This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Email this article to a colleague Similar articles in this journal Similar articles in Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Web of Science (13) Citing Articles via Google Scholar Google Scholar Articles by Oda, A. Articles by Dohi, S. Search for Related Content PubMed PubMed Citation Articles by Oda, A. Articles by Dohi, S.

---

REGIONAL ANESTHESIA AND PAIN MEDICINE

Characteristics of Ropivacaine Block of Na+ Channels in Rat Dorsal Root Ganglion Neurons

Akiyoshi Oda, MD^*, Hidenori Ohashi, DVM, PhD, Seiichi Komori, DVM, PhD, Hiroki Iida, MD, PhD^*, and Shuji Dohi, MD, PhD^*

*Department of Anesthesiology and Critical Care Medicine, and Laboratory of Pharmacology, Veterinary Science, Faculty of Agriculture, Gifu University, Gifu City, Gifu, Japan

Address correspondence and reprint requests to Shuji Dohi, MD, PhD, Department of Anesthesiology and Critical Care Medicine, Gifu University School of Medicine, 40 Tsukasa-machi, Gifu City, Gifu 500-8705, Japan. Address e-mail to shu-dohi{at}cc.gifu-u.ac.jp

	Abstract

Top Abstract Introduction Methods Results Discussion References

 
When used for epidural anesthesia, ropivacaine can produce a satisfactory sensory block with a minor motor block. We investigated its effect on tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) Na⁺ currents in rat dorsal root ganglion (DRG) neurons to elucidate the mechanisms underlying the above effects. Whole-cell patch-clamp recordings were made from enzymatically dissociated neurons from rat DRG. A TTX-S Na⁺ current was recorded preferentially from large DRG neurons and a TTX-R Na⁺ current preferentially from small ones. Ropivacaine shifted the activation curve for the TTX-R Na⁺ channel in the depolarizing direction and the inactivation curve for both types of Na⁺ channel in the hyperpolarizing direction. Ropivacaine blocked TTX-S and TTX-R Na⁺ currents, but its half-maximum inhibitory concentration (IC₅₀) was significantly lower for the latter current (116 ± 35 vs 54 ± 14 µM; P < 0.01); similar IC₅₀ values were obtained with the (R)-isomer of ropivacaine. Ropivacaine produced a use-dependent block of both types of Na⁺ channels. Ropivacaine preferentially blocks TTX-R Na⁺ channels over TTX-S Na⁺ channels. We conclude that because TTX-R Na⁺ channels exist mainly in small DRG neurons (which are responsible for nociceptive sensation), such selective action of ropivacaine could underlie the differential block observed during epidural anesthesia with this drug.

Implications: Whole-cell patch-clamp recordings of tetrodotoxin-sensitive and tetrodotoxin-resistant Na⁺ currents in rat dorsal root ganglion neurons showed ropivacaine preferentially blocked tetrodotoxin-resistant Na⁺ channels over tetrodotoxin-sensitive Na⁺ channels. This could provide a desirable differential sensory blockade during epidural anesthesia using ropivacaine.

	Introduction

Top Abstract Introduction Methods Results Discussion References

 
Ropivacaine (1-propyl-2',6'-pipecoloxylidide) is a new amino-amide local anesthetic similar in structure to bupivacaine; it is prepared as the (S)-isomer. Several clinical studies of epidural anesthesia have shown that although ropivacaine exerts a blocking effect on sensory nerves equivalent to that of bupivacaine, its motor block is slower in onset, less intense, and of shorter duration than that of bupivacaine (1–3). However, neither the mechanisms underlying these effects, nor the pharmacokinetics of ropivacaine’s effects on Na⁺ channels in sensory neurons, have been studied.

The major sites at which local anesthetics act to produce epidural or spinal anesthesia are considered to be: 1) on the mixed nerves in the paravertebral spaces after their passage through the intervertebral foramina, 2) on the dorsal root ganglia (DRG), 3) on the intradural spinal roots, and 4) within the spinal cord. Neurons in the DRG express an unusually slow, tetrodotoxin-resistant (TTX-R) Na⁺ channels as well as a tetrodotoxin-sensitive (TTX-S) Na⁺ channels (4,5). Lidocaine and bupivacaine have both been reported to block TTX-S Na⁺ channels more potently than TTX-R Na⁺ channels in rat DRG neurons (6). Peripheral sensory neurons have their cell bodies in the DRG and connect synaptically to the neurons of the sensory pathways that run in the central nervous system (CNS) (7). Thus, DRG neurons function as key elements in the transmission of sensory stimuli to the CNS. Small myelinated A fibers and unmyelinated C fibers, which transmit nociceptive sensation, are believed to arise from small DRG neurons, whereas large myelinated A and Aß fibers arise from large DRG neurons (8). Previous studies have revealed that small DRG neurons tend to express a Na⁺ current that has slower kinetics of activation and inactivation and is less sensitive (resistant) to TTX, whereas large DRG neurons tend to express a Na⁺ current that has faster kinetics of activation and inactivation and is sensitive to tetrodotoxin (TTX) (4,6).

The present study was designed to examine the effects of ropivacaine on TTX-S and TTX-R Na⁺ channels in rat DRG neurons using whole-cell patch-clamp techniques. In so doing, we hoped to provide a mechanistic explanation for the observation that ropivacaine can bring about an adequate pain block with only a minor motor block when it is used for epidural anesthesia.

	Methods

Top Abstract Introduction Methods Results Discussion References

 
DRG neurons were isolated as described previously (9). Briefly, young Sprague-Dawley rats (7–17 days postnatal, either sex) were killed by decapitation, and DRGs were rapidly removed along the cervical, thoracic, and lumbar sections of the spinal cord. The DRGs were incubated at 37°C for 23–30 min in Tyrode solution (for composition, see below) containing 2 mg/mL collagenase (Type 1; Sigma, St Louis, MO) and 5 mg/mL dispase II (Boehringer Mannheim, Indianapolis, IN). After washing three times with fresh, enzyme-free Tyrode solution, single neuronal cells were obtained by gentle agitation in Tyrode solution through a small-bore Pasteur pipette. After filtration of the cell suspension, the collected cells were resuspended in Tyrode solution, placed on glass coverslips, and incubated in a humidified atmosphere of 5% CO₂ at 37°C for 2–8 h before being used for the patch-clamp experiments.

A coverslip carrying cells was placed in a small organ bath on the stage of an inverted microscope (TMD; Nikon, Tokyo, Japan). Recordings of whole-cell membrane currents were made at the averaged experimental temperature of 23° ± 2°C (n = 45) by using patch-clamp techniques. Patch pipettes were made from glass capillaries using a two-step vertical puller (PP-830; Narishige, Tokyo, Japan), their tips being fire-polished by using a microforge (MF-830; Narishige) to give a final resistance of 1.0–2.0 M. Membrane currents were amplified by using a current amplifier (EPC-8; HEKA electronik, Lambrecht, Germany) and stored on a pulse code modulation data recorder (RD-130T; TEAC, Tokyo, Japan). Data analysis was performed on a personal computer (PowerBook 1400cs; Apple Computer, Cupertino, CA) by using a data acquisition and analysis instrument (MacLab8; AD Instruments, Castle Hill, NSW, Australia). The current signals were filtered at a cutoff frequency of 3 kHz and digitized with a sampling rate of 40 kHz. Series resistance was compensated by 50%–80%. Experimental data were discarded when the voltage error exceeded 5 mV after compensation of the series resistance.

The Tyrode solution was of the following composition (mM): NaCl 140.0, KCl 4.0, MgCl₂ 2.0, glucose 10.0, HEPES 10.0, and it was adjusted to pH 7.4 with NaOH. As reported by Song et al. (5), the pipette solution was of the following composition (mM): CsF 135.0, NaCl 10.0, HEPES 5.0 (adjusted to pH 7.0 with CsOH). The external solution was as follows: NaCl 25.0, tetramethylammonium chloride 75.0, tetraethylammonium chloride 20.0, CsCl 5.0, CaCl₂ 1.8, MgCl₂ 1.0, glucose 25.0, HEPES 5.0 (adjusted to pH 7.4 with tetraethylammonium-OH). Lanthanum solution (LaCl₃, 3 mM) was added to the external solution to give a final concentration of 3 µM, sufficient to block calcium channel currents. A sodium-free external solution was prepared by replacing the NaCl of the external solution with equimolar tetramethylammonium chloride.

The drugs used were the (S)-isomer and the (R)-isomer of ropivacaine (Astra, Södertälje, Sweden) each in the form of the hydrochloride monohydrate, bupivacaine (Sigma, St Louis, MO) in the form of the hydrochloride and tetrodotoxin (Sigma). Extracellular application of drugs was achieved by replacing the bath solution in the recording chamber (0.8 mL) with drug-containing solution 5–7 times within 20 s.

Analyses were performed as described previously (5). The Na⁺ conductance (g_na) of the membrane was calculated by using the equation:

where I_na is the peak amplitude of the Na⁺ current, Vg is the membrane potential achieved using a step pulse, and V_r is the reversal potential for Na⁺ (23.1 mV).

Activation curves were fitted with the Boltzmann equation:

where _maxg_na is the maximal value for g_na, V_g0.5 is the potential at which g_na is half of _maxg_na,, and k_g is the slope factor.

Inactivation curves were also drawn according to the Boltzmann equation:

where _maxI_na is the maximal value for I_na, Vh is the membrane potential achieved using a prepulse (conditioning) potential, Vh_0.5 is the potential at which I_na is half of _maxI_na, and k_h is the slope factor.

The dose-response curves for the blocking action of ropivacaine on TTX-S and TTX-R currents were fitted to the Hill equation:

where IC₅₀ is half-maximum concentration for the inhibitory action of ropivacaine, [Ropivacaine] is the concentration of ropivacaine, and h is the Hill coefficient.

All values in the text are expressed as mean ± SD. Statistical significance was assessed by using a Student’s paired or unpaired t-test; differences were considered significant when P < 0.05.

	Results

Top Abstract Introduction Methods Results Discussion References

 
DRG neurons were held under voltage clamp at -100 mV, and the whole cell membrane Na⁺ currents of DRG neurons were evoked by stepping to 0 mV for 50 ms. Na⁺ current appeared as a brief inward current that was activated and then inactivated within 5 ms or as a longer inward current that persisted for 20 ms or more (Fig. 1, A and B). The short-lasting Na⁺ current was almost completely blocked by 0.2 µM TTX (Fig. 1A), indicating that it is a result of activation of TTX-S Na⁺ channels. The long-lasting Na⁺ current was resistant to or only partly blocked by 0.2 µM TTX (Fig. 1B). The remaining Na⁺ current disappeared when tetramethylammonium chloride was substituted for Na⁺ ions in the bath solution. Thus, the long-lasting Na⁺ current was considered to be a result of activation of TTX-R Na⁺ channels alone or a mixture of TTX-S and TTX-R Na⁺ channels.

View larger version (21K): [in this window] [in a new window]   Figure 1. Characterization of Na+ current in dorsal root ganglion (DRG) neurons. A 50-ms step pulse to 0 mV from the holding potential (-100 mV) was applied to activate Na+ current in the absence (control) and presence of 0.2 µM tetrodotoxin (TTX). A and B, Recording traces of a short-lasting (A) and a long-lasting (B) Na+ current from different cells; the former current was completely blocked by 0.2 µM TTX, which indicates that it is a result of activation of TTX-sensitive (TTX-S) Na+ channels, and the latter current was only partly blocked by 0.2 µM TTX, which indicates that it is a result of activation of a mixture of TTX-S and TTX-resistant (TTX-R) Na+ channels. C, Histogram of distribution of the diameter of DRG neurons that exhibited the short-lasting (hatched columns) and the long-lasting (open columns) Na+ currents. See text for details.

In the following experiments, the short-lasting Na⁺ currents were defined as TTX-S Na⁺ current and long-lasting Na⁺ currents detected in the presence of 0.2 µM TTX as TTX-R Na⁺ current.

To examine the effect of ropivacaine on TTX-S and TTX-R Na⁺ currents, we used the holding potentials -120 mV and -100 mV respectively, according to Song et al. (5). At these holding potentials, both types of Na⁺ channels were completely released from the inactivation state (see Fig. 2). Ropivacaine was used at 300 µM for TTX-S Na⁺ channels and 100 µM for TTX-R Na⁺ channels as the concentration needed to produce approximately half-maximum block (see Fig. 3).

View larger version (25K): [in this window] [in a new window]   Figure 2. A, Effect of ropivacaine on the voltage-dependent activations of tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) Na+ channels. The ratio of Na+ conductance to maximal Na+ conductance (gna/maxgna) was plotted against the membrane potential (Test Potential) attained by stepping from each holding potential. Points were fitted by the Boltzmann equation. Each point represents the mean ± SD of seven measurements. Note that the conductance-voltage relationship for TTX-R Na+ currents was shifted by the drug by 9.1 mV in the positive membrane potential direction. B, Effect of ropivacaine on the voltage-dependent inactivation of TTX-S and TTX-R Na+ channels. Conditioning pulses stepped (for 150 ms) from -120 mV to -20 mV in 10-mV increments were followed by a 5-ms test pulse stepped to -10 mV from the various prepulse potentials for TTX-S Na+ channels. And for TTX-R Na+ channels, the conditioning pulses stepped (for 150 ms) from -100 mV to 0 mV in 10-mV increments were followed by a 5-ms test pulse stepped to 0 mV from the various prepulse potentials in the presence of 0.2 µM tetrodotoxin (TTX). The peak amplitudes of the Na+ currents evoked by test pulses without any preceding conditioning pulse were normalized as 1.0 both in the absence and presence of the drug and the relative peak amplitude of the Na+ current was plotted against the membrane potential. Same symbols as above were used. Points were fitted by the Boltzmann equation. Each point represents the mean ± SD of six measurements. The voltage-dependence of the current inactivation was shifted by the drug by a similar amount in the negative membrane potential direction, irrespective of the type of Na+ current.

View larger version (17K): [in this window] [in a new window]   Figure 3. A, Concentration-response curves for the blocking action of ropivacaine on the tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) Na+ current. Currents were evoked by stepping (for 50 ms) from -120 mV to -10 mV for TTX-S Na+ channels, or from -100 mV to 0 mV for TTX-R Na+ channels in the presence of 0.2 µM tetrodotoxin (TTX). B, Concentration-response curves for the inhibitory effects of ropivacaine, the (R)-isomer of ropivacaine and bupivacaine on TTX-S and TTX-R Na+ current. Currents were evoked by stepping (for 50 ms) from a holding potential of -80 mV to -10 mV for TTX-S Na+ channels, or to 0 mV for TTX-R Na+ channels in the presence of 0.2 µM TTX. Abscissa: log concentration of drugs. Ordinate: % inhibition of the peak amplitude of the current (the peak amplitude of the currents elicited in the absence of drugs were given the value of 0%). Each point represents the mean ± SD of six measurements.

View larger version (22K): [in this window] [in a new window]   Figure 4. Current-voltage relationships for tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) Na+ currents. A and B, TTX-S Na+ currents evoked by stepping (50 ms duration; one step every 7 s) to potentials of -60 to + 30 mV in 10-mV increments from a holding potential of -120 mV in the absence (A) or presence of 300 µM ropivacaine (B). D and E, TTX-R Na+ currents evoked by stepping (50-ms duration; one step every 7 s) to potentials of -50 to + 30 mV in 10 mV increments from a holding potential of -100 mV in the absence (D) or presence of 100 µM ropivacaine (E). C and F, Current-voltage relationships for TTX-S and TTX-R Na+ currents (C and F, respectively) in the absence () or presence of ropivacaine (•), and after removal of ropivacaine (). The amplitude of the Na+ currents was taken as the difference between the peak current and the final level of the current achieved by using the same pulse protocol after replacement of the bath solution by Na+ -free solution. Current traces were obtained by smoothing of original data by analysis software.

Either type of Na⁺ currents were repetitively elicited every 7 s by using a 50-ms step pulse, during which ropivacaine was applied at stepwise increased concentrations every 2 min. The cumulative application of ropivacaine decreased the peak amplitude of the Na⁺ currents in a concentration-dependent manner. The decrease of current amplitude was reversed by removing the drug from the bath solution. Data from such cells that did not exhibit peak current recovery exceeding 50% of the initial value 5 min after the removal of ropivacaine were discarded. The IC₅₀ value for the TTX-S Na⁺ channel block was significantly higher than the value for the TTX-R Na⁺ channel block (Fig. 3A and Table 1).

Na⁺ currents were elicited repeatedly by applying successive depolarizing pulses at 0.2, 5, or 20 Hz in the absence and presence of 30 µM ropivacaine for TTX-S and 10 µM for TTX-R Na⁺ channels; 30 µM and 10 µM ropivacaine produced approximately 15% inhibition of peak currents of TTX-S and TTX-R Na⁺ channels, respectively (Fig. 3B). Currents were evoked repeatedly by stepping (for 10 ms) to 0 mV from a holding potential of -80 mV at one of three different frequencies. TTX-R Na⁺ currents were evoked in the presence of 0.2 µM TTX. The peak amplitude of the current evoked by the first step pulse was normalized as 100% in the absence and presence of ropivacaine. When the current amplitude evoked by the 20th stimulus at a given frequency in the presence of ropivacaine was compared with the corresponding value obtained in its absence, an appreciable drug effect was detected. One min or so was allowed to elapse between one pulse train and the next at each frequency. The current amplitude was invariably smaller in the presence of ropivacaine, and the differences were statistically significant at 5 and 20 Hz in both types of Na⁺ channel (Table 2).

	Discussion

Top Abstract Introduction Methods Results Discussion References

Small DRG neurons play an important role specific to nociceptive transmission in physiologic and pathophysiologic sensory processing (13,14). Action potentials and excitatory postsynaptic potentials recorded from cells in the isolated DRG and dorsal horn that were derived from excitation of C fibers were resistant to TTX (15). The conduction of bradykinin-induced nociceptive sensation in rat primary afferent fibers was resistant to TTX (16). In compound action potentials recorded from isolated nerve fascicles of the human sural nerve, the C-fiber components were TTX-resistant (17). In addition, a recent study using gene-targeting technology to delete TTX-R Na⁺ channels in sensory neurons demonstrated that this type of Na⁺ channel has a specialized function in pain pathways (14). Considering our results with these previous observations, we conclude that small DRG neurons from which A and C fibers arise as afferent sensory fibers (18) may preferentially express TTX-R Na⁺ channels, whereas large DRG neurons giving rise to A and Aß fibers may preferentially express TTX-S Na⁺ channels. If this is so, the present findings can be taken as being consistent with the selective inhibition by ropivacaine of pain sensation that is observed when it is used for epidural anesthesia.

The local anesthetics, lidocaine, etidocaine, and bupivacaine have all been reported to be more potent at blocking TTX-S Na⁺ currents than TTX-R Na⁺ currents in rat DRG neurons (Table 1). Studies of compound action potentials evoked in rat vagal nerve (19) and in rabbit vagal and sciatic nerves (20) have revealed that with respect to the ratio between the smallest concentrations required for reliable block of A fibers and C fibers (C fibers : A fibers), the rank order was lidocaine > etidocaine > bupivacaine > ropivacaine. This order shows a positive correlation with the rank order reported so far for DRG neurons in respect of the ratio between the IC₅₀ values for TTX-S Na⁺ currents and TTX-R Na⁺ current: lidocaine etidocaine > bupivacaine > ropivacaine (Table 1). Two studies indicate that although not differentiated in respect of their TTX sensitivity, the Na⁺ channels in toad sciatic fibers (21) and rat dorsal horn neurons (22) have different sensitivities to a given local anesthetic. In experiments on the Na⁺ channels in membrane patches of toad sciatic neurons, the following IC₅₀ values were determined: tetracaine 0.7 µM, etidocaine 18 µM, bupivacaine 27 µM, lidocaine 204 µM (21). Although there was species difference, some of these show considerable differences from those determined for DRG neurons, as shown in Table 1. This suggests that there may be considerable differences in Na⁺ channel characteristics among primary afferent fibers, DRG neurons, and spinal cord neurons, all of which are in a compartment that can be affected by epidural anesthesia.

It is unclear what factors are involved in the differential blockade of Na⁺ channels produced by individual drugs. The blocking potency of local anesthetics on TTX-R Na⁺ channels in DRG neurons is independent of their lipophilicity (23). Local anesthetics could reach the binding sites not only along the hydrophobic pathway, by which lipid-soluble local anesthetics are thought to come and go from their binding sites via the hydrophobic regions of the membrane, but also along the hydrophilic pathway, by which charged and less lipid-soluble substances pass via a hydrophilic region (the inner channel mouth) (24). When ropivacaine [the (S)-isomer] was compared with the (R)-isomer in terms of the blocking action on evoked action potentials in isolated frog sciatic nerve and the ability to produce infiltration anesthesia after injection into the space surrounding the sciatic nerve in the guinea-pig, the (S)-isomer produced a longer duration of action, although the smallest concentration needed to reliably produce an effect was not different between the two isomers (25). Because the present results showed no appreciable difference between the (S)- and (R)-isomers of ropivacaine for potency at blocking the two types of Na⁺ current, it is suggested that stereospecificity is unlikely to be a factor in the differential block of TTX-S and TTX-R Na⁺ channels in sensory neurons.

An interesting feature of the effect of ropivacaine was the apparent change in the ratio between the IC₅₀ values for TTX-S and TTX-R Na⁺ current when a different holding potential was used. At a holding potential of -80 mV, some 20% of TTX-S Na⁺ channels were inactivated. This situation is consistent with the hypothesis that the binding affinity of a given local anesthetic for a Na⁺ channel may vary depending on the channel state (26). Ropivacaine, like other local anesthetics (24), seems likely to increase the probability that the Na⁺ channel in question is in the inactive state, irrespective of its sensitivity to TTX, as judged by the shift in Vh_0.5 in the hyperpolarizing direction along the voltage axis (Fig. 2B). Ropivacaine was also effective in producing a significant shift of the activation curve for TTX-R Na⁺ channels in the depolarizing direction, but this was not seen for TTX-S Na⁺ channels (Fig. 2A). This suggests that the drug reduces the functional fraction of TTX-R Na⁺ channels present at a given level of membrane depolarization (i.e., decreases the number of channels that can open in response to each depolarizing pulse), which in turn leads to a decreased excitability of the membrane. This effect of ropivacaine may also account for production by the drug of a differential Na⁺ channel block.

A use-dependent block of TTX-R Na⁺ channels has been demonstrated for several local anesthetics in various preparations (4,6,23). The most plausible explanation for this effect is that the local anesthetics reach their site of action more easily when Na⁺ channels are opened (27). The use-dependent block of ropivacaine may also contribute to the ropivacaine-inhibition of Na⁺ currents in small DRG neurons associated with nociceptive C fibers, many of which are activated by tissue damage and involved in the development of some chronic pain syndromes.

In conclusion, ropivacaine preferentially blocks TTX-R Na⁺ channels, expressed primarily in rat DRG neurons from which nociceptive C fibers arise. The differential block of TTX-S and TTX-R Na⁺ channels could provide a desirable differential sensory blockade during epidural anesthesia using ropivacaine. Ropivacaine may also reduce the functional availability of TTX-R Na⁺ channels in nociceptive C fibers through the generation of a use-dependent block and a reduction in voltage sensitivity.

	Acknowledgments

 
Supported, in part, by Grant-in-Aid for Scientific Research No.0847405.

	References

Top Abstract Introduction Methods Results Discussion References

 

1. Datta S, Camann W, Bader A, VanderBurgh L. Clinical effects and maternal and fetal plasma concentrations of epidural ropivacaine versus bupivacaine for cesarean section. Anesthesiology 1995; 82: 1346–52.[Web of Science][Medline]
2. Griffin RP, Reynolds F. Extradural anaesthesia for caesarean section: a double-blind comparison of 0.5% ropivacaine with 0.5% bupivacaine. Br J Anaesth 1995; 74: 512–6.[Abstract/Free Full Text]
3. Brockway MS, Bannister J, McClure JH, et al. Comparison of extradural ropivacaine and bupivacaine. Br J Anaesth 1991; 66: 31–7.[Abstract/Free Full Text]
4. Roy ML, Narahashi T. Differential properties of tetrodotoxin-sensitive and tetrodotoxin-resistant sodium channels in rat dorsal root ganglion neurons. J Neurosci 1992; 12: 2104–11.[Abstract]
5. Song JH, Huang CS, Nagata K, et al. Differential action of riluzole on tetrodotoxin-sensitive and tetrodotoxin-resistant sodium channels. J Pharmacol Exp Ther 1997; 282: 707–14.[Abstract/Free Full Text]
6. Scholz A, Kuboyama N, Hempelmann G, Vogel W. Complex blockade of TTX-resistant Na⁺ currents by lidocaine and bupivacaine reduce firing frequency in DRG neurons. J Neurophysiol 1998; 79: 1746–54.[Abstract/Free Full Text]
7. Levine JD, Fields HL, Basbaum AI. Peptides and the primary afferent nociceptor. J Neurosci 1993; 13: 2273–86.[Abstract]
8. Harper AA, Lawson SN. Conduction velocity is related to morphological cell type in rat dorsal root ganglion neurones. J Physiol (Lond) 1985; 359: 31–46.[Abstract/Free Full Text]
9. Cardenas CG, Del Mar LP, Cooper BY, Scroggs RS. 5HT4 receptors couple positively to tetrodotoxin-insensitive sodium channels in a subpopulation of capsaicin-sensitive rat sensory neurons. J Neurosci 1997; 17: 7181–9.[Abstract/Free Full Text]
10. Amir R, Michaelis M, Devor M. Membrane potential oscillations in dorsal root ganglion neurons: role in normal electrogenesis and neuropathic pain. J Neurosci 1999; 19: 8589–96.[Abstract/Free Full Text]
11. Safronov BV, Bischoff U, Vogel W. Single voltage-gated K ⁺ channels and their functions in small dorsal root ganglion neurones of rat. J Physiol (Lond) 1996; 493: 393–408.[Abstract/Free Full Text]
12. MeLean MJ, Bennett PB, Thomas RM. Subtypes of dorsal root ganglion neurons based on different inward currents as measured by whole-cell voltage clamp. Mol Cell Biochem 1988; 80: 95–107.[Web of Science][Medline]
13. Sangameswaran L, Delgado SG, Fish LM, et al. Structure and function of a novel voltage-gated, tetrodotoxin-resistant sodium channel specific to sensory neurons. J Biol Chem 1996; 271: 5953–6.[Abstract/Free Full Text]
14. Akopian AN, Souslova V, Engl S, et al. The tetrodotoxin-resistant sodium channel SNS has a specialized function in pain pathways. Nature Neuroscience 1999; 2: 541–8.[Web of Science][Medline]
15. Jeftinija S. The role of tetrodotoxin-resistant sodium channels of small primary afferent fibers. Brain Res 1994; 639: 125–34.[Web of Science][Medline]
16. Jeftinija S. Bradykinin excites tetrodotoxin-resistant primary afferent fibers. Brain Res 1994; 665: 69–76.[Web of Science][Medline]
17. Quasthoff S, Grosskreutz J, Schroder JM, et al. Calcium potentials and tetrodotoxin-resistant sodium potentials in unmyelinated C fibres of biopsied human sural nerve. Neuroscience 1995; 69: 955–65.[Web of Science][Medline]
18. Strichartz G, Berde C. Local anesthetics. In: Miller R, Cucchiara R, Miller ED Jr, eds. Anesthesia. New York: Churchill Livingstone, 1994: 489–521.
19. Gissen AJ, Covino BG, Gregus J. Differential sensitivities of mammalian nerve fibers to local anesthetic agents. Anesthesiology 1980; 53: 467–74.[Web of Science][Medline]
20. Rosenberg PH, Heinonen E. Differential sensitivity of A and C nerve fibres to long-acting amide local anaesthetics. Br J Anaesth 1983; 55: 163–7.[Abstract/Free Full Text]
21. Brau ME, Vogel W, Hempelmann G. Fundamental properties of local anesthetics: half-maximal blocking concentrations for tonic block of Na⁺ and K ⁺ channels in peripheral nerve. Anesth Analg 1998; 87: 885–9.[Abstract/Free Full Text]
22. Olschewski A, Hempelmann G, Vogel W, Safronov BV. Blockade of Na⁺ and K ⁺ currents by local anesthetics in the dorsal horn neurons of the spinal cord. Anesthesiology 1998; 88: 172–9.[Web of Science][Medline]
23. Brau ME, Elliott JR. Local anaesthetic effects on tetrodotoxin-resistant Na⁺ currents in rat dorsal root ganglion neurones. Eur J Anaesthesiol 1998; 15: 80–8.[Web of Science][Medline]
24. Hille B. Local anesthetics: hydrophilic and hydrophobic pathways for the drug-receptor reaction. J Gen Physiol 1977; 69: 497–515.[Abstract/Free Full Text]
25. Akerman B, Hellberg IB, Trossvik C. Primary evaluation of the local anaesthetic properties of the amino amide agent ropivacaine (LEA 103). Acta Anaesthesiol Scand 1988; 32: 571–8.[Web of Science][Medline]
26. Butterworth JF 4th, Strichartz GR. Molecular mechanisms of local anesthesia: a review. Anesthesiology 1990;72:711–34.
27. Hille B. Ionic channels of excitable membranes. Sunderland: Sinauer, 1984.

This article has been cited by other articles:

				P. Lirk, I. Haller, H. P. Colvin, L. Lang, B. Tomaselli, L. Klimaschewski, and P. Gerner In Vitro, Inhibition of Mitogen-Activated Protein Kinase Pathways Protects Against Bupivacaine- and Ropivacaine-Induced Neurotoxicity Anesth. Analg., May 1, 2008; 106(5): 1456 - 1464. [Abstract] [Full Text] [PDF]

				C. Sato, A. Sakai, Y. Ikeda, H. Suzuki, and A. Sakamoto The Prolonged Analgesic Effect of Epidural Ropivacaine in a Rat Model of Neuropathic Pain Anesth. Analg., January 1, 2008; 106(1): 313 - 320. [Abstract] [Full Text] [PDF]

				Y. Osawa, A. Oda, H. Iida, S. Tanahashi, and S. Dohi The Effects of Class Ic Antiarrhythmics on Tetrodotoxin-Resistant Na+ Currents in Rat Sensory Neurons Anesth. Analg., August 1, 2004; 99(2): 464 - 471. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Email this article to a colleague Similar articles in this journal Similar articles in Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Web of Science (13) Citing Articles via Google Scholar Google Scholar Articles by Oda, A. Articles by Dohi, S. Search for Related Content PubMed PubMed Citation Articles by Oda, A. Articles by Dohi, S.