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The Journal of Neuroscience, July 26, 2006, 26(30):7984-7994; doi:10.1523/JNEUROSCI.2211-06.2006
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Cellular/Molecular
Sodium Channel  2 Subunits Regulate Tetrodotoxin-Sensitive Sodium Channels in Small Dorsal Root Ganglion Neurons and Modulate the Response to Pain
Luis F. Lopez-Santiago,1
Marie Pertin,2,3
Xavier Morisod,2,3
Chunling Chen,1
Shuangsong Hong,4
John Wiley,4
Isabelle Decosterd,2,3 * and
Lori L. Isom1 *
1Department of Pharmacology, University of Michigan, Ann Arbor, Michigan 48109-0632, 2Anesthesiology Pain Research Group, Anesthesiology Department, Lausanne University Hospital (Centre Hospitalier Universitaire Vaudois), CH-1011 Lausanne, Switzerland, 3Department of Cell Biology and Morphology, Faculty of Biology and Medicine, University of Lausanne, CH-1005 Lausanne, Switzerland, and 4Department of Internal Medicine and General Clinical Research Center, University of Michigan, Ann Arbor, Michigan 48109-0108
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Abstract
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Voltage-gated sodium channel (Nav1)  2 subunits modulate channel gating, assembly, and cell-surface expression in CNS neurons in vitro and in vivo. 2 expression increases in sensory neurons after nerve injury, and development of mechanical allodynia in the spared nerve injury model is attenuated in 2-null mice. Thus, we hypothesized that 2 modulates electrical excitability in dorsal root ganglion (DRG) neurons in vivo. We compared sodium currents (INa) in small DRG neurons from 2+/+ and 2–/– mice to determine the effects of 2 on tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) Nav1 in vivo. Small-fast DRG neurons acutely isolated from 2–/– mice showed significant decreases in TTX-S INa compared with 2+/+ neurons. This decrease included a 51% reduction in maximal sodium conductance with no detectable changes in the voltage dependence of activation or inactivation. TTX-S, but not TTX-R, INa activation and inactivation kinetics in these cells were slower in 2–/– mice compared with controls. The selective regulation of TTX-S INa was supported by reductions in transcript and protein levels of TTX-S Nav1s, particularly Nav1.7. Low-threshold mechanical sensitivity was preserved in 2–/– mice, but they were more sensitive to noxious thermal stimuli than wild type whereas their response during the late phase of the formalin test was attenuated. Our results suggest that 2 modulates TTX-S Nav1 mRNA and protein expression resulting in increased TTX-S INa and increases the rates of TTX-S Nav1 activation and inactivation in small-fast DRG neurons in vivo. TTX-R INa were not significantly modulated by 2.
Key words: sodium channel; subunit; dorsal root ganglion; tetrodotoxin sensitive; nociception; mouse
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Introduction
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Nav1s are composed of a central, pore-forming  subunit and one or two subunits that modulate channel expression levels, voltage dependence, and kinetics (Catterall, 2000 ). 1, 1A, 2, 3, and 4 subunits regulate channel gating, assembly, and cell-surface expression in vitro (Goldin, 1993; Isom et al., 1994; Isom, 2000; Yu et al., 2003; McEwen et al., 2004). 1 and 2 also participate in homophilic (Malhotra et al., 2000) and/or heterophilic (Ratcliffe et al., 2001; Kazarinova-Noyes et al., 2001; McEwen et al., 2004) cell adhesion. 1 and 2 can also interact with extracellular matrix molecules (Srinivasan et al., 1998; Xiao et al., 1999), ankyrin (Srinivasan et al., 1988, 1998; Xiao et al., 1999; Malhotra et al., 2000), and/or receptor tyrosine phosphatase (Ratcliffe et al., 2000).
The effects of 2 on Nav1 cell-surface expression have been well established in primary CNS neuronal cultures in vitro (Schmidt et al., 1985; Schmidt and Catterall, 1986) and in brains of 2–/– mice in vivo (Chen et al., 2002). In primary CNS neuronal cultures, the expression of 2 results in increased levels of Nav1 at the cell surface. The loss of 2 results in a negative shift in the voltage dependence of Nav1 inactivation as well as significant decreases in INa density in acutely dissociated hippocampal neurons of 2–/– mice (Chen et al., 2002). 3H-saxitoxin binding experiments showed that, although the total cellular level of channels is similar in 2+/+ and 2–/– neurons, there is a 42% reduction in the level of plasma membrane channels in 2–/– neurons, consistent with the observed decreases in INa density. The integral of the optic nerve compound action potential is reduced 30% in 2–/– mice compared with control, and its dependence on stimulus strength is shifted to stronger stimuli, consistent with the reduction in channel cell-surface expression. These data clearly demonstrate that 2 plays a critical role in regulating Nav1 density and functional expression in CNS neurons in vivo.
The purpose of this study was to investigate the role of 2 in sensory dorsal root ganglion (DRG) neurons. 2 expression increases in sensory neurons after nerve injury, and 2–/– mice develop less mechanical allodynia than their wild-type counterparts in the spared nerve injury (SNI) model (Pertin et al., 2005). We hypothesized that 2 may modulate electrical excitability in DRG neurons in vivo. To test this hypothesis, we investigated both tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) INa in DRG neurons isolated from 2+/+ and 2–/– mice. We report that TTX-S but not TTX-R INas are reduced  50% in "small-fast" DRG neurons isolated from 2–/– mice compared with wild-type littermates. In addition, the activation and inactivation kinetics of TTX-S INa are slowed significantly. Behavioral studies show acute thermal hypersensitivity and reduced sensitivity to the inflammatory phase of the formalin test in 2 null mice with no measurable differences in sensitivity to mechanical stimulation. We conclude that 2 plays critical roles in electrical excitability in sensory neurons. Furthermore, 2 may differentially affect subclasses of sensory neurons in the DRG.
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Materials and Methods
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Preparation of DRG neurons.
The generation of 2–/– (Scn2b null) mice was described previously (Chen et al., 2002). Animals used in the present study were bred from a congenic strain of 2+/– mice that had been backcrossed repeatedly to C57BL/6 for at least 10 generations. All animal experiments were performed in accordance with the guidelines of the University of Michigan Committee on Use and Care of Animals or the Committee on Animal Experimentation of the Canton of Vaud, Lausanne, Switzerland. DRG neurons were acutely dissociated from adult male (8–12 weeks old) 2+/+ or 2–/– littermate mice. Briefly, the mice were killed with 100% CO2 inhalation for 1–2 min in a closed chamber. The vertebral column was then removed and cut longitudinally, and the DRGs on both sides from L4 and L5 spinal cord segments were removed. These ganglia were selected because 98% of sensory fibers in the sciatic nerve have cell bodies in the L4 and L5 DRGs (Swett et al., 1991). The DRGs were placed in minimal essential medium plus glutamine (Invitrogen, Carlsbad, CA) supplemented with 16.5 mM NaHCO3, 28.2 mM glucose, and distilled water or in DMEM/F-12 (Invitrogen) supplemented with 22 mM glucose, to give a final osmolarity of 320 mOsm/l. The medium was filtered through a 0.2 µm filter flask. The DRGs were minced two to three times before being incubated in medium with enzymes in a culture dish at 37°C. The total incubation time was 50 min with enzymes added according to the following protocol: collagenase type II (3 mg/ml; Worthington, Lakewood, NJ) was present for the entire 50 min incubation; DNase type I (0.05 mg/ml; Sigma, St. Louis, MO) and trypsin type I (1 mg/ml; Sigma) were added for the last 20 and 10 min of incubation, respectively. The dish was agitated every 10 min during the incubation. The enzymatic digestion was stopped by the addition of medium with BSA (20 mg/ml) before being replaced with fresh medium (0.5 ml) lacking BSA. Cells remaining in tissue fragments were dispersed into the medium using sterile fire-polished and silicon-coated Pasteur pipettes (6–12 strokes up and down). The cellular suspension was plated on collagen-coated 35 mm culture dishes or glass coverslips and incubated at 37°C in a humidified atmosphere of 95% air plua 5% CO2 for 1 h. Finally, 2 ml of medium supplemented with 10% fetal bovine serum were added. The cells were incubated for 2–10 h before recording, with the first 2 h used to allow cells to settle and adhere to the bottom of the culture dishes or coverslips. The remaining 8 h recording period was sufficiently short enough to minimize changes in electrical properties that may occur in long-term cultures.
Voltage-clamp recording.
Voltage-clamp recordings were performed in the standard whole-cell configuration (Hamill et al., 1981), using an Axopatch 200B voltage-clamp amplifier (Molecular Devices, Union City, CA). The cell capacitance (Cm) was calculated by integrating the area under capacitive transients as described previously (Meza et al., 1994) or read directly from the amplifier. Isolated INa were recorded from single small DRG neurons (12 pF < Cm < 42 pF) at 21°C in the presence of a bath solution that contained the following (in mM): 80 NaCl, 50 choline-Cl, 30 TEA-Cl, 2 CaCl2, 0.2 CdCl2, 10 HEPES, and 5 glucose, pH 7.3 with NaOH. For some cells, the solution contained 40 mM NaCl and 90 mM choline-Cl. Fire-polished patch pipettes were generated from borosilicate glass capillaries (Warner Instruments, Hamden, CT) using a Sutter P-87 puller (Sutter Instruments, Novato, CA) and were filled with an internal solution containing the following (in mM): 70 CsCl, 30 NaCl, 30 TEA-Cl, 10 EGTA, 1 CaCl2, 2 MgCl2, 2 Na2ATP, 0.05 GTP, 10 HEPES, and 5 glucose, pH 7.3 with CsOH. Glass coverslips to which the cells were attached were removed from the incubator and placed into a small-volume recording chamber (250 µl). Alternatively, if the cells were plated directly onto a 35 mm culture dish, 1 ml of bath solution was used. All cells were subsequently examined within 10–60 min.
Currents were low-pass filtered at 5 kHz with a four-pole Bessel filter and digitally sampled at 20 or 40 kHz. Capacitive transients were canceled with the amplifier circuitry, and linear leakage currents were digitally subtracted on-line with P/4 routines (Armstrong and Bezanilla, 1977). The use of the transient cancellation feature on the amplifier provided estimates for Cm and series resistance. The Cm estimated in this way was similar to that estimated by the integrating method (see above). Patch electrodes had resistances of 0.8–2.5 M , and the series resistance was typically in the range 1–5 M. When appropriate, this was reduced by 40–60% using the compensation circuit of the amplifier. The holding potential was always –80 mV. Recordings were performed using pClamp 8 and 9 software (Molecular Devices).
To analyze the voltage dependence of channel activation, the sodium conductance (GNa) was calculated. Peak current data for each cell were divided by the respective driving force (Vm – Vrev), plotted against Vm, and fit to a Boltzmann equation of the following form:
where Gmax is the maximum GNa, V1/2 is the voltage at which 50% of the Nav1 are activated, and k is the slope of the curve. Steady-state inactivation was measured by applying a double-pulse protocol, consisting of a 500 ms prepulse ranging from –120 to 20 mV (in 5 and 10 mV increments), followed by a test pulse to 0 mV. Each data set (a plot of peak INa during the 0 mV test pulse vs prepulse voltage) was fit with the summation of two Boltzmann equations of the following form:
where F1 and F2 are the fractions of the first and second components of inactivation, respectively. The most negative component (component 1) results from the TTX-S INa whereas the other results from TTX-R INa. V1/2 is the potential at which half of the INa was inactivated, and k is the slope factor for each component. The sum of both fractions is the calculated maximum INa (F1 + F2 = Imax). Data points were then normalized with respect to Imax to obtain the inactivation curve. An alternate calculation method, yielding similar results, is included in the on-line supplemental material (available at www.jneurosci.org).
To examine the rate of channel recovery from inactivation, a protocol was designed comprising a 500 ms prepulse to –120 mV, followed by a test pulse to 0 mV, and then returning to –120 mV for a variable time period (0.25, 0.5, 1, 2, 4, 6, 8, 10, 20, 30, 40, 50, 75, 100, 200, 300, 400, 500, and 750 ms) before application of a second test pulse to 0 mV. The INa amplitude from the second 0 mV pulse was divided by the amplitude of the corresponding first test pulse to obtain the fraction of INa recovered after the recovery time. The data were fit with a double-exponential equation of the following form:
where INa p2/INa p1 is the fraction of current recovered; f1 and f2 are the fractions of the fast and slow recovery components, respectively; t is recovery time; and  1 and 2 are the time constants for each recovery component.
Analysis of electrophysiological data.
Data were analyzed using pClamp 8 and 9 (Molecular Devices) and SigmaPlot 7 (SPSS, Chicago, IL). The statistical significance of differences between mean values for 2+/+ and 2–/– neurons was evaluated by Student’s unpaired t test, with p < 0.050 considered significant. Results are presented as means ± SEM.
Real-time reverse transcription-PCR.
Unilateral L3–L5 DRGs from 2+/+ and 2–/– mice were rapidly dissected and collected in RNA-later solution (Qiagen AG, Basel, Switzerland). Each individual sample consisted of a pool of six DRGs dissected from two animals. Total RNA was isolated from each individual sample using the RNeasy Mini kit (Qiagen AG) with a DNase step (RNase free DNase set; Qiagen AG) on the column. After RNA quality and quantity were assessed by electrophoresis and spectrophotometry, 1.5 µg of RNA for each sample was reverse transcribed using Omniscript reverse transcriptase following the manufacturer’s instructions (Qiagen AG). Beacon Designer 3.0 software (Primer Biosoft International, Palo Alto, CA) was used to design primer and probe sequences (Table 1) according to SYBR green specifications (Vandesompele et al., 2002). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was chosen as an endogenous control to normalize expression levels of the different Nav1 and subunits. Real-time PCRs were performed in a 20 µl total volume containing 50 ng of cDNA, 300 nM of each primer, 10 µl of 2x iQ SYBR green mix containing nucleotides, iTaq DNA polymerase, SYBR green, and fluorescein (Bio-Rad, Reinach, Switzerland) using the MyiQ Single Color Real-Time PCR Detection System (Bio-Rad). The amplification protocol was as follows: 3 min at 95°C, 45 cycles of 10 s at 95°C for denaturation, and 45 s at 60°C for annealing and extension; specificity was assessed using a DNA melting curve by measuring fluorescence during gradual temperature increments (0.5°C) from 55 to 95°C.
To determine the profile of expression of different Nav1 subunits (Nav1.1, Nav1.2, Nav1.3, Nav1.6, Nav1.7, Nav1.8, and Nav1.9 and 1, 2, 3, and 4) in 2–/– and 2+/+ mice, a pool of three samples of each cDNA was used. PCR was performed in triplicate.
The level of expression of Nav1 and subunits detected in 2–/– and 2+/+ mice was determined using four individual samples of total RNA from each genotype; each amplification was performed in triplicate for each target mRNA. The efficiency of amplification was determined by serial dilution of starting DNA, and standard curves were constructed from the respective mean critical threshold (CT) value for GAPDH and Nav1 transcripts. The relative expression of each target gene was calculated based on real-time PCR efficiencies and the threshold value of the unknown sample versus the standard sample (Pfaffl, 2001).
Western blot analysis of DRG membrane preparations.
DRGs were removed and stored in ice-cold dissection medium. After a brief centrifugation at 4°C, the supernatant was discarded and the DRGs were rinsed two times in Tris-EGTA buffer (50 mM Tris, pH 8.0 with NaOH, and 10 mM EGTA) containing Complete protease inhibitors (Roche, Indianapolis, IN) at twice the recommended concentration. The DRGs were homogenized and centrifuged at 3000 x g for 5 min at 4°C to remove nuclei. The supernatant was ultra-centrifuged at 4°C for 10 min at 195,000 x g, and the pellets resuspended in 60 µl of Tris-EGTA buffer containing Complete protease inhibitors. One microliter of the suspension was used for protein assay. Equal amounts of protein (100 µg) were loaded on 4–15% polyacrylamide gradient gels and separated by SDS-PAGE. Proteins were then transferred to nitrocellulose as described previously (McEwen et al., 2004). Blots were probed with specific Nav1 subunit antibodies. Anti-Nav1.1 (Chemicon, Temecula, CA) was used at a 1:500 dilution, anti-Nav1.7 (Chemicon) was used at a 1:500 dilution, and anti-Nav1.6 (Neuromab) was used at a 1:100 dilution. All blots were subsequently probed with anti--tubulin diluted 1:5000 (Cedarlane Laboratories, Hornby, Ontario, Canada). The immune signal from this housekeeping protein was used to control for loading differences. Densitometric measurement of the immunoreactive Nav1 bands was performed using Scion (Frederick, MD) Image. Each Nav1 band was first normalized to the corresponding -tubulin band for each lane on the gel. Nav1 expression levels in the 2–/– lanes were then calculated as a percentage of the corresponding wild-type levels, using the -tubulin normalized 2+/+ Nav1 levels as 100%.
Behavioral tests.
Eight-week-old male 2–/– or 2+/+ mice were habituated to the environment, the tester, and the apparatus for at least 2 weeks before testing. All behavioral testing was performed by an observer blinded to the mouse genotype.
The hot-plate assay was conducted by placing the animals (n = 9 in each group) on the hot-plate surface set at varying temperatures (49, 52, and 55°C) (Cao et al., 1998). The latency of response was determined by a clear hindpaw lick. The cutoff was adjusted for each temperature to avoid tissue damage: 60 s for 49°C, 30 s for 52°C, and 20 s for 55°C.
The tail-flick assay was conducted using a tail-flick analgesia meter (Columbus Instrument, Columbus, OH), and the mice were gently restrained in a conic plastic cloth. The latency of response was recorded manually at two different light-beam intensities (4 and 7; n = 4 in each group) with a cutoff at 20 s (Wilson and Mogil, 2001).
Mechanical sensitivity assessment was performed by applying an ascending series of non-noxious Von Frey monofilaments (Stoelting, Wood Dale, IL) to the plantar surface of each hindpaw (n = 9 in each group). For this purpose, mice were placed on an elevated platform with a delicate wire netting floor. The withdrawal threshold (in grams) was defined as the lowest force that evoked a brisk withdrawal response to at least 2 of 10 stimuli (Suter et al., 2003).
For the formalin test, 10 µl of 5% formalin (formaldehyde; Sigma, St. Louis, MO) was injected subcutaneously in the left hindpaw (n = 6 in each group). The time the animal spent shaking/flinching and licking the injected paw was recorded for each 5 min interval, from the injection time up to 80 min (Wei et al., 2001).
Analysis of behavioral data.
Data are represented as mean ± SEM. Differences between groups were compared using one- or two-way ANOVA for unpaired variables, followed by post hoc Bonferroni’s correction when appropriate. Von Frey series present logarithmic differences between hairs, and logarithmic-transformed values were used for the analysis, enabling ANOVA tests (Suter et al., 2003). Statistical analyses were performed using JMP statistical software (version 5.01; SAS Institute, Cary, NC). A p value  0.05 was considered statistically significant.
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Results
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Identification and definition of neuronal size
Using criteria described in previous studies (Abdulla and Smith, 2001, 2002), DRG neurons were first assigned to "small," "medium," or "large" groups on the basis of their Cm: <42, 42–72, and >72 pF, respectively (2+/+ cells, n = 89; 2–/– cells, n = 85). There were no differences in the proportions of small, medium, and large cells between the two strains (data not shown). We focused this study on small neurons, with a mean Cm of 24.9 ± 1.7 pF (n = 35) for 2+/+ neurons and 23.9 ± 1.4 pF (n = 32) for 2–/– neurons. Assuming a specific Cm of 1 µF/cm2 (Hille, 2001), and that the cells have a spherical shape without invaginations, cell-surface area and diameter can be estimated. Thus, the mean Cm corresponds to cell diameters of 28.1 and 27.6 µm, respectively, for 2+/+ and 2–/– neurons. Neurons with diameters <30 µm were considered nociceptive neurons, or type C cells (Study and Kral, 1996; Flake et al., 2004).
Total INa, the sum of TTX-S and TTX-R INa (Roy and Narahashi, 1992; Rush et al., 1998; Abdulla and Smith, 2002; Dib-Hajj et al., 2002), was recorded using a series of depolarizing voltage commands from a holding potential of –80 mV. To explore the voltage dependence of INa in DRG neurons of 2+/+ and 2–/– mice, we took advantage of the previously described activation and inactivation properties of peripheral nerve Nav1 (Cummins and Waxman, 1997; Akopian et al., 1996, 1999). Thus, a current–voltage (I–V) protocol with a 500 ms prepulse to –120 or –50 mV, followed by a test pulse from –100 to +40 mV was applied, with steps of 5 and 10 mV, waiting 10 s between each step (Fig. 1A, inset). When the I–V protocol with a prepulse to –120 mV was applied, the total INa was obtained (Fig. 1A, top traces). A second I–V protocol was subsequently applied to the same cell, but with a prepulse to –50 mV, to inactivate TTX-S INa and thus record only the TTX-R component (Fig. 1A, middle traces). Finally, the TTX-S component was obtained by digitally subtracting the data obtained with the second protocol from the first (Fig. 1A, bottom traces). Similar results were obtained using 300 nM TTX on some cells (data not shown). This protocol has the advantage of obtaining separate I–V relationships for both TTX-S and TTX-R INa in the absence of TTX, thus saving time and reducing cell deterioration. The two I–V curves allow us to classify the small neurons into two subgroups: small-fast and "small-slow" DRG neurons. When the maximum amplitude of TTX-S INa was >70% of the total INa, the cells were placed in the small-fast subgroup (Abdulla and Smith, 2002). These cells made up 49% (17 of 35) of the 2+/+ small cell population and 53% (17 of 32) of the 2–/– small neurons. Cells placed in the second subgroup had TTX-R INa >70% of the total INa. These cells made up 46% (16 of 35) of the 2+/+ and 41% (13 of 32) of the 2–/– small cells. INa in the 6% of cells remaining from the total cell population, two cells of each genotype, did not clearly fall into either category. Thus, these cells were not included in our analysis.
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Figure 1. Current–voltage relationships. A, Protocol for separation of TTX-R and TTX-S INa. A 500 ms prepulse to –120 or –50 mV was applied before a 50 ms test pulse from –100 to 40 mV with steps of 5 or 10 mV (inset). Currents evoked from one 2–/– small-fast DRG neuron by test pulses from –50 to 0 mV are shown. Both TTX-S and TTX-R INa were apparent after the –120 mV prepulse (top traces); only TTX-R INa were obtained after the –50 mV prepulse (middle traces), and the TTX-S component was obtained (bottom traces) by digitally subtracting the TTX-R INa from the total INa B, Average peak INa density–voltage relationships for TTX-R INa (circles) and TTX-S INa (squares) of small-fast DRG neurons (means ± SEM), 2+/+ (closed symbols; n = 15), or 2–/– (open symbols; n = 14). Smooth lines are I–V curves generated using the Boltzmann fit parameters of the respective activation curves. Inset, I–V curves of total INa from the same cells as in A, 2+/+ (closed symbols) and 2–/– (open symbols). C, Similar to B, but for small-slow neurons, 2+/+ (closed symbols; n = 14), or 2–/– (open symbols; n = 11).
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The absence of 2 results in reduced TTX-S INa
Plots of peak INa density versus command voltage for small-fast and small-slow DRG neurons are shown in Figure 1, B and C, respectively. For both 2+/+ (n = 15) and 2–/– (n = 14) small-fast DRG neurons, the main INa was TTX-S, as expected by definition. The activation threshold for this INa was between –55 and –50 mV, and the maximum inward INa fell between –30 and –20 mV. For both 2+/+ (n = 14) and 2–/– (n = 11) small-slow neurons, the main INa was TTX resistant, detectable between –40 and –30 mV with maximal inward INa at approximately –10 mV. All currents measured displayed a reversal potential (Vrev) of 25 mV, corresponding to the calculated equilibrium potential for sodium ions under these recording conditions (ENa = 25 mV). The I–V curves for small-fast neurons from 2–/– mice show a significant reduction in TTX-S INa density compared with those neurons isolated from 2+/+ mice (Fig. 1B). This reduction can also be clearly observed directly in the total INa (Fig. 1B, inset).
To better compare the voltage dependence of channel activation, the sodium conductance (GNa) was calculated as described in Materials and Methods. For 2+/+ and 2–/– small-slow DRGs neurons (Fig. 2C,D), V1/2 and k were similar for both TTX-R and TTX-S INa; only Gmax for TTX-S INa showed a reduction of 29% in 2–/– neurons compared with 2+/+ neurons, however this reduction was not significant (p = 0.665). For small-fast cells, the mean value of V1/2 and k were also similar for TTX-R and TTX-S INa between groups (Fig. 2B). The Gmax for TTX-R INa was 30% smaller in 2–/– cells compared with 2+/+ (Fig. 2A, dashed lines), but, again, this difference was not significant (p = 0.083). In contrast, the Gmax for TTX-S INa measured in 2–/– neurons was 51% smaller than that observed in 2+/+ cells (Fig. 2A, solid lines), and this reduction was statistically significant (p = 0.001). These results suggest that 2 subunits regulate cell-surface levels of TTX-S but not TTX-R Nav1 in DRG neurons. Alternatively, it is possible that 2 alters TTX-S Nav1 single-channel conductance; however, based on our previous results (Isom et al., 1995b; Chen et al., 2002), we propose that the former is the most likely mechanism of 2 action.
2 does not affect the voltage dependence of INa inactivation
Steady-state inactivation was measured as described in Materials and Methods. An example of the INa obtained from a typical small-fast 2+/+ neuron in response to a test pulse to 0 mV is shown in the inset to Figure 3A. In this example, as in practically all small-fast DRG neurons tested, the fast INa (TTX-S) is inactivated at more negative voltages than the slow INa (TTX-R). The mean of the individual curves are shown in Figure 3A for small-fast neurons and in Figure 3C for small-slow neurons. The corresponding V1/2 and k are compared in Figure 3, B and D. The voltage dependence of TTX-S and TTX-R INa inactivation in small 2–/– neurons was nearly identical to that of TTX-S and TTX-R INa of small 2+/+ neurons (Fig. 3), therefore the 2 subunit does not regulate the INa voltage dependence in small DRG neurons.
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Figure 3. Voltage dependence of inactivation. A, Peak INa at 0 mV, normalized to its maximal value (inset), as a function of voltage during a 500 ms prepulse. INa were measured from 2+/+ (n = 13; closed circles) and 2–/– (n = 10; open circles) small-fast DRG neurons. Each data set was fit with a double Boltzmann function (lines). The inset is an example of INa at 0 mV, from one small-fast 2+/+ cell, after 500 ms prepulses from –90 to –10 mV. B, Parameters of fitted inactivation curves shown in A; circles represent the first component (TTX-S), and squares represent the second component (TTX-R). C, D, Inactivation curves and parameters, respectively, for small-slow 2+/+ (n = 12) and 2–/– (n = 8) neurons; symbols are as in A and B. Error bars indicate SEM.
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According to our classification of cells into small-fast and small-slow based on the proportion of TTX-S INa to that of TTX-R INa, the average values of F1 and F2 (the proportion of TTX-S and TTX-R INa) are close to 0.7 and 0.3, respectively, for both 2+/+ and 2–/– small-fast cells. For small-slow cells, F1 and F2 are 0.1 and 0.9, respectively. Thus, the main INa for small-fast cells is TTX-S, whereas for small-slow cells it is TTX-R.
Effects of 2 on INa kinetics
The rate of channel recovery from inactivation was measured as described in Materials and Methods. A typical set of INa traces obtained using this protocol is shown in the inset of Figure 4. In agreement with previous reports, the recovery from inactivation curve shows two components: the TTX-R INa shows fast recovery, and the TTX-S INa shows slow recovery (Cummins and Waxman, 1997; Rush et al., 1998). Time constants for 2+/+ were 1 = 1.3 ± 0.2 ms and 2 = 95.3 ± 17.8 ms (n = 6), and time constants for 2–/– were 1 = 1.5 ± 0.5 ms and 2 = 83.3 ± 29.4 ms (n = 6); the fraction of TTX-S INa was 0.88 ± 0.03 and 0.80 ± 0.12 for 2+/+ and 2–/–, respectively. There were no significant differences between groups.
Superimposition of INa obtained from small-fast 2+/+ and 2–/– neurons suggested different rates of channel activation and inactivation (Fig. 5A). To evaluate the activation kinetics, two points on each time course were measured: the time to achieve 50% of the maximum current after onset of the test pulse to 0 mV (T1/2 peak), and the time to achieve the peak INa with the same test pulse (Tpeak). Comparing both groups of small-fast neurons, these times were significantly different (Fig. 5B) with T1/2peak and Tpeak for 2–/– (n = 12), 60 and 69% longer than 2+/+ (n = 16) neurons, respectively. To evaluate the inactivation kinetics for the same INa, the decaying phase of the INa was fit using a double-exponential function, obtaining two inactivation time constants (fast and slow). Only fast was affected by the loss of 2 expression (Fig. 5C): fast = 1.25 ± 0.17 ms for 2+/+ neurons compared with 2.13 ± 0.23 ms for 2–/– neurons, an increase of 70%. In contrast, slow was the same for both groups (Fig. 5C). We performed similar analyses for small-slow 2+/+ (n = 11) and 2–/– (n = 9) neurons. Neither activation nor inactivation kinetics were statistically different (data not shown).
Nav1 mRNA levels are regulated by 2
To begin to investigate the molecular basis for the observed reduction in TTX-S INa in 2–/– neurons, we measured Nav1 and subunit mRNA levels in DRGs from both wild-type and null mice. The profile of expression and relative abundance of Nav1 subunit transcripts (Nav1.1, Nav1.2, Nav1.3, Nav1.6, Nav1.7, Nav1.8, and Nav1.9 and 1, 2, 3, and 4) (Table 1) were evaluated independently in 2+/+ and 2–/– mice by real-time reverse transcription (RT)-PCR. Nav1 subunit mRNA levels were normalized to the highest Nav1 subunit transcript expressed in 2+/+ DRGs: Nav1.7 for subunits and 1 for subunits (Fig. 6). In both genotypes, the order of the Nav1 subunit transcript expression levels was the same, with the exception of 2 in the null mice: Nav1.7 > Nav1.8 > Nav1.9 > Nav1.6 > Nav1.1 > Nav1.3 > Nav1.2 and 1 > 4 > 3 > 2. In 2+/+ DRGs, the Nav1.7 transcript was the most abundant. It was twice as abundant as Nav1.6, 7 times more abundant than Nav1.3 or Nav1.1, and 25 times more abundant than Nav1.2, all TTX-S channels. In addition, Nav1.7 was more abundant than transcripts for the TTX-R channels Nav1.8 and Nav1.9 (Fig. 6A). For mRNA of subunits, 1 was 6–10 times more abundant than the other subunits (Fig. 6B). Interestingly, 2 appeared to be the lowest expressed subunit mRNA; however, as shown above, it plays a major functional role in DRG neurons.
In 2–/– DRGs, the profile of and subunit expression did not change, with the exception of 2. However, the abundance of some subunit transcripts did change, notably Nav1.7 (p = 0.043) (Fig. 6). Experiments were then performed to determine the ratios of specific subunit mRNA expression levels in null versus wild-type neurons. 2+/+ and 2–/– DRGs were assessed by real-time RT-PCR and normalized to GAPDH. The ratios of Nav1 and subunit transcript levels in 2–/–/2+/+ DRGs are shown in Figure 7. The expression levels of three TTX-S Nav1 transcripts, Nav1.3, Nav1.6 and Nav1.7, were significantly reduced (p < 0.05) 20–25% by the 2 null mutation (Fig. 7A); similar reductions were found for 3 and 4 mRNAs (Fig. 7B), whereas no changes were observed for the other subunits.
TTX-S Nav1 protein is regulated by 2
Western blot analyses of membrane preparations of DRG neurons for the TTX-S subunits Nav1.7, Nav1.6, and Nav1.1, all normalized to -tubulin as a housekeeping protein loading control, were performed to determine whether the observed changes in mRNA expression were reflected at the level of protein. Changes in mRNA levels are not always reflected by altered protein expression, with Nav-2 subunits serving as an example of this phenomenon (Malhotra et al., 2001; Pertin et al., 2005). We observed that Nav1.1 and Nav1.7 protein levels were reduced in 2–/– DRG neurons compared with 2+/+, whereas the levels of Nav1.6 did not appear to change (Fig. 8A). Similar results to those shown in the figure were obtained from three or four independent experiments. Immunoreactive bands were normalized to -tubulin by densitometry, and changes in 2–/– levels were expressed as a percentage of wild-type levels for each Nav1 (Fig. 8B). These calculations showed that Nav1.1 (p = 0.082; n = 3) and Nav1.7 (p < 0.001; n = 3) were reduced in the null DRGs compared with wild type. The values for Nav1.6 were not significantly different (p = 0.23; n = 4). We propose that the observed reduction in mRNA and protein levels of Nav1.7 (and possibly the reduction in Nav1.1 protein expression) may underlie the reduction in TTX-S INa measured in 2–/– neurons.
2–/– mice show increased thermal but not mechanical sensitivity
Because TTX-S Nav1s are involved in nociceptive transmission, we hypothesized that 2–/– mice would exhibit altered responses to noxious stimuli. We showed previously that 2 expression is upregulated in sensory neurons in neuropathic pain and that development of mechanical allodynia in the SNI model is attenuated in 2–/– mice compared with wild type (Pertin et al., 2005). To determine whether 2 also plays a role in acute pain pathways, we compared the responses of the 2–/– mice to acute thermal and mechanical stimuli with those of their wild-type littermates. In the hot-plate test at 49°C (Fig. 9A), 2+/+ mice exhibited a latency of response of 34.9 ± 3.0 s. In contrast, 2–/– mice displayed a significantly shorter response latency of 27.4 ± 1.5 s (p < 0.05; n = 9 in each group). At higher temperatures, no difference was observed between groups (p  0.05). To confirm the hot-plate test results and to determine whether thermal hypersensitivity in 2–/– mice was induced by spinal processes, we evaluated the simple tail-flick reflex response to a radiant heat beam focused on the tail. At a low-intensity setting (4) of the tail-flick analgesia meter (Fig. 9B), the latency was significantly shorter in 2–/– mice (5.5 ± 1.1 s) compared with 2+/+ mice (9.1 ± 1.2 s) (p < 0.01; n = 4 in each group). At a higher intensity (7), a difference between the two groups was not discernible (p 0.05) (Fig. 9B). To investigate whether compensatory upregulation of heat-sensing genes occurred in the 2–/– mice and thus could be responsible for the observed thermal hypersensitivity, we measured the levels of TRPV1 and TRPV2 in DRGs isolated from 2+/+ and 2–/– mice. No differences were detected between the two genotypes (TRPV1: ratio 2–/–/2+/+ was 0.95, p = 0.57; TRPV2: ratio 2–/–/2+/+ was 0.99, p = 0.95).
Mechanical withdrawal threshold responses to a series of calibrated monofilaments applied to both paws in both groups were also recorded. Deletion of 2 did not modify the animals’ response, and we did not observe any significant differences between groups (p 0.05; n = 9 in each group) (Fig. 10). For 2+/+ mice, the values were 0.246 ± 0.05 g (left hindpaw) and 0.249 ± 0.07 g (right hindpaw) compared with 0.235 ± 0.05 g (left hindpaw) and 0.283 ± 0.08 g (right hindpaw) for 2–/– mice.
2–/– mice show reduced response to inflammatory pain
We next performed an extended formalin test to determine the role of 2 in this model of acute and inflammatory pain (Tjolsen et al., 1992; Wei et al., 2001). During the initial phase of acute pain, the responses of 2–/– and 2+/+ mice were similar (Fig. 11). After the initial phase, an early second phase from 10 to 55 min and a later second phase from 55 to 80 min have been described previously (Wei et al., 2001). These late phases are considered to be models of inflammatory pain (Tjolsen et al., 1992). During the late phase, the behavioral response of 2–/– mice was significantly attenuated when compared with 2+/+ mice (Fig. 11).
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Discussion
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Nav1s in sensory neurons control membrane excitability and contribute to the transmission of nociceptive information to the spinal cord. Both TTX-S and TTX-R Nav1s are expressed in DRG neurons (Baker and Wood, 2001), as are 1, 1A, 2, 3, and 4 (Black et al., 1996; Kazen-Gillespie et al., 2000; Yu et al., 2003). The subunit cDNAs express functional Nav1 in heterologous expression systems. However, for TTX-S subunits, the currents characteristic of these channels expressed in isolation are quite different from native currents. Coexpression of the subunits with these channels results in shifts in the voltage dependence of activation and inactivation, changes in channel modal gating behavior resulting in increases in the rates of inactivation and recovery from inactivation (Isom et al., 1994), and increases in channel expression at the plasma membrane as assessed by 3H-saxitoxin binding (Isom et al., 1995a; Kazarinova-Noyes et al., 2001; McEwen et al., 2004). The subunit-mediated effects on TTX-R Nav1 expressed in heterologous systems are not well understood (Sangameswaran et al., 1996; Malhotra et al., 2001; Vijayaragavan et al., 2001, 2004), and the functional effects of Nav subunits on are dependent on the recipient cell type in vitro (Chen et al., 2002; Meadows and Isom, 2005). Thus, the use of in vivo models is critical to the understanding of their physiological roles. The present studies using 2 null mice make important and novel contributions to elucidating the function of 2 in electrical signal transduction in sensory neurons and are the first report of the differential effects of 2 on TTX-S versus TTX-R channels.
In the present study, we compared INa in DRG neurons isolated from 2+/+ and 2–/– mice to determine the effects of 2 on TTX-S and TTX-R Nav1s in vivo. Small-fast DRG neurons acutely isolated from 2–/– mice showed significant decreases in TTX-S but not TTX-R INa compared with DRG neurons isolated from wild-type littermates. This decrease was not a result of changes in the voltage dependence of activation or inactivation of TTX-S Nav1s. TTX-S, but not TTX-R, INa activation and inactivation kinetics in small-fast DRG neurons were significantly slower in 2–/– mice compared with 2+/+. Our results predict that 2 expression results in increased levels of TTX-S Nav1 mRNA and protein expression, particularly Nav1.7, increased levels of TTX-S Nav1 cell-surface expression, and increased rates of TTX-S INa activation and inactivation in small-fast DRG neurons in vivo. TTX-R INa in small-slow and small-fast DRG neurons are insensitive to modulation by 2.
Interestingly, the 2 null mutation affects TTX-S INa in small-slow versus small-fast DRG neurons differently. TTX-S INa are dramatically reduced in small-fast neurons but are essentially unaffected in small-slow neurons. A possible explanation for this observation is that small-slow and small-fast DRG neurons may express vastly different levels of 2 protein. Alternatively, perhaps these two neuronal populations express different profiles of TTX-S Nav1s that are differentially regulated by 2 in vivo. Previously, we reported that 2 protein expression is low in normal DRG neurons, although immunocytochemical staining could be detected in small, medium, and large cells (Pertin et al., 2005). Unfortunately, this low level of 2 staining precluded our ability to perform a more complete analysis of differential expression in small neurons in the present study. Because 2 mRNA levels are poor predictors of 2 protein levels (Malhotra et al., 2001; Pertin et al., 2005 |
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Abstract
Introduction
