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1 Universität Ulm, Abteilung für Angewandte Physiologie, Albert-Einstein-Allee 11, D-89069 Ulm, Germany
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Abstract |
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(Received 13 August 2004;
accepted after revision 28 October 2004;
first published online 4 November 2004)
Corresponding author W. Melzer: University of Ulm, Department of Applied Physiology, Albert-Einstein-Allee 11, D-89069 Ulm, Germany. Email: werner.melzer{at}medizin.uni-ulm.de
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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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).
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.
Some of the results have been presented previously as an abstract (Ursu et al. 2004).
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Methods |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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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).
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):
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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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.
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.
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.
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).
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| (1) |
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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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.
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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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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| (2) |
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).
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.
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).
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Results |
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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.
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.
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).
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.
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.
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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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).
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.
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.
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).
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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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.
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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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).
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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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).
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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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.
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.
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Discussion |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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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).
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.
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).
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.
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).
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).
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.
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.
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 & Schneide
