of passive force for each given condition, expressed relative to the control-rigor period in the same fibre (see Fig. 5). The number of fibres is shown with each mean. 1.2, 8 and 40 µM denote the free [Ca2+] for the 3 min exposure period. leu and calp, the presence of 1 mM leupeptin and 10 µM calpastatin, respectively. 40 µM+ DTDP, pretreatment of the fibre with dithiodipyridine before stretching and treating it with 40 µM Ca2+. Homogenate, exposure to Ca2+-activated homogenate with final [Ca2+] of 0.4 µM. B, data in A expressed as absolute values of force decline per minute, with each fibre standardized by the force level immediately prior to the first exposure to rigor solution, and with the paired rate of decline during the control-rigor subtracted from it. In some instances the fibres included in B were not included in the normalized data in A because force decline during the control rigor period was approximately zero. *Significantly steeper decline than paired control-rigor period (P < 0.05). #Significantly smaller rate of decline than with 40 µM Ca2+ without calpastatin (P < 0.05).

Effect of calpain inhibitors.  The effect of 8 and 40 µM Ca²⁺ in increasing the rate of decline of passive force was completely abolished by adding 1 mM leupeptin to the rigor solutions (e.g. Fig. 5C). For both concentrations of Ca²⁺, the rate of force decline during the Ca²⁺-rigor period with leupeptin present was not significantly different from that during the corresponding control-rigor period with leupeptin present (P > 0.2, Fig. 6). When the leupeptin was washed out and the fibre again exposed to elevated [Ca²⁺] (40 µM), the rate of force decline increased in every fibre examined (e.g. second 40 µM Ca²⁺-rigor period in Fig. 5C), demonstrating that Ca²⁺ indeed had a detrimental effect in these fibres if leupeptin was absent.

Calpastatin inhibits calpain-1 and calpain-2, but not calpain-3. In contrast to leupeptin, the presence of a relatively high concentration of calpastatin (10 µM) in the rigor-solutions did not prevent the deleterious effect of elevated [Ca²⁺] on passive force (Fig. 6, ‘40 µM+ calp’).

The rate of force decline during the 40 µM Ca²⁺-rigor period with calpastatin present was on average 4.3 times that during the control-rigor period measured in the same fibre (Fig. 6A). This was not significantly different from that found with 40 µM Ca²⁺ in the absence of calpastatin (Fig. 6A). However, if the rate of passive force decline is expressed in absolute terms (percentage initial force per minute) (see Fig. 6B), the decline in 40 µM Ca²⁺ with calpastatin was significantly smaller than that without calpastatin (P < 0.05). Thus, the presence of a high concentration of calpastatin seems to reduce the effect of Ca²⁺ on passive force to some extent, though it was by no means a potent inhibitor.

Effect of oxidation with DTDP.  In fibres that were pretreated for 5 min in relaxing solution with 100 µM dithiodipyridine (DTDP), a highly reactive disulphide (Zaidi et al. 1989), and then stretched to 2.0 times resting length, there was no Ca²⁺-dependent decline in passive force; the rate of force decline during exposure to 40 µM Ca²⁺ was indistinguishable from the rate during control-rigor (n= 6, P > 0.5) (Fig. 6). When the oxidative effect of DTDP was reversed by treating the same fibres with the reducing agent DTT (10 mM for 5 min) (see Posterino & Lamb, 1996; Lamb & Posterino, 2003), passive force was rendered again sensitive to elevated [Ca²⁺], with the decline in force in 40 µM Ca²⁺ being significantly greater than that observed before the DTT treatment (1.9 ± 0.4 times greater, n= 6, P < 0.005).

Effect of membrane removal with Triton X-100.  Some skinned fibres were pretreated with Triton X-100, in order to have the contractile apparatus devoid of membranes. These fibres still showed a significant increase in the rate of decline of passive force during subsequent exposure to 40 µM Ca²⁺-rigor solution whilst stretched (2.6 ± 0.6 times and 5.5 ± 1.4% min^–1 greater than control rigor, both P < 0.05, n= 4). This effect was substantially less than that occurring in freshly mounted fibres exposed to 40 µM Ca²⁺ (rate of decline: 6.1 ± 0.9 times and 21.2 ± 4.2% min^–1 greater than control-rigor, n= 9; Fig. 6), but much of this difference was attributable simply to the longer washing period before the Ca²⁺ treatment, as it was found that the Ca²⁺ treatment was also less effective in paired fibres given the same duration (10 min) of extra prewashing in relaxing solution (rate of decline: 4.0 ± 0.6 times and 9.1 ± 1.4% min^–1 greater than control-rigor, n= 4). These results indicate that at least some of the Ca²⁺-dependent factor involved in the reduction of passive force is bound to contractile or structural proteins and is only gradually lost from the skinned fibres over tens of minutes.

Effect of Ca²⁺ exposure on active force and Ca²⁺-sensitivity of the contractile apparatus

In order to examine whether the Ca²⁺ exposure deleteriously affected active force production by the contractile apparatus, skinned fibres were given repeated 1 min periods of maximum Ca²⁺ activation in an ATP-containing solution. Fibres were stretched to 1.2 times resting length, the same as in the EC coupling experiments, and exposed repeatedly to the maximum activation solution (free [Ca²⁺] very heavily buffered at 20 µM (pCa 4.7)). Maximum Ca²⁺-activated force declined substantially over the course of the first 1 min activation period (e.g. Fig. 7A). In the six fibres examined, the mean total decline over this first 1 min period was 25 ± 1%, expressed as a percentage of the initial level, with the majority of this (20 ± 1%) occurring over the first 30 s. In contrast, when a second 1 min maximal activation period was imposed after keeping the fibre relaxed for 2 min, force initially peaked at just below the level reached at the end of the first activation period, and then showed almost no further decline over the activation period (e.g. Fig. 7A). This effect can also be seen in Fig. 7C, where the peak of the force at the start of each maximal activation period is shown for fibres that were subjected to five or more such periods; the initial peak force was greatly reduced in the second period compared with the first period, but then it decreased relatively little over the subsequent periods. The presence of leupeptin reduced the decline in active force occurring during the first 1 min maximal activation period (e.g. Fig. 7B). (The rise in baseline force after the prolonged activation in Fig. 7B is commonly seen in toad fibres at this temperature (Rees & Stephenson, 1987) and is not an effect of leupeptin). With 0.5 mM leupeptin present, the decline in force during the first period of maximal activation was 9 ± 3 and 12 ± 2% over the first 30 s and 60 s, respectively, in the six fibres examined. These values were approximately half of that occurring in fibres in the absence of leupeptin present (P < 0.01) (also see Fig. 7C).

View larger version (13K): [in this window] [in a new window]   Figure 7.  Leupeptin reduces the initial decay in maximal active force A, force responses to first two periods in maximal Ca2+-activation solution (pCa 4.7, 50 mM CaEGTA-EGTA). The skinned fibre was at 120% of resting length, and had not been activated or exposed to any Ca2+ beforehand. The fibre was relaxed in relaxing solution (pCa > 9) and transferred to solution with 0.5 mM EGTA (pCa 8) for 30 s before each activation period. Initial peak force defined as 100%. B, similar procedure in another fibre with 0.5 mM leupeptin present in all solutions, showing smaller decline in force during the first activation period. (See text regarding change in baseline force). C, mean relative force (±S.E.M.) reached at the start of each successive 1 min maximal activation period. The force is expressed relative to the initial peak force reached during the first activation period. Open and filled symbols indicate presence and absence of 1 mM leupeptin, respectively (4 fibres each).

Calpain-3 in muscle homogenates and skinned fibres

Exposure of stretched fibres to a muscle homogenate that had been preactivated with Ca²⁺ (final [Ca²⁺]0.4 µM) increased the relative rate of decline of passive force to 2.3 times of that occurring in control-rigor solution (Fig. 6A). This deleterious effect on passive force was considerably larger in absolute terms than that seen when applying the 1.2 µM free-Ca²⁺ treatment (Fig. 6B), despite the [Ca²⁺] being much lower, indicating that there was some activated diffusible substance with proteolytic activity present in the homogenate. This diffusible substance was likely to be a calpain, because addition of 1 mM leupeptin to the activated homogenate completely inhibited the effect of the homogenate (Fig. 6).

Western blots were used to determine whether calpain-3 was present in the muscle homogenate. As calpain-3 protein has not previously been reported in toad skeletal muscle, we compared protein bands identified using the anticalpain-3 antibody in rat and toad skeletal muscle (Fig. 8A). In mammalian muscle calpain-3 is a 94 kDa protein which breaks down via autolysis to proteins of 60–55 kDa (Kinbara et al. 1998; Branca et al. 1999; Taveau et al. 2003). Previous characterization of the monoclonal 12A2 anticalpain-3 antibody showed that in addition to protein bands at 94 and 58 kDa, Western blotting of muscle homogenates from rat, mouse, hamster and chicken detects a band at 82 kDa (Anderson et al. 1998; Kramerova et al. 2004). We identified two bands (at 94 and 82 kDa) in rat skeletal muscle homogenate, and corresponding bands were observed in toad muscle. The 94 kDa band disappeared following treatment with Ca²⁺ in skeletal muscle homogenates from both rat (data not shown) and toad (Fig. 8A). In toad preparations, we consistently found some of the 60 kDa protein band in control muscle preparations. The intensity of this band seemed augmented when the muscle had been exposed to Ca²⁺, suggesting that it is the autolysed calpain-3. The 82 kDa band did not appear to be sensitive to the Ca²⁺ treatment in either toad muscle (Fig. 8A) or rat muscle.

View larger version (52K): [in this window] [in a new window]   Figure 8.  Calpain-3 Western blots of muscle homogenates and skinned fibres A, muscle homogenates from rat muscle under control conditions (Rat CON) and toad muscle under control conditions (Toad CON) or after exposure to 5 mM Ca2+ for 1 min (Toad Ca2+) were analysed for calpain-3. The 94 kDa band disappeared after exposing toad muscle to Ca2+. B, groups of three mechanically skinned single fibres were exposed to 40 µM Ca2+ for 1 min, 8 µM Ca2+ for 3 min and 2 µM Ca2+ for 3 min or to the control-rigor solution (Rig) under paraffin oil. Each fibre had been initially washed in the same droplet of control-rigor solution (wash) for 1–7 min (time matched for all fibre groups); proteins found in this solution were those readily washed out of the 12 fibres. The density of the 94 kDa calpain-3 band in lanes 1–4 was normalized by the 42 kDa protein present (likely to be actin, relative density range 70–100%), and expressed as a percentage of that for the control-rigor treatment was 100, 19, 14 and 0% for control rigor, 2, 8 and 40 µM Ca2+, respectively. Whole, homogenate of toad whole muscle. Note that the sensitive streptavidin method required with the single fibres in B resulted in nonspecific detection of additional protein bands.

Titin

Separation of titin from toad muscle resulted in a band of similar size to that seen in rat soleus muscle (Fig. 9A). Full-length titin (T1) readily degraded to a smaller form (T2) in both rat and toad muscle (Fig. 9B and C). There was some degradation of titin in all toad single fibres mounted on the transducer and exposed to the control or Ca²⁺-rigor solutions for 3 min, but this was greatest in fibres that were exposed to 40 µM Ca²⁺ whilst being stretched (Fig. 9D and E). When leupeptin (1 mM) was present in the 40 µM Ca²⁺ rigor solution, the degradation of titin on average was no different from that in fibres exposed to the control-rigor solution. In contrast to titin, there was no obvious change in the size or density of the nebulin band in the fibres investigated. In some of the fibres treated whilst stretched with 40 µM Ca²⁺, a new band appeared on the gel approximately midway between the titin and nebulin (not shown), similar to that seen with titin degradation in rabbit and chicken muscle (Matsuura et al. 1991; Ohtsuka et al. 1992). Attempts to examine the amount of titin degradation in fibres given longer exposure to 40 µM Ca²⁺ (e.g. 5 min) were unsuccessful, as these fibres always broke whilst stretched on the transducer.

View larger version (45K): [in this window] [in a new window]   Figure 9.  Silver stained 2.8% SDS-PAGE gel of skeletal muscle A, homogenates from rat soleus and toad iliofibularis muscle. Full-length titin (T1, 3700 kDa) is similar in size in both samples, whereas nebulin (N) in rat soleus muscle (800 kDa) is slightly larger than that in toad iliofibilaris muscle (700 kDa). B, when toad muscle was warmed and left at room temperature for a period (RT, see Methods), much of the titin present in control conditions (Con) degraded to titin 2 (T2). C, similar results in rat muscle. D, single skinned fibres from toad iliofibilaris muscle that had been exposed to control-rigor (Rig), 40 µM Ca2+ (Ca2+) or 40 µM Ca2++1 mM leupeptin (Leu) for 3 min whilst stretched on the force transducer at 2.0 resting length (see Methods and Fig. 3). The ratio of T1 to T2 was greatest for the control-rigor case and least for the fibre exposed to 40 µM Ca2+. E, mean densitometry values (±S.E.M.) of the T2 band (as a percentage of T1 + T2) in single skinned fibres (number in parentheses) exposed as in D, with ‘unstretched’ denoting single fibres exposed to Ca2+ as in Fig. 3C. *Significant difference compared to Rig and Leu (one-way analysis of variance, P < 0.05).

	Discussion

Top Abstract Introduction Methods Results Discussion References

Passive force production in stretched fibres of toad muscle declined irreversibly when the cytoplasmic [Ca²⁺] was raised above 1 µM (Figs 3–6). This occurred independently of the possible small reversible effects that Ca²⁺ may have on titin (Labeit et al. 2003) and on actin–titin interactions (Kulke et al. 2001). The decline of passive force was evidently caused by proteolysis of titin (Fig. 9) by one or more types of endogenous calpain, as indicated by the following: firstly, calpains are known to proteolyse titin, and the effects here were observed at a [Ca²⁺] (1 µM and above) that is just sufficient to activate two of the calpains present in skeletal muscle, calpain-1 (Goll et al. 2003) and calpain-3 (also called p94) (Sorimachi et al. 1993; Branca et al. 1999; Ono et al. 2004) (calpain-2 requires much higher [Ca²⁺] for activation), and the effect on passive force could be stopped and then restarted by removing and reapplying Ca²⁺ in a manner consistent with the Ca²⁺ dependency of calpain activity (Goll et al. 2003; Diaz et al. 2004); secondly, the deleterious effect of raised [Ca²⁺] could be completely stopped as follows: (i) by 1 mM leupeptin (Figs 4–6), which is known to fully inhibit calpain-1 and calpain-2 (Goll et al. 2003), and also to inhibit calpain-3 once it has been activated by Ca²⁺-dependent autolysis (Ono et al. 2004; Diaz et al. 2004), and (ii) by oxidizing free sulphydryls with DTDP (Fig. 6), a treatment that should render inactive any cysteine proteases such as calpain, provided that the oxidant has access to the free cysteine at the active site, as is the case at least for calpain-1 (Inomata et al. 1984); finally, it was shown that calpain-3 was present in both the skinned fibres and the muscle homogenate and that it was activated by the Ca²⁺ treatment (Fig. 8), and that the homogenate contained a readily diffusible Ca²⁺-activated factor that evidently proteolysed titin and could be inhibited by leupeptin (Fig. 6).

The proteolysis of titin was probably mediated primarily by calpain-3 rather than calpain-1, as indicated by the following: (i) calpain-3 is 10-fold more abundant in skeletal muscle than the other calpains (as judged by mRNA levels, Kinbara et al. 1997); (ii) the threshold [Ca²⁺] for initially activating calpain-3 is slightly lower [Ca²⁺] than that for calpain-1 (0.5 µMversus3 µM; Branca et al. 1999; Goll et al. 2003; Ono et al. 2004); and (iii) the loss of passive force was not greatly reduced by a relatively high concentration of calpastatin (Fig. 6), which is known to fully inhibit calpain-1 (Uemori et al. 1990; Goll et al. 2003) but have little or no effect on calpain-3 even in its activated state (Ono et al. 2004). In mammalian muscle, calpain-3 is predominantly bound to the N2 region of titin (Sorimachi et al. 1995; Keira et al. 2003), which is located on either side of the Z-disk near the A–I boundary in close proximity to the position of the triad junctions in mammalian muscle (Franzini-Armstrong & Jorgensen, 1994); calpain-1 binds in the Z-disk region and I-band (Kumamoto et al. 1992; Goll et al. 2003). The location of calpain binding in amphibian muscle fibres is not known. In the toad fibres here, most of the calpain-3 was bound rather than free in the cytoplasm (Fig. 8), and Triton X-100 treatment indicated that much of the calpain was bound to structural proteins rather than membranes.

It has been reported that titin in both mammalian and chicken muscle degrades from the full-length T1 form (also called -connectin) to a smaller T2 form (ß-connectin) when the muscle is kept for long periods, and that this occurs by cleavage in the I-band near the N2 region (Matsuura et al. 1991; Ohtsuka et al. 1992; Kimura et al. 1992). Exogenous calpain-2 causes similar proteolysis of titin, with longer treatment resulting in further degradation of titin and also degradation of nebulin (Hu et al. 1989). Significantly, Ca²⁺ exposure without exogenous calpain caused degradation of titin from the T1 to the T2 form, and had no effect on nebulin (Hu et al. 1989). This fits well with our findings (Figs 8 and 9) as it can be explained by the Ca²⁺ activating an endogenous calpain, which is likely to be calpain-3 bound at the N2 line, which preferentially cleaves titin at a site nearby. Recently, Kramerova et al. (2004) have shown that calpain-3 cleaves titin at the PEVK region adjacent to the N2 line. This is also relevant to our finding that the loss of passive force production was far greater if the fibre was stretched during the Ca²⁺ exposure than if it was returned to close to resting length (Figs 3 and 4). As the PEVK region is elongated during large stretches (Gautel & Goulding, 1996; Linke et al. 1996), it could be that such stretch renders it more susceptible to being proteolysed by calpain-3. Ca²⁺ activation of calpain-3 itself does not require stretching of the fibre (Fig. 8).

Ca²⁺-dependent loss of EC coupling

Prolonged exposure to [Ca²⁺] above 1 µM also caused an irreversible interruption to EC coupling in the toad fibres here (Figs 1 and 2). It seems very likely that this effect is also the result of the activation of endogenous calpains because of the following: (i) the evidence given above shows that calpains are indeed activated within the fibre over this exact [Ca²⁺] range; (ii) the progression of Ca²⁺-dependent uncoupling can be stopped by lowering the [Ca²⁺] and restarted again by raising the [Ca²⁺] (data not shown), as found with titin proteolysis by calpains; (iii) the uncoupling can be inhibited by leupeptin, provided the activating [Ca²⁺] is relatively low (e.g. 2 µM); (iv) the uncoupling is slowed by more than 10-fold by lowering the temperature from 23 to 3°C, and by more than threefold by reducing the pH from 7.1 to 5.8 (Lamb et al. 1995), consistent with effects of such changes on calpains (Ono et al. 2004); and (v) the uncoupling is triggered by Sr²⁺ at 10-fold higher concentration than by Ca²⁺ (Lamb et al. 1995), consistent with the known Sr²⁺ activation characteristics of calpains (Inomata et al. 1984; Sargianos et al. 1995).

The above points together strongly suggest that calpain(s) have some role in the Ca²⁺-dependent uncoupling in toad fibres. Chin & Allen (1996) suggested that calpains are not involved in the uncoupling process because they found that the membrane-permeable calpain inhibitor, calpeptin, did not prevent a reduction in Ca²⁺ release in intact murine fibres subjected to a treatment involving repeated stimulation in the presence of caffeine. However, it seems that the calpeptin itself may have had direct effects on Ca²⁺ movements in the fibres because the resting [Ca²⁺] rose significantly during the treatment in calpeptin (Chin & Allen, 1996), and it was found here to have direct inhibitory effects on EC coupling in the skinned fibres. Consequently, the reduced Ca²⁺ release that Chin & Allen (1996) observed following the high [Ca²⁺]–calpeptin treatment was possibly a direct effect of the calpeptin itself.

A further line of evidence implicating Ca²⁺-dependent proteolysis as the cause of the uncoupling is that the treatment appears to cause physical damage to the triad junction. Electron microscopy of skinned fibres given a brief (10 s) Ca²⁺-treatment sufficient to cause full uncoupling, showed that the triad junctions were distorted or in some instances completely severed (Lamb et al. 1995). However, the protein target involved is not known. It is not the Ca²⁺ release channel ryanodine receptor (RyR), the voltage-sensor (dihydropyridine receptor) or triadin, as these proteins were not proteolysed in fibres uncoupled by the Ca²⁺ treatment (Lamb et al. 1995). Evidently it is a protein that holds the triad junction together in its normal shape, perhaps being important for keeping the voltage sensors in the T-system in close apposition with the Ca²⁺ release channels in the SR.

Comparison of Ca²⁺ effects on EC uncoupling and titin proteolysis

Although prolonged Ca²⁺ exposure can lead eventually to many structural changes in muscle fibres (Duncan, 1987; Kasuga & Umazume, 1990), and perhaps involves the proteolysis of many different proteins, it seems that voltage sensor coupling to Ca²⁺ release is one of the first processes affected. At a given [Ca²⁺] complete uncoupling occurred considerably more rapidly than did the loss of passive force production; depolarization-induced force responses were fully abolished by a 3 min exposure to 3.5 µM Ca²⁺ (Fig. 2), whereas interpolation of the data shown in Fig. 6B indicates that that [Ca²⁺] only reduced passive force in stretched fibres by 12% in 3 min (i.e. at a rate of 4% min^–1). The loss of EC coupling was also more steeply dependent on the [Ca²⁺] than was the loss of passive force (compare Fig. 2 with effect of 1.2–40 µM Ca²⁺ seen in Fig. 6B). These data indicate that raised [Ca²⁺] causes a proportionately faster and greater level of proteolytic damage to the triad junction than it does to titin, even when the fibre is stretched and titin is most susceptible. This might be the result of calpains being initially bound close to the triad junction, as is apparently the case at least in mammalian muscle fibres (see above). Ca²⁺ causes autolytic activation of single calpain-3 molecules, which can then proteolytically activate other nearby calpain-3 molecules in a strongly self-reinforcing activation cascade (Taveau et al. 2003). The activation cascade behaviour of calpain-3 may also explain why it is not possible to stop EC uncoupling with leupeptin when the [Ca²⁺] is appreciably above the threshold level for calpain activation; it would only be possible to stop such an ‘explosive’ self-reinforcing effect if initial Ca²⁺ activation event proceed slowly so that there was a sufficient chance for the leupeptin to bind to the small number of activated calpain-3 molecules before they had time to activate neighbouring calpain-3 molecules. Here it is important to bear in mind that calpain-3 cannot bind leupeptin at its active site until it has been activated (Diaz et al. 2004), and so pretreatment with leupeptin has no effect on the pool of calpain-3 available for activation.

Physiological relevance

The Ca²⁺-dependent loss of EC coupling reported here seems likely to be important in normal muscle function and in muscle diseases where intracellular [Ca²⁺] is raised excessively. It was found here that the [Ca²⁺] reached during a tetanus is sufficient to activate calpains present endogenously in the fibres, and hence if the [Ca²⁺] is raised for a sufficiently long period there will be an appreciable level of proteolytic disruption of the triad junction and a consequent decrease in EC coupling. It is possible that chaperone molecules normally present in intact fibres could be washed out of the skinned fibres leaving them more susceptible to Ca²⁺-dependent damage. Nevertheless, our previous study showed that such damage still occurs if the intracellular [Ca²⁺] is raised whilst the fibres are intact (Lamb et al. 1995). Moreover, a similar Ca²⁺-dependent uncoupling has also been reported in intact murine fibres (Chin & Allen, 1996; Chin et al. 1997), where a 45% reduction in Ca²⁺ release and a 70% reduction in force were found after repeated tetani that elevated the intracellular [Ca²⁺] to 2 µM for a total period of 41 s, which is quite comparable to the uncoupling found in the toad skinned fibres here (Fig. 2). The present findings further indicate that the uncoupling proceeds proportionally more rapidly at higher [Ca²⁺], consistent with the properties of calpain activation over this [Ca²⁺] range (Ono et al. 2004). Depolarization-induced responses were reduced by 50% with a 180 s exposure to 1.9 µM Ca²⁺ (Fig. 2), whereas when the [Ca²⁺] was raised to 10 µM it only required 10 s exposure for such uncoupling (Lamb et al. 1995). Thus, raising the [Ca²⁺]fivefold in this range (from 1.9 to 10 µM) causes uncoupling to proceed 18 times faster. This indicates that the peak [Ca²⁺] reached in a fibre during a tetanus is likely to be far more important in causing uncoupling than is any increase in resting [Ca²⁺]. This Ca²⁺-dependent uncoupling seems to account for the phenomenon of low-frequency fatigue that occurs when muscles are activated repeatedly for prolonged periods (Edwards et al. 1977; Chin & Allen, 1996). We suggest that the uncoupling phenomenon could function as an important protective feedback mechanism in muscle fibres, interrupting the coupling and Ca²⁺ release at any triad junction where Ca²⁺ release is excessive or too prolonged, thereby preventing widespread Ca²⁺-activated proteolytic damage to other structural elements in the fibre that may otherwise ensue. The finding that titin was most susceptible to proteolysis when the fibre was stretched to twice resting length suggests that during normal concentric contractions in vivo, titin may be relatively protected from proteolytic damage. However, this property could be particularly important for the remodelling of ‘popped’ (elongated) sarcomeres that occur with eccentric muscle damage (Morgan & Proske, 2004).

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