Correspondence to: Jeffrey R. Balser, Room 560, MRB II, Vanderbilt University School of Medicine, Nashville, TN 37232. Fax:(615) 936-0456 E-mail:jeff.balser{at}mcmail.vanderbilt.edu.
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Abstract |
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Voltage-gated sodium (Na+) channels are a fundamental target for modulating excitability in neuronal and muscle cells. When depolarized, Na+ channels may gradually enter long-lived, slow-inactivated conformational states, causing a cumulative loss of function. Although the structural motifs that underlie transient, depolarization-induced Na+ channel conformational states are increasingly recognized, the conformational changes responsible for more sustained forms of inactivation are unresolved. Recent studies have shown that slow inactivation components exhibiting a range of kinetic behavior (from tens of milliseconds to seconds) are modified by mutations in the outer pore P-segments. We examined the state-dependent accessibility of an engineered cysteine in the domain III, P-segment (F1236C; rat skeletal muscle) to methanethiosulfonate-ethylammonium (MTSEA) using whole-cell current recordings in HEK 293 cells. F1236C was reactive with MTSEA applied from outside, but not inside the cell, and modification was markedly increased by depolarization. Depolarized F1236C channels exhibited both intermediate (IM;
30 ms) and slower (IS;
2 s) kinetic components of slow inactivation. Trains of brief, 5-ms depolarizations, which did not induce slow inactivation, produced more rapid modification than did longer (100 ms or 6 s) pulse widths, suggesting both the IM and IS kinetic components inhibit depolarization-induced MTSEA accessibility of the cysteine side chain. Lidocaine inhibited the depolarization-dependent sulfhydryl modification induced by sustained (100 ms) depolarizations, but not by brief (5 ms) depolarizations. We conclude that competing forces influence the depolarization-dependent modification of the cysteine side chain: conformational changes associated with brief periods of depolarization enhance accessibility, whereas slow inactivation tends to inhibit the side chain accessibility. The findings suggest that slow Na+ channel inactivation and use-dependent lidocaine action are linked to a structural rearrangement in the outer pore.
Key Words: local anesthetic, gating, cysteine mutagenesis, lidocaine, electrophysiology
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INTRODUCTION |
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Local anesthetic compounds, such as lidocaine, suppress the ionic current through Na+ channels. By attenuating central and peripheral neuronal excitability, these compounds enjoy widespread use in the treatment of epilepsy and the relief of pain. In addition, local anesthetic compounds block Na+ channels in skeletal and cardiac muscle, and are used to treat neuromuscular diseases and cardiac arrhythmias. An essential characteristic of the local anesthetic action is use dependence, which is the sustained loss of excitability lasting many hundreds of milliseconds that is induced only when the drug-exposed channel is depolarized (
When briefly depolarized, Na+ channels inactivate rapidly within a few milliseconds (fast inactivation), a process mediated by residues situated near the cytoplasmic face of the channel (
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Investigating the structural basis of slow inactivation is hampered by its electrical silence, and requires a means to detect relatively slow changes in the conformational architecture of a nonconducting channel. Cysteine substitution of a P-segment residue in domain III (see Fig 1, F1236C) yields a Na+ channel reactive with the positively charged methanethiosulfonate (MTS)1 reagent MTS-ethylammonium (MTSEA) applied from outside the cell, yet the residue is entirely resistant to MTSEA applied from inside the cell (
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MATERIALS AND METHODS |
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Molecular Biology and Heterologous Expression
For heterologous expression, the wild-type rat skeletal muscle Na+channel (µ1) subunit was subcloned into the HindIII-XbaI site of the vector GFP-IRS for bicistronic expression of the channel protein and GFP reporter as previously described (
subunits were transiently transfected into HEK 293 (human embryonic kidney cell line) cells using lipofectamine (GIBCO BRL), and were cultured in MEM medium supplemented with 10% fetal bovine serum and 1% pen-strep in a 5% CO2 incubator at 37°C for 13 d. In all cases, the HEK cells were cotransfected with the Na+ channel ß1 subunit (provided by Dr. Alfred George, Vanderbilt University). Cells exhibiting green fluorescence were chosen for electrophysiological analysis.
Electrophysiology and Data Analysis
Whole-cell Na+ currents (INa) were recorded (Axopatch 200B; Axon Instruments) using electrodes with resistances of 13 M when filled with a pipet solution containing (in mM): 140 NaF, 10 NaCl, 5 EGTA, 10 HEPES, pH 7.40. Replacing the intracellular K+ with Na+ eliminated the time-dependent K+ currents in our HEK cell recordings. Experiments were conducted at room temperature. Current magnitudes were 12 nA, and 85% of the series resistance was compensated, yielding a maximum voltage error of
1 mV. The bath solution contained (in mM): 150 NaCl, 4.5 KCl, 1.5 CaCl2, 1 MgCl2, 10 HEPES (titrated to pH 7.40 with NaOH). MTSEA, MTSES, and MTSET (Toronto Research Chemicals) were kept at 4°C as high concentration stock solutions and were diluted to 25100 µM in the appropriate bath solution immediately before use. The disulfide reducing agents dithiothreitol (DTT) and glutathione were dissolved directly in the extracellular solution at a concentration of 5 mM (titrated to pH 7.4 with NaOH). Lidocaine HCl (Sigma-Aldrich) or QX-314 (Almone Labs) were diluted from stock solutions to the bath concentrations indicated in the text.
Cells were dialyzed for a 15-min equilibration period before recording data. To avoid junction potentials with solution changes, a 3-M KCl agar bridge was used. Inactivation gating kinetics and use-dependent block were assessed using the voltage-clamp protocols described in the text and figure legends. Whole-cell currents were sampled at 20 kHz (DigiData 1200 A/D converter; Axon Instruments) and low passfiltered at 5 kHz. The data were acquired and analyzed using pClamp8.0 software (Axon Instruments). The results are expressed as mean ± SEM, and statistical comparisons were made using One-Way ANOVA (Microcal Origin) with P < 0.05 indicating significance. Multiexponential functions were fitted to the data using nonlinear least-squares methods (Origin).
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RESULTS |
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We first examined the accessibility of the F1236C (Fig 1) cysteine side chain to sulfhydryl modification using 100 µM MTSEA. A 3-min exposure to MTSEA during hyperpolarization (-100 mV; Fig 2 A, protocol I) reduced F1236C peak INa by 31 ± 3% (after MTSEA washout; summary data Fig 2 B). This exceeded wild-type modification (13 ± 3%, P < 0.05; Fig 2A and Fig B), but was less than that previously seen with F1236C using a much higher concentration of MTSEA (49 ± 15%, 2.5 mM;
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The additional covalent modification afforded by depolarization was partially reversed by the hydrophobic reducing agent DTT. Fig 2 B indicates that after 3 min of depolarization-induced MTSEA modification, the fractional reduction in peak INa (relative to pre-MTSEA) after an additional 20 min of DTT exposure is reduced to 49 ± 5% (P < 0.005 versus the 73% pre-DTT value), indicating the INa reduction associated with MTSEA exposure results from formation of a reducible disulfide bond. The prolonged time (20 min) required for only partial DTT reversal suggests the reducing agent accesses the disulfide bond with some difficulty, perhaps through a hydrophobic pathway. We could not achieve reversal using glutathione, a larger, more hydrophilic, and generally less reactive reducing agent (Fig 2 B). Considered together, the data (Fig 2) suggest that although depolarization increases the accessibility of the F1236C side chain (allowing enhanced modification by MTSEA), the residue still lies at a relatively inaccessible position in the outer pore, limiting the effects of the larger sulfhydryl modification compounds or reducing agents.
The rate of F1236C INa decay during the depolarizing pulse did not differ from wild type (Fig 2 A), suggesting the mutant had no marked effect on the fast inactivation gating process. To examine the kinetics of slow inactivation, wild-type and F1236C peak INa were measured after prepulses of incremental duration (Fig 3, inset, clamp protocol). Each pair of depolarizations was separated by a brief, 20-ms hyperpolarization that allowed channels to recover fully from fast, but not slow, inactivation. Wild-type and F1236C slow inactivation were described by biexponential functions with similar time constants (parameters given in legend), corresponding to intermediate (IM,
30 ms) and slower (IS,
2 s) kinetic components of slow inactivation described in previous studies of µ1 expressed in Xenopus oocytes (
2, from 2,519 to 1,255 ms; see Fig 3 legend). Conversely, the rate of development of slow inactivation in F1236C was similar at -20 mV and +20 mV (Fig 3 B; amplitude of the IM component was
10%). Hence, consistent with previous evidence that P-segment substitutions alter slow inactivation gating processes in Na+ channels (
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A meaningful analysis of the depolarization-dependent rate of MTSEA modification of the F1236C side chain must recognize the distinctive kinetic features of slow inactivation, including the biexponential characteristics noted in Fig 3. Therefore, we used parameters derived from the kinetic analysis of F1236C slow inactivation (-20 mV; Fig 3 legend) to select depolarization pulse widths that would maximize the intermediate and slow kinetic components, IM (100 ms; 3 x
M) and IS (6 s;
3 x
S). In addition, a brief, 5-ms pulse width was used to examine depolarization-induced MTSEA modification in the complete absence of slow inactivation (Fig 3 B). Notably, while the pulse widths are selected to bias the channel toward distinct kinetic components of slow inactivation, caution is needed when relating these components to occupancy of individual gated states. The inactivated state dwell-times depend on many factors, including the detailed connectivity of the kinetic states involved.
Fig 4 (A and B) shows the time-dependent reduction of INa due to MTSEA modification during trains of either 5- or 100-ms pulses (-20 mV). In all experiments, MTSEA was allowed to equilibrate in the bath for at least 1 min before application of the first depolarization pulse. For these studies, the MTSEA was lowered to 25 µM (vs. 100 µM in Fig 2) to minimize tonic, depolarization-independent modification; as such, the reduction in INa during the first pulse after MTSEA addition was consistently 5% (Fig 4A and Fig B), compared with 31% at the higher concentration (Fig 2 B). In Fig 4, currents are plotted from every hundredth pulse (A) or every fifth pulse (B), and the currents shown in the two panels are aligned to reflect matching cumulative depolarization time.
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A comparison of A and B in Fig 4 reveals that extending the depolarization duration from 5 to 100 ms markedly reduced the rate of modification. Fig 4 C plots the time-dependent reduction in INa, due to MTSEA modification for a number of cells, as a function of matching (cumulative) depolarization time, allowing direct visual comparison of the MTSEA modification rates for the three pulse widths (5 ms, 100 ms, and 6 s). The repolarization intervals between pulses (200 ms, 4 s, and 240 s, respectively) were sufficiently long to prevent cumulative reduction in the current because of slow inactivation, and also were chosen such that the cumulative depolarization time increased as a function of total experimental time at the same rate for all three pulse widths. At the 6-s pulse width, the total (18 s) depolarization period was generated by only three pulses (Fig 4 C). Hence, in contrast to the 100-ms pulse train, channels transiently occupy the IM component, but spend a much higher percentage of their total depolarized time in the IS kinetic component. Nonetheless, the rate of depolarization-dependent MTSEA modification is still reduced compared with the brief, 5-s depolarizations (Fig 4 C), indicating that pulses recruiting IS (like IM) reduce MTSEA accessibility of the cysteine side chain. The data in Fig 4 C were fitted to an exponential function (solid line) to determine reaction rates (kon; Fig 4 D, see legend) for each depolarization pulse width; increasing the pulse width to either 100 ms or 6 s significantly reduced kon. Fig 4 D also indicates that the effects of pulse width (5 and 100 ms) were insensitive to changing the depolarization voltage from -20 to +20 mV, which is consistent with the similar rate of slow inactivation for F1236C (in contrast to wild type) at these two membrane potentials (Fig 3 B).
The rate of MTSEA modification changes with the depolarization pulse width; the extent of modification also appears to change. Incomplete elimination of the current could partly result from residual current flow through MTSEA-modified channels. Even under conditions where slow inactivation is prevented (i.e., brief 5-ms pulses), the rate of depolarization-dependent modification is still 100-fold slower than in previous studies examining MTS modification of freely accessible cysteinyl groups (i.e., IIIIV linker 1304C;
We next considered whether accessibility of F1236C to sulfhydryl modification was modified by use-dependent lidocaine block. Since the F1236C mutation alone modifies slow inactivation under control conditions (Fig 3), we first established whether lidocaine would induce significant depolarization-dependent INa suppression in this mutant. Bath exposure to 100 µM lidocaine (Fig 5 A) reduced INa availability mainly by increasing the amplitude of an intermediate ( = 68 ms) kinetic component (see Fig 5 legend, lidocaine increased A1 from 0.08 to 0.65). Hence, the mutant channel retains use-dependent lidocaine sensitivity typical for skeletal muscle Na+ channels expressed in HEK cells (
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To determine whether the effects of lidocaine on use-dependent loss of INa availability could be linked to a structural change in the outer pore, we examined voltage-dependent MTSEA modification during lidocaine exposure. Fig 5 B shows the rate of MTSEA modification induced by 100- ms depolarizing pulses to -20 mV during MTSEA exposure, alone and with lidocaine superfusion. For these experiments, we raised the MTSEA concentration from 25 to 50 µM to allow significant modification of the cysteinyl side chain using a 100-ms pulse width in lidocaine-free conditions (note, for comparison, only minimal modification using a 100-ms pulse width in Fig 4 C). Notably, in lidocaine-free solutions, the rate of MTSEA modification (kon, 103 M-1s-1) using 100-ms pulses (Fig 5 B, 4.4 ± 0.6) was slower than with 5-ms pulses (Fig 5 C, 9.6 ± 1.4, P = 0.004 vs. 100-ms pulses) even though the MTSEA concentration was raised in Fig 5 B. This finding supports the results of Fig 4, indicating a slower rate of depolarization-dependent modification when the pulse duration is lengthened. The data also reveal a marked reduction in MTSEA modification with lidocaine exposure; in Fig 5 B (100-ms pulse width), kon in lidocaine was as decreased to 0.7 ± 0.3 (P = 0.0002). In contrast, lidocaine exposure had no effect on the rate of MTSEA modification when a brief pulse width was used (Fig 5 C, 5 ms). Hence, the protection from sulfhydryl modification afforded by lidocaine is use-dependent and requires a sustained depolarization. This pulse width sensitivity suggests the observed lidocaine effect on MTSEA modification was not a nonspecific, gating-unrelated antagonism between the local anesthetic the MTS reagent.
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DISCUSSION |
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Based on evidence that enzymes (
These results do not resolve whether the impeded sulfhydryl modification rate during slow inactivation results from movement of the domain III P-segment to a less accessible position, or rather from movement of other pore structures into positions that somehow protect the F1236C side chain from MTSEA. Nonetheless, a conceptual model based upon recent studies of the Na+ channel pore demonstrating exceptional mobility of the P-segments (
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The 1236 cysteine side chain is accessible only to modification from outside the cell (
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Footnotes |
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1 Abbreviations used in this paper: DTT, dithiothreitol; HEK, human embryonic kidney; INa, Na+ channel; MTS, methanethiosulfonate; MTSEA, MTS-ethylammonium; MTSET, MTS-ethyltrimethylammonium.
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Acknowledgements |
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We wish to thank Drs. Steve Cannon and Paul Bennett for valuable criticism of the manuscript.
This work was supported by grants from the National Institutes of Health R01 GM56307 (to J.R. Balser) and R01 HL 50411 (to G.F. Tomaselli).
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References |
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