Dissociation of charge movement from calcium release and calcium current in skeletal myotubes by gabapentin

Kris J. Alden and Jesús García

Department of Physiology and Biophysics, University of Illinois at Chicago College of Medicine, Chicago, Illinois 60607


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The skeletal muscle L-type calcium channel or dihydropyridine receptor (DHPR) plays an integral role in excitation-contraction (E-C) coupling. Its activation initiates three sequential events: charge movement (Qr), calcium release, and calcium current (ICa,L). This relationship suggests that changes in Qr might affect release and ICa,L. Here we studied the effect of gabapentin (GBP) on the three events generated by DHPRs in skeletal myotubes in culture. GBP specifically binds to the alpha 2/delta 1 subunit of the brain and skeletal muscle DHPR. Myotubes were stimulated with a protocol that included a depolarizing prepulse to inactivate voltage-dependent proteins other than DHPRs. Gabapentin (50 µM) significantly increased Qr while decreasing the rate of rise of calcium transients. Gabapentin also reduced the maximum amplitude of the ICa,L (as we previously reported) without modifying the kinetics of activation. Exposure of GBP-treated myotubes to 10 µM nifedipine prevented the increase of Qr promoted by this drug, indicating that the extra charge recorded originated from DHPRs. Our data suggest that GBP dissociates the functions of the DHPR from the initial voltage-sensing step and implicates a role for the alpha 2/delta 1 subunit in E-C coupling.

dihydropyridine receptor; excitation-contraction coupling; calcium channels; calcium transients; skeletal muscle


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

EXCITATION-CONTRACTION (E-C) coupling in skeletal muscle depends on the activation of dihydropyridine receptors (DHPRs). The skeletal muscle DHPR is composed of the pore-forming, voltage-sensing alpha 11.1-subunit and the auxiliary subunits alpha 2/delta 1, beta 1, and gamma 1 (9, 40). Upon membrane depolarization, DHPRs undergo conformational changes that give rise to three sequential events: charge movement (Q), calcium release, and calcium current (ICa,L) (16). Charge movement is the electrical manifestation of the movement of the voltage sensor in response to changes in membrane potential. Further conformational changes in the alpha 11.1 protein lead to activation of the sarcoplasmic reticulum (SR), calcium release channel, or ryanodine receptor type 1 (RyR1) (16, 36). The portion of the alpha 11.1 protein involved in controlling calcium release from the SR is the cytoplasmic loop connecting repeats II and III (10, 23, 36). The last event, ICa,L, occurs when the alpha 11.1 has suffered additional changes that allow channel opening and calcium influx.

The sequential relationship of the events resulting from DHPR activation would predict that any maneuvers that alter the conformational changes in alpha 11.1 would have a similar effect on Q, calcium release, and ICa,L. In support of this hypothesis, many experiments have demonstrated that inhibition of the DHPR voltage sensor (e.g., maintained depolarizations, DHPR antagonists, or changes in calcium levels) reduces Q, calcium release, and ICa,L with a similar time course and magnitude (5, 12, 13, 15, 21, 30). However, other studies have revealed alternative effects on E-C coupling. For example, the voltage dependence of charge movement and calcium release (or contraction properties) is shifted in the hyperpolarizing direction without changes in the voltage dependence of ICa,L in the presence of perchlorate (6, 11, 19) or the R615C mutation in the RyR1 (7). In addition, the DHP nifedipine is also able to induce calcium release (39) and modify contractile activity (22) while blocking ICa,L. The differential effects of perchlorate and nifedipine on the events generated by DHPRs indicate that E-C coupling can be modulated at different steps, leading to the generation of charge movement, calcium release, and ICa,L.

We have recently reported that the analgesic and antiepileptic drug gabapentin (GBP) causes a modest reduction of ICa,L in mouse skeletal myotubes (2). In the present paper we have extended those studies and examined the effect of GBP on charge movement and calcium release in mouse myotubes. Interestingly, GBP binds to alpha 2/delta 1-subunits from brain and skeletal muscle with similar kinetics and high affinity (kd = 38 nM in brain and 29 nM in skeletal muscle) (17, 34). Gee et al. (17) have also shown that, from 14 different tissues examined in the rat, the highest level of GBP-binding sites occurs in skeletal muscle. Therefore, this agent is unique because it binds to a subunit of the DHPR complex other than the alpha 11.1, which is considered to be the voltage sensor. Our present studies demonstrate that GBP increases charge movement and decreases the rate of calcium release, suggesting that the ability of the DHPR to trigger calcium release and to function as a calcium channel is impaired by this agent.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The experiments were approved by the Animal Care and Use Committee of the University of Illinois at Chicago and were conducted according to the Guide for the Care and Use of Laboratory Animals (National Academy of Sciences, Washington, DC, 1996).

Skeletal myotube cultures. Primary cultures of skeletal muscle tissues were prepared as previously described (14). Skeletal muscle of newborn mice was removed and finely minced. The small pieces of muscle were incubated at 37°C for 30-45 min in Ca2+, Mg2+-free rodent Ringer (in mM: 155 NaCl, 5 KCl, 11 glucose, and 10 HEPES, pH 7.4) containing collagenase type IA (1 mg/ml) (Sigma Chemical, St. Louis, MO). Dissociated muscle was triturated with a Pasteur pipette in plating medium [vol/vol, 80% Dulbecco's modified Eagle's medium (DMEM) with 4.5 g/l glucose, 10% horse serum, and 10% calf serum]. Large debris was removed from the solution by filtration and centrifugation, and a suspension of myocytes was obtained. Cultures were maintained in a 37°C incubator with a gas mixture of 95% air and 5% CO2. Skeletal myotubes were studied at 7-10 days after initial plating, and all experiments were preformed at room temperature.

Charge movement measurements. The whole cell configuration of the patch-clamp technique (20) was used for measurement of intramembrane charge movement, calcium transients, and ICa,L in skeletal myotubes. Data acquisition was synchronized with pulse generation by a personal computer-controlled 12-bit analog-to-digital/digital-to-analog Digidata 1200A converter. Linear components of the membrane were digitally subtracted by appropriate scaling and subtracting negative control currents that do not activate ionic conductances. Membrane capacitance was measured by integrating the area under the capacity transient before series resistance compensation and was used to normalize the charge moved and ICa,L measurements obtained from different myotubes. Data acquisition and processing were performed with pCLAMP 7.0 software (Axon Instruments). Recording electrodes were pulled from borosilicate glass and had resistances between 1.6 and 2.0 MOmega when filled with a solution containing (in mM) 140 Cs-aspartate, 5 Mg2Cl, 10 Cs-EGTA, and 10 HEPES, pH 7.4 adjusted with CsOH. The extracellular solution used to record charge movement contained (in mM) 145 TEACl, 2 CaCl2, 10 HEPES, 8 MgCl2, 0.5 CdCl2, 0.1 LaCl3, and 0.003 TTX, pH 7.4 adjusted with CsOH. For calcium transients and ICa,L recording, the extracellular solution contained 145 TEACl, 10 CaCl2, 10 HEPES, and 0.003 TTX, pH 7.4 adjusted with CsOH. Gating currents were elicited with 15-ms test pulses delivered from a holding potential of -80 mV in 10-mV increments from -40 to +60 mV. To isolate gating currents due to DHPRs and minimize contributions of gating currents from T-type, potassium, and sodium channels, we used a prepulse protocol, as described in Adams et al. (1). In this protocol, a 1-s depolarizing pulse to -30 mV is followed by a 25-ms repolarization to -50 mV and test pulses of varying amplitude. This voltage-dependent mechanism was used to partially immobilize the movement of positively charged amino acids or voltage sensors. The portion of charge movement resistant to the effect of a prepulse, herein referred to as Qr, represents the gating current of the DHPR, whereas the component of total charge (Qt) that is sensitive to the prepulse (Qs) presumably represents gating currents from T-type, potassium, and sodium channels (1). To determine the nifedipine-sensitive component of the gating current, paired recordings were performed in the absence and presence of 10 µM nifedipine (Sigma Chemical). GBP was kindly provided by Parke-Davis Research Laboratories (Warner-Lambert, Ann Arbor, MI). Skeletal muscle cells were incubated with 50 µM GBP for 1 h before examination, and it was maintained in the external recording solution.

Figure 1 shows the dose-response curve of the inhibition of ICa,L amplitude by GBP. From the normalized inhibition, we calculated an IC50 of 13.9 µM. This value of IC50 is comparable to those previously reported. Stefani et al. (32) determined an IC50 of 3.65 µM for isolated cortex neurons, 13.84 µM for striatum neurons, and 4.52 µM for globus pallidus neurons. Wamil and McLean (37) determined that the value of IC50 decreased with time of exposure to GBP. The IC50 was 130 µM for recordings obtained before 60 s after the application of GBP, 19 µM for recordings obtained between 10 and 60 min (this time and concentration are equivalent to our studies), and 4 µM for 12-48 h.


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Fig. 1.   Dose-response curve for inhibition of calcium current (ICa,L) by gabapentin (GBP). Points show the mean values for 5 (100 nM), 8 (10 µM), 24 (50 µM), and 5 (100 µM) myotubes. The effect was calculated from the maximum amplitude of the ICa,L at 20 mV, which is the peak of the current-voltage relationship. The smooth curve is the best least-squares fit to a one-to-one receptor-drug interaction.

Calcium transient measurements. Changes in intracellular calcium concentration were measured with the cell-impermeant fluorescent dye K5Fluo-3 (200 µM; Molecular Probes), as previously described (14). Individual cells were directly loaded through the patch pipette with Fluo-3 added to the internal solution (contents listed above). Fluorescence emission was collected with a photomultiplier tube mounted to the side port of a Nikon Diaphot 300 inverted microscope. The set of filters used for calcium measurements was as follows: excitation centered at 470 nm (±20 nm); dichroic long-pass mirror centered at 510 nm; and long-pass emission filter centered at 520 nm. The background fluorescence was measured for each myotube in the cell-attached mode before opening of the patch pipette. Measurement of changes in intracellular calcium concentration was expressed as Delta F/F (14), where Delta F denotes an increase in fluorescence from baseline values and F is baseline fluorescence measured before depolarization. Transients were recorded simultaneously with ICa,L and were elicited with 100-ms test pulses using the prepulse protocol.

Curve fitting and statistical analysis. For each cell, the voltage dependence of charge movement, calcium transient (Delta F/F), and calcium conductance (G) was fitted to a Boltzmann distribution
Y=Y<SUB>max</SUB><IT>/</IT>{1<IT>+</IT>exp[−(<IT>V−V</IT><SUB>½</SUB>)<IT>/k</IT>]} (1)
where Ymax was either Qmax, (Delta F/F)max, or Gmax; V1/2 is the membrane potential where Y = Ymax/2, and k is the slope factor. Data were analyzed using analysis of variance with repeated measures and are expressed as means ± SE. Statistical analysis was performed using Statistica 5.1 (StatSoft, Tulsa, OK). Significance for all data was set at P <=  0.05.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Effect of depolarizing prepulse on charge movement, calcium transients, and ICa,L. We used a prepulse protocol to isolate the events resulting from activation of DHPRs. Figure 2A shows records of charge movement obtained from skeletal myotubes in the absence and presence of a prepulse at different test potentials. The voltage dependence of the total charge movement and the charge resistant to a depolarizing prepulse are shown in the graph in Fig. 2A. The difference between the total and resistant charges represents the charge that can be inactivated by the prepulse and corresponds to voltage-sensitive channels other than the DHPR (1). The average amount of Qt was 9.5 ± 0.9 nC/µF, and the average of Qr was 5.2 ± 0.6 nC/µF (n = 27). These values of charge movement are in close agreement with values previously reported (1, 16, 28). The curve corresponding to Qt has a more negative V1/2 (-16.8 ± 1.1 mV) than the Qr curve (-6.2 ± 1.7 mV) because it contains charge from voltage-dependent proteins that activate at lower membrane potentials than the DHPR. The average value of k was 10.21 ± 0.6 and 12.36 ± 0.5 mV in the absence and presence of the prepulse, respectively.


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Fig. 2.   Effect of prepulse on gating current, calcium transients, and ICa,L measurements as a function of membrane potential. A: representative records of gating currents obtained in the absence (-) or presence (+) of a depolarizing prepulse to -30 mV. Membrane potential for both sets of records is indicated at left. The graph (right) shows that the average gating current densities elicited in the absence of a prepulse (; n = 31) were significantly reduced compared with gating currents elicited with a 1-s prepulse to -30 mV (black-triangle; n = 27) (*P < 0.05). Q, charge movement. B: calcium transients (left) elicited in the absence of a prepulse were not significantly different from those elicited with a 1-s depolarizing prepulse to -30 mV (n = 7). The graph (right) shows the voltage dependence of calcium transients (Delta F/F) in the absence () and presence (black-triangle) of a prepulse. C: ICa,L were elicited using voltage steps with (black-triangle) and without () (n = 14) a 1-s depolarizing prepulse to -30 mV. Maximum ICa,L densities were not significantly different between the groups, as shown by the graph (right).

In contrast to the decrease of charge movement with a depolarizing prepulse, neither calcium transients (Delta F/F) nor ICa,L showed changes in amplitude or voltage dependence. The graph in Fig. 2B shows calcium transients in the absence and presence of a prepulse and the fluorescence-voltage relationship. The average maximum Delta F/F was 1.37 ± 0.12 and 1.34 ± 0.13 (n = 7) in the absence and presence of a prepulse, respectively. Figure 2C shows the data corresponding to ICa,L. The current-voltage relationships overlap over the entire voltage range tested. The maximum ICa,L recorded was, on average, -9.6 ± 0.9 and -10.0 ± 1.0 pA/pF (n = 14) in the absence or presence of the prepulse, respectively. Thus, as previously suggested (16), these data indicate that the immobilization-resistant charge movement (Qr) is closely related to calcium release and ICa,L in skeletal muscle. We therefore used this experimental paradigm to study potential effects of GBP on Qr, calcium transients, and ICa,L.

Increase of immobilization-resistant charge movement by GBP. When skeletal myotubes were exposed to GBP, we found a large increase in Qr compared with untreated myotubes. Figure 3A shows representative records of Qr obtained in the absence and presence of GBP. The values of Qr were averaged at each membrane potential and used to construct the voltage dependence shown in Fig. 3B. The increase in Qr in the presence of GBP was observed for membrane potentials from -30 to 60 mV. The average maximum Qr in GBP-treated and control myotubes was 8.9 ± 0.9 (n = 23) and 5.1 ± 0.6 nC/µF (n = 31), respectively (P < 0.05). The average values of the other parameters describing the voltage dependence in GBP-treated myotubes were V1/2 = -8.0 ± 3.8 mV and k = 13.2 ± 1.3 mV (n = 23), whereas the corresponding values for control myotubes were V1/2 -6.2 ± 1.9 mV and k = 12.36 ± 0.5 mV (n = 31). The difference in the parameters of voltage dependence between the two conditions was not significant (P > 0.05). The graph in Fig. 3B shows normalized Qr to the maximum value for GBP-treated and control myotubes, where the similarity of the voltage dependence can be better appreciated. Because Qr is thought to represent the movement of the DHPR voltage sensor in the membrane, the increase in Qr indicates that GBP facilitates the movement of the receptor, yet it does not modify the voltage dependence.


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Fig. 3.   Voltage dependence of resistant charge movement. A: charge movement (Qr) records obtained from untreated (control) and GBP-treated myotubes, both in the presence of a prepulse to -30 mV. Numbers at left indicate the test potentials for both sets of records. B: Qr was significantly enhanced with 50 µM GBP treatment (black-triangle; n = 23) compared with control cells (; n = 31) at membrane potentials from -30 to 60 mV (*P < 0.05). C: Qr normalized to its maximum (Qmax) for 50 µM GBP-treated (black-triangle; n = 23) and control myotubes (; n = 31) demonstrate similar voltage dependence across membrane potentials from -40 to 60 mV.

Effect of GBP on calcium transients. Figure 4 shows typical calcium transients elicited from a control myotube (Fig. 4A) and a GBP-treated myotube (Fig. 4B) at -30, 0, and 50 mV. Calcium transients were also elicited with the prepulse protocol. The maximum amplitude of the calcium transients, measured at the end of the pulse, in GBP-treated myotubes (Delta F/F = 1.15 ± 0.19; n = 14) was similar to the amplitude of the transients recorded from control myotubes (1.25 ± 0.14; n = 20) (P > 0.05). The average values of V1/2 and k for GBP-treated myotubes were -0.56 ± 1.7 and 6.23 ± 0.4 mV (n = 14), respectively, and for control myotubes they were -4.78 ± 1.0 and 4.56 ± 0.4 mV (n = 20). Figure 4, C and D, shows the voltage dependence of calcium transients in absolute and normalized values, respectively, to demonstrate that the parameters of activation were not modified (P > 0.05).


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Fig. 4.   Effect of GBP on calcium transients and rate of calcium release. A: typical calcium transients (Delta F) and their first derivatives (delta F/delta t) for control cells at -30, 0, and 50 mV. B: typical calcium transients and their first derivatives for GBP-treated cells at -30, 0, and 50 mV. C: calcium transients (Delta F/F) are plotted as a function of membrane potential for control (; n = 22) and GBP-treated cells (black-triangle; n = 15). No significant changes were detected in the voltage dependence or maximum amplitude of the calcium transients. D: calcium transients normalized to maximum fluorescence (Delta F/Fmax) as a function of membrane potential. No alterations were detected with GBP treatment. E: first derivative of the calcium transient is plotted as a function of membrane potential in control (; n = 22) and GBP-treated cells (black-triangle; n = 15). The rate of rise of the Ca2+ transient was significantly reduced in GBP-treated cells at membrane potentials of 10-60 mV (*P < 0.05).

We further analyzed calcium transients by calculating the derivative of the records. Under our recording conditions, the derivative provides an accurate approximation of the rate of release in skeletal myotubes (27). It has been shown that myotubes have a negligible calcium removal flux (14), which can be neglected if a large concentration of calcium buffers is present in the sarcoplasm. In our experiments, the intracellular solution contained 10 mM EGTA and 0.2 mM Fluo-3, which represent a high concentration of calcium buffers in these cells. Moreover, the rate of calcium release from the SR is much faster than the rate of removal (25). Derivative traces are shown below the corresponding calcium transient records in Fig. 4, A and B. As seen from these traces, GBP caused a reduction in the rate of rise of calcium transients, reflecting a smaller rate of release, compared with control myotubes. The graph in Fig. 4E shows that the reduction in the rate of rise of the transient was significant at membrane potentials from 10 to 60 mV. These data demonstrate that, despite the increase in Qr, GBP did not affect the maximum amplitude or the voltage dependence of the transients but caused a significant decrease in the rate of calcium release.

Effect of GBP on ICa,L and conductance. GBP caused a small but significant reduction of ICa,L at membrane potentials between 20 and 40 mV compared with control cells, as shown in Fig. 5. Figure 5A shows typical records of ICa,L in untreated and GBP-treated myotubes at different membrane potentials. The conductance of the calcium channel was calculated by fitting the ICa,L data to a Boltzmann equation: I(V) = GmaxL · (V - Vr)/[1 + exp(VL - V)/kL], where I(V) is the maximum ICa,L at a given test potential, GmaxL is the maximum L-type channel conductance, Vr is the reversal potential for calcium, V is the test potential, VL is the half-maximal activation potential for the L-type channel, and kL is the slope factor. Comparison of the conductance-voltage (G-V) relationships is shown in Fig. 5B. As demonstrated, conductance values were significantly decreased in GBP-treated cells at positive membrane potentials compared with control cells. However, the reduction of calcium conductance was not accompanied by a significant change of the activation parameters, as shown by the graph in Fig. 5C. The average values of the activation parameters were as follows: in control myotubes, VL = 9.4 ± 1.0 mV, kL = 4.3 ± 0.2 mV, and Vr = 79.2 ± 1.4 mV (n = 41); in GBP-treated myotubes, VL = 10.5 ± 1.5 mV, kL = 4.6 ± 0.5 mV, and Vr = 80.0 ± 1.3 mV (n = 24) (P > 0.05).


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Fig. 5.   Effect of GBP on calcium conductance and activation kinetics. A: ICa,L records obtained from untreated (control) and GBP-treated myotubes at the test potentials indicated between the traces. B: absolute conductance was significantly reduced in GBP-treated cells (black-triangle; n = 24) compared with control cells (; n = 41) at membrane potentials (20-40 mV) (*P < 0.05). C: normalized conductance (G/Gmax) plotted as a function of membrane potential shows no significant changes in activation parameters with GBP treatment (black-triangle; n = 24) compared with control cells (; n = 41). D: examples of the fitting of ICa,L records to a second-order exponential function. The fitted curves are superimposed on currents obtained at 30 mV. E: ICa,L activation is shown for slow (tau slow) and fast (tau fast) time constants of activation for GBP-treated and control myotubes. No significant differences were detected in tau slow and tau fast with GBP treatment.

To examine the time course of activation of ICa,L, the rising phase of the traces was fitted to a second-order exponential function. Because the activation of ICa,L mediated by DHPRs in skeletal myotubes is voltage independent (8), the analysis was performed at the membrane potential where the current was maximal for each cell. Figure 5D shows an example of the curve fitting superimposed on original ICa,L records for a control and a GBP-treated myotube. In both cases, the curves entirely overlapped the ICa,L traces. We found that neither the slow (tau slow) nor the fast time constants (tau fast) of activation were significantly different between the control and GBP-treated myotubes (Fig. 5E). The average values of tau slow were 57.6 ± 1.76 and 54.93 ± 1.67 ms, and for tau fast the values were 6.65 ± 0.42 and 7.66 ± 0.86 ms, for control (n = 41) and GBP-treated (n = 24) myotubes, respectively.

Nifedipine blocks enhancement of Qr by GBP. Because GBP increased Qr while decreasing both the rate of calcium release and ICa,L, it could be argued that a voltage-sensitive membrane protein other than the DHPR produced the additional charge movement recorded in GBP-treated cells. We tested this hypothesis by using nifedipine, a specific DHPR blocker. Recordings of Qr in the absence and the presence of 10 µM nifedipine were obtained from control and GBP-treated myotubes, as shown in Fig. 6A. On average, nifedipine decreased the maximum Qr to a larger extent in GBP-treated myotubes (60 ± 3%, n = 16) compared with the reduction in untreated myotubes (25 ± 3.5%; n = 25). Thus the application of nifedipine equalized the amount of maximum Qr in control (3.9 ± 0.3 nC/µF; n = 25) and GBP-treated myotubes (3.6 ± 0.3 nC/µF; n = 16), as shown in Fig. 6B. The percent reduction in Qr by nifedipine in control myotubes is consistent with other studies. For example, Strube et al. (33) showed a 33% reduction in charge movement (Qt) from developing skeletal muscle. The decrease of Qr in myotubes treated with GBP and nifedipine thus indicates that the extra charge recorded from GBP-treated myotubes is indeed generated by the DHPR.


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Fig. 6.   Effect of nifedipine (10 µM) on charge movement density. A: Qr records obtained from representative untreated (control) and GBP-treated myotubes at the membrane potentials indicated between the traces. Solid lines correspond to Qr before the application of nifedipine to the extracellular solution. Nifedipine caused a larger reduction of Qr in GBP-treated myotubes. B: Qr density is plotted as a function of membrane potential in control cells treated with nifedipine (10 µM) (black-down-triangle ; n = 25) and GBP (50 µM)-treated cells in the presence of 10 µM nifedipine (; n = 16). No significant differences were detected between control and GBP-treated cells with nifedipine C: Qr normalized to its maximum demonstrates no alteration in voltage dependence between control cells treated with nifedipine (black-down-triangle ) and GBP-treated cells in the presence of nifedipine ().

Figure 6B shows the Qr-voltage relationships obtained for untreated and GBP-treated myotubes. The amount of Qr elicited at each membrane potential in the presence of nifedipine was similar in both groups. Figure 6C shows the data normalized to the maximum Qr as a function of membrane potential. The voltage dependence of Qr was unaltered by nifedipine. The average values of V1/2 and k for myotubes treated with nifedipine alone were -12.8 ± 1.4 and 11.6 ± 0.4 mV (n = 25), respectively. For myotubes treated with GBP and nifedipine, the average values were V1/2 = -10.41 ± 2.0 and k = 11.8 ± 0.7 mV (n = 16).

GBP decreases the effectiveness of the DHPR in E-C coupling. To have an objective measure of the dissociation of Qr and calcium release and ICa,L caused by GBP, we compared the rate of release and conductance as a function of Qr. These data are shown in Fig. 7 for control and GBP-treated myotubes. As shown, GBP shifted the curves to the right, which indicates that more immobilization-resistant charge is needed to attain similar levels of rate of release or conductance. The amount of Qr required for the same value of rate of release or conductance was roughly two times greater in the presence of GBP than the amount of Qr required in control myotubes. Moreover, in GBP-treated myotubes, the maximum rate of release did not reach control levels. Thus this analysis indicates that GBP decreased the effectiveness of the DHPR as a voltage sensor necessary for calcium release and as a calcium channel.


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Fig. 7.   Relationships between calcium release and calcium conductance with Qr. A: rate of calcium release from the sarcoplasmic reticulum (delta F/delta t) is plotted as a function of resistant charge movement (Qr). GBP-treated cells (black-triangle) demonstrated a lower rate of calcium release with greater amounts of charge moved compared with control cells (). B: calcium channel conductance (G) is plotted as a function of charge movement (Qr). GBP-treated cells (black-triangle) demonstrated a twofold increase in charge movement relative to similar conductance compared with control cells ().


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We report a novel regulatory effect of GBP on the E-C coupling mechanism in skeletal muscle. The effect is characterized by dissociation of Qr from calcium release and ICa,L. Two lines of evidence indicate that GBP exerted its effects by acting on the DHPR. First, GBP did not modify the midpoint potential of activation (V1/2) or the slope (k) of the curves of any of the events produced by activation of the DHPR. Second, the increase of Qr caused by GBP was antagonized by nifedipine, such that the amount of Qr remaining in the presence of GBP and nifedipine was similar to that in myotubes treated with nifedipine alone.

Based on the voltage dependence and time course of activation of Qr, calcium release, and ICa,L, a sequential scheme of the conformational changes in the DHPR has been proposed (16). This model would predict that any agent that modifies Qr would have a similar effect on calcium release and ICa,L. However, other studies have shown that perchlorate and nifedipine may also have differential effects on the events generated by the DHPR (6, 22). The target proteins involved in the actions of perchlorate are thought to be the alpha 11.1-subunit of the DHPR and/or the RyR1 (24), whereas nifedipine acts on the alpha 11.1-subunit (39), where its binding site is localized (26). In this respect, the effect of GBP on E-C coupling is unique because the alpha 2/delta -subunit has been identified as the GBP receptor (17), and its binding site has been localized on the extracellular portion of the alpha 2/delta 1-subunit of the DHPR (4, 38).

The remote possibility exists that the effect of GBP on E-C coupling is unrelated to its interaction with the alpha 2/delta 1-subunit and that GBP may also bind to the alpha 11.1 subunit (in addition to the alpha 2/delta 1-subunit) because of the concentration of the drug used in these experiments (50 µM). However, calculation of the slope of the curve fitted to the data in Fig. 1 resulted in a value of 1.05, indicating that GBP bound to a single population of sites in myotubes. This result agrees with previous studies by Gee et al. (17) using rat muscle membranes, where GBP also bound to one population of sites. Furthermore, radioligand binding and immunoblotting studies of fractionated L-type calcium channels isolated from skeletal muscle determined that GBP bound to the alpha 2/delta 1-subunit even at high concentrations. Thus it is conceivable that the effects of GBP on E-C coupling we have elucidated are mediated by the alpha 2/delta 1-subunit.

Our results suggest that GBP binding to the alpha 2/delta 1-subunit would affect the conformational changes experienced by the alpha 11.1-subunit during DHPR activation. Only a few amino acids of the alpha 2/delta 1 protein are found on the cytoplasmic side of the membrane, the remainder being located either within the membrane or on the extracellular side (18). For this reason, it is attractive to hypothesize that the GBP-alpha 2/delta 1-subunit complex acts on the transmembrane repeats and perhaps the outer portion of the alpha 11.1-subunit. Initially the interaction between the GBP-alpha 2/delta 1-subunit complex and alpha 11.1 would favor the movement of the repeats resulting in a larger Qr. Subsequently, the complex would prevent the fast and complete change of repeats II and III (and the II-III cytoplasmic loop). This would result in slower activation of the RyR1 and an attenuated rate of calcium release. Finally, the decrease in ICa,L amplitude may be explained by a blocking effect of the GBP-alpha 2/delta 1-subunit complex. The block may occur on the extracellular side of the calcium channel because it has been shown that the external portion of the alpha 2/delta 1-subunit is required for current stimulation (18). Although repeat I in the alpha 11.1-subunit is thought to set the activation rate of the ICa,L (35), we do not believe the complex is modifying the movement of repeat I because the time constants of activation of the current were similar in untreated and GBP-treated myotubes.

Whether the GBP-binding properties to the alpha 2/delta 1-subunit in skeletal myotubes are independent of the presence or absence of a prepulse cannot be determined from our experiments. As Adams et al. (1) have previously shown and we mentioned earlier, we need to use a prepulse to record the activity of the DHPR separate from other voltage-dependent proteins in skeletal myotubes. However, we have previously shown that the effect of GBP on neuronal calcium channels is noticed only after the application of a prepulse (2).

In recent studies using expression systems, other investigators have shown that charge movement arising from the cardiac alpha 11.2-subunit is modulated by coexpression of the alpha 2/delta -subunit (3, 29, 31). Although the experimental systems between those reports and this manuscript are different and calcium transients were not studied earlier, they agree with our finding that the alpha 2/delta 1-subunit may control the gating of the alpha 11.1-peptide. However, we further consider that the in situ function of the alpha 2/delta 1-subunit may be to effectively couple gating current with calcium release and channel opening.

Taken together the data presented in this work suggest the presence of several closed states before the DHPR is fully activated and that these states can be affected by GBP. Our results are also the first to provide information about the involvement of the alpha 2/delta 1-subunit in skeletal muscle E-C coupling.


    ACKNOWLEDGEMENTS

We thank Dr. Tord D. Alden for help with the statistical analysis and Tanvi M. Shah for laboratory assistance.


    FOOTNOTES

This work was supported by National Science Foundation Grant IBN-9733570 (to J. García). K. Alden was partially supported by National Institute of Diabetes and Digestive and Kidney Diseases Training Grant T32 DK-07739-02.

Address for reprint requests and other correspondence: J. García, Dept. of Physiology and Biophysics, Univ. of Illinois at Chicago College of Medicine, 900 S. Ashland Ave., M/C 902, Chicago, IL 60607 (E-mail: garmar{at}uic.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

May 15, 2002;10.1152/ajpcell.00004.2002

Received 4 January 2002; accepted in final form 14 May 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Adams, BA, Tanabe T, Mikami A, Numa S, and Beam KG. Intramembrane charge movement restored in dysgenic skeletal muscle by injection of dihydropyridine receptor cDNAs. Nature 346: 569-572, 1990[ISI][Medline].

2.   Alden, KJ, and García J. Differential effect of gabapentin on neuronal and muscle calcium currents. J Pharmacol Exp Ther 297: 727-735, 2001[Abstract/Free Full Text].

3.   Bangalore, R, Mehrke K, Gingrich K, Hofmann F, and Kass RS. Influence of L-type Ca channel alpha 2delta -subunit on ionic and gating current in transiently transfected HEK 293 cells. Am J Physiol Heart Circ Physiol 270: H1521-H1528, 1996[Abstract/Free Full Text].

4.   Brown, JP, and Gee NS. Cloning and deletion mutagenesis of the alpha 2delta calcium channel subunit from porcine cerebral cortex. Expression of a soluble form of the protein that retains [3H]gabapentin binding activity. J Biol Chem 273: 25458-25465, 1998[Abstract/Free Full Text].

5.   Chandler, WK, Rakowski RF, and Schneider MF. Effects of glycerol treatment and maintained depolarization on charge movement in skeletal muscle. J Physiol 254: 285-316, 1976[Abstract].

6.   Csernoch, L, Kovacs L, and Szucs G. Perchlorate and the relationship between charge movement and contractile activation in frog skeletal muscle fibres. J Physiol 390: 213-227, 1987[Abstract].

7.   Dietze, B, Henke J, Eichinger HM, Lehmann-Horn F, and Melzer W. Malignant hyperthermia mutation Arg615Cys in the porcine ryanodine receptor alters voltage dependence of Ca2+ release. J Physiol 526: 507-514, 2000[Abstract/Free Full Text].

8.   Dirksen, RT, and Beam KG. Single calcium channel behavior in native skeletal muscle. J Gen Physiol 105: 227-247, 1995[Abstract].

9.   Eberst, R, Dai S, Klugbauer N, and Hofmann F. Identification and functional characterization of a calcium channel gamma subunit. Pflügers Arch 433: 633-637, 1997[ISI][Medline].

10.   El-Hayek, R, Antoniu B, Wang J, Hamilton SL, and Ikemoto N. Identification of calcium release-triggering and blocking regions of the II-III loop of the skeletal muscle dihydropyridine receptor. J Biol Chem 270: 22116-22118, 1995[Abstract/Free Full Text].

11.   Feldmeyer, D, and Lüttgau HC. The effect of perchlorate on Ca currents and mechanical force in skeletal muscle fibres (Abstract). Pflügers Arch 411: R190, 1988.

12.   Feldmeyer, D, Melzer W, and Pohl B. Effects of gallopamil on calcium release and intramembrane charge movements in frog skeletal muscle fibres. J Physiol 421: 343-362, 1990[Abstract].

13.   García, J, Avila-Sakar AJ, and Stefani E. Differential effects of ryanodine and tetracaine on charge movement and calcium transients in frog skeletal muscle. J Physiol 440: 403-417, 1991[Abstract].

14.   García, J, and Beam KG. Measurement of calcium transients and slow calcium current in myotubes. J Gen Physiol 103: 107-123, 1994[Abstract].

15.   García, J, Pizarro G, Rios E, and Stefani E. Effect of the calcium buffer EGTA on the "hump" component of charge movement in skeletal muscle. J Gen Physiol 97: 885-896, 1991[Abstract].

16.   García, J, Tanabe T, and Beam KG. Relationship of calcium transients to calcium currents and charge movements in myotubes expressing skeletal and cardiac dihydropyridine receptors. J Gen Physiol 103: 125-147, 1994[Abstract].

17.   Gee, NS, Brown JP, Dissanayake VU, Offord J, Thurlow R, and Woodruff GN. The novel anticonvulsant drug, gabapentin (Neurontin), binds to the alpha 2delta subunit of a calcium channel. J Biol Chem 271: 5768-5776, 1996[Abstract/Free Full Text].

18.   Gurnett, CA, De Waard M, and Campbell KP. Dual function of the voltage-dependent Ca2+ channel alpha 2delta subunit in current stimulation and subunit interaction. Neuron 16: 431-440, 1996[ISI][Medline].

19.   Gyorke, S, and Palade P. Effects of perchlorate on excitation-contraction coupling in frog and crayfish skeletal muscle. J Physiol 456: 443-451, 1992[Abstract].

20.   Hamill, OP, Marty A, Neher E, Sakmann B, and Sigworth FJ. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflügers Arch 391: 85-100, 1981[ISI][Medline].

21.   Huang, CL. Charge movements in intact amphibian skeletal muscle fibres in the presence of cardiac glycosides. J Physiol 532: 509-523, 2001[Abstract/Free Full Text].

22.   Kitamura, N, Ohta T, Ito S, and Nakazato Y. Effects of nifedipine and Bay K 8644 on contractile activities in single skeletal muscle fibers of the frog. Eur J Pharmacol 256: 169-176, 1994[ISI][Medline].

23.   Leong, P, and MacLennan DH. A 37-amino acid sequence in the skeletal muscle ryanodine receptor interacts with the cytoplasmic loop between domains II and III in the skeletal muscle dihydropyridine receptor. J Biol Chem 273: 7791-7794, 1998[Abstract/Free Full Text].

24.   Ma, J, Anderson K, Shirokov R, Levis R, Gonzalez A, Karhanek M, Hosey MM, Meissner G, and Rios E. Effects of perchlorate on the molecules of excitation-contraction coupling of skeletal and cardiac muscle. J Gen Physiol 102: 423-448, 1993[Abstract].

25.   Melzer, W, Rios E, and Schneider MF. Time course of calcium release and removal in skeletal muscle fibers. Biophys J 45: 637-641, 1984[Abstract].

26.   Mitterdorfer, J, Wang Z, Sinnegger MJ, Hering S, Striessnig J, Grabner M, and Glossmann H. Two amino acid residues in the IIIS5 segment of L-type calcium channels differentially contribute to 1,4-dihydropyridine sensitivity. J Biol Chem 271: 30330-30335, 1996[Abstract/Free Full Text].

27.   Nabhani, T, Zhu X, Simeoni I, Sorrentino V, Valdivia HH, and García J. Imperatoxin A enhances Ca2+ release in developing skeletal muscle containing ryanodine receptor type 3. Biophys J 82: 1319-1328, 2002[Abstract/Free Full Text].

28.   Nakai, J, Dirksen RT, Nguyen HT, Pessah IN, Beam KG, and Allen PD. Enhanced dihydropyridine receptor channel activity in the presence of ryanodine receptor. Nature 380: 72-75, 1996[ISI][Medline].

29.   Platano, D, Qin N, Noceti F, Birnbaumer L, Stefani E, and Olcese R. Expression of the alpha 2delta subunit interferes with prepulse facilitation in cardiac L-type calcium channels. Biophys J 78: 2959-2972, 2000[Abstract/Free Full Text].

30.   Rios, E, and Brum G. Involvement of dihydropyridine receptors in excitation-contraction coupling in skeletal muscle. Nature 325: 717-720, 1987[ISI][Medline].

31.   Shirokov, R, Ferreira G, Yi J, and Ríos E. Inactivation of gating currents of L-type calcium channels. Specific role of the alpha 2delta subunit. J Gen Physiol 111: 807-823, 1998[Abstract/Free Full Text].

32.   Stefani, A, Spadoni F, and Bernardi G. Gabapentin inhibits calcium currents in isolated rat brain neurons. Neuropharmacology 37: 83-91, 1998[ISI][Medline].

33.   Strube, C, Bournaud R, Inoue I, and Shimahara T. Intramembrane charge movement in developing skeletal muscle cells from fetal mice. Pflügers Arch 421: 572-577, 1992[ISI][Medline].

34.   Suman-Chauhan, N, Webdale L, Hill DR, and Woodruff GN. Characterisation of [3H]gabapentin binding to a novel site in rat brain: homogenate binding studies. Eur J Pharmacol 244: 293-301, 1993[Medline].

35.   Tanabe, T, Adams BA, Numa S, and Beam KG. Repeat I of the dihydropyridine receptor is critical in determining calcium channel activation kinetics. Nature 352: 800-803, 1991[ISI][Medline].

36.   Tanabe, T, Beam KG, Adams BA, Niidome T, and Numa S. Regions of the skeletal muscle dihydropyridine receptor critical for excitation-contraction coupling. Nature 346: 567-569, 1990[ISI][Medline].

37.   Wamil, AW, and McLean MJ. Limitation by gabapentin of high frequency action potential firing by mouse central neurons in cell culture. Epilepsy Res 17: 1-11, 1994[ISI][Medline].

38.   Wang, M, Offord J, Oxender DL, and Su TZ. Structural requirement of the calcium-channel subunit alpha 2delta for gabapentin binding. Biochem J 342: 313-320, 1999[ISI][Medline].

39.   Weigl, LG, Hohenegger M, and Kress HG. Dihydropyridine-induced Ca2+ release from ryanodine-sensitive Ca2+ pools in human skeletal muscle cells. J Physiol 525: 461-469, 2000[Abstract/Free Full Text].

40.   Witcher, DR, De Waard M, Sakamoto J, Franzini-Armstrong C, Pragnell M, Kahl SD, and Campbell KP. Subunit identification and reconstitution of the N-type Ca2+ channel complex purified from brain. Science 261: 486-489, 1993[ISI][Medline].


Am J Physiol Cell Physiol 283(3):C941-C949
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