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Lidocaine Promotes the Trafficking and Functional Expression of Na_v1.8 Sodium Channels in Mammalian Cells

¹Le Centre de Recherche Université Laval Robert-Giffard and Department of Medicine, Laval University, Quebec, Quebec, Canada; and ²Jefferson Medical College, Jefferson University, Philadelphia, Pennsylvania

Submitted 2 February 2007; accepted in final form 9 May 2007

	ABSTRACT

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Nociceptive neurons of the dorsal root ganglion (DRG) express a combination of rapidly gating TTX-sensitive and slowly gating TTX-resistant Na currents, and the channels that produce these currents have been cloned. The Na_v1.7 and Na_v1.8 channels encode for the rapidly inactivating TTX-sensitive and slowly inactivating TTX-resistant Na currents, respectively. Although the Na_v1.7 channel expresses well in cultured mammalian cell lines, attempts to express the Na_v1.8 channel using similar approaches has been met with limited success. The inability to heterologously express Na_v1.8 has hampered detailed characterization of the biophysical properties and pharmacology of these channels. In this study, we investigated the determinants of Na_v1.8 expression in tsA201 cells, a transformed variant of HEK293 cells, using a combination of biochemistry, immunochemistry, and electrophysiology. Our data indicate that the unusually low expression levels of Na_v1.8 in tsA201 cells results from a trafficking defect that traps the channel protein in the endoplasmic reticulum. Incubating the cultured cells with the local anesthetic lidocaine dramatically enhanced the cell surface expression of functional Na_v1.8 channels. The biophysical properties of the heterologously expressed Na_v1.8 channel are similar but not identical to those of the TTX-resistant Na current of native DRG neurons, recorded under similar conditions. Our data indicate that the lidocaine acts as a molecular chaperone that promotes efficient trafficking and increased cell surface expression of Na_v1.8 channels.

	INTRODUCTION

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Voltage-gated Na channels are membrane proteins that play an important role in the rapid electrical signaling in nerve fibers (Hille 2001). The nociceptive neurons of the dorsal root ganglia (DRG) express a unique combination of tetrodotoxin (TTX)-sensitive and -resistant Na channels that play a key role in the physiology and pathophysiology of pain (Waxman et al. 2000). Several Na channels have been cloned from peripheral nerve; Na_v1.7 (PN1 for peripheral nerve Na channel 1) encodes for a rapidly gating TTX-sensitive Na channel that is broadly expressed in both large and small DRG neurons (Sangameswaran et al. 1997). Na_v1.8 (PN3) encodes for a slowly gating TTX-resistant Na channel that is highly localized within the small nociceptive neurons of the DRG neurons (Sangameswaran et al. 1996). Differences in the biophysical properties and the pharmacology of these Na channels contribute to the unique electrical properties of sensory nerve fibers.

Several distinct components of TTX-R Na current have been observed in DRG sensory neurons (Elliott and Elliott 1993; Kostyuk et al. 1981; Ogata and Tatebayashi 1993; Roy and Narahashi 1992; Rush et al. 1998; Scholz et al. 1998). One component displays a low threshold for activation and rapid kinetics and is believed to be produced by the Na_v1.5 channel, which is highly expressed in embryonic sensory neurons and at a lower level in adult neurons (Renganathan et al. 2002). A second component of TTX-R current activates at relatively depolarized voltages, has comparatively slow gating kinetics, and rapidly recovers at hyperpolarized voltages (Elliott and Elliott 1993; Kostyuk et al. 1981; Ogata and Tatebayashi 1993; Roy and Narahashi 1992; Rush et al. 1998). These properties are similar to what has been described for the Na_v1.8 channel (Akopian et al. 1999). A third component of TTX-R current has a relatively hyperpolarized threshold for activation (approximately equal to –80 mV) and displays little inactivation (Cummins et al. 1999). The Na_v1.9 channel appears to underlie this component of DRG Na current (Cummins et al. 1999; Fjell et al. 2000).

The presence of multiple overlapping components of TTX-R Na current in small DRG neurons has complicated the detailed biophysical characterization of the underlying Na channels. The cloning of the Na_v1.8 channel raised the prospect of its in vitro expression that would permit detailed electrophysiological characterization of the isolated channels (Akopian et al. 1996; Sangameswaran et al. 1996). The Na_v1.8 channel has been expressed in Xenopus oocytes, and the properties of the heterologously expressed channels are in good agreement with those of the native TTX-R Na current of DRG neurons (Akopian et al. 1996; Sangameswaran et al. 1996; Vijayaragavan et al. 2001). By contrast, attempts to heterologously express the Na_v1.8 channel in cultured mammalian cells have been met with limited success (John et al. 2004; Vijayaragavan et al. 2004). Although the reasons for the poor expression of the Na_v1.8 channels in cultured cells are not known, it is generally believed to result from a trafficking defect that reduces the cell surface expression of functional channels (Okuse et al. 2002).

A number of factors appear to regulate the expression of Na_v1.8 channel in mammalian cell lines. Interaction of Na_v1.8 with accessory proteins appears to either promote the translocation to the plasma membrane or stabilize channels within the plasma membrane. Annexin II light chain (p11) enhances Na_v1.8 expression by directly interacting with the cytoplasmic N terminus of the channel protein resulting in translocation to the plasma membrane (Okuse et al. 2002). Co-expression of the accessory ₃ subunit, annexin II light chain (p11), and clatherin-associated protein-1A (CAP-1A) appear to be important regulators of Na_v1.8 expression (John et al. 2004; Liu et al. 2005; Okuse et al. 2002). Whereas ₃ and p11 were reported to enhance cell surface expression, CAP-1A reduces the density of the expressed Na_v1.8.

Despite the enhanced trafficking produced by accessory proteins (₃ and p11), the current density of the heterologously expressed Na_v1.8 channel remains relatively low by comparison to the native TTX-R current of DRG neurons and the Na_v1.7 channel expressed under similar conditions (Vijayaragavan et al. 2004). As-yet-unidentified factors or regulatory mechanisms appear to contribute to the efficient trafficking and cell surface expression of the Na_v1.8 channel. Increases in nociceptor excitability and changes in the level and distribution of Na_v1.8 expression were associated with acute nerve injury and inflammatory reactions (McCleskey and Gold 1999). Understanding the mechanisms governing Na_v1.8 trafficking therefore has important implications for the peripheral sensitization to painful stimuli linked to tissue damage and nerve injury.

Recently it has been shown that in several channelopathies, mutations in channel proteins that induce misfolding and therefore the retention of the channel in the ER can be rescued by chemical chaperones or pharmacological ligands (Morello et al. 2000). This is the case for mutations that cause the Brugada syndrome an inherent cardiac disorder and that the defect of the misfolding was rescued by the antiarrhythmic drug mexiletine (Valdivia et al. 2002).

Chemical chaperones are small molecules that assist folding and restore the trafficking of receptors and channels (Morello et al. 2000). In this study, we investigated the effects of the local anesthetic lidocaine on the expression of the Na_v1.8 channel in tsA201 cells, a transformed variant of the HEK293 cell line. Our data indicate that lidocaine acts as a chemical chaperon that promotes Na_v1.8 trafficking to the plasma membrane. The biophysical properties of the heterologously expressed Na_v1.8 channels are similar to but not identical to those of the TTX-resistant Na current recorded under identical conditions from DRG neurons.

	METHODS

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Heterologous expression of Nav1.8 in tsA201 cells

The tsA201 cell line is a mammalian cell line derived from human embryonic kidney HEK 293 cells by stable transfection with SV40 large T antigen (Margolskee et al. 1993). Cells were grown in high glucose DMEM supplemented with FBS (10%), L-glutamine (2 mM), penicillin (100 U/ml) and streptomycin (10 mg/ml) (Gibco BRL Life Technologies, Burlington, Ontario, Canada). Cells were incubated in a 5% CO₂ humidified atmosphere. Transfections of tsA201 cells were carried out using the calcium phosphate method as previously described (Margolskee et al. 1993).

The ₁ subunit was co-expressed with Na_v1.8. The human Na⁺ channel ₁-subunit and CD8 were constructed in the piRES vector (piERS/CD8/₁) (Invitrogene Corporation, Carlsbad, CA). Transfected cells were briefly (<5 min) preincubated with CD8 antibody-coated beads prior to recording (Dynabeads M-450 CD8-a). Cells expressing the piRES/CD8/₁ bicistronic vector were decorated with CD8 beads, which were used to identify cells for patch-clamp analysis (Margolskee et al. 1993). Transfected tsA201 cells were then pretreated for 24 h with 1 mM lidocaine. The cells were subsequently incubated for 3–4 h in lidocaine-free media immediately prior to electrophysiological recordings.

Isolation of dorsal root ganglion neurons

Seven-day-old rat pups are anesthetized with isoflurane before decapitation and the dorsal root ganglia from all accessible levels of the spinal cord harvested. Excess connective tissues were removed and the ganglion placed in 3 ml of Hank's balanced salt solution (HBSS, Gibco) supplemented with 10 mM HEPES. The ganglions were incubated for 30 min at 37°C in 2 ml of HBSS/HEPES containing 1.5 mg/ml collagenase (Sigma-Aldrich, St. Louis, MO). The ganglia were washed with HBSS/HEPES before adding 1 mg/ml trypsin (Sigma-Aldrich) and incubating an additional 30 min at 37°C. Trypsin was removed and the ganglia were transferred to L-15 Leibovitz media (Gibco) supplemented with 1% fetal bovine serum (FBS, Gibco), 2 mM glutamine, 24 mM NaHCO₃, 38 mM glucose, 2% penicillin-streptomycin (Gibco), and 50 ng/ml nerve growth factor (Sigma-Aldrich). The ganglia were dissociated by gentle titration using fire-polished Pasteur pipettes, and the dissociated neurons were plated onto 35 mm dishes containing 2 ml of the supplemented Leibovitz media. Dissociated neurons (20 µm, 8–12 pF of capacitance) were suitable for patch-clamp studies 1–2 h after plating. Neonatal rats were killed in accordance with the animal welfare protocols of our institution.

Electrophysiology

Macroscopic Na currents from tsA201 transfected cells were recorded using the whole cell configuration of the patch-clamp technique (Chahine et al. 2004). Command pulses were generated and current recorded using pCLAMP software v8.0 and an Axopatch 200 amplifier (Molecular Devices, Union City, CA). Patch electrodes were fashioned from borosilicate glass (Corning 8161) and coated with silicone elastomer (Sylgard, Dow-Corning, Midland, MI) to minimize stray capacitance. Current recordings were made using low-resistance electrodes (<1M), and the series resistance was compensated at values 80%, to minimize voltage-clamp errors. Whole cell currents were filtered at 5 kHz, digitized at 10 kHz, and stored on a microcomputer equipped with an AD converter (Digidata 1300, Molecular Devices). Data analysis was performed using a combination of pCLAMP software v9.0 (Molecular Devices), Microsoft Excel and SigmaPlot for Windows version 8.0 (SPS, Chicago, IL). The current signal was low-pass filtered at 2 kHz and digitalized at a sampling rate of 100 µs during acquisition. Traces shown were low-pass filtered at 1.5 kHz using a digital filter of Clampfit software.

Solutions and reagents

For whole cell recordings, the patch pipette contained (in mM) 5 NaCl, 135 CsF, 10 EGTA, and 10 Cs-HEPES. The pH was adjusted to 7.4 using 1 N CsOH. The bath solution contained (in mM) 150 NaCl, 2 KCl, 1.5 CaCl₂, 1 MgCl₂ 10 glucose, and 10 Na-HEPES. The pH was adjusted to 7.4 with 1 N NaOH. For Na current recording from DRG neurons, the extracellular solution contained instead 140 mM NaCl and the intracellular solution contained 35 mM NaCl. A correction for the liquid junction potential between the patch pipette and the bath solutions (–7 mV) was applied to the command pulses. The recordings were made 10 min after obtaining the whole cell configuration to allow the current to stabilize and to fully dialyze the cell with pipette solution. The Na_v1.8 -subunit and the ₁ auxiliary subunit were cloned in our laboratory as described previously (Vijayaragavan et al. 2001).

For single-channel recording, 3–5 M patch electrodes were used. Patch electrodes were coated with Sylgard to reduce their capacitance and lower noise emission. The bath solution contained a high concentration of potassium composed of (in mM) 100 K-aspartate, 50 KCl, 1.5 CaCl₂, 1 MgCl₂, 10 glucose, and 10 K-HEPES (pH = 7.4). This solution was used to depolarize the cell, thereby making the applied command potential approximately equal to the voltage across the membrane patch. The patch pipette solution contained (in mM) 150 NaCl, 10 TEA-Cl, (to block endogenous potassium channels), 2 KCl, 1.5 CaCl₂, 1 MgCl₂, 10 glucose, and 10 Na-HEPES (pH = 7.4). Single-channel currents were recorded using an Axopatch 200B amplifier, a Digidata 1200 acquisition system, and pCLAMP v9 (Molecular Devices). Single-channel currents were filtered at 2 kHz and sampled at 100 kHz. Single-channel currents were recorded at room temperature (22–23°C).

Immunocytochemistry

Transfected tsA201 cells were permeabilized using 0.1% Triton in 1 mM PBS-0.5% BSA solution before incubation with antibodies. Cells were fixed using a 1:3 acetone/methanol solution for 20 min. The mouse anti-Na_v1.8 primary antibody (1:100) was used against the Na_v1.8 alpha-subunit (Alomone, Jerusalem). The secondary antibody was a conjugated AffiniPure goat anti-mouse (1:400) (Molecular Probes). Rabbit Anti-Calnexin polyclonal antibody (1:4,000) was used for endoplasmic reticulum (ER) labeling, and was obtained from StressGen Biotechnologies (Victoria, British Columbia, Canada).

Confocal microscopy

Fluorescent probe-labeled tsA201 cells were examined on a Bio-Rad MRC-1024 confocal imaging system equipped with a krypton-argon laser beam mounted on a Zeiss microscope. A x360 oil objective with a 1:4 numerical aperture was used. Confocal settings were as follows: 1-mw laser power, 1.2 zoom, 1 s per scan, Kalman filter, and 4 frames per image. The photomultiplier gain was adjusted and the aperture adjusted for maximum resolution.

Biotinylation of cell surface proteins

tsA201 cells were cultured in 100-mm dishes and transiently transfected with 5 µg of Na_v1.8 and 5 µg of ₁. After transfection (24 h), the cells were treated with 1 mM lidocaine or without (control). After the transfection (48 h), cells were subjected to cell surface biotinylation. Recovery of plasma membrane proteins were carried out using the Pierce cell-surface protein biotinylation and purification kit according to the manufacturer's protocol (Pierce, Rockford, IL). Samples were analyzed by immunoblotting using rabbit anti-Na_v1.8 antibody (Alomone) at 1:200 dilution.

Western blot analysis

tsA201 cells were washed with phosphate-buffered saline (PBS, pH 7.4) and solubilized in 1 ml of STEN buffer consisting of: 0.2% NP40, 1% Triton X-100 and protease inhibitor mixture (Roche Molecular Biochemicals, Mannhein, Germany). Insoluble debris were removed by centrifugation at 13,000 rpm for 30 min. Equivalent amounts of proteins were applied to an SDS-10% PAGE electrophoresis gel, and the separated proteins were transferred to a Hybond P membrane (Amersham Pharmacia Biotech, Piscataway, NJ). After protein transfer, the membrane was blocked for 1 h at room temperature in 5% nonfat milk in PBS-T (0.1% tween –20 in PBS, pH 7.4). The membranes were probed with anti-Na_v1.8 antibody (1/200 dilution; Alomone), followed by horseradish peroxidase-conjugated goat anti-rabbit (1/10,000 dilution; Amersham Pharmacia Biotech) and ECL detection (Amersham Pharmacia Biotech).

	RESULTS

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Lidocaine enhances the cell surface expression of the Na_v1.8 channel

Since the cloning of Na_v1.8 in 1996 (Sangameswaran et al. 1996), the heterologous expression of this Na channel in mammalian cells has proved to be problematic, often resulting in little or no expression of Na current (Okuse et al. 2002; Vijayaragavan et al. 2004). This has complicated efforts to study the biophysical properties of the heterologously expressed channel. The poor expression of Na_v1.8 in cultured cell lines is generally believed to result from a trafficking defect that prevents the channel protein from reaching the plasma membrane (Okuse et al. 2002). To further investigate the underlying mechanism, we used immunohistochemistry and confocal imaging to study the subcellular localization of the Na_v1.8 channel heterologously expressed in tsA201 cells. Immunofluorescence revealed a significant accumulation of Na_v1.8 in perinuclear regions (Fig. 1 A, green staining) that displayed significant overlap with the immunostaining for calnexin, a specific marker of the endoplasmic reticulum (ER; Fig. 1B, red staining). The overlap of the Na_v1.8 and calnexin (Fig. 1C, yellow staining) suggests that the majority of the channel protein is localized within intracellular organelles with little distribution to the plasma membrane.

View larger version (40K): [in this window] [in a new window]   FIG. 1. Subcellular localization the Nav1.8 channel.tsA201 cells were co-transfected with Nav1.8 and the 1 subunit and the subcellular localization of channel protein determined using antibody labeling and confocal microscopy. A: Nav1.8 immunostaining showing perinuclear localization (Green staining). B: immunofluroescence of the same cells as in A showing the distribution of calnexin (red staining), a protein expressed in endoplasmic reticulum. C: superimposition of Nav1.8 (A) and calnexin (B) staining (orange/yellow). D: pattern of Nav1.8 immunofluorescence (green) after incubating the cells for 24 h with 1 mM lidocaine. E: same cell as in D showing that the subcellular localization of calnexin was not altered by lidocaine preincubation. F: superimposition of Nav1.8 (D) and calnexin (E) staining of lidocaine-treated cells.

The immunohistochemistry suggests that lidocaine may act by enhancing the trafficking of the Na_v1.8 channels from the ER to the plasma membrane. Alternatively, the observed changes in Na_v1.8 distribution could have resulted from the synthesis of new channel protein that was specifically targeted to the cell periphery. To distinguish between these alternatives, we used Western blot analysis to investigate the effects of lidocaine on the expression of Na_v1.8 channel protein. Figure 2 A shows the Western blot of the Na_v1.8 channel expressed in tsA201 cells preincubated under control conditions or after 24-h incubation with lidocaine (1 mM). The expression levels of the Na_v1.8 channels were similar under both conditions, suggesting that the increase in peripheral Na_v1.8 immunostaining induced by lidocaine (Fig. 1F) does not result from the synthesis of new channel protein.

View larger version (29K): [in this window] [in a new window]   FIG. 2. Effects of lidocaine on Nav1.8 expression and membrane localization. A: tsA201 cells were co-transfected with Nav1.8 and the 1 subunit. B: total cell extracts were prepared and the expression of Nav1.8 protein evaluated using Western blots. First lane: 10 µg of protein extracted from Nav1.8/1-transfected tsA201 cells nontreated; second lane: equal amount of protein (10 µg) extracted from Nav1.8/1 transfected cells preincubated for 24 h with 1 mM lidocaine. Bottom: Western blot of the same protein extract, probed with -tubilin specific antibody and used for band quantification. Densitometry analysis showed no difference between nontreated and treated cells. B: Nav1.8 expression at the plasma membrane determined by a combination of biotinylation of cell-surface proteins, isolation of biotinylated proteins on biotin binding columns, and Western blotting (see METHODS). Top: Western blot of nontransfected tsA201 cells, Nav1.8/1 transfected cells, and Nav1.8/1 transfected cells after 24 preincubation with lidocaine. Middle: Western blots of total cell lysate of nontransfected cells, Nav1.8/1 transfected drug-free controls, and Nav1.8/1 transfected cells after preincubation with lidocaine. Bottom: Western blot of biotinylated proteins identified using an antibody directed at an intracellular protein (-tubilin).

Lidocaine enhances the expression of functional Na_v1.8 channels in the plasma membrane

Immunohistochemistry indicates that preincubating transfected cells with lidocaine increased the cell surface expression of Na_v1.8 channel protein. If these channels are both functional and associated with the plasma membrane, then these findings predict an increase in the Na current amplitude of cells pretreated with the drug. Figure 3 compares the current-voltage relationships of transfected tsA201 cells measured under control conditions and after preincubation with lidocaine. Lidocaine dramatically increased the peak Na current amplitude (Fig. 3B) by comparison to drug-free controls (Fig. 3A). To quantitate these findings, the peak Na current amplitudes were normalized to the whole cell capacitance of individual cells (Fig. 3, C or D). Preincubating with lidocaine induced a significant sixfold increase in the Na current density (–123.3 ± 6.1 pA/pF, n = 12) by comparison to drug-free controls (–23.3 ± 1.6 pA/pF, n = 9, t-test, P < 0.05). The increase in Na_v1.8 immunostaining along the cell periphery (Fig. 1) and the parallel increase in Na current density are consistent with our working hypothesis that preincubating cells with lidocaine promotes the re-distribution of functional Na_v1.8 channels from intracellular compartments to the plasma membrane.

View larger version (28K): [in this window] [in a new window]   FIG. 3. Lidocaine increased the Na current density of Nav1.8-transfected cells. Nav1.8 and 1 subunits were co-expressed in tsA201 cells. A and B: whole cell Na currents of a drug-free control (A) and after 24 h incubation with 1 mM lidocaine (B). Currents were elicited by depolarizing steps between –100 and +90 mV in 10-mV increments from a holding potential of –140 mV. C: peak Na current at each voltage was normalized to the whole cell capacitance and the current density (pA/pf) plotted vs. the test voltage (control, n = 9; lidocaine treated, n = 12). D: at +20 mV, preincubating with lidocaine produced a significant sixfold increase in the Na current density (–123.3 ± 6.1 pA/pf, n = 12) by comparison to the drug-free controls (–23.3 ± 1.6 pA/pf, n = 9, t-test: P < 0.01).

View larger version (10K): [in this window] [in a new window]   FIG. 4. The effect of 10 µM TTX on Nav1.8 currents. Whole cell Nav1.8 currents were evoked every 4 s by 50-ms pulses from –100 to +95 mV in 5-mV increments. The holding potential is –140 mV. —, control current. · · ·, current in the presence of 10 µM TTX for 10 min, the peak current amplitude were inhibited by 16.02 ± 2.6% (n = 7). - - -, 0 current.

We took advantage of the enhanced expression of Na_v1.8 afforded by lidocaine treatment to further characterize the biophysical properties of the heterologously expressed channel. The time course of the current decay elicited at depolarized voltages was fitted with the sum of two exponentials and the resulting time constants plotted versus voltage (Fig. 5 A). The fast and slow time constants progressively decrease with depolarization, with the fast component accounting for 60–80% of the current decay for voltages between –40 and +70 mV (Fig. 5A, inset). At depolarized voltages where Na_v1.8 channels predominately inactivate from the open state (>0 mV), both fast and slow inactivation significantly contribute to the time course of the current decay, similarly to what has been previously described for the native TTX-R current of DRG sensory neurons (Elliott and Elliott 1993).

View larger version (28K): [in this window] [in a new window]   FIG. 5. Properties of the heterologously expressed Nav1.8 channels. Nav1.8 and 1 subunits were co-expressed in tsA201 cells and preincubated 24 h with lidocaine prior to recording Na current. Test pulses were applied to voltages between –50 and +70 mV and time course of the current decay with a sum of 2 exponential components. A: plot of the fast (fast) and slow (slow) time constants of current decay (n = 8). Inset: relative amplitude of the fast component (Afast) as a function of test potential. B: comparison of the steady-state activation and inactivation of controls and after 24-h preincubation with lidocaine. The Na conductance was calculated from the peak Na current measured over a range of potentials, normalized to the conductance at +95 mV, and plotted vs. the test voltage. The smooth curves are fits to a Boltzmann function with midpoints and slope factors (V1/2/kv) of 1.5 ± 2.3 mV/17.5 ± 0.9 mV for the Nav1.8/1 transfected controls (n = 9) and –0.6 ± 1.9 mV/13.6 ± 1.0 mV for cells preincubated with lidocaine (n = 12). Steady-state inactivation was determined using 500-ms prepulses to voltages between –140 and +5 mV. Steady-state availability was assayed using a standard test pulse (15 mV/20 ms). The test current amplitudes were normalized to the test currents measured after prepulses to –140 mV (I/Imax) and plotted vs. voltage. The smooth curves are fits to a double Boltzmann function {I/Imax = a*(1/{1+exp[(V – V1/21)/kv1]}) + [(1 – a)*(1/{1+exp[(V – V1/22)/kv2]})]} with midpoints and slope factors (V1/2/kv) of –71.6 ± 1.2 mV/9.4 ± 0.7 mV for the more hyperpolarized component and –32.9 ± 1.8 mV/9.4 ± 1.0 mV for depolarized component (n = 14). Also plotted is the steady-state availability of the drug-free controls, which had a midpoints and slope factors of –73.4 ± 2.8 mV/10.8 ± 1.3 mV for the more hyperpolarized component and –31.3 ± 3.4 mV/11.5 ± 1.9 mV for the depolarized component (n = 7).

View larger version (25K): [in this window] [in a new window]   FIG. 6. Slow inactivation is an important determinant of steady-state availability. Nav1.8 and 1 subunits were expressed in tsA201 cells and exposed to 1 mM lidocaine for 24 h prior to recording. The steady-state availability was determined by applying prepulses of variable duration (50, 100, or 500 ms) to voltages between –140 and +5 mV. A standard test pulse (20 mv/20 ms) was then used to assay availability. The test currents were normalized and plotted vs. the prepulse voltage. A: steady-state availability was determined using the 50-, 100-, and 500-ms prepulse durations. The smooth curves are fits to a double Boltzmann function with the parameters listed in Table 1. B: steady-state slow inactivation was determined by applying 500 ms to voltages between –140 and +5 mV. A short hyperpolarizing pulse (10 ms/–140 mV) was applied to permit recovery from fast inactivation and standard test pulse used to assay availability (+15 mV/20 ms). The test currents were normalized and plotted vs. voltage. The smooth curve is a fit to a Boltzmann function [I/Imax = ((1 – A)/{1 + exp[(V – V1/2)/kv]} + A)] with midpoint (V1/2), slope factor (kv) and steady-state availability (A) of –71.6 ± 1.8 mV, 11.8 ± 1.0 mV, and 0.61 ± 0.02, respectively (n = 12).

View larger version (24K): [in this window] [in a new window]   FIG. 7. Time course of Nav1.8 closed-state inactivation and recovery. A: development of closed-state inactivation was determined by depolarizing to voltages between –80 and –60 mV for variable intervals (1 s) from a holding potential of –120 mV. The fractional inactivation was assessed by applying a standard test pulse (+15 mV/20 ms), and the normalized test currents plotted vs. the prepulse duration. The smooth curves are the sum of 2 exponential components with fast and slow time constants of 16.4 ± 2.2 ms and 188.1 ± 19.8 at –80 mV (n = 19) with the amplitude of the fast components, Af = 0.21 ± 0.01; 15.2 ± 1.7 ms and 211.1 ± 22.2 ms at –70 mV (n = 18), Af = 0.31 ± 0.03; and 11.0 ± 0.8 ms 161.2 ± 22.7 ms at –60 mV (n = 20), Af = 0.24 ± 0.04. B: time course of recovery from inactivation was determined by first applying depolarizing prepulses (+15 mV/20 ms) before returning to a hyperpolarized potential (Vrec = –120, –110, –100 mV) for a variable interval (1 ms –1 s) to promote recovery. The fractional recovery at each time point was assessed with a standard test pulse (+15 mV/20 ms). The test current amplitudes were normalized and plotted versus the recovery interval. The smooth curves fits of the data to the sum of 2 exponential components with fast and slow time constants of: 1.8 ± 0.3 and 12.7 ± 3.4 ms at –120 mV (Af = 0.53, n = 11), 2.2 ± 0.3 ms and 20.6 ± 4.3 at –110 mV (Af = 0.6, n = 11), and 2.7 ± 0.5 ms and 29.8 ± 6.1 ms at –100 mV (Af = 0.6, n = 11).

The biexponential kinetics of closed-state inactivation (Fig. 7A), the recovery (Fig. 7B), and the biphasic Boltzmann relationship (Fig. 6A) indicate that both fast and slow inactivation are important determinants of the steady-state availability of the Na_v1.8 channel.

Na_v1.8 single-channel currents

Figure 8, A and B, shows single-channel recordings of the Na_v1.8 channel heterologously expressed in tsA201 cells. At –10 mV, the channels rapidly open near the beginning of the voltage pulses and appear to close and repeatedly re-open during the 160-ms depolarization. Figure 8C shows the ensemble average reconstructed from the single-channel openings at –10 mV, which displayed a typical slow inactivation time course similar to what was observed for the macroscopic Na_v1.8 current (Fig. 3B). Repeated opening and comparatively long open times accounts for the slow time-dependent current decay and may underlie the persistent component of Na_v1.8 current observed at depolarized voltages (Fig. 3A). This contrasts with test pulses to –30 mV, which are near the foot of the conductance-voltage relationship (Fig. 5B). Consequently the channels tend to open later during the voltage pulse, the first latency at –10 mV, was estimated to 6.4 ± 1.0 ms, and have shorter open times (0.50 ± 0.01 ms), presumably due to rapid closing at the more hyperpolarized potential.

View larger version (29K): [in this window] [in a new window]   FIG. 8. Single-channel properties of heterologously expressed Nav1.8. Nav1.8 and 1 subunit were co-expressed in tsA201 cells and exposed to 1 mM lidocaine for 24 h prior to recording. A and B: representative single-channel currents recorded in cell-attached configuration at –10 mV (A) and –30 mV (B). Currents were elicited from a holding potential of –120 mV. C: ensemble average of the single-channel openings measured at –10 mV. D: all-points histogram of the single-channel current amplitudes at –10 mV. The smooth curve is a Gaussian fit with amplitude peaks at 0.75 and –10 mV. E: plot of the current-voltage relationship. The straight line is a linear regression yielding a unitary conductance of 11.0 ± 0.4 pS. Data represents mean amplitudes obtained from 9 to 14 individual patches.

Properties of the TTX-resistant Na current of DRG neurons

Preincubating with lidocaine increased the cell surface expression (Fig. 1) and Na current density (Fig. 3) of tsA201 cells transiently transfected with the Na_v1.8 channel. Although these data are consistent with an increase in the expression of functional Na_v1.8 channels in the plasma membrane, it is important to ascertain that the resulting Na currents have properties that are similar to the native TTX-R current of DRG neurons. We therefore directly compared the properties of the heterologously expressed Na_v1.8 and native DRG Na current. Figure 9 shows whole cell Na current recordings from a dissociated DRG neuron before and after external application of TTX (Fig. 9). TTX reduced the peak current amplitude and slowed the kinetics of current decay, consistent with the selective inhibition of the rapidly gating TTX-S Na current expressed in these neurons. Figure 9D shows the current-voltage relationships of these Na currents. Consistent with what has been previously reported, the TTX-R current of DRG neurons activates at more depolarized voltages and displays slow gating kinetics by comparison to the TTX-S current (Elliott and Elliott 1993).

View larger version (33K): [in this window] [in a new window]   FIG. 9. TTX-resistant Na current of dorsal root ganglion (DRG) neurons. Neurons were isolated from the rat DRG. A: whole cell Na current of a small (17 pf) DRG neuron elicited by depolarizing pulses between –70 and +55 mV in 5-mV increments from a holding potential of –80 mV. B: TTX-resistant (TTX-R) current measured after the bath application of 300 nM TTX. C: TTX-sensitive (TTX-S) current obtained by subtracting the TTX-R component from the total current. D: current-voltage relationships of the Total, TTX-R and TTX-S components of DRG Na current (n = 11).

View larger version (23K): [in this window] [in a new window]   FIG. 10. Voltage-dependent gating of the TTX-R Na current of DRG neurons. A: conductance [G = I/(V – Vr)] was determined from the peak Na current (I) at each test voltage (V) and the reversal potential (Vr) determined from the current-voltage relationships (Fig. 9D). The conductance was normalized to that determined at +60 mV and plotted vs. voltage. The smooth curve is a fit to a Boltzmann function with midpoint (V1/2) and slope factor (kv) of –13.8 ± 1.7 and 6.0 ± 0.6 mV, respectively (n = 11). Steady-state inactivation was determined using 500-ms prepulses to the indicated voltages. The smooth curve is a fit of the data to a double Boltzmann function with V1/2 and kv of –77.13 ± 4.0 and 16.0 ± 0.51 mV for the hyperpolarized component (AH = 0.12 ± 0.02) and –31.6 ± 1.5 and 4.1 ± 0.2 mV for the more depolarized component (n = 8). Parameters are listed in Table 1. B: time course of recovery from inactivation. Depolarizing prepulses (20 mV/75 ms) were applied before returning to a holding potential of –120 mV for a variable interval (0.05–1 s). The fractional recovery was determined using a standard test pulse (20 mV/10 ms). The test currents were normalized to the current measured after >1 s at –120 mV and plotted vs. recovery interval. The smooth curve is a fit to the sum of 2 exponentials with fast and slow time constants () of 0.9 ± 0.38 and 74.5 ± 12.1 ms (Af = 0.74 ± 0.05, n = 3). C: time course of rapid inactivation. The decay phase of the TTX-R current measured at voltages more depolarized than –25 mV was fitted with the sum of 2 exponential components and the resulting fast (fast) and slow (slow) time constants plotted vs. voltage (n = 8).

	DISCUSSION

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The Na_v1.8 channel is preferentially expressed in unmyelinated (C-type) and thinly myelinated (A) fibers of peripheral nerve and significantly contributes to the rapid upstroke and conduction of nociceptor action potentials (Renganathan et al. 2001). Insights into the biophysical properties of the Na_v1.8 channel have been largely obtained from studies of the endogenous TTX-R Na current of small cell bodies isolated from the DRG (Elliott and Elliott 1993). Unfortunately, DRG sensory neurons express multiple components of TTX-R Na currents that complicate the detailed characterization of voltage dependence and kinetics of the individual currents (Renganathan et al. 2001). Establishing an approach for heterologously expressing Na_v1.8 in cells that are devoid of endogenous Na current would enable a more-detailed characterization of the channel. Unfortunately, the heterologously expression of the cloned Na_v1.8 channel in mammalian cell lines has proved to be problematic (Choi et al. 2004; Fitzgerald et al. 1999; John et al. 2004; Okuse et al. 2002; Zhou et al. 2003; Vijayaragavan et al. 2001; Vijayaragavan et al. 2004). Na_v1.8 transfection of cultured cell lines often results in peak Na currents (<1 nA) that are considerably smaller than the endogenous TTX-R current of isolated DRG sensory neurons (>10 nA). Although the reason for this low level of expression is not known, it may be caused by a trafficking defect that prevents the incorporation of Na_v1.8 channels into the plasma membrane (Okuse et al. 2002). Co-expression of accessory proteins have been shown to increase heterologously expressed Na_v1.8 currents (John et al. 2004; Vijayaragavan et al. 2001); however, the mechanisms that govern the cell surface expression of these channels remains poorly understood.

In this study, we found that preincubating Na_v1.8-transfected tsA201 cells with lidocaine produced a sixfold increase in the amplitude of the TTX-R Na current. To further investigate the underlying mechanism, we directly compared the biophysical properties of the TTX-R currents of lidocaine-treated tsA201 cells with the native TTX-R Na current of dissociated rat DRG neurons. Both the Na_v1.8 and native TTX-R Na currents activated at voltages around –50 mV. The time constants of current decay were similar for both Na currents indicating that the underlying channels slowly inactivate at depolarized voltages where the channels were predominately open (Figs. 5C and 10, A and B). Single-channel recordings of the heterogously expressed channel displayed repetitive opening during sustained depolarization that appears to account for the slow time course of inactivation and the small persistent currents observed in macroscopic recordings (Fig. 3B). The single conductance of heterologously expressed Na_v1.8 was 11 ± 1.0 pS (Fig. 8D) and is in good agreement with that previously reported for the endogenous TTX-resistant Na current of DRG neurons (Rush et al. 1998).

The activation of the heterologously expressed Na_v1.8 and native TTX-R Na currents had midpoints (V_1/2) of –0.6 and –13.8 mV, respectively (Table 1). The relatively hyperpolarized midpoint of activation observed for the endogenous TTX-R current is similar to what has been previously reported (Akiba et al. 2003; Liu et al. 2006). We speculate that the difference in the activation of the heterologously expressed and endogenous Na currents may be related to the expression of additional components of TTX-R current (Na_v1.5 and Na_v1.9) in DRG neurons but not tsA201 cells (Fang et al. 2002; Renganathan et al. 2002). Alternatively, DRG neurons may express additional subunits or accessory proteins that regulate the gating of the Na_v1.8 channel.

The steady-state inactivation of heterologously expressed Na_v1.8 obtained using 500-ms prepulses was fitted by a double Boltzmann function with midpoints of –75.1 ± 1.4 and –34.4 ± 3.3 mV, indicating that two distinct components of inactivation contribute to the availability of the heterologously expressed channels (Table 1). We found that shortening the depolarizing prepulses from 500 to 50 ms had no effect on the midpoints (V_1/2) of inactivation but significantly reduced the relative amplitude of the more hyperpolarized component (Fig. 6A). The data suggested that the longer prepulses recruited channels into slow inactivated states that inactivated over a more hyperpolarized range of voltages. To test this mechanism, we directly measured the steady-state slow inactivation of Na_v1.8 channels (Fig. 6B), which had a midpoint (V_1/2 = –71.6 ± 1.8 mV) that was nearly identical to the more hyperpolarized component of inactivation (V_1/2 = –75.1 ± 1.4 mV) obtained using the conventional protocols for measuring steady-state availability (Fig. 5D). The data indicate that Na_v1.8 channels rapidly enter into slow inactivated states at hyperpolarized voltages and that both fast and slow inactivation are important determinants of the steady-state availability of Na_v1.8 channels. Consistent with this hypothesis is the time course of the development of inactivation at hyperpolarized voltages where Na_v1.8 channels do not open before inactivating, which was found to be biexponential (Fig. 5A). The time constant of the slow component was in a range of 161–211 ms and appears to be associated with the entry of Na_v1.8 channels into a slow inactivated state. The hallmark of Na_v1.8 is its insensitivity to TTX, still our data show that the expressed Na channels were resistant to block by TTX (Fig. 4).

The steady-state availability of the endogenous TTX-R current of DRG neurons was fitted by a double Boltzmann with midpoints of –77.1 and –31.6 mV, similar to what was observed for the heterologously expressed channels (Fig. 10). Although the midpoints of fast and slow inactivation of the Na_v1.8 and TTX-R currents were similar, the relative contributions of these components to the steady-state availability substantially differed. When applying 500-ms depolarizing prepulses the majority (88%) of the native TTX-R current was associated with the more depolarized component (Fig. 10A) versus only 24% for heterologously expressed Na_v1.8 (Fig. 6A). By comparison to the heterologously expressed channels, the endogenous TTX-R channel of DRG neurons appeared to be more resistant to entry into the slow inactivated state. We speculate that this may result from differential regulation of the expressed and endogenous channels by accessory subunits or intracellular signaling pathways that are unique to DRG neurons.

Overall, the slow open-state inactivation (Fig. 5A), the relatively depolarized midpoints of activation and inactivation (Fig. 5B), and the rapid recovery from inactivation (Fig. 7B) are similar for the heterologous expressed Na_v1.8 and native TTX-R Na currents. These data indicate that preincubating tsA201 cells with lidocaine selectively increased the cell surface expression of Na_v1.8 channels but did not significantly alter their biophysical properties. The activation and steady-state inactivation of the Na_v1.8 channel considerably overlap between –60 and 0 mV suggesting that depolarizing window currents may contribute to the resting membrane potential of DRG neurons expressing this channel.

The increase in Na_v1.8 current density produced by lidocaine suggested that preincubating with the drug promoted the translocation of Na_v1.8 channels to the plasma membrane. To further investigate the underlying mechanism, we examined the subcellular distribution of the heterologously expressed Na_v1.8 channels using confocal fluorescent imaging. In control experiments, the Na_v1.8 immunostaining largely overlapped with that of calnexin, a specific marker of endoplasmic reticulum (Fig. 1). When expressed in tsA201 cells the majority of the Na_v1.8 channel protein appears to be trapped within the endoplasmic reticulum, which would account for the relatively low Na current density of the transfected cells (Fig. 3A). Preincubating the transfected cells with lidocaine resulted in a substantial shift of Na_v1.8 immunofluorescence from the cytoplasm to the cell membrane that correlated with an increase in the Na current density (Fig. 3D). Western blot analysis indicated that the increase in cell surface immunostaining and Na current density induced by lidocaine was not associated with a parallel increase in the expression of the Na_v1.8 channel protein (Fig. 2). Rather these findings indicate that the increase in functional Na channels results from the redistribution of preexisting Na_v1.8 channels from the cytoplasmic compartment to the plasma membrane. These data suggest that the overall poor expression of the Na_v1.8 in tsA201 cells, and possibly other mammalian cell lines (Fitzgerald et al. 1999; Okuse et al. 2002), results from a trafficking defect that causes channel protein to become trapped in the endoplasmic reticulum. The local anesthetic lidocaine is a potent inhibitor of the Na_v1.8 channel that is known to act by stabilizing Na channels in nonconducting inactivated states (Chevrier et al. 2004). However, we cannot rule out the possibility that lidocaine can interact with other proteins such as GPCRs (Hollmann et al. 2001). Although the state of Na channel in the ER is not known, we speculate that the lidocaine-modified Na_v1.8 channels are stabilized in a conformational that facilitates the trafficking of the channels to the plasma membrane.

	GRANTS

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The study was supported by the Heart and Stroke Foundation of Québec, Canadian Institutes of Health Research Grant MT-13181. Dr. M. Chahine is an Edwards Senior Investigator (Joseph C. Edwards Foundation).

	FOOTNOTES

 
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.

Address for reprint requests and other correspondence: M. Chahine, Le Centre de Recherche Université Laval Robert-Giffard, 2601 Chemin de la Canardière, Québec, Québec G1J 2G3, Canada (E-mail: mohamed.chahine{at}phc.ulaval.ca)

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