SEARCH

SEARCH BY CITATION

Abstract

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

Single-point missense mutations in the peripheral neuronal voltage-gated sodium channel Nav1.7 are implicated in the painful inherited neuropathy paroxysmal extreme pain disorder (PEPD). The Nav1.7 PEPD mutations are located in regions of the channel suggested to play important roles in fast inactivation. PEPD mutations in the putative inactivation gate have been reported to significantly impair fast inactivation, resulting in pronounced persistent currents. However, PEPD mutations in the S4–S5 linker of domain 3 (D3/S4–S5) had not been characterized and the roles of specific residues in this linker in channel gating are unclear. We functionally characterized two of the D3/S4–S5 PEPD mutations (V1298F and V1299F) and compared their effects on gating to an adjacent non-PEPD mutation (V1300F) and the I1461T PEPD mutation, located in the putative inactivation gate. The primary effect of the V1298F and V1299F mutations is to shift the voltage dependence of fast inactivation by ∼20 mV in the depolarizing direction. We observed a similar effect with the PEPD mutation I1461T. Interestingly, while all three PEPD mutations increased persistent currents, the relative amplitudes (∼6% of peak) were much smaller than previously reported for the I1461T mutation. In contrast, the main effect of the V1300F mutation was a depolarizing shift in the voltage dependence of activation. These data demonstrate that (1) mutations within D3/S4–S5 affect inactivation of Nav1.7 in a residue-specific manner and (2) disruption of the fast-inactivated state by PEPD mutations can be more moderate than previously indicated, which has important implications for the pathophysiology of PEPD.

Voltage-gated sodium channels (VGSCs) are dynamic transmembrane proteins that selectively conduct sodium ions through an aqueous pore. In response to changes in the local electric potential across the cell membrane, these pore-forming proteins undergo specific conformational (gating) modifications between ion conducting (open) and non-conducting (closed and inactivated) states. VGSCs are critical in determining the firing threshold and the upstroke of action potentials (APs) in excitable tissues (Hodgkin & Huxley, 1952; Rush et al. 2007). VGSCs are thought to contain multiple subunits. The α-subunit (220–260 kDa) is important for drug binding, ion selectivity and pore formation (Catterall, 2000) whereas auxiliary β-subunits (32–36 kDa) have been suggested to play roles in channel modulation and trafficking (Isom et al. 1992, 1995a,b). The structure of the highly conserved pore-forming α-subunit consists of four homologous domains (D1–D4) each containing six membrane-spanning segments (S1–S6). Electrophysiological studies have shown that mutations within VGSCs can affect their gating properties and, thus, alter tissue excitability (Dib-Hajj et al. 2005; Rush et al. 2006; Kalume et al. 2007). Nine distinct α-subunits (Nav1.1–9) and four β-subunits (β1–4) have been identified. The nine α-subunits are differentially expressed in excitable tissues. For example, the tetrodotoxin (TTX) resistant Nav1.5 is predominantly expressed in cardiac tissue, whereas the TTX-sensitive Nav1.7 is expressed at high levels in the peripheral nervous system, including nociceptive neurons (Catterall et al. 2005). Both TTX-sensitive and TTX-resistant VGSCs expressed in pain-sensing (nociceptive) dorsal root ganglion (DRG) neurons are hypothesized to play critical roles in pain mechanisms (Cummins et al. 2007). The importance of Nav1.7 in pain has been confirmed; individuals with a complete inability to perceive pain have distinct homozygous nonsense mutations in SCN9A (the gene encoding Nav1.7) resulting in loss of channel function. These individuals showed no additional signs of nervous system defects (Cox et al. 2006). Recently, specific single-point missense mutations in SCN9A, clustered in highly conserved regions, have been linked to the distinct neuropathic pain syndromes inherited erythromelalgia (IE) and paroxysmal extreme pain disorder (PEPD) (Drenth et al. 2005; Fertleman et al. 2006). Both IE and PEPD are inherited autosomal dominant disorders. IE is characterized by intermittent attacks of extreme pain that are associated with red, warm and swollen hands, feet, arms and torso (Drenth & Michiels, 1992; Orstavik et al. 2004). In contrast, patients with PEPD, previously known as familial rectal pain, report severe bouts of intense burning pain in the rectum, eye and jaw (Hayden & Grossman, 1959; Fertleman et al. 2007). The IE mutations selectively affect activation and deactivation properties of the channel, resulting in a lowered threshold for activation (Cummins et al. 2004; Dib-Hajj et al. 2005; Choi et al. 2006). A majority of the Nav1.7 PEPD mutations are localized to the cytosolic regions of D3 and D4 and are predicted to play an important role in transition to the inactivated state. Although it has been proposed that all PEPD mutations will selectively impair fast inactivation (Fertleman et al. 2006), several of the identified PEPD mutations had not been functionally characterized.

In this study we investigated the biophysical properties of two single-point PEPD mutations (V1298F and V1299F) within the D3/S4–S5 linker region that had not been previously characterized. Previous studies on rat Nav1.2 channels expressed in Xenopus oocytes suggested the D3/S4–S5 linker may serve as a critical ‘docking’ site for the putative inactivation gate (IFMT) of the D3–D4 cytosolic linker via van der Waals interactions between a cluster of hydrophobic residues (Smith & Goldin, 1997). However, S4–S5 linkers have also been implicated in the activation gating of voltage-gated ion channels (Long et al. 2005a) and questions remain regarding the role of the D3/S4–S5 linker in VGSC gating. We compared the effects of the V1298F and V1299F PEPD mutations with the I1461T PEPD mutation, which is located within the putative inactivation gate of the D3–D4 cytosolic linker and has been reported to markedly disrupt Nav1.7 inactivation (Fertleman et al. 2006). We also mutated an adjacent and identical residue in the D3/S4–S5 linker, V1300F (which has not been implicated in PEPD), to evaluate the residue-specific effects of D3/S4–S5 linker mutations on channel gating. Results from our studies demonstrate that PEPD mutations within the D3/S4–S5 linker alter the stability of the inactivated state and reveal this destabilization is due, in part, to the location and orientation of the specific mutations within the D3/S4–S5 cytosolic linker. Additionally, our research offers new insight into the pathophysiology of pain associated with PEPD by showing that disruption of the fast-inactivated state by PEPD mutations is not necessarily as marked as previously reported.

Methods

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

Ethical information

Human embryonic kidney (hEK293) cells were obtained from ATCC, Manassas, VA, USA. Use of the hEK293 cells was approved by the Institutional Biosafety Committee and conformed to the ethical guidelines for the National Institutes of Health for the use of human-derived cell lines.

Transfections

The human Nav1.7 (hNav1.7 wild-type, V1298F, V1299F, V1300F, or I1461T) channels were transiently cotransfected with human β1 (hβ1) and β2 subunits (hβ2) (Lossin et al. 2002) into human embryonic kidney (hEK293) cells using the calcium phosphate precipitation technique. For a single set of experiments we also used EGFP (pIRES2-EGFP) as a transfection control. Mutagenic primers were designed to introduce the correct base pair change into the hNav1.7 channel using Vector NTI Advance 10 (Invitrogen, Carlsbad, CA, USA). Mutations inserted into the hNav1.7 channel cDNA (Klugbauer et al. 1995) construct were produced using the QuickChange XL site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA) according to the manufacturer's protocol. 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 (FBS). 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 24–72 h after transfection.

Whole-cell patch-clamp recordings

Whole-cell patch-clamp recordings were conducted at room temperature (∼21°C) after obtaining a gigaohm seal (1–20 GΩ) using a HEKA EPC-10 amplifier. Data were acquired on a Windows-based Pentium IV computer using the Pulse program (v. 8.65, HEKA Electronik, Germany). Fire-polished electrodes (0.9–1.4 MΩ) were fabricated from 1.7 mm capillary glass using a Sutter P-97 pullter (Novato, CA, USA). The standard CsF dominant electrode solution consisted of (in mm): 140 CsF, 10 NaCl, 1.1 EGTA and 10 Hepes, pH 7.3. The standard CsCl/caesium aspartate dominant solution consisted of (in mm): 50 CsCl, 70 caesium aspartate, 10 NaCl, 10 Hepes, 11 EGTA, 1 CaCl2, and 2 Mg-ATP, pH 7.3. The standard bathing solution consisted of (in 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 bathing solution. Cells were selected based on their ability to express EGFP. Series resistance errors were always compensated to be less than 3 mV during voltage-clamp recordings. Data were not recorded before three minutes after whole-cell configuration had been established to allow adequate time for the electrode (intracellular) solution(s) to equilibrate. Data recordings did not last more than 45 min and cells were not held in the standard bathing solution for more than 1 h.

Data analysis

Voltage-clamp experimental data were analysed using the Pulsefit (v. 8.65, HEKA Electronik, Germany), Origin (v. 7.0, OriginLab Corp., Northampton, MA, USA), and Microsoft Excel software programs. Normalized conductance–voltage (G–V) relationships were derived using:

  • image(1)

where GNa is macroscopic sodium conductance, Imax is calculated as peak current in response to the test pulse, Vm is the test pulse voltage, and ENa is the measured Na+ equilibrium potential. Normalized availability curves were fitted using a standard single-phase Boltzmann distribution for G–V and steady-state fast inactivation (h) and a double-phase Boltzmann distribution for steady-state slow-inactivation (s) data. Midpoint (V1/2) and slope factors (Z) of (activation) conductance-voltage (G–V) and voltage-dependent steady-state fast inactivation curves were calculated using a standard single-phase Boltzmann distribution fit according to:

  • image(2)

All data are shown as means ±s.e.m. Statistical significance was assessed with Microsoft Excel using Student's unpaired t test. Statistical significance of difference was accepted at P < 0.05.

Results

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

PEPD mutations within the D3/S4–S5 linker have small effects on the voltage dependence of channel conductance but do not alter deactivation time constants

Rapidly activating and inactivating TTX-sensitive sodium currents were observed in hEK293 cells transiently transfected with cDNA for wild-type (WT) and PEPD mutant hNav1.7 channels (Figs 1 and 2). Surprisingly, initial comparison of the current traces did not suggest any major differences in channel kinetics (Fig. 2A) or current–voltage (I–V) properties (Fig. 2B). However, expression levels of V1298F (−144.4 ± 32.3 pA pF−1, n= 7) mutant channels were significantly (P < 0.05) lower than WT (−332.6 ± 59.4 pA pF−1, n= 11), V1299F (−213.3 ± 63.6 pA pF−1, n= 9), and I1461T (−321.3 ± 63.8 pA pF−1, n= 14) channels as determined by current density. Voltage-dependent gating properties were investigated to determine the effects of D3/S4–S5 PEPD mutations on transitions between conducting and non-conducting channel states. The V1298F and V1299F mutations (Fig. 1A) caused small, but significant (P < 0.05, Table 1), depolarizing shifts in the voltage dependence of channel conductance (G–V) and an increase in conductance slope factor for Nav1.7 channels. These shifts were parameterized by fitting data to a single-phase Boltzmann function and comparing the voltage at which 50% of the channel population (V1/2) was conducting Na+ and the fitted slope factor (Z) at this voltage (Fig. 3A). We did not observe a significant (P > 0.05) shift in the V1/2 of channel conductance or Z for the PEPD mutation (I1461T) in the D3–D4 linker located in the putative inactivation gate, which suggests this mutation does not directly affect gating charge movement. We also evaluated the effects of the PEPD mutations on deactivation, which involves rapid, voltage-dependent transitions from open to closed states via an integrated return of the S4 ‘voltage sensors’ to a primed position (Oxford, 1981; Kuo & Bean, 1994; Featherstone et al. 1998). Interestingly, the voltage-dependent deactivation time constants (τd), examined by eliciting tail currents at a range of potentials after briefly activating the channels for the mutant PEPD channels, were not significantly (P > 0.05) affected when compared to WT (Fig. 3B).

image

Figure 1. Diagrammatic and sequence schemes of mutated regions within VGSC Nav1.7 A, linear representation of VGSC α-subunit structure with putative inactivation motif labelled with amino acid residues IFMT. Mutations within the D3/S4–S5 and D3–D4 linker implicated in PEPD are indicated with open square symbols. An adjacent and identical mutation (V1300F), not implicated in PEPD, is indicated with a filled square. B, sequence alignment of human voltage-gated sodium channels. Mutated residues implicated in PEPD are emphasized with bold lettering. Underlined are regions of Nav1.7 mutated in this study which include adjacent hydrophobic valine residues (V1298, V1299, and V1300) within the D3/S4–S5 cytosolic linker and the isoleucine within the putative inactivation motif (I1461) within the D3–D4 cytosolic linker.

Download figure to PowerPoint

image

Figure 2. Comparison of whole-cell ionic current traces and normalized current–voltage properties in CsF dominant electrode solution A, representative traces from WT and PEPD mutant Nav1.7 channels coexpressed with hβ1 and hβ2 subunits in hEK293 cells. B, normalized current–voltage (I–V) properties were assessed using depolarizing step pulses. Cells were held at 120 mV. The currents were elicited by 100 ms test pulses to various potentials from 80 to +90 mV stepped in increments of 5 mV. The peak current evoked by each pulse, normalized to the maximum peak current, is plotted versus the pulse voltage. C, two-dimensional representation of native amino acid residues and mutated side chain moieties implicated in PEPD.

Download figure to PowerPoint

Table 1.  Boltzmann parameters of channel activation and steady-state inactivation curves for WT and PEPD mutant channels
ChannelVoltage-dependence of activationVoltage-dependence of steady-state fast-inactivation
V1/2 (mV)Slope (mV/e-fold)nV1/2 (mV)Slope (mV/e-fold)n
  1. Values derived for V1/2, the voltage of half-maximal activation and inactivation, and the slopes were derived from Boltzmann distribution fits to the averaged and normalized (±s.e.m.) voltage dependence of activation and steady-state inactivation curves.

WT28.9 ± 0.25.9 ± 0.21177.6 ± 0.16.1 ± 0.111
V1298F22.6 ± 0.26.4 ± 0.2 757.8 ± 0.26.2 ± 0.2 7
V1299F24.4 ± 0.26.7 ± 0.2 956.6 ± 0.26.1 ± 0.2 9
I1461T26.4 ± 0.26.0 ± 0.214−58.8 ± 0.26.8 ± 0.214
image

Figure 3. Effects of PEPD mutations within D3/S4–S5 and D3–D4 linkers on voltage-dependent gating properties of Nav1.7 A, comparison of the whole-cell conductance–voltage properties (G–V), calculated from current–voltage (I–V) relationship. B, voltage-dependent time constants for tail current deactivation (τd) at repolarization potentials ranging from 40 to 100 mV. The kinetics of Na+ channel deactivation from the open state were assessed by eliciting tail currents at a range of potentials after briefly activation the channels (0 mV for 0.5 ms). C, comparison of steady-state inactivation for WT and mutant channels. The protocol is comprised of an incrementally depolarizing prepulse from 130 mV, 500 ms in duration, followed immediately by a depolarizing step to +10 mV from a holding potential of 120 mV. D, time constants for open-state fast inactivation as a function of voltage for WT and mutant channels. The decay phases of currents elicited during channel activation protocol were fitted with a Hodgkin–Huxley type m3h model to estimate open-state inactivation time constants (τh). E, representative normalized currents from whole-cell recordings of cells expressing either WT or PEPD mutant channels at +10 mV. F, averaged ramp current (Iramp) traces (n= 6–14) elicited with a slow depolarizing (0.27 mV ms−1) stimulus from a holding potential of 120 mV to +40 mV for PEPD mutant and WT channels. Ramp current amplitude is expressed as a percentage of the peak transient current elicited with a standard I–V protocol. Plotted values for A and C were normalized and fitted with a single-phase Boltzmann distribution for WT and PEPD mutant channels. Voltage protocols are displayed within the insets.

Download figure to PowerPoint

Mutations within the D3/S4–S5 linker, implicated in PEPD, significantly shift the voltage dependence of the steady-state fast inactivation profile

The voltage dependence of WT and PEPD mutant channel fast inactivation was tested using a two-step protocol (Fig. 3C) to determine the fraction of channels transitioning to an inactivated (non-conducting) state in response to changes in membrane potential. To ensure steady-state conditions (h) over the full range of potentials tested, we used 500 ms conditioning pulses. The PEPD mutations (V1298F, V1299F and I1461T) caused a significant (P < 0.05) depolarizing shift in the V1/2 of fast inactivation with no significant (P > 0.05) change in Z. The V1/2 of inactivation (defined as the voltage at which 50% of the channel population has transitioned to a non-conducting state) was depolarized by ∼20 mV for each mutant channel (Table 1) compared to WT, thus resulting in an increased fraction of mutant channels available to conduct Na+ at more depolarized potentials likely to be due to decreased stabilization of the inactivated states. To evaluate the contribution of decreased stabilization on the kinetics of the Na+ current decay phase, we calculated the time constants for inactivation (τh) during depolarization to various potentials (Fig. 3D). At positive potentials (beyond −10 mV) the inactivation time constants for the PEPD mutant channels were significantly (P < 0.05) larger compared to WT channels. Therefore, we chose to compare the decay phase of raw current traces for the PEPD mutants at +10 mV, a potential where the mutant channel time constants seemed to plateau in response to further depolarizing steps in voltage (Fig. 3E). The decay phase of the current elicited, which represents a decrease in open channel probability (increased transition to a fast-inactivated state), for PEPD mutant traces was broader compared to WT and revealed a small persistent component (∼6% normalized current remaining at 8 ms). Thus, it is possible PEPD mutations disrupt the molecular interactions required to stabilize the fast-inactivated state.

Although we observe impaired fast inactivation with the PEPD mutations, the severity of the impairment observed with the I1461T mutation is much less than that reported for this mutation by Fertleman et al. (2006). One possible reason for the differences observed was that we cotransfect the Nav1.7 cDNAs with auxiliary β1 and β2 subunits and they apparently did not. Therefore, we cotransfected hEK293 cells with WT or I1461T constructs with and without β-subunits (using EGFP as our transfection control) to determine if this might influence the size of the non-inactivating component. The inclusion or exclusion of β-subunits during hEK293 transfection did not increase the non-inactivating component of WT or I1461T currents (see online Supplemental material, Supplemental Fig. 1). Another possible factor was that we used 140 mm CsF in our electrode solution and Fertleman et al. used 13 mm CsF. Fluoride has been reported to alter persistent sodium currents in Nav1.3 channels (Chen et al. 2000; Meadows et al. 2002). Therefore, we performed additional recordings of V1299F, I1461T and WT channels using a CsCl/caesium aspartate dominant electrode solution that did not contain fluoride (Supplemental Fig. 2). The voltage dependence of conductance and steady-state fast inactivation obtained with this solution are depolarized compared to those obtained with fluoride (Supplemental Table 1), but we do not see evidence of incomplete steady-state inactivation. However, the amount of persistent current observed at the end of 25 ms depolarizing pulses was slightly greater in the absence of fluoride for I1461T (∼8% of peak) and V1299F (∼15% of peak). These data are consistent with observations suggesting VGSCs can mediate more pronounced persistent currents when expressed in hEK293 cells in the absence of intracellular fluoride ions (Chen et al. 2000; Meadows et al. 2002).

PEPD mutant channels reveal an increased persistent component in response to a slow-duration depolarizing stimulus

Destabilization of the inactivated state can affect the region where steady-state channel activation and inactivation overlap, resulting in an increased probability of channel reopening before inactivating near resting membrane potential (RMP), and the development of persistent ‘window’ currents (Hodgkin & Huxley, 1952). Furthermore, it has been proposed that because persistent neuronal currents, or ramp currents (Iramp), are activated near RMP, these currents may influence action potential (AP) threshold and cellular excitability (Crill, 1996). Therefore, in order to better quantify the persistent component observed above in the PEPD mutant channel traces, we decided to examine WT and PEPD mutant Nav1.7 currents evoked by a slow depolarizing ramp (0.27 mV ms−1) stimulus from a holding potential of −120 mV. We observed a significant (P < 0.05) increase in the amplitude of the ramp currents elicited especially at more depolarized potentials for the mutations implicated in PEPD within the D3/S4–S5 and D3–D4 intracellular linkers compared to WT channels in response to a slow depolarizing ramp stimulus (Fig. 3F). Interestingly, the Iramp characteristics for the PEPD mutations revealed an impaired transition to a non-conducting state, confirming a destabilized channel configuration at more depolarized potentials.

PEPD mutations within the D3/S4–S5 linker differentially affect development of CSI compared to the I1461T PEPD mutation

Before reaching an open (ion conducting) state, VGSCs can transition to an inactivated state during small depolarizing steps, to relatively negative potentials, thereby reducing the population of channels available for activation by stronger depolarizing steps (Oxford & Pooler, 1975; Bean, 1981). Furthermore, it has been demonstrated that transition to the closed-inactivated state can play an important role in Na+ channel response during subthreshold depolarizations (Cummins et al. 1998). Therefore, we tested the effects the PEPD mutations have on development of closed-state inactivation (CSI). We used a time varied conditioning pulse to −60 mV, from a holding potential of −120 mV, followed by a test pulse to 0 mV for 20 ms, to determine the fraction of channels available. We chose −60 mV because this potential is near resting membrane potential (RMP) for nociceptive sensory neurons, and thus changes in VGSC gating properties near this potential may have a large impact on AP threshold (Herzog et al. 2001; Renganathan et al. 2001; Rush et al. 2006; Rush et al. 2007). Consistent with our steady-state inactivation data, a decrease in the fraction of mutant channels transitioning to a closed-inactivated state at this potential was observed compared to WT (Fig. 4A). Additionally, we noticed a significant (P < 0.05) difference in the time constants (τ−60) for CSI between PEPD mutations in the D3/S4–S5 and D3–D4 (inactivation gate) linker (Table 2). Surprisingly, the PEPD mutations within the D3/S4–S5 linker (V1298F and V1299F) displayed time constants smaller than those for both the WT channels and channels with the mutation within the putative inactivation gate (I1461T), indicating a faster transition to closed-inactivated states for the D3/S4–S5 PEPD mutants.

image

Figure 4. Effects of D3/S4–S5 PEPD mutations on inactivation kinetics A, development of closed-state inactivation (CSI) near resting membrane potential (n= 5–16). The standard voltage protocol for development of CSI involved prepulsing the cells from a holding potential of 120 mV to a relatively hyperpolarized potential (60 mV) for increasing time durations and then quickly testing for the fraction of current available using a strong depolarizing stimulus to 0 mV for 20 ms. B, time course for recovery from CSI near resting membrane potential (n= 7–10). Recovery from CSI was examined using a conditioning pulse to 60 mV for 500 ms quickly followed by a hyperpolarizing pulse to 90 mV for increasing time durations to allow adequate recovery then tested for the fraction of current available by depolarizing the cells to 0 mV for 20 ms. C, recovery from open-state inactivation (OSI) to primed channel state (n= 46). To test for channel recovery from OSI the cells were prepulsed to a relatively depolarized potential of 0 mV for 20 ms then hyperpolarized to 90 mV for increasing time increments and were then tested for current available using a strong depolarizing stimulus to 0 mV for 20 ms. Plotted values were fitted with a single exponential function to determine time constant values (τ). Development and recovery protocols are displayed within the insets.

Download figure to PowerPoint

Table 2.  Development and recovery inactivation time constant values for WT and PEPD mutant and channels
ChannelDevelopment time constant (τ) (ms)nRecovery time constant (τ) (ms)n
  1. Time constant values for development (τ−60) of and recovery (τ−90) from closed-state inactivation (CSI). Additionally, we obtained time constant values for development (τ0) of and recovery (τ−90) from open-state inactivation (OSI). Voltage-derived time constant values for development of OSI at 0 mV were determined by fitting the decay phases of currents elicited for WT and mutant channels using a Hodgkin–Huxley m3h function. Development of CSI and recovery from OSI and CSI data were fitted using a single-exponential function and time constant values were calculated and averaged from individual cell recordings.

Closed-state inactivation (CSI)
 WT74.2 ± 8.21646.9 ± 3.36
 V1298F49.3 ± 2.7 4 9.7 ± 0.97
 V1299F31.7 ± 3.5 8 7.3 ± 1.18
 I1461T68.8 ± 3.2 515.6 ± 1.210 
 WT 0.6 ± 0.11130.6 ± 5.66
Open-state inactivation (OSI)
 V1298F 1.0 ± 0.03 7 9.6 ± 0.74
 V1299F 0.8 ± 0.1 9 4.2 ± 0.75
 I1461T 0.8 ± 0.031413.0 ± 2.15

Mutations associated with PEPD disrupt the stability of inactivation by increasing the recovery rate from CSI

The decrease in the inactivation time constant (τ−60) and incomplete development of CSI for the PEPD mutations within the D3/S4–S5 linker are intriguing because they suggest that these residues may play a direct role in stabilizing the closed-inactivated state of Nav1.7. Therefore, we next examined the inactivation kinetics by evaluating the effects the PEPD mutations had on recovery from CSI by pulsing cells to −60 mV for 500 ms and then allowing channels to recover at −90 mV for varied time increments before testing for available current (Fig. 4B). All of the PEPD mutant channels tested (V1298F, V1299F, and I1461T) recovered from CSI faster than WT. The recovery time constants (τ−90) for the PEPD mutant channels were significantly (P < 0.05) faster (3–8 fold) than those for WT channels (Table 2). These results likely contribute to the incomplete development of CSI for the PEPD mutant channels at this potential.

D3/S4–S5 and D3–D4 PEPD linker mutations allow Nav1.7 channels to recover from OSI faster than WT channels

Because we observed a depolarizing shift in the V1/2 of steady-state inactivation (h) and changes in the kinetics of CSI for PEPD mutant channels compared to WT we also examined channel recovery from open-state inactivation (OSI). VGSCs quickly transition to an open-inactivated state after reaching an ion-conducting (open) state (Hodgkin & Huxley, 1952; Hoyt, 1971; Goldman & Schauf, 1972; Armstrong et al. 1973; Armstrong & Bezanilla, 1977; Bezanilla & Armstrong, 1977). To examine recovery from OSI a two-step protocol was implemented. Cells were conditioned at depolarized potentials (0 mV, 20 ms) before hyperpolarizing the membrane potential (−90 mV) for varied times and then testing for available current (0 mV, 20 ms). We chose a recovery potential of −90 mV based on our steady-state inactivation (h) data which shows that a large population (∼90%) of WT channels are available for ion conductance at this potential. All PEPD mutant channels recovered from OSI faster than WT channels with significant (P < 0.05) decreases in recovery time constants (τ−90) (Fig. 4C). This is consistent with a decreased fraction of mutant channels transitioning to the fast open inactivated states at 0 mV.

An adjacent and identical mutation within the D3/S4–S5 linker, not implicated in PEPD, displays unique voltage-dependent and kinetic properties compared to neighbouring PEPD mutations

Based on amino acid sequence analysis and helical wheel projections, it has been proposed that the D1–D4 S4–S5 linkers form a highly conserved α-helical secondary structure (Lerche et al. 1997; Filatov et al. 1998). The secondary structure of the linkers would indicate that residues may interact based on their orientation around the α-helix. To test this possibility, we mutated an adjacent and identical residue (V1300F), not implicated in PEPD, within the S4–S5 linker of D3 to examine the location-specific effects of the valine to phenylalanine substitutions within the S4–S5 linker. This mutant channel produced currents with rapid conductance and inactivation characteristics (Fig. 5A). We further examined the I–V properties of the V1300F mutant in comparison to WT channels (Fig. 5B). The V1300F mutation resulted in a decreased probability of the channel population opening at voltages near threshold (−40 mV), which were sufficient to allow sodium flux through WT channels. This suggests the V1300F mutation decreases the initial voltage sensitivity required for D3 activation, and thus impairs the fraction of channels available to open. As such, we evaluated the voltage-dependent properties and kinetics of gating for the V1300F mutant channel in a similar manner to that described for the PEPD mutations. When comparing WT and mutant G–V characteristics, we observed a significant (P < 0.05) depolarizing shift in the V1/2 of conductance (−18.50 ± 0.20 mV) and Z (6.92 ± 0.18 mV/e-fold) for the V1300F mutant channel and a very slight, but significant (P < 0.05), hyperpolarizing shift in V1/2 of steady-state inactivation (−81.96 ± 0.15 mV) with no significant (P > 0.05) change in Z (6.39 ± 0.13 mV/e-fold) (Fig. 5C). Additionally, in contrast to the adjacent PEPD mutations, the V1300F mutation significantly (P < 0.05) increased the voltage-dependent deactivation time constants (τd) at −60 and −40 mV (Fig. 5D) and had little effect on OSI at potentials near +10 mV (Fig. 5E and F). We also compared Iramp elicited using a slow depolarizing stimulus (0.27 mV ms−1) for WT and mutant channels (Fig. 5G) and noticed a significant decrease (P < 0.05) in the percentage peak current remaining for the V1300F mutant channels compared to WT (Fig. 5H). This was in contrast to the adjacent PEPD mutations that significantly (P < 0.05) increased ramp current elicited. Consistent with this, we observed an enhanced transition to CSI for the V1300F mutant channels, and a significant (P < 0.05) decrease in the time constant (τ−60) for CSI compared to WT (Fig. 6A). However, this mutation did not significantly (P > 0.05) alter the time constants for recovery from CSI or OSI compared to WT (Fig. 6B and C).

image

Figure 5. An adjacent and identical mutation within the D3/S4–S5 linker of hNav1.7, not implicated in PEPD, displays altered voltage-dependent channel properties in a manner distinct from PEPD mutations A, representative current trace for the V1300F mutant channels coexpressed with hβ1 and hβ2 obtained using a +5 mV stepwise pulse protocol from 80 to +90 mV for a duration of 100 ms from a holding potential of 120 mV to elicit inward Na+ current in a CsF dominant electrode solution. B, normalized current–voltage (I–V) plot comparison of WT and V1300F mutant channels. C, steady-state gating characteristics of WT and V1300F mutant channels determined from separate activation and inactivation pulse protocols. Voltage-dependent conductance properties were derived from I–V data and the steady-state inactivation profiles were determined using a stepwise conditioning pulse from 150 mV to +10 mV (in +10 mV increments) for 500 ms and followed by a test pulse (0 mV, 20 ms). D, voltage-dependent deactivation characteristics were examined for WT and V1300F channels using the protocol displayed within the inset. E, open-state fast inactivation (OSI) time constants (τh) were determined using a Hodgkin–Huxley (m3h) type fit to the decay phase of individual current traces in a similar manner as described for Fig. 3D. F, representative current trace for WT and mutant channels at +10 mV emphasizing the decay phase for each trace. G, averaged ramp current (Iramp) elicited in response to slow depolarizing stimulus (0.27 mV ms−1) for WT and V1300F channels utilizing the same protocol that was used to examine PEPD mutant channels. Ramp current amplitude is expressed as a percentage of the peak transient current elicited with a standard I–V protocol. H, bar graph interpretation of percentage peak current during the slow depolarizing ramp stimulus for all channels examined in this study (n= 6–14). A significant (P < 0.05) increase in percentage peak current is denoted with a single asterisk (*), whereas a significant (P < 0.05) decrease in percentage peak current is symbolized with a double asterisk (**).

Download figure to PowerPoint

image

Figure 6. Comparison of kinetic profiles for development and recovery from inactivated (OSI and CSI) states for V1300F mutant and WT channels A, development of CSI at 60 mV for V1300F and WT channels was examined using a two-step pulse protocol shown within the inset. B, recovery from CSI was tested experimentally using a two-step pulse protocol shown within the inset to determine the fraction of WT and mutant channels that recover at 90 mV. C, recovery from OSI was evaluated using a two-step protocol with a strong conditioning prepulse to 0 mV followed by a time-varied hyperpolarizing pulse (90 mV) and a test pulse (0 mV) to determine the fraction of channels recovering from OSI.

Download figure to PowerPoint

Voltage-dependent slow-inactivation properties for PEPD mutant channels are altered in a manner distinct from V1300F mutant channels

In addition to transitioning to a fast-inactivated state, VGSCs can transition to an additional inactivated state called slow-inactivation, which occurs over seconds or even minutes, and slowly recovers (for review, see Vilin & Ruben, 2001). As evidence of the physiological importance of slow-inactivation, it has been shown that mutations altering the fraction of channels accumulating in a slow-inactivated state in response to changes in voltage may play an important role in determining cellular excitability in skeletal muscle (Cummins & Sigworth, 1996; Bendahhou et al. 2002) and computer-modelled DRG neurons (Sheets et al. 2007). The effects of the Nav1.7 PEPD mutations on transition to a slow-inactivated state have not been determined. Therefore, we examined the voltage-dependent properties of slow inactivation for WT and each of the mutant channels using an extended (10 s) conditioning pulse followed by a brief hyperpolarizing pulse that allowed for recovery of fast-inactivated channels, before testing for the available population of channels (Fig. 7). Interestingly, all of the PEPD mutant channels decreased the voltage-dependent transition to a slow-inactivated state when compared to WT. Results for the PEPD mutant channels were in contrast to the non-PEPD mutant channel (V1300F), which did not decrease transition to the slow-inactivated state compared to WT. Both WT and V1300F channel availability were fitted well with a double-phase Boltzmann distribution, suggesting that the population of channels transitioned between two slow-inactivated states. However, the PEPD mutant channel transition to slow-inactivation was fitted well with a single-phase Boltzmann distribution. Therefore, it is possible that in addition to their role in impairing fast inactivation, the D3/S4–S5 mutations implicated in PEPD also affect the voltage-dependent transition to a slow-inactivated state.

image

Figure 7. Comparison of the derived voltage-dependent slow-inactivation properties for WT and mutant channels during transition between distinct states A, all D3/S4–S5 (V1298F, V1299F, and V1300F) and D3–D4 (I1461T) linker mutant channels were tested experimentally to determine the fraction of channels transitioning to slow-inactivated states in response to changes in voltage. To examine this transition we used a pulse protocol that included a +10 mV stepwise conditioning pulse from 130 mV to +10 mV for 10 s, and then quickly hyperpolarized cells to 120 mV to allow rapid recovery of fast-inactivated channels before testing for available channels for 20 ms at +10 mV. WT and V1300F channels were fitted using a double-phase Boltzmann distribution whereas PEPD mutant channels were fitted using a single-phase Boltzmann distribution. I–V data points to determine fraction of channels conducting in response to changes in voltage. D, steady-state inactivation for WT and mutant channels determined using the protocol displayed within the inset from a holding potential of 80 mV.

Download figure to PowerPoint

Discussion

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

VGSCs are believed to play important roles in pain (Cummins et al. 2007). Single-point mutations within highly conserved regions of Nav1.7 have been implicated in painful disorders (for review, see Koopmann et al. 2006; Cummins et al. 2007). It has been demonstrated that many of these mutations can preferentially alter particular voltage-dependent gating properties. All of the IE mutations studied to date shift the voltage dependence of activation in the hyperpolarizing direction. Although it has been proposed that all PEPD mutations might selectively cause marked deficits in fast inactivation (Fertleman et al. 2006), only three of the eight known PEPD mutations had been functionally characterized. Therefore, we examined the consequences of three PEPD mutations: the I1461T mutation studied by Fertleman et al. (2006) and two uncharacterized mutations identified within the cytosolic S4–S5 linker of D3, proximal to D3/S4. We demonstrate that the V1298F and V1299F mutations in Nav1.7, implicated in the painful inherited neuropathy PEPD (Fertleman et al. 2006), significantly shift the voltage-dependent transition to a fast-inactivated state in the depolarizing direction and also induce relatively small changes to the voltage-dependent conductance properties. Our data show these changes are likely to involve destabilization of closed and open fast-inactivated states, compared to WT channels. The effects of the V1298F and V1299F mutations on channel inactivation were similar to that of the I1461T PEPD mutation, located within the putative inactivation gate of the D3–D4 cytosolic linker of Nav1.7. As discussed below, this could have important implications for our understanding of the pathophysiology of PEPD. In contrast to the effects caused by the PEPD mutations, a non-PEPD mutation (V1300F) within the D3/S4–S5 linker, adjacent to the V1298F and V1299F mutations (Fig. 1A), had very different effects on Nav1.7 gating. Taken together, these data suggest that (1) the D3/S4–S5 linker of Nav1.7 is a key determinant in stability of the inactivated state, (2) specific residues within this linker play an integral role in stability based on their location and orientation, and (3) alteration of Nav1.7 steady-state inactivation properties by mutations involved in the pathophysiology of PEPD can be more moderate than previously indicated.

Functional consequences of PEPD mutations

Two molecular neuropathic pain syndromes, IE and PEPD, are caused by mutations in Nav1.7, and the consequences of these mutations are distinct. A majority of the SCN9A mutations implicated in IE are found within transmembrane segments and cytoplasmic linkers of D1 and D2 in Nav1.7. D1 and D2 IE mutant channels, expressed in hEK293 cells, cause a lowering of VGSC activation threshold, slowing of deactivation, and increased ramp current amplitudes between −60 and −40 mV (Cummins et al. 2004; Choi et al. 2006). DRG neurons transfected with Nav1.7 channels containing IE mutations increase neuronal excitability (Dib-Hajj et al. 2005; Rush et al. 2006) and computer simulations indicate the hyperpolarizing shift in activation is a major determinant of this hyperexcitability (Sheets et al. 2007). Fertleman et al. (2006) identified eight missense mutations (in 11 families) in SCN9A in patients with PEPD and seven of the identified mutations lie within the highly conserved cytoplasmic linkers, including the putative inactivation gate of D3 and D4 in Nav1.7. Initial functional analysis of three recombinant PEPD mutant channels expressed in hEK293 cells revealed that mutations within the putative inactivation gate and the D4/S4–S5 linker caused major deficits in fast inactivation (Fertleman et al. 2006), leading to a proposal that all PEPD mutations might result in a loss of inactivation.

We investigated whether two of the D3/S4–S5 PEPD mutations (Fig. 1A and B) had similar effects on voltage-dependent gating properties of Nav1.7 channels utilizing whole-cell voltage-clamp electrophysiological techniques. Both D3/S4–S5 PEPD mutations (V1298F and V1299F) significantly shifted the V1/2 of steady-state inactivation by ∼20 mV in the depolarizing direction compared to WT. This was nearly identical to the effect of the I1461T mutation on steady-state fast inactivation we observed. However, our data differ from those of Fertleman et al. (2006), which indicated the I1461T mutation impaired fast inactivation to a much greater extent than we observed. In Fertleman et al. it was shown that ∼40% of the I1461T current failed to inactivate during a steady-state inactivation protocol. Conversely, we observed nearly complete steady-state inactivation with the PEPD mutants (Fig. 3C). One reason for the differences observed in steady-state inactivation is that the protocol used in Fertleman et al. employed 60 ms inactivating prepulses, which, as can be seen in Fig. 4A, is inadequate to obtain steady-state conditions. Nav1.7 channels exhibit slow kinetics for the development of closed state inactivation (Cummins et al. 1998) and therefore 500 ms conditioning pulses are more appropriate to ensure steady-state conditions over the full-range of potentials tested. Our data are consistent with Fertleman et al. in that mutations implicated in PEPD alter the voltage-sensitivity of the fast-inactivated state, resulting in a decreased fraction of channels transitioning to a non-conducting state at potentials between −90 and −20 mV. However, because our data show that PEPD mutations do not substantially increase the non-inactivating component under true steady-state inactivating conditions (with both fluoride containing and non-fluoride containing electrode solutions), this indicates the impairment of fast inactivation by PEPD mutations can be more moderate than previously suggested.

Our data also show that the D3/S4–S5 PEPD mutations, as well as the I1461T mutation, increase the inactivation time constants (τh) compared to WT at potentials positive to −10 mV. This indicates rate transitions for inactivation of mutant channels become less voltage dependent at potentials positive to −10 mV thus, providing further evidence that the PEPD mutations destabilize the inactivated configuration. Upon inspection of the mutant I–V traces at −10 mV, we observed a small persistent component (∼6% of peak) with the three PEPD mutant traces compared to WT (Fig. 3E). By contrast, Fertleman et al. reported that short depolarizing pulses elicited relatively large persistent components for I1461T currents. A mutation that induces non-inactivating components would be expected to have major consequences on the neuronal excitability of every cell that the mutation is expressed in and would therefore be predicted to cause more widespread pain than is typically observed in many individuals with PEPD. Our data, demonstrating that the impairment of inactivation caused by the V1298F, V1299F and I1461T PEPD mutations can be more moderate than previously indicated for the I1461T mutation, help explain why PEPD mutations do not cause pain throughout the body.

We further evaluated the effects the PEPD mutations had on the stability of the inactivated state by testing channel development and recovery from OSI and CSI. All three PEPD mutations increased the recovery rates for CSI and OSI. Surprisingly, the V1298F and V1299F mutations within the D3/S4–S5 linker increased the rate of development of CSI, while the I1461T mutation, within the putative inactivation gate, did not. However, although the rate for development of CSI at −60 mV was increased, the fraction of channels undergoing CSI at −60 mV was greatly reduced for the three PEPD mutant channels transfected, compared to WT channels, which is likely to reflect disruption of strong binding of the inactivation gate by the PEPD mutations. Since it has been demonstrated that variability of CSI kinetics between different VGSC subtypes can influence Iramp amplitudes (Cummins et al. 1998), we examined currents elicited by a slow (0.27 mV ms−1) depolarizing ramp stimulus. We observed a significant increase in the Iramp for all three PEPD mutant channels at potentials positive to −20 mV when compared to WT, which is likely to be due to the decreased rate of OSI at these potentials. While both IE and PEPD mutations are likely to increase ramp current amplitudes, the voltage dependence of the ramp currents observed with the PEPD mutant channels are distinct from the ramp currents observed with several of the Nav1.7 IE mutant channels, which exhibit increased ramp current amplitudes at negative potentials but not at potentials near −10 mV compared to WT channels (Cummins et al. 2004). As such, the PEPD mutations may differentially alter action potential properties when compared to the IE mutations. The PEPD mutations are likely to contribute to broadening of the action potential duration and enhance repetitive or burst firing, but probably do not decrease the threshold for action potential firing and depolarize the resting membrane potential in the same manner as IE mutations (Dib-Hajj et al. 2005; Rush et al. 2006). We also show that PEPD mutations can alter the voltage dependence of slow inactivation of Nav1.7. The impairment of slow inactivation that we observe at potentials between −90 and −60 mV with all three of the PEPD mutations examined in this study is also likely to increase Nav1.7 channel availability near resting potential and therefore contribute to enhanced excitability of sensory neurons.

Structural role of the Nav1.7 D3/S4–S5 cytosolic linker in channel gating

Our data provide additional insight into the role of the D3/S4–S5 cytosolic linker in VGSC gating. Specific regions within each of the four domains (D1–D4) of VGSCs are thought to have distinct, but integrated, roles in channel gating and conformational stability in response to changes in membrane potential. Significant advances in understanding the structural interactions involved in voltage-sensitive channel gating have been demonstrated through the determination of the structure of potassium channels using X-ray crystallography (Doyle et al. 1998; Long et al. 2005a,b, 2007). The modelled crystal structure of the Kv1.2 channel shows the α-helical S4–S5 segments crossing over hydrophobic regions of the S6 segments. It has been proposed that the distal portion of the S6 segments need to pivot or splay during the activating transition to the ion-conducting state and that the voltage-dependent outward displacement of the S4 segments is coupled to channel opening via hydrophobic interactions of the S4–S5 and S6 segments. A crystal structure has not been obtained for mammalian VGSCs. While homology modelling of VGSCs based on potassium channel structures provides some insight, study of disease-associated and laboratory-designed mutations has enhanced our knowledge of the secondary structure and critical features involved in gating of VGSCs (Hille, 2001). Several studies have indicated that residues within the D3/S4–S5 linker (1) act as a part of the unique docking site for the VGSC putative inactivation gate via direct interactions and/or (2) indirectly stabilize the fast-inactivated configuration via cooperative interactions with specific residues located in the D4/S4–S5 cytosolic linker as the inactivation gate (IFMT) docks at a site within the channel pore (Smith & Goldin, 1997; Popa et al. 2004). Our data demonstrate that V1298 and V1299 are important to help stabilize the fast-inactivated states of Nav1.7. Our data on the adjacent and identical mutation V1300F within D3/S4–S5 of Nav1.7 indicate that orientation and/or position-specific interactions are important. In contrast to V1298F and V1299F, the V1300F mutation had very small effects on inactivation, shifting the V1/2 of steady-state inactivation by 4 mV in the hyperpolarizing direction and decreased the percentage of peak current elicited during a slow ramp depolarization compared to WT channels. The V1300F mutation had a more pronounced effect on activation, shifting the V1/2 of channel conductance by ∼10 mV in the depolarizing direction and decreasing the voltage-dependent time constants for deactivation. The distinct effects of adjacent, identical mutations are consistent with the hypothesis that the D3 and D4/S4–S5 linkers of VGSCs retain an α-helical secondary structure (Filatov et al. 1998). These studies are important because they suggest that mutation within the D3 and D4/S4–S5 linkers may, in an isoform-specific manner, predictably disrupt critical stabilizing interactions that decrease the overall affinity of gating reactions, which may offer insight on structural requirements for gating. Thus, our studies confirm an important structural and functional role for D3/S4–S5 of Nav1.7 during stabilization of the inactivated states, such that mutation of residues critical during transition decreases the voltage-dependent probability of channels residing in this configuration.

The aim of this research was to determine the effects that two PEPD mutations within D3/S4–S5 had on voltage-dependent gating properties of Nav1.7. We demonstrate that D3/S4–S5 mutations identified in patients with the inherited molecular neuropathy PEPD result in destabilization of the fast and slow-inactivated states of Nav1.7, thus contributing to depolarizing shifts in the steady-state inactivation profiles and enhancing ramp currents. It is worth noting that at least 11 additional D3/S4–S5 VGSC mutations have been associated with inherited disorders of excitability, including epilepsy (Heron et al. 2002; Berkovic et al. 2004; Meisler & Kearney, 2005), ataxia (Kohrman et al. 1996), long QT syndrome type 3 (Wang et al. 1996; Smits et al. 2005), brugada syndrome (Casini et al. 2007), myotonia (Yang et al. 1994; Richmond et al. 1997; Bouhours et al. 2005), and hypokalaemic paralysis (Sugiura et al. 2003), indicating this region may be a ‘hotspot’ for disruptive mutation. These disorders are a result of single-point mutations occurring in a highly conserved region of various VGSC isoforms. Interestingly, some of these disease-related mutations predominately alter the voltage dependence of channel conductance. Although many of these mutations slow the rate of inactivation, none of them have been shown to substantially increase the percentage of non-inactivating current. Here, our studies show that PEPD mutations also do not necessarily cause loss of fast inactivation. The moderate decrease in the stability of the fast-inactivated configuration we observe might help explain why the pain associated with PEPD mutations in Nav1.7 is typically not associated with all nociceptive neurons expressing Nav1.7.

References

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

Appendix

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

Acknowledgements

This work was supported by National Institutes of Health Research Grant NS053422 (TRC).

Supporting Information

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  8. Supporting Information

Supplemental material, for publication (Table 1, Figures 1-2)

Please note: Blackwell Publishing are not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.

FilenameFormatSizeDescription
TJP_3016_sm_Tab1Figs1-2.pdf548KSupporting info item

Please note: Wiley Blackwell is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.