Nitric oxide synthase inhibition affects sarcoplasmic reticulum Ca2+ release in skeletal muscle fibres from mouse
1 Physiologie Intégrative Cellulaire et Moléculaire, Université Claude Bernard – Lyon 1, UMR CNRS 5123, Villeurbanne, France
Abstract
Nitric oxide (NO) generated by skeletal muscle is believed to regulate force production but how this is achieved remains poorly understood. In the present work we tested the effects of NO synthase (NOs) inhibitors on membrane current and intracellular calcium in isolated skeletal muscle fibres from mouse, under voltage-clamp conditions. Resting [Ca2+] and [Ca2+] transients evoked by large depolarizations exhibited similar properties in control fibres and in fibres loaded with tenth millimolar levels of the NOs inhibitor N-nitro-L-arginine (L-NNA). Yet the voltage dependence of calcium release was found to be shifted by 15 mV towards negative values in the presence of L-NNA. This effect could be reproduced by the other NOs inhibitor S-methyl-L-thiocitrulline (L-SMT). Separate experiments showed that the voltage dependence of charge movement and of the slow calcium current were unaffected by the presence of L-NNA, ruling out an effect on the voltage sensor. A negative shift in the voltage dependence of calcium release with no concurrent alteration in the properties of charge movement was also observed in fibres exposed to the oxidant H2O2 (1 mM). Conversely the reducing agent dithiothreitol (10 mM) had no obvious effect on Ca2+ release. Overall, the results indicate that physiological levels of NO exert a tonic inhibitory control on the activation of the calcium release channels. Changes in the voltage dependence of Ca2+ release activation may be a ubiquitous physiological consequence of redox-related modifications of the ryanodine receptor.
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Introduction
Nitric oxide is a universal signalling molecule with a key role in an extremely diverse variety of physiological and patho-physiological processes. Its relevance has been especially well established in smooth and cardiac muscle (see for recent reviews Ignarro, 2002 and Massion et al. 2003) but NO is also suspected to play an important role in skeletal muscle. Mature skeletal muscle fibres express the three major isoforms of NO synthase with a specific splice variant of the neuronal isoform being predominant (Silvagno et al. 1996). Contraction of isolated muscle preparations can be affected by exogenous NO and by inhibitors of NO synthesis, and both impairment of endogenous NO supply and overproduction of NO are thought to be involved in some skeletal muscle disorders (for reviews see Eu et al. 1999; Stamler & Meissner, 2001). In terms of signalling pathway, the situation is, however, far less clear than in the cardiovascular system where clear-cut NO effects occur through changes in the intracellular cGMP level, a second messenger that is of poorly defined relevance to skeletal muscle function. Rather, a massive body of evidence supports the view that in skeletal muscle, NO could largely act through mechanisms of oxidation and/or S-nitrosylation of free thiols, with many reports pointing to the sarcoplasmic reticulum (SR) calcium release channel as a major potential target protein (Meszaros et al. 1996; Aghdasi et al. 1997; Stoyanovsky et al. 1997; Suko et al. 1999; Eu et al. 2000, 2003; Hart & Dulhunty, 2000; Sun et al. 2001a, 2003). Under normal conditions, SR Ca2+ release by the ryanodine receptors is driven by a voltage-induced conformational change in the dihydropyridine receptors present in the t-tubule membrane. Whether or not NO production can modulate this process in situ and if it does, how this is achieved remains misunderstood. In this context the effects of NO synthesis inhibition are of strong interest: for instance various NO synthase inhibitors were shown to potentially enhance force production in isolated muscle preparations (Kobzik et al. 1994; Gath et al. 1996; Reid et al. 1998). Whether these effects on force production are related to a modulation of the SR Ca2+ release flux has not been established. In a recent study we showed that excess NO applied to an isolated skeletal muscle fibre produces a use-dependent increase in resting [Ca2+] consistent with some calcium release channels remaining locked open at rest (Pouvreau et al. 2004). In the present work we attempted to reveal the role of the endogenously produced NO levels on the control of intracellular [Ca2+]. Results show that inhibitors of NO synthase affect the voltage dependence of Ca2+ release and that a similar effect is produced by the oxidant H2O2.
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Methods
Preparation of the muscle fibres
Experiments were performed on single skeletal fibres isolated from the flexor digitorum brevis (FDB) muscles from 4- to 8-week-old OF1 mice. All experiments were performed in accordance with the guidelines of the French Ministry of Agriculture (87/848) and of the European Community (86/609/EEC). Procedures for enzymatic isolation of single fibres, partial insulation of the fibres with silicone grease and intracellular dye loading, were as previously described (Jacquemond, 1997; Collet et al. 1999, 2004; Collet & Jacquemond, 2002). In brief, mice were killed by cervical dislocation before removal of the muscles. Muscles were treated with collagenase (ICN type 1) for 60–75 min at 37°C in the presence of external Tyrode solution. Single fibres were then obtained by triturating the muscles within the experimental chamber. The major part of a single fibre was electrically insulated with silicone grease so that whole-cell voltage clamp could be achieved on a short portion of the fibre extremity. Prior to voltage clamp, indo-1 was introduced locally into the fibre by pressure microinjection through a micropipette containing 0.5 mM indo-1 dissolved in an intracellular-like solution (see ‘Solutions’). Fibres were then left for 1 h to allow for intracellular equilibration of the dye. The NOs inhibitors N-nitro-L-arginine (L-NNA) and S-methyl-L-thiocitrulline (L-SMT) were in some cases present in the microinjected solution (see ‘Solutions’). Following diffusion and equilibration within the cytoplasm, it was believed that final cytoplasmic concentrations of these compounds within one tenth to one fifth of the initial pipette concentration were achieved (for details concerning microinjections see Csernoch et al. 1998). For each series of measurements under a given condition, control fibres used for the analysis were always issued from the same muscles as the test fibres. In some experiments, the two collagenase-treated FDB muscles from the same mouse were kept overnight at 4°C. One muscle was kept in the Tyrode solution for control measurements. The other muscle was in the Tyrode solution which also contained 5 mM each of L-NNA and L-SMT; L-NNA and L-SMT were present in the solution bathing the fibres during all subsequent steps of the experimental procedure. All experiments were performed at room temperature (20–22°C).
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Electrophysiology
An RK-400 patch-clamp amplifier (Bio-Logic, Claix, France) was used in whole-cell configuration. Command voltage pulse generation and data acquisition were done using commercial software (Biopatch Acquire, Bio-Logic) driving an A/D, D/A converter (LabMaster DMA board, Scientific Solutions Inc., Mentor, OH, USA). Analog compensation was systematically used to decrease the effective series resistance. Voltage clamp was performed with a microelectrode filled with the intracellular-like solution. The tip of the microelectrode was inserted through the silicone, within the insulated part of the fibre. Unless otherwise specified, membrane depolarizations (or series of depolarizations) were applied every 30 s from a holding command potential of –80 mV.
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Measurement of the slow calcium current
Calcium current records were obtained in response to 1 s-long depolarizing steps to various potentials applied every 30 s. The linear leak component of the current was removed by subtracting the adequately scaled value of the steady current measured during a 20 mV hyperpolarizing step applied before each test pulse. The voltage dependence of the peak calcium current density was fitted with the following equation:
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where I(V) is the peak current density at the command potential V, Gmax is the maximum conductance, Vrev is the apparent reversal potential, V0.5 is the half-activation potential and k is a steepness factor.
Measurement and analysis of intramembrane charge movement
Charge movement currents were measured and analysed according to previously described procedures (Collet et al. 2003; Pouvreau et al. 2004). For these measurements the holding potential was set to –100 mV. In brief, adequately scaled control current records elicited by 50 ms-long hyperpolarizing pulses of 20 mV were subtracted from the current elicited by test depolarizing pulses of the same duration to various levels. The amount of charge moved during a test pulse was measured by integrating the ‘on’ portion of the corrected test current records. The calculated charge was normalized to the capacitance of the fibre. The steady-state distribution of the normalized charge was fitted with a two-state Boltzmann function:
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where Qmax corresponds to the maximal available charge, V0.5 to the voltage of equal charge distribution and k to the steepness factor.
Fluorescence measurements
The optical set-up and the procedures used for indo-1 fluorescence measurements were as previously described (Jacquemond, 1997; Collet et al. 1999; Collet & Jacquemond, 2002; Pouvreau et al. 2004). In brief, a Nikon Diaphot epifluorescence microscope was used in diafluorescence mode. The beam of light from a high-pressure mercury bulb set on the top of the microscope was passed through a 335 nm interference filter and focused onto the preparation using a quartz aspherical doublet. The emitted indo-1 fluorescence light was collected by a x 40 objective and simultaneously detected at 405 ± 5 nm (F405) and 470 ± 5 nm (F470) by two photomultipliers. The fluorescence measurement field was 40 μm in diameter and the silicone-free extremity of each tested fibre was placed in the middle of the field. Background fluorescence at both emission wavelengths was measured next to each tested fibre and was then subtracted from all measurements.
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Calibration of the indo-1 response and [Ca2+]i calculation
The standard ratio method was used with the parameters: R = F405/F470, with Rmin, Rmax, KD and having their usual definitions. Results were either expressed in terms of indo-1% saturation or in actual free calcium concentration (for details of calculation, see Jacquemond, 1997; Csernoch et al. 1998). In vivo values for Rmin, Rmax and were measured using procedures previously described (Collet et al. 1999; Collet & Jacquemond, 2002). Unless otherwise specified, no correction was made for indo-1–Ca2+ binding and dissociation kinetics. In vitro measurements of the indo-1 response performed on the experimental set-up (Jacquemond, 1997) showed that the dye response to calcium was unaffected by L-NNA (not illustrated).
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Calculation of the rate of calcium release
The method used to estimate the SR calcium release flux from the indo-1 transients involved taking the time derivative of the total myoplasmic Ca2+ calculated from the occupancy of the main intracellular calcium binding sites (Baylor et al. 1983). The model included troponin C binding sites with a total site concentration TNtotal = 250 μM, an ‘on’ rate constant kon,CaTN = 0.0575 μM–1 ms–1 and an ‘off’ rate constant koff,CaTN = 0.115 ms–1; Ca2+–Mg2+ binding sites on parvalbumin with a total site concentration PVtotal = 700 μM, ‘on’ rate constant for Ca2+ kon,CaPV = 0.125 μM–1 ms–1, ‘off’ rate constant for Ca2+ koff,CaPV = 5 x 10–4 ms–1, ‘on’ rate constant for Mg2+ kon,MgPV = 3.3 x 10–5 μM–1 ms–1, ‘off’ rate constant for Mg2+ koff,MgPV = 3 x 10–3 ms–1. Resting [Mg2+] was assumed to be 1.5 mM. For these calculations free calcium transients were corrected for the Ca2+–indo-1 binding kinetics using values for the ‘on’ and ‘off’ rate constants of the Ca2+–indo-1 binding reaction of 1 x 108 M–1 s–1 and 30 s–1, respectively, by analogy with the values derived for fura-2 by Baylor & Hollingworth (1988).
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Solutions
The intracellular-like solution contained (mM): 120 potassium glutamate, 5 Na2-ATP, 5 Na2-phosphocreatine, 5.5 MgCl2, 5 glucose, 5 Hepes adjusted to pH 7.20 with KOH. In some experiments the NOs inhibitors N-nitro-L-arginine (L-NNA) and S-methyl-L-thiocitrulline (L-SMT) were used. When microinjected, the respective inhibitors were present in the micropipette solution at the following concentrations: L-NNA, 5 mM; L-SMT, 5 or 20 mM. For the incubated fibres L-NNA and L-SMT were present in the extracellular solution at a concentration of 5 mM. The standard extracellular solution contained (mM): 140 TEA-methanesulphonate, 2.5 CaCl2, 2 MgCl2, 10 TEA-Hepes and 0.002 tetrodotoxin, pH 7.20. For measurements of the slow calcium current the extracellular solution also contained 4-aminopyridine (1 mM) and the fibres were pressure-microinjected with a solution containing 50 mM EGTA diluted in the intracellular-like solution. For measurements of charge movement the extracellular solution contained 140 TEA-methanesulphonate, 0.5 CaCl2, 4 MgCl2, 1 4-aminopyridine, 0.5 or 1 CdCl2, 0.3 LaCl3, 10 TEA-Hepes and 0.002 tetrodotoxin, pH 7.20. In addition fibres were pressure-microinjected with a solution containing 10 mM EGTA diluted in the intracellular-like solution.
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Statistics
Least-squares fits were performed using a Marquardt-Levenberg algorithm routine included in Microcal Origin (OriginLab, Northampton, MA, USA). Data values are presented as means ± S.E.M. for n fibres, where n is specified in Results. Statistical significance was determined using Student's t test assuming significance for P < 0.05.
Results
Calcium transients in response to depolarizing pulses of increasing duration in the presence of the NOs inhibitor L-NNA
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Figure 1A shows mean (continuous line) ± S.E.M. (grey shading) indo-1 saturation traces elicited from control fibres (n = 20) and from fibres injected with L-NNA (n = 11) in response to successive depolarizing pulses of 20, 70 and 120 ms duration to +10 mV. In each panel the inset shows an example of traces from an individual fibre in the given condition. Obviously the presence of L-NNA did not dramatically affect the qualitative features of the transients, that is a rise triggered by the depolarization with an initial very rapid phase, and a decline to the resting level that immediately follows membrane repolarization. The decaying phase of the transients appeared though to be somewhat faster in the presence of L-NNA than in control conditions. Figure 1B presents mean values from that series of measurements for resting [Ca2+], peak [Ca2+] and time constant of [Ca2+] decay. Measurements were performed on individual [Ca2+] traces calculated from each fibre. The time constant of decay was obtained from fitting a single exponential plus constant function to the [Ca2+] decline as previously described (Collet et al. 1999; Pouvreau et al. 2004). The resting [Ca2+] and peak [Ca2+] values did not significantly differ between the two groups. The mean value for the time constant of [Ca2+] decay was 25–30% lower in the presence of L-NNA, the difference being significant only for the shortest pulse duration (P = 0.02). Overall, these results indicate that the presence of a potent inhibitor of nNOS activity within the intracellular medium did not impair the ability of the fibres to regulate [Ca2+] at rest and to produce robust calcium transients in response to membrane depolarization. The reason for the slightly faster decay of the transients in response to short pulses in the presence of L-NNA is unclear. It may be the consequence of a chronic effect of NO on a component of the cytoplasmic Ca2+ removal systems. This effect was not further investigated within the present work.
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A, mean (continuous lines) ± S.E.M. (grey shading) indo-1 saturation signals obtained in response to depolarizations of 20, 70 and 120 ms duration to +10 mV in control fibres and in fibres injected with L-NNA. In each panel the inset shows an example of traces measured from a single fibre under the corresponding condition. B, mean ± S.E.M. values of resting [Ca2+], peak change in [Ca2+] and time constant of [Ca2+] decay measured from the individual control fibres and L-NNA-injected fibres under the conditions shown in A.
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Alteration of the voltage dependence of Ca2+ release activation in the presence of NOs inhibitors
Figure 2 illustrates what we believe is the prevailing effect of NOs inhibition on the control of intracellular calcium. Indo-1 calcium transients were examined in control fibres and in fibres treated with the NOs inhibitors L-NNA or (and) L-SMT using the pulse protocol shown above the corresponding series of records in Fig. 2. The protocols consisted of three consecutive depolarizations of 50 ms duration to the indicated values; each pulse was then incremented by 5 mV and the sequence repeated. Within five runs this allowed an explicit examination of the voltage dependence of the properties of the transients. In addition, two levels of depolarizations (–30 and –10 mV) were tested twice, allowing the stability of the preparation to be checked during the course of the protocol. The interval between two pulses within a run was either 0.5 s (as in the top series of records) or 1 s; this did not affect the mean values for the parameters that were measured from the data under a given condition (not illustrated). Qualitatively, traces in Fig. 2A and B show that, as compared to the control conditions (left series of records), the indo-1 transients elicited in the fibres injected with L-NNA or L-SMT tended to emerge for lower levels of membrane depolarization and the maximal amplitude was also reached for less depolarized levels. Figure 2C illustrates the fact that this was also the case in fibres that were kept for several hours in the presence of L-NNA and L-SMT before taking measurements (see Methods). In order to quantitatively estimate this effect, free calcium transients and corresponding release fluxes were calculated from the indo-1 transients as described in Methods. An example of the [Ca2+] trace and of the corresponding release flux is shown in Fig. 3A and C, respectively. Figure 3B shows the voltage dependence of the mean normalized peak [Ca2+] transients from control fibres (filled symbols, n = 19), from fibres injected with L-NNA (, n = 8), from fibres injected with L-SMT (, n = 7) and from fibres incubated in the presence of both inhibitors (, n = 4). In each fibre all values for peak [Ca2+] were normalized to the maximal peak value that was reached during the protocol. For the fibres incubated with L-NNA and L-SMT, control data that are shown (filled squares, n = 3) were from fibres from the same batch, incubated during the same amount of time in control solution (see Methods). Figure 3D shows the mean voltage dependence of the peak release flux calculated from the same data. A negative shift in the voltage dependence of release activation was manifest under the three tested conditions of NOs inhibition. The individual release flux versus voltage relationship from each individual fibre was fitted with a Boltzmann function. Mean values for the maximal release flux and for the steepness factor were 4.6 ± 0.6 μM ms–1 and 6.0 ± 0.3 mV in control fibres (n = 19), 4.9 ± 0.6 μM ms–1 and 6.6 ± 0.2 mV in L-NNA-injected fibres (n = 8) and 3.4 ± 0.6 μM ms–1 and 7.0 ± 0.5 mV in L-SMT-injected fibres (n = 7), respectively. Both parameters were not significantly different between control and treated fibres. The same was true for the batch of incubated fibres: mean values for the maximal release flux and steepness factor were 3.8 ± 0.9 μM ms–1 and 6.7 ± 0.6 mV in control fibres (n = 3) and 3.2 ± 0.5 μM ms–1 and 6.2 ± 0.2 mV in fibres incubated with L-NNA and L-SMT (n = 4), respectively. Conversely mean values for the midpoint voltage were significantly more negative in the NOs inhibitor-treated fibres than in the control fibres: they were –17.0 ± 2.0, –29.2 ± 2.3 and –27.4 ± 2.1 mV in control fibres (n = 19), fibres injected with L-NNA (n = 8, P = 0.002) and in fibres injected with L-SMT (n = 7, P = 0.008), respectively. Accordingly the mean midpoint voltage from fibres incubated overnight in the presence of both inhibitors was –31.8 ± 2.4 mV (n = 4) as compared to –17.6 ± 1 mV (n = 3, P = 0.005) for the concurrent batch of control fibres. It should be mentioned that the estimation of the half-activation potential in Fig. 3B was certainly more accurate in the NOs inhibitor-treated fibres (for which saturation was reached early within the explored voltage range) than in the control ones. The actual value of the half-activation voltage in the control fibres may be slightly more positive than estimated from our fits and this would tend to a somewhat underestimation of the difference with the fibres treated with the NOs inhibitors.
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A, B and C, indo-1 saturation signals elicited in three distinct control fibres. D, E and F, indo-1 saturation signals elicited in a L-NNA-injected fibre, in a L-SMT-injected fibre and in a fibre incubated in the presence of L-NNA and L-SMT prior to the experiment, respectively (see text and Methods for details). The pulse protocol that was used is shown above each series (A and D) or pair of series (B and C, E and F) of records; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV.
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A, example of [Ca2+] trace calculated from an indo-1 signal elicited by consecutive depolarizing pulses to –30, –10 and +10 mV (protocol shown in Fig. 2). C, corresponding Ca2+ release flux trace calculated as described in Methods. B, voltage dependence of the mean relative peak change in [Ca2+] in control fibres (, n = 19), in L-NNA-injected fibres (, n = 8), in L-SMT-injected fibres (, n = 7) and in fibres incubated in the presence of L-NNA and L-SMT (, n = 4). Control data for the fibres incubated with L-NNA and L-SMT were from fibres incubated for the same amount of time in control solution (, n = 3). In each fibre the value for peak [Ca2+] elicited by a given pulse was normalized to the maximum measured value. D, corresponding voltage dependence of the mean peak Ca2+ release flux. The superimposed continuous lines correspond to the results from fitting a Boltzmann function to the mean data points: corresponding values for maximum release flux, midpoint voltage and steepness factor in the two batches of control fibres were 4.8 μM ms–1, –15 mV and 8.6 mV () and 3.8 μM ms–1, –17.4 mV and 6.8 mV (), respectively. Values from the fits to data points in L-NNA-injected fibres, in L-SMT-injected fibres and in fibres incubated in the presence of L-NNA and L-SMT were 4.9 μM ms–1, –28.4 mV and 7.5 mV, 3.41 μM ms–1, –25.6 mV and 7.8 mV, and 3.2 μM ms–1, –31.1 mV and 6.3 mV, respectively.
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Function of the dihydropyridine receptor in the presence of a NOs inhibitor
The effects of NOs inhibitors on Ca2+ release activation could be due to a modification of the voltage-sensing steps of excitation–contraction (E–C) coupling. In order to check for this, we looked at the calcium channel and voltage-sensing functions of the dihydropyrdine receptor. We first examined the membrane current from the experiments described in relation to Fig. 3. An inward current was clearly flowing during the most depolarized values of step potential, indicative of the activation of the slow inward calcium current (not shown). Analysis of the voltage dependence of the end-pulse current in the control fibres, in the fibres injected with L-NNA, and in the fibres injected with L-SMT revealed that at –20 mV the current was significantly larger in the group of L-NNA-injected fibres (–0.96 ± 0.3, n = 8) than in the control fibres (–0.34 ± 0.14, n = 17). However, for other potential values there was no significant difference between control fibres and either L-NNA-injected fibres or L-SMT-injected fibres.
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In order to more carefully assess a possible effect of NOs inhibition on the channel function of the dihydropyridine receptor, we performed a specific series of calcium current measurements in a batch of control and L-NNA-injected fibres using 1 s-long depolarizations (see Methods). Figure 4A shows calcium current traces measured from a control fibre (left) and from an L-NNA-injected fibre (right) obtained in response to membrane depolarizations ranging between –50 and +70 mV with a 5 mV increment. There was no striking difference between the two. Figure 4B shows the voltage dependence of the mean peak calcium current density from control fibres (, n = 7) and L-NNA-injected fibres (, n = 9). Fitting the individual I–V curves as described in Methods gave mean values for Gmax, Vrev, V0.5 and k of 142 ± 12 S F–1, 62.2 ± 3 mV, –1.2 ± 2 mV and 6.7 ± 0.6 mV in control fibres, and 127 ± 7 S F–1, 60.6 ± 3.5 mV, 1.6 ± 2 mV and 6.8 ± 0.5 mV in L-NNA-injected fibres, respectively. None of these parameters significantly differ between the two groups.
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A, calcium current records obtained in response to 1 s-long depolarizing pulses to values between –50 and +70 mV in a control fibre (left) and in a fibre pressure-microinjected with L-NNA (right). B, mean voltage dependence of the peak calcium current amplitude in control fibres (, n = 7) and in L-NNA-injected fibres (, n = 9). The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details).
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In order to further explore the possibility that the voltage sensor may be affected by the treatment with NOs inhibitors we performed a specific series of experiments designed to measure intramembrane charge movement in control and in L-NNA-injected fibres. Figure 5A shows representative traces of intramembrane charge current from a control fibre (left) and from an L-NNA-injected fibre (right). There was no obvious difference between the two series. Figure 5B shows the mean voltage distribution of the ‘on’ charge in the two groups of fibres. Fitting a two-state Boltzmann distribution to the amount of ‘on’ charge versus voltage data in each fibre gave mean values for Qmax, V0.5 and k of 24 ± 3 nC μF–1, –30.7 ± 2 mV and 11 ± 1 mV in control fibres (n = 6), and 24 ± 4 nC μF–1, –30.8 ± 2 mV and 11.3 ± 2 mV in L-NNA-injected fibres (n = 7), respectively. Obviously, none of the parameters was significantly different between the two conditions.
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A, charge movement records from a control fibre and from an L-NNA-injected fibre, measured in response to 50 ms-long depolarizing steps to the indicated values of command potential. B, mean voltage distribution of the ‘on’ charge in control fibres (n = 6) and in L-NNA-injected fibres (n = 7). Superimposed continuous lines correspond to the result of fitting a two-state Boltzmann distribution.
Effect of H2O2 on voltage-activated calcium release
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Many previously reported effects of the NO signalling pathway on skeletal muscle contractility, intracellular calcium regulation and on the activity of the isolated ryanodine receptor were either shown or at least suspected to be intimately linked to redox-based modulation of protein function (for review see Reid, 2001; Stamler & Meissner, 2001). For instance it was shown that low concentrations of NO could prevent activation of the calcium release channel by oxidation (Aghdasi et al. 1997). Within the present context it was therefore of particular interest to study how calcium release is affected by the presence of an oxidant under the same conditions as above. This was done by exposing isolated fibres to H2O2. Figure 6A shows indo-1 saturation traces obtained in response to the same protocol as in Fig. 2 in a control fibre and in a fibre bathed in the presence of H2O2 (1 mM). As observed in the presence of NOs inhibitors, transients were detected for lower voltage levels in the fibre exposed to H2O2 than in the control fibre and the maximal peak amplitude of the transient also tended to be reached at lower levels of membrane depolarization. Figure 6B shows the voltage dependence of the mean normalized peak change in [Ca2+] (left) and of the corresponding mean peak release flux (right) in both groups of fibres. There was a definite leftward shift in the voltage dependence of Ca2+ release in the presence of H2O2, but this was also associated with a reduction of the maximal peak release flux. The individual peak release flux versus voltage relationship from each fibre was fitted with a Boltzmann function. Mean values for the maximal release flux, midpoint voltage and steepness factor were 7.7 ± 0.4 μM ms–1, –18.8 ± 1.6 mV and 6.0 ± 0.4 mV in control fibres (n = 10), and 4.2 ± 0.5 μM ms–1, –26.9 ± 3 mV and 7.4 ± 0.7 mV in fibres exposed to H2O2 (n = 7), respectively. Values for maximal flux and for midpoint voltage were significantly different.
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A, indo-1 saturation signals elicited by a series of 50 ms-long depolarizing pulses of increasing amplitude in a control fibre and in a fibre bathed in the presence of 1 mM H2O2. The pulse protocol that was used is shown on the top of the panel; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV. B, voltage dependence of the mean relative peak change in [Ca2+] and of the corresponding mean peak Ca2+ release flux in control fibres (, n = 10) and in fibres exposed to H2O2 (, n = 7). Four fibres were bathed in the presence of the oxidant for 15–30 min before starting to take the measurements and three fibres were bathed in H2O2 for a longer period of time (1 h or more). The mean values for the Boltzmann parameters obtained from fitting the voltage dependence of the peak release flux in each fibre did not differ between these two sets of H2O2-treated fibres (not illustrated). Continuous lines superimposed on the release flux data correspond to the result of fitting a Boltzmann function to the mean data points: this gave values for maximum release flux, midpoint voltage and steepness factor of 7.8 μM ms–1, –19.4 mV and 6.9 mV in control fibres, and of 4.0 μM ms–1, –28.6 mV and 7.9 mV in fibres exposed to H2O2. It should be noted that this series of experiments was performed on fibres issued from a different batch of mice, and at a different period of time as compared to other results presented in this paper; this may be the reason for the difference in absolute values of peak release flux between control fibres from this series as compared to control data presented in relation to Fig. 3.
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Function of the dihydropyridine receptor in the presence of H2O2
In order to test whether the effects of H2O2 on Ca2+ release could be due to an alteration of the voltage-sensing functions of E–C coupling we measured, in separate sets of experiments, the slow calcium current and the intramembrane charge movements in the presence of the oxidant. Results are illustrated in Fig. 7. Figure 7A shows a series of calcium current records elicited by 1 s-long membrane depolarizations of increasing amplitude in a fibre exposed to 1 mM H2O2, whereas Fig. 7B shows the mean peak current versus voltage (I–V) relationships obtained from this series of measurements in four control fibres () and seven fibres exposed to H2O2 (). The peak current was somewhat reduced in the presence of H2O2. Fitting the individual I–V curves as described in Methods gave mean values for Gmax, Vrev, V0.5 and k of 176 ± 27 S F–1, 56.4 ± 2.9 mV, –0.02 ± 1.6 mV and 7.8 ± 0.9 mV in control fibres (n = 4), and 113 ± 11 S F–1, 56.6 ± 3.7 mV, –6.4 ± 3.6 mV and 8.2 ± 1.1 mV in H2O2-treated fibres (n = 7), respectively. The mean maximum conductance was reduced by 35% in the presence of H2O2 (P = 0.03). Other parameters did not differ significantly.
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A, calcium current records obtained in response to 1 s-long depolarizing pulses to values between –50 and +70 mV in a fibre exposed to 1 mM H2O2. B, mean voltage dependence of the peak calcium current amplitude in control fibres (, n = 4) and in H2O2-exposed fibres (, n = 7). The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details). C, charge movement records from a fibre exposed to 1 mM H2O2 measured in response to 50 ms-long depolarizing steps to values ranging between –80 and +15 mV. D, mean voltage distribution of the ‘on’ charge in control fibres (n = 6) and in fibres exposed to 1 mM H2O2 (n = 7). For these two sets of measurements H2O2-treated fibres were exposed to the oxidant for 15 min before starting the experiment.
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Figure 7C shows an example of charge movement traces obtained from a fibre bathed in the presence of 1 mM H2O2. Traces were obtained in response to 50 ms-long depolarizing pulses to voltages ranging between –80 mV (bottom) and +15 mV (top) with a 5 mV increment from a holding potential of –100 mV. There was no sign of gross alteration of the properties of the ‘on’ and ‘off’ charge transients. Figure 7D shows the mean voltage dependence of the ‘on’ charge in the H2O2-treated fibres (, n = 7) and in control fibres isolated from the same mice (, n = 6). There was no difference between the two populations. Fitting the individual series of data points with a Boltzmann function gave mean values for Qmax, V0.5 and k of 26.3 ± 3.2 nC –F–1, –29.5 ± 3.7 mV and 14.1 ± 2.8 mV in control fibres, and 27.0 ± 1.9 nC μF–1, –31.1 ± 2.1 mV and 13.1 ± 1.7 mV in H2O2-treated fibres, respectively.
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The reason for the H2O2-induced reduction of the calcium current conductance is unclear. It may be due to the existence of redox-sensitive uncharged transitions leading to the channel opening. In any case this effect cannot be taken as responsible for the reduced calcium release observed in the presence of H2O2 as it is well established that the fraction of calcium entering a muscle fibre through this pathway is not significant compared to the amount released from the SR. Overall this set of data indicates that the effects of H2O2 on calcium release are not related to the voltage-sensing steps of E–C coupling and are thus very likely to be due to specific alterations of the ryanodine receptor.
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Voltage-activated calcium release in the presence of dithiothreitol
We also tested the possibility that the ryanodine receptor would be in an oxidized state under our experimental conditions of ambient PO2 (see Eu et al. 2003) by measuring calcium release in the presence of the reducing agent dithiothreitol (DTT). For this calcium transients were measured first in response to 50 ms-long pulses of increasing amplitude (same 3-step protocol as in Figs 2 and 6) under control conditions; the extracellular solution was then replaced by a 10 mM DTT-containing one and the protocol was re-applied. Figure 8A illustrates the results from such an experiment. Traces shown underneath the pulse protocol correspond to indo-1 saturation signals measured in the control solution while bottom traces were obtained from the same fibre, 10 min after exchanging the extracellular solution for the DTT-containing solution. There was no obvious difference between the two series of traces. Figure 8B shows the voltage dependence of the mean normalized peak change in [Ca2+] (left) and of the corresponding mean peak release flux (right) from six fibres under control conditions () and after 10 min exposure to 10 mM DTT (). The mean values for maximal release flux, midpoint voltage and steepness factor obtained from fitting a Boltzmann function to the voltage dependence of Ca2+ release under control conditions were 9.6 ± 1 μM ms–1, –15.2 ± 2.2 mV and 6.1 ± 0.6 mV. Corresponding values in the presence of DTT were 8.5 ± 0.7 μM ms–1, –19.9 ± 3.2 mV and 6.0 ± 0.3 mV, respectively. None of the parameters significantly differ between the two conditions.
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A, indo-1 saturation signals elicited by a series of 50 ms-long depolarizing pulses of increasing amplitude in the same fibre under control conditions (upper records) and in the presence of 10 mM DTT (lower records). The pulse protocol that was used is shown at the top of the panel; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV. B, voltage dependence of the mean relative peak change in [Ca2+] and of the corresponding mean peak Ca2+ release flux from the same fibres (n = 6) under control conditions () and after 10 min exposure to 10 mM DTT (). Continuous lines superimposed on the release flux data correspond to the result of fitting a Boltzmann function to the mean data points: this gave values for maximum release flux, midpoint voltage and steepness factor of 9.5 μM ms–1, –15 mV and 6.5 mV under control conditions, and of 8.6 μM ms–1, –19.5 mV and 7.5 mV in the presence of DTT.
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Discussion
The present work provides further insights into the role of NO in the physiological regulation of SR Ca2+ release. Mainly we demonstrate here that conditions designed to block endogenous production of NO in an intact muscle fibre shift the voltage dependence of SR Ca2+ release activation towards more negative values. The most straightforward inference is that physiological intracellular levels of NO tend to reduce the sensitivity of calcium release activation to voltage, and thus exert a chronic negative effect on the ‘efficiency’ of the E–C coupling process. The effect was observed with two potent inhibitors of constitutive isoforms of nitric oxide synthase, L-NNA and L-SMT, used at sufficiently large concentrations to presumably ensure a strong inhibition of the enzyme activity (see Boer et al. 2000; Alderton et al. 2001).
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The first thinkable candidate responsible for a shift in voltage dependence of calcium release would be the voltage-sensing dihydropyridine receptor in the t-tubule membrane. However, the properties of intramembrane charge movement (including the voltage dependence) were unaffected by L-NNA which suggests that it was the calcium release channel that was involved in the present effects of NOs inhibition. This is in agreement with the results from Meszaros et al. (1996) showing that nitro-L-arginine methyl ester blocked a NO (generated from L-arginine)-induced inhibition of Ca2+ release in skeletal muscle homogenates. More generally this is also in line with many previous studies on fragmented muscle preparations that highlighted the ryanodine receptor as a potential physiological target for NO (see Stamler & Meissner, 2001 for review). This brings us back to the questions concerning the relevance and the mechanisms of modulation of the calcium release channel activity by endogenous diffusible effectors, beyond the tight control exerted by the voltage sensor. Our results would be consistent with such a role for NO; one may even speculate that changes within the physiological range of intracellular NO levels may finely tune the sensitivity of the calcium release channel activation to the changes in t-tubule membrane voltage, so as to adjust the efficiency of the coupling process. Importantly, a shift in the voltage dependence of Ca2+ release with no modification in voltage-sensing steps was previously shown to occur under other types of conditions of alteration of the release channel function (see for instance Klein et al. 1990; Dietze et al. 2000). The affected E–C coupling step then probably takes place beyond charge movement, in a voltage-independent reaction step of the ryanodine receptor, as for instance modelled by Dietze et al. (2000).
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According to many previous works, one expects the modulation of the calcium release channel activity by NO to result from oxidation and/or S-nitrosylation of certain classes of thiols within the channel protein (Suko et al. 1999; Eu et al. 2000; Hart & Dulhunty, 2000; Sun et al. 2001a,b, 2003; Aracena et al. 2003). These studies also pointed out the potential complexity of this regulation because NO can result in either activation or inhibition of the isolated channel depending on various parameters, including its concentration. The precise mechanism (including the classes of thiols that react with NO) responsible for the effects described here is then certainly different from the mechanism triggered by large concentrations of NO which produce an increased SR calcium leak (Pouvreau et al. 2004). In particular, the use-dependence of the high NO-induced SR Ca2+ leak implied that the release channels' reactivity to large levels of NO was conditioned by prior opening of the channels. Rather, the present results suggest that the release channel protein also has some constant reactivity versus endogenous levels of NO, and that this tends to maintain calcium release in a chronically somewhat reduced level of sensitivity to activation by the voltage sensors.
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The increased voltage sensitivity of Ca2+ release by NOs inhibition provides a consistent explanation for the previously reported enhanced contractile function of various skeletal muscle preparations by NOs inhibitors (Kobzik et al. 1994; Gath et al. 1996; Reid et al. 1998; Richmonds & Kaminski, 2001). However, one has to be aware that the relevance of these previous works and of the present one is challenged by the potential influence of the PO2. Indeed carrying out experiments on isolated muscle preparations at ambient non-physiological oxygen tension when skeletal muscle PO2 is in the range of 4–20 mmHg (0.5–2.5% O2) is a definite limitation in these studies. This is particularly important to consider in light of the results from Eu et al. (2003) showing that in the presence of physiological (low) O2 concentration twitch force was reduced by NOs inhibition, whereas at increased O2 concentration (ambient conditions) twitch force was enhanced by NOs inhibition. This latter effect was believed to be unlikely to involve a direct modulation of Ca2+ release because of a presumably poor responsiveness of the ryanodine receptor to NO at supranormal PO2. This clearly was not true here because, despite the experimental ambient O2 environment, Ca2+ release was clearly affected by NOs inhibition. Nevertheless, we have no right to extrapolate our conclusions to the physiological PO2 situation.
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According to Eu et al. (2000, 2003) the normal NO-responsive configuration of the ryanodine receptor is achieved at physiological O2 concentrations because of the reduced redox state of a small subset of thiol groups. These thiols would be oxidized at supranormal PO2, making the channel insensitive to activation by endogenous NO. This, for instance, would be responsible for the observed 20–45% reduced force production at 20% versus 1% O2 in isolated muscle preparations (Eu et al. 2003). Whether under our experimental conditions the direction and/or extent of the observed shift in voltage sensitivity of Ca2+ release would differ depending on the O2 tension level remains undetermined and would be hard to test. However, relevant information was obtained from the effect of dithiothreitol (DTT), under the assumption that oxidized thiols (due to the ambient O2 level) within the ryanodine receptor protein would be reduced by application of this agent. DTT was indeed previously shown to reverse thiol oxidation on the skeletal muscle ryanodine receptor (Aghdasi et al. 1997; Haarmann et al. 1999) and to mimic the effects of reduced PO2 on the activity of various ion channels (Park et al. 1995; Perez-Garcia et al. 1999; Hool, 2000, 2004). Furthermore, we showed that DTT was effective under our experimental conditions in reversing the effects of large levels of NO on intracellular Ca2+ homeostasis (Pouvreau et al. 2004).
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However, we already showed that DTT had no obvious effect on the properties of the voltage-activated Ca2+ transient elicited by a short depolarizing pulse to +10 mV under our control conditions (see Fig. 8 in Pouvreau et al. 2004), whereas according to Eu et al. (2003) one may reasonably have expected a substantial increase in Ca2+ release under these conditions since force is enhanced under low PO2. Furthermore, we now demonstrate that DTT in addition does not affect the voltage dependence of Ca2+ release (see Fig. 8). Although one cannot exclude the possibility of PO2-sensitive disulphide bridges that are resistant to DTT reduction, these data do not favour an oxidized conformation of the ryanodine receptor under our experimental conditions.
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Along these lines we show that the oxidant H2O2 also produces a leftward shift in the voltage dependence of release activation without concurrent change in intramembrane charge movement. This effect, which is thus most likely to be related to an alteration of the ryanodine receptor function, also indicates that, despite the ambient conditions of O2 tension, the ryanodine receptor can be further oxidized. This result is consistent with data from Posterino et al. (2003) in skinned muscle fibres showing that oxidative treatment increased the effectiveness of voltage sensor activation of Ca2+ release when using submaximal conditions of depolarization. In agreement with their conclusions, less voltage sensor activation is indeed needed to open the release channels in the presence of H2O2, but our results indicate that this is also true when NO production is inhibited. The two phenomena may be mechanistically intimately linked, for instance if the presence of NO protects the channel against the effects of oxidation, as suggested by Aghdasi et al. (1997). Overall, it may also be speculated that changes in the voltage sensitivity of Ca2+ release channel activation may be a ubiquitous consequence of redox-related modifications of the ryanodine receptor protein, under physiological conditions.
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It should be noted that, under our conditions, the peak amplitude of the release was also substantially reduced in the presence of H2O2. Although we have no straightforward explanation for this, the strong oxidizing condition could have had additional, maybe deleterious, effects on the channel protein or on other components of the E–C coupling process (see also Posterino et al. 2003).
In conclusion, the present data provide important information concerning the mechanism of regulation of the excitation–contraction coupling process by NO. Although the overall picture is still not entirely clear, this work definitely contributes to our understanding of how NO affects skeletal muscle function. It may also prove helpful for resolving issues concerning diseases related to dysfunction of the NO signalling pathway in this tissue.
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Abstract
Nitric oxide (NO) generated by skeletal muscle is believed to regulate force production but how this is achieved remains poorly understood. In the present work we tested the effects of NO synthase (NOs) inhibitors on membrane current and intracellular calcium in isolated skeletal muscle fibres from mouse, under voltage-clamp conditions. Resting [Ca2+] and [Ca2+] transients evoked by large depolarizations exhibited similar properties in control fibres and in fibres loaded with tenth millimolar levels of the NOs inhibitor N-nitro-L-arginine (L-NNA). Yet the voltage dependence of calcium release was found to be shifted by 15 mV towards negative values in the presence of L-NNA. This effect could be reproduced by the other NOs inhibitor S-methyl-L-thiocitrulline (L-SMT). Separate experiments showed that the voltage dependence of charge movement and of the slow calcium current were unaffected by the presence of L-NNA, ruling out an effect on the voltage sensor. A negative shift in the voltage dependence of calcium release with no concurrent alteration in the properties of charge movement was also observed in fibres exposed to the oxidant H2O2 (1 mM). Conversely the reducing agent dithiothreitol (10 mM) had no obvious effect on Ca2+ release. Overall, the results indicate that physiological levels of NO exert a tonic inhibitory control on the activation of the calcium release channels. Changes in the voltage dependence of Ca2+ release activation may be a ubiquitous physiological consequence of redox-related modifications of the ryanodine receptor.
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Introduction
Nitric oxide is a universal signalling molecule with a key role in an extremely diverse variety of physiological and patho-physiological processes. Its relevance has been especially well established in smooth and cardiac muscle (see for recent reviews Ignarro, 2002 and Massion et al. 2003) but NO is also suspected to play an important role in skeletal muscle. Mature skeletal muscle fibres express the three major isoforms of NO synthase with a specific splice variant of the neuronal isoform being predominant (Silvagno et al. 1996). Contraction of isolated muscle preparations can be affected by exogenous NO and by inhibitors of NO synthesis, and both impairment of endogenous NO supply and overproduction of NO are thought to be involved in some skeletal muscle disorders (for reviews see Eu et al. 1999; Stamler & Meissner, 2001). In terms of signalling pathway, the situation is, however, far less clear than in the cardiovascular system where clear-cut NO effects occur through changes in the intracellular cGMP level, a second messenger that is of poorly defined relevance to skeletal muscle function. Rather, a massive body of evidence supports the view that in skeletal muscle, NO could largely act through mechanisms of oxidation and/or S-nitrosylation of free thiols, with many reports pointing to the sarcoplasmic reticulum (SR) calcium release channel as a major potential target protein (Meszaros et al. 1996; Aghdasi et al. 1997; Stoyanovsky et al. 1997; Suko et al. 1999; Eu et al. 2000, 2003; Hart & Dulhunty, 2000; Sun et al. 2001a, 2003). Under normal conditions, SR Ca2+ release by the ryanodine receptors is driven by a voltage-induced conformational change in the dihydropyridine receptors present in the t-tubule membrane. Whether or not NO production can modulate this process in situ and if it does, how this is achieved remains misunderstood. In this context the effects of NO synthesis inhibition are of strong interest: for instance various NO synthase inhibitors were shown to potentially enhance force production in isolated muscle preparations (Kobzik et al. 1994; Gath et al. 1996; Reid et al. 1998). Whether these effects on force production are related to a modulation of the SR Ca2+ release flux has not been established. In a recent study we showed that excess NO applied to an isolated skeletal muscle fibre produces a use-dependent increase in resting [Ca2+] consistent with some calcium release channels remaining locked open at rest (Pouvreau et al. 2004). In the present work we attempted to reveal the role of the endogenously produced NO levels on the control of intracellular [Ca2+]. Results show that inhibitors of NO synthase affect the voltage dependence of Ca2+ release and that a similar effect is produced by the oxidant H2O2.
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Methods
Preparation of the muscle fibres
Experiments were performed on single skeletal fibres isolated from the flexor digitorum brevis (FDB) muscles from 4- to 8-week-old OF1 mice. All experiments were performed in accordance with the guidelines of the French Ministry of Agriculture (87/848) and of the European Community (86/609/EEC). Procedures for enzymatic isolation of single fibres, partial insulation of the fibres with silicone grease and intracellular dye loading, were as previously described (Jacquemond, 1997; Collet et al. 1999, 2004; Collet & Jacquemond, 2002). In brief, mice were killed by cervical dislocation before removal of the muscles. Muscles were treated with collagenase (ICN type 1) for 60–75 min at 37°C in the presence of external Tyrode solution. Single fibres were then obtained by triturating the muscles within the experimental chamber. The major part of a single fibre was electrically insulated with silicone grease so that whole-cell voltage clamp could be achieved on a short portion of the fibre extremity. Prior to voltage clamp, indo-1 was introduced locally into the fibre by pressure microinjection through a micropipette containing 0.5 mM indo-1 dissolved in an intracellular-like solution (see ‘Solutions’). Fibres were then left for 1 h to allow for intracellular equilibration of the dye. The NOs inhibitors N-nitro-L-arginine (L-NNA) and S-methyl-L-thiocitrulline (L-SMT) were in some cases present in the microinjected solution (see ‘Solutions’). Following diffusion and equilibration within the cytoplasm, it was believed that final cytoplasmic concentrations of these compounds within one tenth to one fifth of the initial pipette concentration were achieved (for details concerning microinjections see Csernoch et al. 1998). For each series of measurements under a given condition, control fibres used for the analysis were always issued from the same muscles as the test fibres. In some experiments, the two collagenase-treated FDB muscles from the same mouse were kept overnight at 4°C. One muscle was kept in the Tyrode solution for control measurements. The other muscle was in the Tyrode solution which also contained 5 mM each of L-NNA and L-SMT; L-NNA and L-SMT were present in the solution bathing the fibres during all subsequent steps of the experimental procedure. All experiments were performed at room temperature (20–22°C).
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Electrophysiology
An RK-400 patch-clamp amplifier (Bio-Logic, Claix, France) was used in whole-cell configuration. Command voltage pulse generation and data acquisition were done using commercial software (Biopatch Acquire, Bio-Logic) driving an A/D, D/A converter (LabMaster DMA board, Scientific Solutions Inc., Mentor, OH, USA). Analog compensation was systematically used to decrease the effective series resistance. Voltage clamp was performed with a microelectrode filled with the intracellular-like solution. The tip of the microelectrode was inserted through the silicone, within the insulated part of the fibre. Unless otherwise specified, membrane depolarizations (or series of depolarizations) were applied every 30 s from a holding command potential of –80 mV.
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Measurement of the slow calcium current
Calcium current records were obtained in response to 1 s-long depolarizing steps to various potentials applied every 30 s. The linear leak component of the current was removed by subtracting the adequately scaled value of the steady current measured during a 20 mV hyperpolarizing step applied before each test pulse. The voltage dependence of the peak calcium current density was fitted with the following equation:
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where I(V) is the peak current density at the command potential V, Gmax is the maximum conductance, Vrev is the apparent reversal potential, V0.5 is the half-activation potential and k is a steepness factor.
Measurement and analysis of intramembrane charge movement
Charge movement currents were measured and analysed according to previously described procedures (Collet et al. 2003; Pouvreau et al. 2004). For these measurements the holding potential was set to –100 mV. In brief, adequately scaled control current records elicited by 50 ms-long hyperpolarizing pulses of 20 mV were subtracted from the current elicited by test depolarizing pulses of the same duration to various levels. The amount of charge moved during a test pulse was measured by integrating the ‘on’ portion of the corrected test current records. The calculated charge was normalized to the capacitance of the fibre. The steady-state distribution of the normalized charge was fitted with a two-state Boltzmann function:
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where Qmax corresponds to the maximal available charge, V0.5 to the voltage of equal charge distribution and k to the steepness factor.
Fluorescence measurements
The optical set-up and the procedures used for indo-1 fluorescence measurements were as previously described (Jacquemond, 1997; Collet et al. 1999; Collet & Jacquemond, 2002; Pouvreau et al. 2004). In brief, a Nikon Diaphot epifluorescence microscope was used in diafluorescence mode. The beam of light from a high-pressure mercury bulb set on the top of the microscope was passed through a 335 nm interference filter and focused onto the preparation using a quartz aspherical doublet. The emitted indo-1 fluorescence light was collected by a x 40 objective and simultaneously detected at 405 ± 5 nm (F405) and 470 ± 5 nm (F470) by two photomultipliers. The fluorescence measurement field was 40 μm in diameter and the silicone-free extremity of each tested fibre was placed in the middle of the field. Background fluorescence at both emission wavelengths was measured next to each tested fibre and was then subtracted from all measurements.
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Calibration of the indo-1 response and [Ca2+]i calculation
The standard ratio method was used with the parameters: R = F405/F470, with Rmin, Rmax, KD and having their usual definitions. Results were either expressed in terms of indo-1% saturation or in actual free calcium concentration (for details of calculation, see Jacquemond, 1997; Csernoch et al. 1998). In vivo values for Rmin, Rmax and were measured using procedures previously described (Collet et al. 1999; Collet & Jacquemond, 2002). Unless otherwise specified, no correction was made for indo-1–Ca2+ binding and dissociation kinetics. In vitro measurements of the indo-1 response performed on the experimental set-up (Jacquemond, 1997) showed that the dye response to calcium was unaffected by L-NNA (not illustrated).
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Calculation of the rate of calcium release
The method used to estimate the SR calcium release flux from the indo-1 transients involved taking the time derivative of the total myoplasmic Ca2+ calculated from the occupancy of the main intracellular calcium binding sites (Baylor et al. 1983). The model included troponin C binding sites with a total site concentration TNtotal = 250 μM, an ‘on’ rate constant kon,CaTN = 0.0575 μM–1 ms–1 and an ‘off’ rate constant koff,CaTN = 0.115 ms–1; Ca2+–Mg2+ binding sites on parvalbumin with a total site concentration PVtotal = 700 μM, ‘on’ rate constant for Ca2+ kon,CaPV = 0.125 μM–1 ms–1, ‘off’ rate constant for Ca2+ koff,CaPV = 5 x 10–4 ms–1, ‘on’ rate constant for Mg2+ kon,MgPV = 3.3 x 10–5 μM–1 ms–1, ‘off’ rate constant for Mg2+ koff,MgPV = 3 x 10–3 ms–1. Resting [Mg2+] was assumed to be 1.5 mM. For these calculations free calcium transients were corrected for the Ca2+–indo-1 binding kinetics using values for the ‘on’ and ‘off’ rate constants of the Ca2+–indo-1 binding reaction of 1 x 108 M–1 s–1 and 30 s–1, respectively, by analogy with the values derived for fura-2 by Baylor & Hollingworth (1988).
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Solutions
The intracellular-like solution contained (mM): 120 potassium glutamate, 5 Na2-ATP, 5 Na2-phosphocreatine, 5.5 MgCl2, 5 glucose, 5 Hepes adjusted to pH 7.20 with KOH. In some experiments the NOs inhibitors N-nitro-L-arginine (L-NNA) and S-methyl-L-thiocitrulline (L-SMT) were used. When microinjected, the respective inhibitors were present in the micropipette solution at the following concentrations: L-NNA, 5 mM; L-SMT, 5 or 20 mM. For the incubated fibres L-NNA and L-SMT were present in the extracellular solution at a concentration of 5 mM. The standard extracellular solution contained (mM): 140 TEA-methanesulphonate, 2.5 CaCl2, 2 MgCl2, 10 TEA-Hepes and 0.002 tetrodotoxin, pH 7.20. For measurements of the slow calcium current the extracellular solution also contained 4-aminopyridine (1 mM) and the fibres were pressure-microinjected with a solution containing 50 mM EGTA diluted in the intracellular-like solution. For measurements of charge movement the extracellular solution contained 140 TEA-methanesulphonate, 0.5 CaCl2, 4 MgCl2, 1 4-aminopyridine, 0.5 or 1 CdCl2, 0.3 LaCl3, 10 TEA-Hepes and 0.002 tetrodotoxin, pH 7.20. In addition fibres were pressure-microinjected with a solution containing 10 mM EGTA diluted in the intracellular-like solution.
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Statistics
Least-squares fits were performed using a Marquardt-Levenberg algorithm routine included in Microcal Origin (OriginLab, Northampton, MA, USA). Data values are presented as means ± S.E.M. for n fibres, where n is specified in Results. Statistical significance was determined using Student's t test assuming significance for P < 0.05.
Results
Calcium transients in response to depolarizing pulses of increasing duration in the presence of the NOs inhibitor L-NNA
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Figure 1A shows mean (continuous line) ± S.E.M. (grey shading) indo-1 saturation traces elicited from control fibres (n = 20) and from fibres injected with L-NNA (n = 11) in response to successive depolarizing pulses of 20, 70 and 120 ms duration to +10 mV. In each panel the inset shows an example of traces from an individual fibre in the given condition. Obviously the presence of L-NNA did not dramatically affect the qualitative features of the transients, that is a rise triggered by the depolarization with an initial very rapid phase, and a decline to the resting level that immediately follows membrane repolarization. The decaying phase of the transients appeared though to be somewhat faster in the presence of L-NNA than in control conditions. Figure 1B presents mean values from that series of measurements for resting [Ca2+], peak [Ca2+] and time constant of [Ca2+] decay. Measurements were performed on individual [Ca2+] traces calculated from each fibre. The time constant of decay was obtained from fitting a single exponential plus constant function to the [Ca2+] decline as previously described (Collet et al. 1999; Pouvreau et al. 2004). The resting [Ca2+] and peak [Ca2+] values did not significantly differ between the two groups. The mean value for the time constant of [Ca2+] decay was 25–30% lower in the presence of L-NNA, the difference being significant only for the shortest pulse duration (P = 0.02). Overall, these results indicate that the presence of a potent inhibitor of nNOS activity within the intracellular medium did not impair the ability of the fibres to regulate [Ca2+] at rest and to produce robust calcium transients in response to membrane depolarization. The reason for the slightly faster decay of the transients in response to short pulses in the presence of L-NNA is unclear. It may be the consequence of a chronic effect of NO on a component of the cytoplasmic Ca2+ removal systems. This effect was not further investigated within the present work.
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A, mean (continuous lines) ± S.E.M. (grey shading) indo-1 saturation signals obtained in response to depolarizations of 20, 70 and 120 ms duration to +10 mV in control fibres and in fibres injected with L-NNA. In each panel the inset shows an example of traces measured from a single fibre under the corresponding condition. B, mean ± S.E.M. values of resting [Ca2+], peak change in [Ca2+] and time constant of [Ca2+] decay measured from the individual control fibres and L-NNA-injected fibres under the conditions shown in A.
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Alteration of the voltage dependence of Ca2+ release activation in the presence of NOs inhibitors
Figure 2 illustrates what we believe is the prevailing effect of NOs inhibition on the control of intracellular calcium. Indo-1 calcium transients were examined in control fibres and in fibres treated with the NOs inhibitors L-NNA or (and) L-SMT using the pulse protocol shown above the corresponding series of records in Fig. 2. The protocols consisted of three consecutive depolarizations of 50 ms duration to the indicated values; each pulse was then incremented by 5 mV and the sequence repeated. Within five runs this allowed an explicit examination of the voltage dependence of the properties of the transients. In addition, two levels of depolarizations (–30 and –10 mV) were tested twice, allowing the stability of the preparation to be checked during the course of the protocol. The interval between two pulses within a run was either 0.5 s (as in the top series of records) or 1 s; this did not affect the mean values for the parameters that were measured from the data under a given condition (not illustrated). Qualitatively, traces in Fig. 2A and B show that, as compared to the control conditions (left series of records), the indo-1 transients elicited in the fibres injected with L-NNA or L-SMT tended to emerge for lower levels of membrane depolarization and the maximal amplitude was also reached for less depolarized levels. Figure 2C illustrates the fact that this was also the case in fibres that were kept for several hours in the presence of L-NNA and L-SMT before taking measurements (see Methods). In order to quantitatively estimate this effect, free calcium transients and corresponding release fluxes were calculated from the indo-1 transients as described in Methods. An example of the [Ca2+] trace and of the corresponding release flux is shown in Fig. 3A and C, respectively. Figure 3B shows the voltage dependence of the mean normalized peak [Ca2+] transients from control fibres (filled symbols, n = 19), from fibres injected with L-NNA (, n = 8), from fibres injected with L-SMT (, n = 7) and from fibres incubated in the presence of both inhibitors (, n = 4). In each fibre all values for peak [Ca2+] were normalized to the maximal peak value that was reached during the protocol. For the fibres incubated with L-NNA and L-SMT, control data that are shown (filled squares, n = 3) were from fibres from the same batch, incubated during the same amount of time in control solution (see Methods). Figure 3D shows the mean voltage dependence of the peak release flux calculated from the same data. A negative shift in the voltage dependence of release activation was manifest under the three tested conditions of NOs inhibition. The individual release flux versus voltage relationship from each individual fibre was fitted with a Boltzmann function. Mean values for the maximal release flux and for the steepness factor were 4.6 ± 0.6 μM ms–1 and 6.0 ± 0.3 mV in control fibres (n = 19), 4.9 ± 0.6 μM ms–1 and 6.6 ± 0.2 mV in L-NNA-injected fibres (n = 8) and 3.4 ± 0.6 μM ms–1 and 7.0 ± 0.5 mV in L-SMT-injected fibres (n = 7), respectively. Both parameters were not significantly different between control and treated fibres. The same was true for the batch of incubated fibres: mean values for the maximal release flux and steepness factor were 3.8 ± 0.9 μM ms–1 and 6.7 ± 0.6 mV in control fibres (n = 3) and 3.2 ± 0.5 μM ms–1 and 6.2 ± 0.2 mV in fibres incubated with L-NNA and L-SMT (n = 4), respectively. Conversely mean values for the midpoint voltage were significantly more negative in the NOs inhibitor-treated fibres than in the control fibres: they were –17.0 ± 2.0, –29.2 ± 2.3 and –27.4 ± 2.1 mV in control fibres (n = 19), fibres injected with L-NNA (n = 8, P = 0.002) and in fibres injected with L-SMT (n = 7, P = 0.008), respectively. Accordingly the mean midpoint voltage from fibres incubated overnight in the presence of both inhibitors was –31.8 ± 2.4 mV (n = 4) as compared to –17.6 ± 1 mV (n = 3, P = 0.005) for the concurrent batch of control fibres. It should be mentioned that the estimation of the half-activation potential in Fig. 3B was certainly more accurate in the NOs inhibitor-treated fibres (for which saturation was reached early within the explored voltage range) than in the control ones. The actual value of the half-activation voltage in the control fibres may be slightly more positive than estimated from our fits and this would tend to a somewhat underestimation of the difference with the fibres treated with the NOs inhibitors.
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A, B and C, indo-1 saturation signals elicited in three distinct control fibres. D, E and F, indo-1 saturation signals elicited in a L-NNA-injected fibre, in a L-SMT-injected fibre and in a fibre incubated in the presence of L-NNA and L-SMT prior to the experiment, respectively (see text and Methods for details). The pulse protocol that was used is shown above each series (A and D) or pair of series (B and C, E and F) of records; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV.
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A, example of [Ca2+] trace calculated from an indo-1 signal elicited by consecutive depolarizing pulses to –30, –10 and +10 mV (protocol shown in Fig. 2). C, corresponding Ca2+ release flux trace calculated as described in Methods. B, voltage dependence of the mean relative peak change in [Ca2+] in control fibres (, n = 19), in L-NNA-injected fibres (, n = 8), in L-SMT-injected fibres (, n = 7) and in fibres incubated in the presence of L-NNA and L-SMT (, n = 4). Control data for the fibres incubated with L-NNA and L-SMT were from fibres incubated for the same amount of time in control solution (, n = 3). In each fibre the value for peak [Ca2+] elicited by a given pulse was normalized to the maximum measured value. D, corresponding voltage dependence of the mean peak Ca2+ release flux. The superimposed continuous lines correspond to the results from fitting a Boltzmann function to the mean data points: corresponding values for maximum release flux, midpoint voltage and steepness factor in the two batches of control fibres were 4.8 μM ms–1, –15 mV and 8.6 mV () and 3.8 μM ms–1, –17.4 mV and 6.8 mV (), respectively. Values from the fits to data points in L-NNA-injected fibres, in L-SMT-injected fibres and in fibres incubated in the presence of L-NNA and L-SMT were 4.9 μM ms–1, –28.4 mV and 7.5 mV, 3.41 μM ms–1, –25.6 mV and 7.8 mV, and 3.2 μM ms–1, –31.1 mV and 6.3 mV, respectively.
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Function of the dihydropyridine receptor in the presence of a NOs inhibitor
The effects of NOs inhibitors on Ca2+ release activation could be due to a modification of the voltage-sensing steps of excitation–contraction (E–C) coupling. In order to check for this, we looked at the calcium channel and voltage-sensing functions of the dihydropyrdine receptor. We first examined the membrane current from the experiments described in relation to Fig. 3. An inward current was clearly flowing during the most depolarized values of step potential, indicative of the activation of the slow inward calcium current (not shown). Analysis of the voltage dependence of the end-pulse current in the control fibres, in the fibres injected with L-NNA, and in the fibres injected with L-SMT revealed that at –20 mV the current was significantly larger in the group of L-NNA-injected fibres (–0.96 ± 0.3, n = 8) than in the control fibres (–0.34 ± 0.14, n = 17). However, for other potential values there was no significant difference between control fibres and either L-NNA-injected fibres or L-SMT-injected fibres.
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In order to more carefully assess a possible effect of NOs inhibition on the channel function of the dihydropyridine receptor, we performed a specific series of calcium current measurements in a batch of control and L-NNA-injected fibres using 1 s-long depolarizations (see Methods). Figure 4A shows calcium current traces measured from a control fibre (left) and from an L-NNA-injected fibre (right) obtained in response to membrane depolarizations ranging between –50 and +70 mV with a 5 mV increment. There was no striking difference between the two. Figure 4B shows the voltage dependence of the mean peak calcium current density from control fibres (, n = 7) and L-NNA-injected fibres (, n = 9). Fitting the individual I–V curves as described in Methods gave mean values for Gmax, Vrev, V0.5 and k of 142 ± 12 S F–1, 62.2 ± 3 mV, –1.2 ± 2 mV and 6.7 ± 0.6 mV in control fibres, and 127 ± 7 S F–1, 60.6 ± 3.5 mV, 1.6 ± 2 mV and 6.8 ± 0.5 mV in L-NNA-injected fibres, respectively. None of these parameters significantly differ between the two groups.
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A, calcium current records obtained in response to 1 s-long depolarizing pulses to values between –50 and +70 mV in a control fibre (left) and in a fibre pressure-microinjected with L-NNA (right). B, mean voltage dependence of the peak calcium current amplitude in control fibres (, n = 7) and in L-NNA-injected fibres (, n = 9). The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details).
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In order to further explore the possibility that the voltage sensor may be affected by the treatment with NOs inhibitors we performed a specific series of experiments designed to measure intramembrane charge movement in control and in L-NNA-injected fibres. Figure 5A shows representative traces of intramembrane charge current from a control fibre (left) and from an L-NNA-injected fibre (right). There was no obvious difference between the two series. Figure 5B shows the mean voltage distribution of the ‘on’ charge in the two groups of fibres. Fitting a two-state Boltzmann distribution to the amount of ‘on’ charge versus voltage data in each fibre gave mean values for Qmax, V0.5 and k of 24 ± 3 nC μF–1, –30.7 ± 2 mV and 11 ± 1 mV in control fibres (n = 6), and 24 ± 4 nC μF–1, –30.8 ± 2 mV and 11.3 ± 2 mV in L-NNA-injected fibres (n = 7), respectively. Obviously, none of the parameters was significantly different between the two conditions.
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A, charge movement records from a control fibre and from an L-NNA-injected fibre, measured in response to 50 ms-long depolarizing steps to the indicated values of command potential. B, mean voltage distribution of the ‘on’ charge in control fibres (n = 6) and in L-NNA-injected fibres (n = 7). Superimposed continuous lines correspond to the result of fitting a two-state Boltzmann distribution.
Effect of H2O2 on voltage-activated calcium release
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Many previously reported effects of the NO signalling pathway on skeletal muscle contractility, intracellular calcium regulation and on the activity of the isolated ryanodine receptor were either shown or at least suspected to be intimately linked to redox-based modulation of protein function (for review see Reid, 2001; Stamler & Meissner, 2001). For instance it was shown that low concentrations of NO could prevent activation of the calcium release channel by oxidation (Aghdasi et al. 1997). Within the present context it was therefore of particular interest to study how calcium release is affected by the presence of an oxidant under the same conditions as above. This was done by exposing isolated fibres to H2O2. Figure 6A shows indo-1 saturation traces obtained in response to the same protocol as in Fig. 2 in a control fibre and in a fibre bathed in the presence of H2O2 (1 mM). As observed in the presence of NOs inhibitors, transients were detected for lower voltage levels in the fibre exposed to H2O2 than in the control fibre and the maximal peak amplitude of the transient also tended to be reached at lower levels of membrane depolarization. Figure 6B shows the voltage dependence of the mean normalized peak change in [Ca2+] (left) and of the corresponding mean peak release flux (right) in both groups of fibres. There was a definite leftward shift in the voltage dependence of Ca2+ release in the presence of H2O2, but this was also associated with a reduction of the maximal peak release flux. The individual peak release flux versus voltage relationship from each fibre was fitted with a Boltzmann function. Mean values for the maximal release flux, midpoint voltage and steepness factor were 7.7 ± 0.4 μM ms–1, –18.8 ± 1.6 mV and 6.0 ± 0.4 mV in control fibres (n = 10), and 4.2 ± 0.5 μM ms–1, –26.9 ± 3 mV and 7.4 ± 0.7 mV in fibres exposed to H2O2 (n = 7), respectively. Values for maximal flux and for midpoint voltage were significantly different.
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A, indo-1 saturation signals elicited by a series of 50 ms-long depolarizing pulses of increasing amplitude in a control fibre and in a fibre bathed in the presence of 1 mM H2O2. The pulse protocol that was used is shown on the top of the panel; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV. B, voltage dependence of the mean relative peak change in [Ca2+] and of the corresponding mean peak Ca2+ release flux in control fibres (, n = 10) and in fibres exposed to H2O2 (, n = 7). Four fibres were bathed in the presence of the oxidant for 15–30 min before starting to take the measurements and three fibres were bathed in H2O2 for a longer period of time (1 h or more). The mean values for the Boltzmann parameters obtained from fitting the voltage dependence of the peak release flux in each fibre did not differ between these two sets of H2O2-treated fibres (not illustrated). Continuous lines superimposed on the release flux data correspond to the result of fitting a Boltzmann function to the mean data points: this gave values for maximum release flux, midpoint voltage and steepness factor of 7.8 μM ms–1, –19.4 mV and 6.9 mV in control fibres, and of 4.0 μM ms–1, –28.6 mV and 7.9 mV in fibres exposed to H2O2. It should be noted that this series of experiments was performed on fibres issued from a different batch of mice, and at a different period of time as compared to other results presented in this paper; this may be the reason for the difference in absolute values of peak release flux between control fibres from this series as compared to control data presented in relation to Fig. 3.
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Function of the dihydropyridine receptor in the presence of H2O2
In order to test whether the effects of H2O2 on Ca2+ release could be due to an alteration of the voltage-sensing functions of E–C coupling we measured, in separate sets of experiments, the slow calcium current and the intramembrane charge movements in the presence of the oxidant. Results are illustrated in Fig. 7. Figure 7A shows a series of calcium current records elicited by 1 s-long membrane depolarizations of increasing amplitude in a fibre exposed to 1 mM H2O2, whereas Fig. 7B shows the mean peak current versus voltage (I–V) relationships obtained from this series of measurements in four control fibres () and seven fibres exposed to H2O2 (). The peak current was somewhat reduced in the presence of H2O2. Fitting the individual I–V curves as described in Methods gave mean values for Gmax, Vrev, V0.5 and k of 176 ± 27 S F–1, 56.4 ± 2.9 mV, –0.02 ± 1.6 mV and 7.8 ± 0.9 mV in control fibres (n = 4), and 113 ± 11 S F–1, 56.6 ± 3.7 mV, –6.4 ± 3.6 mV and 8.2 ± 1.1 mV in H2O2-treated fibres (n = 7), respectively. The mean maximum conductance was reduced by 35% in the presence of H2O2 (P = 0.03). Other parameters did not differ significantly.
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A, calcium current records obtained in response to 1 s-long depolarizing pulses to values between –50 and +70 mV in a fibre exposed to 1 mM H2O2. B, mean voltage dependence of the peak calcium current amplitude in control fibres (, n = 4) and in H2O2-exposed fibres (, n = 7). The superimposed curves were calculated using the mean parameters obtained from fitting the voltage dependence in each fibre (see text for details). C, charge movement records from a fibre exposed to 1 mM H2O2 measured in response to 50 ms-long depolarizing steps to values ranging between –80 and +15 mV. D, mean voltage distribution of the ‘on’ charge in control fibres (n = 6) and in fibres exposed to 1 mM H2O2 (n = 7). For these two sets of measurements H2O2-treated fibres were exposed to the oxidant for 15 min before starting the experiment.
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Figure 7C shows an example of charge movement traces obtained from a fibre bathed in the presence of 1 mM H2O2. Traces were obtained in response to 50 ms-long depolarizing pulses to voltages ranging between –80 mV (bottom) and +15 mV (top) with a 5 mV increment from a holding potential of –100 mV. There was no sign of gross alteration of the properties of the ‘on’ and ‘off’ charge transients. Figure 7D shows the mean voltage dependence of the ‘on’ charge in the H2O2-treated fibres (, n = 7) and in control fibres isolated from the same mice (, n = 6). There was no difference between the two populations. Fitting the individual series of data points with a Boltzmann function gave mean values for Qmax, V0.5 and k of 26.3 ± 3.2 nC –F–1, –29.5 ± 3.7 mV and 14.1 ± 2.8 mV in control fibres, and 27.0 ± 1.9 nC μF–1, –31.1 ± 2.1 mV and 13.1 ± 1.7 mV in H2O2-treated fibres, respectively.
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The reason for the H2O2-induced reduction of the calcium current conductance is unclear. It may be due to the existence of redox-sensitive uncharged transitions leading to the channel opening. In any case this effect cannot be taken as responsible for the reduced calcium release observed in the presence of H2O2 as it is well established that the fraction of calcium entering a muscle fibre through this pathway is not significant compared to the amount released from the SR. Overall this set of data indicates that the effects of H2O2 on calcium release are not related to the voltage-sensing steps of E–C coupling and are thus very likely to be due to specific alterations of the ryanodine receptor.
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Voltage-activated calcium release in the presence of dithiothreitol
We also tested the possibility that the ryanodine receptor would be in an oxidized state under our experimental conditions of ambient PO2 (see Eu et al. 2003) by measuring calcium release in the presence of the reducing agent dithiothreitol (DTT). For this calcium transients were measured first in response to 50 ms-long pulses of increasing amplitude (same 3-step protocol as in Figs 2 and 6) under control conditions; the extracellular solution was then replaced by a 10 mM DTT-containing one and the protocol was re-applied. Figure 8A illustrates the results from such an experiment. Traces shown underneath the pulse protocol correspond to indo-1 saturation signals measured in the control solution while bottom traces were obtained from the same fibre, 10 min after exchanging the extracellular solution for the DTT-containing solution. There was no obvious difference between the two series of traces. Figure 8B shows the voltage dependence of the mean normalized peak change in [Ca2+] (left) and of the corresponding mean peak release flux (right) from six fibres under control conditions () and after 10 min exposure to 10 mM DTT (). The mean values for maximal release flux, midpoint voltage and steepness factor obtained from fitting a Boltzmann function to the voltage dependence of Ca2+ release under control conditions were 9.6 ± 1 μM ms–1, –15.2 ± 2.2 mV and 6.1 ± 0.6 mV. Corresponding values in the presence of DTT were 8.5 ± 0.7 μM ms–1, –19.9 ± 3.2 mV and 6.0 ± 0.3 mV, respectively. None of the parameters significantly differ between the two conditions.
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A, indo-1 saturation signals elicited by a series of 50 ms-long depolarizing pulses of increasing amplitude in the same fibre under control conditions (upper records) and in the presence of 10 mM DTT (lower records). The pulse protocol that was used is shown at the top of the panel; the value of membrane potential indicated next to each series of steps corresponds to the first potential level that was applied from –80 mV. B, voltage dependence of the mean relative peak change in [Ca2+] and of the corresponding mean peak Ca2+ release flux from the same fibres (n = 6) under control conditions () and after 10 min exposure to 10 mM DTT (). Continuous lines superimposed on the release flux data correspond to the result of fitting a Boltzmann function to the mean data points: this gave values for maximum release flux, midpoint voltage and steepness factor of 9.5 μM ms–1, –15 mV and 6.5 mV under control conditions, and of 8.6 μM ms–1, –19.5 mV and 7.5 mV in the presence of DTT.
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Discussion
The present work provides further insights into the role of NO in the physiological regulation of SR Ca2+ release. Mainly we demonstrate here that conditions designed to block endogenous production of NO in an intact muscle fibre shift the voltage dependence of SR Ca2+ release activation towards more negative values. The most straightforward inference is that physiological intracellular levels of NO tend to reduce the sensitivity of calcium release activation to voltage, and thus exert a chronic negative effect on the ‘efficiency’ of the E–C coupling process. The effect was observed with two potent inhibitors of constitutive isoforms of nitric oxide synthase, L-NNA and L-SMT, used at sufficiently large concentrations to presumably ensure a strong inhibition of the enzyme activity (see Boer et al. 2000; Alderton et al. 2001).
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The first thinkable candidate responsible for a shift in voltage dependence of calcium release would be the voltage-sensing dihydropyridine receptor in the t-tubule membrane. However, the properties of intramembrane charge movement (including the voltage dependence) were unaffected by L-NNA which suggests that it was the calcium release channel that was involved in the present effects of NOs inhibition. This is in agreement with the results from Meszaros et al. (1996) showing that nitro-L-arginine methyl ester blocked a NO (generated from L-arginine)-induced inhibition of Ca2+ release in skeletal muscle homogenates. More generally this is also in line with many previous studies on fragmented muscle preparations that highlighted the ryanodine receptor as a potential physiological target for NO (see Stamler & Meissner, 2001 for review). This brings us back to the questions concerning the relevance and the mechanisms of modulation of the calcium release channel activity by endogenous diffusible effectors, beyond the tight control exerted by the voltage sensor. Our results would be consistent with such a role for NO; one may even speculate that changes within the physiological range of intracellular NO levels may finely tune the sensitivity of the calcium release channel activation to the changes in t-tubule membrane voltage, so as to adjust the efficiency of the coupling process. Importantly, a shift in the voltage dependence of Ca2+ release with no modification in voltage-sensing steps was previously shown to occur under other types of conditions of alteration of the release channel function (see for instance Klein et al. 1990; Dietze et al. 2000). The affected E–C coupling step then probably takes place beyond charge movement, in a voltage-independent reaction step of the ryanodine receptor, as for instance modelled by Dietze et al. (2000).
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According to many previous works, one expects the modulation of the calcium release channel activity by NO to result from oxidation and/or S-nitrosylation of certain classes of thiols within the channel protein (Suko et al. 1999; Eu et al. 2000; Hart & Dulhunty, 2000; Sun et al. 2001a,b, 2003; Aracena et al. 2003). These studies also pointed out the potential complexity of this regulation because NO can result in either activation or inhibition of the isolated channel depending on various parameters, including its concentration. The precise mechanism (including the classes of thiols that react with NO) responsible for the effects described here is then certainly different from the mechanism triggered by large concentrations of NO which produce an increased SR calcium leak (Pouvreau et al. 2004). In particular, the use-dependence of the high NO-induced SR Ca2+ leak implied that the release channels' reactivity to large levels of NO was conditioned by prior opening of the channels. Rather, the present results suggest that the release channel protein also has some constant reactivity versus endogenous levels of NO, and that this tends to maintain calcium release in a chronically somewhat reduced level of sensitivity to activation by the voltage sensors.
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The increased voltage sensitivity of Ca2+ release by NOs inhibition provides a consistent explanation for the previously reported enhanced contractile function of various skeletal muscle preparations by NOs inhibitors (Kobzik et al. 1994; Gath et al. 1996; Reid et al. 1998; Richmonds & Kaminski, 2001). However, one has to be aware that the relevance of these previous works and of the present one is challenged by the potential influence of the PO2. Indeed carrying out experiments on isolated muscle preparations at ambient non-physiological oxygen tension when skeletal muscle PO2 is in the range of 4–20 mmHg (0.5–2.5% O2) is a definite limitation in these studies. This is particularly important to consider in light of the results from Eu et al. (2003) showing that in the presence of physiological (low) O2 concentration twitch force was reduced by NOs inhibition, whereas at increased O2 concentration (ambient conditions) twitch force was enhanced by NOs inhibition. This latter effect was believed to be unlikely to involve a direct modulation of Ca2+ release because of a presumably poor responsiveness of the ryanodine receptor to NO at supranormal PO2. This clearly was not true here because, despite the experimental ambient O2 environment, Ca2+ release was clearly affected by NOs inhibition. Nevertheless, we have no right to extrapolate our conclusions to the physiological PO2 situation.
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According to Eu et al. (2000, 2003) the normal NO-responsive configuration of the ryanodine receptor is achieved at physiological O2 concentrations because of the reduced redox state of a small subset of thiol groups. These thiols would be oxidized at supranormal PO2, making the channel insensitive to activation by endogenous NO. This, for instance, would be responsible for the observed 20–45% reduced force production at 20% versus 1% O2 in isolated muscle preparations (Eu et al. 2003). Whether under our experimental conditions the direction and/or extent of the observed shift in voltage sensitivity of Ca2+ release would differ depending on the O2 tension level remains undetermined and would be hard to test. However, relevant information was obtained from the effect of dithiothreitol (DTT), under the assumption that oxidized thiols (due to the ambient O2 level) within the ryanodine receptor protein would be reduced by application of this agent. DTT was indeed previously shown to reverse thiol oxidation on the skeletal muscle ryanodine receptor (Aghdasi et al. 1997; Haarmann et al. 1999) and to mimic the effects of reduced PO2 on the activity of various ion channels (Park et al. 1995; Perez-Garcia et al. 1999; Hool, 2000, 2004). Furthermore, we showed that DTT was effective under our experimental conditions in reversing the effects of large levels of NO on intracellular Ca2+ homeostasis (Pouvreau et al. 2004).
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However, we already showed that DTT had no obvious effect on the properties of the voltage-activated Ca2+ transient elicited by a short depolarizing pulse to +10 mV under our control conditions (see Fig. 8 in Pouvreau et al. 2004), whereas according to Eu et al. (2003) one may reasonably have expected a substantial increase in Ca2+ release under these conditions since force is enhanced under low PO2. Furthermore, we now demonstrate that DTT in addition does not affect the voltage dependence of Ca2+ release (see Fig. 8). Although one cannot exclude the possibility of PO2-sensitive disulphide bridges that are resistant to DTT reduction, these data do not favour an oxidized conformation of the ryanodine receptor under our experimental conditions.
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Along these lines we show that the oxidant H2O2 also produces a leftward shift in the voltage dependence of release activation without concurrent change in intramembrane charge movement. This effect, which is thus most likely to be related to an alteration of the ryanodine receptor function, also indicates that, despite the ambient conditions of O2 tension, the ryanodine receptor can be further oxidized. This result is consistent with data from Posterino et al. (2003) in skinned muscle fibres showing that oxidative treatment increased the effectiveness of voltage sensor activation of Ca2+ release when using submaximal conditions of depolarization. In agreement with their conclusions, less voltage sensor activation is indeed needed to open the release channels in the presence of H2O2, but our results indicate that this is also true when NO production is inhibited. The two phenomena may be mechanistically intimately linked, for instance if the presence of NO protects the channel against the effects of oxidation, as suggested by Aghdasi et al. (1997). Overall, it may also be speculated that changes in the voltage sensitivity of Ca2+ release channel activation may be a ubiquitous consequence of redox-related modifications of the ryanodine receptor protein, under physiological conditions.
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It should be noted that, under our conditions, the peak amplitude of the release was also substantially reduced in the presence of H2O2. Although we have no straightforward explanation for this, the strong oxidizing condition could have had additional, maybe deleterious, effects on the channel protein or on other components of the E–C coupling process (see also Posterino et al. 2003).
In conclusion, the present data provide important information concerning the mechanism of regulation of the excitation–contraction coupling process by NO. Although the overall picture is still not entirely clear, this work definitely contributes to our understanding of how NO affects skeletal muscle function. It may also prove helpful for resolving issues concerning diseases related to dysfunction of the NO signalling pathway in this tissue.
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