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1Department of Neurology and 2Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven; and 3Rehabilitation Research Center, Veterans Administration Connecticut Healthcare System, West Haven, Connecticut
Submitted 27 September 2006; accepted in final form 10 November 2006
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ABSTRACT |
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INTRODUCTION |
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). Peripherally, these two subpopulations terminate in different epidermal strata (Zylka et al. 2005). Centrally, IB4+ neurons project to dorsal horn inner lamina II and are responsive to glial cell line–derived neurotrophic factor (GDNF), whereas IB4– neurons project to lamina I and outer lamina II and are responsive to NGF (Snider and McMahon 1998). Using genetically regulated transneuronal tracer, Braz et al. (2005) showed that IB4+ and IB4– nociceptors signal pain through distinct parallel central pathways. Stucky and Lewin (1999) reported that IB4+ neurons have longer-duration action potentials (APs), higher AP threshold, and produce larger tetrodotoxin-resistant (TTX-R) currents than IB4– neurons in response to a long step current injection stimulus. Fang et al. (2006), studying functionally identified C-nociceptor DRG neurons, found not only that strongly IB4+ cells have longer AP durations and rise times, but also that these cells have slower conduction velocities and more negative resting membrane potentials than those of IB4– neurons. Taken together, these neurochemical, neuroanatomical, and electrophysiological differences suggest that IB4+ and IB4– neurons may be functionally distinct.
Nav1.8, a TTX-R sensory neuron-specific voltage-gated sodium channel, produces a slowly inactivating sodium current characterized by depolarized voltage dependency (Akopian et al. 1996; Sangameswaran et al. 1996) and rapid recovery from fast inactivation (Cummins and Waxman 1997; Elliott and Elliott 1993; Schild and Kunze 1997). Nav1.8 channels produce the majority of the inward current during the AP upstroke in the DRG neurons in which they are present (Blair and Bean 2002; Renganathan et al. 2001), most of which are nociceptive (Djouhri et al. 2003). TTX-R currents attributable to Nav1.8 enter rapidly into, and recover slowly from, slow inactivation even during short depolarizing pulses from holding potentials near resting membrane potential, thereby contributing to adaptation of firing in response to capsaicin application (Blair and Bean 2003). However, the degree of use-dependent reduction of TTX-R current that could contribute to adaptation is quite variable (35–70%) within DRG neurons (Blair and Bean 2003; Gold and Thut 2001; Roy and Narahashi 1992; Rush et al. 1998). In this study we present data that show that use-dependent reduction and the kinetics of slow inactivation of Nav1.8 current are distinct in IB4+ and IB4– subpopulations of rat small DRG neurons and suggest that these differences contribute to differences in high-frequency firing properties in these two groups of DRG neurons.
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METHODS |
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DRG cultures followed the protocol of Rizzo et al. (1994). Briefly, adult Sprague–Dawley rats (1–2 mo old) were decapitated and L4/L5 DRGs were quickly removed and desheathed in sterile complete saline solution (CSS), pH 7.2, enzymatically digested for 25 min with collagenase A (1 mg/ml; Roche, Indianapolis, IN) and then with collagenase D (1 mg/ml; Roche) and papain (30 U/ml; Worthington, Lakewood, NJ) in CSS at 37°C, and gently centrifuged (100 x g for 3 min). Pellets were triturated in DRG media (1:1 DMEM/F12, 10% FCS, 100 U/ml penicillin, and 0.1 mg/ml streptomycin) containing 1.5 mg/ml BSA (Fraction V; Sigma, St. Louis, MO) and 1.5 mg/ml trypsin inhibitor (Sigma). Cells were plated on poly-ornithine-laminin-coated glass coverslips, flooded with DRG media after 1 h, and incubated at 37°C (humidified 95% O2-5% CO2). Immediately before recording, neurons were incubated with 3 µg/ml IB4-Alexa Fluor 488 (Molecular Probes, Eugene, OR) in the full-strength Na+ bath solution (see following text) for 5 min and rinsed twice for 2 min to wash out nonspecific binding. IB4-Alexa Fluor 488 was previously shown to have no effect on DRG neuron excitability (Rush et al. 2005; Vydyanathan et al. 2005).
Electrophysiological recordings
Whole cell patch-clamp recordings were made from small (
25 µm diameter) DRG neurons at room temperature (21–25°C) within 8 h after plating, using Axopatch 200B amplifiers (Axon Instruments, Foster City, CA). For currents >20 nA, we switched to a 50-M
feedback resistor (
of 0.1), which can pass 200 nA. Micropipettes (0.6–0.9 M) were pulled from capillary glass (PG10165-4; WPI, Sarasota, FL) with a Flaming-Brown P80 puller (Sutter, Novato, CA), and polished on a microforge. Pipette tips were wrapped with Parafilm to reduce capacitance, permitting fast current clamp with low-resistance pipettes. Cells were not considered for analysis if they had high leakage currents (holding current >0.5 nA at –70 mV of holding potential) or an access resistance >2 M. For current-clamp recording, the pipette solution contained (in mM): 140 KCl, 1 EGTA, 10 NaCl, 2 Mg-ATP, and 10 HEPES, pH 7.3 (adjusted to 310 mOsm/l with sucrose). The full-strength Na+ bath solution contained (in mM): 140 NaCl, 3 KCl, 1 MgCl2, 1 CaCl2, 10 glucose, and 10 HEPES, pH 7.3 (adjusted to 320 mOsm/l with sucrose). To isolate Nav1.8 currents in voltage-clamp recording, 20 mM TEA-Cl, 0.1 mM CdCl2, and 300 nM TTX were included in the bath solution to inhibit endogenous K+, Ca2+, and TTX-S Na+ currents, and the 140 mM KCl in the pipette solution was replaced with 140 mM CsF; fluoride in the pipette solution is associated with a fast time-dependent shift of voltage dependency of activation and inactivation of Nav1.9 in the hyperpolarizing direction, although Nav1.8 currents are unaffected (Coste et al. 2004). Mg-ATP (2 mM) was omitted in voltage clamp. Pipette potential was zeroed before seal formation and voltages were not corrected for liquid junction potential. Whole cell Na+ currents and APs were filtered at 5 and 10 kHz, and acquired at 50 and 100 kHz, respectively, using Clampex 8.2 (Axon Instruments). Capacity transients were cancelled before switching to current-clamp mode; series resistance was compensated (90%) in all experiments.
Protocols and data analysis
For voltage-clamp recording, DRG neurons were held at –70 mV. Recording was started 10 min after establishing whole cell configuration to allow currents to stabilize and minimize contamination by residual persistent TTX-R Nav1.9 currents (Choi et al. 2006; Coste et al. 2004; Cummins et al. 1999). Leakage current was subtracted using hyperpolarizing control pulses, applied after the test pulse (P/N subtraction). Use-dependent reduction was determined using 60 repetitive 100-ms depolarizing pulses to –10 mV from a holding potential of –70 mV at 1 Hz.
The kinetics of entry of Nav1.8 channel into slow inactivation were fitted with a single-exponential function
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, t is duration of the conditioning pulse, and C is a steady-state asymptote.
Recovery from slow inactivation of Nav1.8 channel was fitted with single- and double-exponential functions
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1 and 2, respectively; t is recovery duration; and C is maximal recovery asymptote. To eliminate cell-to-cell variations in resting potential for current clamp, steady polarizing currents were applied to set a holding potential of –60 mV, when required. APs were elicited by three types of current injection: short (0.5 ms), long (400 ms), and ramp (0 to 1, or to 2 nA in 400 ms). To determine voltage threshold, AP peak, and duration, short current injections were used so that the AP was uncontaminated by injected current. Long current injections were used to measure current threshold and the number of APs evoked by injecting a 500-pA current. Responses to ramp current injection were also analyzed.
Student’s unpaired t-tests were used (criterion for statistical significance, 0.05). Descriptive data are presented as means ± SE. Data were analyzed using Clampfit 8.2 (Axon Instruments) and Origin 6.1 (Microcal, Northampton, MA).
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RESULTS |
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The largest peak TTX-R sodium current amplitude recorded from acutely isolated DRG neurons in the presence of 300 nM TTX was 39 nA. To voltage clamp these large currents adequately, we used low-resistance pipettes (about 0.8 M) and 90% series resistance compensation (Fig. 1 A). For our characterization of the voltage-dependent and kinetic properties of Nav1.8 current, cells were held at –70 mV for 10 min using fluoride-based pipette solution to inactivate Nav1.9 current (Choi et al. 2006; Coste et al. 2004). Because Nav1.9 channels produce persistent currents during the depolarization pulses of 100 ms (Dib-Hajj et al. 2002) and are more heavily expressed in IB4+ than in IB4– cells (Fang et al. 2006; Fjell et al. 1999; Rush et al. 2005) we estimated the extent of possible contamination of Nav1.9 current using the ratio of maximum late current amplitude at the end of 100-ms pulses to peak current amplitude. The maximal percentage of late current amplitudes to peak current for IB4+ and IB4– neurons was 5.0 ± 0.5 and 3.0 ± 0.5%, respectively, and were significantly different (P < 0.01). The late current under our recording conditions could also have come from noninactivated Nav1.8 channels at voltages near the potential at which the channel activates (e.g., at –25 mV). The small difference of the amplitude of the late current (roughly 2%) between IB4+ and IB4– neurons and the possibility of its contamination by noninactivating Nav1.8 channels suggest that the effect of the apparent contamination of Nav1.9 currents on the measured kinetic properties of Nav1.8 is negligible.
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and the estimated maximum voltage error after series resistance compensation was 3.3 ± 0.3 mV. Voltage dependencies of activation of Nav1.8 in IB4+ and IB4– neurons are shown in Fig. 1B. Reversal potentials of Nav1.8 peak currents were 71.7 ± 1.0 mV for IB4+ neurons and 72.1 ± 1.1 mV for IB4–, close to the calculated Nernst potential (69.9 mV). The V1/2 for activation of Nav1.8 in IB4+ neurons was –16.7 ± 0.2 mV and the slope k was 4.1 ± 0.2 mV. The V1/2 for activation of Nav1.8 in IB4– neurons was –15.7 ± 0.3 mV and the slope value was 4.2 ± 0.2 mV. The midpoint values were not significantly different (P > 0.05).
Steady-state fast inactivation curves for IB4+ and IB4– cells, measured with 500-ms depolarizing prepulses, are shown in Fig. 1B. The V1/2 of steady-state fast inactivation was –31.8 ± 0.2 mV and the slope factor k was 4.9 ± 0.2 mV for IB4+ neurons and –31.9 ± 0.1 and 4.2 ± 0.1 mV, respectively, for IB4– neurons. The midpoint values for inactivation were not significantly different (P > 0.05).
Stronger use-dependent reduction of Nav1.8 current in IB4+ DRG neurons than in IB4–
Nav1.8 channels in DRG neurons show a broad spectrum of use-dependent current reduction (Blair and Bean 2003; Gold and Thut 2001; Roy and Narahashi 1992; Rush et al. 1998), but the basis for this variability has not been fully understood. We found that use-dependent reduction of Nav1.8 current was significantly different (P < 0.001) in IB4+ and IB4– small DRG neurons (Fig. 1, C and D). Sixty 100-ms depolarizing pulses to –10 mV from a holding potential of –70 mV were applied at 1 Hz (Fig. 1C). Residual peak Nav1.8 current amplitude at the 60th depolarizing pulse decreased to 31.3 ± 2.2% in IB4+ neurons and only to 60.0 ± 2.7% in IB4– neurons (Fig. 1D).
Nav1.8 channels in IB4+ DRG neurons enter quickly into slow inactivation
To determine whether the differences in use-dependent reduction of Nav1.8 currents in IB4+ and IB4– neurons reflect differences in the degree of slow inactivation elicited by short depolarizations, we measured the entry of Nav1.8 into slow inactivation using 2.5- to 1,500-ms conditioning pulses to –10 mV from a holding potential of –70 mV (Fig. 2, A and B). The conditioning pulse was followed by a 40-ms step to –70 mV for recovery from fast inactivation. When the conditioning pulse was lengthened to 100 ms at –10 mV, the same depolarizing pulse for the use-dependent protocol, the degree of slow inactivation of Nav1.8 in IB4+ neurons was 62.8 ± 1.6% (n = 9), whereas that of IB4– neurons was 16.4 ± 2.1% (n = 6). The entry of Nav1.8 into slow inactivation was well fitted by a single-exponential function (Fig. 2B). Time constants for entry of Nav1.8 into slow inactivation in IB4+ and IB4– neurons were 137.0 ± 22.4 ms (n = 9) and 544.8 ± 58.9 ms (n = 6), respectively. Based on a single-exponential fit, 23.4 ± 2.8% of Nav1.8 channels in IB4– neurons are expected not to enter the slow inactivation state (as reflected by the offset, i.e., steady-state asymptote in the plot of inactivation vs. conditioning pulse duration), compared with only 5.8 ± 0.7% of Nav1.8 channels in IB4+ neurons. The time constant and offset of entry into slow inactivation state for Nav1.8 current in IB4+ and IB4– neurons were significantly different at all voltages (Fig. 2, C and D; P < 0.01).
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We analyzed recovery from slow inactivation of Nav1.8 by measuring peak current after a step depolarization to –10 mV. Slow inactivation was elicited with a 5-s pulse to +10 mV, enough to induce steady-state slow inactivation in both IB4+ and IB4– neurons, and was followed by a recovery potential (–50, –70, –90, or –110 mV) of variable duration (Fig. 3, A and B). All currents were normalized to the maximal current after 90 s at a conditioning recovery potential of –110 mV. The kinetics of recovery from slow inactivation at recovery potentials of –50 and –70 mV in both IB4+ and IB4– neurons were well fitted by single-exponential functions (dotted lines), whereas data from recovery potentials of –90 and –110 mV were better fitted by double-exponential functions (solid lines). The absence of a good fit for recovery from inactivation at hyperpolarized potentials might be explained by the recovery from ultraslow inactivation of Nav1.8 current and/or contamination by recovery from inactivation of Nav1.9 current at hyperpolarized potentials. The recovery from slow inactivation of Nav1.8 current in IB4– neurons at all voltages was significantly faster than that in IB4+ neurons (Fig. 3, C and D, P < 0.01), except for the recovery potential of –50 mV, a recovery potential at which the offset in IB4+ neurons is significantly smaller than that in IB4– neurons. The fast component of recovery from slow inactivation in IB4– neurons fitted by double-exponential functions at –90 and –110 mV was also faster than that in IB4+ neurons (P < 0.05).
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Because slow inactivation of TTX-R current was previously shown to contribute to adaptation of AP firing in DRG neurons (Blair and Bean 2003), we asked whether there are differences in excitability and AP adaptation in IB4+ versus IB4– neurons. APs were recorded from 27 IB4+ and 25 IB4– small (diameter 25 µm) DRG neurons in current-clamp mode without addition of internal GTP (which is widely used in pipette solution for current-clamp but not for voltage-clamp studies). The rationale for excluding GTP arises from the fact that intracellular GTP increases DRG neuron excitability by upregulation of Nav1.8 (Saab et al. 2003) and of persistent Nav1.9 current (Baker et al. 2003), which is predominantly present in IB4+ neurons (Cummins et al. 2000; Fjell et al. 2000; Rush et al. 2005). To exclude additional effects of upregulated TTX-R currents and to minimize possible time-dependent channel rundown in the absence of GTP in the pipette, recordings were made within 15 min after establishing whole cell configuration. Mean cell capacitance and mean input resistance of IB4+ and IB4– neurons, measured in voltage clamp, were not significantly different (Table 1). The average resting membrane potential of IB4– neurons (–48.9 ± 1.6 mV) was significantly depolarized compared with IB4+ cells (–59.9 ± 1.9 mV; P < 0.001). To avoid cell-to-cell variations, neurons were held at –60 mV. We measured APs using short (0.5-ms) current injections to minimize the effect of the injected stimulus current (Fig. 4, A and B). Current threshold, i.e., the current required to generate the all-or-none AP, was significantly lower in IB4– (1.8 ± 0.1 nA) than in IB4+ neurons (2.3 ± 0.2 nA; P < 0.05). However, the mean AP peak, AP duration at 0 mV, and mean voltage threshold (i.e., voltage for take-off of an all-or-none AP) were not significantly different in IB4+ and IB4– neurons (Table 1).
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To further compare adaptation of AP firing in IB4+ and IB4– DRG neurons, current stimuli were injected for a longer period (400 ms) (Fig. 4, C and D). The mean current threshold for AP generation in IB4+ neurons under these conditions was significantly higher than that in IB4– neurons (Table 1). However, voltage threshold was the same as that with 0.5-ms current injection (about 30 mV) and did not show a significant difference between the two subpopulations. Only 22% (six of 27) of IB4+ neurons generated multiple (two to 11 APs) APs at 500 pA stimulus current for 400 ms, whereas 60% (15 of 25) of IB4– neurons produced multiple APs (two to 19 APs) under the same stimulation protocol.
We also used 400-ms ramp current injections from 0 pA to 1 nA to examine the adaptation of APs (Fig. 5, A and B). Because APs elicited by ramp current injection showed especially strong adaptation in IB4+ neurons we counted only overshooting APs (i.e., APs with peaks >0 mV). Most IB4+ neurons (20 of 27) generated two or more overshooting APs in response to this stimulus, but in all of the cells APs quickly adapted, showing rapid decrement of amplitude (Fig. 5A). Nearly all IB4– neurons (24 of 25) produced two or more APs and more than half (13 of 24) of these neurons did not show the decline of AP amplitude during a 400-ms ramp current injection (Fig. 5B). To measure the adaptation, we compared the peak of the last AP-like response near the end of ramp current injection to the first AP peak elicited by the ramp depolarization, normalizing both values to the holding potential of –60 mV. As shown in Fig. 5E (left), adaptation was significantly stronger in IB4+ neurons (P < 0.01). Increasing ramp current injection to 2 nA generated stronger AP adaptation of the same neurons in both groups (Fig. 5, C and D). As with 1-nA stimuli, adaptation occurred more rapidly in IB4+ neurons. The average adaptation in response to 2-nA ramp injection current was significantly stronger for IB4+ neurons than for IB4– neurons (Fig. 5E, right; P < 0.01).
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DISCUSSION |
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; Waxman 1999). Nav1.8 currents contribute substantially to spontaneous activity in damaged sensory axons (Roza et al. 2003) and were previously shown to respond to the application of inflammatory agents (Black et al. 2004; Gold et al. 1996; Jin and Gereau 2006; Okuse et al. 1997; Tanaka et al. 1998), suggesting that this channel is involved in inflammation-evoked pain (Wood and Waxman 2005; Wood et al. 2004). We show in this study that Nav1.8 channels in IB4+, but not IB4– small DRG neurons, enter quickly into, and recover slowly from, slow inactivation, a characteristic associated with greater use-dependent reduction of the Nav1.8 current amplitude in IB4+ compared with IB4– neurons. We also observed a lower current threshold for AP generation, an increased tendency to generate multiple APs, and less decrement of AP amplitude during repetitive firing in IB4– compared with IB4+ neurons. These data suggest that differential modulation of Nav1.8 channels contributes to the functional difference between these two DRG neuron subtypes.
Small-diameter DRG neurons are notable in displaying a broad range of use-dependent reduction of TTX-R current (Blair and Bean 2003; Gold and Thut 2001; Roy and Narahashi 1992; Rush et al. 1998). The present study demonstrates a cell background-dependent contribution to this variability and indicates that differences in slow inactivation of Nav1.8 between IB4+ and IB4– neurons can contribute to the spectrum of use-dependent reduction of TTX-R current in small DRG neurons. In these earlier studies, differences in the duration of the depolarizing pulse, holding potential, and cellular background may also have contributed to the differences that were observed in the degree of use-dependent reduction of Nav1.8 current.
Nav1.8 produces the majority of the current underlying the depolarizing phase of AP and is critically important for production of multiple APs during high-frequency activity (Blair and Bean 2002; Renganathan et al. 2001). We show in the present study that 60% of IB4– neurons produce multiple APs in response to a sustained strong (500-pA) step current injection, whereas only 22% of IB4+ neurons generate multiple APs. Moreover, IB4– neurons were able to generate sustained AP activity without decrement of AP amplitude, whereas IB4+ neurons displayed pronounced decrement in AP overshoot. Our results suggest that the basis for the differential response of these two subpopulations of DRG neurons to a sustained stimulus is the slower entry of the channel into slow inactivation in IB4– neurons compared with that in IB4+ neurons. This effect of slow inactivation of Nav1.8 on adaptation of DRG neurons is in agreement with the previously published data from Bair and Bean (2002) and extends these findings to demonstrate that this effect is dependent on the neuronal background. Our finding also suggests that entry of Nav1.8 into slow inactivation is a biophysical property of the channel that is regulated in a cell type–dependent manner.
Although the molecular basis for differential use-dependent reduction of Nav1.8 currents in IB4+ and IB4– neurons is not fully understood, it should not be too surprising that the distinct cell backgrounds of IB4+ and IB4– neurons influence the expression and/or properties of Nav1.8. It is well established that these two subpopulations are characterized by distinct neurochemical profiles and different responses to neurotrophic factors (Snider and McMahon 1998). It was recently shown that IB4+ neurons, in contrast to IB4– neurons, possess a PKA-independent, c-AMP–activated signaling pathway through Epac, a cAMP-activated guanine exchange factor that, together with PKC
, mediates the neuropathic pain response to the activation of -adrenergic receptor (Hucho et al. 2005). Additionally, the Nav1.8 current density is reduced by half in IB4+, but not IB4–, DRG neurons from contactin-null mice (Rush et al. 2005). Importantly, we recently showed that prevention of calmodulin binding to the C-terminus of Nav1.8 increases the frequency-dependent inhibition of Nav1.8 (Choi et al. 2006). Although calmodulin, a major calcium-binding protein, is ubiquitously present in all cell types, it is possible that its ability to bind to Nav1.8 is impaired in IB4+ neurons by unfavorable local structure of the Nav1.8 C-terminus, by binding of another channel partner, or by posttranslational modification (such as phosphorylation) of the channel.
We show in this study that the resting membrane potential of IB4+ neurons is more hyperpolarized than that of IB4– neurons. This finding is surprising, given that Nav1.9, a channel that is predominantly expressed in IB4+ neurons (Cummins et al. 2000; Fjell et al. 1999), produces a persistent inward current at rest that can acutely shift resting potential in a depolarizing direction (Baker et al. 2003; Herzog et al. 2001). Our data in this regard are similar to results recently reported by Fang et al. (2006). One possible explanation is that, without added GTP, the Nav1.9 persistent current is too low to effect membrane potential (Baker et al. 2003). Another contributing factor might be the dependency of the Na/K-ATPase electrogenic pump, which is known to be present within DRG neurons (Dobretsov et al. 1999; Hamada et al. 2003; Mata et al. 1991), on persistent Na+ influx that maintains intracellular Na+ at levels required for Na/K-ATPase activity. Consistent with this mechanism, in several neural systems TTX produces a hyperpolarizing shift in resting potential as the depolarizing effect of the resting sodium conductance is attenuated, followed by a progressive depolarization as ATPase activity is abolished (Sontheimer et al. 1994; Stys et al. 1993). Also consistent with this mechanism, it was previously reported (Morisset et al. 2005) that the resting potential of small DRG neurons is shifted in a depolarizing direction in Nav1.9-null mice.
In this study we used high potassium intracellular solution excluding GTP, which is frequently added in pipette solution for current-clamp recording. Intracellular GTP at the concentration often used for current clamp (100–500 µM) upregulates Nav1.8 (Saab et al. 2003) as well as Nav1.9, which is predominantly present in IB4+ neurons (Fang et al. 2002; Fjell et al. 2000), where it produces persistent sodium current that increases excitability (Baker et al. 2003). Use of GTP in the recording pipette may explain the results of Vydyanathan et al. (2005), who reported similar resting potentials in IB4– and IB4+ neurons, even though IB4+ neurons have a higher voltage-gated potassium current density, which might be expected to have a hyperpolarizing influence on resting potential.
Although we cannot formally rule out the possibility that differential expression of sodium channels—including the presence of the TTX-R cardiac channel Nav1.5—might influence the present study, it is not likely to have had a major effect on our analysis for the following reasons. First, the Nav1.5 current was detected in a very small percentage (3%) of adult DRG neurons and, when present, the current has a significantly lower density compared with that of Nav1.8 (Renganathan et al. 2002). Second, and most important, the cells in this study were held at –70 and –60 mV before applying a depolarizing stimulus in voltage- and current-clamp experiments, respectively, and most of the Nav1.5 channels would have been inactivated under these conditions (V1/2 of steady-state inactivation = –83 mV) (Renganathan et al. 2002). Thus the differential use-dependent reduction of sodium current in IB4+ and IB4– neurons that we report in this study is likely to be regulated by factors that affect the properties of Nav1.8 channel rather than the differential expression of different sodium channels in these neurons.
It is now clear that Nav1.8 plays a major role in determining the firing properties of DRG neurons (Blair and Bean 2002, 2003; Renganathan et al. 2001; Rush et al. 2006). Our results show that, in addition to being distinct subpopulations of DRG neurons with different neurochemical and action potential characteristics (Stucky and Lewin 1999), and different synaptic projections (Braz et al. 2005) and peripheral termination sites (Zylka et al. 2005), IB4+ and IB4– DRG neurons display significantly different activity-dependent responses. We propose that use-dependent reduction of Nav1.8 current by slow inactivation modulates the excitability of nociceptive DRG neurons and suggest that the distinct use-dependent current reduction in IB4+ and IB4– nociceptive DRG neurons contributes to different firing patterns in these two groups of cells that may be important after inflammation or injury. The putative differential regulation of Nav1.8 by a calcium sensor, calmodulin, in these two subpopulations of DRG neurons could shape the integration of the sensory response to external stimuli including nociception.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: S. G. Waxman, Dept. of Neurology, Yale School of Medicine, 333 Cedar Street, LCI-707, New Haven, CT 06510 (E-mail: Stephen.Waxman{at}yale.edu)
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); V1/2 = –16.7 ± 0.2 mV, k = 4.1 ± 0.2 mV (n = 16) for activation, V1/2 = –31.8 ± 0.2 mV, k = 4.9 ± 0.2 mV (n = 6) for inactivation, IB4– (
); V1/2 = –15.7 ± 0.3 mV, k = 4.2 ± 0.2 mV (n = 13) for activation, V1/2 = –31.9 ± 0.1 mV, k = 4.2 ± 0.1 mV (n = 6) for inactivation. C: Nav1.8 channels in IB4+ and IB4– neurons were subjected to 60 repetitive 100-ms depolarizing pulses to –10 mV from a Vholding of –70 mV at 1 Hz. Representative traces show 1st to 5th, 10th, and 60th sweeps. D: distribution of normalized Nav1.8 current of the 60th depolarization pulse at 1 Hz in IB4+ (
