Voltage-controlled Ca2+ release and entry flux in isolated adult muscle fibres of the mouse
1 Universitt Ulm, Abteilung für Angewandte Physiologie, Albert-Einstein-Allee 11, D-89069 Ulm, Germany
Abstract
The voltage-activated fluxes of Ca2+ from the sarcoplasmic reticulum (SR) and from the extracellular space were studied in skeletal muscle fibres of adult mice. Single fibres of the interosseus muscle were enzymatically isolated and voltage clamped using a two-electrode technique. The fibres were perfused from the current-passing micropipette with a solution containing 15 mM EGTA and 0.2 mM of either fura-2 or the faster, lower affinity indicator fura-FF. Electrical recordings in parallel with the fluorescence measurements allowed the estimation of intramembrane gating charge movements and transmembrane Ca2+ inward current exhibiting half-maximal activation at –7.60 ± 1.29 and 3.0 ± 1.44 mV, respectively. The rate of Ca2+ release from the SR was calculated after fitting the relaxation phases of fluorescence ratio signals with a kinetic model to quantify overall Ca2+ removal. Results obtained with the two indicators were similar. Ca2+ release was 2–3 orders of magnitude larger than the flux carried by the L-type Ca2+ current. At maximal depolarization (+50 mV), release flux peaked at about 3 ms after the onset of the voltage pulse and then decayed in two distinct phases. The slower phase, most likely resulting from SR depletion, indicated a decrease in lumenal Ca2+ content by about 80% within 100 ms. Unlike in frog fibres, the kinetics of the rapid phase of decay showed no dependence on the filling state of the SR and the results provide little evidence for a substantial increase of SR permeability on depletion. The approach described here promises insight into excitation–contraction coupling in future studies of genetically altered mice.
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Introduction
Mechanical force in muscle is controlled by rapid changes in the myoplasmic calcium concentration that result from Ca2+ release and uptake mechanisms of the sarcoplasmic reticulum (SR) (for overviews see for instance Rüegg, 1988; Melzer et al. 1995; Bers, 2001). The release of Ca2+ ions stored in the SR is activated by a depolarization of the transverse tubular membrane (Ríos & Brum, 2002). The fundamentals of Ca2+ release and its voltage control have been elucidated in voltage clamp experiments mainly performed on isolated frog muscle fibres (see summaries by Ríos & Pizarro, 1991; Schneider, 1994; Melzer et al. 1995). Analysis of the Ca2+ signals recorded with optical indicator dyes revealed a characteristic time course of the global efflux of calcium from the SR during a step depolarization. An early large but transient flux component (peak), attributed to Ca2+-induced activation and inactivation (Melzer et al. 1987; Schneider & Simon, 1988; Csernoch et al. 1993), could be distinguished from a rather persistent plateau component of Ca2+ release flux. The latter showed a slow decline attributed to progressive store depletion (Schneider et al. 1987).
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In recent years, cellular studies on muscle excitation–contraction (EC) coupling shifted their focus from amphibian preparations to the experimentally more challenging mammalian muscle cells. Of central importance are muscle cells of mice with genetic alterations of EC coupling proteins. In particular, certain null-mutant mice proved to be extremely useful. Much experimental work focused on myotubes of such mutants for studies on the molecular physiology of EC coupling (e.g. Beam et al. 1986; Tanabe et al. 1988; Adams & Beam, 1990; Nakai et al. 1996; Powell et al. 1996; Strube et al. 1996; Beurg et al. 1997; Dietze et al. 1998; Protasi et al. 1998, 2000; Ursu et al. 2001).
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On the other hand, only a few studies are available that investigated Ca2+ currents or Ca2+ signals under voltage clamp control in mature mouse muscle fibres (Jacquemond, 1997; Friedrich et al. 1999, 2004; Wang et al. 1999; Szentesi et al. 2001; Collet & Jacquemond, 2002). Ca2+ release flux properties have not yet been assessed in voltage-clamped mouse fibres.
In the present investigation, we measured global Ca2+ signals and Ca2+ currents during step depolarizations in isolated adult mouse fibres voltage clamped with a two-electrode technique and loaded with high concentrations of EGTA. The EGTA buffering improved the stability of the cells while still permitting the detection of clean Ca2+ signals with fluorescent indicators. To determine the input flux of Ca2+ underlying the measured Ca2+ signals, we used a general approach applied previously to EGTA-loaded cut muscle fibres of frog and rat (Gonzalez & Ríos, 1993; Shirokova et al. 1996) and mouse myotubes (Schuhmeier & Melzer, 2004). By combining these methods and using two ratiometric indicator dyes of different affinity, we achieved a quantification of the fluxes of both Ca2+ release and Ca2+ entry in mouse muscle fibres. The experiments also provided the first information on Ca2+ release properties in this preparation under conditions of substantial SR depletion and allowed the estimation of fractional changes of the SR Ca2+ content during step depolarization.
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Some of the results have been presented previously as an abstract (Ursu et al. 2004).
Methods
Preparation
129SvJ mice were bred and kept at the Animal Research Centre of the University of Ulm. The age of the specimens used for experiments varied between 16 and 43 weeks. Animals were killed in accordance with the guidelines of the local animal care committee (exposure to a rising concentration of CO2 followed by cervical dislocation).
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Interosseus muscles were dissected in Krebs-Ringer solution and incubated in ‘dissociation solution’ at 37°C. After 60 min of gentle rotation (100 min–1) the dissociation solution was replaced with normal Krebs-Ringer solution. Isolated fibres were stored at 4°C in Krebs-Ringer solution during the day of the experiment. In some experiments fibres were used on the next day and found to be fully functional.
Solutions
The following solutions were used for the experiments (concentrations in mM):
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Krebs-Ringer solution for muscle dissection and fibre storage: 118 NaCl, 3.4 KCl, 0.8 MgSO4, 1.2 KH2PO4, 11.1 glucose, 25 NaHCO3, 2.5 CaCl2, pH 7.4.
Dissociation solution for muscle fibre isolation: Krebs-Ringer solution containing 2 mg ml–1 collagenase.
External (bathing) solution for voltage clamp experiments: 135 TEA-OH, 135 HCH3SO3, 2 MgCl2, 10 CaCl2, 5 4-aminopyridine (4-AP), 10 Hepes, 0.001 TTX, 5 glucose, 0.05 N-benzyl-p-toluene sulphonamide (BTS), pH 7.4.
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Internal (pipette) solution for intracellular perfusion: 145 CsOH, 135 aspartic acid, 0.75 Na2ATP, 4.25 MgATP (resulting in 1 mM free Mg2+), 15 EGTA, 1.5 CaCl2 (resulting in 20 nM free Ca2+), 10 Hepes, 0.2 fura-2 or fura-FF, 5 sodium creatine phosphate, pH 7.2.
Voltage clamp
The experiments were performed at room temperature (20–23°C) in external solution containing 50 μM of the myosin II ATPase inhibitor BTS to suppress contractions (Cheung et al. 2002; Shaw et al. 2003). To ensure reliable voltage control, a two-microelectrode technique was used in the present experiments (Friedrich et al. 1999). Fibres were voltage clamped using an Axoclamp 2B amplifier (Axon Instruments, Union City, USA). The voltage recording electrodes were filled with 3 M KCl and had resistances between 4.8 and 7.5 M when immersed in external solution. The current-passing electrodes were low resistance (2.5–3.8 M) suction pipettes as used in conventional whole-cell recordings and permitted diffusional exchange of the intracellular constituents with an artificial intracellular solution (Fig. 1A). The high concentration of EGTA in this pipette solution suppressed contraction and created optimal conditions for the quantification of Ca2+ release flux (Gonzalez & Ríos, 1993; Pape et al. 1995; Shirokova et al. 1996; Song et al. 1998; Schuhmeier & Melzer, 2004).
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A, fluorescence emission was recorded from enzymatically isolated muscle fibres with a photomultiplier tube attached to the output port of an inverted fluorescence microscope. P1, voltage-recording intracellular micropipette filled with 3 M KCl; P2, current-passing electrode filled with internal solution containing the indicator dye; R, reference electrode; L, excitation light source; FC, filter changer; S, shutter; ExF, excitation filters; BS, beam splitter; EmF, emission filter; PMT, photomultiplier tube; A1, A2, voltage clamp amplifiers. B, loading of the fibre with fura-2 recorded by measuring the resting fluorescence F at 360 and 380 nm excitation (see Methods). Two fluorescence data points measured before start of perfusion (arrow) are outside the displayed voltage range. C, ratios of F380 and F360 after subtracting the extrapolated background fluorescence at time zero (dotted lines in B) from all measurement points, resulting in a mean free Ca2+ concentration of 46 nM.D, example of voltage clamp pulse (0 mV, 100 ms), L-type Ca2+ inward current and fura-2 fluorescence ratio signal (F380/F360). E, voltage dependence of activation of Ca2+ conductance () and Ca2+ signal () using more measurements of the experiment in D. Fit parameter values for description of current–voltage data (maximal Ca2+ conductance, reversal potential, voltage of half-maximal activation and steepness factor; for equations see Schuhmeier & Melzer, 2004): gCa,max= 186 S F–1, VCa= 73.2 mV, V1/2= 6.8 mV, k= 5.2 mV; fit parameters for the fluorescence data: V1/2=–15.2 mV, k= 5.5 mV. The interval between two consecutive recordings was 60 s.
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Fibres were measured at slack length (sarcomere length of about 2 μm as determined with a confocal microscope). Length and width of the fibres were measured by microscopic inspection using an eye-piece microscale. In 16 fibres, width and length were determined to be 58.8 ± 2.8 μm and 624 ± 12 μm, respectively. The experimental solutions strongly reduce all major ion conductance pathways. Quantitative considerations by Friedrich et al. (1999) using the same preparation under similar experimental conditions indicate essentially uniform polarization. The additional intracellular perfusion of our fibres with solutions containing impermeant ions provides even better conditions for space clamping.
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To determine the voltage dependence of Ca2+ release and Ca2+ entry, 100 ms depolarizing voltage steps with increasingly larger amplitude were applied. The interval between pulses was 60 s. The series of pulses was bracketed by voltage steps to +20 mV to assess any changes in reproducibility. To reduce the amplitude of the capacitive current transients, the command voltage was rounded by low-pass filtering at 500 Hz using an 8-pole Bessel filter (Geitmann, Menden, Germany). Capacitive transient and leak current were electrically compensated using subtraction of a signal generated by a transient generator with two time constants.
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Data acquisition
Electrophysiological data (current, voltage) and fluorescence were recorded simultaneously at 2 kHz sampling frequency using a CED 1401+ interface (Cambridge Electronic Design, Cambridge, UK) connected to a AMD K6-2 computer. For data acquisition, software written in Delphi 3 (Borland International, Scotts Valley, USA) was used. Macro routines implemented in Excel (Microsoft) and Delphi programs were used for data analysis.
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Ca2+ recording with fura-2 and fura-FF
The cells were loaded with the indicator dye by diffusion of the artificial internal solution from the current-passing electrode as described in Results. Fluorescence was detected using a photomultiplier system (PMT R268, Hamamatsu, Herrsching, Germany) attached to the bottom (Keller) output of the inverted epifluorescence microscope (Axiovert 135 TV, Zeiss, Oberkochen, Germany). Fluorescence emission recorded from the fibre section in the field of view was filtered with a 510 nm bandpass interference filter (510W B40, Omega Optical, Brattleboro, VT, USA). A fast electromagnetic shutter (VS 25, Uniblitz, Rochester, USA) was used to control irradiation from the xenon arc source (XBO, 75 W, Zeiss). The shutter was opened during a 1.5 s interval for each measurement. A home-made electromagnetic filter changer was used for alternating irradiation near 380 nm (Ca2+ signals) and 360 nm (isosbestic point) using interference filters 380.1/14.8 and 358.2/9.2, respectively (Schott, Mainz, Germany). We refer to the corresponding fluorescence emission intensities as ‘F380’ and ‘F360’. A beam splitter (FT460, Zeiss) directed the excitation light to the microscope objective (40x/0.75 W, Zeiss). Because of almost identical spectral properties of fura-2 and fura-FF (also named fura-2FF) (Hyrc et al. 2000), the same experimental arrangement could be used for both dyes.
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In vitro and in vivo calibration
The relation between the fluorescence ratio R (=F380/F360) and [Ca2+] is described by eqn (1) (Klein et al. 1988).
Rmin and Rmax values used for the analysis of fura-2 records were determined in quartz capillaries of 50 μm internal diameter (Dynamics Inc., Rockaway, USA) filled with modified internal solutions (15 mM EGTA, 200 μM fura-2) that contained either no added Ca2+ (for Rmin) or were buffered to 100 μM free Ca2+ (for Rmax). The values were 3.53 and 0.41, respectively.
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We also performed an in vivo estimate of Rmax in three muscle fibres perfused with a high-Ca2+ (5 mM), low-EGTA (0.1 mM) solution and stimulated by 100 ms depolarizations to +20 mV to ensure full dye saturation. Because the resultant value Rmax= 0.45 ± 0.05 was not significantly different from the in vitro value, we used the latter for calculations.
The in vivo value of the indicator dissociation rate constant koff,Dye which is essential for the correct deconvolution of fluorescence signals to calculate free [Ca2+] (eqn (1)) was determined in the removal model fit as described below. For KDye we used the value of 276 nM determined by Schuhmeier et al. (2003) with internal solutions of different [Ca2+]. KDye may be higher in the myoplasm (Konishi et al. 1988). However, as described in Results, both koff,Dye (and therefore Ca2+ dynamics) and, most importantly, the Ca2+ flux determination are independent of the choice of KDye when using the removal model fit method (see also Fig. 2 and Schuhmeier & Melzer, 2004).
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Fluorescence ratio signals (B) and calculated free Ca2+ signals (C) obtained with a train of four 50 ms pulses to 0 mV (A) separated by 150 ms intervals. Superimposed on the relaxation intervals after pulse repolarization in B are calculated ratio signals (thick lines) using the model of Schuhmeier & Melzer (2004). For best-fit parameters to calculate the model traces see list under E. D–E, calculation of Ca2+ input flux for a single step of voltage to 0 mV (D). Same experiment as in Fig. 1D and E. E, free Ca2+ concentration calculated for two different KDye values: 276 nM (continuous trace) and 1000 nM (dashed trace). Best-fit parameter values resulting from the two assumptions: koff,Dye= 33.9 s–1, kon,S= 18.5 μM–1 s–1, koff,S= 3.38 s–1, kuptake= 6591 s–1 and koff,Dye= 33.9 s–1, kon,S= 5.12 μM–1 s–1, koff,S= 3.38 s–1, kuptake= 1820 s–1, respectively. Initial values for fits in this and other experiments: koff,Dye= 30 s–1, kon,S= 1.5 μM–1 s–1, koff,S= 0.3 s–1 and kuptake= 1000 s–1 (Smith et al. 1984; Dietze et al. 1998; Schuhmeier & Melzer, 2004). Boundaries set for the parameters in the fitting algorithm were: 103 s–1, 104μM–1 s–1, 103 s–1 and 105 s–1, respectively. Only results that stayed within these limits without touching the boundaries for all four parameters were used here and elsewhere in this investigation. Fixed parameter values of the model used for the fit here and in all other fura-2 analyses: Rmin= 3.53, Rmax= 0.41, [Dye]total= 0.2 mM, [S]total= 15 mM and KDye=koff,Dye/kon,Dye= 0.276 μM (for equations see Schuhmeier & Melzer, 2004). F, calculated Ca2+ input fluxes for the two different assumptions in E leading to identical results. Note: flux amplitudes were scaled by 0.4 to account for smaller [Dye]total and [S]total in the cell (see Methods and Results).
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F360 recordings in the muscle fibres were used for approximate determinations of the intracellular fura-2 concentrations by comparison with F360 recordings in capillaries as described by Klein et al. (1988). Capillaries of two different inner diameters (50 and 100 μm) were used containing 50, 100 and 200 μM of dye in the internal solution.
For the low-affinity indicator fura-FF (concentration 200 μM), dye saturation in vivo could not be achieved without destroying the fibre. Therefore, Rmax was determined in 50 μm microcapillaries using a modified internal solution with high [Ca2+]. Similar values were determined with either 5 mM free Ca2+ and 0.1 mM EGTA (Rmax= 1.66) or 0.1 mM free Ca2+ and 15 mM EGTA (Rmax= 1.50). The latter value was used for the analysis of experimental recordings. Because of the low affinity of the indicator, Rmin was set to Rbaseline in experiments (the mean R value in the baseline before each pulse). A KD value of 6.5 μM was determined in microcapillaries using a commercial calibration kit (kit no. 3, Molecular Probes) and a fura-FF concentration of 10 μM. The pH was 7.2 in all calibration experiments.
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Ca2+ input flux analysis
Background and bleaching corrections were performed as described by Schuhmeier et al. (2003). Free Ca2+ concentration was calculated from voltage-activated changes of R using eqn (1). Ca2+ input flux, i.e. the total flux of Ca2+ into the myoplasm, was derived as described by Schuhmeier & Melzer (2004). Briefly, the relaxation of fluorescence ratio traces obtained in the intervals between repetitive voltage pulses were fitted with a kinetic model for the distribution of released Ca2+ to different compartments (see Fig. 2B). The fit was always started 8 ms after the end of the depolarization to account for the time course of release turn-off. The model fit served to quantify overall myoplasmic Ca2+ removal (Melzer et al. 1986). The model consisted of the indicator dye described by Rmin, Rmax, rate constants kon,Dye, koff,Dye and concentration [Dye]total, of a saturating buffer representing EGTA (parameters kon,S, koff,S and [S]total) and an uptake mechanism (rate constant kuptake). [Dye]total, [S]total and KDye=koff,Dye/kon,Dye were set to fixed values 0.2 mM, 15 mM and 0.276 μM (fura-2) or 6.5 μM (fura-FF), respectively. The fluorescence records during depolarizing pulses and the best-fit values of kinetic constants (koff,Dye, kon,S, koff,S and kuptake) in the removal model were then used to calculate the depolarization-induced Ca2+ flux into the myoplasmic water space. Flux traces calculated under these assumptions were subsequently scaled down by a factor of 0.4 taking into account the estimated mean fraction of cellular loading (see Results for further details). The voltage dependence of the Ca2+ input flux (peak and plateau) were best fitted with the product of a single Boltzmann function and a linear function (see legend of Fig. 4).
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A, calculated Ca2+ entry fluxes (using eqn (2)). B, Ca2+ input fluxes calculated using the removal model fit procedure. Note that the absolute scales in panels A and B differ by a factor of almost 200 (same fibre as in Fig. 1D and E). C, peak () and plateau component () of Ca2+ input flux plotted as functions of voltage. Mean values from 8 experiments are shown. , measurements at +20 mV 1 min before and after the series of different voltage pulses indicating a small run-down in flux. Data were fitted by the product of a single Boltzmann function (1/(1 + exp((V1/2–V)/k))) and a linear function (a+bV). The fit parameters V1/2, k, a and b had mean values of –10.27 ± 1.43 mV, 6.96 ± 0.24 mV, 129.51 ± 12.83 μM ms–1 and 885 ± 210 μM ms–1 V–1, respectively, for the peak and –14.23 ± 2.07 mV, 5.99 ± 0.22 mV, 22.46 ± 1.61 μM ms–1 and –81.1 ± 17.5 μM ms–1 V–1, respectively, for the plateau. The Ca2+ input flux calculations were based on a removal model with the following set of parameters: [fura-2]= 0.2 mM, [EGTA]= 15 mM, Rmin= 3.53, Rmax= 0.41, KDye= 276 nM, koff,Dye= 33.6 ± 2.9 s–1, kon,S= 24.5 ± 7.3 μM–1 s–1, koff,S= 4.32 ± 0.74 s–1, kuptake= 7.3 x 103± 1.6 x 103 s–1. The parameters koff,Dye, kon,S, koff,S and kuptake were optimized by the removal model fit procedure as described in Results. Fluxes were subsequently scaled by 0.4 to account for lower intracellular versus pipette concentrations (see Methods and Results). D, voltage dependence of Ca2+ entry flux derived from the leak-corrected Ca2+ inward current densities. Current data were fitted as described in Methods. The fit parameters gCa,max, VCa, V1/2 and k showed mean values of 112 ± 9 S F–1, 77.2 ± 5.45 mV, 3.0 ± 1.4 mV and 4.91 ± 0.24 mV, respectively. Current densities were scaled with the factor 22.5 μM s–1 A–1 F to obtain fluxes.
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Ca2+ current and Ca2+ entry flux analysis
Ca2+ currents were analysed as described by Schuhmeier & Melzer (2004). Unless otherwise stated, the last 8 ms of the current traces during voltage pulses of 100 ms duration were averaged for constructing current–voltage relations. Ca2+ entry flux (expressed in the same dimension as the input flux, i.e. as total concentration change in the myoplasmic water volume per time) was calculated from the measured Ca2+ current as described by Schuhmeier & Melzer (2004) using eqn (2):
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Here, iCa is the leak-corrected Ca2+ current density, z the valency of the Ca2+ ion, F the Faraday constant, VC the total intracellular volume per membrane capacitance and fV the fraction of the total volume that is immediately accessible to Ca2+.
VC was estimated in a number of fibres by using the measured membrane capacitance and an estimate of the total fibre volume derived from the fibre dimensions assuming cylindrical geometry. The mean value of VC was 0.32 ± 0.02 l F–1(n= 16) and for the same fibres the mean capacitance Cm was 5.73 ± 0.30 nF (ranging between 4.61 and 7.41 nF). The factor fV corrects the total intracellular volume for the fraction of the intracellular space that is occupied by organelles. The space made up by organelles (SR and mitochondria) has been estimated to be about 30% of the fibre volume in frog muscle (i.e. fV= 0.7; Baylor et al. 1983).
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Intramembrane charge movements
Linear capacitive current and leak current, elicited by 10 mV pulses (50 ms) from the holding potential of –80 mV, were compensated by subtracting the signal of an analog transient generator. Non-linear gating charge was determined at the onset of voltage pulses without using further control pulses by integrating the first 8 ms of the corrected current record. The current level measured immediately before the voltage pulse was used as baseline for the non-linear current determination. According to Wang et al. (1999) essentially all non-linear charge moves within this interval in mouse fibres at room temperature. Consistent with this, the estimated charge in the low voltage range did not increase when the integration interval was prolonged. Because of the much slower activation of the L-type current, a kinetic separation of gating current and inward ionic current was also possible at larger depolarizations in this interval. However activation of Ca2+ current made the charge estimates unreliable at longer integration intervals. The charge–voltage relations were fitted by the sum of a Boltzmann function and a linear function to account for any uncompensated linear capacitance. The best fit revealed only a very small linear component corresponding to a charge offset of 0.04% of the maximum of the Boltzmann component.
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Statistics
Unless otherwise stated, averaged data are presented and plotted as means ±S.E.M. (n= number of experiments). Student's two-sided t test was used to test for significant differences of mean values (assuming two independent populations; P= 0.05).
Results
Muscle fibre loading with internal solution
Immediately after inserting the micropipettes, a ‘loading protocol’ was started to observe the progress of intracellular equilibration by recording the change in resting fluorescence at the two different excitation wavelengths (360 and 380 nm). Simultaneously the bath solution was completely exchanged by perfusing the chamber for 2 min at a low rate (3 ml min–1) with fresh external solution (containing 50 μM BTS). This removed any fluorescent dye that had flowed out of the current-injecting electrode. Figure 1B shows the fluorescence at the two excitation wavelengths recorded during a time interval of 71 min. Intracellular diffusion started at time zero (arrow). Figure 1C shows the ratio R (see Methods) for the same time interval after subtracting the background fluorescence levels which are indicated by the dotted lines in Fig. 1B. As in the experiment shown, the resting ratio was usually quite constant over time, despite the clear changes in absolute intracellular concentrations, indicating that a steady state of resting Ca2+ was maintained from the very beginning of the experiments. Using the calibration parameters listed in Methods, a mean value for the initial basal free Ca2+ (baseline before the first pulse) of 58.6 ± 14.4 nM(n= 8) was determined.
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The ‘loading protocol’ was run for 30 min before depolarizing pulses were applied to elicit Ca2+ currents and Ca2+ release (Fig. 1D). Studying the increase of F360 over time to indicate the progress of intracellular perfusion, we found that even in long experiments (as in Fig. 1B) a final saturation could not be obtained. Comparing fluorescence levels in microcapillaries (see Methods and Klein et al. 1988) with the fluorescence recording at 360 nm excitation in seven muscle fibres led to a mean value of 82.6 ± 10.6 μM fura-2 at the time of measuring the voltage dependence (32–45 min after start of loading), corresponding to a fraction of 0.4 of the pipette concentration. Assuming comparable diffusion rates for EGTA and fura-2, the concentration of the chelator was 6 rather than 15 mM during this time. We used these estimates for the quantifications of flux amplitudes described below.
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Activation of Ca2+ current and intracellular Ca2+ signals
Figure 1D shows an example of a simultaneously recorded slow inward current and a fura-2 fluorescence ratio signal at a step depolarization to 0 mV of 100 ms duration. Figure 1E compares, using more measurements of the same experiment, the voltage dependence of Ca2+ conductance and the fractional change of the fluorescence ratio after evaluating the averages of the last 16 measurement points (8 ms) of the pulse during each trace. The conductance signal required 22 mV stronger depolarization for half-maximal activation than the fluorescence signal. The data were fitted with Boltzmann functions (continuous and dotted lines; for best-fit parameters see legend to Fig. 1).
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Calculation of Ca2+ input flux
To determine the time course of the flux of Ca2+ mobilization that causes the fluorescence signals we made use of a method originally described by Melzer et al. (1986, 1987), later adjusted by Gonzalez & Ríos (1993) for use with high intracellular EGTA concentrations and modified for experiments on myotubes by Schuhmeier & Melzer (2004). Under our conditions, the large concentration of EGTA exceeds that of the intrinsic buffers and a simple kinetic model can describe the time course of the Ca2+ transients (Schuhmeier & Melzer, 2004). We used the relaxation phases in a series of four identical pulses applied at relatively high frequency (interval 150 ms; Fig. 2A–C). Theoretical curves generated with the model were fitted simultaneously to a set of relaxation phases of the indicator signals when Ca2+ input flux was turned off by the repolarization. This leads to a reliable description of the overall Ca2+ removal properties for each experiment which could then be used to calculate the Ca2+ input flux from the given fluorescence ratio traces.
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Figure 2B shows the result after convergence of the fitting algorithm. The depolarization-induced changes of free Ca2+ concentration, calculated using the set of best-fit model parameters (Fig. 2C), differ from the fluorescence transients (Fig. 2B) by exhibiting a pronounced peak as the result of the derivative term in eqn (1). The equation shows that the scaling depends on KDye in the cell which may be considerably larger than in vitro (Konishi et al. 1988) but is not accessible to direct measurement. In Fig. 2E free Ca2+ transients are shown assuming two different values for KDye (276 and 1000 nM) in the model fit analysis. The time course of [Ca2+] was not altered (it is determined by koff,Dye) but the amplitude increased in proportion to KDye (see eqn (1)). However, the final result of the analysis, i.e. the Ca2+ input flux, showed both identical time course and identical amplitude. In Fig. 2F both calculated flux traces are superimposed.
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Stability during repetitive pulses
Our results demonstrate that voltage-controlled Ca2+ release flux during a 100 ms depolarization has similar kinetic characteristics in mouse twitch fibres to those originally found in frog muscle fibres (e.g. Melzer et al. 1987; Schneider et al. 1987). There, the initial peak is followed by a rapid decline caused by a fast inactivation process and a much slower decline resulting from a decrease in the SR Ca2+ content. In experiments to determine the voltage dependence of the Ca2+ release flux we therefore used intervals between individual pulses of 60 s to allow the fibre to recover from these changes. Figure 3 demonstrates the reproducibility of the depolarization responses under these conditions. It shows the measured Ca2+ inward current (A) and the calculated Ca2+ release flux (B) for 10 successive applications of a 100 ms voltage pulse to +20 mV.
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Rectangular +20 mV voltage steps of 100 ms duration were applied with a frequency of 1 min–1. A, Ca2+ inward current. B, calculated Ca2+ input flux. The individual responses show only very small differences. Compared with the first pulse, the inward current amplitude for the last pulse was 8% larger and the peak release flux 9% smaller indicating almost full recovery from inactivation and depletion within 1 min intervals.
Contribution of Ca2+ entry flux
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As shown in Fig. 1D, Ca2+ inward currents were measured simultaneously with the fluorescent indicator transients. The Ca2+ current density, i.e. L-type current per linear capacitance of the cell membrane, can be converted to Ca2+ entry flux in the same units as the optically determined Ca2+ input flux by using eqn (2) (see Methods and Schuhmeier et al. 2003). In Fig. 4, Ca2+ entry flux traces (A) are compared with the optically determined Ca2+ input fluxes (B).
, 百拇医药 Scaling of the ordinates in the two panels differs by a factor of almost 200. For the pulse with the largest current (+20 mV), the amplitude of the corresponding Ca2+ entry flux (in the middle of the record, i.e. 25–75 ms during the pulse), was calculated to be about 70 times smaller than the total Ca2+ input flux during the same time interval. For eight fibres the average amplitude ratio (25–75 ms) at +20 mV was 118.53 ± 13.01. This value will vary in proportion to the actual concentration of EGTA in the fibre. Even though this concentration is somewhat uncertain, it is safe to conclude that Ca2+ entry was much smaller than Ca2+ release in these experiments. In the following, we therefore do not make a strict distinction between ‘Ca2+ input’ and ‘Ca2+ release’ in the description of the experiments.
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Figure 4C and D shows the corresponding voltage dependence. The data points were fitted as described in Methods and the continuous curves were drawn by using the mean values of the best fit parameters of the individual experiments. The peak release flux in Fig. 4C did not approach a constant value at large voltages as predicted by a Boltzmann distribution. Instead, it gradually increased with voltage between +20 and +50 mV. This might have been due to a gradual change in release efficiency. This possibility could be ruled out because +20 mV depolarizations applied immediately before and after the test pulse series showed on average very similar responses (, smaller value obtained later) as the +20 mV pulse within the series. Thus, the quasi-linear increase in peak release flux at large voltages seems to be a true property of the voltage dependence of Ca2+ mobilization in mouse fibres.
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Effect of SR depletion
The plateau phase of Ca2+ release flux showed a slow decline whose equivalent in frog fibres was interpreted as the result of progressive depletion of Ca2+ in the SR (Schneider et al. 1987). Consistent with the depletion hypothesis, in our experiments the plateau showed a faster fractional decline at the larger depolarizations that caused larger release flux amplitudes. We therefore subjected the calculated flux records to an analysis procedure that corrects for the effect of putative store depletion to derive the time course of SR Ca2+ permeability (Schneider et al. 1987; Gonzalez & Ríos, 1993). Permeability was calculated as flux divided by Ca2+ content in the SR, both referred to the myoplasmic water volume. Permeability thus results in the dimension time–1 and is independent of the absolute amplitude determination of the release flux. The Ca2+ content is the difference between an initial Ca2+ content and the released amount. Using a modified version of the procedure described by Schneider et al. (1987) and Gonzalez & Ríos (1993) we determined the initial Ca2+ content in the SR, assuming permeability to be constant during the plateau phase (see Schuhmeier & Melzer, 2004).
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Figure 5A shows a release flux trace resulting from a 100 ms depolarization to +20 mV. Figure 5B displays the fractional SR content derived from the analysis for the same record. The analysis indicates that the voltage pulse released about 54% of the initial content. With [S]total= 6 mM (see Methods) the initial SR content (referred to myoplasmic water space) was estimated to be 5.2 mM. Figure 5C (continuous line) shows the calculated permeability change that represents the estimated whole-cell (global) gating kinetics of the release channels in the SR during the depolarization according to published methods (Schneider et al. 1987; Gonzalez & Ríos, 1993; Schuhmeier & Melzer, 2004). The total Ca2+ released during a voltage pulse may not exactly describe the loss of lumenal Ca2+ in the terminal SR cisternae because some of the released Ca2+ is recycled during the pulse. The true amount of recycled Ca2+ is uncertain but the removal model analysis provides an approximation using the component described by the rate constant kuptake. In Fig. 5B and C, fractional depletion and permeability, when taking into account the uptake rate, are indicated by the dotted traces. The correction led to rather similar results. Initial SR content was somewhat lower (4.9 mM) but the fraction of depletion and its time course were almost identical and the calculated permeability was slightly larger (7.3% higher amplitude).
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Ca2+ input flux (A) obtained at +20 mV was analysed as described in Methods and converted to permeability to correct for the influence of depletion in the SR assuming that the quasi-linear slope in the plateau component is caused by a decrease in the lumenal Ca2+ concentration. B, fractional decrease of SR Ca2+ content calculated for the assumption of absence (continuous line) and presence of recycling (superimposed dotted line). The calculated initial SR Ca2+ concentrations (100% values) were 5.2 and 4.9 mM, respectively. C, SR Ca2+ permeability obtained as the result of the depletion correction. D, voltage dependence of permeabilities derived from the data of Fig. 4C. Values of the fit parameters V1/2, k, a and b (see legend of Fig. 4) were –11.32 ± 1.75 mV, 6.94 ± 0.4 mV, 3.92 ± 0.55% ms–1 and 43.08 ± 6.35% ms–1 V–1, respectively, for the peak, and –10.08 ± 2.27 mV, 6.38 ± 0.45 mV, 1.28 ± 0.10% ms–1 and 0.99 ± 1.3% ms–1 V–1, respectively, for the plateau. Estimated SR contents at +10, +20, +30, +40 and +50 mV were 3.44 ± 0.32, 3.28 ± 0.28, 3.17 ± 0.25, 3.06 ± 0.23 and 2.97 ± 0.22 mM, respectively. E, comparison of voltage dependence of intramembrane charge movements (), peak release permeability (, dashed line) and Ca2+ conductance (), presented as fraction of the mean value obtained at +50 mV. Data from a subset of 6 experiments in which charge movements could be evaluated for a total of 8 fura-2 experiments. For charge movements, maximal value Qmax, V1/2 and k were 17.15 ± 2.51 nC μF–1, –7.6 ± 1.3 mV and 12.2 ± 0.7 mV, respectively. For release permeability, V1/2, k, a and b (see above) were –10.33 ± 2.10 mV, 7.24 ± 0.47 mV, 3.84 ± 0.74% ms–1 and 38.5 ± 7.5% ms–1 V–1, respectively. For conductance, gCa,max, V1/2 and k were 106 ± 9.7 S F–1, 2.15 ± 1.81 mV and 4.85 ± 0.32 mV, respectively.
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Voltage dependence of gating
Figure 5D shows mean peak and plateau of the Ca2+ permeability obtained in the set of eight experiments plotted versus pulse voltage. The determination of SR content failed at low voltages in half of the fibres because of noise in the small flux signals. However, all fibres showed reliably similar SR content estimates with small variance at +10 mV and larger. The mean values differed maximally by 14% (see Fig. 5 legend). Therefore, we used the value obtained at +10 mV for the calculations of permeability in the voltage range –60 to +10 mV that are shown in Fig. 5D. In Fig. 5E, the voltage dependence of peak Ca2+ permeability () is shown for a subset of six fibres of the experiments in panel D. It is compared with fractional activation of L-type Ca2+ conductance () obtained by fitting the current–voltage relations as described by Schuhmeier & Melzer (2004). In addition, gating charge movements were determined at the onset of the pulse where a kinetic separation from the slowly activating inward current was possible even at large depolarizations (see Methods). The relative position of the three activation curves obtained simultaneously from the same set of experiments seems consistent with sequential gating schemes in which Ca2+ conductance is activated subsequently to Ca2+ release by a single voltage-sensing process.
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Initial SR content and SR depletion
The determination of SR Ca2+ content gets less precise with smaller depolarizations because of the smaller size of the Ca2+ signals and the correspondingly smaller depletion effect. Nevertheless, in 4 of the 8 fibres the calculated SR content showed very similar values and little variance between –20 and +50 mV. This is demonstrated in Fig. 6A (), where the content estimated at each voltage is presented as normalized to the value obtained at +50 mV. For comparison, the mean fractional activation is shown for the same set of experiments in this panel (). Figure 6B presents the fractional decrease in SR content measured at the end of the 100 ms pulse at each voltage. Thus, the large pulses caused substantial depletion. Estimated for all eight fibres, SR content decreased to 25.5 ± 2.4, 22.3 ± 2.0, 21.2 ± 1.8, 20.4 ± 1.7 and 20.1 ± 1.8% of the initial value at +10, +20, +30, +40 and +50 mV, respectively.
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A, fractional SR Ca2+ content prior to depolarization () and fractional activation of Ca2+ release by the depolarization (), normalized to the values at +50 mV. Subset of four fura-2 experiments in which the depletion analysis could be carried out even at low voltages. B, estimated fractional SR content after 100 ms of depolarization at the different voltages. Same experiments as in A.
Figure 7A shows calculated Ca2+ input fluxes in a sequence of four 50 ms depolarizations to 0 mV separated by 150 ms intervals that was used for the removal fit analysis (Fig. 2). The measurement is from one of the experiments depicted in Fig. 6A and B. The amplitude of the flux decreased dramatically within the sequence. Inactivation and depletion are possible reasons for the decrease. Figure 7B compares the time course of the individual flux traces by superimposing them after normalization to the peak. It shows only a very small decrease in the ratio of peak to end level which is depicted in Fig. 7C as the average of seven experiments. If temporal summation of fast inactivation was the reason for the decline, one should expect (based on the results in frog fibres, Schneider & Simon, 1988) that the peak decreased relative to the plateau more than was actually observed. Thus, the reduced driving force due to SR depletion was suspected to be the main cause of the decline. Figure 7D–I investigates this possibility further. Figure 7D shows the result of applying the depletion correction, described in conjunction with Fig. 5, to each of the four pulses in the sequence individually. We used a 20 ms interval in the plateau, starting 30 ms after the onset of the pulse, for the correction procedure. The SR contents prior to each pulse determined in the analysis are plotted as open circles in Fig. 7E. The continuous line in Fig. 7E shows the predicted SR contents over the whole sequence by using just the correction result from the first pulse. The difference between predicted and determined value at the last pulse is relatively small, comprising about 10% of the initial value in the sequence.
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A, calculated Ca2+ release fluxes for four consecutive pulses (for protocol see Fig. 2). B, normalized time courses of the traces in A. C, ratio of peak to end level of Ca2+ release fluxes as mean values from seven experiments like the one in A. D, calculated permeability changes for the trace in A using depletion analysis for each pulse separately. E, fractional SR content derived for the trace in A based on the depletion analysis of the first pulse (continuous line) and based on each individual depletion analysis (). F, time constants of rapid inactivation of permeability for the four different traces in D. G, mean values of permeabilities of seven experiments (same analysis as in D, same experiments as in C). H, comparison of predicted (dark grey columns) and individually determined (light grey columns) SR contents as described in E, calculated for the seven experiments in G. I, mean time constants of rapid inactivation for the traces in G.
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The depletion-corrected fluxes (Fig. 7D) show a 30% decrease in peak amplitude (last versus first pulse) and a relative constancy of the plateau values. Figure 7G and H presents the result exemplified in Fig. 7D and E as an average from seven experiments. The mean peak amplitude showed a non-significant increase from 32.5 ± 4.3 to 40.7 ± 4.5% ms–1 (first to last pulse). For each pulse, predicted (dark grey columns) and estimated (light grey columns) SR contents show no significant differences (paired t test, P= 0.13). These results seem consistent with a major contribution of SR depletion to the amplitude changes shown in Fig. 7A. In frog fibres and skinned rat fibres (Posterino & Lamb, 2003), strong depletion has been suggested to lead to a compensatory increase in permeability (Pape & Carrier, 1998; Pizarro & Ríos, 2004). The results of Fig. 7G show indeed a slight increase in mean permeability during the pulses, but much less than reported for frog fibres (Pape & Carrier, 1998; Fenelon & Pape, 2002; Pizarro & Ríos, 2004). Fitting the decay after the peak with a single exponential function led to very similar time constants in the example of Fig. 7F and showed no significant differences between the mean time constants (Fig. 7I, paired, t test, P= 0.32). In contrast, frog fibres exhibited a marked increase in the inactivation time constant with increasing SR depletion (Pizarro & Ríos, 2004).
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Determination of Ca2+ input flux using the indicator fura-FF
The analysis of our fura-2 recordings led to Ca2+ release fluxes that showed pronounced peaks early during the depolarizing steps. Because of the high concentrations of EGTA in the intracellular solution the free Ca2+ transient must have a similar time course as the underlying Ca2+ flux (Gonzalez & Ríos, 1993; Song et al. 1998; Schuhmeier et al. 2003). However, because of the relatively slow kinetics of the indicator fura-2 which causes low-pass filtering of the free Ca2+ transients, no peak is seen in the original fluorescence records. Only the kinetic deconvolution (eqn (1)) reveals the phasic time course of the underlying Ca2+ signal (Fig. 2C and E, see also Struk et al. 1998). As an additional check for the validity of our procedure, we performed experiments with an indicator of lower affinity and faster kinetics. We used fura-FF for which considerably higher dissociation constants than for fura-2 have been reported: 6 μM (Hyrc et al. 2000), 13 μM (Weinberg et al. 1997) and 35 μM (Golovina & Blaustein, 1997). Our estimate in microcapillaries was 6.5 μM (see Methods).
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Figure 8A shows a series of fluorescence ratio recordings at different pulse voltages. As in the case of fura-2, fluorescence of fura-FF decreases on Ca2+ binding at 380 nm excitation and therefore the F380/F360 records show the time course of dye-bound Ca2+ in inverted display. Because of the lower Ca2+ affinity of the dye, the signal-to-noise ratio was lower than in the fura-2 records at equal pipette concentrations. However, because of the faster kinetics, the fura-FF records (Fig. 8A), unlike the fura-2 records (Fig. 1D), showed phasic components at the beginning of the pulses resulting from the initial peak of the Ca2+ mobilization rate.
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A, fura-FF fluorescence ratio recordings (F380/F360) obtained at four different voltages (–30, –10, +10 and +20 mV). As in the case of the fura-2 recordings (Fig. 1D) a decrease in the ratio indicates an increase in Ca2+ concentration. B, Ca2+ input flux obtained by removal model fitting as described in the text. Best-fit parameter values: koff,Dye= 170 s–1, kon,S= 7.3 μM–1 s–1, koff,S= 6.7 s–1 and kuptake= 3862 s–1. Fixed parameters here and in all other fura-FF analyses had the following values: Rmin= resting R (see Methods), Rmax= 1.5, [Dye]total= 0.2 mM, [S]total= 15 mM and KDye=koff,Dye/kon,Dye= 6.5 μM. As in fura-2 experiments, flux amplitudes were scaled by 0.4 to account for lower concentrations of [Dye]total and [S]total in the cell (see Methods and Results). C, peak () and plateau component () of Ca2+ input flux derived from six fura-FF experiments plotted as functions of voltage. Data were fitted as described in Fig. 4C. , measurements at +20 mV, 1 min before and after the series of test pulses indicating the degree of run-down in peak flux. The fit parameters V1/2, k, a and b had mean values of –8.03 ± 2.12 mV, 8.54 ± 0.53 mV, 212 ± 65.2 μM ms–1 and 0.12 ± 0.57 μM ms–1 V–1, respectively, for the peak, and –10.87 ± 2.18 mV, 7.48 ± 0.45 mV, 29.7 ± 4.04 μM ms–1 and 0.16 ± 0.04 μM ms–1 V–1, respectively, for the plateau. Estimated SR contents at +10, +20, +30, +40 and +50 mV were 7.77 ± 1.32, 5.98 ± 0.49, 6.46 ± 1.02, 5.6 ± 0.58 and 5.15 ± 0.51 mM, respectively. D, voltage dependence of the corresponding permeabilities derived from the data in C. Values of the fit parameters (see C) were –4.37 ± 4.07 mV, 9.92 ± 0.51 mV, 3.28 ± 0.77% ms–1 and 19.25 ± 12.09% ms–1 V–1, respectively, for the peak, and –7.99 ± 4.64 mV, 7.61 ± 0.68 mV, 0.678 ± 0.137% ms–1 and –0.64 ± 0.99% ms–1 V–1, respectively, for the plateau. The Ca2+ input flux calculations were based on a removal model with the following set of parameters: [fura-FF]= 0.2 mM, [EGTA]= 15 mM, Rmin= resting R, Rmax= 1.50, KDye= 6.5 μM, koff,Dye= 192 ± 30 s–1, kon,S= 11.0 ± 1.8 μM–1 s–1, koff,S= 4.94 ± 1.26 s–1, kuptake= 4.00 x 103± 0.77 x 103 s–1. The parameters koff,Dye, kon,S, koff,S and kuptake were optimized by the removal model fit procedure as described in Results.
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The Ca2+ input flux was determined with the same method as described for fura-2 records. Of the fixed parameters, Rmax and KDye were adjusted to the values determined in calibrations for fura-FF (see Methods and legend of Fig. 8). Because changes of fluorescence relative to the absolute fluorescence at 380 nm were about 5 times smaller for fura-FF than for fura-2, small residual movement artifacts apparently caused convergence problems in the removal model fit in 3 of the 6 fibres tested. For the rest of the cells the algorithm converged to good fits of the fluorescence relaxation phases. The mean values of koff,Dye, koff,S, kon,S and kuptake were 192.48 ± 29.52 s–1, 4.94 ± 1.26 s–1, 11 ± 1.83 μM–1 s–1 and 4003 ± 733 s–1, respectively (n= 3). Figure 8B shows the calculated Ca2+ release rates corresponding to the four ratio records (Fig. 8A) calculated as described before for the fura-2 experiments (Figs 2F and 4B).
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Voltage dependence of Ca2+ input flux derived from fura-FF transients
Figure 8C shows the voltage dependence of the mean values of peak and plateau Ca2+ release flux obtained in six fura-FF experiments calculated by using the best-fit parameter results of the three fibres for which the complete removal analysis could be carried out (see above). These parameters also led to acceptable descriptions of the relaxation phases of the ratio signals in the fibres that contained slight movement artifacts. In Fig. 8D, the corresponding voltage dependence of permeability (peak and plateau) is shown. As in the fura-2 experiments, we used the initial SR Ca2+ content values obtained at +10 mV for the permeability calculation at +10 mV and smaller depolarizations in each experiment. The open diamonds indicate the results of two bracketing +20 mV pulses (lower value obtained at the end of the activation protocol). They indicate a stronger run-down than in the fura-2 experiments. As in the fura-2 measurements, the run-down seems to be at least partly due to loss of Ca2+ from the SR because the +20 mV values are closer together after correction for depletion (Fig. 8D, diamonds). Because of the stronger run-down, the quasi-linear voltage dependence at large depolarizations seen in the fura-2 experiments does not show up in the flux–voltage relation but in the plot of peak permeability versus voltage (Fig. 8D).
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A comparison with the fura-2 results (Fig. 4) shows that the voltage range of activation and maximal amplitudes of Ca2+ input flux are quite similar. The voltage of half-maximal activation, V1/2, was –8.03 ± 2.12 mV (n= 6) for fura-FF compared to –10.17 ± 1.43 mV (n= 8) for fura-2. The maximal amplitudes determined at +50 mV differed by only 20%.
Time course of Ca2+ input flux
The calculated time course of the Ca2+ input flux in the fura-FF experiment was very similar to the one in the fura-2 experiments. Figure 9 shows the average of all records obtained at +20 mV normalized to the mean peak value for a comparison of the calculated Ca2+ input flux (A and E) and Ca2+ permeability traces (B and F) obtained with fura-2 and fura-FF, respectively. The thin lines indicate the point by point calculated S.E.M.s. The peak fluxes were 147.02 ± 15.54 μM ms–1 and 207.9 ± 64.37 μM ms–1. The peak permeabilities were 4.78 ± 0.62% ms–1 (fura-2, Fig. 9B) and 3.48 ± 0.93% ms–1 (fura-FF, Fig. 9F), respectively.
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Averaged traces of the calculated Ca2+ input flux at +20 mV depolarization obtained from 8 experiments performed with fura-2 as the indicator (A) and from 6 experiments performed with fura-FF (E). B and F, averaged traces of the depolarization-induced permeability changes derived from the data in A and E, respectively. Vertical scales were adjusted so that the sizes of the peaks match. Peak values were 147.01 ± 16.65 μM ms–1 (A), 207.93 ± 64.93 μM ms–1 (E), 4.78 ± 0.62% ms–1 (B) and 3.48 ± 0.93% ms–1 (F).Thin lines indicate S.E.M.C, voltage dependence of time-to-peak of input flux of the fura-2 experiments. The dashed line indicates delays caused by the Bessel filters used for command voltage rounding and signal smoothing. D, corresponding mean values of the peak-versus-plateau ratios of flux () and permeability (). Note that peak and plateau were both measured from the baseline level. G, voltage dependence of time-to-peak of input flux for the fura-FF experiments. H, corresponding values of the peak-versus-plateau ratios of flux () and permeability ().
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To compare the time course of Ca2+ release gating determined with the two indicators, we evaluated the half-time of decline after the peak from the permeability records. The mean values for the +20 mV pulses were 9.56 ± 0.42 ms for the fura-2 (n= 8) and 8.02 ± 0.44 ms for the fura-FF recordings (n= 6), respectively. Thus the fura-FF recordings showed slightly faster kinetics of the flux peak component.
Figure 9C shows the voltage dependence of the rise time from the beginning of the voltage pulse to the peak of the release flux (time to peak). It decreased with increasing voltage and reached a minimal value at +50 mV. The scale shows the time from the beginning of the command voltage pulse at the output of the DA converter to the peak. Because the command and the measured signal were both filtered by Bessel filters a fixed delay time estimated to be 2.7 ms has to be subtracted (dashed line). The estimated minimal time to peak after correction for the filter effects was 3.24 ± 0.14 ms for fura-2. Figure 9G shows the voltage dependence for the time to peak input flux for fura-FF. It approaches a very similar value at large depolarizations as in the measurements with fura-2. The mean value at +50 mV after correcting for filter delays was 3.13 ± 0.24 ms. Figure 9D presents the ratio of peak versus plateau for fura-2 for the voltage range –20 to +50 mV for both release flux () and permeability (). The correction for depletion reduced the peak/plateau ratio values and the steepness of their voltage dependence. Figure 9H shows the peak-versus-plateau ratios for fura-FF. When comparing the mean values with those of Fig. 9D, the range of values is quite similar. Except for voltages between –20 and +10 mV values were not significantly different.
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In summary, despite small differences, the calculation results derived with the two indicator dyes, which are the result of different calibrations and different degrees of kinetic deconvolution, are very similar.
Discussion
Removal model fit in voltage-clamped EGTA-loaded mouse muscle fibres
In this study we describe the determination of Ca2+ fluxes in voltage-clamped adult muscle fibres of the mouse. Our method combined the advantages of intracellular perfusion as in whole-cell patch clamp recordings (Wang et al. 1999) with the better voltage control of a two-electrode system (Friedrich et al. 1999). It requires less preparative effort than the silicon grease gap technique described by Jacquemond (1997). A similar approach has recently successfully been applied to flexor digitorum brevis fibres of normal and mdx mice (Woods et al. 2004a).
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The high intracellular EGTA, forming the dominating Ca2+ buffer (denoted S in our removal model), captures almost all the Ca2+ that is released during a depolarization with little alteration to the time course of the release flux (Gonzalez & Ríos, 1993; Pape et al. 1995; Ríos & Brum, 2002). The removal model fit algorithm that we used extracts from the fluorescence ratio records information on the time course of Ca2+ binding to S and determines its rate constants. It cannot, however, provide an unambiguous determination of kon,EGTA in the cell as long as the intracellular dissociation constant of the indicator is not known (Schuhmeier & Melzer, 2004). According to Konishi et al. (1988), it is likely that the dissociation constant for fura-2 and other indicators is higher inside muscle cells than in free solution. On the other hand, the rate constant koff,S is insensitive to the KDye value chosen (Schuhmeier & Melzer, 2004). In agreement with our value of 4.94 s–1 in this study, are several groups, who presented apparent ‘off ’ rate constants of EGTA to fit Ca2+ recordings in skeletal and cardiac myocytes obtained under conditions of millimolar intracellular EGTA. Reported values were between about 3 and 5 s–1 (Gonzalez & Ríos, 1993; Shirokova et al. 1996; Song et al. 1998; Schuhmeier & Melzer, 2004), i.e. about one order of magnitude higher than the values obtained from temperature jump (Naraghi, 1997) and stopped flow experiments (Smith et al. 1984) in free solution which were 0.5 s–1 and 0.3 s–1, respectively. The reason for this discrepancy is not clear. Gonzalez & Ríos (1993) attributed it to possible interactions of EGTA in the myoplasm, gradients of Ca2+ along the sarcomere or an effect of non-linear pumping and binding processes. Perhaps the possibility should also be considered that the in vitro experiments underestimated the true values in free solution.
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As we confirmed using simulated fluorescence traces of the kind shown in Schuhmeier et al. (2003), koff,Dye can reliably be determined by the removal fit procedure, thus providing an in vivo determination of the dynamic properties of the indicator. This parameter is essential for the correct temporal deconvolution of the fluorescence signals (eqn (1), Klein et al. 1988) and therefore for the determination of the time courses of [Ca2+] and Ca2+ input flux. The mean value of 34 s–1 obtained for the fura-2 dissociation rate constant in the present experiments is somewhat smaller but close to the value found in experiments on myotubes (45 s–1Schuhmeier et al. 2003; 46 s–1Schuhmeier & Melzer, 2004). It is 35–40% of the values reported for free solution (84 s–1, Jackson et al. 1987; 97 s–1, Kao & Tsien, 1988) in agreement with the data of Baylor & Hollingworth (1988) and Garcia & Schneider (1993), after correcting for the effect of temperature (Bakker et al. 1997). Consistent with the lower affinity and more rapid kinetics of Ca2+ binding, the removal model fit revealed a 5.7-fold higher off rate constant koff,Dye for fura-FF than for fura-2 (192 versus 34 s–1).
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Ca2+ input flux in mouse muscle fibres
Independent of the rate constants of individual model components determined with the fit algorithm, it is important to note that the final calculation result of the analysis, i.e. the Ca2+ input flux, proved to be insensitive to the choice of the KDye value (Schuhmeier & Melzer, 2004 and Fig. 2F. Consistent with its faster kinetics fura-FF showed faster fluorescence responses than fura-2 with a clear peak component at the beginning and faster decline at the end (Fig. 8A) but led to a very similar time course of Ca2+ input flux as derived from fura-2 records (Fig. 9) supporting the notion that the time course of the Ca2+ input flux is equally well determined by both indicators despite the stronger temporal deconvolution that is necessary for fura-2 records. Fura-FF fluorescence transients, however, showed a lower signal-to-noise ratio and corresponding problems in fitting the traces with the removal model in part of the cells. It was also more difficult to resolve the time course of slow changes after the end of the pulse and during the plateau phase which made the depletion correction less reliable. Thus, for use under high [EGTA] conditions and depending on the focus of the study, dyes with low Ca2+ affinity may not be the optimal choice.
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Even though the estimated input flux amplitude is virtually invariant to assumptions made for KDye in the myoplasmic environment as shown by Schuhmeier & Melzer (2004; see also Fig. 2F), it is proportional to the assumed value of [S]total. The effective [S]total value depends on the fractional loading of the intracellular space with the solution in the pipette. The estimated average value of 40% loading in the present experiments (which all followed the same timing scheme), obtained by comparison with fluorescence recordings from dye-filled microcapillaries, corresponds to a putative [S]total value of 6 mM. Despite some uncertainty in this value it seems clear from the results of Fig. 4 that the amplitude of the Ca2+ entry flux is many times smaller than the release flux and both time course and voltage dependence of Ca2+ input flux indicate that Ca2+ entry is of negligible contribution to the optical signal consistent with investigations on adult frog muscle fibres (Brum et al. 1987, 1988).
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Comparison with previous results on Ca2+ transients in mouse muscle fibres
Several groups have investigated action potential-induced Ca2+ transients in single mouse fibres (e.g. Hollingworth et al. 1996; Liu et al. 1997; Bakker et al. 1997; Westerblad et al. 1997; Tutdibi et al. 1999; Bruton et al. 2003; Baylor & Hollingworth, 2003; Woods et al. 2004b). Hollingworth et al. (1996) estimated Ca2+ release for single action potentials with the fast dye furaptra and reported peak rates of 140–150 μM ms–1. In a more recent paper the same group determined 212 μM ms–1 (Baylor & Hollingworth, 2003). These values are quite close to our peak Ca2+ input fluxes at +50 mV (166 μM ms–1 for fura-2 and 209 μM ms–1 for fura-FF measurements).
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Only a few studies are available investigating Ca2+ currents or Ca2+ signals in mouse fibres under voltage clamp conditions. Friedrich et al. (1999) measured Ca2+ inward currents in unperfused isolated toe muscle fibres bathed in a solution containing 2 mM Ca2+. The activation characteristics appear to be similar to those found by Wang et al. (1999) who likewise measured Ca2+ inward currents in the presence of 2 mM external Ca2+ using mouse flexor digitorum brevis (FDB) and the whole-cell patch clamp technique. This group reports values of maximal Ca2+ conductance (gCa,max) = 85 S F–1 and V1/2=–0.3 mV compared to gCa,max= 55 S F–1 and V1/2=–18.8 mV determined in a double vaseline gap system (Delbono et al. 1997). Using the silicone grease gap technique (Jacquemond, 1997) and 5 mM extracellular Ca2+, Szentesi et al. (2001) determined parameters gCa,max= 200 S F–1, VCa= 73 mV, V1/2=–10.8 mV, k= 7.1 mV, whereas in our experiments (10 mM external Ca2+) we found gCa,max= 112 S F–1, VCa= 77.6 mV, V1/2= 3 mV, k= 4.9 mV.
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Voltage control of Ca2+ release was studied in mouse fibres by Wang et al. (1999) and by Jacquemond (1997). These groups assessed Ca2+ release properties by investigating the amplitude of Ca2+ transients that were measured with different indicators (fluo-3, calcium green-5N or calcium orange-5N and indo-1, respectively). Wang et al. (1999) obtained half-maximal activation of their Ca2+ signals at +6.2 mV whereas the indo-1 signals of Collet et al. (1999) reached their half-maximal value between –30 and –20 mV. In a recent preliminary report Woods et al. (2004a) describe measurements using an experimental arrangement similar to ours with a high concentration of intracellular EGTA and using the Ca2+ indicator oregon green-488–BAPTA-5N (OGB-5N). The voltage of half-maximal activation was –42 mV, which is considerably more negative than in the other studies and in our fura-2 flux measurements (about –10 mV for peak flux). Apparently, parameters of the voltage dependence vary considerably between different studies. The variance is likely to be due to the different methods and solutions used and sets limits to a quantitative comparison of individual parameters among different studies.
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Comparison with input flux estimates in voltage-clamped rat fibres
Ca2+ release flux in voltage-clamped mouse muscle fibres has not yet been investigated by other groups but data are available for cut rat EDL fibres with low and high intracellular EGTA showing amplitudes of up to about 20 μM ms–1 (Garcia & Schneider, 1993; Delbono & Stefani, 1993; Shirokova et al. 1996). Csernoch et al. (1999a,b) used enzymatically isolated cut fibres and reported maximal peak permeabilities of 1.4% ms–1 which amounts to a flux of 25 μM ms–1 with their estimated mean SR content of 1.8 mM. Voltages for half-maximal activation of –4.6 and –17.2 mV were reported. The SR permeability of our mouse fibres was activated in a similar voltage range (V1/2=–11 mV in the fura-2 experiments and V1/2=–4.4 in the fura-FF experiments) but showed up to 4 times larger maximal peak permeabilities, i.e. about 6% ms–1 at +50 mV corresponding to 166.6 μM ms–1 for an estimated SR content of 3 mM. Thus, the isolated mouse fibres under the present experimental conditions appear to be more efficient in releasing Ca2+ from the SR than the previously studied cut rat fibres. Nevertheless, they showed a similar voltage dependence of peak-to-plateau ratio of permeability as reported for this preparation by Shirokova et al. (1996).
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When exploring voltage sensor charge movements and Ca2+ release flux at large depolarizations (between 0 and +60 mV) in frog fibres, Shirokova et al. (1995) made the observation that the amount of sensor charge was not constant but kept increasing in parallel with an increase in release permeability. In mammalian muscle, Ca2+ release flux at large depolarizations has not been investigated yet. In our experiments we observed an almost linear increase in peak input flux and permeability between +20 mV and +50 mV (Figs 4C, 5D and E, 6A and 8D). Only in the fura-FF peak flux–voltage dependence (Fig. 8C) is this behaviour not evident due to the larger run-down indicated by the bracketing +20 mV recordings. Our preliminary estimates of intramembrane charge movements revealed no second voltage-dependent component at large depolarizations (Fig. 5E). However, despite its slow activation, the Ca2+ conductance introduced some uncertainty into charge measurements in this voltage range and maximum charge density was lower than determined previously in a very similar preparation (Wang et al. 1999). This quasi-linear activation of Ca2+ release permeability at large depolarizations deserves further investigation. It may be related to the continuing decrease in the time to peak with increasing depolarization (Fig. 9C), possibly reflecting progressive synchronization of activation of the ryanodine receptors.
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Estimates of SR Ca2+ content
Following procedures introduced by Schneider et al. (1987) we corrected release flux records during a depolarizing voltage pulse for depletion in the SR and converted them to permeability records by scaling them with 1/(Ca0,SR–Flux x dt). The correction procedure led to values of the SR content before the pulse (Ca0,SR) and its fractional change during the pulse (see Fig. 5B). In our fura-2 experiments we estimated a mean value of about 3 mM of the SR Ca2+ content (referred to the myoplasmic water space and using the estimated 6 mM intracellular EGTA). For measurements done on frog fibres under similar conditions (high intracellular EGTA), and using similar calculations, Gonzalez & Ríos (1993) report Ca0,SR values between 0.7 and 4 mM. In a comparative study on frog and rat fibres Shirokova et al. (1996) obtained an average of 3.4 mM for frog and 1.9 mM for rat. Using a pH-metric method and 20 mM intracellular EGTA Pape et al. (1995) estimated the total release of Ca2+ in cut frog fibres caused by trains of 20–40 action potentials at 50 Hz that supposedly led to complete depletion of the SR. They obtained a mean value of 2.7 mM.Posterino & Lamb (2003) measuring force transients of skinned rat EDL fibres reported normal Ca2+ contents between 1.5 and 1.8 mM, very close to results determined in the same preparation with a different method (Fryer & Stephenson, 1996; Owen et al. 1997) and 4–5 mM in maximally loaded fibres. Thus the values of Ca0,SR estimated in our experiments in isolated mouse interosseus fibres are similar to various independent estimates in other vertebrate skeletal muscle cells. According to Baylor et al. (1983) myoplasmic water (exclusive SR and mitochondria) occupies 71% of the total fibre volume. The SR volume is about 10% of the fibre volume (Peachey, 1965; Mobley & Eisenberg, 1975). Thus the total Ca2+ concentration within the SR in our experiments is calculated to be 0.71 x 3 mM/0.1 = 21.3 mM.
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Is SR Ca2+ permeability in mouse fibres dependent on lumenal [Ca2+]
The determinations of SR content (Ca0,SR) are based on the assumption (Schneider et al. 1987) that permeability is constant during the time of the plateau of the Ca2+ release flux, i.e. that the slope measured during the plateau results exclusively from the decrease of the Ca2+ concentration in the SR. Experiments on voltage-clamped frog fibres (e.g. Pape & Carrier, 1998; Pizarro & Rios, 2004) indicated that permeability does not stay constant in the plateau of the release flux when fast inactivation is over. Instead it was reported to increase with progressive depletion, thus providing a feedback to stabilize release (see also Posterino & Lamb, 2003). In this model, the almost linear decline in flux during the time of the flux plateau is thought to be not only caused by a decrease in the Ca2+ gradient driving the flux but also to be modulated by a superimposed change in permeability that is not immediately evident from the flux time course. We determined that a 100 ms voltage pulse that maximally activates Ca2+ release leads to about 80% reduction in the initial SR content. This substantial decrease might cause a significant depletion-dependent permeability change which we ignore in our analysis.
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Systematic analysis of depletion and its putative effect on permeability under voltage clamp conditions has recently been performed on frog skeletal muscle fibres (Pizarro & Rios, 2004). A decrease of SR content to 25% was reported to cause an increase in plateau permeability by more than 2-fold. Other authors reported even larger changes (e.g. Pape & Carrier, 1998). With the present experiments, we cannot rule out an effect of SR content on SR permeability but it seems that in the mouse it is less substantial and possibly even absent. Consistent with the constant permeability assumption, we found that the estimated Ca0,SR values were very similar over a wide range of voltages that released different amounts of Ca2+ and generated different slopes in the flux plateau (see Fig. 6A). If the applied assumption was not correct, i.e. if the SR permeability was critically dependent on the lumenal concentration of Ca2+, we would have expected to calculate more strongly deviating values for Ca0,SR at different voltages. Furthermore, the analysis in Fig. 7 indicates strong depletion during the sequence of identical pulses but only a small increase in permeability, which may also have resulted from slightly underestimating the recycling of Ca2+ to the SR lumen. The data presented in Fig. 7B and C show other differences to the frog experiments. In particular, the peak of the flux did not decrease in relation to the plateau as reported for frog fibres and the decay from the peak to the plateau was not significantly slowed (Fig. 7B and C) as found in frog fibres (Pizarro & Rios, 2004).
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If the constant permeability assumption is not made, an exact and independent quantification of recycling and remnant Ca2+ in the SR becomes extremely important. When the lumenal SR content reaches low values, small errors in the estimated concentration will lead to large errors in the calculated permeability value. The SR content is usually deduced from the cytoplasmic Ca2+ signal or a signal related to it (e.g. pH changes) and includes certain assumptions about the Ca2+ recycling activity. A direct measurement of Ca2+ in the SR seems therefore necessary to obtain exact information on the permeability time course for each pulse and to verify a possible dependence of permeability on the filling state of the SR.
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Conclusion
In the present study we assessed previously unavailable properties of the SR calcium release mechanism in single voltage-clamped muscle fibres of the mouse. We investigated in detail voltage-controlled global Ca2+ fluxes originating from extracellular and intracellular sources. The results provide information on the time course and voltage dependence of the Ca2+ fluxes. Our study also supplies data on the loading state of the sarcoplasmic reticulum and its dynamic change during depolarizations of different strengths. It does not provide clear evidence for an effect of lumenal Ca2+ on SR permeability. The growing importance of mouse models with defined alterations in skeletal muscle proteins calls for adequate functional investigation. Thus the approach applied here will be of benefit for future investigations of genetically altered mice to study changes in excitation–contraction coupling and properties of storing and buffering Ca2+ in the SR.
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Abstract
The voltage-activated fluxes of Ca2+ from the sarcoplasmic reticulum (SR) and from the extracellular space were studied in skeletal muscle fibres of adult mice. Single fibres of the interosseus muscle were enzymatically isolated and voltage clamped using a two-electrode technique. The fibres were perfused from the current-passing micropipette with a solution containing 15 mM EGTA and 0.2 mM of either fura-2 or the faster, lower affinity indicator fura-FF. Electrical recordings in parallel with the fluorescence measurements allowed the estimation of intramembrane gating charge movements and transmembrane Ca2+ inward current exhibiting half-maximal activation at –7.60 ± 1.29 and 3.0 ± 1.44 mV, respectively. The rate of Ca2+ release from the SR was calculated after fitting the relaxation phases of fluorescence ratio signals with a kinetic model to quantify overall Ca2+ removal. Results obtained with the two indicators were similar. Ca2+ release was 2–3 orders of magnitude larger than the flux carried by the L-type Ca2+ current. At maximal depolarization (+50 mV), release flux peaked at about 3 ms after the onset of the voltage pulse and then decayed in two distinct phases. The slower phase, most likely resulting from SR depletion, indicated a decrease in lumenal Ca2+ content by about 80% within 100 ms. Unlike in frog fibres, the kinetics of the rapid phase of decay showed no dependence on the filling state of the SR and the results provide little evidence for a substantial increase of SR permeability on depletion. The approach described here promises insight into excitation–contraction coupling in future studies of genetically altered mice.
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Introduction
Mechanical force in muscle is controlled by rapid changes in the myoplasmic calcium concentration that result from Ca2+ release and uptake mechanisms of the sarcoplasmic reticulum (SR) (for overviews see for instance Rüegg, 1988; Melzer et al. 1995; Bers, 2001). The release of Ca2+ ions stored in the SR is activated by a depolarization of the transverse tubular membrane (Ríos & Brum, 2002). The fundamentals of Ca2+ release and its voltage control have been elucidated in voltage clamp experiments mainly performed on isolated frog muscle fibres (see summaries by Ríos & Pizarro, 1991; Schneider, 1994; Melzer et al. 1995). Analysis of the Ca2+ signals recorded with optical indicator dyes revealed a characteristic time course of the global efflux of calcium from the SR during a step depolarization. An early large but transient flux component (peak), attributed to Ca2+-induced activation and inactivation (Melzer et al. 1987; Schneider & Simon, 1988; Csernoch et al. 1993), could be distinguished from a rather persistent plateau component of Ca2+ release flux. The latter showed a slow decline attributed to progressive store depletion (Schneider et al. 1987).
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In recent years, cellular studies on muscle excitation–contraction (EC) coupling shifted their focus from amphibian preparations to the experimentally more challenging mammalian muscle cells. Of central importance are muscle cells of mice with genetic alterations of EC coupling proteins. In particular, certain null-mutant mice proved to be extremely useful. Much experimental work focused on myotubes of such mutants for studies on the molecular physiology of EC coupling (e.g. Beam et al. 1986; Tanabe et al. 1988; Adams & Beam, 1990; Nakai et al. 1996; Powell et al. 1996; Strube et al. 1996; Beurg et al. 1997; Dietze et al. 1998; Protasi et al. 1998, 2000; Ursu et al. 2001).
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On the other hand, only a few studies are available that investigated Ca2+ currents or Ca2+ signals under voltage clamp control in mature mouse muscle fibres (Jacquemond, 1997; Friedrich et al. 1999, 2004; Wang et al. 1999; Szentesi et al. 2001; Collet & Jacquemond, 2002). Ca2+ release flux properties have not yet been assessed in voltage-clamped mouse fibres.
In the present investigation, we measured global Ca2+ signals and Ca2+ currents during step depolarizations in isolated adult mouse fibres voltage clamped with a two-electrode technique and loaded with high concentrations of EGTA. The EGTA buffering improved the stability of the cells while still permitting the detection of clean Ca2+ signals with fluorescent indicators. To determine the input flux of Ca2+ underlying the measured Ca2+ signals, we used a general approach applied previously to EGTA-loaded cut muscle fibres of frog and rat (Gonzalez & Ríos, 1993; Shirokova et al. 1996) and mouse myotubes (Schuhmeier & Melzer, 2004). By combining these methods and using two ratiometric indicator dyes of different affinity, we achieved a quantification of the fluxes of both Ca2+ release and Ca2+ entry in mouse muscle fibres. The experiments also provided the first information on Ca2+ release properties in this preparation under conditions of substantial SR depletion and allowed the estimation of fractional changes of the SR Ca2+ content during step depolarization.
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Some of the results have been presented previously as an abstract (Ursu et al. 2004).
Methods
Preparation
129SvJ mice were bred and kept at the Animal Research Centre of the University of Ulm. The age of the specimens used for experiments varied between 16 and 43 weeks. Animals were killed in accordance with the guidelines of the local animal care committee (exposure to a rising concentration of CO2 followed by cervical dislocation).
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Interosseus muscles were dissected in Krebs-Ringer solution and incubated in ‘dissociation solution’ at 37°C. After 60 min of gentle rotation (100 min–1) the dissociation solution was replaced with normal Krebs-Ringer solution. Isolated fibres were stored at 4°C in Krebs-Ringer solution during the day of the experiment. In some experiments fibres were used on the next day and found to be fully functional.
Solutions
The following solutions were used for the experiments (concentrations in mM):
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Krebs-Ringer solution for muscle dissection and fibre storage: 118 NaCl, 3.4 KCl, 0.8 MgSO4, 1.2 KH2PO4, 11.1 glucose, 25 NaHCO3, 2.5 CaCl2, pH 7.4.
Dissociation solution for muscle fibre isolation: Krebs-Ringer solution containing 2 mg ml–1 collagenase.
External (bathing) solution for voltage clamp experiments: 135 TEA-OH, 135 HCH3SO3, 2 MgCl2, 10 CaCl2, 5 4-aminopyridine (4-AP), 10 Hepes, 0.001 TTX, 5 glucose, 0.05 N-benzyl-p-toluene sulphonamide (BTS), pH 7.4.
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Internal (pipette) solution for intracellular perfusion: 145 CsOH, 135 aspartic acid, 0.75 Na2ATP, 4.25 MgATP (resulting in 1 mM free Mg2+), 15 EGTA, 1.5 CaCl2 (resulting in 20 nM free Ca2+), 10 Hepes, 0.2 fura-2 or fura-FF, 5 sodium creatine phosphate, pH 7.2.
Voltage clamp
The experiments were performed at room temperature (20–23°C) in external solution containing 50 μM of the myosin II ATPase inhibitor BTS to suppress contractions (Cheung et al. 2002; Shaw et al. 2003). To ensure reliable voltage control, a two-microelectrode technique was used in the present experiments (Friedrich et al. 1999). Fibres were voltage clamped using an Axoclamp 2B amplifier (Axon Instruments, Union City, USA). The voltage recording electrodes were filled with 3 M KCl and had resistances between 4.8 and 7.5 M when immersed in external solution. The current-passing electrodes were low resistance (2.5–3.8 M) suction pipettes as used in conventional whole-cell recordings and permitted diffusional exchange of the intracellular constituents with an artificial intracellular solution (Fig. 1A). The high concentration of EGTA in this pipette solution suppressed contraction and created optimal conditions for the quantification of Ca2+ release flux (Gonzalez & Ríos, 1993; Pape et al. 1995; Shirokova et al. 1996; Song et al. 1998; Schuhmeier & Melzer, 2004).
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A, fluorescence emission was recorded from enzymatically isolated muscle fibres with a photomultiplier tube attached to the output port of an inverted fluorescence microscope. P1, voltage-recording intracellular micropipette filled with 3 M KCl; P2, current-passing electrode filled with internal solution containing the indicator dye; R, reference electrode; L, excitation light source; FC, filter changer; S, shutter; ExF, excitation filters; BS, beam splitter; EmF, emission filter; PMT, photomultiplier tube; A1, A2, voltage clamp amplifiers. B, loading of the fibre with fura-2 recorded by measuring the resting fluorescence F at 360 and 380 nm excitation (see Methods). Two fluorescence data points measured before start of perfusion (arrow) are outside the displayed voltage range. C, ratios of F380 and F360 after subtracting the extrapolated background fluorescence at time zero (dotted lines in B) from all measurement points, resulting in a mean free Ca2+ concentration of 46 nM.D, example of voltage clamp pulse (0 mV, 100 ms), L-type Ca2+ inward current and fura-2 fluorescence ratio signal (F380/F360). E, voltage dependence of activation of Ca2+ conductance () and Ca2+ signal () using more measurements of the experiment in D. Fit parameter values for description of current–voltage data (maximal Ca2+ conductance, reversal potential, voltage of half-maximal activation and steepness factor; for equations see Schuhmeier & Melzer, 2004): gCa,max= 186 S F–1, VCa= 73.2 mV, V1/2= 6.8 mV, k= 5.2 mV; fit parameters for the fluorescence data: V1/2=–15.2 mV, k= 5.5 mV. The interval between two consecutive recordings was 60 s.
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Fibres were measured at slack length (sarcomere length of about 2 μm as determined with a confocal microscope). Length and width of the fibres were measured by microscopic inspection using an eye-piece microscale. In 16 fibres, width and length were determined to be 58.8 ± 2.8 μm and 624 ± 12 μm, respectively. The experimental solutions strongly reduce all major ion conductance pathways. Quantitative considerations by Friedrich et al. (1999) using the same preparation under similar experimental conditions indicate essentially uniform polarization. The additional intracellular perfusion of our fibres with solutions containing impermeant ions provides even better conditions for space clamping.
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To determine the voltage dependence of Ca2+ release and Ca2+ entry, 100 ms depolarizing voltage steps with increasingly larger amplitude were applied. The interval between pulses was 60 s. The series of pulses was bracketed by voltage steps to +20 mV to assess any changes in reproducibility. To reduce the amplitude of the capacitive current transients, the command voltage was rounded by low-pass filtering at 500 Hz using an 8-pole Bessel filter (Geitmann, Menden, Germany). Capacitive transient and leak current were electrically compensated using subtraction of a signal generated by a transient generator with two time constants.
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Data acquisition
Electrophysiological data (current, voltage) and fluorescence were recorded simultaneously at 2 kHz sampling frequency using a CED 1401+ interface (Cambridge Electronic Design, Cambridge, UK) connected to a AMD K6-2 computer. For data acquisition, software written in Delphi 3 (Borland International, Scotts Valley, USA) was used. Macro routines implemented in Excel (Microsoft) and Delphi programs were used for data analysis.
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Ca2+ recording with fura-2 and fura-FF
The cells were loaded with the indicator dye by diffusion of the artificial internal solution from the current-passing electrode as described in Results. Fluorescence was detected using a photomultiplier system (PMT R268, Hamamatsu, Herrsching, Germany) attached to the bottom (Keller) output of the inverted epifluorescence microscope (Axiovert 135 TV, Zeiss, Oberkochen, Germany). Fluorescence emission recorded from the fibre section in the field of view was filtered with a 510 nm bandpass interference filter (510W B40, Omega Optical, Brattleboro, VT, USA). A fast electromagnetic shutter (VS 25, Uniblitz, Rochester, USA) was used to control irradiation from the xenon arc source (XBO, 75 W, Zeiss). The shutter was opened during a 1.5 s interval for each measurement. A home-made electromagnetic filter changer was used for alternating irradiation near 380 nm (Ca2+ signals) and 360 nm (isosbestic point) using interference filters 380.1/14.8 and 358.2/9.2, respectively (Schott, Mainz, Germany). We refer to the corresponding fluorescence emission intensities as ‘F380’ and ‘F360’. A beam splitter (FT460, Zeiss) directed the excitation light to the microscope objective (40x/0.75 W, Zeiss). Because of almost identical spectral properties of fura-2 and fura-FF (also named fura-2FF) (Hyrc et al. 2000), the same experimental arrangement could be used for both dyes.
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In vitro and in vivo calibration
The relation between the fluorescence ratio R (=F380/F360) and [Ca2+] is described by eqn (1) (Klein et al. 1988).
Rmin and Rmax values used for the analysis of fura-2 records were determined in quartz capillaries of 50 μm internal diameter (Dynamics Inc., Rockaway, USA) filled with modified internal solutions (15 mM EGTA, 200 μM fura-2) that contained either no added Ca2+ (for Rmin) or were buffered to 100 μM free Ca2+ (for Rmax). The values were 3.53 and 0.41, respectively.
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We also performed an in vivo estimate of Rmax in three muscle fibres perfused with a high-Ca2+ (5 mM), low-EGTA (0.1 mM) solution and stimulated by 100 ms depolarizations to +20 mV to ensure full dye saturation. Because the resultant value Rmax= 0.45 ± 0.05 was not significantly different from the in vitro value, we used the latter for calculations.
The in vivo value of the indicator dissociation rate constant koff,Dye which is essential for the correct deconvolution of fluorescence signals to calculate free [Ca2+] (eqn (1)) was determined in the removal model fit as described below. For KDye we used the value of 276 nM determined by Schuhmeier et al. (2003) with internal solutions of different [Ca2+]. KDye may be higher in the myoplasm (Konishi et al. 1988). However, as described in Results, both koff,Dye (and therefore Ca2+ dynamics) and, most importantly, the Ca2+ flux determination are independent of the choice of KDye when using the removal model fit method (see also Fig. 2 and Schuhmeier & Melzer, 2004).
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Fluorescence ratio signals (B) and calculated free Ca2+ signals (C) obtained with a train of four 50 ms pulses to 0 mV (A) separated by 150 ms intervals. Superimposed on the relaxation intervals after pulse repolarization in B are calculated ratio signals (thick lines) using the model of Schuhmeier & Melzer (2004). For best-fit parameters to calculate the model traces see list under E. D–E, calculation of Ca2+ input flux for a single step of voltage to 0 mV (D). Same experiment as in Fig. 1D and E. E, free Ca2+ concentration calculated for two different KDye values: 276 nM (continuous trace) and 1000 nM (dashed trace). Best-fit parameter values resulting from the two assumptions: koff,Dye= 33.9 s–1, kon,S= 18.5 μM–1 s–1, koff,S= 3.38 s–1, kuptake= 6591 s–1 and koff,Dye= 33.9 s–1, kon,S= 5.12 μM–1 s–1, koff,S= 3.38 s–1, kuptake= 1820 s–1, respectively. Initial values for fits in this and other experiments: koff,Dye= 30 s–1, kon,S= 1.5 μM–1 s–1, koff,S= 0.3 s–1 and kuptake= 1000 s–1 (Smith et al. 1984; Dietze et al. 1998; Schuhmeier & Melzer, 2004). Boundaries set for the parameters in the fitting algorithm were: 103 s–1, 104μM–1 s–1, 103 s–1 and 105 s–1, respectively. Only results that stayed within these limits without touching the boundaries for all four parameters were used here and elsewhere in this investigation. Fixed parameter values of the model used for the fit here and in all other fura-2 analyses: Rmin= 3.53, Rmax= 0.41, [Dye]total= 0.2 mM, [S]total= 15 mM and KDye=koff,Dye/kon,Dye= 0.276 μM (for equations see Schuhmeier & Melzer, 2004). F, calculated Ca2+ input fluxes for the two different assumptions in E leading to identical results. Note: flux amplitudes were scaled by 0.4 to account for smaller [Dye]total and [S]total in the cell (see Methods and Results).
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F360 recordings in the muscle fibres were used for approximate determinations of the intracellular fura-2 concentrations by comparison with F360 recordings in capillaries as described by Klein et al. (1988). Capillaries of two different inner diameters (50 and 100 μm) were used containing 50, 100 and 200 μM of dye in the internal solution.
For the low-affinity indicator fura-FF (concentration 200 μM), dye saturation in vivo could not be achieved without destroying the fibre. Therefore, Rmax was determined in 50 μm microcapillaries using a modified internal solution with high [Ca2+]. Similar values were determined with either 5 mM free Ca2+ and 0.1 mM EGTA (Rmax= 1.66) or 0.1 mM free Ca2+ and 15 mM EGTA (Rmax= 1.50). The latter value was used for the analysis of experimental recordings. Because of the low affinity of the indicator, Rmin was set to Rbaseline in experiments (the mean R value in the baseline before each pulse). A KD value of 6.5 μM was determined in microcapillaries using a commercial calibration kit (kit no. 3, Molecular Probes) and a fura-FF concentration of 10 μM. The pH was 7.2 in all calibration experiments.
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Ca2+ input flux analysis
Background and bleaching corrections were performed as described by Schuhmeier et al. (2003). Free Ca2+ concentration was calculated from voltage-activated changes of R using eqn (1). Ca2+ input flux, i.e. the total flux of Ca2+ into the myoplasm, was derived as described by Schuhmeier & Melzer (2004). Briefly, the relaxation of fluorescence ratio traces obtained in the intervals between repetitive voltage pulses were fitted with a kinetic model for the distribution of released Ca2+ to different compartments (see Fig. 2B). The fit was always started 8 ms after the end of the depolarization to account for the time course of release turn-off. The model fit served to quantify overall myoplasmic Ca2+ removal (Melzer et al. 1986). The model consisted of the indicator dye described by Rmin, Rmax, rate constants kon,Dye, koff,Dye and concentration [Dye]total, of a saturating buffer representing EGTA (parameters kon,S, koff,S and [S]total) and an uptake mechanism (rate constant kuptake). [Dye]total, [S]total and KDye=koff,Dye/kon,Dye were set to fixed values 0.2 mM, 15 mM and 0.276 μM (fura-2) or 6.5 μM (fura-FF), respectively. The fluorescence records during depolarizing pulses and the best-fit values of kinetic constants (koff,Dye, kon,S, koff,S and kuptake) in the removal model were then used to calculate the depolarization-induced Ca2+ flux into the myoplasmic water space. Flux traces calculated under these assumptions were subsequently scaled down by a factor of 0.4 taking into account the estimated mean fraction of cellular loading (see Results for further details). The voltage dependence of the Ca2+ input flux (peak and plateau) were best fitted with the product of a single Boltzmann function and a linear function (see legend of Fig. 4).
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A, calculated Ca2+ entry fluxes (using eqn (2)). B, Ca2+ input fluxes calculated using the removal model fit procedure. Note that the absolute scales in panels A and B differ by a factor of almost 200 (same fibre as in Fig. 1D and E). C, peak () and plateau component () of Ca2+ input flux plotted as functions of voltage. Mean values from 8 experiments are shown. , measurements at +20 mV 1 min before and after the series of different voltage pulses indicating a small run-down in flux. Data were fitted by the product of a single Boltzmann function (1/(1 + exp((V1/2–V)/k))) and a linear function (a+bV). The fit parameters V1/2, k, a and b had mean values of –10.27 ± 1.43 mV, 6.96 ± 0.24 mV, 129.51 ± 12.83 μM ms–1 and 885 ± 210 μM ms–1 V–1, respectively, for the peak and –14.23 ± 2.07 mV, 5.99 ± 0.22 mV, 22.46 ± 1.61 μM ms–1 and –81.1 ± 17.5 μM ms–1 V–1, respectively, for the plateau. The Ca2+ input flux calculations were based on a removal model with the following set of parameters: [fura-2]= 0.2 mM, [EGTA]= 15 mM, Rmin= 3.53, Rmax= 0.41, KDye= 276 nM, koff,Dye= 33.6 ± 2.9 s–1, kon,S= 24.5 ± 7.3 μM–1 s–1, koff,S= 4.32 ± 0.74 s–1, kuptake= 7.3 x 103± 1.6 x 103 s–1. The parameters koff,Dye, kon,S, koff,S and kuptake were optimized by the removal model fit procedure as described in Results. Fluxes were subsequently scaled by 0.4 to account for lower intracellular versus pipette concentrations (see Methods and Results). D, voltage dependence of Ca2+ entry flux derived from the leak-corrected Ca2+ inward current densities. Current data were fitted as described in Methods. The fit parameters gCa,max, VCa, V1/2 and k showed mean values of 112 ± 9 S F–1, 77.2 ± 5.45 mV, 3.0 ± 1.4 mV and 4.91 ± 0.24 mV, respectively. Current densities were scaled with the factor 22.5 μM s–1 A–1 F to obtain fluxes.
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Ca2+ current and Ca2+ entry flux analysis
Ca2+ currents were analysed as described by Schuhmeier & Melzer (2004). Unless otherwise stated, the last 8 ms of the current traces during voltage pulses of 100 ms duration were averaged for constructing current–voltage relations. Ca2+ entry flux (expressed in the same dimension as the input flux, i.e. as total concentration change in the myoplasmic water volume per time) was calculated from the measured Ca2+ current as described by Schuhmeier & Melzer (2004) using eqn (2):
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Here, iCa is the leak-corrected Ca2+ current density, z the valency of the Ca2+ ion, F the Faraday constant, VC the total intracellular volume per membrane capacitance and fV the fraction of the total volume that is immediately accessible to Ca2+.
VC was estimated in a number of fibres by using the measured membrane capacitance and an estimate of the total fibre volume derived from the fibre dimensions assuming cylindrical geometry. The mean value of VC was 0.32 ± 0.02 l F–1(n= 16) and for the same fibres the mean capacitance Cm was 5.73 ± 0.30 nF (ranging between 4.61 and 7.41 nF). The factor fV corrects the total intracellular volume for the fraction of the intracellular space that is occupied by organelles. The space made up by organelles (SR and mitochondria) has been estimated to be about 30% of the fibre volume in frog muscle (i.e. fV= 0.7; Baylor et al. 1983).
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Intramembrane charge movements
Linear capacitive current and leak current, elicited by 10 mV pulses (50 ms) from the holding potential of –80 mV, were compensated by subtracting the signal of an analog transient generator. Non-linear gating charge was determined at the onset of voltage pulses without using further control pulses by integrating the first 8 ms of the corrected current record. The current level measured immediately before the voltage pulse was used as baseline for the non-linear current determination. According to Wang et al. (1999) essentially all non-linear charge moves within this interval in mouse fibres at room temperature. Consistent with this, the estimated charge in the low voltage range did not increase when the integration interval was prolonged. Because of the much slower activation of the L-type current, a kinetic separation of gating current and inward ionic current was also possible at larger depolarizations in this interval. However activation of Ca2+ current made the charge estimates unreliable at longer integration intervals. The charge–voltage relations were fitted by the sum of a Boltzmann function and a linear function to account for any uncompensated linear capacitance. The best fit revealed only a very small linear component corresponding to a charge offset of 0.04% of the maximum of the Boltzmann component.
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Statistics
Unless otherwise stated, averaged data are presented and plotted as means ±S.E.M. (n= number of experiments). Student's two-sided t test was used to test for significant differences of mean values (assuming two independent populations; P= 0.05).
Results
Muscle fibre loading with internal solution
Immediately after inserting the micropipettes, a ‘loading protocol’ was started to observe the progress of intracellular equilibration by recording the change in resting fluorescence at the two different excitation wavelengths (360 and 380 nm). Simultaneously the bath solution was completely exchanged by perfusing the chamber for 2 min at a low rate (3 ml min–1) with fresh external solution (containing 50 μM BTS). This removed any fluorescent dye that had flowed out of the current-injecting electrode. Figure 1B shows the fluorescence at the two excitation wavelengths recorded during a time interval of 71 min. Intracellular diffusion started at time zero (arrow). Figure 1C shows the ratio R (see Methods) for the same time interval after subtracting the background fluorescence levels which are indicated by the dotted lines in Fig. 1B. As in the experiment shown, the resting ratio was usually quite constant over time, despite the clear changes in absolute intracellular concentrations, indicating that a steady state of resting Ca2+ was maintained from the very beginning of the experiments. Using the calibration parameters listed in Methods, a mean value for the initial basal free Ca2+ (baseline before the first pulse) of 58.6 ± 14.4 nM(n= 8) was determined.
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The ‘loading protocol’ was run for 30 min before depolarizing pulses were applied to elicit Ca2+ currents and Ca2+ release (Fig. 1D). Studying the increase of F360 over time to indicate the progress of intracellular perfusion, we found that even in long experiments (as in Fig. 1B) a final saturation could not be obtained. Comparing fluorescence levels in microcapillaries (see Methods and Klein et al. 1988) with the fluorescence recording at 360 nm excitation in seven muscle fibres led to a mean value of 82.6 ± 10.6 μM fura-2 at the time of measuring the voltage dependence (32–45 min after start of loading), corresponding to a fraction of 0.4 of the pipette concentration. Assuming comparable diffusion rates for EGTA and fura-2, the concentration of the chelator was 6 rather than 15 mM during this time. We used these estimates for the quantifications of flux amplitudes described below.
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Activation of Ca2+ current and intracellular Ca2+ signals
Figure 1D shows an example of a simultaneously recorded slow inward current and a fura-2 fluorescence ratio signal at a step depolarization to 0 mV of 100 ms duration. Figure 1E compares, using more measurements of the same experiment, the voltage dependence of Ca2+ conductance and the fractional change of the fluorescence ratio after evaluating the averages of the last 16 measurement points (8 ms) of the pulse during each trace. The conductance signal required 22 mV stronger depolarization for half-maximal activation than the fluorescence signal. The data were fitted with Boltzmann functions (continuous and dotted lines; for best-fit parameters see legend to Fig. 1).
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Calculation of Ca2+ input flux
To determine the time course of the flux of Ca2+ mobilization that causes the fluorescence signals we made use of a method originally described by Melzer et al. (1986, 1987), later adjusted by Gonzalez & Ríos (1993) for use with high intracellular EGTA concentrations and modified for experiments on myotubes by Schuhmeier & Melzer (2004). Under our conditions, the large concentration of EGTA exceeds that of the intrinsic buffers and a simple kinetic model can describe the time course of the Ca2+ transients (Schuhmeier & Melzer, 2004). We used the relaxation phases in a series of four identical pulses applied at relatively high frequency (interval 150 ms; Fig. 2A–C). Theoretical curves generated with the model were fitted simultaneously to a set of relaxation phases of the indicator signals when Ca2+ input flux was turned off by the repolarization. This leads to a reliable description of the overall Ca2+ removal properties for each experiment which could then be used to calculate the Ca2+ input flux from the given fluorescence ratio traces.
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Figure 2B shows the result after convergence of the fitting algorithm. The depolarization-induced changes of free Ca2+ concentration, calculated using the set of best-fit model parameters (Fig. 2C), differ from the fluorescence transients (Fig. 2B) by exhibiting a pronounced peak as the result of the derivative term in eqn (1). The equation shows that the scaling depends on KDye in the cell which may be considerably larger than in vitro (Konishi et al. 1988) but is not accessible to direct measurement. In Fig. 2E free Ca2+ transients are shown assuming two different values for KDye (276 and 1000 nM) in the model fit analysis. The time course of [Ca2+] was not altered (it is determined by koff,Dye) but the amplitude increased in proportion to KDye (see eqn (1)). However, the final result of the analysis, i.e. the Ca2+ input flux, showed both identical time course and identical amplitude. In Fig. 2F both calculated flux traces are superimposed.
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Stability during repetitive pulses
Our results demonstrate that voltage-controlled Ca2+ release flux during a 100 ms depolarization has similar kinetic characteristics in mouse twitch fibres to those originally found in frog muscle fibres (e.g. Melzer et al. 1987; Schneider et al. 1987). There, the initial peak is followed by a rapid decline caused by a fast inactivation process and a much slower decline resulting from a decrease in the SR Ca2+ content. In experiments to determine the voltage dependence of the Ca2+ release flux we therefore used intervals between individual pulses of 60 s to allow the fibre to recover from these changes. Figure 3 demonstrates the reproducibility of the depolarization responses under these conditions. It shows the measured Ca2+ inward current (A) and the calculated Ca2+ release flux (B) for 10 successive applications of a 100 ms voltage pulse to +20 mV.
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Rectangular +20 mV voltage steps of 100 ms duration were applied with a frequency of 1 min–1. A, Ca2+ inward current. B, calculated Ca2+ input flux. The individual responses show only very small differences. Compared with the first pulse, the inward current amplitude for the last pulse was 8% larger and the peak release flux 9% smaller indicating almost full recovery from inactivation and depletion within 1 min intervals.
Contribution of Ca2+ entry flux
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As shown in Fig. 1D, Ca2+ inward currents were measured simultaneously with the fluorescent indicator transients. The Ca2+ current density, i.e. L-type current per linear capacitance of the cell membrane, can be converted to Ca2+ entry flux in the same units as the optically determined Ca2+ input flux by using eqn (2) (see Methods and Schuhmeier et al. 2003). In Fig. 4, Ca2+ entry flux traces (A) are compared with the optically determined Ca2+ input fluxes (B).
, 百拇医药 Scaling of the ordinates in the two panels differs by a factor of almost 200. For the pulse with the largest current (+20 mV), the amplitude of the corresponding Ca2+ entry flux (in the middle of the record, i.e. 25–75 ms during the pulse), was calculated to be about 70 times smaller than the total Ca2+ input flux during the same time interval. For eight fibres the average amplitude ratio (25–75 ms) at +20 mV was 118.53 ± 13.01. This value will vary in proportion to the actual concentration of EGTA in the fibre. Even though this concentration is somewhat uncertain, it is safe to conclude that Ca2+ entry was much smaller than Ca2+ release in these experiments. In the following, we therefore do not make a strict distinction between ‘Ca2+ input’ and ‘Ca2+ release’ in the description of the experiments.
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Figure 4C and D shows the corresponding voltage dependence. The data points were fitted as described in Methods and the continuous curves were drawn by using the mean values of the best fit parameters of the individual experiments. The peak release flux in Fig. 4C did not approach a constant value at large voltages as predicted by a Boltzmann distribution. Instead, it gradually increased with voltage between +20 and +50 mV. This might have been due to a gradual change in release efficiency. This possibility could be ruled out because +20 mV depolarizations applied immediately before and after the test pulse series showed on average very similar responses (, smaller value obtained later) as the +20 mV pulse within the series. Thus, the quasi-linear increase in peak release flux at large voltages seems to be a true property of the voltage dependence of Ca2+ mobilization in mouse fibres.
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Effect of SR depletion
The plateau phase of Ca2+ release flux showed a slow decline whose equivalent in frog fibres was interpreted as the result of progressive depletion of Ca2+ in the SR (Schneider et al. 1987). Consistent with the depletion hypothesis, in our experiments the plateau showed a faster fractional decline at the larger depolarizations that caused larger release flux amplitudes. We therefore subjected the calculated flux records to an analysis procedure that corrects for the effect of putative store depletion to derive the time course of SR Ca2+ permeability (Schneider et al. 1987; Gonzalez & Ríos, 1993). Permeability was calculated as flux divided by Ca2+ content in the SR, both referred to the myoplasmic water volume. Permeability thus results in the dimension time–1 and is independent of the absolute amplitude determination of the release flux. The Ca2+ content is the difference between an initial Ca2+ content and the released amount. Using a modified version of the procedure described by Schneider et al. (1987) and Gonzalez & Ríos (1993) we determined the initial Ca2+ content in the SR, assuming permeability to be constant during the plateau phase (see Schuhmeier & Melzer, 2004).
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Figure 5A shows a release flux trace resulting from a 100 ms depolarization to +20 mV. Figure 5B displays the fractional SR content derived from the analysis for the same record. The analysis indicates that the voltage pulse released about 54% of the initial content. With [S]total= 6 mM (see Methods) the initial SR content (referred to myoplasmic water space) was estimated to be 5.2 mM. Figure 5C (continuous line) shows the calculated permeability change that represents the estimated whole-cell (global) gating kinetics of the release channels in the SR during the depolarization according to published methods (Schneider et al. 1987; Gonzalez & Ríos, 1993; Schuhmeier & Melzer, 2004). The total Ca2+ released during a voltage pulse may not exactly describe the loss of lumenal Ca2+ in the terminal SR cisternae because some of the released Ca2+ is recycled during the pulse. The true amount of recycled Ca2+ is uncertain but the removal model analysis provides an approximation using the component described by the rate constant kuptake. In Fig. 5B and C, fractional depletion and permeability, when taking into account the uptake rate, are indicated by the dotted traces. The correction led to rather similar results. Initial SR content was somewhat lower (4.9 mM) but the fraction of depletion and its time course were almost identical and the calculated permeability was slightly larger (7.3% higher amplitude).
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Ca2+ input flux (A) obtained at +20 mV was analysed as described in Methods and converted to permeability to correct for the influence of depletion in the SR assuming that the quasi-linear slope in the plateau component is caused by a decrease in the lumenal Ca2+ concentration. B, fractional decrease of SR Ca2+ content calculated for the assumption of absence (continuous line) and presence of recycling (superimposed dotted line). The calculated initial SR Ca2+ concentrations (100% values) were 5.2 and 4.9 mM, respectively. C, SR Ca2+ permeability obtained as the result of the depletion correction. D, voltage dependence of permeabilities derived from the data of Fig. 4C. Values of the fit parameters V1/2, k, a and b (see legend of Fig. 4) were –11.32 ± 1.75 mV, 6.94 ± 0.4 mV, 3.92 ± 0.55% ms–1 and 43.08 ± 6.35% ms–1 V–1, respectively, for the peak, and –10.08 ± 2.27 mV, 6.38 ± 0.45 mV, 1.28 ± 0.10% ms–1 and 0.99 ± 1.3% ms–1 V–1, respectively, for the plateau. Estimated SR contents at +10, +20, +30, +40 and +50 mV were 3.44 ± 0.32, 3.28 ± 0.28, 3.17 ± 0.25, 3.06 ± 0.23 and 2.97 ± 0.22 mM, respectively. E, comparison of voltage dependence of intramembrane charge movements (), peak release permeability (, dashed line) and Ca2+ conductance (), presented as fraction of the mean value obtained at +50 mV. Data from a subset of 6 experiments in which charge movements could be evaluated for a total of 8 fura-2 experiments. For charge movements, maximal value Qmax, V1/2 and k were 17.15 ± 2.51 nC μF–1, –7.6 ± 1.3 mV and 12.2 ± 0.7 mV, respectively. For release permeability, V1/2, k, a and b (see above) were –10.33 ± 2.10 mV, 7.24 ± 0.47 mV, 3.84 ± 0.74% ms–1 and 38.5 ± 7.5% ms–1 V–1, respectively. For conductance, gCa,max, V1/2 and k were 106 ± 9.7 S F–1, 2.15 ± 1.81 mV and 4.85 ± 0.32 mV, respectively.
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Voltage dependence of gating
Figure 5D shows mean peak and plateau of the Ca2+ permeability obtained in the set of eight experiments plotted versus pulse voltage. The determination of SR content failed at low voltages in half of the fibres because of noise in the small flux signals. However, all fibres showed reliably similar SR content estimates with small variance at +10 mV and larger. The mean values differed maximally by 14% (see Fig. 5 legend). Therefore, we used the value obtained at +10 mV for the calculations of permeability in the voltage range –60 to +10 mV that are shown in Fig. 5D. In Fig. 5E, the voltage dependence of peak Ca2+ permeability () is shown for a subset of six fibres of the experiments in panel D. It is compared with fractional activation of L-type Ca2+ conductance () obtained by fitting the current–voltage relations as described by Schuhmeier & Melzer (2004). In addition, gating charge movements were determined at the onset of the pulse where a kinetic separation from the slowly activating inward current was possible even at large depolarizations (see Methods). The relative position of the three activation curves obtained simultaneously from the same set of experiments seems consistent with sequential gating schemes in which Ca2+ conductance is activated subsequently to Ca2+ release by a single voltage-sensing process.
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Initial SR content and SR depletion
The determination of SR Ca2+ content gets less precise with smaller depolarizations because of the smaller size of the Ca2+ signals and the correspondingly smaller depletion effect. Nevertheless, in 4 of the 8 fibres the calculated SR content showed very similar values and little variance between –20 and +50 mV. This is demonstrated in Fig. 6A (), where the content estimated at each voltage is presented as normalized to the value obtained at +50 mV. For comparison, the mean fractional activation is shown for the same set of experiments in this panel (). Figure 6B presents the fractional decrease in SR content measured at the end of the 100 ms pulse at each voltage. Thus, the large pulses caused substantial depletion. Estimated for all eight fibres, SR content decreased to 25.5 ± 2.4, 22.3 ± 2.0, 21.2 ± 1.8, 20.4 ± 1.7 and 20.1 ± 1.8% of the initial value at +10, +20, +30, +40 and +50 mV, respectively.
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A, fractional SR Ca2+ content prior to depolarization () and fractional activation of Ca2+ release by the depolarization (), normalized to the values at +50 mV. Subset of four fura-2 experiments in which the depletion analysis could be carried out even at low voltages. B, estimated fractional SR content after 100 ms of depolarization at the different voltages. Same experiments as in A.
Figure 7A shows calculated Ca2+ input fluxes in a sequence of four 50 ms depolarizations to 0 mV separated by 150 ms intervals that was used for the removal fit analysis (Fig. 2). The measurement is from one of the experiments depicted in Fig. 6A and B. The amplitude of the flux decreased dramatically within the sequence. Inactivation and depletion are possible reasons for the decrease. Figure 7B compares the time course of the individual flux traces by superimposing them after normalization to the peak. It shows only a very small decrease in the ratio of peak to end level which is depicted in Fig. 7C as the average of seven experiments. If temporal summation of fast inactivation was the reason for the decline, one should expect (based on the results in frog fibres, Schneider & Simon, 1988) that the peak decreased relative to the plateau more than was actually observed. Thus, the reduced driving force due to SR depletion was suspected to be the main cause of the decline. Figure 7D–I investigates this possibility further. Figure 7D shows the result of applying the depletion correction, described in conjunction with Fig. 5, to each of the four pulses in the sequence individually. We used a 20 ms interval in the plateau, starting 30 ms after the onset of the pulse, for the correction procedure. The SR contents prior to each pulse determined in the analysis are plotted as open circles in Fig. 7E. The continuous line in Fig. 7E shows the predicted SR contents over the whole sequence by using just the correction result from the first pulse. The difference between predicted and determined value at the last pulse is relatively small, comprising about 10% of the initial value in the sequence.
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A, calculated Ca2+ release fluxes for four consecutive pulses (for protocol see Fig. 2). B, normalized time courses of the traces in A. C, ratio of peak to end level of Ca2+ release fluxes as mean values from seven experiments like the one in A. D, calculated permeability changes for the trace in A using depletion analysis for each pulse separately. E, fractional SR content derived for the trace in A based on the depletion analysis of the first pulse (continuous line) and based on each individual depletion analysis (). F, time constants of rapid inactivation of permeability for the four different traces in D. G, mean values of permeabilities of seven experiments (same analysis as in D, same experiments as in C). H, comparison of predicted (dark grey columns) and individually determined (light grey columns) SR contents as described in E, calculated for the seven experiments in G. I, mean time constants of rapid inactivation for the traces in G.
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The depletion-corrected fluxes (Fig. 7D) show a 30% decrease in peak amplitude (last versus first pulse) and a relative constancy of the plateau values. Figure 7G and H presents the result exemplified in Fig. 7D and E as an average from seven experiments. The mean peak amplitude showed a non-significant increase from 32.5 ± 4.3 to 40.7 ± 4.5% ms–1 (first to last pulse). For each pulse, predicted (dark grey columns) and estimated (light grey columns) SR contents show no significant differences (paired t test, P= 0.13). These results seem consistent with a major contribution of SR depletion to the amplitude changes shown in Fig. 7A. In frog fibres and skinned rat fibres (Posterino & Lamb, 2003), strong depletion has been suggested to lead to a compensatory increase in permeability (Pape & Carrier, 1998; Pizarro & Ríos, 2004). The results of Fig. 7G show indeed a slight increase in mean permeability during the pulses, but much less than reported for frog fibres (Pape & Carrier, 1998; Fenelon & Pape, 2002; Pizarro & Ríos, 2004). Fitting the decay after the peak with a single exponential function led to very similar time constants in the example of Fig. 7F and showed no significant differences between the mean time constants (Fig. 7I, paired, t test, P= 0.32). In contrast, frog fibres exhibited a marked increase in the inactivation time constant with increasing SR depletion (Pizarro & Ríos, 2004).
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Determination of Ca2+ input flux using the indicator fura-FF
The analysis of our fura-2 recordings led to Ca2+ release fluxes that showed pronounced peaks early during the depolarizing steps. Because of the high concentrations of EGTA in the intracellular solution the free Ca2+ transient must have a similar time course as the underlying Ca2+ flux (Gonzalez & Ríos, 1993; Song et al. 1998; Schuhmeier et al. 2003). However, because of the relatively slow kinetics of the indicator fura-2 which causes low-pass filtering of the free Ca2+ transients, no peak is seen in the original fluorescence records. Only the kinetic deconvolution (eqn (1)) reveals the phasic time course of the underlying Ca2+ signal (Fig. 2C and E, see also Struk et al. 1998). As an additional check for the validity of our procedure, we performed experiments with an indicator of lower affinity and faster kinetics. We used fura-FF for which considerably higher dissociation constants than for fura-2 have been reported: 6 μM (Hyrc et al. 2000), 13 μM (Weinberg et al. 1997) and 35 μM (Golovina & Blaustein, 1997). Our estimate in microcapillaries was 6.5 μM (see Methods).
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Figure 8A shows a series of fluorescence ratio recordings at different pulse voltages. As in the case of fura-2, fluorescence of fura-FF decreases on Ca2+ binding at 380 nm excitation and therefore the F380/F360 records show the time course of dye-bound Ca2+ in inverted display. Because of the lower Ca2+ affinity of the dye, the signal-to-noise ratio was lower than in the fura-2 records at equal pipette concentrations. However, because of the faster kinetics, the fura-FF records (Fig. 8A), unlike the fura-2 records (Fig. 1D), showed phasic components at the beginning of the pulses resulting from the initial peak of the Ca2+ mobilization rate.
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A, fura-FF fluorescence ratio recordings (F380/F360) obtained at four different voltages (–30, –10, +10 and +20 mV). As in the case of the fura-2 recordings (Fig. 1D) a decrease in the ratio indicates an increase in Ca2+ concentration. B, Ca2+ input flux obtained by removal model fitting as described in the text. Best-fit parameter values: koff,Dye= 170 s–1, kon,S= 7.3 μM–1 s–1, koff,S= 6.7 s–1 and kuptake= 3862 s–1. Fixed parameters here and in all other fura-FF analyses had the following values: Rmin= resting R (see Methods), Rmax= 1.5, [Dye]total= 0.2 mM, [S]total= 15 mM and KDye=koff,Dye/kon,Dye= 6.5 μM. As in fura-2 experiments, flux amplitudes were scaled by 0.4 to account for lower concentrations of [Dye]total and [S]total in the cell (see Methods and Results). C, peak () and plateau component () of Ca2+ input flux derived from six fura-FF experiments plotted as functions of voltage. Data were fitted as described in Fig. 4C. , measurements at +20 mV, 1 min before and after the series of test pulses indicating the degree of run-down in peak flux. The fit parameters V1/2, k, a and b had mean values of –8.03 ± 2.12 mV, 8.54 ± 0.53 mV, 212 ± 65.2 μM ms–1 and 0.12 ± 0.57 μM ms–1 V–1, respectively, for the peak, and –10.87 ± 2.18 mV, 7.48 ± 0.45 mV, 29.7 ± 4.04 μM ms–1 and 0.16 ± 0.04 μM ms–1 V–1, respectively, for the plateau. Estimated SR contents at +10, +20, +30, +40 and +50 mV were 7.77 ± 1.32, 5.98 ± 0.49, 6.46 ± 1.02, 5.6 ± 0.58 and 5.15 ± 0.51 mM, respectively. D, voltage dependence of the corresponding permeabilities derived from the data in C. Values of the fit parameters (see C) were –4.37 ± 4.07 mV, 9.92 ± 0.51 mV, 3.28 ± 0.77% ms–1 and 19.25 ± 12.09% ms–1 V–1, respectively, for the peak, and –7.99 ± 4.64 mV, 7.61 ± 0.68 mV, 0.678 ± 0.137% ms–1 and –0.64 ± 0.99% ms–1 V–1, respectively, for the plateau. The Ca2+ input flux calculations were based on a removal model with the following set of parameters: [fura-FF]= 0.2 mM, [EGTA]= 15 mM, Rmin= resting R, Rmax= 1.50, KDye= 6.5 μM, koff,Dye= 192 ± 30 s–1, kon,S= 11.0 ± 1.8 μM–1 s–1, koff,S= 4.94 ± 1.26 s–1, kuptake= 4.00 x 103± 0.77 x 103 s–1. The parameters koff,Dye, kon,S, koff,S and kuptake were optimized by the removal model fit procedure as described in Results.
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The Ca2+ input flux was determined with the same method as described for fura-2 records. Of the fixed parameters, Rmax and KDye were adjusted to the values determined in calibrations for fura-FF (see Methods and legend of Fig. 8). Because changes of fluorescence relative to the absolute fluorescence at 380 nm were about 5 times smaller for fura-FF than for fura-2, small residual movement artifacts apparently caused convergence problems in the removal model fit in 3 of the 6 fibres tested. For the rest of the cells the algorithm converged to good fits of the fluorescence relaxation phases. The mean values of koff,Dye, koff,S, kon,S and kuptake were 192.48 ± 29.52 s–1, 4.94 ± 1.26 s–1, 11 ± 1.83 μM–1 s–1 and 4003 ± 733 s–1, respectively (n= 3). Figure 8B shows the calculated Ca2+ release rates corresponding to the four ratio records (Fig. 8A) calculated as described before for the fura-2 experiments (Figs 2F and 4B).
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Voltage dependence of Ca2+ input flux derived from fura-FF transients
Figure 8C shows the voltage dependence of the mean values of peak and plateau Ca2+ release flux obtained in six fura-FF experiments calculated by using the best-fit parameter results of the three fibres for which the complete removal analysis could be carried out (see above). These parameters also led to acceptable descriptions of the relaxation phases of the ratio signals in the fibres that contained slight movement artifacts. In Fig. 8D, the corresponding voltage dependence of permeability (peak and plateau) is shown. As in the fura-2 experiments, we used the initial SR Ca2+ content values obtained at +10 mV for the permeability calculation at +10 mV and smaller depolarizations in each experiment. The open diamonds indicate the results of two bracketing +20 mV pulses (lower value obtained at the end of the activation protocol). They indicate a stronger run-down than in the fura-2 experiments. As in the fura-2 measurements, the run-down seems to be at least partly due to loss of Ca2+ from the SR because the +20 mV values are closer together after correction for depletion (Fig. 8D, diamonds). Because of the stronger run-down, the quasi-linear voltage dependence at large depolarizations seen in the fura-2 experiments does not show up in the flux–voltage relation but in the plot of peak permeability versus voltage (Fig. 8D).
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A comparison with the fura-2 results (Fig. 4) shows that the voltage range of activation and maximal amplitudes of Ca2+ input flux are quite similar. The voltage of half-maximal activation, V1/2, was –8.03 ± 2.12 mV (n= 6) for fura-FF compared to –10.17 ± 1.43 mV (n= 8) for fura-2. The maximal amplitudes determined at +50 mV differed by only 20%.
Time course of Ca2+ input flux
The calculated time course of the Ca2+ input flux in the fura-FF experiment was very similar to the one in the fura-2 experiments. Figure 9 shows the average of all records obtained at +20 mV normalized to the mean peak value for a comparison of the calculated Ca2+ input flux (A and E) and Ca2+ permeability traces (B and F) obtained with fura-2 and fura-FF, respectively. The thin lines indicate the point by point calculated S.E.M.s. The peak fluxes were 147.02 ± 15.54 μM ms–1 and 207.9 ± 64.37 μM ms–1. The peak permeabilities were 4.78 ± 0.62% ms–1 (fura-2, Fig. 9B) and 3.48 ± 0.93% ms–1 (fura-FF, Fig. 9F), respectively.
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Averaged traces of the calculated Ca2+ input flux at +20 mV depolarization obtained from 8 experiments performed with fura-2 as the indicator (A) and from 6 experiments performed with fura-FF (E). B and F, averaged traces of the depolarization-induced permeability changes derived from the data in A and E, respectively. Vertical scales were adjusted so that the sizes of the peaks match. Peak values were 147.01 ± 16.65 μM ms–1 (A), 207.93 ± 64.93 μM ms–1 (E), 4.78 ± 0.62% ms–1 (B) and 3.48 ± 0.93% ms–1 (F).Thin lines indicate S.E.M.C, voltage dependence of time-to-peak of input flux of the fura-2 experiments. The dashed line indicates delays caused by the Bessel filters used for command voltage rounding and signal smoothing. D, corresponding mean values of the peak-versus-plateau ratios of flux () and permeability (). Note that peak and plateau were both measured from the baseline level. G, voltage dependence of time-to-peak of input flux for the fura-FF experiments. H, corresponding values of the peak-versus-plateau ratios of flux () and permeability ().
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To compare the time course of Ca2+ release gating determined with the two indicators, we evaluated the half-time of decline after the peak from the permeability records. The mean values for the +20 mV pulses were 9.56 ± 0.42 ms for the fura-2 (n= 8) and 8.02 ± 0.44 ms for the fura-FF recordings (n= 6), respectively. Thus the fura-FF recordings showed slightly faster kinetics of the flux peak component.
Figure 9C shows the voltage dependence of the rise time from the beginning of the voltage pulse to the peak of the release flux (time to peak). It decreased with increasing voltage and reached a minimal value at +50 mV. The scale shows the time from the beginning of the command voltage pulse at the output of the DA converter to the peak. Because the command and the measured signal were both filtered by Bessel filters a fixed delay time estimated to be 2.7 ms has to be subtracted (dashed line). The estimated minimal time to peak after correction for the filter effects was 3.24 ± 0.14 ms for fura-2. Figure 9G shows the voltage dependence for the time to peak input flux for fura-FF. It approaches a very similar value at large depolarizations as in the measurements with fura-2. The mean value at +50 mV after correcting for filter delays was 3.13 ± 0.24 ms. Figure 9D presents the ratio of peak versus plateau for fura-2 for the voltage range –20 to +50 mV for both release flux () and permeability (). The correction for depletion reduced the peak/plateau ratio values and the steepness of their voltage dependence. Figure 9H shows the peak-versus-plateau ratios for fura-FF. When comparing the mean values with those of Fig. 9D, the range of values is quite similar. Except for voltages between –20 and +10 mV values were not significantly different.
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In summary, despite small differences, the calculation results derived with the two indicator dyes, which are the result of different calibrations and different degrees of kinetic deconvolution, are very similar.
Discussion
Removal model fit in voltage-clamped EGTA-loaded mouse muscle fibres
In this study we describe the determination of Ca2+ fluxes in voltage-clamped adult muscle fibres of the mouse. Our method combined the advantages of intracellular perfusion as in whole-cell patch clamp recordings (Wang et al. 1999) with the better voltage control of a two-electrode system (Friedrich et al. 1999). It requires less preparative effort than the silicon grease gap technique described by Jacquemond (1997). A similar approach has recently successfully been applied to flexor digitorum brevis fibres of normal and mdx mice (Woods et al. 2004a).
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The high intracellular EGTA, forming the dominating Ca2+ buffer (denoted S in our removal model), captures almost all the Ca2+ that is released during a depolarization with little alteration to the time course of the release flux (Gonzalez & Ríos, 1993; Pape et al. 1995; Ríos & Brum, 2002). The removal model fit algorithm that we used extracts from the fluorescence ratio records information on the time course of Ca2+ binding to S and determines its rate constants. It cannot, however, provide an unambiguous determination of kon,EGTA in the cell as long as the intracellular dissociation constant of the indicator is not known (Schuhmeier & Melzer, 2004). According to Konishi et al. (1988), it is likely that the dissociation constant for fura-2 and other indicators is higher inside muscle cells than in free solution. On the other hand, the rate constant koff,S is insensitive to the KDye value chosen (Schuhmeier & Melzer, 2004). In agreement with our value of 4.94 s–1 in this study, are several groups, who presented apparent ‘off ’ rate constants of EGTA to fit Ca2+ recordings in skeletal and cardiac myocytes obtained under conditions of millimolar intracellular EGTA. Reported values were between about 3 and 5 s–1 (Gonzalez & Ríos, 1993; Shirokova et al. 1996; Song et al. 1998; Schuhmeier & Melzer, 2004), i.e. about one order of magnitude higher than the values obtained from temperature jump (Naraghi, 1997) and stopped flow experiments (Smith et al. 1984) in free solution which were 0.5 s–1 and 0.3 s–1, respectively. The reason for this discrepancy is not clear. Gonzalez & Ríos (1993) attributed it to possible interactions of EGTA in the myoplasm, gradients of Ca2+ along the sarcomere or an effect of non-linear pumping and binding processes. Perhaps the possibility should also be considered that the in vitro experiments underestimated the true values in free solution.
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As we confirmed using simulated fluorescence traces of the kind shown in Schuhmeier et al. (2003), koff,Dye can reliably be determined by the removal fit procedure, thus providing an in vivo determination of the dynamic properties of the indicator. This parameter is essential for the correct temporal deconvolution of the fluorescence signals (eqn (1), Klein et al. 1988) and therefore for the determination of the time courses of [Ca2+] and Ca2+ input flux. The mean value of 34 s–1 obtained for the fura-2 dissociation rate constant in the present experiments is somewhat smaller but close to the value found in experiments on myotubes (45 s–1Schuhmeier et al. 2003; 46 s–1Schuhmeier & Melzer, 2004). It is 35–40% of the values reported for free solution (84 s–1, Jackson et al. 1987; 97 s–1, Kao & Tsien, 1988) in agreement with the data of Baylor & Hollingworth (1988) and Garcia & Schneider (1993), after correcting for the effect of temperature (Bakker et al. 1997). Consistent with the lower affinity and more rapid kinetics of Ca2+ binding, the removal model fit revealed a 5.7-fold higher off rate constant koff,Dye for fura-FF than for fura-2 (192 versus 34 s–1).
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Ca2+ input flux in mouse muscle fibres
Independent of the rate constants of individual model components determined with the fit algorithm, it is important to note that the final calculation result of the analysis, i.e. the Ca2+ input flux, proved to be insensitive to the choice of the KDye value (Schuhmeier & Melzer, 2004 and Fig. 2F. Consistent with its faster kinetics fura-FF showed faster fluorescence responses than fura-2 with a clear peak component at the beginning and faster decline at the end (Fig. 8A) but led to a very similar time course of Ca2+ input flux as derived from fura-2 records (Fig. 9) supporting the notion that the time course of the Ca2+ input flux is equally well determined by both indicators despite the stronger temporal deconvolution that is necessary for fura-2 records. Fura-FF fluorescence transients, however, showed a lower signal-to-noise ratio and corresponding problems in fitting the traces with the removal model in part of the cells. It was also more difficult to resolve the time course of slow changes after the end of the pulse and during the plateau phase which made the depletion correction less reliable. Thus, for use under high [EGTA] conditions and depending on the focus of the study, dyes with low Ca2+ affinity may not be the optimal choice.
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Even though the estimated input flux amplitude is virtually invariant to assumptions made for KDye in the myoplasmic environment as shown by Schuhmeier & Melzer (2004; see also Fig. 2F), it is proportional to the assumed value of [S]total. The effective [S]total value depends on the fractional loading of the intracellular space with the solution in the pipette. The estimated average value of 40% loading in the present experiments (which all followed the same timing scheme), obtained by comparison with fluorescence recordings from dye-filled microcapillaries, corresponds to a putative [S]total value of 6 mM. Despite some uncertainty in this value it seems clear from the results of Fig. 4 that the amplitude of the Ca2+ entry flux is many times smaller than the release flux and both time course and voltage dependence of Ca2+ input flux indicate that Ca2+ entry is of negligible contribution to the optical signal consistent with investigations on adult frog muscle fibres (Brum et al. 1987, 1988).
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Comparison with previous results on Ca2+ transients in mouse muscle fibres
Several groups have investigated action potential-induced Ca2+ transients in single mouse fibres (e.g. Hollingworth et al. 1996; Liu et al. 1997; Bakker et al. 1997; Westerblad et al. 1997; Tutdibi et al. 1999; Bruton et al. 2003; Baylor & Hollingworth, 2003; Woods et al. 2004b). Hollingworth et al. (1996) estimated Ca2+ release for single action potentials with the fast dye furaptra and reported peak rates of 140–150 μM ms–1. In a more recent paper the same group determined 212 μM ms–1 (Baylor & Hollingworth, 2003). These values are quite close to our peak Ca2+ input fluxes at +50 mV (166 μM ms–1 for fura-2 and 209 μM ms–1 for fura-FF measurements).
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Only a few studies are available investigating Ca2+ currents or Ca2+ signals in mouse fibres under voltage clamp conditions. Friedrich et al. (1999) measured Ca2+ inward currents in unperfused isolated toe muscle fibres bathed in a solution containing 2 mM Ca2+. The activation characteristics appear to be similar to those found by Wang et al. (1999) who likewise measured Ca2+ inward currents in the presence of 2 mM external Ca2+ using mouse flexor digitorum brevis (FDB) and the whole-cell patch clamp technique. This group reports values of maximal Ca2+ conductance (gCa,max) = 85 S F–1 and V1/2=–0.3 mV compared to gCa,max= 55 S F–1 and V1/2=–18.8 mV determined in a double vaseline gap system (Delbono et al. 1997). Using the silicone grease gap technique (Jacquemond, 1997) and 5 mM extracellular Ca2+, Szentesi et al. (2001) determined parameters gCa,max= 200 S F–1, VCa= 73 mV, V1/2=–10.8 mV, k= 7.1 mV, whereas in our experiments (10 mM external Ca2+) we found gCa,max= 112 S F–1, VCa= 77.6 mV, V1/2= 3 mV, k= 4.9 mV.
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Voltage control of Ca2+ release was studied in mouse fibres by Wang et al. (1999) and by Jacquemond (1997). These groups assessed Ca2+ release properties by investigating the amplitude of Ca2+ transients that were measured with different indicators (fluo-3, calcium green-5N or calcium orange-5N and indo-1, respectively). Wang et al. (1999) obtained half-maximal activation of their Ca2+ signals at +6.2 mV whereas the indo-1 signals of Collet et al. (1999) reached their half-maximal value between –30 and –20 mV. In a recent preliminary report Woods et al. (2004a) describe measurements using an experimental arrangement similar to ours with a high concentration of intracellular EGTA and using the Ca2+ indicator oregon green-488–BAPTA-5N (OGB-5N). The voltage of half-maximal activation was –42 mV, which is considerably more negative than in the other studies and in our fura-2 flux measurements (about –10 mV for peak flux). Apparently, parameters of the voltage dependence vary considerably between different studies. The variance is likely to be due to the different methods and solutions used and sets limits to a quantitative comparison of individual parameters among different studies.
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Comparison with input flux estimates in voltage-clamped rat fibres
Ca2+ release flux in voltage-clamped mouse muscle fibres has not yet been investigated by other groups but data are available for cut rat EDL fibres with low and high intracellular EGTA showing amplitudes of up to about 20 μM ms–1 (Garcia & Schneider, 1993; Delbono & Stefani, 1993; Shirokova et al. 1996). Csernoch et al. (1999a,b) used enzymatically isolated cut fibres and reported maximal peak permeabilities of 1.4% ms–1 which amounts to a flux of 25 μM ms–1 with their estimated mean SR content of 1.8 mM. Voltages for half-maximal activation of –4.6 and –17.2 mV were reported. The SR permeability of our mouse fibres was activated in a similar voltage range (V1/2=–11 mV in the fura-2 experiments and V1/2=–4.4 in the fura-FF experiments) but showed up to 4 times larger maximal peak permeabilities, i.e. about 6% ms–1 at +50 mV corresponding to 166.6 μM ms–1 for an estimated SR content of 3 mM. Thus, the isolated mouse fibres under the present experimental conditions appear to be more efficient in releasing Ca2+ from the SR than the previously studied cut rat fibres. Nevertheless, they showed a similar voltage dependence of peak-to-plateau ratio of permeability as reported for this preparation by Shirokova et al. (1996).
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When exploring voltage sensor charge movements and Ca2+ release flux at large depolarizations (between 0 and +60 mV) in frog fibres, Shirokova et al. (1995) made the observation that the amount of sensor charge was not constant but kept increasing in parallel with an increase in release permeability. In mammalian muscle, Ca2+ release flux at large depolarizations has not been investigated yet. In our experiments we observed an almost linear increase in peak input flux and permeability between +20 mV and +50 mV (Figs 4C, 5D and E, 6A and 8D). Only in the fura-FF peak flux–voltage dependence (Fig. 8C) is this behaviour not evident due to the larger run-down indicated by the bracketing +20 mV recordings. Our preliminary estimates of intramembrane charge movements revealed no second voltage-dependent component at large depolarizations (Fig. 5E). However, despite its slow activation, the Ca2+ conductance introduced some uncertainty into charge measurements in this voltage range and maximum charge density was lower than determined previously in a very similar preparation (Wang et al. 1999). This quasi-linear activation of Ca2+ release permeability at large depolarizations deserves further investigation. It may be related to the continuing decrease in the time to peak with increasing depolarization (Fig. 9C), possibly reflecting progressive synchronization of activation of the ryanodine receptors.
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Estimates of SR Ca2+ content
Following procedures introduced by Schneider et al. (1987) we corrected release flux records during a depolarizing voltage pulse for depletion in the SR and converted them to permeability records by scaling them with 1/(Ca0,SR–Flux x dt). The correction procedure led to values of the SR content before the pulse (Ca0,SR) and its fractional change during the pulse (see Fig. 5B). In our fura-2 experiments we estimated a mean value of about 3 mM of the SR Ca2+ content (referred to the myoplasmic water space and using the estimated 6 mM intracellular EGTA). For measurements done on frog fibres under similar conditions (high intracellular EGTA), and using similar calculations, Gonzalez & Ríos (1993) report Ca0,SR values between 0.7 and 4 mM. In a comparative study on frog and rat fibres Shirokova et al. (1996) obtained an average of 3.4 mM for frog and 1.9 mM for rat. Using a pH-metric method and 20 mM intracellular EGTA Pape et al. (1995) estimated the total release of Ca2+ in cut frog fibres caused by trains of 20–40 action potentials at 50 Hz that supposedly led to complete depletion of the SR. They obtained a mean value of 2.7 mM.Posterino & Lamb (2003) measuring force transients of skinned rat EDL fibres reported normal Ca2+ contents between 1.5 and 1.8 mM, very close to results determined in the same preparation with a different method (Fryer & Stephenson, 1996; Owen et al. 1997) and 4–5 mM in maximally loaded fibres. Thus the values of Ca0,SR estimated in our experiments in isolated mouse interosseus fibres are similar to various independent estimates in other vertebrate skeletal muscle cells. According to Baylor et al. (1983) myoplasmic water (exclusive SR and mitochondria) occupies 71% of the total fibre volume. The SR volume is about 10% of the fibre volume (Peachey, 1965; Mobley & Eisenberg, 1975). Thus the total Ca2+ concentration within the SR in our experiments is calculated to be 0.71 x 3 mM/0.1 = 21.3 mM.
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Is SR Ca2+ permeability in mouse fibres dependent on lumenal [Ca2+]
The determinations of SR content (Ca0,SR) are based on the assumption (Schneider et al. 1987) that permeability is constant during the time of the plateau of the Ca2+ release flux, i.e. that the slope measured during the plateau results exclusively from the decrease of the Ca2+ concentration in the SR. Experiments on voltage-clamped frog fibres (e.g. Pape & Carrier, 1998; Pizarro & Rios, 2004) indicated that permeability does not stay constant in the plateau of the release flux when fast inactivation is over. Instead it was reported to increase with progressive depletion, thus providing a feedback to stabilize release (see also Posterino & Lamb, 2003). In this model, the almost linear decline in flux during the time of the flux plateau is thought to be not only caused by a decrease in the Ca2+ gradient driving the flux but also to be modulated by a superimposed change in permeability that is not immediately evident from the flux time course. We determined that a 100 ms voltage pulse that maximally activates Ca2+ release leads to about 80% reduction in the initial SR content. This substantial decrease might cause a significant depletion-dependent permeability change which we ignore in our analysis.
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Systematic analysis of depletion and its putative effect on permeability under voltage clamp conditions has recently been performed on frog skeletal muscle fibres (Pizarro & Rios, 2004). A decrease of SR content to 25% was reported to cause an increase in plateau permeability by more than 2-fold. Other authors reported even larger changes (e.g. Pape & Carrier, 1998). With the present experiments, we cannot rule out an effect of SR content on SR permeability but it seems that in the mouse it is less substantial and possibly even absent. Consistent with the constant permeability assumption, we found that the estimated Ca0,SR values were very similar over a wide range of voltages that released different amounts of Ca2+ and generated different slopes in the flux plateau (see Fig. 6A). If the applied assumption was not correct, i.e. if the SR permeability was critically dependent on the lumenal concentration of Ca2+, we would have expected to calculate more strongly deviating values for Ca0,SR at different voltages. Furthermore, the analysis in Fig. 7 indicates strong depletion during the sequence of identical pulses but only a small increase in permeability, which may also have resulted from slightly underestimating the recycling of Ca2+ to the SR lumen. The data presented in Fig. 7B and C show other differences to the frog experiments. In particular, the peak of the flux did not decrease in relation to the plateau as reported for frog fibres and the decay from the peak to the plateau was not significantly slowed (Fig. 7B and C) as found in frog fibres (Pizarro & Rios, 2004).
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If the constant permeability assumption is not made, an exact and independent quantification of recycling and remnant Ca2+ in the SR becomes extremely important. When the lumenal SR content reaches low values, small errors in the estimated concentration will lead to large errors in the calculated permeability value. The SR content is usually deduced from the cytoplasmic Ca2+ signal or a signal related to it (e.g. pH changes) and includes certain assumptions about the Ca2+ recycling activity. A direct measurement of Ca2+ in the SR seems therefore necessary to obtain exact information on the permeability time course for each pulse and to verify a possible dependence of permeability on the filling state of the SR.
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Conclusion
In the present study we assessed previously unavailable properties of the SR calcium release mechanism in single voltage-clamped muscle fibres of the mouse. We investigated in detail voltage-controlled global Ca2+ fluxes originating from extracellular and intracellular sources. The results provide information on the time course and voltage dependence of the Ca2+ fluxes. Our study also supplies data on the loading state of the sarcoplasmic reticulum and its dynamic change during depolarizations of different strengths. It does not provide clear evidence for an effect of lumenal Ca2+ on SR permeability. The growing importance of mouse models with defined alterations in skeletal muscle proteins calls for adequate functional investigation. Thus the approach applied here will be of benefit for future investigations of genetically altered mice to study changes in excitation–contraction coupling and properties of storing and buffering Ca2+ in the SR.
, 百拇医药
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