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SKELETAL MUSCLE AND EXERCISE |
1 Laboratory of Cell Physiology, Université catholique de Louvain (UCL5540), Brussels B-1200, Belgium
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Abstract |
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(Received 9 June 2006;
accepted after revision 3 July 2006;
first published online 6 July 2006)
Corresponding author P. Gailly: Laboratory of Cell Physiology, Université catholique de Louvain, UCL/FYCL 5540 av. Hippocrate, 55, B-1200 Brussels, Belgium. Email: gailly{at}fycl.ucl.ac.be
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Introduction |
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So far, the TRP channels identified as possibly involved in store-operated influxes of Ca2+ belong to the TRPC and TRPV subfamilies. Indeed, using a methodology of heterologous expression, TRPC1, 2, 3, 4 and 5, and TRPV6, have been shown to be possibly activated by store depletion (Groschner et al. 1998; Philipp et al. 1998; Vannier et al. 1999; Warnat et al. 1999; Liu et al. 2000; Yue et al. 2001). However, other authors showed that heterologous expression of these proteins also gave channels whose activation was independent of store depletion (Zitt et al. 1997; Hofmann et al. 1999; Schaefer et al. 2000; Wu et al. 2000; Bodding et al. 2003). This might be due to the fact that the mechanism of channel activation depends on their expression level (Vazquez et al. 2003). Strategies using repression with antisense oligonucleotides were then used to demonstrate the possible involvement of TRPC1, 2 and 3, and TRPC4, in store-operated entry of Ca2+ (Philipp et al. 2000; Wu et al. 2000; Gailly & Colson-Van Schoor, 2001). Finally, the involvement of TRPC4 in store-operated Ca2+ current was demonstrated by showing that mice deficient in TRPC4 lack this current in endothelial cells and exhibit reduced vasorelaxation (Freichel et al. 2001).
In a previous work, we have shown that store-dependent channels are present in skeletal muscles and that their activity is abnormally increased in Duchenne muscular dystrophy (Vandebrouck et al. 2002), a myopathy due to the lack of a cytoskeletal protein called dystrophin.
TRP channels also respond to stimuli other than store depletion. Indeed, TRPC proteins have been shown to respond to agonists independently of store depletion, and TRPV proteins are sensitive to heat and cold, pH changes, osmolarity or volume changes, or are constitutively active and responsible for Ca2+ reabsorption in kidney and duodenum (TRPV5 and 6) (reviewed in Benham et al. 2002; Minke & Cook, 2002). In the present study, we characterize the pharmacological profile and the different modes of activation of voltage-independent Ca2+ channels present in adult skeletal muscle fibres, and we assess their possible involvement in the normal mechanical response of muscle to electrical stimulation.
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Methods |
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Adult wild-type (C57) mice were deeply anaesthetized with a solution (10 ml kg–1, intraperitoneally) containing ketamine (10 mg ml–1) and xylazine (1 mg ml–1) in order to preserve muscle perfusion during dissection of both soleus muscles. Depth of anaesthesia was assessed by the abolition of eyelid and pedal reflexes. After dissection, the animals were killed by rapid neck dislocation. This protocol has been approved by the Animal Ethics Committee of the Catholic University of Louvain, Brussels. Soleus muscles were bathed in a 1 ml horizontal chamber continuously superfused with oxygenated Krebs solution (O2 95%/CO2 5%) containing (mM): NaCl 118, NaHCO3 25, KCl 5, KH2PO4 1, CaCl2 2.5, MgSO4 1, glucose 5, maintained at a temperature of 20 ± 0.1°C. One end of the muscle was tied to an isometric force transducer, and the other end to an electromagnetic motor and length transducer. Stimulation was delivered through platinum electrodes running parallel to the muscles. Muscle length was carefully adjusted for maximal isometric force using 0.35 s maximally fused tetani. Force was recorded on a high-speed pen recorder (Sanborn model 320).
Isolation of adult skeletal muscle fibres
The flexor digitorum brevis (FDB) muscles were removed and incubated for 38 min at 37°C in an oxygenated ‘Krebs–Hepes’ solution (see composition below) containing 0.2% collagenase type IV (Sigma, St Louis, MO, USA). Muscles were then washed twice in Krebs buffer, suspended in DMEM/HAM F12 (Sigma) supplemented with 2% fetal bovine serum (Sigma), and mechanically dissociated by repeated passages through fire-polished Pasteur pipettes of progressively decreasing diameter. Dissociated fibres were plated onto tissue culture dishes coated with extracellular matrix basement membrane (Harbor Bio-Products, Norwood, MA, USA) and allowed to adhere to the bottom of the dish for 2 h. For Ca2+ measurements, cells were plated on circular glass coverslips. Culture dishes were kept in an incubator, with 5% CO2 at 30°C.
Measurements of cytosolic [Ca2+]
Muscle fibres were loaded for 1 h at room temperature with the membrane-permeant Ca2+-indicator Fura-PE3/AM (1 µM) and Pluronic F-127 (0.004%). Fura-PE3/AM was preferred to Fura-2/AM as it is stable during long-lasting experiments, with little or no compartmentation (Vorndran et al. 1995). Fibres were illuminated through an inverted Nikon microscope (x40-magnification objective) alternately at 340 and 380 nm, and the fluorescent light emitted at 510 nm was measured using a Deltascan spectrofluorimeter (Photon Technology International). The ratio R340/380 of the fluorescence intensity emitted at the two excitation wavelengths was calculated, and cytosolic concentration of Ca2+ ([Ca2+]i) was determined with a calibration previously described (Vandebrouck et al. 2002). The Krebs–Hepes solution contained (mM): NaCl 135.5, MgCl2 1.2, KCl 5.9, glucose 11.5, Hepes 11.5, CaCl2 1.8 (pH 7.3). When necessary, CaCl2 was omitted and replaced by 50 µM sodium EGTA, and osmolarity was adjusted with sucrose. The potassium aspartate solution contained (mM): potassium aspartate 150, MgCl2 5, EGTA 10 and Hepes 10 (pH 7.3).
Electrophysiological methods
Single-channel activity was recorded from cell-attached patches using the technique described by Hamill et al. (1981). Patch electrodes were pulled on a DMZ-Universal (Zeitz-Instruments) puller in three stages from borosilicate glass capillaries (1.5 mm in diameter; Harvard Apparatus) to a tip diameter of 1–2 µm. Patch electrodes had a resistance of 2–5 M
. Cells were viewed under phase contrast with a Diaphot Nikon inverted microscope. The activity was recorded at a constant holding potential of –60 mV and at room temperature, using a HEKA EPC-9 amplifier. This holding potential value takes into account the basal membrane potential (measured independently at –50 mV) (Cahalan & Neher, 1992). Current records were filtered with a Bessel filter at 3 kHz and digitized at 10 kHz. Data were analysed using Pulse-Fit, Pulse-Tools and Origin 6.1 software. Most of the patches contained more than one channel. Therefore, the global open probability (n.Po) was calculated (ratio of total open time to total time for a given patch).
The intrapipette solution contained (mM): CaCl2 110, Hepes 10 and DIDS (4,4'-diisothyocyanatostilbene-
2,2'-disulphonic acid) 0.1. The bathing solution contained (mM): NaCl 124, MgCl2 1.2, KCl 5.9, glucose 11.5, HepesNa 11.5, EGTA 10. The osmolarity (measured with a microosmometer type 13/13DR Roebling) of these solutions was adjusted to 320–330 mosmol l–1 by adding water or sucrose, and adjusted to pH 7.3 with NaOH. DIDS was used to block possible chloride conductances.
Thapsigargin (TG) was dissolved in DMSO and diluted 1:2000 into the bath to a final concentration of 1 µM. Channels activity was recorded during 90 s (30 sweeps of 3 s) in the absence of TG (control condition) and was prolonged for 3–5 min after application of the drug on the same patch. The solvent alone had no effect on channel activity. Mechanical stimulation was performed by applying on the patch pipette a suction the pressure of which was measured with a mercury-filled U-shape manometer. Channel mechanosensitivity was therefore only studied under stationary conditions. As the response to stretch was fast, channels activity was only recorded during 30–60 s for each chosen pressure (one or two series of 10 sweeps of 3 s).
Reagents
The GsMTx4 toxin, isolated from Grammostola spatulata spider (Suchyna et al. 2000), was obtained from Peptides International (Louisville, KY, USA); SKF-96365 and 2-APB were from Alexis Corporation (Lausen, Switzerland); and Fura-PE3/AM was from Calbiochem (Darmstadt, Germany). All other reagents were of analytical grade and purchased from Sigma. Channel inhibitors were added to the pipette solution in order to have access to the external face of the channel in cell-attached configuration.
Statistics
Data are presented as means ±
S.E.M.). The 
2 test, Fisher exact test, ANOVA and Student's t test were used to determine statistical significance.
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Results |
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In a previous paper we showed the presence, in muscle fibres, of voltage-independent Ca2+ channels that are activated by store depletion. Under the conditions used here (cell-attached configuration, muscle fibres in physiological medium), we detected a basal Ca2+ channel activity with a unitary conductance of 7.4 ± 0.47 pS (n = 7, in 110 mM Ca2+) and a reversal potential at +43 mV (Fig. 1A and B). TG was used to deplete the stores by inhibiting the sarco-endoplasmic reticulum Ca2+-ATPase (SERCA). After 90 s of patch recording under control conditions, 1 µM TG was added to the bath and recording was prolonged by 3 min. As previously shown (Vandebrouck et al. 2002), TG did not induce significant modifications of the properties of these channels (unitary conductance of 7.1 ± 0.68 pS, n = 20, reversal potential of +48 mV). However, their occurrence (number of patches in which a Ca2+ current is recorded/number of patches sampled) was significantly increased by a factor of 2.8 in the presence of TG (the channel activity appearing in patches devoid of activity in the absence of TG, Fig. 1C). The open channel probability (Po) was also significantly increased after TG application (0.05 ± 0.02 (n = 7) versus 0.19 ± 0.08 (n = 20)) and the quantity of charge passing through the channels (integration of the current extrapolated over a period of 120 s of observation) approximately doubled (Fig. 1C). The current passing through these channels was efficiently inhibited by 50 µM Gd3+, 50 µM La3+ or 30 µM SKF-96365, but not by 100 µM 2-APB (Fig 1A, D and E). A dose–response relationship is presented for Gd3+, showing an IC50 value of around 30 µM (Fig. 1D). All these properties were consistent with previously described properties of voltage-independent Ca2+ channels found in skeletal muscle (Vandebrouck et al. 2002), and presented some similarities with the channels studied previously by Franco-Obregon and Lansman and reported to be mechanosensitive (Franco-Obregon & Lansman, 1994). This prompted us to investigate whether the store-operated channels detected here were also mechanosensitive.
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Figure 2 shows that applying a negative pressure to the patch electrode increased the Po of voltage-independent Ca2+ channels but had no significant effect on their conductance (7.1 ± 0.39 pS, n = 6) or on their reversal potential (+52 mV), which were similar to those described above (Fig. 2A and B). The relationship between the amount of pressure applied to the patch electrode and channel Po was well fitted by a Boltzmann equation (Fig. 2C). In contrast, the occurrence of channel activity was not significantly modified by mechanical stretch (Fig. 2D). We also compared the response to mechanical stretch in the presence and the absence of TG. Figure 2C shows that channel activity recorded from muscle fibres under the two experimental conditions increased with the pressure of suction applied to the patch electrode. The occurrence (Fig. 2D) and the total quantity of charge (not shown) also significantly increased. To further determine whether store-dependent channels and mechanosensitive channels belong to the same population, we studied the sensitivity of these channels to pharmacological agents. We found that inhibitors of store-operated channels, such as Gd3+ (50 µM, i.e. just above the EC50 value for SOC inhibition, see above) and SKF-96365 (30 µM) which inhibited, respectively, 78 and 50% of the current induced by TG (Fig. 1E) also inhibited 78 and 69%, respectively, of the current triggered by pressure application (Fig. 2F). We also tested GsMTx4, a peptide toxin from the tarantula Grammostola spatulata, reported to specifically block mechanosensitive channels (Suchyna et al. 2000). In the presence of 5 µM GsMTx4, no basal activity was detected. As expected, 5 µM GsMTx4 inhibited the response to stretch (suction from –20 to –60 mmHg), but interestingly, also completely abolished the response to TG (no activity detected in eight experiments, representing a total recording of more than 43 min in the presence of TG; Fig. 3). We concluded that adult muscle fibres have SOCs and SACs that share similar pharmacological profiles. In particular, both channels are inhibited by the GsMTx4 toxin which has been shown to inhibit TRPC1 channels.
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Experimentally, sarcoplasmic reticulum can be fully depleted by a combined and prolonged action of 20 mM caffeine and 1 µM TG in the absence of external Ca2+ (10 mM EGTA). Re-introduction of Ca2+ in the external medium induces a large entry of Ca2+ which is inhibited by Gd3+ (data not shown). But do the channels studied here function as store-operated channels in vivo? The following experiments were designed to examine the level of store-depletion necessary to activate voltage-independent Ca2+ channels. Ca2+ measurements and patch-clamp studies were performed in parallel on cells maintained in Krebs medium that were first transferred to potassium aspartate medium. Depolarization in this solution induced a [Ca2+]i transient that was taken as an index of the amount of releasable Ca2+ (Fig. 5A, procedure no.1). Fibres were then kept for 5 min in this solution in the absence or in the presence of 1 µM thapsigargin (Fig. 5A and C, procedures no.2 and no. 4); alternatively, fibres were repolarized rapidly after the first peak of Ca2+ and TG was applied for 5 min in a Krebs solution (Fig. 5B, procedure no.3). Whatever the treatment, the fibres were then rinsed in Krebs medium (to prepare for the next depolarization) containing 50 µM EGTA to avoid any refilling of the stores during this period of time. The return to potassium aspartate solution produced a second peak of [Ca2+]i the amplitude of which could be compared with the initial transient of [Ca2+]i in order to estimate the decrease of releasable Ca2+ staying in the stores. Compared with the control situation, these three different procedures significantly reduced the content of releasable Ca2+ to 69, 34 and 18% of their initial content (Fig. 5D). The level of activity of Ca2+ channels was studied under similar conditions and a relationship between the content of releasable Ca2+ versus the Po of store-operated channels is presented in Fig. 5D. It turns out that a threshold of 30% of depletion seems sufficient to activate store-operated channels.
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Muscles stimulated maximally and repeatedly present a progressive decrease in tension production, in shortening velocity and in relaxation speed. These observations, commonly grouped under the concept of muscle fatigue, were classically attributed to the accumulation of intracellular lactic acid (Hill & Kupalov, 1929) resulting in acidosis, but it is now clear that this effect contributes little to muscle fatigue (Allen, 2004) and that intracellular ionic changes also play major roles in this process (see Discussion; Stephenson et al. 1998). The following experiments were designed to assess the role of the influx of Ca2+ in muscle fatigue. Soleus muscles were chosen for their dependence on oxidative metabolism, thus limiting the role of anaerobic processes in ionic changes and their contribution to muscle fatigue. These muscles were subjected to 50 Hz stimulation trains of 500 ms duration at 1 s intervals (50% duty cycle). After 2 min, the muscles were allowed to recover and were stimulated at 10 s intervals for 10 min. Under control conditions, maximal force was maintained during the first 30 s and then progressively declined. At the same time, relaxation became incomplete during the 0.5 s separating two successive stimulation periods (Fig. 6A). Recovery occurred during the first 5 min after the protocol of fatigue. This recovery was complete and a second fatigue protocol could be performed without any modification in comparison to the first protocol. In the absence of extracellular Ca2+, the maximal force was not maintained, declined significantly faster and the relaxation stayed complete during the whole protocol (Fig. 6A). Recovery was significantly slower and less complete (data not shown). In order to investigate the possible involvement of the store-dependent and mechanosensitive channels, the same protocol was followed in the presence of extracellular Ca2+ but in the absence and then in the presence of SKF-96365, Gd3+ or GsMTx4 toxin (paired experiments). The presence of Gd3+ (50 µM) significantly accentuated the decrease of force during the fatigue protocol (Fig. 6B; P < 0.05, t test on paired experiments). The presence of SKF-96365 also accentuated the decrease of force albeit a little later (effect significant after 30 s). Finally, the GsMTx4 toxin (5 µM; tested only two times because of its cost) had a similar effect and partially mimicked the absence of extracellular Ca2+ (Fig. 6B).
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Discussion |
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Possible molecular identity of SOCs and SACs
SACs in muscle fibres might be constituted of the TRPC1 isoform, which has been shown to form such stretch-activated cation channels in vertebrate cells (Maroto et al. 2005). Accordingly, this channel is completely inhibited by the spider venom toxin GsMTx4, which, to date, has only been shown to block SACs encoded by TRPC1 (Suchyna et al. 2000; Gottlieb et al. 2006). We have previously suggested the involvement of this protein as a constituent of SOCs in normal and dystrophic skeletal muscle fibres (Vandebrouck et al. 2002). The similarity of the properties of SACs and SOCs, in particular their similar sensitivity to GsMTx4 toxin, suggests that both channels might be constituted, at least partially, of TRPC1 protein. Similarly, SACs and SOCs might also involve TRPV2 protein (previously named GRC or VRL1), a channel which translocates to the membrane upon IGF-1 stimulation and which is elevated in the membrane of dystrophic patients (Kanzaki et al. 1999; Iwata et al. 2003; Muraki et al. 2003; Yeung et al. 2005). Finally, they might implicate the TRPV4 isoform which is activated by cell swelling through a phospholipase A2 (PLA2)-dependent mechanism (Vriens et al. 2004). Interestingly, we have previously shown that the channels described here have an abnormally high open probability in dystrophic muscle fibres (Gailly, 2002; Vandebrouck et al. 2002), a situation in which the activity of PLA2 has been reported to be increased by a factor of 40 (Lindahl et al. 1995). Precise identification of the isoform(s) involved is under study.
Functional role of SACs and SOCS in skeletal muscle fibres
It is not obvious that store depletion occurs in vivo because adult muscle fibres do not exchange much Ca2+ with the extracellular medium. Indeed, skeletal muscle fibres have huge amounts of sarcoplasmic reticulum which is extremely rich in Ca2+-pumps (SERCA) and which contains a high buffering capacity (calsequestrin). Ca2+ extrusion through the plasma membrane seems also very slow (reviewed in Martonosi & Pikula, 2003). So almost all of the Ca2+ released from the sarcoplasmic reticulum is rapidly restored to the sarcoplasmic reticulum after stimulation; accordingly, twitch contractions can thus be produced repeatedly in the absence of extracellular Ca2+ (Armstrong et al. 1972). However, evaluations of Ca2+ influx using the Ca45 uptake technique indicate that each twitch contraction induces a small increase of Ca2+ entry (Bianchi & Shanes, 1959), the mechanism of which is unknown. Here, we show that a decrease of the order of 30% of the Ca2+ stores is sufficient to induce an entry of Ca2+. Interestingly, it has been evaluated that a single action potential triggers the release of 0.2–0.3 mM from the sarcoplasmic reticulum to the cytoplasm (Baylor et al. 1983), which corresponds to more than a quarter of the Ca2+ contents present in the sarcoplasmic reticulum (Endo, 1977). Thus it seems reasonable to think that store-operated channels are indeed activated during in vivo contraction. This is corroborated by the fatigue experiments. In the absence of external Ca2+ or when SOCs are inhibited, a faster decline of force is observed, suggesting that a sustained activity, such as the one observed in tonic muscles as soleus, requires a constant repletion of the stores by a subsequent entry of Ca2+. A partial failure of the sarcoplasmic reticulum to release Ca2+ during tetanus has been proposed as a possible cause of fatigue (Allen & Westerblad, 2001; Allen, 2004). This seems to be due to the accumulation of Ca2+ phosphate in the sarcoplasmic reticulum (Kabbara & Allen, 1999; Dutka et al. 2005). It is interesting to note that, at rest, the sarcoplasmic reticulum of slow twitch fibres (as in soleus muscle) is saturated with Ca2+ while the sarcoplasmic reticulum of fast twitch muscle fibres is only about one-third full (Fryer & Stephenson, 1996), suggesting that the amount of Ca2+ in the stores might be more critical for the physiological function of slow twitch fibres. Our results emphasize the importance of the entry of Ca2+ during sustained trains of contractions of slow twitch muscle. Such importance of Ca2+ handling in muscle fatigue has also been suggested in fast and slow muscles deficient in mitsugumin, a protein expressed at the triad junction, the lack of which leads to a disorganization of the T-tubules and sarcoplasmic relationship and to a susceptibility to fatigue and to a dysfunction of store-operated entry of Ca2+ (observed in embryonic and neonatal muscles but not in adult fast fibres) (Nishi et al. 1999; Nagaraj et al. 2000; Kurebayashi & Ogawa, 2001; Kurebayashi et al. 2003; Ma & Pan, 2003).
Whether the channels studied here are also activated by stretch during contraction is difficult to evaluate. Indeed, the force applied to membrane patches in the experiments presented here represents only about 5% of the force developed by muscle during contraction. However, there are no data on the possible transmission of force (produced during contraction) to the cell membrane.
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Footnotes |
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) and TG-treated cells (
). Voltages indicated were determined from the applied holding potential and the estimated resting potential (–50 mV). C, the proportion of patches with a channel activity (occurrence), the open state probability (n.Po), and the quantity of charge obtained by integration of the current observed extrapolated over a period of 120 s under control condition and after application of 1 µM TG (same patch studied). *Significant difference (P < 0.05) between control and TG-treated cells; the number of patches under each condition is indicated above the histograms. 
Significantly different (P < 0.05) from –20 mmHg pressure (Fisher exact test). The proportion of patches in which an activity is detected is indicated above each column of the histogram. E, pharmacological characterization of stretch-activated single-channel currents. Examples of current traces (patch-clamp; cell-attached configuration at –60 mV holding potential) recorded under control condition (during a –20 mmHg suction) or in the presence of Gd3+ (50 µM) and SKF-96365 (25 µM). Mean unitary currents of 0.87 pA. F, response (in percentage) to –20 mmHg stretch in the absence and presence of 50 µM Gd3+ or 30 µM SKF-96365. *Significantly different (P < 0.05) from control conditions (stretch without inhibitors; one-way ANOVA followed by a Tukey test).
