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NEUROPHARMACOLOGY

Use-Dependent Block by Lidocaine but Not Amitriptyline Is More Pronounced in Tetrodotoxin (TTX)-Resistant Na_v1.8 Than in TTX-Sensitive Na⁺ Channels

Department of Anesthesiology, Friedrich-Alexander-University Erlangen-Nuremberg, Erlangen, Germany

Received June 7, 2006; accepted September 26, 2006.

	Abstract

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The excitability of sensory neurons depends on the expression of various voltage-gated Na⁺ channel isoforms. The tetrodotoxin-resistant (TTXr) Na⁺ channel Na_v1.8 accounts for the electroresponsiveness of nociceptive neurons and contributes to inflammatory and neuropathic pain. Na⁺ channel blockers are clinically employed for chronic pain management, but side effects limit their use. There is conflicting information whether their potency to block tetrodotoxin-sensitive (TTXs) and TTXr Na⁺ channels differs. We analyzed the action of lidocaine and amitriptyline on TTXr Na_v1.8 heterologously expressed in ND7/23 cells in comparison with TTXs Na⁺ channels endogenously expressed in ND7/23 cells. TTXr Na_v1.8 and TTXs currents were investigated under whole-cell voltage-clamp. At a holding potential of –80 mV, lidocaine was 5-fold and amitriptyline 8-fold more potent to tonically block TTXs than Na_v1.8 currents. This was due to a higher percentage of TTXs channels residing in the inactivated, high-affinity state at this potential. Tonic block of either resting or inactivated channels by lidocaine or amitriptyline revealed little differences between TTXs and Na_v1.8 channels. Use-dependent block by amitriptyline was similar in TTXs and Na_v1.8 channels. Surprisingly, use-dependent block by lidocaine was more pronounced in Na_v1.8 than in TTXs channels. This result was confirmed in dorsal root ganglion neurons and is associated with the greater tendency of Na_v1.8 to enter a slow inactivated state. Our data suggest that lidocaine could selectively block Na_v1.8-mediated action potential firing. It is conceivable that the expression pattern of Na⁺ channels in sensory neurons might influence the efficiency of Na⁺ channel blockers used for chronic pain management.

Detailed pharmacological characterization of recombinant Na_v1.8 is rare due to difficulties to functionally express this channel in mammalian cell lines. Recently, ND7/23 cells have been used successfully to express recombinant Na_v1.8 channels (Choi et al., 2004; John et al., 2004). In the present study, we employed this expression system and asked whether Na_v1.8 and TTXs channels indeed display different sensitivities toward two drugs used clinically for the treatment of neuropathic pain, the local anesthetic lidocaine and the tricyclic antidepressant amitriptyline.

	Materials and Methods

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Transient Transfection. Rat Na_v1.8-cDNA was subcloned into the mammalian expression vector pCMV-Script (Stratagene, La Jolla, CA). The dorsal root ganglion neuroblastoma hybridoma cell line ND7/23 was purchased from the European Collection of Animal Cell Cultures (Porton Down, UK). ND7/23 cells were maintained in Dulbecco's modified Eagle's medium (GIBCO-Invitrogen, Karlsruhe, Germany), supplemented with 100 U/ml penicillin/streptomycin (GIBCO-Invitrogen), 25 mM HEPES (GIBCO-Invitrogen), 10% heat-inactivated fetal bovine serum (GIBCO-Invitrogen), and 3 mM taurine (Sigma-Aldrich Chemie GmbH, Steinheim, Germany) at 37°C and 5% CO₂. Cells were transiently transfected with rat Na_v1.8-pCMV-Script (5–10 µg), rat 1-pcDNA1/Amp (5 µg), and CD8-pih3m reporter plasmid (1 µg), or 1-pcDNA1/Amp and reporter plasmid alone (to investigate endogenously expressed TTXs channels) by the calcium phosphate precipitation method. After incubation for 12 to 15 h, cells were replated in 35-mm culture dishes. Transfected cells were used for experiments within 3 to 4 days. Transfection-positive cells were identified by immunobeads (anti-CD-8 Dynabeads; Dynal Biotech, Oslo, Norway). Transfection efficiency was 20% on average for Na_v1.8. Activation and inactivation kinetics were also studied in untransfected ND7/23 cells that were maintained as described above and replated in 35-mm culture dishes the day preceding the experiments.

Cell Culture. Littermates of heterozygous C57/BL6 Na_v1.8-null mice were genotyped using commercially available primers. Animals were killed by CO₂ asphyxiation. DRGs were removed from adult (20–30 g) C57/BL6 wild-type or Na_v1.8-null mice and collected in Dulbecco's modified Eagle's medium. The DRGs were treated with papain (20 U/ml; Sigma) for 20 min followed by collagenase (0.28 U/ml; Biochrom, Berlin, Germany) for 30 min. Cells were dispersed by trituration in Ham's F12 medium supplemented with 10% horse serum, penicillin (100 U/ml), streptomycin (100 µg/ml), and L-glutamine (3.0 mM). Cells were plated on glass coverslips coated with poly-L-lysine and kept at 37°C and 5% CO₂. Cells were used for experiments within 24 h after plating.

Electrophysiological Technique and Data Acquisition. Whole-cell patch-clamp recordings were conducted at room temperature (21°C). Patch pipettes were pulled from borosilicate glass tubes (TW150F-3; World Precision Instruments, Berlin, Germany) and heat-polished at the tip to give a resistance of 0.8 to 1.5 M. Currents were acquired with an Axopatch 200B patch-clamp amplifier (Axon Instruments/Molecular Devices, Sunnyvale, CA), filtered at 5 kHz, and sampled at 20 kHz. The offset potential was zeroed before the electrode was attached to the cell. Voltage errors were minimized using 70 to 80% series resistance compensation, and the capacitance artifact was canceled by using the computer-controlled circuitry of the patch clamp amplifier. Linear leak subtraction, based on resistance estimates from four hyperpolarizing pulses applied before the test pulse, was used for all voltage-clamp recordings except for the experiments on use-dependent block. pCLAMP 8.2 software (Axon Instruments/Molecular Devices) was used for acquisition and analysis of currents. Microcal Origin 6.1/7 software (OriginLab Corp., Northampton, MA) was used to perform least-squares fitting and to create figures. Data are presented as mean ± S.E.M. or fitted value ± S.E. of the fit. An unpaired Student's t test (SigmaStat; SPSS Science, Chicago, IL) was used to evaluate the significance of changes in mean values. p < 0.05 was considered statistically significant.

Chemicals and Solutions. Experiments on ND7/23 cells were performed with an external solution containing 65 mM NaCl, 85 mM choline Cl, 2 mM CaCl₂, and 10 mM HEPES (adjusted to pH 7.4 with tetramethylammonium hydroxide) and a pipette solution containing 100 mM NaF, 30 mM NaCl, 10 mM EGTA, and 10 mM HEPES (adjusted to pH 7.2 with CsOH). The reversed Na⁺ gradient was used to minimize the series resistance artifact, which is less serious with outward currents. In some experiments, more physiological ionic conditions were employed that generate inward currents at depolarizing pulses more negative than 60 to 70 mV. In these experiments, the external solution contained 140 mM NaCl, 3 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, and 10 mM HEPES (adjusted to pH 7.4 with tetramethylammonium hydroxide), and the pipette solution contained 140 mM CsF, 1 mM EGTA, 10 mM NaCl, and 10 mM HEPES (adjusted to pH 7.2 with CsOH). Experiments on DRG neurons were performed with an external solution containing 40 mM NaCl, 100 mM choline Cl, 3 mM KCl, 1 mM MgCl₂, 1 mM CaCl₂, and 10 mM HEPES (adjusted to pH 7.4 with tetramethylammonium hydroxide) and a pipette solution containing 140 mM CsF, 1 mM EGTA, 10 mM NaCl, and 10 mM HEPES (adjusted to pH 7.2 with CsOH). The osmolarity of all solutions was adjusted to 300 to 310 mOsm. Lidocaine hydrochloride and amitriptyline hydrochloride (both Sigma-Aldrich) were dissolved in dimethyl sulfoxide to give stock solutions of 100 mM. The highest dimethyl sulfoxide concentration obtained was 1% and had no effect on Na⁺ currents. TTX (Tocris, Ellisville, MO) was dissolved in water to give a stock solution of 5 mM. Control and test solutions were applied through a gravity-driven polytetrafluoroethylene-glass multiple-barrel perfusion system allowing superperfusion of the cell under investigation.

	Results

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Biophysical Properties of Heterologously Expressed Na_v1.8 and Endogenously Expressed TTXs Channels in ND7/23 Cells. As an expression system for this study, we chose the ND7/23 hybrid cell line derived from neonatal rat DRG neurons fused with the mouse neuroblastoma N18Tg2, that exhibit sensory neuron-like properties (Wood et al., 1990). ND7/23 cells endogenously express sodium currents with rapid kinetics that are completely blocked by 250 nM TTX. The molecular identities of these TTXs currents are unknown; however, they exhibit similar kinetics and activation and inactivation properties to the TTXs currents in DRGs (John et al., 2004).

View larger version (30K): [in this window] [in a new window]   Fig. 1. TTXr Nav1.8 and TTXs Na+ currents. Representative TTXr Na+ currents in ND7/23 cells following transient transfection of Nav1.8 and 1-subunit, recorded in the presence of 250 nM TTX in the extracellular solution (A), and TTXs Na+ currents in ND7/23 cells transfected with 1-subunit only (B). Vh was –140 mV. The standard ionic conditions used in this study generate outward currents at depolarizing pulses more positive than –20 to –10 mV (calculated ENa = –17.5 mV) (A and B). In the second line of traces, TTXre-Nav1.8 currents (C) and TTXs Na+ currents (D) are shown recorded at Vh = –80 mV and under ionic conditions that generate inward currents at depolarizing pulses more negative than 60 to 70 mV (calculated ENa = 67 mV). Voltage dependence of activation (E) and steady-state inactivation (F) of Nav1.8 () and TTXs currents in cells transfected with 1 unit () or untransfected cells () at Vh = –140 mV. For activation, currents were evoked by 5-ms-long test pulses from –120 to 60 mV in steps of 10 mV. Conductance was determined at a given voltage step from the equation gm = INa/(Em – Erev), where INa is the peak current, Em is the amplitude of the voltage step, and Erev is the estimated reversal potential. The data were fitted with a Boltzmann function. Note the drop in the normalized peak Na+ conductance in TTXs currents at potentials >+30 mV. The average midpoint voltages and slope factors are given in Table 1. The h curves were determined by a two-pulse protocol: 100-ms-long prepulses from –120 to 40 mV in steps of 5 mV were followed by a test pulse to 50 mV. Peak currents were measured at the test pulse, normalized, and plotted against the prepulse potential. The data were fitted with a Boltzmann function.

Activation and inactivation kinetics were characterized by standard pulse protocols. Holding potential (V_h) was –140 mV. The standard ionic conditions used in this study generated outward currents at depolarizing pulses more positive than –20 to –10 mV [calculated sodium equilibrium potential (E_Na) = –17.5 mV] (Fig. 1, A and B). Activation and inactivation curves are shown in Fig. 1, E and F, and mid-point potentials of peak Na⁺ conductance and of steady-state inactivation are given in Table 1. Voltage dependence of activation and steady-state inactivation were similar when cells were held at V_h = –80 mV or when more physiological ionic conditions were employed that generated inward currents at depolarizing pulses more negative than 60 to 70 mV (E_Na = 67 mV) (Fig. 1, C and D; Table 1). Coexpression of 1 did not influence activation or inactivation kinetics of TTXs currents in ND7/23 cells. The data for activation and steady-state inactivation found in the present study are in reasonable agreement with values found previously for Na_v1.8 heterologously expressed in ND7/23 cells (Choi et al., 2004; John et al., 2004) and closely resemble those for native TTXr currents in DRG neurons (Cummins and Waxman, 1997; John et al., 2004).

View larger version (28K): [in this window] [in a new window]   Fig. 2. Tonic block of Nav1.8 and TTXs channels in ND7/23 cells by lidocaine and amitriptyline. Representative traces of Nav1.8 (A) and TTXs (B) currents in the absence and presence of various concentrations of lidocaine (Vh = –80 mV). Currents were activated by 5-ms-long test-pulses to 50 mV in intervals of 30 s. Concentration-dependent block of Nav1.8 and TTXs currents by lidocaine (C and E) and amitriptyline (D and F) at Vh = –80 (C and D) or –140 (E and F) mV. The peak amplitudes of Na+ currents were measured in different drug concentrations, normalized with respect to the peak amplitude in control, and plotted against the drug concentration. Solid lines, fits of the data to the Hill equation. IC50 values and Hill coefficients are given in Table 1.

View larger version (19K): [in this window] [in a new window]   Fig. 3. State-dependent block of Nav1.8 and TTXs channels by amitriptyline. A and B, normalized Nav1.8 (squares) and TTXs currents (circles) as a function of conditioning prepulse potentials (A and B). The pulse protocol is shown below. In detail, 10-s conditioning prepulses from –160 to –30 mV were applied. Intervals of 100 ms at the holding potential of –140 mV were inserted before test pulses to 50 mV were applied to activate Na+ currents. Pulses were applied at intervals of 30 s. Control currents (open symbols) were normalized to the current obtained with a prepulse to –160 mV. Currents obtained in the presence of 1 µM amitriptyline (solid symbols) were normalized to the current obtained in control with the corresponding prepulse potential. Solid lines, fits of the data to a Boltzmann function. B, concentration-dependent block of inactivated Nav1.8 and TTXs channels by amitriptyline. Na+ currents were activated by 5-ms-long test pulses to 50 mV after 10 s long prepulses to –40 mV for Nav1.8 or –70 mV for TTXs channels and a 100-ms-long interval at the holding potential. IC50 values were calculated as described in Fig. 2.

To assess in detail state-dependent block of Na_v1.8 and TTXs channels by amitriptyline, a three-step pulse protocol was applied as described previously (Wright et al., 1997; Nau et al., 1999). Cells were held at –140 mV, and 10-s-long conditioning prepulses between –160 and –30 mV were applied to allow amitriptyline binding to Na⁺ channels to reach steady state. After a 100-ms-long interval at –140 mV, inserted to allow drug-free channels to recover from fast inactivation, drug-free channels were activated by test pulses to 50 mV. In Fig. 3A, normalized peak currents of Na_v1.8 and TTXs channels in control and in the presence of 1 µM amitriptyline are plotted as a function of the conditioning prepulse potentials. In control solution, a small fraction of Na⁺ channels exhibited slow inactivation after prepulses > –90 mV. Block by 1 µM amitriptyline was absent or small at prepulses <–100 mV for Na_v1.8 and <–110 mV for TTXs channels, respectively. This block is deemed to correspond to block of resting channels. Block by 1 µM amitriptyline reached a second plateau after prepulse >–40 mV for Na_v1.8 and >–70 mV for TTXs channels. This block is deemed to correspond to high-affinity binding and block of inactivated channels (Fig. 3A). State-dependent block by amitriptyline could be fitted by a Boltzmann equation with midpoint (millivolt) and slope factor (millivolt per e-fold) values, respectively, of –75.9 ± 0.8 and 11.3 ± 0.7 for Na_v1.8 and –91.7 ± 0.9 and 7.7 ± 0.8 for TTXs currents, respectively. Note that the midpoint values critically depend on the fractional distributions of resting and inactivated channels at a given conditioning potential (Wright et al., 1999). To more accurately determine the sensitivities of inactivated channels to amitriptyline, we measured the concentration dependence of block with conditioning prepulses to –40 mV for Na_v1.8 and to –70 mV for TTXs channels and determined the IC₅₀ values (Fig. 3B). As demonstrated in Fig. 3B, inactivated TTXs currents displayed an approximately 1.3-fold higher sensitivity (p < 0.05) to amitriptyline compared with Na_v1.8 (Table 1).

The recovery time course of Na⁺ channels from the inactivated drug-bound state is reported to be slow in the presence of amitriptyline (Nau et al., 2000; Wang et al., 2004). Because recovery from block by lidocaine is much faster, significant amounts of lidocaine unbinding from inactivated channels would occur during the 100-ms-long interval at –140 mV between the conditioning prepulse and the test pulse (Wright et al., 1997), leading to an underestimation of block of inactivated channels when using the pulse protocol as described above. To confirm this prediction under our experimental conditions, we determined the recovery time of inactivated Na_v1.8 and TTXs channels from block by amitriptyline and lidocaine (Fig. 4) by applying a 5-ms test pulse to +50 mV from a holding potential of –140 mV at various times after a 10-s conditioning prepulse to –40 mV for Na_v1.8 and to –70 mV for TTXs channels (Fig. 4, inset). Figure 4, A and B, show normalized peak currents of Na_v1.8 and TTXs channels in control and in the presence of 1 µM amitriptyline (n = 5 and 9, respectively) and 50 µM lidocaine (n = 8 and 7, respectively) as a function of the interpulse duration.

View larger version (21K): [in this window] [in a new window]   Fig. 4. Recovery of inactivated Nav1.8 (A) and TTXs channels (B) from block by amitriptyline and lidocaine. The pulse protocol is shown below. In detail, cells were conditioned by 10-s-long prepulses to –40 mV for Nav1.8 and to –70 mV for TTXs channels. Currents were activated by applying a 5-ms test pulse to +50 mV at various times after the conditioning pulse. The peak currents evoked by the test pulse were measured, normalized with respect to the peak amplitude of the test pulse obtained after 50-s recovery time, and plotted against the recovery time. The data were well fitted by a double-exponential function.

The time constants are given in Fig. 4, A and B. It is apparent that in the control, a greater fraction of TTXr Na_v1.8 than TTXs channels recovers from inactivation with a second time constant. This indicates that TTXr Na_v1.8 channels are more prone to enter a slow inactivated state than TTXs channels. The fractional amplitudes of the second phase of recovery of Na_v1.8 currents under amitriptyline and lidocaine were 64 and 65%, respectively (Fig. 4A). The fractional amplitudes of the second phase of recovery of TTXs currents under amitriptyline and lidocaine were 81 and 24%, respectively.

These results clearly show that at 100-ms recovery time, little recovery of inactivated and blocked Na_v1.8 or TTXs channels had occurred in the presence of amitriptyline. In the presence of lidocaine, fast and slow components of recovery could not be separated adequately. This confirms the presumed significant differences in kinetics of block by amitriptyline and lidocaine and validates the pulse protocol used to estimate block of inactivated channels by amitriptyline but not lidocaine. To estimate block of inactivated channels by lidocaine, we employed an indirect approach, which is based on the concentration-dependent shift of the steady-state inactivation curve (Bean et al., 1983; Leuwer et al., 2004) and can be expressed by the equation:

(1)

where V_0.5 is the shift in midpoint of the steady-state inactivation curve, k is the slope factor of the steady-state inactivation curve derived from a Boltzmann fit, [L] is the concentration of lidocaine applied, IC_50R is the IC₅₀ value for resting channels, obtained in concentration-inhibition experiments at V_h = –140 mV, and K_I is the dissociation constant for block of inactivated channels by lidocaine. We measured steady-state inactivation of Na_v1.8 and TTXs channels at V_h = –140 mV as described in Fig. 1F in control and in the presence of 50 µM lidocaine (data not shown). 50 µM lidocaine induced a reversible hyperpolarizing shift of the midpoint by –9.7 mV in Na_v1.8 and by –5.8 mV in TTXs channels. The slope factors k were not changed significantly; thus, the mean from control and lidocaine data were taken to calculate K_I, yielding estimated values of 41 and 46 µM for Na_v1.8 and TTXs channels, respectively. The differences of these K_I values from the IC₅₀ value for block of TTXs channels by lidocaine obtained at V_h = –80 mV (30 ± 2 µM) might be explained by the fact that the model tends to overestimate the effect of low drug concentrations and to underestimate the effect of high drug concentrations. However, as measured for amitriptyline, the data demonstrate that inactivated Na_v1.8 and TTXs channels displayed similar sensitivities to lidocaine as well.

View larger version (28K): [in this window] [in a new window]   Fig. 5. Use-dependent block of Nav1.8 and TTXs channels. Representative currents of Nav1.8 (A) and TTXs (B) channels and both channel populations together (C) evoked by 25-ms-long pulses to 50 mV at a frequency of 10 Hz in the presence of 50 µM lidocaine. Use-dependent block was examined at 2 and 10 Hz with 60 5- or 25-ms-long test pulses to 50 mV from Vh = 140 mV. Development of use-dependent block (25-ms-long test pulses) at 2 and 10 Hz in control solution (D), with 1 µM amitriptyline (E), and with 50 µM lidocaine (F). Peak currents were measured, normalized to the current of the first pulse, and plotted against the pulse number. Block at pulse number 60 with respect to the current of the first pulse (5 or 25 ms) in control solution (G), with 1 µM amitriptyline (H), and with 50 µM lidocaine (I). An unpaired Student's t test (SigmaStat; SPSS Science) was used to evaluate the significance of changes in mean values. *, p < 0.05 was considered statistically significant.

We additionally investigated use-dependent block by lidocaine or amitriptyline on both channel populations in the absence of TTX (Fig. 5C). Currents clearly showed two components with a fast one due to TTXs channels and a slower one due to TTXr Na_v1.8 channels. Quantifying use-dependent block by measuring peak currents revealed results similar to the results obtained with TTXs currents (Fig. 5, D–I, gray symbols and bars). Quantifying use-dependent block by measuring current amplitudes 0.8 ms after current activation, a time point at which TTXs channels are already inactivated, revealed results similar to the results obtained with TTXr Na_v1.8 channels (data not shown). Apparently, lidocaine as well as amitriptyline affect TTXs and TTXr Na_v1.8 channels independently.

View larger version (26K): [in this window] [in a new window]   Fig. 6. Concentration dependence of use-dependent block of Nav1.8 and TTXs channels. Development of use-dependent block (25-ms-long test pulses, 10 Hz) in control solution and with 0.2, 1, and 10 µM amitriptyline (A) or 10, 50, and 500 µM lidocaine (B). Decaying normalized peak currents were fitted by a single-exponential function after subtracting respective control currents. Time constants of current decays are given in the figure. Block at pulse number 60 with respect to the current of the first pulse by various concentrations of amitriptyline (C) or lidocaine (D), respectively. An unpaired Student's t test (SigmaStat; SPSS Science) was used to evaluate the significance of changes in mean values. *, p < 0.05 was considered statistically significant.

Comparable with results obtained with 1 µM amitriptyline, Na_v1.8 and TTXs channels exhibited a similar degree of use-dependent block in the presence of 0.2 and 10 µM amitriptyline, respectively (Fig. 6C). In addition, comparable with results obtained with 50 µM lidocaine, Na_v1.8 channels exhibited a higher degree of use-dependent block compared with TTXs channels in the presence of 10 and 500 µM lidocaine, respectively (Fig. 6D).

To further confirm disparate use-dependent blocking properties of lidocaine and amitriptyline toward TTXs and TTXr Na_v1.8 channels under more native conditions, we also investigated use-dependent block of Na⁺ currents in DRG wild-type neurons in the presence of TTX, in DRG neurons of Na_v1.8 knockout mice, and in DRG wild-type neurons in the absence of TTX (Fig. 7). The results obtained by these experiments resemble those obtained by experiments in ND7/23 cells. Specifically, use-dependent block by lidocaine was more pronounced in TTXr channels of wild-type mice than in TTXs channels of Na_v1.8 knockout mice, whereas use-dependent block by amitriptyline was similar in these channel populations.

View larger version (27K): [in this window] [in a new window]   Fig. 7. Use-dependent block of TTXr and TTXs channels in DRG neurons. Representative TTXr currents (A), Na+ currents in Nav1.8 –/– neurons (B) and in wild-type neurons (C) evoked by 25-ms-long pulses to 0 mV at a frequency of 10 Hz in the presence of 50 µM lidocaine. Use-dependent block was examined at 2 and 10 Hz with 60 5- or 25-ms-long test pulses to 0 mV from Vh = 140 mV. Development of use-dependent block (25-ms-long test pulses) at 2 and 10 Hz in control solution (D), with 1 µM amitriptyline (E), and with 50 µM lidocaine (F). Peak currents were measured, normalized to the current of the first pulse, and plotted against the pulse number. Block at pulse number 60 with respect to the current of the first pulse (5 or 25 ms) in control solution (G), with 1 µM amitriptyline (H), and with 50 µM lidocaine (I). An unpaired Student's t test (SigmaStat; SPSS Science) was used to evaluate the significance of changes in mean values. *, p < 0.05 was considered statistically significant.

	Discussion

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Expression of Na_v1.8 in the ND7/23 Cell Line and Coexpression of the 1 Subunit. The TTXr Na_v1.8 channel is the molecular entity underlying the slowly inactivating TTXr current of DRG neurons. Unlike other sodium channels, recombinant Na_v1.8 is poorly expressed in most mammalian cell lines, even in the presence of accessory -subunits. Annexin II light chain (p11), among other interacting proteins, has been identified as an essential regulatory factor that directly binds to the N terminus of Na_v1.8 and promotes translocation of the channel to the plasma membrane (Okuse et al., 2002). The hybrid cell line ND7/23 derived from neonatal rat DRG neurons fused with the mouse neuroblastoma N18Tg2 has proven to be a suitable heterologous host cell line for sufficient expression of recombinant Na_v1.8 channels (Choi et al., 2004; John et al., 2004). This cell line might provide the necessary cellular neuronal environment for functional expression of recombinant Na_v1.8.

Four -subunits have been described to date that potentially associate with Na⁺ channel -subunits (Catterall et al., 2003; Yu et al., 2003). -Subunits affect Na⁺ channel gating and cell surface expression when expressed heterologously. However, the effect of coexpression of -subunits depends critically on the cell type in which the expression experiments are performed. In contrast to results in Xenopus oocytes, expression of 1-subunits with Na_v1.2 or Na_v1.4 in human embryonic kidney cells have little or no effect on the kinetics of sodium currents and cause positive shifts in the voltage dependence of activation and inactivation (Wright et al., 1999; Qu et al., 2001). ND7/23 cells reportedly express 1- and 3-subunits (John et al., 2004). We obtained larger Na_v1.8 and TTXs current amplitudes by coexpression of 1. Thus, additional recombinant 1-subunit expression can enhance recombinant or constitutive Na⁺ currents. Expression of Na_v1.8 without coexpression of 1 was rather poor, and detailed analysis of the effect of 1 on current kinetics and voltage dependence of activation and inactivation was hampered. Surprisingly, coexpression of 1 did not influence current kinetics or activation and inactivation properties of TTXs currents in ND7/23 cells.

It was demonstrated previously that tonic block of resting and inactivated channels by the local anesthetic cocaine is not changed significantly by coexpression of the 1-subunit in mammalian cells (Wright et al., 1999). To our knowledge, no data are available on the effect of 1 on use-dependent block of recombinant Na⁺ currents in mammalian cells. Because the 1-subunit has the tendency to reduce the proportion of slow gating, use-dependent block might be enhanced in the absence of 1, especially in Na_v1.8. Clearly, further studies are needed to investigate interaction and effect of 1 and 3 subunits on Na_v1.8 channels.

Similarities in Affinities of Resting and Inactivated Channels. Block of voltage-gated Na⁺ channels by local anesthetics is highly state-dependent with a preference for open and inactivated rather than for resting channel states. In this study, we have demonstrated that tonic block of resting TTXs and TTXr Na_v1.8 channels by lidocaine and amitriptyline, measured at a holding potential of V_h = –140 mV, reveal little differences. At V_h =–80 mV, the fraction of channels residing in the inactivated, high-affinity state is higher for TTXs compared with TTXr Na_v1.8 channels. Consequently, Na⁺ channel blockers are more potent to tonically block TTXs than TTXr currents at a holding potential of V_h = –80 mV, as demonstrated before (Roy and Narahashi, 1992; Scholz et al., 1998). This difference is therefore due to state-dependent drug affinity rather than to binding site differences of TTXr Na_v1.8 and TTXs channel isoforms.

It was demonstrated previously that the binding affinity of inactivated channels could be estimated directly for Na⁺ channel blockers that unbind slowly from inactivated channels with the three-step protocol shown in Fig. 3 (Wright et al., 1997). For amitriptyline, this pulse protocol was applicable because the recovery time course from inactivated drug-bound channel states was slow, as reported before (Nau et al., 2000; Wang et al., 2004). In this study, block of inactivated TTXs and TTXr Na_v1.8 channels by amitriptyline was estimated with conditioning prepulses to –70 and –40 mV, respectively. These prepulses induced some amount of slow inactivated channels with a higher percentage in TTXr Na_v1.8 than in TTXs channels. Consequently, slow inactivated TTXr Na_v1.8 channels might have contributed to block by amitriptyline measured with this approach. Still, TTXs channels displayed approximately 1.3-fold higher sensitivity to amitriptyline than TTXr Na_v1.8 channels.

A different approach was required to estimate block of inactivated channels by lidocaine due to the fast recovery of inactivated channels from block by lidocaine. This approach exploits the concentration-dependent shift of the steady-state inactivation curve by Na⁺ channel blockers (Bean et al., 1983; Leuwer et al., 2004). The K_i values of 41 and 46 µM calculated for Na_v1.8 and TTXs channels, respectively, are higher than the IC₅₀ value measured for block of TTXs channels in concentration-inhibition experiments at V_h =–80 mV (30 ± 2 µM). The same phenomenon was observed when employing this approach for estimating block of inactivated channels by amitriptyline (data not shown) and might be explained by the fact that the model tends to overestimate the effect of low drug concentrations and to underestimate the effect of high drug concentrations. However, based on the evidence demonstrated, we conclude that sensitivities for lidocaine or amitriptyline reveal little differences between inactivated Na_v1.8 and TTXs channels.

Stronger Use-Dependent Inhibition of Na_v1.8 by Lidocaine. The most interesting finding in this study is the significant difference in use-dependent block of TTXr Na_v1.8 and TTXs channels by lidocaine but not by amitriptyline. This phenomenon holds true for several drug concentrations and could be observed in ND7/23 cells expressing TTXs and TTXr Na_v1.8 channels and in DRG neurons of wild-type and Nav1.8 knockout mice. A more pronounced use-dependent block by lidocaine has been observed before for TTXr compared with TTXs channels in native DRG neurons (Roy and Narahashi, 1992) and for TTXr Na_v1.8 compared with TTXs Na_v1.7 channels coexpressed with the 1 subunit in Xenopus oocytes (Chevrier et al., 2004). However, it has not been noticed explicitly that a more pronounced use-dependent block of TTXr Na_v1.8 might be a feature of some but not all Na⁺ channel blockers.

Use-dependent block arises from binding of local anesthetics to inactivated channels recruited during repetitive pulses and from dissociation of local anesthetics from inactivated states with a time constant slower than the frequency of the pulses. Alternatively, it was suggested that use-dependent block results from slow recovery of drug-bound channels due to an interaction between local anesthetics and slow inactivated states (Ong et al., 2000).

Indeed, slow inactivation is more pronounced in TTXr compared with TTXs channels (Rush et al., 1998; Blair and Bean, 2003; Chevrier et al., 2004). Moreover, slow inactivation in TTXr channels was suggested to play a major role in controlling adaptation of action potential firing in nociceptive sensory neurons (Blair and Bean, 2003).

In the present study, slow inactivation in TTXr Na_v1.8 channels is responsible for the greater fraction of TTXr Na_v1.8 channels recovering from inactivation with a slow time constant in control (Fig. 4) and for the more pronounced use-dependent inhibition of TTXr Na_v1.8 currents in the absence of blockers (Figs. 5, D and G, and 6, D and G). Preferential binding of lidocaine to slow inactivated states of TTXr Na_v1.8 might indeed explain the more pronounced use-dependent block of TTXr Na_v1.8 compared with TTXs channels.

In contrast to lidocaine, amitriptyline is a potent use-dependent blocker of both TTXs and TTXr Na_v1.8 currents. Several possible mechanisms could account for this difference in use-dependent block by amitriptyline. The slow recovery time course of fast-inactivated drug-bound Na⁺ channels in the presence of amitriptyline might cover or overlap any interactions with slow inactivated states. Amitriptyline might inhibit any interactions with slow inactivated states. Amitriptyline might prevent Na_v1.8 channels to enter slow inactivated states. Clearly, further studies are needed to support or reject these hypotheses.

Clinical Relevance of the Findings. Following injury of peripheral nerves, the composition of Na⁺ channel isoforms expressed in sensory neurons undergoes significant changes (Lai et al., 2003). Subsequent electrophysiological changes might poise cells to fire spontaneously or at inappropriately high frequencies. Among the changes following nerve injury is an increase in the expression of TTXs Na_v1.3 in cell bodies of sensory neurons (Cummins and Waxman, 1997) and a redistribution of Na_v1.8 and Na_v1.9 from cell bodies of sensory neurons to the peripheral axon at the site of injury (Lai et al., 2002; Gold et al., 2003).

Contribution of Na_v1.8 to alteration of pain threshold is unclear to date. Studies on Na_v1.8 knockout mice indicated that Na_v1.8 might not be involved (Kerr et al., 2001). In addition, neuropathic pain develops normally in mice lacking both Na_v1.7 and Na_v1.8 (Nassar et al., 2005). In contrast, antisense oligonucleotides directed against Na_v1.8 administered i.t. completely reversed neuropathic pain behavior (Lai et al., 2002). Furthermore, Na_v1.8 might play a role in spontaneous neuropathic pain because neuromas in Na_v1.8 knockout mice display less ectopic discharges than in wild-type mice (Roza et al., 2003).

Altogether, it is generally conceived that isoform-specific Na⁺ channel blockers, especially those targeting Na_v1.8 and Na_v1.3, could be useful analgesics. Our data, however, confirm the presumption that it is highly unlikely to find isoform-specific Na⁺ channel blockers interacting with the local anesthetic binding site. On the other hand, they demonstrate that there is significant functional selectivity by lidocaine but not amitriptyline toward slow inactivated Na_v1.8 channel states. This phenomenon could explain the potential of lidocaine to inhibit ectopic activity in peripheral nerves after injury at concentrations that do not block nociception (Devor et al., 1992). Furthermore, the disparate use-dependent drug effects found in this study could be exploited to search for other compounds with more favorable therapeutic profiles. Finally, it is conceivable that the efficiency of Na⁺ channel blockers in chronic pain management might be determined by the distinct and individual expression pattern of Na_v1.8 and TTXs channels in sensory neurons.

	Acknowledgements

 
Na_v1.8-cDNA and heterozygous breeding pairs of Na_v1.8-null mice were kindly provided by John Wood (University College London, London, UK). We thank Miriam Hacker-Rogner for excellent technical assistance and J. Schüttler, H. Schwilden, P. Reeh, and H. O. Handwerker for continuous support.

	Footnotes

 
This work was supported by Deutsche Forschungsgemeinschaft Grants NA 350/2-5 (Emmy Noether-Programm; to C.N.) and NA 350/3-2 (KFO130 to C.N. and A.L.).

Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.

doi:10.1124/jpet.106.109025.

ABBREVIATIONS: TTX, tetrodotoxin; TTXr, tetrodotoxin-resistant; TTXs, tetrodotoxin-sensitive; DRG, dorsal root ganglion.

Address correspondence to: Dr. Carla Nau, Department of Anesthesiology, Friedrich-Alexander-University Erlangen-Nuremberg, Krankenhausstrasse 12, 91054 Erlangen, Germany. E-mail: Carla.Nau{at}kfa.imed.uni-erlangen.de

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