Effects of local anesthetics on Na+ channels containing the equine hyperkalemic periodic paralysis mutation

Rajan L. Sah, Robert G. Tsushima, and Peter H. Backx

Departments of Medicine and Physiology, University of Toronto, Toronto, Ontario, Canada M5G 1L7

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

We examined the ability of local anesthetics to correct altered inactivation properties of rat skeletal muscle Na+ channels containing the equine hyperkalemic periodic paralysis (eqHPP) mutation when expressed in Xenopus oocytes. Increased time constants of current decay in eqHPP channels compared with wild-type channels were restored by 1 mM benzocaine but were not altered by lidocaine or mexiletine. Inactivation curves, which were determined by measuring the dependence of the relative peak current amplitude after depolarization to -10 mV on conditioning prepulse voltages, could be shifted in eqHPP channels back toward that observed for wild-type (WT) channels using selected concentrations of benzocaine, lidocaine, and mexiletine. Recovery from inactivation at -80 mV (50-ms conditioning pulse) in eqHPP channels followed a monoexponential time course and was markedly accelerated compared with wild-type channels (tau WT = 10.8 ± 0.9 ms; tau eqHPP = 2.9 ± 0.4 ms). Benzocaine slowed the time course of recovery (tau eqHPP,ben = 9.6 ± 0.4 ms at 1 mM) in a concentration-dependent manner. In contrast, the recovery from inactivation with lidocaine and mexiletine had a fast component (tau fast,lid = 3.2 ± 0.2 ms; tau fast,mex = 3.1 ± 0.2 ms), which was identical to the recovery in eqHPP channels without drug, and a slow component (tau slow,lid = 1,688 ± 180 ms; tau slow,mex = 2,323 ± 328 ms). The time constant of the slow component of the recovery from inactivation was independent of the drug concentration, whereas the fraction of current recovering slowly depended on drug concentrations and conditioning pulse durations. Our results show that local anesthetics are generally incapable of fully restoring normal WT behavior in inactivation-deficient eqHPP channels.

sodium channel; inactivation

    INTRODUCTION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

NORMAL ACTION POTENTIALS in nerve, heart, and skeletal muscle cells depend critically on a transient influx of Na+ through voltage-gated Na+ channels. After membrane depolarization, Na+ channels open or "activate" and subsequently rapidly inactivate. Channel openings are not possible again until channels "recover" from inactivation, a process requiring several milliseconds after membrane repolarization. A number of mutations of the alpha -subunit of skeletal and cardiac Na+ channels disrupt inactivation, thereby causing disease. For example, several mutations of the human skeletal muscle gene (SCN4A) have been associated with heritable muscle diseases including hyperkalemic periodic paralysis (HPP), paramyotonia congenita, and Na+ channel myotonia (18, 33, 40, 42), whereas mutations in cardiac Na+ channel genes result in a form of the long Q-T syndrome (7, 28, 48). Functional studies of muscle-related Na+ channel mutations have shown that many of these mutations disrupt normal Na+ channel inactivation, either alone or in combination with other functional defects (7, 9, 13, 16, 18, 32, 35, 51).

Local anesthetic and antiarrhythmic drugs, such as lidocaine, act by blocking Na+ flux through voltage-gated Na+ channels and have been shown to bind preferentially to inactivated conformation of the cardiac Na+ channel (3-6, 8, 17, 24, 36, 47, 49). Therefore, local anesthetic agents might be expected to reconstitute inactivation in Na+ channels with defective inactivation (18, 25). Within the "ball and chain" paradigm of Na+ channel inactivation, preferential binding of local anesthetics to the inactivated state appears to stabilize the interaction of the inactivation gate with the receptor, thereby locking channels into the inactivated conformation (3-6, 8, 18, 22, 24, 47). This mechanism of local anesthetic action could explain the clinical use of these agents in patients with HPP (18, 25).

In the present study, we examined the effects of local anesthetics on heterologously expressed rat skeletal muscle Na+ channels containing the equine HPP (eqHPP) mutation (F1412L), located in the third transmembrane spanning region (S3) of domain IV (43). The equine mutation is characterized by episodes of myotonia, weakness, and ultimately flaccid paralysis in association with elevated serum K+ levels (39). We and others (11, 23) have shown that the eqHPP mutation disrupts Na+ channel inactivation. Specifically, heterologously expressed rat skeletal muscle Na+ channels containing the F1412L mutation showed slowed inactivation kinetics, a rightwardly shifted steady-state inactivation curve, and accelerated rates of recovery from inactivation compared with wild-type channels (23). Na+ channels recorded from skeletal muscle myocytes from horses with eqHPP also showed evidence for altered modal behavior (11). The objectives of our studies were to examine whether the local anesthetics could promote inactivation and whether the application of these agents could, under the appropriate conditions, restore normal Na+ channel gating properties.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Site-directed mutagenesis. Site-directed mutagenesis of the rat skeletal muscle Na+ channel (rSkM1) (46) was performed to create the F1412L constructs. The mutations were introduced into a 2.5-kb Sph I-Kpn I cassette subcloned into pGEM7f+ (Promega, Madison, WI) using the oligonucleotide containing the appropriate base substitution. Phenotypic selection was performed using the Kunkel method (30), and the mutation was confirmed by dideoxy sequencing (45). The cassette was subcloned into rSkM1 in pGW1H (British Biolabs, Oxford, UK) for intranuclear injection of cDNA.

Expression of Na+ channels in Xenopus oocytes. Oocytes were removed from adult female Xenopus laevis frogs (NASCO, Fort Atkinson, WI; XENOPUS, Ann Arbor, MI) anesthetized by immersion for 10-25 min in a 0.25% solution of tricaine (Sigma Chemical, St. Louis, MO) in tap water. Oocytes were digested with 2 mg/ml collagenase (type 1A, Sigma) in OR-2 containing (in mM) 88 NaCl, 2 KCl, 1 MgCl2, and 5 mM HEPES (pH 7.6). Oocytes were stored at room temperature in ND96 containing (in mM) 96 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2, and 5 HEPES (pH 7.6) supplemented with 50 µg/ml gentamicin, 5 mM pyruvate, and 0.5 mM theophylline. The nuclei of oocytes at stages IV-VI were coinjected with 0.2 ng of alpha -subunit cDNA and 4.0 ng of beta 1-subunit cDNA (26). The 20-fold excess of beta -subunits compared with alpha -subunits was used to minimize the altered gating properties of Na+ channels alpha -subunits observed when expressed in oocytes (4, 12, 27). Injected oocytes were incubated at 22°C for 24-48 h before recording.

Electrophysiological recording. Whole cell currents were recorded at room temperature (21-23°C) using two-electrode voltage-clamp techniques (Oocyte Clamp OC-725A, Warner Instruments, Hampden, CT) with 3 M KCl in the pipette and a bath solution (ND96). Electrode pipettes were fabricated from 1.2-mm-OD thin-walled borosilicate glass (TW120F-6, World Precision Instruments, Sarasota, FL) pulled on a Sutter puller (model P-87). Pipette tips were plugged with 1% agarose (in 3 M KCl) and had a final resistance of 1-4 MOmega for the voltage-sensing electrode and <2 MOmega for the current-passing electrode. Leak subtraction was accomplished using a P/8 protocol from a holding potential of -120 mV. Currents were filtered at 2 kHz and digitized at 10 kHz using an IBM-compatible computer, analog-to-digital interface (Warner model PP-50 Lab1), and custom acquisition software. To minimize difficulties associated with adequately voltage-clamping oocytes which expressed large numbers of channels, whole cell recordings were limited to oocytes expressing <5 µA of peak current. The local anesthetics lidocaine (Sigma), mexiletine (Boehringer-Ingelheim, Ingelheim, Germany), and benzocaine (ICN Biomedicals, Mississauga, Canada) were introduced at the desired concentration in ND96 to the bath by perfusion with at least 30 ml (bath volume approx 0.6 ml). After total bath exchange (requiring <3 min), the drug was allowed to equilibrate with the oocyte for a minimum of 6 min before recording.

Voltage protocols. In the absence of drug, steady-state fast-inactivation curves were constructed by plotting the relative peak current amplitude elicited in response to "test" depolarizations (to -10 mV for 50 ms) as a function of conditioning prepulse voltages (ranging from -120 to -10 mV for 50 ms) that were applied immediately before the test pulse. Previous studies in rSkM1 channels have established that 50-ms prepulses are sufficiently long to allow equilibration between the closed and inactivated states of the channel (19). Using similar voltage protocols, we also examined the effects of local anesthetics on the relative current in test pulses as a function of conditioning prepulse voltage (from -120 to -10 mV). The prepulse durations in the presence of drug were either 50 or 500 ms in length. These measurements allow the assessment of the changes in the voltage and time dependence of channels entering into the inactivation state produced by local anesthetics. In these experiments, the voltage pulses were applied every 10 s to ensure complete recovery from inactivation between recordings.

Recovery from inactivation was assessed using a two-pulse protocol, in which (conditioning) depolarizing pulses to -10 mV for either 50 or 500 ms were followed by repolarization to -80 mV (recovery potential) for a variable duration, and this was immediately followed by a (test) pulse to -10 mV (50 ms). The test current was normalized to the conditioning current and plotted as a function of the duration at the recovery potential. A repetition frequency of 0.1 Hz was used for the recovery from inactivation protocols to ensure complete recovery of channels between recordings. Previous studies have established that 50-ms prepulses are sufficient to allow equilibration between closed and fast inactivated states of rSkM1 channels (19). In the presence of polar anesthetics like lidocaine and mexiletine, equilibration of drug binding to the inactivated channels is dose dependent but is largely complete within 500 ms in both expressed cardiac and skeletal muscle Na+ channels with -20 mV prepulses in the presence of 100 µM lidocaine (47).

Statistics and curve fitting. The Marquardt-Levenberg algorithm in conjunction with a nonlinear least-squares procedure was used to fit the functions shown below to the experimental data. Data from experiments involving voltage protocols using conditioning prepulses (i.e., relative peak current measured in a test pulse as a function of conditioning prepulse voltage) were fit with Boltzmann functions (24)
<IT>h</IT><SUB>∞</SUB> = 1/{1 + exp[(<IT>V</IT> − <IT>V</IT><SUB>½</SUB>)/<IT>k</IT>]}
where V is the step or conditioning potential, V1/2 is the voltage midpoint of the function, and k is the slope factor. In the remainder of the paper, the curves generated from these fits are referred to as "steady-state inactivation" curves in the absence of drug (17) and simple "inactivation curves" when drug is present. This distinction emphasizes the possibility that, with conditioning prepulses of short duration used in our studies, incomplete equilibration of drug binding to inactivated channels occurs.

Monoexponential or biexponential functions were used to fit recovery from inactivation data and kinetic decay of the whole cell currents after depolarization. For fits to the recovery from inactivation data, we used the following biexponential function
<IT>I</IT>/<IT>I</IT><SUB>0</SUB> = <IT>A</IT><SUB>fast</SUB>[1 − exp(−<IT>t</IT>/&tgr;<SUB>fast</SUB>)] + <IT>A</IT><SUB>slow</SUB>[1 − exp(−<IT>t</IT>/&tgr;<SUB>slow</SUB>)]
where Afast and Aslow are the amplitudes of the fast and slow components of the recovery, respectively; tau fast and tau slow are the time constants for the fast and slow components for recovery, respectively; and t is the time spent at the recovery potential (see above). When the recovery data were fit with a monoexponential, Afast = Aslow = A and tau fast = tau slow = tau . For the recovery from inactivation, a time delay was commonly observed, particularly with benzocaine. Therefore, when fitting these data, we routinely included a time delay to account for this effect as described previously (37). For these fits, the F-statistic and F-distribution (P < 0.05) were used to assess whether biexponential fits to the data gave significantly superior fits compared with the monoexponential fits (21).

In the experiments using mexiletine and lidocaine, the amplitude of the slow component of recovery from inactivation (i.e., Aslow) was fit with the binding isotherm equation
<IT>A</IT><SUB>slow</SUB> = [LA]<SUP><IT>n</IT></SUP>/{[LA]<SUP><IT>n</IT></SUP> + (IC<SUB>50,LA</SUB>)<SUP><IT>n</IT></SUP>}
where [LA] is the concentration of either lidocaine or mexiletine, n is the Hill coefficient, and IC50,LA is the concentration of drug resulting in 50% of the inactivated channels being drug bound.

Combined data are presented as means ± SE. Statistical significance was determined using an unpaired Student's t-test (21) and a confidence limit of 95%.

    RESULTS
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Abstract
Introduction
Materials & Methods
Results
Discussion
References

Phenotypic alteration of eqHPP channels. The electrophysiological changes associated with eqHPP channels are illustrated in Fig. 1A, showing scaled raw Na+ current traces recorded from wild-type (WT) and eqHPP channels after step depolarizations to -10 mV from a holding potential of -120 mV. Notice that WT Na+ currents decay more rapidly than eqHPP channels: tau WT = 0.97 ± 0.08 ms (n = 6) vs. tau eqHPP = 2.14 ± 0.07 ms (n = 15). Slowed current decay in eqHPP channels compared with WT can originate from either slowed entry into the inactivated state (2, 23, 32, 34, 38) or voltage shifts in channel activation (14, 16, 24). Figure 1B shows that there was a significant rightward shift in the steady-state inactivation curve of eqHPP mutant channels (solid squares) by ~6 mV relative to WT channels (open squares) (V1/2,eqHPP -47.6 ± 0.6 mV, n = 10 vs. V1/2,WT = -53.1 ± 0.4 mV, n = 7; P < 0.05), suggesting that the eqHPP mutation has altered channel inactivation (23). This is further supported by the observations summarized in Fig. 1C, illustrating that the recovery from inactivation (after 50-ms conditioning pulse) is accelerated more than threefold for eqHPP mutant channels compared with WT channels at -80 mV (i.e., tau eqHPP = 2.9 ± 0.4 ms, n = 7 vs. tau WT = 10.8 ± 0.9 ms, n = 7; P < 0.01).


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Fig. 1.   A: whole cell currents recorded from wild-type (WT) and equine hyperkalemic periodic paralysis (eqHPP) mutant Na+ channels expressed in oocytes. Currents were evoked by a 50-ms voltage step to -10 mV from a holding potential of -120 mV. Currents have been normalized to have same peak amplitudes; peak inward current amplitude was 3.4 µA for eqHPP trace and 2.9 µA for WT trace. eqHPP current displays a slowed rate of decay relative to decay of WT current (tau eqHPP = 2.14 ± 0.07 ms, n = 15; tau WT = 0.97 ± 0.08 ms, n = 6). B: steady-state inactivation curves constructed by plotting relative current magnitude as a function of 50-ms prepulse voltages for WT (; n = 7) and eqHPP (; n = 10) channels. Inactivation was induced by 50-ms conditioning prepulses from -100 to -10 mV in 5-mV increments. Extent of inactivation was then assessed by a step depolarization to -10 mV. C: recovery from inactivation of WT (open circle ; n = 9) and eqHPP mutant (; n = 7) was assayed using a 2-pulse protocol at a recovery potential and holding potential of -80 mV. On average, eqHPP mutant recovers significantly faster from inactivation (tau eqHPP = 2.9 ± 0.4 ms) than WT channel (tau WT = 10.8 ± 0.9 ms) (P < 0.05).

Effect of local anesthetic on inactivation properties of eqHPP. Because local anesthetics selectively bind to the inactivated state of Na+ channels and thereby stabilize the inactivated state (3-6, 8, 17, 22, 24, 36), we tested the effects of lidocaine, mexiletine, and benzocaine on the inactivation properties of eqHPP channels. Figure 2A shows raw current traces recorded in response to 50-ms step depolarizations to -10 mV from a holding potential of -80 mV before and after the application of lidocaine, mexiletine, and benzocaine. At the indicated dosages, lidocaine, mexiletine, and benzocaine reduced the peak current of eqHPP channels. This reduction of peak current is generally referred to as tonic block and probably represents binding of these drugs to the closed state or possibly rapid block of the open state (5, 24). Normalization of current traces in the presence of the drug to current traces in the absence of drug, as illustrated in Fig. 2B, shows that 600 µM lidocaine and mexiletine do not noticeably accelerate the rate of current decay (tau eqHPP = 2.14 ± 0.07 ms, n = 15; tau lid = 2.29 ± 0.21 ms, n = 5; tau mex = 2.43 ± 0.16 ms, n = 5), whereas 1 mM benzocaine clearly enhances the rate of current decline (tau ben = 1.11 ± 0.06 ms, n = 5; P < 0.05). The dependence of the time constant for whole cell current decline recorded after depolarizations to -10 mV as a function of the concentration of drug is illustrated in Fig. 2C. Lidocaine and mexiletine caused a very modest and statistically insignificant acceleration in the rate of current decay of the Na+ current as a function of the drug concentration (P > 0.05) as previously demonstrated (47). For example, after the application of 600 µM lidocaine, the time constant tau  for decay decreased from 2.33 ± 0.17 to 1.95 ± 0.19 ms. In contrast, the time constant of Na+ current reduction decreased with elevated benzocaine concentrations, asymptotically reaching a limiting value of 1.05 ± 0.03 ms at 1 mM benzocaine. These differences between lidocaine and mexiletine compared with benzocaine likely result from differences in hydrophobicity between these different local anesthetics (8, 17, 24, 41).


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Fig. 2.   A: reduction of peak current of eqHPP mutant in presence of local anesthetics. Currents were evoked by 50-ms step depolarizations to -10 mV from a holding potential of -80 mV. Only first 30 ms are shown. B: current traces from A normalized with respect to peak eqHPP mutant current. Neither 300 µM lidocaine nor 300 µM mexiletine accelerated rate of current decay of eqHPP mutant, whereas 1 mM benzocaine markedly increased rate of decay in this oocyte. C: time constants (tau h) of current decay of eqHPP mutant vs. concentration of lidocaine (left), mexiletine (middle), and benzocaine (right). Lidocaine and mexiletine data were well fit by a linear regression function [r2 (lidocaine) = 0.95; r2 (mexiletine) = 0.95] with a slope that was not significantly different from zero. Benzocaine data were fit to a monoexponential function.

As a result of the ability of benzocaine to enhance the rate of current decline, we speculated that, at the appropriate concentration, this agent might reconstitute WT behavior to eqHPP channel with respect to other aspects of inactivation. Figure 3A shows superimposed current traces of WT channels and of HPP channels plus 1 mM benzocaine after depolarization to -10 mV; the current traces were scaled to match the peak currents. In the presence of 1 mM benzocaine, the current decay of HPP channels measured at -10 mV mimic that of WT channels without benzocaine (tau WT = 0.97 ± 0.08 ms, n = 6 vs. tau ben = 1.11 ± 0.06 ms, n = 5). However, as illustrated in Fig. 3B, the time constants for whole cell current decay become largely voltage independent in the presence of 1 mM benzocaine in eqHPP channels (solid triangles) compared with WT (open circles) or eqHPP (solid squares) channels without drug. As a result, the relationship between the inactivation time constants and voltage in eqHPP channels treated with benzocaine cross over the corresponding relationship in WT channels at around -10 mV. Therefore, although benzocaine does reverse the slowed inactivation rate of eqHPP at high concentrations (i.e., 1 mM), it does not generally confer WT behavior to eqHPP channels at all voltages. In addition, as discussed more fully below but not shown in Fig. 3, the peak current magnitude was reduced by ~50% with this dose of benzocaine (Table 1).


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Fig. 3.   A: superimposed current traces of WT channel and eqHPP mutant in presence of 1 mM benzocaine normalized with respect to peak current. Currents were evoked as described in Fig. 2. Peak current in WT traces was 3.3 µA, and current in benzocaine-treated eqHPP channels was 2.3 µA. eqHPP mutant in presence of 1 mM benzocaine closely approximates current decay of WT channel (tau WT = 0.97 ± 0.08 ms, n = 6; tau eqHPP,ben = 1.11 ± 0.06 ms, n = 5). B: time constants of current decay vs. voltage of WT channels (open circle ; n = 6) and eqHPP mutant channels in absence (; n = 6) and presence of 1 mM benzocaine (black-triangle; n = 5). Notice that little voltage dependence is observed in time constant of current decay of eqHPP mutant channels when 1 mM benzocaine is present.

                              
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Table 1.   Tonic block of eqHPP by local anesthetics

The rightward shift of the steady-state inactivation curve along the voltage axis (Fig. 1B) of eqHPP channels relative to WT channels requires that more positive prepulses are necessary to inactivate the same proportion of eqHPP channels in comparison to WT channels (23). Therefore, we examined the effects of local anesthetics on the relative peak current amplitude recorded during test depolarizations (-10 mV for 50 ms) as a function of the conditioning prepulse voltage having a duration of 50 or 500 ms. The relative peak current amplitudes as a function of prepulse voltage are plotted in Fig. 4 for WT channels (solid circles) and eqHPP channels in the absence (solid squares) and presence of drug (open symbols). The estimated V1/2 values, obtained by fitting the data for these experiments to the Boltzmann equation (see MATERIALS AND METHODS) for these experiments, are summarized in Table 2. With 50-ms conditioning pulses, 600 µM lidocaine (Fig. 4A), 300 µM mexiletine (Fig. 4B), and 30 µM benzocaine (Fig. 4C) shifted the inactivation curves of eqHPP channels sufficiently leftward to cause superposition with the WT steady-state inactivation curves. These results demonstrate that with short conditioning pulses, benzocaine is far more potent than lidocaine or mexiletine in promoting entry into the inactivated state. With longer conditioning prepulses (500 ms), illustrated in Fig. 4, right, 100 µM lidocaine and mexiletine shifted the dependence of the relative current on prepulse voltage for eqHPP sufficiently leftward to approximately overlay the WT curve, whereas 30 µM benzocaine was again similar to WT (Table 2). Thus increasing the conditioning pulse duration from 50 to 500 ms had little effect on benzocaine binding, consistent with rapid equilibration of drug binding to the channel (8, 17, 36, 41). In contrast, prolongation of the conditioning pulse duration in the presence of lidocaine or mexiletine did markedly enhance drug binding as expected for the slow binding kinetics displayed by polar local anesthetics (5, 17, 41, 47).


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Fig. 4.   Effect of local anesthetics on voltage dependence of channel inactivation in response to conditioning prepulses. Relative peak current observed in WT channels (bullet ) and eqHPP channels in absence () and presence (open symbols) of lidocaine (A), mexiletine (B), and benzocaine (C) after conditioning prepulses to various voltages with a duration of either 50 ms (left) or 500 ms (right). Solid lines were obtained from best fits to experimental data using a Boltzmann equation. In absence of drug, these plots measure steady-state inactivation properties of WT and eqHPP channels (19). Lidocaine, mexiletine, and benzocaine shift voltage dependence of these inactivation curves to more hyperpolarized potentials. With the use of 500-ms conditioning pulses, 30 µM benzocaine, 100 µM lidocaine, or 100 µM mexiletine is able to shift inactivation curves measured in eqHPP channels so that they nearly overlay that recorded in WT channels without drug. rSkM1, rat skeletal muscle Na+ channel.

                              
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Table 2.   Estimates of V1/2 and k derived from fits to measured relative peak current amplitude as a function of 500-ms conditioning prepulse voltages for WT channels and eqHPP channels in the absence and presence of drug

Further information about local anesthetic effects on the stability of the inactivated state of eqHPP channels can be derived from the rate at which channels recover from inactivation in the presence of drug. The kinetics of recovery from inactivation in oocytes were assessed using a two-pulse protocol wherein conditioning pulses to -10 mV for 50 or 500 ms were followed by repolarization to -80 mV for a variable period before the introduction of a second test depolarization to -10 mV for 50 ms. Figure 5 summarizes the effects of lidocaine and mexiletine on the kinetics of recovery from inactivation using 50-ms conditioning pulses in WT channels (solid circles) and eqHPP channels with (open symbols) and without (solid squares) lidocaine (left) or mexiletine (right). On the brief time scale in Fig. 5, A and B, it can be seen that the degree of current recovery in eqHPP channels after 20 ms of repolarization to -80 mV was depressed by both lidocaine and mexiletine in a dose-dependent manner. Furthermore, Fig. 5, A and B, demonstrates that during the first 20 ms, the time course of recovery from inactivation of eqHPP in the presence of lidocaine or mexiletine is reasonably well described by a monoexponential function (solid lines). Indeed, inclusion of a second exponential component did not improve the quality of the fit (see MATERIALS AND METHODS). Despite the incomplete recovery of current on the time scale displayed in Fig. 5, A and B, the time constants estimated from the monoexponential fits did not change as the drug concentration was varied from 0 to 600 µM (see Fig. 5, E and F). As a result, the time course of recovery of eqHPP channels with drug application (Fig. 5, open symbols) never matched the recovery kinetics of WT channels (Fig. 5, solid circle). Extension of the recovery period as shown in Fig. 5, C (lidocaine) and D (mexiletine), reveals that full recovery from inactivation of eqHPP channels requires longer than 2 s in the presence of these agents. The full inactivation recovery curves shown in Fig. 5, C and D, required a biexponential function to accurately fit the experimental data (P < 0.05). As summarized in Fig. 5, E and F, the fast time constants for recovery (solid squares) in the presence of either lidocaine (Fig. 5E) or mexiletine (Fig. 5F), estimated from fits to the data in Fig. 5, C and D, did not vary with concentration; that is, the slopes of the relationship between tau fast and drug concentration were not significantly different from zero (P > 0.20 for lidocaine and P > 0.32 for mexiletine). Additionally, the fast time constants for recovery were not found to be statistically different from the time constant for recovery of eqHPP channels in the absence of drug (tau fast,lid = 3.2 ± 0.2 ms, n = 15 and tau fast,mex = 3.1 ± 0.2 ms, n = 15 vs. tau eqHPP = 2.9 ± 0.4 ms, n = 7). Lidocaine and mexiletine application did, however, induce the appearance of a much slower time constant for recovery from inactivation not observed in the untreated eqHPP channels. It is apparent from inspection of Fig. 5, C and D, that the relative amount of recovery occurring on a slow time scale (i.e., Aslow) increased with elevated drug levels (also see Fig. 6). On the other hand, the slope of the relationship between the slow time constant (i.e., tau slow) for recovery and the drug concentration was not significantly different from zero (Fig. 5, E and F, solid circles), again indicating independence of recovery kinetics on drug levels.


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Fig. 5.   A and B: recovery from inactivation over 20 ms of WT channels (bullet , n = 9) and of eqHPP channels in absence () and presence (open symbols) of lidocaine (A, n = 3-7) and mexiletine (B, n = 3-7) using 50-ms conditioning pulses. Data were fit with a monoexponential function. C and D: recovery from inactivation/block of WT and eqHPP mutant channels shown in A and B extended over 2 s in absence and presence of lidocaine (C) and mexiletine (D). Data over 2-s period were best fit with a biexponential function (solid lines). E and F: average fast (tau fast, ) and slow (tau slow, bullet ) time constants for recovery from inactivation estimated by fitting data in C and D to a biexponential function plotted as a function of lidocaine (E) or mexiletine (F) concentration. Note that neither time constant depended on drug concentration.


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Fig. 6.   Recovery from inactivation/block over 2 s of WT channel and eqHPP mutant in absence and presence of lidocaine (A) and mexiletine (B) using a 500-ms conditioning pulse. Again, data were best fit with a biexponential function (solid lines). Symbols are as indicated in Fig. 5. C and D: magnitude of slow component of recovery from block of eqHPP mutant, estimated from biexponential fits, as a function of lidocaine (C) and mexiletine (D) concentration using 50-ms () or 500-ms (open circle ) conditioning pulses. Data were fit with a binding isotherm using a nonlinear least-squares algorithm (see MATERIALS AND METHODS). IC50 values and Hill coefficients (n) for lidocaine and mexiletine using 50-ms conditioning pulse duration were estimated as follows: IC50,lid = 230 ± 12 µM, nlid = 1.6 ± 0.2, n = 4; and IC50,mex = 293 ± 9 µM, nmex = 1.3 ± 0.1, n = 4. With 500-ms conditioning pulse durations, values were as follows: IC50,lid = 44 ± 6 µM, nlid = 1.5 ± 0.1, n = 4; IC50,mex = 44 ± 8 µM, nmex = 1.7 ± 0.1, n = 4.

By comparison, Fig. 6 summarizes the kinetics of recovery from inactivation measured in eqHPP channels in the presence of lidocaine (A) or mexiletine (B) using conditioning pulses with a 500-ms duration. Inspection of Fig. 6, A and B, shows that the magnitude of the slow and fast time constants of recovery were not notably different when 500- vs. 50-ms conditioning pulses were used. However, the amplitude of the slow component for any given concentration of drug was much greater using 500-ms conditioning pulses compared with 50-ms pulses. For example, in the presence of 100 µM lidocaine, only 30% of the overall recovery occurs slowly when 50-ms conditioning pulses were used (Fig. 5C, open inverted triangles) compared with ~80% with 500-ms conditioning pulses (Fig. 6A, open inverted triangles). Similar effects of conditioning pulse duration can be seen with mexiletine by comparing Figs. 5D and 6B. This dependence of the relative amplitude of the slowly recovering component (Aslow) on conditioning pulse duration [i.e., 50 ms (solid squares) and 500 ms (open circles)] for different drug concentrations is plotted in Fig. 6C for lidocaine and in Fig. 6D for mexiletine application. Prolonging the conditioning pulse from 50 to 500 ms resulted in about a fivefold shift in the estimated IC50 for both lidocaine [Fig. 6, left, IC50,lid = 230 µM (50 ms) or 44 µM (500 ms)] and mexiletine [Fig. 6, right, IC50,mex = 293 µM (50 ms) or 44 µM (500 ms)] binding to eqHPP channels.

The data in Figs. 5 and 6 are consistent with previous studies in cardiac Na+ channels using lidocaine (5, 6) and suggest that recovery from inactivation involves two populations of channels: one drug-free population recovering at rates indistinguishable from untreated channels and one drug-attached population whose recovery is nearly 500-fold slower (5, 6). The differences in the number of drug-bound channels observed as a function of the conditioning pulse duration is also consistent with the previous studies using polar local anesthetics like lidocaine and mexiletine (6, 8, 17, 41, 47). The existence of two populations of channels in the presence of lidocaine and mexiletine underlies the complete lack of correspondence between the recovery time course of drug-treated eqHPP channels (open symbols) and either WT (solid circles) or untreated eqHPP (solid squares) shown in Figs. 5 and 6.

In contrast, Fig. 7 shows that the recovery properties of benzocaine-treated eqHPP channels using 50-ms (A) and 500-ms (B) conditioning pulses are very different from lidocaine- or mexiletine-treated channels. Figure 7, A and B, shows that the fractional recovery from inactivation of eqHPP channels at different benzocaine concentrations using either 50- or 500-ms conditioning pulse durations follows a monoexponential time course as expected from the rapid kinetics of benzocaine binding to Na+ channels (see DISCUSSION). Figure 7, A and B, further demonstrates that the kinetics of recovery from inactivation for WT channels closely matches eqHPP recovery in the presence of 1 mM and 300 µM benzocaine when 50- and 500-ms conditioning pulses were used, respectively. The dependence of the time constants for recovery from inactivation with increasing amounts of benzocaine is presented in Fig. 7, C and D, which shows that 1 mM benzocaine causes a nearly three- to sixfold slowing in the rate of recovery from inactivation for eqHPP channels. Although it appears from Fig. 7, A and B, that benzocaine, at a concentration between 300 µM and 1 mM, causes eqHPP channels to mimic the recovery properties of WT channels, it should be remembered that normalization of the data has eliminated the effects tonic block by benzocaine as illustrated in Fig. 2A. The concentration dependence of tonic block for benzocaine, as well as lidocaine and mexiletine, are summarized in Table 1.


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Fig. 7.   A and B: recovery from inactivation of WT channel (bullet ; n = 4-9) and eqHPP mutant in absence (; n = 7) and presence of 300 µM (triangle ; n = 10) and 1 mM (down-triangle; n = 7) benzocaine recorded for 50-ms (A) and 500-ms (B) conditioning pulses. Data were fit with a monoexponential function. Note that time delays were required to achieve adequate fits in presence of benzocaine (31). C and D: plots of recovery time constants of eqHPP channels as a function of benzocaine concentration with 50-ms (C) and 500-ms (D) conditioning pulse durations.

    DISCUSSION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References

Our data show that lidocaine, mexiletine, and benzocaine selectively modified inactivation properties of eqHPP channels. These observations are consistent with data from many previous studies which have established that local anesthetics preferentially bind to the inactivated as well as possibly the open conformation of Na+ channels (1-6, 8, 17, 18, 22, 24, 47) while binding very weakly to the closed conformations (6, 15, 47). We postulated that local anesthetics might be capable of restoring inactivation to inactivation-deficient Na+ channels carrying the equine periodic paralysis mutation (i.e., F1412L). Because local anesthetics are currently used clinically in the treatment of some forms of muscle myotonias, we examined whether these agents could confer the normal inactivation properties of WT channels onto eqHPP channels.

Neither lidocaine, mexiletine, nor benzocaine could fully bestow normal WT inactivation properties to eqHPP channels under a specific set of experimental conditions. For example, benzocaine at a concentration of 1 mM slows the rate of recovery from inactivation with 50-ms prepulses of eqHPP channels, making recovery kinetics nearly identical to WT channels. Moreover, benzocaine restores WT behavior with respect to the rate of Na+ current inactivation in eqHPP channels after depolarization to -10 mV, but this property was not observed at other membrane potentials, particularly below -20 mV (see Fig. 3B). At concentrations of benzocaine above 30 µM, the inactivation curves (measured as the dependence of peak current amplitude on prepulse voltage) for eqHPP channels were located leftward of that observed in WT channels without drug. Therefore, despite benzocaine's ability to correct the major inactivation defects observed in eqHPP channels, no single dose of this agent could fully restore WT inactivation behavior.

In comparison with benzocaine, lidocaine and mexiletine did not significantly alter the rate of Na+ current inactivation even at doses as high as 600 µM lidocaine (47). With the use of a 500-ms conditioning pulse protocol, the application of 100 µM lidocaine or mexiletine causes the eqHPP inactivation curves to overlap with the WT curve, whereas at lower and higher dosages, the inactivation curves for eqHPP channels were located rightward and leftward relative to WT channels, respectively. When examined over a 2-s period, recovery from inactivation occurred in two distinct phases at all concentrations of lidocaine and mexiletine studied. The amplitude of the slow component of recovery depended on drug levels and could be fit using a binding equation with a Hill coefficient between 1.3 and 1.7 (Fig. 6), which is different from the value of 1 reported previously in cardiac Na+ channels (5). These discrepancies might reflect genuine differences in the mechanism of block by local anesthetics between eqHPP, rSkM1, and cardiac Na+ channels (37, 47). Alternatively, this lack of correspondence could arise from differences in the experimental conditions. For example, Bean et al. (5) used 5-s conditioning prepulses, which is sufficiently long to induce slow inactivation (6, 19, 47). Hill coefficients greater than one might also reflect incomplete equilibration between the drug and channels in our experiments (47). Regardless, our studies demonstrate that lidocaine and mexiletine cannot faithfully restore normal inactivation properties in eqHPP channels at any drug concentration.

Differences in the effects of lidocaine and mexiletine vs. benzocaine probably reflect the faster binding and unbinding rates of benzocaine compared with lidocaine and mexiletine to the open and inactivated states of channels (8, 17, 24, 36, 41). With 50-ms conditioning pulses, a 10-fold lower concentration of benzocaine compared with lidocaine or mexiletine was required to produce the same degree of shift in the inactivation curves (i.e., same change in V1/2). We initially used conditioning pulses of 50-ms duration to crudely mimic the late phase of action potential duration observed in skeletal muscle (20, 29). However, a nearly threefold lower concentration of lidocaine and mexiletine is required to produce the same change in V1/2 (Fig. 4) when 500- vs. 50-ms conditioning pulses are used, reflecting incomplete equilibrium with short prepulses (5, 6, 17, 24, 49). Interestingly, prolonging the prepulse duration beyond 500 ms (data not shown) did not result in further measurable shifts in V1/2 as reported previously (47).

The initial focus of our experiments was to examine the ability of local anesthetic to modify and correct fast inactivation properties in eqHPP channels. Although the kinetics of mexiletine and lidocaine interaction with Na+ channels are too slow to confer WT kinetics onto eqHPP channels, they are nevertheless used clinically to abolish the multiple repetitive firing of skeletal muscle in human myotonia patients (25) at doses well below those used in our study. Cannon et al. (10) used a mathematical two-compartment model of skeletal muscle to demonstrate that incomplete inactivation of subpopulations of (mutant) Na+ channels within muscle cells was sufficient to cause repetitive trains of action potentials and, therefore, myotonia in patients with HPP. Thus preferential blockade of persistently active Na+ channels rather than reconstitution of inactivation might be key for clinical utility. In this regard, previous studies in cardiac channels containing mutations causing long Q-T syndrome (1) and inactivation-deficient rSkM1 channels (3) have demonstrated that lidocaine blocks noninactivating currents with a much higher affinity than the peak current. On the other hand, many Na+ channel mutations associated with skeletal muscle disease (9, 13, 11, 14, 16, 32, 34, 35, 39, 40, 42, 43), including eqHPP (23), and long Q-T syndrome in heart (7, 28, 48) significantly accelerate the rate of recovery from inactivation and shorten the effective refractory period. This feature of mutant channels has also been suggested to contribute to repetitive depolarizations in tissues containing mutant channels (1, 14). Therefore, slowing the rate of recovery, as we observed with lidocaine, mexiletine, and benzocaine, could also be an important element of drug action in disease treatment.

On the basis of the above discussion, fast agents like benzocaine might be preferred in the treatment of some forms of muscle myotonia by more rapidly promoting inactivation and blockade of non-inactivated channels. Slower binding drugs would leave a subpopulation of drug-free channels with persistent activity and reduced refractoriness (10, 34). Alternatively, when excessive numbers of Na+ channels are drug-bound, as might occur during periods of high muscle activity, sluggish agents like lidocaine and mexiletine are expected to remain bound to channels for longer periods thus promoting muscle inexcitability. However, the applicability of our results to the clinical treatment of muscle disorders is limited for a number of reasons. First, our investigations were restricted to the effects of local anesthetics on fast inactivation. It has been argued previously that impairment of slow inactivation is necessary for clinical disease (44). Moreover, use-dependent binding of lidocaine to slow inactivated channels is potentiated (4). Second, an important consideration guiding the choice of the Na+ channel modifiers in the treatment of muscle disorders is their relative tissue-specificity or their ability to discriminate between normal and inactivation-deficient channels. Indeed, major limitations of using Na+ channel modifiers in muscle disease treatment are side effects associated with drug action on normal channels (25). In this regard, the drug concentrations we used far exceed the dosages commonly used clinically (25).

It is important to note that two analogs of lidocaine (i.e., mexiletine and tocainide) are used clinically in the treatment of Na+ channel-based skeletal muscle disease like paramyotonia congenita as well as some forms of hyperkalemic paralysis/paramyotonia but not in the treatment of HPP (25). This observation is surprising, since the phenotypic changes occurring with different disease-causing Na+ channel mutations often have common biophysical properties (9, 13, 11, 23, 32, 34, 51). Our results show that local anesthetics can differentially modify inactivation properties in mutant channels associated with periodic paralysis in horses. Therefore, because disrupted function in eqHPP channels is not unlike that seen in many other Na+ channel mutations (9, 13, 32, 34, 51), these studies may provide some insights into the mechanism of local anesthetic action in the treatment of muscle diseases.

    ACKNOWLEDGEMENTS

We thank G. Mandel and W. A. Catterall for kindly providing the alpha -subunit of the rat skeletal muscle Na+ channel and the beta 1-subunit of the rat brain Na+ channel, respectively.

    FOOTNOTES

This work was supported by the Muscular Dystrophy Association of Canada and the Medical Research Council of Canada (to P. H. Backx). P. H. Backx holds a Medical Research Council of Canada scholarship award. R. L. Sah was supported by a Muscular Dystrophy of Canada Summer Studentship. We are grateful for support for equipment provided by the Alan Tiffin Trust, Toronto, Ontario, Canada.

Address for reprint requests: P. H. Backx, The Toronto Hospital, General Division, CCRW 3-802, 101 College St., Toronto, Ontario, Canada M5G 1L7.

Received 26 February 1997; accepted in final form 27 April 1998.

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