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NEUROSCIENCE |
1 Department of Pharmacology and Toxicology, Stark Neurosciences Research Institute, Indiana University School of Medicine, Indianapolis, IN 46202, USA
2 Department of Neurology
3 Center for Neuroscience and Regeneration Research, Yale University School of Medicine, New Haven, CT 06510, USA
4 Rehabilitation Research Center, Veterans Administration Connecticut Healthcare System, West Haven, CT 06516, USA
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Abstract |
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(Received 21 December 2006;
accepted after revision 2 April 2007;
first published online 12 April 2007)
Corresponding author T. R. Cummins: Department of Pharmacology and Toxicology, Stark Neurosciences Research Institute, Indiana University School of Medicine, 950 West Walnut St, R2 468, Indianapolis, IN 46202, USA. Email: trcummin{at}iupui.edu.
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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One hereditary erythromelalgia mutation that has been associated with a very severe phenotype is the N395K Nav1.7 mutation (Drenth et al. 2005), located in the S6 segment of domain I. The residue corresponding to N395 in other voltage-gated sodium channels is involved in the interaction with local anaesthetics. Lysine substitution at the corresponding residue (N434K) in the rat Nav1.4 sodium channel reduces local anaesthetic block of Nav1.4 (Nau et al. 1999). Therefore, we predicted that the N395K mutation in Nav1.7 alters interaction of local anaesthetics with the channel. Unfortunately both of the affected individuals in the original family with the N395K mutation (Drenth et al. 2005) are now deceased and data on sensitivity of these patients to local anaesthetics is not available. Interestingly, one study has shown that treatment of erythromelalgia with lidocaine relieves pain in only 55% of afflicted patients (Davis & Sandroni, 2005). Due to the multiple mutations involved in hereditary erythromelalgia, it is possible that some but not all of these mutations alter local anaesthetic binding to Nav1.7, and this may contribute to the variability in patients responding to lidocaine. Therefore, the goals of this study were to (1) examine electrophysiological changes in Nav1.7 produced by the N395K mutation and how they affect the excitability of a model sensory neuron and (2) determine if the N395K mutation in Nav1.7 changes the effects of lidocaine on sodium channel current.
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Methods |
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The human Nav1.7 (wild-type, N395K, or F216S) channels were cotransfected with the human 
1 and 2 subunits into human embryonic kidney (HEK293) cells using the calcium phosphate precipitation technique. The N395K and F216S mutations were produced using the QuickChange XL Site-Directed Mutagenesis Kit (Stratagene; La Jolla, CA, USA). HEK293 cells were grown under standard tissue culture conditions (5% CO2; 37°C) in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum. The calcium phosphate–DNA mixture was added to the cell culture medium and left for 3 h, after which the cells were washed with fresh medium. Sodium currents were recorded 40–72 h after transfection.
Chemicals and solutions
Lidocaine hydrochloride (Sigma Aldrich, St Louis, MO, USA) was dissolved in standard bathing solution to give a stock solution of 100 mM. Subsequent dilutions were performed in standard bathing solution to give concentrations of (mM): 0.1, 0.3, 1, 3, 10 and 30. Lidocaine solutions were made fresh before patch-clamp recordings.
Whole-cell patch-clamp recordings
Whole-cell patch-clamp recordings were conducted at room temperature (
21°C) using a HEKA EPC-10 amplifier. Data were acquired on a Windows-based Pentium IV computer using the Pulse program (v 8.65, HEKA Electronic, Germany). Fire-polished electrodes (0.9–1.3 M
) were fabricated from 1.7 mm capillary glass using a Sutter P-97 puller (Novato, CA, USA). The standard pipette solution contained (mM): 140 CsF, 10 NaCl, 1.1 EGTA and 10 Hepes, pH 7.3. The standard bathing solution contained (mM): 140 NaCl, 1 MgCl2, 3 KCl, 1 CaCl2 and 10 Hepes, pH 7.3 (adjusted with NaOH). Cells on glass coverslips were transferred to a recording chamber containing 250 µl of standard bathing solution. Protocols were performed 4 min after whole-cell configuration had been established to allow for adequate equilibration of the intracellular solution. Lidocaine solutions were added to the bath compartment by first withdrawing 25 µl of bathing solution, then adding 25 µl of 10-fold concentrated lidocaine and mixing 10–15 times with a 100 µl pipettor. Final concentrations of lidocaine in the bath compartment were in the range of 10 µM to 10 mM. Series resistance errors were compensated to be less than 3 mV during voltage-clamp recordings.
Data analysis
Voltage clamp experimental data were analysed using the Pulsefit (v 8.65, HEKA Electronic), Origin (OriginLab Corp., Northhampton, MA, USA), and Microsoft Excel software programs. Slope factors of conductance–voltage and steady-state inactivation curves were calculated using the general Boltzmann function:
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Computer simulations
The electrical properties of small sensory neurons were simulated using the NEURON program (version 5.9) (Hines & Carnevale, 1997). Sodium and potassium conductances were modelled as described below using Hodgkin & Huxley-type (HH) descriptions (Hodgkin & Huxley, 1952) of the various voltage-dependent currents. Action potential firing was simulated using a single compartment cylindrical model of length 37.75 µm and radius 20 µm, simulating a small sensory neuron with a 3000 µm2 surface area and 24.3 pF capacitance. The integration method was Backward Euler at an integration time step dt of 0.025 ms.
Simulations were performed assuming free ionic concentrations of sodium ([Na+]o
= 145 mM; [Na+]i
= 12 mM) and potassium ([K+]o
= 8 mM; [K+]i
= 118 mM) which were used to calculate their Nernst reversal potential of +63.4 mV (ENa) and –68.5 mV (EK), respectively. By analogy to the HH model of action potential electrogenesis, the linear leakage current was defined as ILeak
=
gLeak(V
–
ELeak), where gLeak is the leak conductance, V is the membrane potential, and ELeak is the reversal potential for the leak current. ELeak was set at –55 mV. The size of the current was adjusted so that its amplitude corresponded to an input resistance of 706 M: gLeak
= 0.0000425 S cm–2, similar to that reported for small sensory neurons (Wang et al. 2005).
The DRG neuron model included a leak conductance, two potassium conductances, Nav1.7 conductance(s) (wild-type and mutant) and Nav1.8, slowly inactivating TTX-R conductance. The predominant potassium conductances in small DRG neurons are a sustained (delayed rectifier type) conductance and a transient (A-type) conductance (Gold et al. 1996). Therefore we introduced into the model a delayed rectifier potassium current (IKDR) and an A-type potassium current (IKA). The majority of small (< 25 µm diameter) DRG neurons exhibit both TTX-S and -R currents (Cummins & Waxman, 1997). Although many small DRG neurons express the mRNA for more than one TTX-S sodium channel isoform (Black et al. 1996), the major TTX-S current in the majority of small DRG neurons closely resembles the current produced by Nav1.7 channels (Cummins et al. 1998). Therefore the only TTX-S conductances included in our model simulate Nav1.7 currents. Both slowly inactivating (Nav1.8-like) and persistent (Nav1.9-like) TTX-R currents can be recorded from many small DRG neurons (Cummins et al. 1999). The persistent TTX-R current is subject to significant modulation and tends to be negligible when the cells are held at
–60 mV (Cummins et al. 1999) and therefore we only included the Nav1.8-like TTX-R current in our model neuron. For specific details of the current models see online Supplemental material.
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Results |
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Wild-type (WT) hNav1.7 or the mutant derivative channel N395K were transiently expressed with h-1 and h-2 subunits in HEK293 cells. An initial comparison of current traces from both channels did not suggest any major differences in channel kinetics (Fig. 1A and B). The voltage dependence of activation was measured for each channel using a series of depolarizing test pulses from –100 to +70 mV. N395K channels exhibited a hyperpolarized current–voltage (I–V) and conductance–voltage (G–V) curve compared to wild-type channels (Fig. 1C and D). The midpoint of activation (estimated using a Boltzmann function) was significantly (P < 0.001) more negative (–28.0 ± 1.1 mV; slope: 6.7 ± 0.1; n
= 18) for N395K channels than for WT channels (–20.3 ± 0.8 mV; slope:7.27 ± 0.1; n
= 19; Student's unpaired t test). The voltage dependence of steady-state fast inactivation was also evaluated for each channel and, in contrast to activation, did not differ between the WT and mutant channels (Fig. 2A). The midpoint of steady-state fast inactivation for the WT channels (–71.3 ± 0.8 mV; slope: 6.79 ± 0.2; n
= 19) was not significantly different (P
= 0.76) from that of the N395K channels (–70.9 ± 1.1 mV; slope: 6.14 ± 0.1; n
= 18; Student's unpaired t test).
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The kinetics of deactivation for WT and N395K channels was also examined by eliciting tail currents at a range of potentials after brief activation of the channels (–20 mV for 0.5 ms). Deactivation reflects the transition of the channel from the open to the closed state, and defects in skeletal muscle sodium channel deactivation have been implicated in paramyotonia congenita (Featherstone et al. 1998). The time constant of deactivation (measured with single exponential fits) was slower for the N395K channel than that of the WT channel at potentials ranging from –100 mV to –40 mV (Fig. 2C). The change in the kinetics of deactivation is consistent with the shift in the voltage dependence of activation.
A computer model of a sensory neuron expressing the electrophysiological properties of the N395K channel displays increased excitability
Our data show that the N395K mutation has two predominant effects on channel gating: it shifts the voltage dependence of activation in the negative direction and impairs slow inactivation of Nav1.7 channels. To investigate how these alterations in sodium channel gating influence action potential firing, we simulated sensory neuron membrane conductances and firing properties in the single compartment model using the NEURON modelling program (Hines & Carnevale, 1997). Hodgkin–Huxley-type simulations of Nav1.7, Nav1.8, a delayed rectifier potassium current and an A-type potassium current were developed based on our sodium current recordings and descriptions in the literature (Gold et al. 1996) to approximate voltage-gated channel behaviour (Fig. 3A–D). The effects of several different mutant conductances were simulated. An activation mutant had its voltage dependence of activation shifted in the negative direction by 10 mV. A negative shift in activation should theoretically increase window current amplitudes. In the model the –10 mV shift increased window currents from 0.05% to 0.2% of the peak current amplitude. A 0.2% window current has an amplitude of 5 pA for a peak current of 2.5 nA. Therefore, while the model does predict that the mutation induces an increase in window currents, a current this size would be difficult to measure experimentally in HEK293 cells. In the model we also simulated a slow-inactivation mutant that had its steady-state slow inactivation altered to mimic the effect of the N395K mutation on this parameter. The simulated N395K mutant conductance combined these two alterations (Fig. 3E). Finally, because some erythromelalgia mutants shift activation in the negative direction but enhance steady-state slow inactivation (Cummins et al. 2004; Choi et al. 2006) we also included a simulated conductance where activation was shifted in the negative direction by 10 mV and slow inactivation was enhanced by increasing the transition rates 2-fold and shifting the midpoint of steady-state slow inactivation by 16 mV in the negative direction. This combination approximates that observed with the F216S erythromelalgia mutation (Choi et al. 2006).
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The N395K mutation decreases Nav1.7 sensitivity to lidocaine
Lysine substitution at the residue corresponding to N395 in rat skeletal muscle Nav1.4 channel (N434) has been shown to reduce the affinity for both amitriptyline and local anaesthetics for inactivated channels (Nau et al. 1999; Wang et al. 2004; Nau & Wang, 2004). Therefore we tested the impact of the N395K mutation on local anaesthetic binding to Nav1.7. Inhibition of channel current by a range of lidocaine concentrations (10 µM to 10 mM) was examined for WT and N395K channels in the inactivated and resting state. Lidocaine binding to the inactivated state of the channels was tested by holding the cells at –120 mV and stepping to an inactivating pre-pulse (–50 mV) for 10 s, which is sufficient to inactivate all of the channels. This was followed by 100 ms pulses to –120 mV to allow recovery from fast inactivation, and the fraction of current remaining was measured in the absence and presence of lidocaine with a 20 ms test pulse to –10 mV.
Inhibition of channel current by lidocaine was attenuated for inactivated N395K channels compared to WT (Fig. 5A). Using a one-site binding fit, the IC50 of lidocaine for inactivated WT channels was estimated at 500 µM while the IC50 for inactivated N395K channels was estimated at 2.8 mM. However, the one-site binding model did not produce a good fit for the dose–inhibition relationship of lidocaine and inactivated WT channels. The data show that between 100 and 300 µM, lidocaine inhibition of WT inactivated channels reaches a plateau before increasing at doses of 1 mM and higher (Fig. 5A). We hypothesized that this plateau in the relationship was due to more than one site being available for lidocaine binding. A two-site binding model gave a much better fit to the WT data. This fit indicated that there are possibly two populations of WT sodium channels at –50 mV, one that exhibits high affinity lidocaine binding while the other exhibits low affinity binding. Based on the two-site fit, the IC50 of lidocaine for the high affinity and low affinity binding to WT channels was 210 µM and 4.3 mM, respectively. The high affinity and low affinity populations may represent channels in different inactivated conformations (see Discussion). These data show that the N395K mutation significantly reduces lidocaine inhibition of Nav1.7 channels at –50 mV, especially the high affinity binding.
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As local anaesthetics are known to exhibit use-dependent inhibition of sodium currents, use-dependent effects of lidocaine on the WT and N395K channels were also studied. We investigated this by pulsing the transfected cells to –10 mV at a frequency of 5 Hz in the absence and presence of drug. Figure 6A and B displays examples of results under control conditions (left) and in the presence of 300 µM lidocaine (right) for both the WT and N395K channels. WT current amplitude decreased 10.4 ± 3.3% (n = 6), 18.1 ± 3.2% (n = 6), 27.8 ± 3.3% (n = 11) and 42.7 ± 2.0% (n = 7) from the first pulse to the last pulse of the protocol in the presence of 10, 30, 100 and 300 µM lidocaine, respectively (Fig. 6C). Decreases in N395K current amplitude were significantly (P < 0.01) smaller (one-way ANOVA follwed by Tukey's comparison test) at 30, 100 and 300 µM lidocaine (Fig. 6C). Decreases observed for the N395K channel were 2.4 ± 0.6% (n = 6), 1.7 ± 0.6% (n = 5), 4.3 ± 0.6% (n = 6) and 11.0 ± 2.6% (n = 5) for 10, 30, 100 and 300 µM lidocaine, respectively.
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Discussion |
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The first goal of this study was to examine electrophysiological differences in Nav1.7 caused by the N395K mutation. The hyperpolarizing shift in activation and slower onset of deactivation caused by N395K is similar to findings involving other hereditary erythromelalgia mutations (Cummins et al. 2004; Han et al. 2006; Harty et al. 2006). All of the other erythromelalgia mutations studied to data (F216S, S241T, I848T, L858H, L858F, F1449V and A863P) shift the voltage dependence of activation in the negative direction by 7–14 mV (Cummins et al. 2004; Dib-Hajj et al. 2005; Choi et al. 2006; Han et al. 2006; Harty et al. 2006; Lampert et al. 2006). All of the mutations, with the exception of F1449V, also slow deactivation, but they differ in the manner in which they affect deactivation. The N395K mutation, like the F216S mutation, simply shifts the voltage dependence of the deactivation time constant curve in the negative direction in a manner that is consistent with the shift in activation. By contrast many of the other mutations, such as the L858H mutation, alter deactivation in a more pronounced voltage-dependent fashion (see Fig. 1D of Cummins et al. 2004). This suggests that the different erythromelalgia mutations alter Nav1.7 gating in at least two distinct ways. The N395K mutation also caused impairment of steady-state slow inactivation which is seen with sodium channel mutations associated with disorders such as epilepsy and hyperkalaemic periodic paralysis, as well as hereditary erythromelalgia (Cummins & Sigworth, 1996; Bendahhou et al. 1999, 2002; Cummins et al. 2004; Rhodes et al. 2005). However, only the I848T and N395K erythromelalgia mutations impair slow inactivation. The F1449V mutation does not alter slow inactivation and the other mutations enhance slow inactivation. This differential effect on slow inactivation might indicate that slow inactivation may not be important for the pathophysiology of erythermalgia.
Interestingly, a recent study (Fertleman et al. 2006) reported that three mutations linked to PEPD, a familial rectal pain syndrome, do not cause a hyperpolarizing shift in activation, but rather impair fast inactivation. It is intriguing that Nav1.7 mutations predominantly associated with rectal pain may consistently impair fast inactivation while mutations predominantly associated with burning pain sensations in the hands and feet cause a hyperpolarizing shift in activation. It is not known if the PEPD mutations alter deactivation or slow inactivation of Nav1.7. A total of eight different missense mutations were identified in families with PEPD but it is not known if all eight mutations alter predominantly fast inactivation or if any of the PEPD mutations increase excitability of sensory neurons.
To examine the relative impact of the changes in activation and slow inactivation associated with erythromelalgia mutations on neuronal excitability, we performed simulations using the NEURON computer program (Hines & Carnevale, 1997). Our simulations indicate that the shift in activation is sufficient to decrease the threshold for action potential generation and increase repetitive firing rates, changes that have been observed when recombinant F1449V, L858H, and A863P sodium channels were transfected into cultured DRG neurons (Dib-Hajj et al. 2005; Harty et al. 2006; Rush et al. 2006). The simulations also indicated that while changes in slow inactivation may not have major influences on neuronal excitability by themselves, they can significantly modulate the degree to which the shift in activation impacts excitability. These data suggest that mutations with impaired slow inactivation are more likely to be associated with a severe clinical phenotype. The HH formulations used in the simulations presented here are not adequate to determine whether the differential effects of the mutations on deactivation kinetics can also modulate excitability and therefore we did not address this in the present study. Nonetheless, these results, based on observations of a hereditary erythromelalgia mutation, may provide insight into how electrophysiological changes in Nav1.7 contribute to pain in humans. For example, our findings should help in determining whether changes in Nav1.7 properties caused by modulation (e.g. phosphorylation) are likely to cause increased sensitization of nociceptive neurons and contribute to increased pain sensations.
The second goal of this study was to determine whether the N395K mutation changes the effects of lidocaine on Nav1.7 current. The N395 residue is located in the S6 segment of domain I and lies within the local anaesthetic-binding site of Nav1.7. One study has shown that lidocaine relieved pain in only 55% of erythromelalgia patients (Davis & Sandroni, 2005). This raises the possibility that specific mutations such as N395K in Nav1.7 might render patients relatively unresponsive to lidocaine treatment. Therefore, our hypothesis was that mutation of this residue would reduce the inhibitory effects of lidocaine on channel current. Interestingly, we observed a two-phase inhibition of lidocaine on 1.7 WT-inactivated channels. The dose–response curve for this inhibition was best fitted by using a two-site binding model. One possibility for this phenomenon is that holding the cell at –50 mV for prolonged periods of time transitions a subset of channels into a high affinity binding state and a subset into a low affinity binding state. These two distinct states could involve closed-state and open-state inactivation which may have different affinities for lidocaine. A recent study in the giant squid axon showed that at –50 mV total inactivation is comprised of both closed-state and open-state inactivation (Armstrong, 2006). The possibility that the ratio of closed-state versus open-state inactivated channels during the prolonged –50 mV pre-pulse might explain the two-phase dose–inhibition curve for lidocaine on 1.7 WT-inactivated channels, while outside the scope of our current study, deserves further investigation. Nonetheless, our data clearly demonstrate an attenuation of lidocaine inhibition on inactivated N395K channels at –50 mV. This reduction of the lidocaine effect was seen at all concentrations tested including concentrations comparable to total lidocaine serum concentrations (10–16 µM) that have been shown to cause analgesia in neuropathic pain patients (Ferrante et al. 1996).
One explanation for the reduction of the lidocaine effect on the N395K channel is that the mutation impairs steady-state slow inactivation and thus there are fewer channels in the inactivated state. This could decrease the number of channels that preferentially bind lidocaine. Indeed, based on recordings from Xenopus oocytes, Chevrier et al. (2004) proposed that lidocaine might alter slow inactivation of Nav1.7 channels. However, F216S channels display enhanced steady-state inactivation at –50 mV (Choi et al. 2006) and in the present study, F216S channels did not show enhanced inhibition with lidocaine in the inactivated state compared to wild-type. Furthermore, WT channels pre-pulsed to –70 mV for 30 s and N395K channels pre-pulsed to –50 mV for 30 s both have approximately 21% of their channels slow inactivated (Fig. 1C), yet WT channels still displayed a higher inhibition of channel current by lidocaine with a –70 mV pre-pulse than N395K channels with a –50 mV pre-pulse (data not shown). This indicates that the N395K mutation in Nav1.7 alters the direct interaction of lidocaine with the inactivated state of the channel, and suggests that changes in steady-state slow inactivation do not affect lidocaine inhibition of the channel. In addition, the inhibitory effect of lidocaine at high concentrations on the resting N395K channel was reduced compared to the resting WT channel. This further confirms that the N395K mutation affects the direct interaction of lidocaine with Nav1.7 because the percentage of total channels in the resting state would be the same for N395K and WT using a –140 mV pre-pulse.
A recent study has reported that a loss of functional Nav1.7 channels in humans due to homozygous nonsense mutations leads to a complete inability to sense pain (Cox et al. 2006). Surprisingly, individuals with this profound inability to sense pain were deemed to be healthy in virtually every other way. Although Nav1.7 is believed to be present in both sensory and sympathetic ganglion neurons, none of the individuals studied by Cox et al. exhibited symptoms of autonomic nervous system dysfunction. As non-nociceptive sensory functions appeared normal in the individuals with the homozygous nonsense mutations that eliminated functional Nav1.7 currents, the study by Cox et al. indicates that drugs that specifically target Nav1.7 could have the potential to selectively ameliorate pain. It is important to note that the individuals studied by Cox et al. (2006) were all children between the ages of 6 and 14, so it is not yet clear if they have a complete inability to develop neuropathic pain. It has been reported that mice that selectively lack Nav1.7 in their sensory neurons can develop neuropathic pain (Nassar et al. 2004) suggesting that Nav1.7 may not be essential for neuropathic pain. However, mice that have a global deficit in Nav1.7 die just after birth (Nassar et al. 2004), indicating that the role(s) of Nav1.7 could be different between mice and humans. Although there is now clear evidence that at least two types of chronic painful neuropathies, hereditary erythromelalgia and PEPD, can be caused by mutations in SCN9A that alter the gating properties of Nav1.7, it is not known if Nav1.7 is important in other human chronic pain syndromes. Drugs that specifically block Nav1.7 currents could help determine if Nav1.7 channels play a major role in common neuropathic pain syndromes such as diabetic neuropathy or pain associated with post-herpetic neuralgia.
Our results suggest that variability in the response of erythromelalgia patients to local anaesthetics that target sodium channels may be due, at least in part, to differing effects of the various hereditary erythromelalgia mutations in Nav1.7 on lidocaine sensitivity. Our data predict that the pain symptoms in individuals with the N395K mutation are likely to be more intense and more resistant to lidocaine treatment than for individuals with the F216S mutation. Genotype–phenotype correlations are needed to help determine if this is indeed the case. Although none of the other hereditary erythromelalgia mutations identified thus far are located in the local anaesthetic-binding site, the possibility that one or more of these also alters lidocaine sensitivity merits examination in future experiments. Interestingly, all of the hereditary erythromelalgia mutations studied to date, including the N395K mutation studied here, induce a hyperpolarizing shift in the voltage dependence of activation of Nav1.7 currents, suggesting that this alteration is crucial to the development of the pain associated with hereditary erythromelalgia. We speculate that other manipulations, such as phosphorylation, that also induce hyperpolarizing shifts in activation may also contribute to increased pain sensations. Thus our observations on the N395K mutation provide insight into the changes in Nav1.7 channel activity that may contribute to abnormal pain sensations in erythromelalgia. Our results also suggest that research focusing on the function and pharmacology of the Nav1.7 channel is crucial to the understanding of pain pathophysiology and the advancement of pain management therapies.
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Footnotes |
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References |
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) current only one action potential was generated for current injections of 100–350 pA. The response was virtually identical when 50% of the Nav1.7 current had impaired slow inactivation (
). Repetitive firing was increased to different extents when 50% of the Nav1.7 current had hyperpolarized activation (
), hyperpolarized activation combined with impaired slow inactivation (N395K mutant; 
) or hyperpolarized activation combined with enhanced slow inactivation (F216S mutant; 
). B–F, action potentials simulated in response to a 10 s injection of 350 pA are shown for the different combinations of Nav1.7 wild-type and mutant currents.
