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Departments of
1 Anaesthesiology
2 Pharmacology, Vanderbilt University, School of Medicine, Nashville, TN 37232, USA
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Abstract |
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(Received 17 December 2004;
accepted after revision 24 January 2005;
first published online 27 January 2005)
Corresponding author P. C. Viswanathan: 560, Preston Research Building, 2220 Pierce Avenue, Vanderbilt University School of Medicine, Nashville, TN 37232-6602, USA. Email: prakash.viswanathan{at}vanderbilt.edu
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Introduction |
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With prolonged depolarizations, Na+ channels progressively enter ‘slow inactivated’ states with lifetimes ranging from hundreds of milliseconds to several seconds (Adelman & Palti, 1969). Pore mutations and ionic conditions that modify slow inactivation have parallel effects on Na+ channel sensitivity to use-dependent block by LAs (Kambouris et al. 1998; Chen et al. 2000). In Shaker K+ channels, conformational changes in the pore structure have been linked to slow ‘C-type’ inactivation (Choi et al. 1991; Hoshi et al. 1991; Lopez-Barneo et al. 1993; Yellen et al. 1994). In Na+ channels, linkages between the position of pore residues near the selectivity filter, DEKA (Heinemann et al. 1992), and slow inactivation have also been identified (Benitah et al. 1999; Todt et al. 1999; Ong et al. 2000; Hilber et al. 2001). A recent study suggests pore residues C-terminal to the selectivity filter are not involved in slow inactivation (Struyk & Cannon, 2002). Hence, it appears that residues in or N-terminal to the Na+ channel selectivity filter may represent the narrowest regions of the pore and are the most affected by slow inactivation gating.
Recently, we showed that slow inactivation protected an engineered cysteine at the 1236 position of the rat skeletal muscle Na+ channel (µ1: F1236C) from depolarization-dependent MTSEA modification (Ong et al. 2000). Lidocaine-induced use-dependent block substantially increased cysteinyl protection during pulse protocols optimized to elicit slow inactivation, but not fast inactivation. However, to clearly establish a causal link between slow inactivation and use-dependent block, it is necessary to show that the LA-associated protection of P-segment cysteines tracks the use-dependent properties of LA compounds with distinctive kinetic signatures such as lidocaine and bupivacaine.
Here we assayed the state-dependent accessibility of two cysteines engineered into the domain III, P-segment (µ1: F1236C, K1237C) to sulfhydryl modification with a methanethiosulphonate (MTS) reagent, MTS-ethylammonium (MTSEA). The rat skeletal muscle sodium channel (µ1) was used because this isoform has been well characterized especially with regard to slow inactivation and local anaesthetic block. Furthermore, previous studies have shown that these channels respond linearly to MTSEA reagents (Perez-Garcia et al. 1997). Using lidocaine and bupivacaine in a high-speed solution exchange system we are able to show, under identical pulse-train conditions, that sulfhydryl protection is conditionally dependent on the degree of use-dependent block. We propose that LA compounds may produce their kinetically distinct voltage-dependent behaviour by modulating slow inactivation gating to varying degrees.
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Methods |
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The rat skeletal muscle Na+ channel 
subunit (µ1) was subcloned into the HindIII–XbaI site of the vector green fluorescent protein with an internal ribosomal entry site (pGFPires) as previously described (Johns et al. 1997). Mutagenesis, F1236C or K1237C, was performed on the rat Na+ channel (Yamagishi et al. 1997) using a PCR-based method (QuickChange site-directed mutagenesis kit; Stratagene, La Jolla, CA, USA). All mutations were confirmed by dideoxynucleotide sequencing. Wild-type and mutated channels were expressed in HEK 293 cells using lipofectamine (Invitrogen Co., Carlsbad, CA, USA) along with Na+ channel ß1 subunit (provided by Dr Alfred George, Vanderbilt University). For experiments with F1304Q (provided by Dr Gordon Tomaselli, Johns Hopkins University), cells were cotransfected with rat Na+ channel ß1 subunit, and GFP. Transfected cells were incubated at 37°C in 95% O2–5% CO2 for 24–72 h before electrical recordings. Cells exhibiting green fluorescence were chosen for electrophysiological analysis.
Electrophysiology
All recordings were performed at room temperature (20–22°C). Whole-cell Na+ currents (INa) were obtained using an Axopatch 200B amplifier (Axon Instruments, Union City, CA, USA) and were acquired using pCLAMP 8 (Axon Instruments). Patch pipettes were pulled from borosilicate glass (Warner Instrument Inc., Hamden, CT, USA) with a Model P-97 Flaming-Brown micropipette puller (Sutter Instruments, San Rafael, CA, USA). Pipette resistance was 1–3 M
when filled with a pipette solution containing (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. For experiments involving F1304Q channels, pipette solution contained (mM): 10 NaF, 110 CsF, 20 CsCl, 10 EGTA, 10 Hepes, pH 7.35 with CsOH. The bath solution in both cases contained (mM): 150 NaCl, 4.5 KCl, 1.5 CaCl2, 1 MgCl2, 10 Hepes (pH 7.4 with NaOH). In all recordings, 80% of the series resistance was compensated, yielding a maximum voltage error of 
1 mV. MTS reagents: MTSEA, MTS-ethylsulphonate (MTSES), and MTS-ethyltrimethylammonium (MTSET) (Toronto Research Chemicals, Toronto, Ontario, Canada) were kept on ice as high concentration stock solutions and were diluted to 200 µM, 300 µM, or 5 mM in 5 ml bath solution immediately before use. Lidocaine HCl (Sigma Chemical Co., St Louis, MO, USA) or bupivacaine HCl (Sigma Chemical Co.) were diluted from stock solutions to the bath concentrations indicated in the text. Cells were clamped in whole-cell mode for at least 10 min before recording data. Whole-cell currents were sampled at 20 kHz (DigiData 1200 A/D converter; Axon instruments) and low pass-filtered at 5 kHz.
Rapid solution exchange
Rapid solution exchange was achieved using a computer-controlled High Speed Solution Exchange system (HSSE) (ALA Scientific Instruments, Westbury, NY, USA). HSSE has two orthogonal microbore Teflon® output tubes (300 µm inner diameter) which face each other at 90 deg in the same plane (as depicted in Fig. 1A). The pipette (along with the cell attached) was positioned close to the tips of the tubes. The solution flow through the two tubes was controlled using pCLAMP 8.0 software (Axon instruments). The time resolution of HSSE in our set-up was determined by exposing a high Na+ bath solution (150 mM NaCl) and a low Na+ bath solution (10 mM NaCl and 135 mM N-methyl-D-glucamine+) alternatively to wild-type Na+ channels using the protocol shown in Fig. 1B (inset). Briefly, a test pulse to –20 mV was applied in the presence of high Na+. After 50 ms at –120 mV the cell was exposed to the low Na+ solution for varying durations (P1). As seen in Fig. 1B, the direction of the Na+ current fully reversed by the time the cell was exposed to low Na+ for 30 ms, suggesting that solution exchange was completed within this time frame. The HSSE system was used only in experiments shown in Figs 4 and 5.
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The data were acquired and analysed using pCLAMP 8.0 software (Axon instruments). All results are expressed as means ± S.E.M. and statistical comparisons were made using one-way ANOVA (Origin, OriginLab Corp., Northampton, MA, USA) with P < 0.05 indicating significance. Multi-exponential functions were fitted to the data using nonlinear least-squares methods (Origin).
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Results |
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Depolarization-dependent accessibility of pore cysteines to MTS reagents
To assess conformational changes in the pore, we first examined depolarization-dependent accessibility of pore cysteine mutations to sulfhydryl reagents. For comparison to our prior studies (Ong et al. 2000), we retested the accessibility of a cysteine engineered at the 1236 locus (F1236C), to sulfhydryl reagents using the same protocols from Ong et al. (Fig. 2A). Since prior studies (Perez-Garcia et al. 1997) showed that MTSEA modification reduced peak INa in a voltage-independent manner, all of our experiments were conducted with depolarizations to –20 mV, a potential at which peak inward current was maximum (data not shown). A 3 min exposure to 200 µM MTSEA during prolonged depolarization (protocol II) reduced F1236C peak sodium current (31 ± 3%) to a greater extent than hyperpolarization (protocol I) (20 ± 1%, P < 0.05). While these results reconfirm that depolarization increases the accessibility of the cysteine side chain (see Fig. 2 of Ong et al. 2000), they do not distinguish between changes in MTS reactivity due to fast or slow inactivation. The higher concentration of MTSEA required for depolarization-dependent modification of F1236C in this study was probably due to variation in the potency of the compound.
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As shown previously for the 1236 locus (Ong et al. 2000), lidocaine exposure attenuated the modification of K1237C by MTSEA during depolarization (Fig. 2B and C, 20 ± 3%, versus control depolarization, P < 0.05). For consistency with prior work, the effect of lidocaine to attenuate MTSEA modification of the 1236 locus was also reconfirmed (Fig. 2C). In addition, bupivacaine, a LA with far slower use-dependent recovery kinetics, also attenuated MTSEA modification of K1237C (Fig. 2C, 19 ± 2%, versus control depolarization, P < 0.05). These results indicate that inhibition of the depolarization-dependent MTSEA modification is not specific to lidocaine, but also holds true for other LA compounds.
Kinetic differences between local anaesthestics
To explore whether kinetic differences between LA compounds in use-dependent block correlate with cysteine side chain accessibility in the P-loop during gating, we first evaluated the rate at which K1237C channels recovered from block by either lidocaine or bupivacaine (Fig. 3A). The recovery kinetics were fitted to a biexponential function (Table 1), and the drug concentrations were selected to produce slow recovery components of similar magnitude (A2). Lidocaine (100 µM) and bupivacaine (30 µM) increased the amplitude of the slow recovery component to a similar degree (A2: 0.78 ± 0.01 for lidocaine, 0.79 ± 0.04 for bupivacaine). However, the time constant of recovery of the slow component (
slow) in bupivacaine was much slower than in lidocaine (Fig. 3A: slow; 8744 ± 444 ms versus 314 ± 14 ms, P < 0.0001). The slower recovery from bupivacaine block is in agreement with previous findings (Clarkson & Hondeghem, 1985; Berman & Lipka, 1994).
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slow) remains long, and far slower than in lidocaine (Fig. 3B: slow; 5088.5 ± 442.5 ms versus 167 ± 30.3 ms, P < 0.01). Moreover, recovery of the F1304Q slow component in lidocaine was still delayed compared to drug-free conditions (see Table 1). These results show that distinctive recovery components induced by these LA agents do not require intact fast inactivation, and corroborate the findings of Vedantham & Cannon (1999). Slow inactivation protects MTS modification
Figure 3A also shows that 3 s after the end of the 1 s conditioning pulse, almost all K1237C channels had fully recovered (–120 mV) in the presence of lidocaine, while only 40–50% of channels recovered in the presence of bupivacaine (vertical bar, Fig. 3A). We made use of this difference to identify pulse train conditions that render cumulative block in bupivacaine, but not lidocaine. The protocol (Fig. 4A) applied repetitive 1 s depolarizing pulses to –20 mV with an interpulse repolarization to –120 mV for 3 s, allowing nearly complete recovery between pulses in lidocaine, but only partial recovery in bupivacaine. Figure 4B shows the representative current (every 10th pulse) in the presence of lidocaine or bupivacaine, in the absence of MTSEA. Peak INa is reduced by 50% at steady state in the presence of bupivacaine (right), but only by 10% in the presence of lidocaine (left), consistent with the distinctive use-dependent characteristics of the two agents, as also reflected in Fig. 3A.
We then examined whether marked differences in cumulative block by bupivacaine and lidocaine predictably track inactivation-dependent sulfhydryl modification of the 1237 cysteinyl. To selectively examine accessibility of the 1237 cysteinyl during inactivation, a rapid solution exchange system was utilized (Fig. 1). MTSEA was applied from 100 ms before the depolarization to 400 ms after depolarization (a total of 500 ms), as indicated in the protocol shown in Fig. 4A. While these exposure periods include both fast and slow inactivation gating components, our results in F1304Q (Fig. 3B) effectively exclude fast inactivation as the gating component primarily responsible for lidocaine–bupivacaine kinetic differences. In these experiments, the concentration of MTSEA was increased to 5 mM to allow substantial modification during these brief exposure periods. Since MTSEA was applied only during the period shown using the HSSE system, non-specific effects were minimized. Although depolarization in control, lidocaine, or bupivacine would lead to slow inactivation of Na+ channels during the initial 400 ms of each depolarization, we hypothesized that cumulative use-dependent bupivacaine block (under the conditions of the protocol in Fig. 4A) would ‘protect’ the K1237C residue from modification. In contrast, the complete recovery from inactivation in drug-free conditions or in lidocaine would allow MTS modification of unblocked channels at the onset of each depolarization. The measured currents, recorded before (Pre) and 60 s after the 100 depolarizing pulses (Post) allow for complete recovery from use-dependent drug block, so that any current reduction reflects MTS modification (superimposed in Fig. 4C). The data from all experiments are summarized in the bar graph (Fig. 4D), which shows the percentage of channels that have been modified by MTSEA. We measured less modification of K1237C by MTSEA in bupivacaine (39 ± 4%) than in lidocaine or control (52 ± 3% and 57 ± 3%, respectively, both P < 0.05 versus bupivacaine). This result suggests that accessibility of the 1237 cysteine to MTSEA is more restricted under kinetic conditions where use-dependent bupivacaine block is maintained, but use-dependent lidocaine block is not.
A gating-independent explanation for this result is that bupivacaine somehow provides better protection of the cysteinyl side chain, independent of its effect on inactivation gating. In this case, we would anticipate greater protection from MTSEA modification by bupivacaine under conditions where use-dependent INa reduction is similar to that evoked by lidocaine. Figure 5A shows the development of use-dependent block, in the absence or presence of lidocaine or bupivacaine, evaluated using the protocol shown in the inset. The concentrations of lidocaine (300 µM) and bupivacaine (30 µM) were chosen so that most of the channels (> 85%) were blocked by 500 ms or longer depolarizations to –20 mV (boxed region in Fig. 5A). To evaluate cysteine modification, a repetitive 1 s depolarizing pulse was applied, and MTSEA was selectively exposed for 500 ms in the latter portion of the 1 s depolarization pulse (Fig. 5B, upper panel), a period when most of the Na+ channels were blocked by lidocaine or bupivacaine (boxed region, Fig. 5A). As in Fig. 4, a 3 s interpulse repolarization to –120 mV was applied between each depolarization. The optimal approach would allow 1 min or more for full recovery of all channels from bupivacaine block (Fig. 3A); however, given the lengthy duration of the experiment, this was technically not feasible. While a 3 s interpulse period allowed only 40–50% of channels to recover from bupivacaine block (versus 100% in lidocaine), during all subsequent depolarizations, the channels are nonetheless fully blocked by either LA during the critical period (the latter 500 ms of Pn) when MTSEA is repeatedly exposed. Hence, the protocol (Fig. 5B) does provide conditions where MTSEA is exposed to channels when the extent of slow inactivation is saturated for lidocaine and bupivacaine, and thus similar, particularly in contrast to the conditions of Fig. 4.
The modification by MTSEA (Fig. 5B, bottom panel) was assessed before (Pre) and 60 s after the pulse train (Post). As before, this allowed for complete recovery from use-dependent lidocaine and bupivacaine block, so that any current reduction reflects MTS modification alone. Lidocaine significantly inhibited the modification of MTSEA compared to control (Fig. 5C: 17 ± 3% versus 27 ± 3%, P < 0.05), and the reduction of MTSEA modification by lidocaine was comparable to that of bupivacaine (14 ± 3%, P < 0.05 with control, NS versus lidocaine). Hence, similar degrees of use-dependent block have comparable effects on cysteinyl modification. The findings suggest that the differences found in Fig. 4D relate to drug-induced differences in gating (differential recovery from slow inactivation), as opposed to direct LA-specific interference of cysteine modification.
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Discussion |
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Voltage-gated Na+ channels are dynamic molecules that modify their conformational state in response to changes in membrane potential. Depolarization leads to channel inactivation within a few milliseconds (fast inactivation), a process involving residues situated near the cytoplasmic face of the channel (West et al. 1992). Prolonged depolarization induces channels to progressively occupy more stable slow-inactivated states. Numerous studies have linked conformational changes in the pore-lining segments of Na+ channels to slow inactivation (Balser et al. 1996a; Kambouris et al. 1998; Mitrovic et al. 2000; Xiong et al. 2003). The same residues, particularly K1237C, play a critical role in ionic selectivity of the pore for Na+ over Ca2+ (Favre et al. 1996; Tsushima et al. 1997).
Here we show that slow inactivation alters the accessibility of a cysteine engineered at the 1237 locus to sulfhydryl modification (Fig. 2), supporting prior studies indicating the deepest region of Na+ channel pore has restrictive access and undergoes conformational rearrangement during slow inactivation (Benitah et al. 1999; Ong et al. 2000). Unlike the K1237E or K1237S (Todt et al. 1999), K1237C in the present experiment did not produce ‘ultra slow’ inactivation ( on the order of 100 s), but rather, exhibited normal recovery kinetics in the absence of MTSEA (Fig. 3A), suggesting cysteine substitution alone did not significantly alter the channel gating. A recent study (Struyk & Cannon, 2002) showed that MTSET accessibility to pore cysteine mutations C-terminal to the selectivity filter did not change during slow inactivation. In our studies, MTSET (or MTSES) accessibility to K1237C was also not affected during depolarization (Fig. 2C) consistent with a previous study (Chiamvimonvat et al. 1996). Hence, pore rearrangements during slow gating appear to be focused on the narrow selectivity filter locus (K1237C) and residues immediately N-terminal (F1236C) (Ong et al. 2000) that is not accessible to the larger MTSET or negatively charged MTSES.
Local anaesthetics and slow inactivation
LAs exert their therapeutic effects by suppressing the ionic currents through Na+ channels. While the efficacy of LA action is enhanced by membrane depolarization (use dependence), the mechanistic basis for compound-specific differences in use-dependent drug block remains debated. Enzymes and mutations that remove fast inactivation attenuate use-dependent suppression of sodium channels by lidocaine and other Na+ channel blockers (Cahalan, 1978; Yeh & Tanguy, 1985; Bennett et al. 1995), suggesting an interaction between LAs and the fast-inactivated state. However, recent studies have questioned the postulated linkage between fast inactivation and use-dependent block by demonstrating that structural components of the Na+ channel involved in fast inactivation gating are not trapped in the fast-inactivated state by lidocaine (Vedantham & Cannon, 1999). In the present work, removal of fast inactivation (using a fast inactivation deficient mutant) did not eliminate the slow component of recovery seen in lidocaine and bupivacaine. Moreover, lidocaine did not inhibit the covalent modification of 1237C when MTSEA was exposed during the initial 400 ms of depolarization, a period that includes fast inactivation (Fig. 4), further supporting the findings of Vedantham & Cannon (1999). At the same time, bupivacaine elicits a slowly recovering block component within this time frame (Fig. 3A), and protected the 1237 cysteine from MTSEA modification (Fig. 4C and D).
Using MTSEA modification of the adjacent cysteine (F1236C), we recently (Ong et al. 2000) showed that lidocaine attenuates MTSEA modification during sustained depolarizations, but not brief, 5 ms depolarizations, also suggesting that pore rearrangements linked to slow inactivation and use-dependent lidocaine action are mechanistically linked (Ong et al. 2000). In this study, we provide further insight into the relationship between the pore rearrangement and use-dependent LA action by leveraging the kinetic differences between lidocaine and bupivacaine. Specifically, these data suggest that pore conformational changes that alter MTSEA accessibility track the differences in rate of recovery from use-dependent block for the two LAs and implicate slow inactivation in the compound-specific differences in use-dependent block.
We postulate the conceptual model shown in Fig. 6. While K1237C is relatively inaccessible during hyperpolarization (Fig. 6, left), depolarization leads to conformational changes in the pore-lining segments that increase MTSEA accessibility (Fig. 2). LAs do not inhibit accessibility of MTSEA during opening or fast inactivation, presumably because LAs are not trapped in the pore (Fig. 6, middle) (Vedantham & Cannon, 1999). However, with the accumulation of use-dependent drug block, the pore rearranges such that LA binding is stabilized and MTSEA accessibility of K1237C is reduced. Recently Bai et al. (2003) reported that mutations in the S6-domain LA binding sites affect slow inactivation, strengthening the notion that the binding of LAs may stabilize a structural rearrangement related to slow inactivation. Moreover, our findings showing the recovery kinetics of bupivacaine and lidocaine track the MTSEA accessibility of the 1237 cysteine side chain (Fig. 4) suggest that the various LAs may stabilize the slow-inactivated conformational state to differing degrees. As such, measured differences in rate of recovery from use-dependent block may reflect differences in LA affinity for the rearranged pore (Fig. 6, right).
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electron interactions (Ragsdale et al. 1994). In this regard, bupivacaine has two aromatic rings while lidocaine has only one. Moreover, three hydrophobic aromatic residues face the pore lumen in the putative S6 LA binding site (Ragsdale et al. 1994). Since bupivacaine has two aromatic rings, it is possible that electron interactions are enhanced, and stabilize slow inactivation, thus delaying recovery from drug block. We postulate that LAs stabilize structural rearrangements coupled to slow inactivation by interaction with pore-lining side chains in or near the selectivity filter, and that use-dependent kinetic differences between LAs may partly result from differences in the stability of this drug-bound conformational state.
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References |
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) and 30 µM bupivacaine (
) produced similar levels of block of INa (shaded region) after prolonged depolarizations (500–1000 ms). B, peak INa recorded using the protocol shown in the panel. In both LAs 50 repetitive pulses were applied to –20 mV with a 3 s interpulse recovery at –120 mV. Here, MTSEA (5 mM) was applied for the latter 500 ms of each 1 s depolarization, using the HSSE system, to expose cells to MTSEA during similar LA block. Plotted are currents recorded before (Pre) and 60 s after pulse train (Post), overlaid for comparison. C, fractional modification by MTSEA is summarized in the bar graph as the percentage reduction of peak INa (1 – Post/Pre). The number of cells recorded in each condition is shown in parentheses. For similar levels of slow inactivation, bupivacaine and lidocaine protected K1237C residue from MTS modification to a similar extent (NS with each other but P < 0.05 with control).
) or 100 µM lidocaine (