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Copyright © 1999 The Biophysical Society. All rights reserved.
Biophysical Journal, Volume 76, Issue 2, 861-868, 1 February 1999

doi:10.1016/S0006-3495(99)77249-8

Channels, Receptors, and Transporters

Enhanced Slow Inactivation by V445M: A Sodium Channel Mutation Associated with Myotonia

Masanori P. Takahashi^* and Stephen C. Cannon^*, #, ,  

^* Department of Neurology, Massachusetts General Hospital, Boston, MA 02114 USA
^# Department of Neurobiology, Harvard Medical School, Boston, MA 02114 USA

Address reprint requests to Dr. Stephen C. Cannon, EDR 413, Massachusetts General Hospital, Boston, MA 02114. Tel.: 617-724-3531; Fax: 617-726-3926.

Voltage-gated Na channels are the primary determinant of excitability in skeletal and heart muscle and in neurons. Excitability is strongly dependent on intrinsic gating properties of Na channels, and inactivation—a decline in Na current despite a maintained membrane depolarization—is a universal feature of all voltage-gated Na channels. Fast inactivation operates on a time scale of milliseconds and alters the availability of Na channels over the time course of a single action potential. Slow inactivation occurs on a time scale of seconds to minutes and may modulate excitability in response to slow shifts in the resting potential (Chandler and Meves, 1970,Almers et al,Ruff et al). Defects of inactivation have recently been identified in mutant human Na channels that cause disorders of skeletal muscle (Cannon, 1998) or heart (Ackerman, 1998). In this study we have identified a novel enhancement of slow inactivation by V445M, a missense mutation associated with myotonia that is predicted to lie in the S6 segment of domain I (Rosenfeld et al).

More than 20 missense mutations in the α subunit of the adult human skeletal muscle Na channel (hSkM1) are known to cause several heritable muscle diseases including hyperkalemic periodic paralysis (HyperPP), paramyotonia congenita (PMC), and potassium-aggravated myotonia (PAM). The functional consequences of these mutations have been investigated in order to understand the pathophysiological basis for the enhanced excitability in myotonia and the inexcitability during attacks of periodic paralysis. Fast inactivation is partially disrupted by every mutation tested to date (Cannon, 1997) and a subset of mutations also cause a hyperpolarized shift in activation (Cummins et al,Mitrovic et al,Green et al,Plassart-Schiess et al). In model simulations these functional defects are sufficient to cause the repetitive discharges that give rise to myotonia and may cause paralysis in the more severely disrupted mutants by a depolarization-induced loss of excitability (Cannon et al,Hayward et al). Slow inactivation has recently been recognized as an additional determinant of the pathophysiology of these disorders (Ruff, 1994,Cannon, 1996,Cummins and Sigworth, 1996). Mutations that impair slow inactivation and alter fast-gating transitions occur in families in which weakness is a prominent feature (HyperPP). This association and model simulations both suggest that slow inactivation may normally protect muscle from prolonged depolarized shifts in the resting potential caused by a persistent Na current conducted by Na channels with altered fast-gating (Cummins and Sigworth, 1996,Hayward et al).

A novel missense mutation was recently identified in a family with recurrent attacks of painful myotonic stiffness but no episodic weakness (Rosenfeld et al). Valine 445, which lies in S6 of domain I and is conserved in the Na channels of most species from jellyfish to humans, was mutated to methionine. None of the other 20 missense mutations in hSkM1 associated with diseases of skeletal muscle occurs in domain I. We have transiently expressed V445M in mammalian cells and have detected changes in gating behavior. In agreement with a preliminary study by Bennett et al, we observed a mild disruption of fast inactivation and a small hyperpolarized shift of activation. More importantly, we also identified a pronounced enhancement of slow inactivation produced by an impediment to recovery (hyperpolarized shift in the voltage dependence of recovery) and a faster entry rate (∼1.6-fold). This is the first example of a disease-related mutation that enhances slow inactivation and has implications for the pathogenesis of myotonia and weakness. Moreover, V445 is the second site in IS6 that augments slow inactivation when mutated. Wang and Wang, 1997 showed that N434A in rat SkM1 (equivalent to N440A in hSkM1) speeds entry and slows recovery from slow inactivation. Although the molecular mechanism of slow inactivation remains unknown, these studies provide new evidence that IS6 has an important role in slow inactivation.

The adult isoform of the human skeletal muscle sodium channel α-subunit, hSkM1 (George et al), and the mutant V445M were provided by Al George, Jr. These cDNAs were subcloned between the NotI and XbaI sites of the mammalian expression vector pRc/CMV. The human β subunit cDNA (McClatchey et al) was subcloned into the EcoRI site of the mammalian expression vector pcDNAI (Invitrogen, San Diego, CA).

Culture of human embryonic kidney (HEK) cells and their transient transfection were performed as described previously (Hayward et al). In brief, plasmid DNAs encoding wild-type (WT) or mutant human Na channel α subunits (0.9μg/35-mm dish), the human Na channel β subunit (fourfold molar excess over α subunit DNA), and a CD8 marker (0.175μg) were cotransfected by the calcium phosphate method. At 1–3 days after transfection, the HEK cells were trypsinized briefly and passaged to 22-mm round glass coverslips for electrophysiological recording. Individual transfection-positive cells were identified by labeling with anti-CD8 antibody cross-linked to microbeads (Dynal, Great Neck, NY) (Jurman et al).

Na currents were measured using conventional whole-cell recording techniques as described previously (Hayward et al). Recordings were made with an Axopatch 200A amplifier (Axon Instruments, Foster City, CA). The output was filtered at 5kHz and digitally sampled at 40kHz using an LM900 interface (Dagan, Minneapolis, MN). Data were stored to a 486-based computer using a custom AxoBasic (Axon Instruments, Foster City, CA) data acquisition program. More than 80% of the series resistance was compensated by the analog circuitry of the amplifier and the leakage conductance was corrected by digital scaling and subtraction of the passive current elicited by a 20-mV depolarization from the holding potential. Cells with peak currents of<1 nA or>20 nA upon step depolarization from −120mV to −10mV were excluded. After initially establishing whole-cell access, we often observed leftward shifts in the voltage dependence of gating, an increase in the size of the peak current, and a decrease in the amplitude of persistent Na current. To minimize these effects, we waited at least 10min for equilibration after gaining access to the cells.

Patch electrodes were fabricated from borosilicate capillary tubes with a multistage puller (Sutter, Novato, CA). The shank of the pipette was coated with Sylgard and the tip was heat-polished to a final tip resistance (in bath solution) of 0.5–2.0 MΩ. The pipette (internal) solution contained 105mM CsF, 35mM NaCl, 10mM EGTA, and 10mM Cs-HEPES (pH 7.4). Fluoride was used in the pipette to prolong seal stability. The bath contained 140mM NaCl, 4mM KCl, 2mM CaCl₂, 1mM MgCl₂, 5mM glucose, and 10mM Na-HEPES (pH 7.4). Recordings were made at room temperature (21–23 C°). Tetrodotoxin (TTX) was purchased from Sigma (St. Louis, MO).

Curve fitting was performed manually off-line using AxoBasic, SigmaPlot (Jandel Scientific, San Rafael, CA), or Origin (Microcal, Northhampton, MA). Conductance was calculated as G(V)=I_peak(V)/(V−E_rev), where the reversal potential, E_rev, was measured experimentally for each cell. Steady-state fast and slow inactivation were fitted to a Boltzmann function with a nonzero pedestal, I_o, calculated as I/I_peak=(1−I_o)/[1+exp((V−V_1/2)/k)]+I_o, where V_1/2 is the half-maximum voltage and k is the slope factor (Table 1). Symbols with error bars indicate means±SEM. Statistical significance was determined by the unpaired t-test with p values noted in text.

The kinetics of Na channel gating were characterized by recording whole-cell currents from HEK cells transiently transfected with cDNAs encoding WT or mutant (V445M) hSkM1, and the human isoform of the β subunit. Bennett et al have reported, in abstract form, that fast inactivation is altered by V445M. We confirmed many of their observations, as briefly described below.

The voltage dependence of steady-state fast inactivation, h_∞(υ), was measured with a 100-ms conditioning pulse, in order to minimize the effect of entry to the slow inactivated state (see below). The relative peak currents elicited at −10mV were fit with a Boltzmann function, and the estimated parameter values are listed in Table 1. Mutant V445M had a −5mV leftward (hyperpolarized) shift in the midpoint (p<0.0001), whereas there was no difference in the steepness.

The amplitude of the persistent Na current, I_ss was evaluated by measuring the TTX-sensitive component with a subtraction protocol. The current elicited by a 10-ms step depolarization from −120mV to −10mV in the presence of 5μM TTX was subtracted from that in normal external solution. The amplitude of steady-state current during the last 0.2ms of the pulse was averaged and normalized to the peak value. I_peak V445M channels had an increased persistent current at 10ms (0.70%) compared to WT (0.05%).

The voltage dependence of activation was measured by applying step depolarizations from −120mV. The Na conductance was estimated from the peak current and the measured reversal potential, E_rev, as G(V)=I_peak(V)/(V−E_rev). The conductance data were fit with a Boltzmann function and the estimated parameters are listed in Table 1. The V445M mutant had a −4mV leftward (hyperpolarized) shift in the midpoint (p<0.002).

Thus far, our observations on the effects of V445M on fast-gating are in agreement with those reported by Bennett et al. In addition, Bennett et al found that recovery from inactivation was profoundly slowed for V445M. This was unexpected because other myotonia-associated mutations in hSkM1 accelerated the recovery from fast inactivation, whereas a sluggish recovery rate reduces the excitability of the cell. In their recovery protocol, Bennett et al used a 500-ms conditioning pulse, which would cause a significant degree of slow, as well as fast, inactivation. We used a series of conditioning pulse durations to separate the effects of V445M on fast and slow inactivation.

Recovery from inactivation was tested using the three-pulse protocol shown in the inset of Figure 1A. Cells were held at −120mV, and a conditioning pulse to −10mV was applied for a preset duration. The conditioning pulse was followed by recovery at −120mV for 0.05 to 10,000ms. Peak Na current was measured in response to a subsequent test depolarization to −10mV. Channel availability was measured as the ratio of the peak current during the test depolarization to that during the reference pulse. Traces in Figure 1A show example data for WT and V445M channels obtained with 300ms conditioning pulses. The I_Na elicited at −10mV after recovery times of 0.1, 3, 15, 70, and 10,000ms were normalized to the peak of reference current and superimposed. Within 15ms, 65% of the current recovered in WT channels but only 50% was available for V445M channels. The time course of recovery at −120mV, after a series of conditioning pulses to −10mV, is shown for WT and V445M in Figure 1B. Solid lines show fits to a two-exponential relaxation for visual guidance. For brief (30-ms) conditioning pulse intervals, more than 90% of the current for both WT and V445M channels recovered within 15ms (inset, Figure 1B), indicating that the rapid monoexponential recovery from fast inactivation was indistinguishable for the two channel types. With longer conditioning pulse intervals (300 and 3000ms), however, the time course of recovery of V445M channels was slower than that of WT. Recovery followed a multi-exponential time course after these prolonged conditioning pulses due to slow inactivation of channels. The sluggishness of the V445M channels was caused by a higher proportion of channels being slow-inactivated (lower amplitude of the fast component of recovery, Figure 1B inset) and by a slower recovery rate from the slow-inactivated state (Fig. 1 B). These data suggest that V445M alters slow inactivation, rather than a slowing of the recovery from fast inactivation suggested by Bennett et al.

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Figure 1

Recovery from inactivation was slowed for V445M channels. (A) A three-pulse protocol (inset) was used to measure recovery from inactivation as the fraction of available I_Na. Traces show currents for wild-type and V445M channels after a 300-ms conditioning pulse at −10mV. I_Na elicited at −10mV, after recovery times of 0.1, 3, 15, 70, and 10,000ms, was normalized to the peak of reference current (12.4 nA for WT and 16.9 nA for V445M). (B) Relative current from WT and V445M channels is plotted as a function of the duration of recovery at −120mV. Smooth curves are χ² minimized fits with double-exponential relaxation to averaged data and are shown for visual guidance.

The voltage dependence of entry to slow inactivation was tested using the three pulse protocol shown in the inset of Fig. 2. Cells were held at −120mV, and a conditioning pulse with varying duration was applied. Channel availability was measured as the ratio of the peak current during the test depolarization to that during the reference pulse. Between the reference and conditioning pulse the cell was held at −120mV for 500ms. Three different conditioning potentials (−90, −70, and −40mV) were tested. The conditioning and test pulses were separated by a 20-ms hyperpolarization to −120mV to allow recovery from fast inactivation. The kinetics of entry to slow inactivation at different membrane potentials are shown in Fig. 2. The entry to slow inactivation for both the WT and V445M channels was quicker and more complete at more depolarized potentials. For the V445M channels, however, slow inactivation developed more quickly than for WT channels. The time course of entry was fit to a single-exponential for each cell. The means of the estimated parameter values were used to generate curves in Fig. 2 and are plotted in Figure 5C. The time constant of WT channels at −90mV is not shown in Figure 5C, because a reliable fit was difficult due to the small amplitude of the decay (see Fig. 2).

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Figure 2

Entry to slow inactivation was enhanced for V445M channels. Entry to slow inactivation was measured as the peak I_Na elicited at −10mV, after a conditioning pulse of variable duration (inset). Current amplitudes were normalized to the reference peak current elicited from a holding potential of −120mV. The conditioning and test pulses were separated by a 20-ms hyperpolarization to −120mV to allow channels to recover from fast inactivation. Symbols denote different conditioning voltages, and the smooth curves are generated with mean parameter values from χ² minimized fits of a single-exponential relaxation plus a constant to the data of each cell.

Data from the entry to slow inactivation protocol also enabled us to define the voltage dependence of steady-state slow inactivation, S_∞. As shown in Fig. 2, the extent of entry to slow inactivation approached a constant value within 60s. The voltage dependence of S_∞ is shown for WT and V445M mutant channels in Fig. 3. Steady-state slow inactivation was measured using a 60-s prepulse, followed by a 20-ms gap at −120mV to allow recovery from fast inactivation, before the −10mV test pulse (inset). The voltage dependence of slow inactivation in V445M channels was steeper, shifted to the left (hyperpolarized) and more complete than WT. The data in Fig. 3 were fitted by a single Boltzmann plus a constant term, and estimated parameter values were listed in Table 1.

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Figure 3

Steady-state slow inactivation was augmented for V445M channels. Steady-state slow inactivation was measured as the peak I_Na elicited at −10mV after a 60-s conditioning pulse (inset). Current amplitudes were normalized to the reference peak current elicited from a holding potential of −120mV. The conditioning and test pulses were separated by a 20ms hyperpolarization to −120mV to allow channels to recover from fast inactivation. Smooth curves show χ² minimized fits of the data by a Boltzmann function plus a constant term; WT: V_1/2=−64.2±1.0mV, k=7.9±0.5mV, S₀=0.16±0.04; V445M: V_1/2=−73.6±1.4mV, k=6.6±0.5mV, S₀=0.08±0.01 .

The voltage dependence of recovery from slow inactivation was measured for WT and V445M channels at −80, −100, and −120mV. The voltage protocol is shown in the inset of Fig. 4. In this protocol, the holding and recovery potentials were identical so that the reference I_Na and test I_Na were elicited from the same starting potential. The recovery from slow inactivation in both the WT and V445M channels was quicker and more complete at more hyperpolarized potentials. For the WT channels, however, recovery from slow inactivation was faster than for V445M channels. A quantitative comparison was made by fitting the time course of the recovery data to a single exponential. Because this protocol measures the total recovery (both nonslow and slow inactivated channels), a fractional offset was used in the exponential fit. The fraction of channels not slow inactivated was set equal to the mean of the S_∞ data at −10mV (0.15 for WT and 0.07 for V445M). The smooth curves were generated with mean parameter values from fitting the data of each cell to a single-exponential relaxation. This single-exponential approximation for recovery was suboptimal, as discussed previously (Hayward et al), and a two- or three-exponential approximation was more accurate. We employed a single-exponential approximation, however, because it is representative of the majority of the current recovery and allows a single parameter comparison. The mean values of the time constant at −120, −100, and −80mV, obtained from fits to data from 4 to 9 cells, are plotted in Fig. 5 C.

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Figure 4

Recovery from slow inactivation was slowed for V445M channels. Relative current from WT and V445M channels is plotted as a function of recovery duration on a logarithmic time axis. Inset shows the voltage protocol. Smooth curves are drawn using mean parameter values obtained from χ² minimized fits of a single exponential plus a constant (to account for the rapid recovery of channels not slow inactivated) to the data for each cell.

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Figure 5

The enhanced slow inactivation for V445M was due primarily to a reduced rate of recovery, produced by a left (hyperpolarized) shift in β(V). (A) The entry (α) and recovery (β) rates were computed from the steady-state, S_∞, and relaxation, τ_s, data as: α=(1−S_∞)/τ_s) and β=S_∞/τ_s. The voltage dependencies of the rates were fit to exponential functions that saturated at depolarized potentials: α(V)=α_o/(1+e^−(V−V_α^)/k_α) and β(V) =β_o (1+e^−(V−V_β^)/k_β). The parameter values were WT: α_o=0.60s⁻¹, V_α=−52.5mV, k_α=8.6mV, β_o=0.11s⁻¹, V_β=−75.0mV, k_β=9.6mV; V445M: α_o=0.99s⁻¹, V_α=−53.3mV, k_α=9.0mV, β_o=0.086s⁻¹, V_β=−84.9mV, k_β=10.9mV. Based on these fits for α(V) and β(V), the calculated fraction of channels not slow inactivated (S_∞=β/(α+β), lines in B) and the calculated time constant (τ_s=1/(α+β), lines in C) are comparable to the data.

To define further the mechanism by which V445M enhances slow inactivation, we examined the combined kinetic and steady-state behaviors in relation to a two-state model for slow inaction:

The rates of entry (α) and recovery (β) were computed from the relaxation time constant, τ_s, and the fraction of channels not slow inactivated, S_∞, as

To assess whether the differences in slow inactivation between WT and V445M channels might alter the availability of Na channels in a use-dependent manner, we measured the peaks of Na currents elicited during a train of depolarizations. 20-ms depolarizations to −10mV were applied at a frequency of 10Hz from a holding potential of −90mV. The amplitude of the peak I_Na measured during each pulse was normalized by the amplitude of first pulse. The mean of the normalized amplitudes from 8 WT cells and 6 V445M mutants is plotted as a function of the pulse number within a train (Fig. 6). The use-dependent reduction in peak I_Na was more pronounced for V445M than WT channels. The augmented use-dependent inhibition for V445M channels is attributable to differences in slow inactivation because recovery from fast inactivation was complete within 80ms at −90mV for both WT and mutant V445M channels (data not shown).

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Figure 6

Use-dependent reduction of I_Na was greater for V445M than WT channels. Sodium currents were elicited by depolarizations to −10mV from a holding potential of −90mV. The 20-ms depolarizations were applied at a rate of 10Hz. The peak current amplitude of each pulse was normalized with respect to the amplitude of the first pulse and plotted against the pulse number.

The gating behavior of heterologously expressed human Na channels was compared for WT and V445M, a missense mutation in the S6 segment of domain I that causes a dominantly inherited from of myotonia. Functional characterization of V445M is of particular interest because none of the other 20 missense mutations in hSkM1 associated with myotonia or periodic paralysis occur in domain I and very few site-directed mutations have been studied in this region. In agreement with a preliminary report by Bennett et al, we found that the gating of rapid transitions was altered by V445M in a manner that would increase excitability. The persistent Na current was increased, and the G(V) curve was left-shifted. The left shift of the h_∞(V), however, is predicted to reduce the tendency for repetitive myotonic discharges. Like Bennett et al, we also observed a slowed recovery from inactivation for V445M channels. Our study differs, however, in that we conclude there is no significant difference in recovery from fast inactivation between WT and V445M channels (cf. Fig. 1, 30-ms conditioning pulse). The difference in recovery occurred because slow inactivation was augmented by V445M. At depolarized potentials, slow inactivation was accelerated and was more complete for V445M mutants. Repolarization after prolonged depolarization showed that V445M channels recover about twofold more slowly than WT. The steady-state voltage-dependence of slow inactivation for V445M mutants was more complete and was shifted toward hyperpolarized potentials. In a two-state gating scheme, these changes can be reconciled by a large (10mV) hyperpolarized shift in the voltage-dependence of the recovery rate and a modest (∼ 1.6-fold) increase in the rate of entry to slow inactivation.

Slow inactivation of the skeletal muscle Na channel has recently been recognized as an important determinant in the propensity for periodic paralysis. Ruff, 1994 proposed that a defect of slow inactivation must occur in hSkM1 mutations that cause depolarization-induced periodic paralysis. Otherwise, slow inactivation would shut off the aberrant Na current produced by impaired fast inactivation and the muscle would repolarize. Indeed, slow inactivation is partially disrupted in the two most commonly occurring mutations found in families with HyperPP: T704M (Cummins and Sigworth, 1996,Hayward et al) and M1592V (Hayward et al). However, other hSkM1 mutations associated with HyperPP (M1360V) or with PMC in which prolonged episodes of weakness may occur (T1313M, R1448C) have no detectable alteration in slow inactivation (Hayward et al,Richmond et al). Conversely, defects in slow inactivation have never been identified in functional studies of hSkM1 mutants associated with pure myotonia without weakness (Hayward et al,Richmond et al). Based on these results and a model simulation, Hayward et al suggested that a defect of slow inactivation increases the likelihood of paralysis but is not a necessary condition. A corollary is that intact slow inactivation might protect against attacks of paralysis. V445M is the only disease-associated mutation of hSkM1 found to enhance slow inactivation. The observation that patients with V445M have painful disabling myotonia, presumably caused by the observed impairment of fast inactivation and left shift of activation (Bennett et al), but never have attacks of episodic weakness (Rosenfeld et al) is consistent with our proposed role for slow inactivation. The enhanced slow inactivation of V445M is predicted to reduce the risk of depolarization-induced attacks of weakness, but is still too sluggish to prevent transient runs of repetitive discharges that give rise to myotonic stiffness.

The mechanism by which Na channels slow inactivate and the critical regions of the protein for this process remain unknown. V445M is the second instance of a missense mutation in a Na channel α subunit that enhances slow inactivation. Wang and Wang, 1997 reported that slow inactivation is enhanced by a nearby residue in IS6, N434A, in the rat isoform of SkM1 (corresponding to N440 in human SkM1). In contrast, divergent effects on fast gating were observed for mutations in this region. V445M shifted the G(V) relation by −4mV whereas rN434A caused a +24mV shift, and V445M mildly slowed the rate of fast inactivation (increased the limiting value of τ_h for strong depolarizations) whereas rN434A accelerated fast inactivation nearly twofold. The divergent effects on fast gating processes and the comparable enhancement of slow inactivation implies the cytoplasmic end of IS6 is important for slow inactivation. In addition, the S6 segment is known to contribute to various forms of slow inactivation in other voltage-gated channels. C-type inactivation of Shaker K channels is strongly influenced by mutations in S6 (Hoshi et al,Boland et al). In Ca channels, variations in inactivation kinetics have been attributed to differences in residues within IS6 or in the flanking extracellular and cytoplasmic regions (Zhang et al).

An augmentation of slow inactivation has also been observed when fast inactivation is severely disabled, either by internal proteases (Rudy, 1978) or missense mutations within the III-IV loop (Featherstone et al). The enhanced slow inactivation observed for V445M and rN434A is not likely to be a consequence of altered fast inactivation for several reasons. First, a nearly complete abolition of fast inactivation is required for significant enhancement of slow inactivation. Second, for channels with abolished fast inactivation the augmentation of slow inactivation occurs solely by an increased rate of entry (Rudy, 1978), whereas missense mutations in IS6 dramatically slow the rate of recovery from slow inactivation. Third, the accelerated entry to slow inactivation was several times faster for rN434A than WT channels, after fast inactivation was abolished in both channel types by treatment with chloramine-T. Fourth, we have recently demonstrated that movement of the fast inactivation gate is not tightly coupled to slow inactivation (Vedantham and Cannon, 1998).

Several other regions of the Na channel have been shown to influence slow inactivation. Mutations at the cytoplasmic end of IIS5 (rT698M) or IVS6 (rM1585V) partially disrupt slow inactivation (Cummins and Sigworth, 1996,Hayward et al). Taken together, the studies on disease-associated missense mutations suggest that a conformational change at the inner vestibule of the pore (cytoplasmic ends of IS6, IIS5, and IVS6) might occur during slow inactivation. On the other hand, slow inactivation is resistant to cytoplasmic application of proteases (Rudy, 1978). Other data have implicated the extracellular face of the channel. Townsend and Horn, 1997 demonstrated that slow inactivation of cardiac Na channels is impeded by elevated extracellular levels of alkali metal cations but not by larger organic cations, suggesting that cation binding near the outer mouth of the pore inhibits closing of the slow inactivation gate. Finally, the voltage-sensing segments are also thought to influence slow inactivation. Slow inactivation appears to be coupled to activation (Ruben et al), and a mutation in IIS4 of the rat brain IIa channel (L860F) disrupts the slow mode of inactivation in the oocyte expression system (Fleig et al).

In contrast to fast inactivation, no mutation or physiochemical manipulation has been identified that completely abolishes slow inactivation. This failure may be an ascertainment bias, since fast inactivation is studied more easily and more commonly. A more likely possibility is that slow inactivation does not arise from closure of a single gate or hinged lid. A more global conformational change, involving distant regions of the primary α subunit structure, appears to occur. Whatever the mechanism, our data and that of Wang and Wang, 1997 clearly demonstrate that the IS6 region must play a role in slow inactivation.