Address correspondence to Werner Melzer, University of Ulm, Dept. of Applied Physiology, Albert-Einstein-Allee 11, D-89069 Ulm, Germany. Fax: 49-731-500-23260; email: werner.melzer{at}medizin.uni-ulm.de
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ABSTRACT |
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Key Words: mammalian skeletal muscle excitationcontraction coupling accessory subunits knockout mouse voltage-dependent inactivation
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INTRODUCTION |
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The dihydropyridine-sensitive 1S polypeptide of skeletal muscle is stably associated with four auxiliary subunits that are also found in the purified channel complex, ß1,
1,
, and
2. The latter two originate from the same gene and are postranslationally cleaved, but remain linked by disulfide bridges (Arikkath and Campbell, 2003
).
1 is most specific for skeletal muscle (Biel et al., 1991
). It is a polypeptide of 32 kD molecular weight, consisting of 222 amino acid residues, that is encoded by a gene with four translated exons residing as a single copy in the haploid mouse genome (Powers et al., 1993
; Wissenbach et al., 1998
). It exhibits four putative membrane-spanning
helices (Bosse et al., 1990
; Jay et al., 1990
) and both NH2 and COOH termini are located on the cytoplasmic side.
Because it proved difficult to express the skeletal muscle 1S subunit in heterologous expression systems, functional coexpression studies of the
1 subunit have been performed together with the cardiac
1C pore-forming subunit (Singer et al., 1991
; Wei et al., 1991
; Lerche et al., 1996
; Eberst et al., 1997
; Sipos et al., 2000
). These investigations indicated alterations in steady-state inactivation caused by
1 (shift to more negative potentials) and enhanced activation and inactivation kinetics. The experiments could, however, only provide indirect clues as to possible functions of
1 in skeletal muscle. Neither could one be sure that
1S resembles
1C regarding interaction with
1 nor did these experiments provide information on a possible modulation of Ca2+ release.
Two laboratories have independently generated mice lacking expression of the 1 subunit (Ahern et al., 2001
; Freise et al., 2000
). Unlike
1S/ and ß1/ mice that die at birth because of complete failure of EC coupling,
/ mice showed no obvious deviation from the normal phenotype. Nevertheless, in myotubes derived from the
-knockout mice, several functional differences have been described in comparison to myotubes of control animals (Freise et al., 2000
; Ahern et al., 2001
; Ursu et al., 2001
). Changes in Ca2+ inward current densities were reported using neonatal or embryonic mice as the source of myoblasts for myotube cultures (Freise et al., 2000
; Ahern et al., 2001
; Held et al., 2002
). The experiments indicated a partial suppression of L-type current by the
subunit. In contrast, myotubes of older
/ mice (4 wk and more) showed no significant deviations from controls in the L-type current densities, and a difference in sensitivity to 8-Br-cAMP observed in myotubes of young mice was not found either (Ursu et al., 2001
; Held et al., 2002
). This points to a
1-controlled cAMP modulation of the channel restricted to an early period of development. In addition, compatible with the coexpression studies in nonmuscle cells, a shift to more positive potentials of the curve that describes steady-state inactivation of the Ca2+ channels as a function of voltage was found in myotubes (Freise et al., 2000
; Ahern et al., 2001
). This effect was independent of cAMP and of the age of the mice (Held et al., 2002
).
Because the primary function of the DHPR protein in skeletal muscle is voltage sensing to control Ca2+ release from the SR (Melzer et al., 1995), excitationcontraction coupling events were investigated by Ursu et al. (2001)
and Ahern et al. (2001)
in
1/ myotubes and by Ursu et al. (2001)
in isolated fast and slow twitch muscles. The flux of Ca2+ underlying the depolarization-induced Ca2+ transients was found to be slightly larger in
/ myotubes (Ursu et al., 2001
) but voltage sensor charge movements were not statistically different (Ahern et al., 2001
). Neither were contractile properties of extensor digitorum longus (EDL) and soleus, investigated by single twitches and tetani both under normal and fatiguing conditions, found to be different (Ursu et al., 2001
). Yet, voltage clamp studies of Ca2+ currents and EC coupling events in mature fibers of
1 knockout mice have not yet been performed. Therefore, the purpose of the present investigation was to study functional effects of eliminating the
1 subunit on Ca2+ inward currents and Ca2+ release under voltage clamp conditions in enzymatically isolated adult skeletal muscle fibers of
1-deficient mice. The results show that of the different effects attributed to the
1 subunit, the alteration of voltage-dependent inactivation prevails in mature muscle. In particular, we demonstrate that
1 enhances inactivation of SR Ca2+ release in a voltage-dependent manner.
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MATERIALS AND METHODS |
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Solutions
Solutions used for the experiments had the following compositions (concentrations in mM): Krebs-Ringer solution for muscle dissection, fiber storage, and contraction experiments, 118 NaCl, 3.4 KCl, 0.8 MgSO4, 1.2 KH2PO4, 11.1 glucose, 25 NaHCO3, 2.5 CaCl2, pH 7.4; contracture solution, 2 TES, 1 MgSO4, 11 glucose, 2.5 CaSO4, 4.8 KCl, 57.6 K2SO4, 40 Na2SO4, pH 7.4; Ca2+-free contracture solution, 2 TES, 11 glucose, 3.5 MgSO4, 4.8 KCl, 57.6 K2SO4, 40 Na2SO4, pH 7.4; dissociation solution for muscle fiber isolation, Krebs-Ringer solution containing 2 mg/ml 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; internal (pipette) solution for intracellular perfusion, 145 CsOH, 135 aspartic acid, 0.75 Na2ATP, 4.25 MgATP (5.16 mM total Mg2+, resulting in 1 mM free Mg2+), 1.5 CaCl2 (resulting in 20 nM free Ca2+), 10 HEPES, 15 EGTA, 0.2 fura-2, 5 Na2creatinePO4, pH 7.2.
Contractions
Force recordings were performed at 25°C as described previously (Ursu et al., 2001). Extracellular electrical field stimulation was performed by applying supramaximal shocks of 1 ms duration. The experiments started in normal Krebs-Ringer solution by eliciting a twitch and a tetanus (500 ms, 125 Hz) that was used for contracture force normalization. To permanently depolarize the muscle fibers, the volume of the recording chamber (40 ml) was replaced with contracture solution containing 120 mM [K+] (at constant [K+]x[Cl]) immediately after the test tetanus. As the result, a contracture developed, followed by a slow spontaneous relaxation caused by inactivation. After 5 min, the high [K+] solution was washed out with normal Krebs-Ringer solution.
Voltage Clamp and Data Acquisition
Single cell experiments were performed at room temperature (2023°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
). Fibers were voltage clamped with two microelectrodes using an Axoclamp 2B amplifier (Axon Instruments, Inc.). Micropipettes were fabricated from borosilicate glass (GB150TF-10; Science Products). The voltage recording electrodes were filled with 3 M KCl and had a mean resistance of 6.76 ± 0.51 M
(n = 20) when measured in extracellular solution.
The current passing electrodes were filled with artificial internal solution containing 15 mM EGTA and 0.2 mM fura-2 and had a mean resistance of 2.79 ± 0.09 M (n = 20). After inserting the voltage-recording electrode, the control voltage was set to 80 mV with the voltage clamp circuit at minimum gain (30). Then the current-passing electrode was gently sealed to the membrane and the previously applied positive pressure was released, which usually led to establishing the contact to the cytoplasm. The amplifier gain was then increased to the final value of 800 used in all experiments.
The progress of diffusion of intracellular solution into the fiber was observed by measuring the increase in the resting fluorescence of fura-2 at 360 nm excitation (see also Schuhmeier et al., 2003). To study voltage-dependent activation of slow Ca2+ inward current and Ca2+ release, fibers were stimulated with 100-ms depolarizing pulses of increasing amplitude separated by intervals of 60 s (activation protocol). Voltage activation was started after 30 min of loading. [Ca2+]-dependent fura-2 fluorescence changes were recorded at 380 nm excitation and the ratio R (=F380/F360) calculated. For the fura-2 concentration at the time when voltage activation was performed, we used a mean estimate of 83 µM, obtained with a method described by Klein et al. (1988)
. The estimate resulted from F360 recordings in fibers that were compared with equivalent measurements in quartz microcapillaries of two different inner diameters (50 and 100 µm) containing 50, 100, and 200 µM of dye in internal solution. Assuming comparable diffusion rates for EGTA and fura-2, the concentration of the chelator in the fibers corresponding to the dye concentration is 6 mM, which we used for the calculations.
Current, voltage, and fluorescence were recorded simultaneously at 2 kHz sampling frequency using a CED 1401+ interface (Cambridge Electronic Design) connected to an AMD K6-2 computer. For data acquisition and analysis, we used own software written in Delphi (Borland International).
Ca2+ Current Analysis
Currentvoltage relations were least-squares fitted using
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Here, gleak and Vleak are conductance and reversal potential of a linear leak component and gCa,max and VCa are maximal conductance and reversal potential of the Ca2+ current. gleak and gCa,max are normalized by the linear capacitance. The gating function f(V) is defined by Eq. 2. V0.5, k, and F are the voltage of half-maximal activation, the voltage sensitivity, and the maximal value, respectively. F was unity for f(V) to describe fractional activation of conductance.
Ca2+ Input and Entry Flux Analysis
Free Ca2+ was calculated from voltage-activated changes of R (see above) using Eq. 3 after background and bleaching corrections (Klein et al., 1988; Schuhmeier et al., 2003
).
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Ca2+ input flux, i.e., the total flux of Ca2+ into the myoplasm, was derived as described by Schuhmeier and Melzer (2004). In brief, a kinetic model describing the removal of released Ca2+ to different compartments (see Melzer et al., 1986
) was fitted to the repolarization intervals of four consecutive depolarizing voltage pulses (50 ms, 0 mV, intervals 150 ms) usually applied 2 min after the last pulse of the activation protocol. The model consisted of the indicator dye described by Rmin, Rmax (3.52 and 0.41, respectively, determined for this setup), rate constants kon,Dye, koff,Dye and concentration [Dye]total, of a saturating buffer (parameters kon,S, koff,S, and [S]total), and an uptake mechanism (rate constant kuptake). KDye = koff,Dye/kon,Dye was set to 0.276 µM determined in vitro (Schuhmeier et al., 2003
). The best fit values of kinetic constants (koff,Dye, kon,S, koff,S, and kuptake) in the removal model were used to calculate the depolarization-induced Ca2+ flux into the myoplasm from other voltage-activated fluorescence records in the experiment.
Ca2+ entry flux was calculated from the measured Ca2+ current as described by Schuhmeier and Melzer (2004) assuming a fractional fiber volume for Ca2+ distribution of fV = 0.71 (see Baylor et al., 1983
) and a volume capacitance ratio VC = 0.32 liter F1 (mean value obtained from simultaneous volume and capacitance measurements).
Depletion Correction and Conversion of Release Flux to Permeability
We subjected the calculated Ca2+ input flux records to an analysis procedure that corrects for the effect of store depletion caused by the release to derive the time course of SR Ca2+ permeability during a depolarizing voltage step (Gonzalez and Ríos, 1993; Schneider et al., 1987
). Permeability was calculated as flux divided by the time-dependent Ca2+ content in the SR, both referred to the myoplasmic water volume. The Ca2+ content is the difference between an initial Ca2+ content and the released amount. The procedure assumes that permeability is constant during the plateau phase of the flux trace and determines the initial Ca2+ concentration in the SR that leads to zero slope in the plateau phase of the calculated permeability traces (see Schuhmeier and Melzer, 2004
).
Statistics
Unless otherwise stated, averaged data are presented and plotted as means ± SEM (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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Fig. 2 shows averaged records and their point-by-point SEM of the calculated fluxes at several different test voltages for the 16 experiments on +/+ and the 19 experiments on
/ cells. In both groups of cells, the time course of the calculated Ca2+ input flux shows a fast (arrow 1 in A) and a slow decay component (arrow 2 in A) after the initial peak as has been reported in many previous publications for other muscle preparations (e.g., Schneider et al., 1987
; Gonzalez and Ríos, 1993
; Csernoch et al., 1999b
; Schuhmeier and Melzer, 2004
). Comparing the averaged traces at each voltage showed no significant differences in time course and scale of the flux responses.
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The mean result of this calculation, performed for each individual experiment is presented in Fig. 2 in comparison to the uncorrected Ca2+ input fluxes (B versus A for +/+ and D versus C for
/). The result shows a very similar time course in both types of fibers, consistent with Ca2+ permeability reaching a maximum early during the voltage pulse and then decaying to a lower value due to rapid partial inactivation as has been described for other muscle preparations (Schneider et al., 1987
; Gonzalez and Ríos, 1993
; Csernoch et al., 1999b
; Schuhmeier and Melzer, 2004
).
Fig. 3 compares the voltage dependence of peak and end level both for Ca2+ input flux (A and B) and for permeability (C and D). Fig. 3 (A and B) is normalized to the maximal value at +50 mV in the controls (+/+). Absolute amplitudes of the release flux estimates depend on the assumed fractional loading of the cell with the EGTA in the pipette solution (see Schuhmeier and Melzer, 2004
). The estimation of intracellular fura-2 concentration at the time of the measurements (see MATERIALS AND METHODS) indicated a mean fractional loading of 41%. With this value, the peak amplitudes of Ca2+ input flux at +50 mV were 144.03 ± 17.08 µM/ms and 108.83 ± 17.81 µM/ms for
+/+ and
/, respectively. The calculated peak permeabilities at +50 mV were very similar, showing values of 5.19 ± 0.43%/ms and 4.94 ± 0.68%/ms, respectively. For comparison, using the procedure described in MATERIALS AND METHODS, we estimated the size of the maximal flux of Ca2+ entry (at +10 mV) from the data of Fig. 1 (C and G). The values were 0.18 ± 0.01 µM·ms1 and 0.19 ± 0.01 µM·ms1 for
+/+ and
/, respectively. Thus, the Ca2+ input flux is essentially identical to the Ca2+ release flux in both preparations and has only a small contribution from Ca2+ entry. The peak release flux values were larger than reported previously for voltage-clamped rat fibers (e.g., Garcia and Schneider, 1993
; Shirokova et al., 1996
; Csernoch et al., 1999a
,b
) but similar to flux amplitudes reported for action potentialstimulated mouse fibers (e.g., Baylor and Hollingworth, 2003
). Because the rat experiments were performed on cut segments of muscle fibers, the long depolarization during dissection may have contributed to a lower release activity in these experiments.
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In conclusion, the experimental data described so far provide little indication that properties of the voltage-dependent input flux of Ca2+ activated within a 100-ms step of depolarization are changed. Thus, neither entry from the extracellular space nor release from the SR seem to be affected by elimination of the DHPR 1 subunit in adult muscle fibers.
Ca2+ Current Inactivation
A consistent finding in several functional investigations of the 1 subunit coexpressed with the (cardiac)
1 subunit in nonmuscle cells and performed on myotubes of knockout mice (both neonatal and adult) was a change in the "steady-state" voltage dependence of inactivation of Ca2+ conductance (see INTRODUCTION). We therefore investigated whether this functional difference was preserved in the fully differentiated state of the muscle fiber or whether it disappeared with maturation like the modulatory effect on Ca2+ current density.
To study the process of slow voltagedependent inactivation, we used a pulse paradigm in which increasingly depolarizing steady voltage levels were applied. Each new depolarization interval lasted 30 s and was ended with a short (100 ms) test depolarization to a fixed voltage of +20 mV (Fig. 5 A). This cumulative inactivation procedure was chosen to avoid repeated long depolarizations of increasing amplitude that would have led to repeated strong and long-lasting activation of Ca2+ release with a high risk of prematurely destroying the muscle fibers.
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Fig. 5 (D and E) shows the results from the same type of experiment performed in a muscle fiber of a / mouse. When comparing the traces, it becomes evident that small activation by the +20 mV pulse of both Ca2+ current and Ca2+ release is still possible at a holding potential of 20 mV in the
/ fiber. At the same potential, the
+/+ fiber shows no trace of response. Therefore, membrane depolarization appears to be less efficient in inactivating DHPR in the
/ fiber.
The qualitative impression obtained from Fig. 5 is confirmed quantitatively when comparing mean values of the fractional inactivation at different voltages from several fibers. The experimental results of Fig. 6 were obtained from eight +/+ and 11
/ fibers using the experimental protocol of Fig. 5 A.
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Inactivation of Ca2+ Flux
Because inactivation of the DHPR-mediated Ca2+ current (slow L-type current) was affected by 1 knockout and because the DHPR voltage sensor also controls Ca2+ release from the SR, it was important to determine whether the Ca2+ input flux derived from the fura-2 ratio signals was changed by the knockout in a similar way. The example of Fig. 5 indicates that this is indeed the case and the statistical evaluation of the same
+/+ and
/ fibers that were used for Fig. 6 A confirmed this notion. In Fig. 6 B, we plotted the mean peak values of the Ca2+ input flux traces versus conditioning voltage. Again, the results of the two groups showed significant differences. V0.5 exhibited a mean value of 36.09 ± 1.36 mV in
/ versus 50.21 ± 2.79 mV in
+/+ and the sensitivity parameter k was 4.96 ± 0.30 mV in
/ versus 5.98 ± 0.329 mV in
+/+. For the input flux end level, alterations were similar.
Thus, as for the Ca2+ current, the Ca2+ input flux exhibited a shift in its voltage dependence of inactivation to more positive membrane potentials in / fibers. To demonstrate the range in which the test pulse signals are altered more clearly, Fig. 6 (C and D) shows the difference between the inactivation curves of
+/+ and
/ at the various conditioning voltages. The details of the results obtained with the inactivation protocol are summarized in Table II. In summary, the experiments reported here provide first results on voltage-dependent inactivation of Ca2+ release in mammalian muscle and demonstrate that this inactivation is altered in a very similar way as L-type current inactivation in adult muscle fibers of the DHPR
-knockout mouse.
K+ Contractures in EDL Fiber Bundles
Previous measurements in adult -knockout muscle using single twitches or short tetani did not indicate any difference in force responses (Ursu et al., 2001
). However, if the steady-state voltage dependence of inactivation is altered whereas the voltage dependence of activation is unchanged, one might expect differences in the amplitude of the free Ca2+ transient and of the force transient during long depolarizations.
To investigate the effect of long depolarizations well above the activation threshold on force development in intact adult muscle under normal intracellular conditions, we measured the isometric tension of isolated fiber bundles of the EDL stimulated by a strong increase in the extracellular bath concentration of potassium.
The bundles were dissected from / and control mice of comparable size and were trimmed to similar diameter. The force induced by potassium stimulation was normalized by the tetanic force obtained before the potassium contracture.
Fig. 7 shows typical K+ contractures in a +/+ (A) and a
/ preparation (B) displayed in percent of tetanic force. The bath solution (Krebs-Ringer solution) in the recording chamber containing 3.4 mM potassium was replaced (within
15 s) by a solution containing 120 mM potassium. The amplitudes of the slow force transients relative to tetanus force in
/ muscle were consistently larger than in
+/+ muscle as shown in this example. Fig. 7 (C and D) presents the averaged responses from 11
+/+ and 11
/ bundles, respectively. The normalized force amplitude was 18.50 ± 1.41% of tetanic force in
/ muscle compared with 6.37 ± 0.43% in
+/+ muscle.
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DISCUSSION |
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Because of the established bifunctional role of the skeletal muscle DHPR as a voltage-dependent Ca2+ channel and as a voltage sensor for the activation of RyR we combined the recording of Ca2+ inward currents with measurements of intracellular Ca2+ transients. Using the Ca2+ transients, we quantified the flux of Ca2+ from its sources (SR and extracellular space) in the muscle fiber, which is commonly termed "Ca2+ input flux". Since Ca2+ input flux was about two orders of magnitude larger than the flux of Ca2+ entry from the extracellular space and showed an utterly different time course, it likely consists of almost pure Ca2+ release flux from the SR.
In the comparative approach presented here, experimental conditions and analysis procedures were identical for both +/+ and
/. Therefore, possible systematic errors in the quantification are of minor impact. The focus was on differences in the behavior of the two types of cells. In our experimental strategy, we included, in addition to protocols previously applied to myotubes (Ursu et al., 2001
), a new approach to study the effects of long-lasting depolarization on Ca2+ release, which has hitherto not been possible in mammalian muscle preparations.
Activation of Ca2+ Current and Ca2+ Release by Short Depolarizations
Experiments on primary cultured myotubes obtained from myoblasts of neonatal mice indicated an amplitude modulation of the Ca2+ inward current by the 1 subunit. The mean maximal current density was
30% larger in
/ myotubes (Freise et al., 2000
; Held et al., 2002
). Even though small, this difference has potential physiological relevance because it was shown to be modulated by cAMP. Myotubes derived from satellite cells of older animals (14 mo), on the other hand, showed no significant difference in size of the Ca2+ current densities (Ursu et al., 2001
; Held et al., 2002
). Similarly, in the present experiments, current density and voltage dependence of activation were not different in adult muscle fibers of
/ mice as compared with WT.
Investigating EC coupling in / myotubes (from adult animals) with short step depolarizations indicated
30% higher flux of Ca2+ input than for WT (Ursu et al., 2001
), pointing to a certain degree of suppression of voltage-dependent activation of Ca2+ release by the
subunit. This result seemed at first consistent with the increase in force amplitude observed in the present study in potassium contractures. However, in the isolated adult muscle fibers, voltage clamp activation by short depolarizations (100 ms as applied in myotubes) indicated no comparable difference in Ca2+ signals and in the calculated Ca2+ release flux at any voltage (Fig. 3). Thus, the data indicate that the force increase obtained with long-lasting potassium depolarizations in
/ fibers is not the consequence of altered voltage sensor activation. Differences regarding the amplitude of Ca2+ currents and Ca2+ release flux activated by short depolarizations in the fully reprimed state seem to be confined to developmental stages of muscle and to be lost after terminal differentiation. Held et al. (2002)
suggested two independent mechanisms linked to the
1 subunit that affect amplitude and steady-state inactivation, respectively. A bifunctional role has also been attributed to the neuronal
2 subunit (stargazin) that affects the targeting of AMPA receptors in addition to steady-state inactivation (Chen et al., 2000
). For
1, only the mechanism related to inactivation seems to be retained in adult muscle.
Inactivation of Ca2+ Current and Ca2+ Release
Both heterologous expression studies (Singer et al., 1991; Lerche et al., 1996
; Eberst et al., 1997
; Sipos et al., 2000
) and subsequent experiments on
/ myotubes (Freise et al., 2000
; Ahern et al., 2001
) indicated alterations of steady-state inactivation of the L-type Ca2+ current by the
subunit. In cells lacking expression of the subunit, stronger conditioning depolarizations had to be applied than in cells expressing
1 to reach the same degree of inactivation. These findings could be confirmed for adult muscle fibers in the present study. On average, the voltage of half-maximal current inactivation was 16 mV more positive in
/ fibers than in controls. This is quite similar to the shifts reported for myotubes (Freise et al., 2000
; Ahern et al., 2001
; Held et al., 2002
).
However, the main function of the skeletal muscle DHPR is the control of Ca2+ release from the SR. In contrast to Ca2+ inward current, Ca2+ release exhibits two distinct types of inactivation during step depolarization, a fast one operating within tens of milliseconds (Melzer et al., 1984, 1987
; Simon et al., 1991
) and a slow one that takes many seconds for completion (Brum et al., 1988
; Pizarro et al., 1988
; Melzer et al., 1995
). While fast inactivation of Ca2+ release flux appears to be a Ca2+-dependent property of the release channels (Schneider and Simon, 1988
; Jong et al., 1993
), slow inactivation is known to reside in the DHPR and is voltage dependent (Ríos and Pizarro, 1991
; Melzer et al., 1995
).
Slow voltagedependent inactivation of Ca2+ release has been studied in voltage-clamped adult muscle fibers of the frog both by force measurements and by Ca2+ measurements (e.g., Caputo and Fernandez, 1979; Caputo and Bolanos, 1987
; Pizarro et al., 1988
; Schnier et al., 1993
). In adult mammalian muscle, properties of inactivation have been indirectly assessed by K+ depolarization and force measurements (Chua and Dulhunty, 1988
, 1989
; Dulhunty, 1991
), but data on Ca2+ release flux inactivation in mammalian muscle have not been available until now. The higher sensitivity to long-lasting depolarization and the lower robustness of Ca2+ release had also precluded the investigation of Ca2+ release inactivation in myotubes. In the present study, we succeeded to study the properties of Ca2+ release inactivation due to conditioning depolarization and we could demonstrate that this process is affected in a very similar way as is Ca2+ current inactivation by the elimination of the
subunit. Slow inactivation is likely controlled by the same voltage-dependent mechanism that causes L-type current inactivation. The present results are consistent with this hypothesis, even though the strict identity of the DHPRs that generate Ca2+ inward current and those that control Ca2+ release remains to be demonstrated (Lamb, 1992
).
K+ Contractures
Considering the voltage clamp results on interosseus fibers, it seems likely that the observed difference in potassium contracture force in EDL muscle can be attributed to the altered inactivation properties of the DHPR during long depolarization rather than to an alteration in activation. A direct or indirect effect of a Ca2+ inward current showing weaker inactivation can be ruled out because the difference in force responses remained unchanged when extracellular Ca2+ was removed. Ca2+ current as the cause of the increased contracture force seemed also unlikely because of the small estimated size of Ca2+ entry compared with the total amount of Ca2+ mobilization. This leaves as the likely cause for the force difference the lower sensitivity to depolarization of SR Ca2+ release inactivation in the / muscle.
Fig. 9 summarizes the results on voltage-dependent Ca2+ release obtained from the single cell experiments and provides a tentative explanation of the force results obtained with the multicellular muscle preparation. The curves on the right in each panel of Fig. 9 A show the voltage dependence of fractional activation of plateau permeability (obtained from Fig. 3, C and D, squares). This provides an estimate of the steady-state permeability as a function of voltage in the absence of slow inactivation. The curves on the left of each panel show the fractional inactivation of Ca2+ input flux derived from the data in Fig. 6 B. The expected voltage dependence of release permeability in the steady state in the presence of slow inactivation can be calculated as the product of plateau permeability and fractional inactivation. Fig. 9 B compares the result of this calculation for +/+ (left panel) and
/ (right panel), respectively. Because of the selective shift of the inactivation curve caused by the elimination of
1, a "window" of noninactivatable permeability appears with a maximum at 30 mV (Fig. 9 B, right). Thus, a steady depolarization in the voltage range of about 50 to 10 mV can be expected to cause a steady release flux that should lead to a measurable elevation of Ca2+ in the myoplasm. In the inactivation experiments, steady elevations of Ca2+ and a difference between
+/+ and
/ consistent with the predicted voltage dependence of Fig. 9 B were in fact indicated by the fluorescence recordings. Fig. 9 C shows the mean values of the increase above resting levels (measured at 80 mV) of the indicator's fractional occupancy by Ca2+. The values were obtained from the baselines immediately before each test pulse at the end of the preceding conditioning period of the inactivation experiments described in conjunction with Figs. 5 and 6. The maximal change was 2.75-fold larger in
/ (Fig. 9 C, right).
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Conclusions
In summary, the results of the present investigation suggest that the slow voltagedependent inactivation of Ca2+ current and Ca2+ release is the main target of functional alteration by the 1 subunit in adult muscle fibers. Short term activation of Ca2+ current and Ca2+ release that had been implicated in
1 effects based on experiments in nonmuscle cells and myotubes were not found to be changed by the elimination of this auxiliary subunit in mature fibers. The finding that inactivation of SR Ca2+ release is affected is important because the SR is the predominant source of Ca2+ during voltage activation of skeletal muscle fibers.
The 1 subunit bears the potential for a modulatory role in EC coupling. By affecting steady-state inactivation, it can control the availability of DHPRs for voltage-dependent activation of Ca2+ release and may change the force response on the single fiber level. Whether
1-mediated modulation is actually used under physiological circumstances remains equally unclear at present as the molecular mechanism of slow inactivation in general. Alterations of the voltage dependence of slow inactivation of K+ contractures have been reported for rat EDL, for instance as a result of chronic administration of tri-iodothyronine (rightward shift; Chua and Dulhunty, 1989
) and of exercise training (leftward shift; Joumaa and Leoty, 2002
).
The shift of the inactivation curve to more negative potentials caused by 1 is reminiscent of the effect of Ca2+-antagonistic drugs on the voltage sensor for EC coupling (Berwe et al., 1987
; Ríos and Brum, 1987
; Pizarro et al., 1988
; Erdmann and Lüttgau, 1989
; Neuhaus et al., 1990
), which has in part been attributed to selective binding of the drugs to the inactivated conformation of the DHPR (Ríos and Pizarro, 1991
).
1 has been shown to interact directly with
1S, probably via the first two transmembrane domains (Arikkath and Campbell, 2003
). This interaction might stabilize the inactivated state in analogy to some Ca2+ antagonists and extracellular metal ion substitutions (e.g., Pizarro et al., 1988
; Erdmann and Lüttgau, 1989
; Feldmeyer et al., 1990
; Ríos and Pizarro, 1991
; Schnier et al., 1993
; Melzer et al., 1995
) and may even influence the force of action potentialinduced twitches or tetani under certain conditions. It seems, therefore, worthwhile to search for possible conditions that increase the strength of the
1
1S interaction beyond the normal level, thus provoking a true Ca2+-antagonistic effect.
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ACKNOWLEDGMENTS |
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We acknowledge a contribution to Dr. W. Melzer from the European Commission for graduate training (HPRN-CT-2002-00331). This work was funded by a grant of the Deutsche Forschungsgemeinschaft to W. Melzer (ME-713/10-3).
Olaf S. Andersen served as editor.
Submitted: 13 August 2004
Accepted: 4 October 2004
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REFERENCES |
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