Lidocaine Increases Intracellular Sodium Concentration through Voltage-dependent Sodium Channels in an Identified Lymnaea Neuron

All animal experiments were approved by the Animal Care Committee of Miyazaki Medical College (Kiyotake, Miyazaki, Japan). The cell culture procedures were adopted from Syed et al.  11Specifically, we used individually identifiable RPeD1 neurons from laboratory-raised L. stagnalis  (fresh water snail) at room temperature. For this study, 2- to 3-month-old snails (shell length, 20– 30 mm) were selected. Snails were deshelled and transferred to a sterile dissection dish in antibiotic normal saline (150 μg/ml Gentamicin [G3632; Sigma Chemical Co., St. Louis, MO] in normal Lymnaea  saline: 51.3 mm NaCl, 1.7 mm KCl, 4.1 mm CaCl², 1.5 mm MgCl², 5.0 mm HEPES, pH 7.9). The central ganglionic rings were isolated using standard dissection procedures and subsequently pinned to the silicone rubber base of a tissue culture plate. The outer connective tissue of the ganglia was removed before enzymatic treatment. Ganglia were treated in defined medium (serum-free 50% Leibovitz L-15 medium [GIBCO-BRL Life Technologies, Burlington, Ontario, Canada] with added inorganic salts, 20 μg/ml gentamycin, pH 7.9) for 25 min with 0.2% trypsin (type III; Sigma Chemical Co.). Subsequently, ganglia were also treated with 0.2% soybean trypsin inhibitor (Sigma Chemical Co.) for 15 min in the defined medium. Before removal of identified neurons, the inner connective tissue sheath was dissected with fine forceps from the ganglia. Neurons were removed by gentle suction with a siliconized, microforge fine-polished pipette with an outside diameter of 1.5 mm (IB-150 F; WPI, Sarasota, FL), and then these neurons were maintained in a high-osmolarity medium containing 30 mm glucose. After this, neurons were transferred to poly-l-lysine–coated culture dishes (Falcon Plastics, Los Angeles, CA) with 3 ml of the defined medium.

The following neuronal activity was monitored using conventional intracellular recording. A glass microelectrode, outside diameter of 1.0 mm (GC100-10; CEI, London, England) was filled with a saturated solution of K²SO⁴, yielding a tip resistance of 20–60 MΩ. A neuron was observed under an inverted microscope (TE-300; Nikon, Tokyo, Japan) and impaled by a microelectrode using a manipulator (MM 202; M 204, Narishige, Japan). Electrical signals were amplified (IR-283; Cygnus Technologies, Delaware Water Gap, PA), displayed on an oscilloscope (VC-11; Nihon Kohden, Tokyo, Japan), and stored on computer (G4; Apple Computer Inc., Cupertino, CA) through an A/D converter (Powerlab; ADInstruments, Colorado Springs, CO) and a thermal alley recorder (RTA1200; Nihon Kohden).

For the measurement of [Na⁺]ⁱ, the radiometric fluorescent indicator, an acetoxymethyl ester form of sodium-binding benzofuran isophthalate (SBFI-AM) (Molecular Probes, Eugene, OR) was used. SBFI-AM, 1 mm, in dimethyl sulfoxide (DMSO) was mixed with the defined medium containing the neuron at a volume of 0.1%. After incubation at 20°C for 60 min, this defined medium was removed, and the neuron was suspended in normal saline for 1 h to take out hydrolysis of AM esters and diacetates. Measurements were made by exciting this indicator dye at 340 nm, where fluorescence is particularly sensitive to the sodium ion concentration, and 390 nm, where is very close to the isosbestic point.20–22Therefore, the intensity of intracellular SBFI fluorescence was measured at two quickly alternating excitation wavelengths (340/390 nm) and continuously recorded at 510 nm by an inverted fluorescent microscope (TE-300), a cooled high-speed charge-coupled device video camera (C-6970; Hamamatsu Photonics, Hamamatsu, Japan), and a fluorescence imaging system (Argus-Hisca; Hamamatsu Photonics). The exposure time for a single image was 159 ms, resulting in a total time of 600 ms needed for a 340/390-nm pair image. Background fluorescent images were subtracted before analysis.

Two fluorescence ratios with SBFI were converted into using the calibration curve for RPeD1 in vivo .23–25The calibration curve for [Na⁺]ⁱwas constructed by plotting the fluorescence ratio versus  Na⁺concentration of the calibration solutions (fig. 1). To equilibrate the [Na⁺]ⁱwith extra ([Na⁺]^o), 30 min before the experiment, 10 μm of the Na⁺ionophore gramicidin D (ICN Biomedicals, Inc., Costa Mesa, CA) was added, and then RPeD1 neurons were exposed to Na⁺-free concentration saline with gramicidin D solution for 30 min. Fluorescent image pairs were taken, and then RPeD1 neurons were exposed to salines of increasing sodium concentration (1, 25, 50, 100, 150 mm) for 10 min each. Image pairs were taken again at the end of each exposure.

Neurons were divided into five trials: normal saline, high-Na⁺saline, Na⁺-free saline, membrane hyperpolarization, and tetrodotoxin (Sankyo, Tokyo, Japan; 30 μm) pretreatment. For the membrane hyperpolarization trials, the membrane potential was held at −80 mV by current injection 2 min before lidocaine perfusion. In the high-Na⁺saline trials, high-Na⁺saline (103 mm NaCl, 1.7 mm KCl, 4.1 mm CaCl², 1.5 mm MgCl², 5.0 mm HEPES, pH 7.9) was used as the extracellular fluid. In the Na⁺-free saline trials, Na⁺-free saline (0 mm NaCl, 50 mm N -methyl-d-glucamine, 1.7 mm KCl, 4.1 mm CaCl², 1.5 mm MgCl², 5.0 mm HEPES, pH 7.9) was used. In the normal saline, membrane hyperpolarization, and tetrodotoxin pretreatment trials, normal saline was used.

In the normal saline trials, lidocaine (0.01, 0.1, and 1 mm) was perfused into the culture dish after measurements of baseline values. In the high-Na⁺saline, Na⁺-free saline, membrane hyperpolarization, and tetrodotoxin pretreatment trials, 1 mm lidocaine was perfused. In the tetrodotoxin pretreatment trials, tetrodotoxin (30 μm) was perfused for 4–6 min before lidocaine perfusion.

In the normal saline trials, membrane potential and [Na⁺]ⁱwere measured for 5 min after each dose of lidocaine. In the high-Na⁺saline and the Na⁺-free saline trials, the normal salines were changed to either high-Na⁺or Na⁺-free saline at 2 min, and membrane potential and [Na⁺]ⁱwere measured for 7 min after lidocaine administration. In the normal saline and membrane hyperpolarization trials, the baseline values were collected during the 4 min before lidocaine perfusion (at 0 min in the figures). In the high-Na⁺saline, Na⁺-free saline, and tetrodotoxin pretreatment trials, the baseline values were measured 2 min before lidocaine perfusion (at 0 min in the figures). In each trial, the membrane potential was measured relative to the resting membrane potential of −60 mV in RPeD1.

Whole cell voltage clamp recordings in RPeD1 neurons26,27were made using an EPC9 amplifier (HEKA elektronik, Lambrecht, Germany). Patch electrodes (tip diameter adjusted to 1.0 μm, resistance of 1–3 MΩ) were pulled from glass tubing (outside diameter of 1.5 mm) with no filament (PG-150T-7.5; Warner Instrument, Hamden, CT) on a vertical pipette puller (Kopf 750, Tujunga, CA). For experiment control and data acquisition, an A/D and D/A converter interface board (PCI-16; HEKA elektronik) was inserted into a personal computer (G4). Data acquisition and analysis were performed using Pulse software (version 8.66; HEKA elektronik). The current was filtered at 1 kHz using a four-pole Bessel filter and digitized at a sampling frequency of 20 kHz. To study INa, pipettes were filled with filtered (0.22-μm filter) cesium pipette solution consisting of 50 mm CsCl, 5 mm EGTA, 5 mm MgCl², 10 mm HEPES, 2 mm ATP-Mg, and 0.1 mm GTP-Tris, and the pH was adjusted to 7.4 (with CsOH). The bath solution consisted of 40 mm NaCl, 10 mm TEACl, 4.0 mm MgCl², 1.0 mm CaCl², 1.0 mm 4-aminopyridine, 1.0 mm CdCl², and 10 mm HEPES (pH 7.9).26,27All experiments were performed at room temperature (20°–22°C). After obtaining a gigaohm seal, the series resistance was compensated approximately 70–80% using the series resistance compensation of the pulse software, and the final series resistance revel was compensated under 1 MΩ. The leak currents were subtracted by leak compensation within the pulse software for each experiment.

In whole cell voltage clamp recordings, neurons were divided into three groups: control, 0.1, and 1 mm lidocaine perfusion groups. Currents were measured 6 min after each dose of lidocaine. The current–voltage curve, steady state activation curve, and steady state inactivation curve were plotted.

For the current–voltage curve, INa was measured in response to 10-mV, 20-ms steps from a holding potential of −120 mV (1 s) and 10-mV incremental steps to potentials between −80 and +40 mV.26,27Currents were normalized to the maximum in control and plotted as a function of membrane potential.

For the steady state activation curve, peak INa was measured in response to 20-ms, 10-mV incremental depolarizing steps from −120 to +40 mV from a holding potential of −120 mV for 1 s. From the peak INa, the conductance values (G) were calculated according to the equation G = INa/(V − ENa), where INa was the peak Na⁺current measured at each membrane potential (V), and ENa is the reversal potential estimated by the current–voltage curve.28Calculated G values were normalized to the maximum of the control or to the maximum of each group and plotted as a function of membrane potential. Plots were fitted to Boltzmann relations of the form y = G^min+ (G^max− G^min)/{1 + exp[(V− V⁵⁰)/K]}, where y is normalized GNa, G^minis the G of baseline, G^maxis the G of maximum, V is each membrane potential, V⁵⁰is the membrane potential at which conductance is halfway between G^minand G^max, and K is the slope factor.10,28,29 

For the steady state inactivation curve, the membrane potential was prepulsed from −120 to 40 mV at 10-mV increments for 5 s. INa was measured with a 20-ms test pulse to −10 mV. Currents were normalized to the maximum in control and plotted as a function of prepulse potential. Plots were fitted to Boltzmann relations of the form y = I^min+ (I^max− I^min)/{1 + exp[(V − V⁵⁰)/K]}, where y is normalized INa of the maximum in control or in each group, I^minis the INa of baseline, I^maxis the INa of maximum, V is each prepulse potential, V⁵⁰is the membrane potential at which conductance is halfway between I^minand I^max, and K is a slope factor.10,28,29In steady state activation and inactivation curves, analyses were performed using Kaleida graph (version 3.52; Synergy Software, Reading, PA).

Results are expressed as mean ± SEM. The results of repeated measurements in each dose in each group of trials were analyzed by repeated-measurement one-way analysis of variance, followed by the Scheffé test. Among the three doses of lidocaine in the normal saline trials and among the high-Na⁺saline, Na⁺-free saline, hyperpolarized, and tetrodotoxin pretreatment trials, the three groups in the current–voltage curve of whole cell patch clamp recordings were analyzed by analysis of variance, followed by the Scheffé test. Stat view (version 4.5; Abacus, Berkeley, CA) was used for these analyses. P < 0.05 was considered statistically significant.

To test whether lidocaine affects neuronal excitability and intracellular Na concentration, intracellular activity of individually isolated neurons were monitored either in the absence or in the presence of lidocaine.

The mean values of the fluorescence ratio and [Na⁺]ⁱat resting membrane potential were 2.4 ± 0.1 and 9 ± 6 mm, respectively. There was no difference in the initial mean values of [Na⁺]ⁱbefore lidocaine or tetrodotoxin administration in each trial. Half of RPeD1 neurons have spontaneous activity at resting membrane potentials.

Figure 2Ashows experimental tracings of membrane potential and [Na⁺]ⁱbefore and after lidocaine administration. After lidocaine perfusion, [Na⁺]ⁱincreased, and membrane potential depolarized. Soon after a high dose of lidocaine (1 mm), [Na⁺]ⁱincreased, membrane potential continued to be depolarized, and action potential frequency increased (maximum frequency of action potentials: from 5 ± 5 to 37 ± 9 rate/min at 0.01 mm, to 80 ± 12 rate/min at 0.1 mm, and to 114 ± 21 rate/min at 1 mm). However, this initial increase in action potential frequency was transient, lasting only a few seconds, followed by a flat membrane potential. In the normal saline trials, lidocaine significantly depolarized the membrane potential (from −59 ± 3 to −55 ± 3 mV at 0.01 mm, to −50 ± 3 mV at 0.1 mm, and to −21 ± 3 mV at 1 mm; fig. 2B). [Na⁺]ⁱwas significantly increased at each lidocaine dose (from 9 ± 6 to 14 ± 6 mm at 0.01 mm, to 16 ± 6 at 0.1 mm, and to 39 ± 9 at 1 mm; fig. 2B). Both the depolarization of the membrane potential and the increase in [Na⁺]ⁱafter lidocaine perfusion were dose dependent.

The membrane potential and the [Na⁺]ⁱwere stabilized for 2 min before high-Na⁺saline application, and also, these were allowed to stabilize in high-Na⁺saline 2 min before lidocaine application. High-Na⁺saline increased [Na⁺]ⁱand depolarized membrane potential. Subsequent application of lidocaine, in the presence of high-Na⁺saline, further increased [Na⁺]ⁱand also further depolarized the membrane. ([Na⁺]ⁱ: from 10 ± 3 to 59 ± 10 mm; membrane potentials: from −58 ± 6 to −14 ± 4 mV; fig. 3). The membrane potential and the intracellular sodium concentration after lidocaine perfusion were compared not only with the data of control but also with the data after high-Na⁺saline perfusion during the same period.

In the Na⁺-free saline trials, the increase in [Na⁺]ⁱafter lidocaine perfusion was significantly lower than that of normal saline trials (from 7 ± 4 to 5 ± 3 mm; fig. 4). Membrane potential was depolarized after lidocaine perfusion (from −58 ± 4 to −44 ± 14 mV), and this increase was significantly less than that of normal saline trials.

In the membrane hyperpolarization trials, where membrane potential was reduced from −60 mV to −80 mV, spontaneous action potentials ceased to occur. In the membrane hyperpolarization trials, the increase in [Na⁺]ⁱafter lidocaine perfusion was significantly lower than that of normal saline trials (from 8 ± 4 to 17 ± 6 mm; fig. 5).

Tetrodotoxin (30 μm) significantly suppressed the lidocaine-induced increase in [Na⁺]ⁱand membrane potential depolarization when compared with normal saline trials ([Na⁺]ⁱ: from 9 ± 2 to 11 ± 3 mm; membrane potentials: from −59 ± 2 to −56 ± 6 mV; fig. 6).

In the presence of normal saline, the threshold for INa was −40 mV, and the maximum current response occurred at −10 mV, with the mean value at −1.73 ± 0.12 nA. The reversal potential of INa was +22 mV. The INa was blocked by 30 μm tetrodotoxin at all potentials.

Lidocaine decreased maximum current response of INa at −10 mV in a dose-dependent manner as compared with control (to 76 ± 4% at 0.1 mm, to 48 ± 4% at 1 mm; figs. 7A–C). However, the typical amount of membrane depolarization resulting from 1 mm lidocaine application (horizontal gray arrow), from the resting membrane potential in the control (shown as the INa at −60 mV in the control with black arrow) to the decreased membrane potential (usually −20 mV), should result in an inward sodium current (vertical gray arrow) (40-fold increase in INa between −60 and −20 mV).

Lidocaine did not apparently effect the steady state activation curve of INa. There was not a significant shift of the steady state activation curve with up to 1 mm lidocaine (V⁵⁰: −24.2 mV at control to −22.3 mV at 0.1 mm, to −19.6 mV at 1 mm); also, there was no significant difference in the slope values up to 1 mm lidocaine (K value: 6.4 ± 0.3 at control to 6.5 ± 0.2 at 0.1 mm, to 7.3 ± 0.2 at 1 mm; figs. 8A–C).

Lidocaine induced a hyperpolarizing shift in the steady state inactivation curve in a dose-dependent manner (V⁵⁰: −55.3 mV at control to −62.3 mV at 0.1 mm, to −71.9 mV at 1 mm; K value: 8.6 ± 0.3 at control to 9.3 ± 0.3 at 0.1 mm, to 9.5 ± 0.3 at 1 mm; figs. 9A–C). With the perfusion of 1 mm lidocaine, 20% of the inward sodium current remained (approximately 80% inactivation) at the resting membrane potential of −60 mV (fig. 9C). From −60 to −20 mV, there are window currents where the activation and inactivation curves overlap, and thus a persistent inward sodium current can occur (slight conductance and incomplete inactivation) (fig. 10). Note that the range of this window current corresponds with the level of depolarization induced by 1 mm lidocaine (fig. 2B).

Our results show that lidocaine increases [Na⁺]ⁱand depolarizes the membrane potential of the isolated RPeD1 respiratory pacemaker neuron at rest.

In the high-Na⁺saline trials, the increases in [Na⁺]ⁱafter lidocaine treatment were significantly higher than that of normal saline. However, in the Na⁺-free saline trials, lidocaine did not produce an enhancement of [Na⁺]ⁱ. Therefore, the increase in [Na⁺]ⁱproduced by lidocaine is likely from the extracellular fluid. These results indicate that the lidocaine-induced enhancement of [Na⁺]ⁱmay result from the activation of Na channels. Consistent with this conclusion are our results that demonstrated that an increase in [Na⁺]ⁱinduced by lidocaine was suppressed by hyperpolarization or after tetrodotoxin pretreatment. Therefore, one of the mechanisms by which lidocaine causes an increase in [Na⁺]ⁱmay involve tetrodotoxin-sensitive voltage-dependent sodium channels. These results are supported by patch clamp data, which show that lidocaine increases INa approximately 40 times greater than observed at resting membrane potentials under control conditions.

In the current study, lidocaine decreased the maximum current response in a dose-dependent manner, and these data are consistent with those reported previously.30–32However, there were some variations with previous reports, one being that lidocaine increased inward sodium currents when compared with the INa of a resting membrane potential. This can be explained by analyzing previous reports where the effects of lidocaine on INa were compared at the same membrane potential. For example, the INa was mostly compared at the membrane potential with an induced maximum current response of approximately −10 to 0 mV. However, there was in fact a difference in membrane potential before and after lidocaine treatments. Therefore, we reason that the currents including INa must be compared for each membrane potential before and after lidocaine or after the blocker treatment. In addition, our results indicate that lidocaine depolarizes membrane potential, increases INa and induces more excitation to each neuron when the membrane potential is in the window currents. However, in contrast, when the membrane potential is high, outside of the window current, as would be the case during an action potential, lidocaine would decrease INa. In the steady state inactivation curve, lidocaine induced a hyperpolarizing shift, which is similar to that of previous reports.10,32,33In these reports, it is concluded that the hyperpolarizing shift in the steady-state inactivation curve is one of the reasons that lidocaine reduces INa. However, in RPeD1, the availability of voltage-dependent sodium channels at −60 mV remains 20% after lidocaine perfusion (fig. 9C). In the range of −60 to −20 mV, which is the same range depolarized by 1 mm lidocaine (fig. 2B), activation and inactivation curves overlap before and after lidocaine perfusion, which is referred to as window currents . Therefore, in this range, depolarization induced by lidocaine will trigger conductance through sodium channels as activation occurs, and inactivation will be incomplete (fig. 10). Thus, depolarization of RPeD1 into the range of the window current allows for a small but persistent sodium influx with slower inactivation, which corresponds with the sodium imaging data of figure 2. In contrary, at maximum membrane potentials (> −10 mV), the availability of voltage-dependent sodium channels for persistent sodium influx is minimum, because inactivation is almost complete from −10 to +20 mV after a 5-s prepulse. This could be one of the reasons why lidocaine shows both inhibition and excitation in pacemaker neurons.

However, it is still unclear why lidocaine increases membrane potential. In the current study, lidocaine induced the hyperpolarizing shift in the steady state inactivation curve. Therefore, it is difficult to consider that INa through voltage-dependent sodium channels is the main cause of depolarization by lidocaine. Rodeau et al.  9reported that procaine (10 mm), another local anesthetic, increases membrane conductance and depolarizes membrane potential. This study concluded that local anesthetics increase membrane conductance by physically passing through channel pores. Procaine is an organic cation, but it moves through the cell membrane in “base” form, not “cation” form. Therefore, it is hard to consider that procaine will produce a major current itself. On the other hand, Ibuski et al.  34reported that the intracellular concentrations of the ionic form of local anesthetics increased to approximately 70% of the extracellular concentration. Thus, depolarization of membrane potential by local anesthetics could be caused by an increase in the cation form of local anesthetics within the intracellular space or the gap between intracellular and extracellular concentration of lidocaine. Our results indicate that lidocaine also depolarized membrane potential when sodium was excluded from the extracellular fluid. However, this depolarization was significantly less than that in normal saline trials. Also, tetrodotoxin pretreatment suppressed membrane depolarization by lidocaine. These results indicate that the lidocaine-induced depolarization of the membrane potential is not solely generated by lidocaine itself, but that Na⁺influx via  the tetrodotoxin-sensitive, voltage-dependent sodium channels promotes this depolarization. Komai and McDowell35reported that local anesthetics inhibit potassium currents and induce depolarization. This also may contribute to the initial membrane depolarization; however, potassium currents were not included in this study.

We choose concentrations of 0.01–1 mm lidocaine in the study because it is reported that lidocaine at doses of 0.07–0.1 mm of blood concentration induce convulsions in humans.36In this study, we used SBFI-AM to measure [Na⁺]ⁱ. To investigate the interference to SBFI-AM by Lymnaea  saline and lidocaine, we measured the fluorescent ratio using the same concentration of SBFI and lidocaine solution, but no major interference or fluorescent loss was observed. In figure 2, the depolarized membrane potential by lidocaine only lasts 3–4 min and then has a tendency to come back to the base level even during the application of lidocaine. Many factors contribute to the maintaining of membrane potential; thus, the inhibitory effect of lidocaine on voltage-dependent sodium channels may not directly result in alternations in membrane potential. Other factors, such as sodium–calcium exchangers and other ionic currents, may be modified by lidocaine application. To determine how lidocaine changes [Na⁺]ⁱ, more investigations that include these exchangers and sodium channels are necessary. The major receptors and ion channels that have been identified in L. stagnalis  are similar to those found in humans, and therefore, L. stagnalis  represents an excellent model to study the effects of lidocaine.

In conclusion, lidocaine increases intracellular sodium concentration and promotes excitation in respiratory pacemaker neurons through voltage-dependent sodium channels by altering membrane potential. This depolarization is enough to activate a persistent sodium influx through the window current, an idea confirmed with sodium imaging.

The authors thank Tyler Dunn, Ph.D. (Respiratory and Neuroscience Research Groups, Departments of Cell Biology and Anatomy/Physiology and Biophysics, Faculty of Medicine, University of Calgary, Calgary, Alberta, Canada), and Naweed I. Syed, M.D, Ph.D. (Associate Professor, Respiratory and Neuroscience Research Groups, Departments of Cell Biology and Anatomy/Physiology and Biophysics, Faculty of Medicine, University of Calgary), for helping with the manuscript preparation.