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- and C-Fiber Conduction and Synaptic Transmission1Instituto de Biologia Molecular e Celular, Universidade do Porto, Porto, Portugal; 2Laboratório de Biologia Celular e Molecular, Faculdade de Medicina, Universidade do Porto, Porto, Portugal; and 3Vollum Institute, Oregon Health and Science University, Portland, Oregon
Submitted 21 August 2007; accepted in final form 2 December 2007
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ABSTRACT |
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- and C-fiber axons as well as on the A- and C-fiber-mediated excitatory postsynaptic currents (EPSCs) in spinal dorsal horn neurons of rat. At room temperature, TTX completely blocked A-fibers (IC50, 5–7 nM) in dorsal roots (central branch) and spinal, sciatic, and sural nerves (peripheral branch). The C-fiber responses were blocked by 85–89% in the peripheral branch and by 65–66% in dorsal roots (IC50, 14–33 nM) with simultaneous threefold reduction in their conduction velocity. At physiological temperature, the degree of TTX block in dorsal roots increased to 93%. The A- and C-fiber-mediated EPSCs in dorsal horn neurons were also sensitive to TTX. At room temperature, 30 nM blocked completely A-input and 84% of the C-fiber input, which was completely suppressed at 300 nM TTX. We conclude that in mammals, the TTX-S Na+ channels dominate conduction in all thin primary afferents. It is the only type of functional Na+ channel in A-fibers. In C-fibers, the TTX-S Na+ channels determine the physiological conduction velocity and control synaptic transmission. TTX-R Na+ channels could not provide propagation of full-amplitude spikes able to trigger synaptic release in the spinal cord. |
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INTRODUCTION |
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- and unmyelinated C-fibers terminate in the superficial dorsal horn of the spinal cord (Brown 1981
; Light and Perl 1977). Many of them convey nociceptive inputs and are involved in the establishment and maintenance of pathophysiological states, including neuropathic pain, caused by abnormal change in the expression of ion channels. A number of studies have shown a pivotal role of voltage-gated Na+ channels in both normal nerve conduction (Hille 2001) and development of neuropathic states (Black et al. 1999; Devor and Seltzer 1999; Gold et al. 2003; Lai et al. 2002; Novakovic et al. 1998; Waxman et al. 1999; Zhang et al. 1997).
Voltage-gated Na+ channels were intensively studied in the soma of peripheral sensory neurons and separated in terms of their sensitivity to tetrodotoxin (TTX) in two basic classes, TTX-sensitive (TTX-S) and TTX-resistant (TTX-R) (Bossu and Feltz 1984; Elliott and Elliott 1993; Kostyuk et al. 1981; Ogata and Tatebayashi 1993). TTX-S Na+ channels are completely blocked by 100 nM TTX, whereas TTX-R current remains insensitive to 75–100 µM TTX (Elliott and Elliott 1993; Ogata and Tatebayashi 1993). TTX-R Na+ channels were found in small-diameter dorsal root ganglion (DRG) neurons giving origin to unmyelinated C- and weakly myelinated A-fiber axons. More recent studies showed that both TTX-S and TTX-R currents are formed by multiple types of Na+ channels with distinct electrophysiological properties (Rush et al. 1998; Scholz et al. 1998) and gene origin (Akopian et al. 1996; Black et al. 1996; Dib-Hajj et al. 1998; Sangameswaran et al. 1996; Tate et al. 1998; Toledo-Aral et al. 1997).
Although TTX-R Na+ channels are expressed at density sufficient to generate an action potential in the soma of small DRG neurons of both A- and C-types (Ritter and Mendell 1992; Villiere and McLachlan 1996; Yoshida and Matsuda 1979), it is still unclear whether they are also capable of generating propagating action potentials in the central and peripheral branches of afferent fibers. Little is also known about the TTX-S and TTX-R Na+ channel distribution along the afferent axon and their involvement in conduction and synaptic transmission. A number of studies used TTX as a tool to address the question whether TTX-R Na+ channels have sufficient density to provide spike propagation in unmyelinated axons. Differences in experimental designs, peripheral nerve regions, sites of TTX application, and the blocker concentrations gave quite contradictory results. On one hand, conduction in C-fibers was found to persist in TTX for frog sciatic nerve (100 µM, Kobayashi et al. 1993; 1 µM, Buchanan et al. 1996), rat dorsal root (0.5 µM, Jeftinija 1994; Jeftinija and Urban 1994; 1 µM, Steffens et al. 2001) and biopsied human sural nerve (1–100 µM) (Grosskreutz et al. 1996; Quasthoff et al. 1995). On the other hand, C-fiber conduction was reported to be blocked by TTX in mouse peripheral nerve (
3 µM) (Yoshida and Matsuda 1979), rat dorsal root, and sciatic nerve (0.3–1 µM) (Villiere and McLachlan 1996). In a similar manner, synaptic transmission from C-fibers to the spinal dorsal horn neurons was found to be blocked by 0.5 µM TTX applied to the spinal cord slice with attached dorsal root (Yoshimura and Jessell 1990) but to persist in 0.5 µM TTX applied to DRG neurons (Jeftinija and Urban 1994).
Therefore we studied the contribution of TTX-S and TTX-R Na+ channels to spike propagation along the fine-caliber A- and C-fiber axons. For this purpose, we measured effects of nanomolar concentrations of TTX on A- and C-fiber responses along the peripheral and central branches of primary afferents as well as on their synaptic transmission in the spinal cord. We found that, regardless of the location along the peripheral and central branches of sensory axons, the TTX-S Na+ channels strongly dominate conduction in both A- and C-fibers and are necessary for synaptic release.
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METHODS |
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Laboratory Wistar rats were killed in accordance with the national guidelines (Direcção Geral de Veterinária, Ministério da Agricultura) after the anesthesia by intraperitoneal injection of Na+-pentobarbital (30 mg/kg). Compound action potential currents were recorded from different regions of the central and peripheral branches of afferent axons (see Fig. 3A) using L5 dorsal root, L5 mixed spinal nerve with a proximal sciatic nerve and sural nerve from 4- to 18-wk-old rats. The nerves and roots were dissected and cleaned from the connective tissue sheath in ice-cold oxygenated artificial cerebrospinal fluid (ACSF). After isolation, the roots and nerves were kept at room temperature of 22–24°C until use. Entire L5 dorsal roots were 13–27 mm long. In the experiments studying the TTX sensitivity in the central branch of afferents, each L5 root was divided in two equal parts, rostral dorsal root (adjacent to the spinal cord) and caudal dorsal root (near the dorsal root ganglion). To record from the proximal part of the peripheral branch of sensory axons, we used the spinal-sciatic nerve preparation that included the L5 mixed spinal nerve and the proximal sciatic nerve. This spinal-sciatic nerve preparation allowed recordings from those sciatic nerve fibers that originated from the L5 segment. In the experiments with TTX, the nerve was shorter (10–13 mm), to reduce the temporal dispersion of the C-fiber response and therefore to increase its amplitude. When the conduction velocity (CV) was measured, longer spinal-sciatic nerve preparation (28–48 mm) was used to increase the accuracy of the measurements. To record from the distal part of the peripheral branch of the axon, we isolated the sural nerves of 8- to 12-mm length.
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) from 3- to 6-wk-old rats. The vertebral column was quickly cut out and immersed in ice-cold oxygenated ACSF. The whole lumbar enlargement with attached 10- to 15-mm-long L5 root was dissected and laterally glued by cyanoacrylate adhesive to a 1-mm-thick metal plate (Fig. 7). One sagittal cut was done using a tissue slicer (Leica VT 1000S, Germany) to create an access to gray matter. After incubation during 45 min at 33°C, the spinal cord with the metal plate was transferred into the recording chamber where the metal plate provided mechanical stability of the preparation. Dorsal horn neurons were visualized in the whole spinal cord by using the light-emitting diode (LED) illumination according to a previous description (Safronov et al. 2007). In this study, we used blue and white LEDs positioned outside the solution meniscus for oblique illumination (Fig. 7). When the preparation was positioned in the recording chamber, care was taken to avoid direct shadow imposed by the dorsal root on the cut spinal cord surface. Tight-seal whole cell recordings from spinal dorsal horn neurons were done as previously described (Melnick et al. 2004; Safronov et al. 1997).
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We used a patch-clamp amplifier to record in nerves and roots the compound action potential currents (CAPCs), which are the currents associated with a propagation of compound action potentials (CAPs). In voltage-clamp mode, the amplifier monitored the extracellular current which had to be injected, to hold at 0 mV the voltage drop on the leakage resistance of the recording suction pipette at the moment when CAP invaded it. As shown in Fig. 1A, CAPCs represented recordings from the same population of conducting fibers as conventional CAPs, and CAPCs could be directly converted to CAPs by multiplication by the leakage resistance. We have chosen to record CAPCs instead of conventional CAPs because of higher resolution limits of patch-clamp amplifiers in voltage-clamp mode (the current-clamp mode resolution was limited by the size of the voltage digitization bin). CAPCs were recorded using EPC9 or EPC10 amplifiers (HEKA, Lambrecht, Germany). In control experiments comparing CAPs and CAPCs (Fig. 1A), we used EPC10 amplifier that had a current-clamp circuitry constructed as a real voltage-follower and thus allowed recording of fast voltage changes without signal distortion (Magistretti et al. 1996,1998). Suction electrodes for stimulation and recording were fabricated from borosilicate glass tubes with 1.5 mm OD and 0.86 or 1.2 mm ID (Modulohm, Denmark) and were fire-polished to fit the size of the nerve. Leakage resistance of the recording suction pipette with inserted nerve was 17–53 k
for dorsal roots, 27–87 k for sural nerves, and 6–28 k for spinal-sciatic nerve with a recording electrode always positioned on narrower mixed spinal nerve. Recording electrodes were voltage-clamped at 0 mV when CAPCs were measured. The voltage error due to resistance in series was supposed to be negligible for extracellular recording. The effective corner frequency of the low-pass filter was 14.3 kHz (with an exception of 2 and 3 recordings filtered at 2.9 and 8.5 kHz, respectively), and the frequency of digitization was 20–200 kHz. The traces with slow C-fiber responses were filtered off-line at 1 kHz. EPSCs evoked by attached root stimulation, were recorded in the spinal dorsal horn neurons using voltage-clamp mode of EPC9 amplifier. The patch pipettes were pulled from thick-walled borosilicate glass tubes with 1.5 mm OD and 0.86 mm ID (Modulohm, Denmark) and had after fire-polishing a resistance of 3–5 M. The effective corner frequency of the low-pass filter was 2.9 kHz. The frequency of digitization was 10 kHz. Offset potentials were compensated directly before formation of a seal. Liquid junction potentials were calculated and corrected for in all experiments. In neurons, the series resistance measured in the current-clamp mode was <14 M and was not compensated. The mean input resistance of the neurons was 2.3 ± 0.4 G (n = 10), and the mean resting potential measured with balanced amplifier input (Santos et al. 2004) was –81.0 ± 3.5 mV (n = 10).
Stimulation
Fibers were stimulated through suction electrodes using an isolated pulse stimulator (2100, A-M Systems). Short pulses of 50 µs were applied to activate CAPCs in A
β- and A-fibers. The intensity of effective stimulation depended on the type of fiber and was in the range of 10–250 µA for dorsal roots, 10–450 µA for spinal-sciatic nerve, and 10–200 µA for sural nerve. These stimulations were not sufficient to activate C-fibers. For activation of CAPCs in C-fibers, 1-ms pulses were used: 20–150 µA for dorsal root, 30–600 µA for spinal-sciatic nerve, and 10–120 µA for sural nerve. In the presence of TTX, the intensity of stimulation was increased as shown in Fig. 3C. CVs were calculated from latencies of corresponding CAPC waves measured from the end of 50-µs pulse for Aβ- and A-fibers and from the middle point of 1-ms pulse for C-fibers. Stimulus utilization time, i.e., the delay between the stimulus and beginning of the spike in the axon (Waddel et al. 1989), was not taken into account, assuming its negligible effect on pharmacological results. The Aβ- and A-fibers were stimulated at 0.5–1 Hz. The C-fibers were stimulated at 0.1 Hz because stimulations of our preparation at higher frequencies of 1, 0.5, and 0.2 Hz evoked responses with latencies progressively increasing with the number of stimulus (Gee et al. 1996). Except those in Figs. 1B and 6C, CAPCs are shown as averages of 
10 traces.
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ACSF contained (in mM) 115 NaCl, 3 KCl, 2 CaCl2, 1 MgCl2, 11 glucose, 1 NaH2PO4, and 25 NaHCO3 (pH 7.4 when bubbled with 95%-5% mixture of O2-CO2). Patch pipettes were filled with a solution containing (in mM) 3 KCl, 150 K-gluconate, 1 MgCl2, 1 BAPTA, and 10 HEPES (pH 7.3 adjusted with KOH, final [K+] was 161 mM).
Evaluation of the effect of TTX
TTX (Alomone labs, Jerusalem, Israel) was applied at 1- to 1,000-nM concentrations. In control experiments, we found that at low TTX concentrations, the nerve conduction block developed during 30–40 min (not shown). Therefore to ensure complete diffusion of the blocker within the nerve, each concentration was applied to the recording chamber for 1 h, and all measurements were done on stabilized responses. The wash out from TTX lasted 1–2 h. The effect of TTX was estimated by measuring reduction in the total area under the "positive" and "negative" deviations of the CAPC. In the spinal cord preparation, the block developed within few minutes (see Fig. 7), indicating that the primary afferent terminals within the spinal cord were likely better exposed to the blocker. Degree of the block was estimated from the area under the EPSC traces. For each measurement, at least five consecutive EPSCs evoked at 0.1 Hz were averaged.
All numbers are given as means ± SE. The values obtained by data fitting with a nonlinear least-squares procedure are given as means ± SE. In all figures, the error bars are shown when exceeding the symbol size. The parameters were compared by paired or independent Student's t-test. The present study is based on recordings from 72 dorsal roots, 23 spinal-sciatic nerves, 23 sural nerves, and 36 superficial dorsal horn neurons. All experiments, except those in Fig. 2 and 5, were carried out at room temperature of 22–24°C. To measure CVs and TTX effects at physiological conditions (Figs. 2 and 5), the bath temperature was increased to 35–37°C using the in-line solution heater (Warner Instruments, Hamden).
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RESULTS |
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To test whether the voltage-clamp mode of the patch-clamp amplifier can be used for adequate measurements of extracellular currents in dorsal roots and nerves, we compared the fast Aβ- and A-components of both compound action potential current (CAPC) and compound action potential (CAP) recorded in L5 dorsal root (Fig. 1A). In these experiments, EPC10 amplifier with a current-clamp circuitry designed as a real voltage-follower was used. Responses of the same root were recorded first under voltage-clamp conditions as CAPCs and then in current-clamp as CAPs. The Aβ-fiber CAPCs had amplitudes of 20–420 nA, and the corresponding CAPs were 
15 mV. To compare the shape of both signals, we scaled the trace of the CAPC by a factor close to the value of the leakage resistance for the recording electrode and superimposed (dotted line) on the CAP trace (Fig. 1A, right). In total five dorsal roots tested both CAPCs and CAPs showed the same kinetics. We therefore concluded that CAPCs recorded under voltage-clamp conditions can be used, instead of traditional CAPs, for adequate study of excitability in the whole nerve preparation. Thus the standard patch-clamp amplifier can be used for intracellular recordings from neurons as well as for study of population nerve action potentials. The following experiments were done in voltage-clamp mode and CAPCs were recorded.
Classification of Aβ-, A-, and C-fiber CAPCs
Stimulation of nerves with short (50 µs) current pulses evoked CAPCs in myelinated Aβ- and A-fibers, without activating slow C-fibers (Fig. 1B, top traces, n = 10). We considered the inflection point on the fast CAPC as the end of the Aβ-component and the beginning of the A-component. The end of the A-component was individually determined for each response by analyzing its decay at higher magnification (not shown). In many cases, Aβ-fibers started to activate at lower stimulation intensities than A-fibers. Stimulation of the nerve with long (1 ms) current pulses activated both the fast Aβ/A- and the slow C-fibers (Fig. 1B, bottom traces). The C-fiber CAPC had a typical biphasic shape; an inward current was followed by an outward current. The C-fiber response was measured from the beginning of the inward current to the end of the outward current. All events observed between the A- and C-responses were classified as A/C-fiber currents (Fig. 1B).
The CVs were measured in long (conduction distance, 24–44 mm) fragments of spinal-sciatic nerve at room temperature of 22°C (n = 5) and physiological temperature of 36–37°C (n = 5), and the results are shown in Fig. 2 and Table 1. The highest and lowest CVs were calculated for each component by dividing its shortest and longest latency times by the conduction distance. At room temperature, CV was 9.4–26.7 m/s (mean slowest CV –mean fastest CV) for Aβ-fibers, 1.1–9.4 m/s for A-fibers and 0.23–0.7 m/s for C-fibers. The CV values increased at 36–37°C to 17.4–44.2 m/s for Aβ-fibers, 1.8–17.4 m/s for A-fibers and 0.34–1.3 m/s for C-fibers. Taking into account that the CV changes along the afferent axons (Waddell et al. 1989), we also did measurements for dorsal roots (Table 2, n = 5; 11–13 mm conduction distance, 22–24°C). In comparison with the spinal-sciatic nerve, the root conducted more slowly giving the CV values of 4.2 to >16 m/s for Aβ-fibers, 0.56–4.2 m/s for A-fibers, and 0.16–0.43 m/s for C-fibers.
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The effect of TTX on CAPCs was studied in four regions along the central and peripheral branches of primary afferent axon (Fig. 3A): the rostral dorsal root, the caudal dorsal root, the mixed spinal nerve with the proximal sciatic nerve and the sural nerve. TTX at 3–1,000 nM blocked the C-fiber CAPCs, simultaneously increasing their latency and temporal dispersion as well as the stimulation intensity needed to evoke the corresponding maximum response (Fig. 3B). To account for the temporal dispersion of the C-fiber CAPC, we analyzed changes in its integrated area as a measure of block (Gokin et al. 2001; Huang et al. 1997). At each TTX concentration, the stimulation with increasing intensity was done and the corresponding maximum response was determined during the off-line analysis (Fig. 3C, all corresponding maximum responses are shown normalized to 1). The C-fibers in all four regions of primary afferents were sensitive to TTX, which at 1,000 nM suppressed 0.85 ± 0.02 (n = 5) of the CAPCs in the sural nerve, 0.89 ± 0.02 (n = 5) in the spinal-sciatic nerve, 0.65 ± 0.03 (n = 5) in the caudal dorsal root, and 0.66 ± 0.05 (n = 5) in the rostral dorsal root (Fig. 3D). The IC50 values obtained by fitting the data points for all regions ranged between 14 and 33 nM. The CAPC block in both regions of the peripheral branch occurred in a narrower concentration range. The remaining TTX-resistant component of the C-fiber CAPC was significantly larger in the central branch of the afferents (mean 0.34 and 0.35 for the rostral and caudal dorsal roots, respectively) compared with the peripheral branch (0.11 and 0.15 for the spinal-sciatic nerve and the sural nerve, respectively). Comparison using the independent Student's t-test gave the following values: P < 0.001 for the caudal dorsal root versus the spinal-sciatic nerve, P < 0.001 for the caudal dorsal root versus the sural nerve, P < 0.003 for the rostral dorsal root versus the spinal-sciatic nerve, and P < 0.01 for the rostral dorsal root versus the sural nerve. The differences in the TTX-resistant CAPC components within the central or peripheral branches were not significant: P = 0.89 for the caudal dorsal root versus the rostral dorsal root and P = 0.27 for the spinal-sciatic nerve versus the sural nerve.
For comparison, we also re-evaluated the data from the Fig. 3D and constructed the concentration-effect curves where the C-fiber CAPCs were measured as a negative peak current amplitude (Fig. 3E). Under these conditions, the apparent TTX block was stronger. The reduction of the peak CAPCs was seen at lower concentrations and the TTX-resistant CAPC components measured at 1,000 nM were smaller.
Control experiments were done to ensure that the C-fiber CAPCs remaining in the presence of 1,000 nM TTX were Na+-dependent. A substitution of 2 mM Ca2+ with a mixture of inorganic Ca2+ channel blockers (2 mM Co2+, 0.1 mM Cd2+, and 2 mM Mg2+) did not reduce the remaining CAPCs (n = 5; 2 rostral dorsal roots, 1 caudal dorsal root, and 2 sciatic nerves, not shown). In absence of TTX, the CAPCs completely disappeared when 115 mM NaCl in ACSF was substituted with 115 mM choline-Cl (n = 4, 2 rostral dorsal roots, 1 caudal dorsal root, 1 entire dorsal root, not shown). Thus it could be concluded that the C-fiber CAPCs were Na+-dependent.
The Aβ- and A-fiber CAPCs in all studied regions of the afferent axons were blocked by TTX in a narrow concentration range of 1–30 nM (Fig. 4, A and B). For this reason, the magnitudes of the Aβ- and A-fiber CAPCs were plotted as a function of TTX concentration for pooled data (n = 8) from two caudal dorsal roots, two rostral dorsal roots, two spinal-sciatic nerves, and two sural nerves (Fig. 4B). The half-maximum block was obtained at IC50 of 5–7 nM TTX.
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; Ogata and Tatebayashi 1993) has a strong effect on conduction in both A- and C-fibers. It could be concluded that the TTX-S Na+ channels blocked by 1–1,000 nM TTX play dominant role in conduction of both myelinated and unmyelinated sensory axons. C-fiber CAPC block by TTX increases with temperature
It has been recently shown for the rat cutaneous C-fiber terminals that the relative contribution of TTX-R Na+ channels to the spike generation was smaller at physiological temperature than at low (10°C) temperature (Zimmermann et al. 2007). Therefore we checked whether the magnitude of the TTX-R component of C-fiber CAPC depends on temperature. These experiments were performed on dorsal roots (n = 6; 8- to 11-mm conduction distance) because of the largest portion of TTX-R response observed in this region (see Fig. 3D).
The C-fiber CAPC was fist recorded at 22°C (Fig. 5A). Following this, the perfusing solution was heated to 35–37°C, and a control recording at the physiological temperature has been done. The CV of the C-fiber CAPC increased with a temperature by a factor of 2.1 ± 0.1 (n = 6, CV = 0.37 ± 0.02 m/s at 22°C vs. CV = 0.76 ± 0.03 m/s at 35–37°C, measured for the peak of negative wave, the time of the peak was determined by the trace fitting with a Gaussian function). Then the temperature returned to 22°C for a new control recording (control*), which was followed by the application of 1,000 nM TTX. In the presence of TTX, the temperature was again increased to 35–37°C (a part of the trace is shown at a higher amplification). The CV of the TTX-R portion of the CAPC increased with temperature by a factor of 1.7 ± 0.03 (n = 6). Still in the presence of TTX, the temperature was lowered to 22°C, and then the blocker was washed out. All changes in the magnitude of the C-fiber CAPCs (integrated areas) during these experiments are shown in Fig. 5B, where all responses were normalized to the first control measurement at 22°C. It is clear from Fig. 5, A and B, that the remaining TTX-R part of the C-fiber response was much smaller at physiological temperature than at room temperature.
Because control CAPCs slightly increased after each heating-cooling cycle (see Fig. 5B), we measured the TTX block by normalizing the responses in 1,000 nM to their nearest (preceding/following) controls (Fig. 5C). At 22°C, the remaining TTX-R responses of 0.29 ± 0.03 (n = 6; TTX 22°C normalized to control* 22°C) and 0.36 ± 0.04 (n = 6, TTX* 22°C normalized to recovery 22°C) were in agreement with our data from Fig. 3D. At 35–37°C, the remaining TTX-R component was 0.07 ± 0.02 (n = 6, TTX 35–37°C normalized to control 35–37°C). This increase in the block at physiological temperature was significant (P < 0.002, paired Student's t-test).
These experiments indicated that the contribution of TTX-R Na+ channels to spike propagation in dorsal root C-fibers at physiological temperature is much smaller than at room temperature.
Slowing of C-fiber conduction by TTX
To study the slowing of the C-fiber conduction with a block of TTX-S Na+ channels by nanomolar TTX, we measured the changes in the latency to the peak of negative wave in different blocker concentrations at room temperature (Fig. 6A, shown for a caudal dorsal root). The exact time moment of the peak of the negative C-wave was determined by locally fitting the data points (±2–30 ms from the peak) with a Gaussian function (see Fig. 6C). Latencies obtained for different TTX concentrations and normalized to those for control are shown in Fig. 6B. The first significant conduction slowing was seen at 3 nM (1.10 ± 0.02, n = 6, P < 0.003, paired Student's t-test). The effect increased with TTX concentration reaching its saturation at 300 nM, where the conduction was slowed by a factor of 2.8 ± 0.1 (n = 6). Fitting the concentration dependence of the conduction slowing with an isotherm gave the half-maximum effect concentration of 26 ± 2 nM (Fig. 6B).
In the following experiments, we tested whether the TTX-dependent increase in the C-fiber CAPC latency is also accompanied by an increase in the latency time variation. Both these parameters are important for defining criteria of identification of monosynaptic EPSCs in spinal dorsal horn neurons in the presence of TTX. We did measurements for 30 nM TTX, a concentration at which the C-fiber CAPC latency was increased twice (2.0 ± 0.1, n = 6, Fig. 6B). In these experiments, individual (not averaged) CAPC traces recorded in control and in 30 nM TTX were fitted with Gaussian function to obtain the current peak times (Fig. 6C, top), which were then normalized to their corresponding mean current (averaged CAPC trace) peak time and plotted as a histogram (Fig. 6C, bottom). In control, the variation of the current peak times was small, and all 120 measurements were within the ±2% range of the mean (1.00) with 114 of them (95%) within narrower range of ±1%. The latency variation substantially increased in 30 nM TTX where 147 of total 150 measurements (98%) were distributed within the ±4% range of the mean.
Our observations that nanomolar TTX blocked the major part of the C-fiber CAPC in dorsal roots and peripheral nerve and reduced by several times the axonal CV raised a question about the functional role of the TTX–R Na+ channels in the C-fiber conduction and synaptic transmission. To answer this question we studied how the application of 30 nM TTX changes the efficiency of synaptic transmission from C-fiber terminals to spinal dorsal horn neurons.
Effect of TTX on synaptic transmission
EPSCs activated by stimulation of attached L5 dorsal root were recorded at room temperature in superficial dorsal horn neurons most of which were located in substantia gelatinosa. The input was classified as A- or C-type on the basis of calculated CV (with a 1-ms allowance for synaptic transmission) and duration of stimulation used for its activation. Of a total of 36 dorsal horn neurons responding monosynaptically to primary afferent stimulation, 4 had only A-input, 15 only C-input, and 17 neurons had both A- and C-fiber inputs. A-fiber-mediated EPSCs were elicited by a short (50 µs) pulse stimulation of dorsal roots and the corresponding CV for the fiber ranged from 0.74 to 3.56 m/s. The activation of C-input always needed a 1-ms stimulation of the root and the calculated CV ranged from 0.19 to 0.65 m/s. In control, the EPSCs were considered as monosynaptic if no failure was seen in 10 consecutive stimulations (0.1 Hz) and a variation of the EPSC latency did not exceed 1 ms. In the case of the multi-component C-fiber-mediated EPSCs, each component was analyzed using these criteria. Polysynaptic responses were not considered.
In all tested neurons, monosynaptic A-inputs were completely and reversibly blocked by 30 nM TTX (Fig. 7A; n = 15). TTX at 30 nM was also applied to 21 neurons with C-fiber input. In seven of them, the C-fiber-mediated EPSCs were completely blocked (Fig. 7A). In all these cases, the C-fiber-mediated EPSCs first showed increased latencies and then completely disappeared within 5–10 min of TTX perfusion. In 14 of 21 neurons with C-fiber input, the EPSCs were incompletely blocked (Fig. 7B). In recordings done during TTX perfusion, these C-fiber-mediated EPSCs showed progressively increasing latencies and latency variations, decreasing magnitudes and increased number of failures. An increase in the stimulation strength did not reduce the number of failures. After the stabilization of the block, the variation of the EPSC latency was within the ±4% range of the mean, in agreements with predictions of our CAPC experiments. The mean increase in the EPSC latency was 1.7 ± 0.1 (n = 14, range: 1.3–2.6). To estimate the degree of the synaptic transmission block, we used the ratio of the mean EPSC areas in at least five consecutive episodes in TTX and control. In 14 neurons with incomplete EPSC block, 30 nM TTX suppressed C-fiber synaptic transmission by 76.4 ± 6.6%. In total 21 neurons tested for 30 nM TTX, the mean block of C-fiber transmission was 84.3 ± 5.0%. Thus 30 nM TTX blocked the major portion of C-fiber input to the spinal cord. To test whether the remaining part was also TTX-sensitive, we applied 300 nM concentration of TTX (still ineffective for TTX-R Na+ channels) (Elliott and Elliott 1993) in 14 neurons, including 2 with the largest EPSCs remaining in 30 nM TTX (77 and 81% of control). In all 14 cases, 300 nM TTX completely blocked the EPSCs within 2–5 min of perfusion (Fig. 7C). Thus the A- and C-fiber inputs to the spinal dorsal horn neurons were blocked with suppression of TTX-S Na+ channels by nanomolar TTX.
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DISCUSSION |
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-fiber conduction as well as synaptic transmission from both A- and C-fibers to the spinal cord. This indicated that spike propagation and synaptic release in both types of thin afferent axons are predominantly based on TTX-S Na+ channels. In unmyelinated C-fibers, TTX-S Na+ channels determine physiological CVs and trigger transmitter release to the second-order sensory neurons in the spinal dorsal horn. This dominance of TTX-S Na+ channels is not changed along the peripheral and central branches of primary afferents. Under physiological conditions, TTX-R Na+ channels seem to be expressed at densities insufficient to provide synaptic transmission to the spinal cord.
We used a voltage-clamp mode of a patch-clamp amplifier to record CAPCs that showed the time course similar to conventional CAPs seen under current-clamp conditions. The voltage-clamp amplifier provided high resolution of CAPCs with amplitudes in nanoampere range, and our classification of A- and C-fibers on the basis of their CVs was in agreement with previous studies done at both physiological and room temperatures (Gokin et al. 2001; Grudt and Perl 2002; Harper and Lawson 1985; Villiere and McLachlan 1996; Waddel et al. 1989).
Both Aβ- and A-fibers were completely suppressed by 30 nM TTX with IC50 of 5–7 nM. TTX-S component of the C-fiber CAPC was blocked in a concentration range between 1 and 100 nM with IC50 of 14–33 nM. These IC50 values are typical for TTX-S Na+ channels in a number of preparations (Ogata and Tatebayashi 1993; Safronov et al. 1997; Takahashi 1990). Slightly higher sensitivity obtained in our experiments for myelinated nerves might reflect structural differences in fiber organization. It is possible that TTX, as a membrane impermeant blocker, has better access to Na+ channels in nodes of Ranvier of myelinated axons than to those in unmyelinated axons grouped in Remak bundles and surrounded by Schwann cell membrane (Murinson and Griffin 2004).
The C-fiber CAPCs were blocked in 1,000 nM TTX by 65–89% at room temperature and by 93% at 35–37°C, implying that the major part of the response was carried through TTX-S Na+ channels. This observation raised a question about the physiological role of TTX-R Na+ channels in unmyelinated C-fiber axons. Presence of a small TTX-R component of CAPC could be explained either by the existence of a small population of C-fibers with pure TTX-R conduction or, alternatively, by the presence of some TTX-R Na+ channels in C-fibers with predominantly TTX-S conductance. Our experiments with a latency increase due to Na+ channel block by TTX have strongly supported the second hypothesis. If C-fibers with pure TTX-R spikes were present in the dorsal root, one would expect that detectable portion of the C-fiber CAPC would not increase its latency in 30–300 nM TTX. In contrast, in all our experiments, the entire C-fiber CAPC was progressively slowed by two to three times in a concentration-dependent manner, indicating that there were no fibers unaffected by low TTX concentrations. Interestingly, the half-maximum effect concentrations for the CAPC suppression and the latency increase were close to each other (14–33 nM for the CAPC block vs. 26 nM for the latency increase), once more indicating that both processes are likely to depend on the block of the same type of channel. Thus one can conclude that TTX-S Na+ channels are critically important for spike propagation and determination of the physiological CV in C-fibers. After block of TTX-S channels, TTX-R Na+ channels can probably provide only slowly propagating spikes in C-fibers.
Our experiments have also shown that the relative contribution of TTX-R Na+ channels to the C-fiber response changes along the axon in such a way that the central branch of the afferent axon had significantly higher percentage of TTX-R component than its peripheral branch. Within the branches, however, the difference in the relative TTX-R channel expression was found to be not significant. This may imply differential mechanisms of regulation of TTX-S/TTX-R Na+ channel expression in the central and peripheral axon branches of primary sensory neuron. However, in spite of this variation in the expression of functional TTX-R Na+ channels along the extension of the axon, its conduction was always dominated by TTX-S Na+ channels.
In the spinal cord preparation, 30 nM TTX had stronger effect on C-fiber-mediated EPSCs (84% block) than on the C-fiber CAPCs in isolated dorsal roots (45% block for rostral and 30% block for caudal, calculated from the fitting curves in Fig. 3D). In the presence of TTX, the EPSCs always showed a typical increase in their latency corresponding to the slowing of conduction observed in the isolated roots. Most axonal spikes reaching the spinal cord became unable to trigger synaptic release to dorsal horn neurons. In 300 nM TTX, the synaptic conduction was completely blocked. This full block of neurotransmitter release could not be attributed to blockade of C-fiber conduction, as 300 nM TTX only reduced responses in dorsal roots to 37–38% of control (fitting curves in Fig. 3D). Thus TTX-R action potentials could still arrive at the terminal, but transmitter release did not occur. One can assume, therefore, that TTX-R Na+ channels alone cannot provide spikes of amplitude sufficient to activate presynaptic high-threshold voltage-gated Ca2+ channels critical for transmitter release from the C-fiber axon terminals (Bao et al. 1998; Heinke et al. 2004). Our data also indicate that 30 nM TTX cannot be used as a tool for separating A- and C-fiber-mediated EPSCs. Separation of these components in 50 nM TTX observed by Yoshimura and Jessell (1990) might be due to incomplete perfusion of the slice with the blocker.
Our data agree with studies showing that under physiological conditions, C-fiber conduction is blocked by TTX (Villiere and McLachlan 1996; Yoshida and Matsuda 1979). Partial resistance of C-fiber-mediated transmission to the spinal cord observed in experiments where TTX was locally applied to primary afferents (Jeftinija 1994; Jeftinija and Urban 1994; Steffens et al. 2001), may be also explained in context of our results, by assuming that in some C-fibers TTX-R Na+ channels have sufficient density to provide slowly propagating low-amplitude spikes able to cross the region of TTX application. In this case, the TTX-S Na+ channels in the dorsal root part rostral to the region of TTX application could restore original spike amplitude needed to trigger synaptic release. In contrast to rat axons, in amphibian sciatic nerve the TTX-R Na+ channels appear to play more important role in C-fiber conduction (Buchanan et al. 1996; Kobayashi et al. 1993). Large TTX-R C-fiber responses were also seen in human sural nerve biopsied from patients with different types of neuropathy (Grosskreutz et al. 1996; Quasthoff et al. 1995). However, it is not clear whether or not this strong expression of TTX-R Na+ channels in unmyelinated C-fiber axons was caused by the neuropathies.
Our results suggest that under normal conditions, an ability of TTX-R Na+ channels to generate action potential is probably restricted to the soma of small primary sensory neurons and their peripheral endings. Indeed, the somata showed intense immunoreactivity for synthesized proteins of the TTX-R Na+ channel types Nav1.8 and Nav1.9 (Amaya et al. 2000; Djouhri et al. 2003; Fang et al. 2002; Novakovic et al. 1998; Tate et al. 1998), and the density of functional membrane-inserted channels was sufficient for spike generation in the presence of TTX (Blair and Bean 2002; Ritter and Mendell 1992; Villiere and McLachlan 1996; Yoshida and Matsuda 1979). In the peripheral endings, the TTX-R Na+ channels were also shown to be capable of generating action potentials (Brock et al. 1998; Carr et al. 2002; Strassman and Raymond 1999; Zimmermann et al. 2007). Nav1.8 and Nav1.9 protein immunoreactivity was also found in axoplasm of myelinated and unmyelinated nerve fibers (Coward et al. 2000; Fjell et al. 2000; Liu et al. 2001) and central primary afferent terminals (Amaya et al. 2000; Novakovic et al. 1998). However, according to our data, the density of functional TTX-R Na+ channels expressed in the axonal membrane is negligible in myelinated A-fibers and is insufficient for full-amplitude spike conduction in unmyelinated C-axons. The role of axonal TTX-R Na+ channels may change under neuropathic conditions when translocation of intracellularly located channel proteins from the soma to the peripheral axon with subsequent accumulation at the site of injury can create the source of ectopic discharge (Devor and Seltzer 1999; Novakovic et al. 1998).
Our results also reveal that the contribution of TTX-R Na+ channels to spike propagation at physiological temperature is surprisingly small. In dorsal roots, the TTX-R component formed only 7% of the C-fiber CAPC at 35–37°C, in comparison with 34–35% measured at 22–24°C. Increasing temperature was reported to have several effects on Na+ channels and passive membrane properties (Zimmermann et al. 2007). It increases the amplitudes of both TTX-S and TTX-R voltage-gated Na+ currents, accelerates their activation kinetics, reduces the steady-state slow inactivation of TTX-S (but not TTX-R) Na+ channels, and decreases membrane input resistance. This overall increase in TTX-S Na+ current together with its acceleration can explain about twofold increase in CV of the C-fiber CAPC (without reduction in magnitude) observed after temperature increase from 22 to 35–37°C in the absence of TTX. By contrast, when temperature was increased in the presence of TTX, the increase in CV for the remaining TTX-R component was accompanied by its four- to fivefold reduction. It can be assumed that a decrease in membrane input resistance at 35–37°C could be compensated by the temperature-dependent augmentation and acceleration of TTX-R Na+ currents only in a few percent of C-fiber axons, whereas in a vast majority of them conduction collapsed. These experiments provided one more evidence that at room temperature TTX-R Na+ channels can only support a propagation of low-amplitude spikes with a low safety factor, which collapse in most C-fibers at physiological temperature.
From the functional point of view, it seems that in mammals, TTX-R Na+ channels are molecules for nociception in the cold and for cold pain (Nav1.8) (Zimmermann et al. 2007) designated for the peripheral sensory nerve endings. Unique properties of the TTX-R Na+ channels, like the cold resistance of slow inactivation, make them suitable for the electrical signal generation at tissue-damaging levels of cold. However, in the peripheral (nonterminal) and central branches of C-fiber axons, which in mammals do not experience temperatures below physiological level, the contribution of the TTX-R Na+ channels to signal conduction is very small and their physiological function remains unclear. In contrast to mammals, TTX-R Na+ channels are strongly involved in the physiological C-fiber conduction in cold-blooded animals (Buchanan et al. 1996; Kobayashi et al. 1993).
In conclusion, our study has shown that TTX-S type of Na+ channels dominates conduction in fine-caliber primary afferents. It was the only type of Na+ channels underlying A-fiber responses. In C-fiber axons, the TTX-S Na+ channels determine the physiological CV and control synaptic transmission. Although TTX-R Na+ channels were revealed at least in a part of C-fibers, they could not provide propagation of full-amplitude spikes able to trigger synaptic release in the spinal cord. In this context, the gain of the function of axonal TTX-R Na+ channels by their modulation, expressional upregulation or redistribution (England et al. 1996; Gold et al. 1996; Khasar et al. 1998; Lai et al. 2002; Novakovic et al. 1998; Waxman et al. 1999; Zhang et al. 1997) may be a key factor in developing diverse pathological states.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: B. V. Safronov, Instituto de Biologia Molecular e Celular - IBMC, Rua do Campo Alegre 823, 4150-180 Porto, Portugal (E-mail: safronov{at}ibmc.up.pt)
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ABSTRACT
INTRODUCTION
) and A
) as a function of TTX concentration. The data were from 2 caudal dorsal roots, 2 rostral dorsal roots, 2 spinal-sciatic nerves, and 2 sural nerves. The points were fitted with parameters IMax = 1.0, IC50 = 6.8 ± 0.7 nM, and n = 1.8 ± 0.3 for the A