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J Physiol (2003), 548.1, pp. 85-96
© Copyright 2003 The Physiological Society
DOI: 10.1113/jphysiol.2002.033084

Negative charges in the DIII-DIV linker of human skeletal muscle Na⁺ channels regulate deactivation gating

James R. Groome*†, Esther Fujimoto† and Peter C. Ruben†

	ABSTRACT

Top Abstract Introduction Methods Results Discussion References

Charge reversing, neutralizing and substituting mutations at D1309 and EE1314,15 in the DIII-DIV linker of the human skeletal muscle sodium channel hNa_v1.4 were constructed and expressed in Xenopus oocytes. The effects of these mutations on conductance, inactivation and deactivation were determined using on-cell macropatches. D1309R caused a depolarizing shift of the conductance-voltage (g(V)) curve and increased the apparent valency of activation. D1309R and EE1314,15RR increased time to peak activation. D1309R caused a depolarizing shift of the steady-state fast inactivation curve, whereas EE1314,15RR produced a hyperpolarizing shift and decreased the apparent valency. Charge reversal at either D1309 or EE1314,15 slowed open-state fast inactivation and accelerated closed-state fast inactivation. D1309R accelerated recovery from fast inactivation, whereas EE1314,15RR and EE1314,15QQ slowed recovery. Deactivation from the inactivated state was determined by the delay in the onset to recovery from fast inactivation. Recovery delay was abbreviated for D1309R but was prolonged for EE1314,15RR and EE1314,15QQ. Open-state deactivation was determined from the time constant of the decay (_D) of tail currents. _D was slowed by D1309R, D1309E, EE1314,15RR and EE1314,15QQ. Our findings suggest an important role in deactivation gating in hNa_v1.4 for the negative cluster of charge at EE1314,15. These and previous findings suggest that clusters of negatively and positively charged residues in the hNa_v1.4 DIII-DIV linker differentially regulate the kinetics of fast inactivation.

(Received 23 September 2002; accepted after revision 16 January 2003; first published online 14 February 2003)
Corresponding author P. C. Ruben: Department of Biology, Utah State University, Logan, UT 84322-5305, USA. Email: pruben{at}biology.usu.edu

	INTRODUCTION

Top Abstract Introduction Methods Results Discussion References

Voltage-gated Na⁺ channels form the pathway for inward ionic current that underlies the rising phase of action potentials in most excitable cells (Catterall, 2000). Membrane depolarization increases Na⁺ permeability through voltage-dependent activation of Na⁺ channels. Subsequent fast inactivation limits the duration of the action potential (Hodgkin & Huxley, 1952), and prolonged depolarization leads to one or more forms of slow inactivation (Vilin & Ruben, 2001). Deactivation from the open state also limits the duration of the action potential (Featherstone et al. 1998) whereas deactivation from the inactivated state is requisite for the re-opening of channels (Kuo & Bean, 1994). Since Na⁺ channels are generally available for activation only from the closed state, recovery from inactivation and deactivation of open or inactivated channels are critical determinants of channel availability (Rayner et al. 1993; Kuo & Bean, 1994). Excitability is therefore regulated by the stability of the inactivated state(s) and the rates at which channels close. Although several structural determinants of fast and slow inactivation have been identified, less is known about Na⁺ channel structures that dictate deactivating transitions from inactivated or open states.

The amino acid sequence of the Na⁺ channel subunit predicts four homologous domains containing six membrane-spanning segments (Noda et al. 1984). Positively charged residues in the fourth segment (S4) of each domain (DI to DIV) are found at regularly spaced intervals along the proposed helix, a feature that predicts their function as voltage sensors (Stühmer et al. 1989; Yang & Horn, 1995; Yang et al. 1996; Horn et al. 2000). Na⁺ channel S4 segments exhibit asymmetry of charge content (Trimmer et al. 1989). Not surprisingly, domain-specific roles for S4 segments have been proposed for activation (Kontis et al. 1997; Mitrovic et al. 1998), deactivation (Kontis et al. 1997; Groome et al. 1999, 2002), fast inactivation (Chahine et al. 1994; Chen et al. 1996; Kontis & Goldin, 1997; Sheets et al. 1999; Horn et al. 2000), slow inactivation (Mitrovic et al. 2000) and charge immobilization (Cha et al. 1999).

Na⁺ channels deactivate from the inactivated or open states. Inactivated-state deactivation (I to C) is a silent transition that appears as a brief delay preceding the recovery from fast inactivation in Na⁺ channels (Kuo & Bean, 1994). Open-state deactivation (O to C) is observed as a decay in tail current following a brief depolarization of the channel. Both transitions involve voltage-dependent translocations of the S4 segments (Groome et al. 2000). However, the two forms of deactivation show markedly different kinetics, such that recovery delay (I to C deactivating transition) is approximately one order of magnitude slower than tail current decay (O to C deactivating transition). These findings suggest that the structural determinants of deactivation comprise domain-specific translocations of the S4 segments that are influenced by factors in addition to the transmembrane electric field.

The cytoplasmic linker between DIII and DIV in Na⁺ channels acts as a 'hinged lid' to block the permeation pathway. Fast inactivation is critically dependent on a tripeptide IFM motif in this linker (Vassilev et al. 1988; West et al. 1992; Kellenberger et al. 1996; Horn et al. 2000). In addition to the IFM inactivation particle, 14 positively charged residues and four negatively charged residues exist in the DIII-DIV linker of Na_v1.4 (George et al. 1993). This region of the channel thus contains a concentration of charged residues similar to that found in the S4 segments, a fact that intuitively suggests a role for linker charges in voltage-dependent channel gating. Although the results of previous studies have been equivocal as to the necessity of charge in the DIII-DIV linker for activation and fast inactivation (Moorman et al. 1990; Patton et al. 1992), a role for linker charges in Na⁺ channel function is suggested by the finding that the kinetics of fast inactivation are significantly accelerated by a reduction of positive charge in this region of the channel (Moorman et al. 1990; Miller et al. 2000).

The role of DIII-DIV linker charges in deactivation gating has not previously been investigated. Therefore, we sought to determine whether charged residues in the linker contribute to the structural determinants of deactivation. Specifically, we tested the hypothesis that, given the proximity of D1309 and EE1314,15 in hNa_v1.4 to the inactivation particle, negatively charged residues in the linker regulate deactivation from the inactivated state but not from the open state. To test this hypothesis, we compared gating properties in hNa_v1.4 to charge reversing, neutralizing and substituting mutations at D1309 and at the EE1314,15 cluster. Our results suggest that mutations at D1309 allosterically alter Na⁺ channel deactivation, whereas charge content at EE1314,15 appears to be an important determinant of deactivation from both the open and inactivated states. Mutations at these loci of negative charge also influence the kinetics of fast inactivation from open and closed states, as well as the recovery of channels from fast inactivation. Some of these results have been reported in abstract form (Groome et al. 2001).

	METHODS

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Site-directed mutagenesis

All mutations were prepared by site-directed mutagenesis using a PCR overlap extension method (Ho et al. 1989) with the appropriate primers containing the mutation. For IFM/QQQ, a 1.2 kb PCR-prepared, Tth111 I-Sac II fragment was cloned into hNa_v1.4/sp64T. For D1309R, D1309Q, D1309E, EE1314,15RR, EE1314,15QQ and EE1314,15DD, a 930 bp Avr II-Sac II fragment was cloned into hNa_v1.4/pgh19. Mutations were verified by sequencing.

Oocyte preparation

Xenopus laevis oocytes were surgically removed from adult frogs anaesthetized with 0.17 % tricaine (3-aminobenzoic acid ethyl ester; Sigma, St Louis, MO, USA) as approved by NIH guidelines and by the Institutional Animal Care and Use Committee at Utah State University. Frogs were humanely killed after the final oocyte collection.

Oocytes were separated using 2 mg ml^-1 collagenase (Sigma) in a solution containing (mM): NaCl 96, KCl 2, MgCl₂ 20 and Hepes 5, pH 7.4. Oocytes were incubated at 18 °C in culture medium containing (mM): NaCl 96, KCl 2, MgCl₂ 1, CaCl₂ 1.8, Hepes 5, sodium pyruvate 2.5, pH 7.4, with 100 mg l^-1 gentamicin and 3 % horse serum (Gibco BRL, Rockville, MD, USA). One day after oocytes were isolated, mRNA for and 1 subunits (1:1 volume, subunit at 1 µg µl^-1, 1 subunit at 3 µg µl^-1) was injected at a total volume of 50 nl oocyte^-1. Prior to recording, vitelline membranes were manually removed with forceps after a 5 min treatment in a hyperosmotic solution containing (mM): NaCl 96, KCl 2, MgCl₂ 20, Hepes 5 and mannitol 400, pH 7.4.

Electrophysiology

All recordings were from cell-attached macropatches (Featherstone et al. 1998). Pipette solution was (mM): NaCl 96, KCl 4, MgCl₂ 1, CaCl₂ 1.8 and Hepes 5, pH 7.4. Bath solution was (mM): NaCl 9.6, KCl 88, EGTA 11 and Hepes 5, pH 7.4. This solution was designed to zero the membrane potential prior to seal formation, such that voltage applied to cell-attached macropatches was thus the actual membrane potential. Voltage clamping and data acquisition were carried out as previously described (Featherstone et al. 1998) using an EPC-9 patch-clamp amplifier (HEKA, Lambrecht, Germany) controlled via Pulse software (HEKA) running on a Power Macintosh G3 or G4. Data were acquired at 5 µs per point and low-pass filtered at 5 kHz during acquisition. Bath temperature was maintained at 15 ± 0.1 °C for all experiments with a Peltier device and HCC-100A temperature controller (Dagan, Minneapolis, MN, USA). Oocyte holding potential was -120 to -150 mV, and leak subtraction was automatically performed using a P/4 protocol. Leak and capacitance subtractions were done upon seal formation and corrected before each voltage-clamp experiment. Analysis and graphing were done using PulseFit (HEKA) and Igor Pro (Wavemetrics, Lake Oswego, OR, USA).

Conductance-voltage (g(V)) relationships were derived using the equation:

g_Na = I_max/(V_M - E_Na),

where g_Na is Na⁺ conductance, I_max is measured as peak current of the test pulse, V_M is the test pulse voltage and E_Na is the measured Na⁺ equilibrium potential. Activation and steady-state fast inactivation (h) curves were fitted by a Boltzmann distribution, as:

I/I_max = 1/(1 + exp(-ze₀(V_M - V_1/2)/kT)),

where I/I_max (normalized current amplitude) was measured during a variable-voltage test pulse or prepulse from a holding potential of -150 mV, V_M is the test pulse potential, z is the apparent valency, e₀ is the elementary charge, V_1/2 is the midpoint voltage, k is the Boltzmann constant and T is temperature in K.

Time constants for entry into fast inactivation were derived from fitting the monoexponential decay of current following activation from a holding potential of -150 mV to voltages from -40 to +20 mV. Time constants of closed-state fast inactivation were obtained by fitting the decay in peak current amplitude evoked with test pulses to -20 mV following prepulses of varying duration to voltages from -100 to -50 mV. Recovery from fast inactivation was determined using a double-pulse protocol (Kuo & Bean, 1994). A 21 ms depolarizing voltage step to 0 mV was used to inactivate channels. This was followed by a command to voltages that ranged between -190 and -90 mV for durations that ranged from 0.05 to 5 ms in steps of 50 µs. For some mutations, interpulse duration was increased to 10 ms at interpulse voltages more depolarized than -130 mV to more accurately determine recovery. The interpulse was followed by a 5 ms recovery test pulse to 0 mV. Time constants of recovery were obtained by fitting the increase in peak current amplitude evoked with these test pulses. Fast inactivation time constants were calculated using the equation:

I = I_ss + a₁exp(-t/),

where I is current amplitude, I_ss is the steady-state current (asymptote), a₁ is the amplitude at time zero and is the time constant.

Descriptions of fast inactivation were further characterized by assuming a first-order, two-state model of the fast inactivation reaction (not inactivated ™ inactivated) and fitting versus voltage curves to the equation:

(V_M) = 1/(k_f + k_b),

where (V_M) represents the time constant of progression to equilibrium of fast inactivation as a function of membrane potential, k_f is the rate of the forward reaction (not inactivated inactivated) and k_b is the rate of the reverse reaction (inactivated not inactivated). Reaction rates were determined according to the equations:

k_f = Aexp + ([z(1 - )(V_M - V_1/2)]/kT),

and

k_b = Aexp - ([z(V_M - V_1/2)]/kT),

where A is the half-rate at V₀, z is the (apparent) total reaction valency (in electronic charge), is the fractional barrier distance, V_M is membrane potential (in mV), V_1/2 is the midpoint potential (in mV), k is the Boltzmann constant and T is temperature (in K).

Descriptions of inactivated-state deactivation rates were derived from measurements of the delay in the onset to recovery from fast inactivation. Recovery current amplitudes were extrapolated with a single-exponential function to the time (t₀) at which current amplitude was zero, which was taken as recovery delay. Descriptions of open-state deactivation rates, given as time constants (_D), were derived with a single-exponential fit of the decay of tail currents. These were evoked by 50 mV, 0.25 ms depolarizations followed by command hyperpolarizations from -140 to -70 mV. In other experiments tail currents were evoked by depolarizations at voltages ranging from -40 to +20 mV for 0.5 ms, followed by command hyperpolarizations as described above.

Statistical analyses were performed using Instat (Graph Pad Software, San Diego, CA, USA). All results are reported as means ± S.E.M. Statistical significance was determined using Student's t tests or, in those cases where there was a statistically significant difference between standard deviations, Welch's alternative t tests. Statistical significance of difference was accepted at P 0.05. For recovery and deactivation kinetics, t tests of mutations at individual voltages were used to compare differences over the voltage range tested. Significant differences of mutations for at least three consecutive voltages were taken as an indication of a significant difference in voltage dependence. Statistical comparisons at individual voltages were also noted where significant differences were not continuous over several voltages.

	RESULTS

Top Abstract Introduction Methods Results Discussion References

Effects of mutations on activation and fast inactivation

Charge reversing, neutralizing and substituting mutations were made at D1309 and at the EE1314,15 cluster in hNa_v1.4. Each mutation was tested for effects on conductance, and kinetics of inactivation and deactivation. Families of Na⁺ currents evoked in response to step depolarizations from -90 to +60 mV from a holding potential of -150 mV are shown in Fig. 1. An examination of current decay following these mutations suggested that fast inactivation was not severely impaired. These results are consistent with earlier findings suggesting that negatively charged residues in the DIII-DIV linker do not constitute part of the inactivation particle (Patton et al. 1992; Kellenberger et al. 1997).

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Figure 1. Na+ currents for hNav1.4 and following mutations in the DIII-DIV linker Channels were held at -150 mV prior to step depolarizations of 20 ms duration over a voltage range from -90 to +60 mV; averaged sweeps are shown for hNav1.4 and charge-neutralizing mutations. Calibration: 5 ms, 1 nA for EE1314,15QQ; 500 pA for hNav1.4, EE1314,15RR, D1309R and D1309Q; and 300 pA for EE1314,15DD.

To determine whether D1309 or EE1314,15 participate in other aspects of Na⁺ channel gating, we first characterized equilibrium properties of mutations at these loci of negative charge. We then used protocols to isolate open-state and closed-state fast inactivation, recovery from fast inactivation, and deactivation from the inactivated and open states.

Conductance and steady-state parameters

Activation (g(V)) and steady-state fast inactivation (h) parameters of hNa_v1.4 and following mutations at D1309 and EE1314,15 are shown in Fig. 2 and Fig. 3, and summarized in Table 1. The midpoint of activation (V_1/2) was right-shifted by D1309R (Fig. 2). The apparent valency (z) of the g(V) curve was increased by D1309R and by EE1314,15QQ. Apparent valency was also increased by the respective charge substitutions at these loci, indicating that these effects were allosteric. Time to peak activation was significantly increased by D1309R, with charge substitution at this residue producing a lesser increase. Time to peak activation was increased by EE1314,15RR and was decreased by charge substitution of these residues.

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Figure 2. Conductance-voltage relationships for hNav1.4 and following mutations in the DIII-DIV linker Channels were depolarized from a holding potential of -150 mV with 20 ms step depolarizations from -90 to +60 mV. The charge reversing mutation D1309R produced a significant depolarizing shift of the activation midpoint.

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Figure 3. Steady-state fast inactivation (h) curves for hNav1.4 and following mutations in the DIII-DIV linker Channels were subjected to 500 ms prepulses at voltages ranging from -170 to -35 mV prior to a 0 mV depolarizing test pulse. A depolarizing shift was observed with charge reversal at D1309 and a hyperpolarizing shift was observed for charge reversal at EE1314,15. The dashed line through the data for hNav1.4 is included for clarity.

The midpoint of the h curve (h_1/2) was right-shifted by D1309R and by D1309E (Fig. 3), whereas a significant left shift of h was observed in EE1314,15RR. The h curve for EE1314,15DD was similar to that for hNa_v1.4. Apparent valency (z) of the h curve for D1309R was similar to that for hNa_v1.4 but was decreased with neutralization of this residue, and was increased by charge substitution. EE1314,15RR decreased the apparent valency of the h curve, but z was not affected by charge substitution in EE1314,15DD. Thus, the effects of mutations at D1309 on steady-state fast inactivation were allosteric whereas the effects of mutations at EE1314,15 were a consequence of alterations in charge content.

Fast inactivation

The rate of open-state fast inactivation was determined from the monoexponential decay of current following activation from a holding potential of -150 mV to voltages ranging from -40 to +20 mV. Time constants of open-state fast inactivation (_h) are shown in Fig. 4. D1309R and D1309E increased _h at voltages more positive than -50 and -40 mV, respectively, indicating that the effect of D1309R on _h was allosteric. EE1314,15RR and EE1314,15QQ increased _h at voltages more positive than -40 and -20 mV, respectively. However, _h for EE1314,15DD was similar to that for hNa_v1.4 over the voltage range tested, indicating that mutations at EE1314,15 altered _h as a consequence of alterations in charge content. The voltage dependence of _h was slightly decreased by charge reversal at D1309 and to a greater extent by charge reversal at EE1314,15.

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Figure 4. Voltage dependence of the kinetics of entry into the fast-inactivated state Time constants of fast inactivation from -40 to +20 mV are shown for hNav1.4 and following mutations at D1309 (A) and at the EE1314,15 cluster (B). Charge reversal at either loci slowed open-state fast inactivation.

Closed-state fast inactivation was measured by the decline in peak current elicited by a depolarizing test pulse following a variable duration, conditioning prepulse (Fig. 5). Closed-state fast inactivation was accelerated by D1309R from -100 to -70 mV and by D1309Q from -100 to -90 mV. In contrast, closed-state fast inactivation was slowed by D1309E from -90 to -70 mV. Closed-state fast inactivation was accelerated by EE1314,15RR from -90 to -60 mV. Time constants of closed-state fast inactivation for EE1314,15QQ and for EE1314,15DD were similar to that for hNa_v1.4. Closed-state inactivation appeared to be less complete for D1309R than for hNa_v1.4, and more complete for EE1314,15RR (Fig. 5). Differential effects of charge reversal at D1309 and EE1314,15 on the completion of closed-state inactivation are consistent with their relative effects on steady-state fast inactivation; D1309R (less complete closed-state fast inactivation) produced a depolarizing shift in h whereas EE1314,15RR (more complete closed-state fast inactivation) produced a hyperpolarizing shift in h (Fig. 3).

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Figure 5. Time course of entry into the fast-inactivated state from the closed state for hNav1.4 and following mutations at D1309 and EE1314,15 Channels were depolarized by prepulses to voltages ranging from -100 to -50 mV for varying durations up to 30 ms. Channel availability at the end of each prepulse was tested by a second depolarization to -20 mV, and the time course of the decrease in evoked current was taken as the kinetics of the C to I transition.

Recovery from fast inactivation was determined using a double-pulse protocol as described in Methods. Recovery currents are shown in Fig. 6 and normalized recovery curves are shown in Fig. 7. Recovery was accelerated for D1309R from -180 to -100 mV, but was slowed for D1309Q and for D1309E from -190 to -170 mV. In contrast to the effects of charge reversal at D1309, recovery was slowed for EE1314,15RR from -190 to -120 mV, and for EE1314,15QQ from -190 to -170 mV and at -130 mV. Recovery was slowed in EE1314,15DD from -140 to -130 mV.

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Figure 6. Double-pulse protocol used to measure recovery rate and delay for hNav1.4 and following mutations of negatively charged residues in the DIII-DIV linker For each trace, every tenth sweep of the 21 ms, 0 mV depolarizing pulse used to inactivate channels and the second 5 ms, 0 mV depolarizing pulse used to assess recovery are shown. The dashed vertical line indicates the end of the first depolarizing pulse for all sweeps. Calibration: 5 ms, 1 nA for hNav1.4, EE1314,15QQ; 500 pA for EE1314,15RR, EE1314,15DD and D1309R.

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Figure 7. Time course of the recovery from fast inactivation from experiments shown in Fig. 6 Charge reversal at D1309 accelerated channel recovery, whereas charge reversal at EE1314,15 slowed the recovery from fast inactivation.

Figure 8 summarizes the effects of mutations at D1309 and at EE1314,15 on fast inactivation, expressed as time constants of recovery from and entry into the fast-inactivated state, obtained from the experiments described in Figs 4-7. Fits to curves assuming first-order, two-state reaction kinetics were used to calculate the parameters shown in Table 2. Time constants used to generate these fits thus represent the progression towards equilibrium of the inactivation reaction, and are dependent upon the state of the channel (inactivated, closed or open) prior to the tests for fast inactivation recovery or onset.

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Figure 8. Fast inactivation kinetics for hNav1.4 and following mutations in D1309 (A) and EE1314,15 (B) A first-order, two-state reaction model of inactivated - not inactivated was used to generate fits (shown by dashed lines) from time constants of inactivation determined from protocols isolating fast inactivation recovery, and closed-state and open-state fast inactivation.

Charge reversal at D1309 had a depolarizing effect on the inactivation reaction midpoint, and decreased the maximum time constant. The effect of D1309 on the reaction midpoint appeared to be due in part to reversal of charge since the reaction midpoint for D1309E was similar to that for hNa_v1.4, whereas the reduction in the maximum time constant by D1309R was an allosteric effect; D1309E increased this parameter with respect to hNa_v1.4. Charge reversal at EE1314,15 decreased the maximum time constant and reduced the fractional barrier distance; both effects appeared to be due in part to a reversal of charge in the EE1314,15RR mutant since each of these parameters for EE1314,15DD were similar to those for hNa_v1.4.

Deactivation from the fast-inactivated state

Recovery currents for hNa_v1.4 and mutants obtained from experiments shown in Fig. 6 and Fig. 7 were used to calculate the delay in the onset to recovery from fast inactivation, as a measure of the rate of the I to C transition. Figure 9 shows the voltage dependence of recovery delay for hNa_v1.4 and linker mutants. Recovery delay was abbreviated by D1309R at all voltages tested (Fig. 9A). In contrast, recovery delay was increased by D1309Q at -190 to -140 mV, at -120 and at -90 mV. Delay was increased by D1309E at -170 to -150 mV but was similar to that for hNa_v1.4 at other voltages. Recovery delay was increased by EE1314,15RR at all voltages tested (Fig. 9B). Delay was increased by EE1314,15QQ at all voltages except -100 mV. EE1314,15DD produced recovery delays similar to that of hNa_v1.4 at most voltages.

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Figure 9. Recovery delays in hNav1.4 and following mutations of negative charges in the DIII-DIV linker Voltage dependence of the delay in the onset of recovery from fast inactivation for hNav1.4 (dashed line is included for clarity) and following mutations at D1309 (A) and at the EE1314,15 cluster (B) in the DIII-DIV linker. Delay was determined as the x intercept of the recovery curve obtained from the double-pulse protocol.

Open-state deactivation

The tail currents shown in Fig. 10 were evoked by 50 mV, 0.25 ms depolarizations followed by command hyperpolarizations from -140 to -70 mV. Tail currents completely decayed in hNa_v1.4, with linker charge mutations, and with the mutation IFM/QQQ. Fast inactivation for IFM/QQQ is completely abolished (Featherstone et al. 1998) and (complete) tail current decay is assumed to represent open-state deactivation. We found that tail currents for IFM/QQQ decayed completely at voltages more negative than -60 mV. Therefore, single-exponential fits to the decay of tail currents observed during command hyperpolarizations from -140 to -70 mV were used to measure the kinetics of channel closure from the open state.

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Figure 10. Tail currents for hNav1.4 and following mutations of negative charges in the DIII-DIV linker Tail currents were evoked by command hyperpolarizations at voltages from -140 to -70 mV following a 50 mV depolarizing pulse for 0.25 ms. Calibration: 500 µs, 500 pA for hNav1.4, EE1314,15QQ, EE1314,15DD; 1 nA for EE1314,15RR and D1309R.

Figure 11 shows the voltage dependence of _D in hNa_v1.4 and mutants during command hyperpolarizations following a 50 mV depolarization for 0.25 ms. IFM/QQQ increased _D compared to hNa_v1.4 at all voltages. _D was significantly increased for D1309R and for D1309E and was similar to that of hNa_v1.4 for D1309Q. _D was increased for EE1314,15RR and for EE1314,15QQ whereas EE1314,15DD decreased _D compared to hNa_v1.4 at voltages more positive than -90 mV.

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Figure 11. Deactivation time constants as a function of command hyperpolarization voltage Voltage dependence of D for hNav1.4 (dashed line is included for clarity), IFM/QQQ (A) and following mutations at D1309 (B) and EE1314,15 (C). D was calculated from single-exponential fits to the decay of tail currents evoked by command hyperpolarizations from -140 to -70 mV, following a depolarizing pulse of 50 mV for 0.25 ms.

Our observation that the IFM/QQQ mutation increased _D suggests that the structural determinants of fast inactivation and open state deactivation may not be independent. We found that charge reversal at D1309 or at EE1314,15 slowed fast inactivation and prolonged tail current decay. To investigate the possibility that tail currents were slowed by these mutations due to their effects on fast inactivation, we compared tail currents for hNa_v1.4, D1309R and EE1314,15RR evoked by hyperpolarizing commands that followed 0.5 ms depolarizations to voltages ranging from -40 to +20 mV. These 'conditioning depolarizations' comprised the same voltage range used to measure _h, as shown in Fig. 4. Tail currents evoked by command hyperpolarizations at -120, -100 and -80 mV using this protocol are shown in Fig. 12. In hNa_v1.4, the rate of tail current decay was not steeply dependent on the voltage of the conditioning depolarization used to open channels over the voltage range of -40 to +20 mV. Tail currents were similar to that of hNa_v1.4 for D1309R and for EE1314,15 with command hyperpolarizations of -120 mV (Fig. 12A). However, charge reversal at either loci produced increases in _D with command hyperpolarizations at -100 mV (Fig. 12B) or -80 mV (Fig. 12C). Thus, charge reversal at D1309 and at EE1314,15 slowed tail currents in protocols using conditioning depolarizations for which the rate of fast inactivation was also slowed by these mutations (Fig. 4).

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Figure 12. Deactivation time constants as a function of prepulse depolarization voltage Kinetics of tail current decay for hNav1.4, D1309R and EE1314,15RR for command hyperpolarizations at -120 mV (A), -100 mV (B) and -80 mV (C) in protocols in which the depolarization used to open channels was varied over the same range as that used to assess open-state fast inactivation. More depolarized conditioning depolarizations increased D for charge reversing mutations at D1309R and EE1314,15 but not for hNav1.4.

	DISCUSSION

Top Abstract Introduction Methods Results Discussion References

Our findings suggest that deactivation from either the inactivated or open state is regulated by the cluster of negative charge at EE1314,15 in the DIII-DIV linker. Charge content at D1309 does not appear to be an important determinant of deactivation, and the effects of mutations at either loci of negative charge on open-state deactivation may be an indirect consequence of the effects of mutations on fast inactivation. Our results are consistent with earlier studies which show that negatively charged residues in the DIII-DIV linker of Na⁺ channels are not required for fast inactivation (Patton et al. 1992; Kellenberger et al. 1997). Complete inactivation was observed during depolarizing test pulses and the steady-state inactivation curve was not affected by mutations at D1309 and EE1314,15. However, these negative charges in the hNa_v1.4 DIII-DIV linker play a role in regulating the rate of entry into and recovery from the fast-inactivated state.

Open-state fast inactivation

Mutations at two loci of negative charge (D1309, EE1314,15) in the DIII-DIV linker of hNa_v1.4 produced significant effects on the rate of entry into the fast-inactivated state. Open-state fast inactivation was slowed by charge reversals at D1309 and at EE1314,15. The effects of D1309R were primarily a consequence of structural alteration, since charge substitution, but not charge neutralization, at D1309 slowed fast inactivation. Charge neutralizations at the analogous residue D1487 in rat Na_v1.4 (rNa_v1.4) have little effect on open-state fast inactivation (Kellenberger et al. 1997), which supports the conclusion that charge content at this residue does not regulate the kinetics of open-state fast inactivation. The effects of mutations at EE1314,15 to slow fast inactivation were consistent with this being a consequence of altering negative charge, since charge reversal and neutralization at this cluster, but not substitution, increased _h.

Our findings, together with results from studies of the effects of mutations of small clusters of positive charge in the DIII-DIV linker in Na_v1.4, support the hypothesis that the charge polarity of residues in this region of the channel determines their effect on the kinetics of open-state fast inactivation. We found that charge reversal or neutralization at EE1314,15 slowed the entry of channels into the fast-inactivated state. In rNa_v1.4, charge neutralizations at KK1317,18 and at KKK1322,23,26 decrease _h (Miller et al. 2000). More rapid entry of channels into the fast-inactivated state is also observed with neutralization of KK1317,18 in hNa_v1.4 (analogous to KK1310,11 in rNa_v1.4; J. R. Groome, unpublished observations). Taken together, these data suggest that the negative charge cluster at EE1314,15 in hNa_v1.4 promotes entry into the fast-inactivated state whereas nearby positive charges retard entry into the fast-inactivated state.

Deactivation

We observed a close correlation between charge content at EE1314,15 and deactivation kinetics. Recovery delay and tail current decay were each prolonged by reversal or neutralization of charge at EE1314,15, whereas deactivation from either state was unaffected by charge substitution. These findings suggest that one or each of the glutamate residues near the IFM motif in hNa_v1.4 comprise or interact with structural determinants of deactivation in Na⁺ channels.

Deactivation from the inactivated state

Recovery delay reflects the deactivating transition from the inactivated to the closed state in Na_v1.4 (Groome et al. 2000). Fast-inactivated channels undergo charge immobilization that must be overcome for channels to become available (Armstrong & Bezanilla, 1977), a finding that may underlie the observation that recovery delay (inactivated-state deactivation) exhibits slower kinetics than the tail current (open-state deactivation). Several immobilizable residues have been identified, including the outermost and central residues in DIVS4 (Cha et al. 1999; Kühn & Greef, 1999). Consistent with those findings, recovery delay is abbreviated by neutralization of the outermost (R1) and central (R5, R6) residues in DIVS4 (Groome et al. 2002). In the present study, we found that an increase in positive charge at EE1314,15 prolonged recovery delay. Thus, charged residues in DIVS4 and in the DIII-DIV linker appear to be important determinants of inactivated-state deactivation.

Open-state deactivation

Charge content at EE1314,15 dictated the effect on tail currents elicited by command hyperpolarizations following brief depolarizations. Deactivation from the open state was slowed by charge reversal or neutralization at EE1314,15, whereas charge substitution of this cluster produced a deactivation profile similar to hNa_v1.4. These effects suggest a role for negative charge at EE1314,15 in the regulation of open-state deactivation. In contrast to a clear dependence on charge content at EE1314,15 for the effects on _D, the effects of mutations at D1309 on _D were allosteric. Thus, charge substitution (D1309E) produced effects on _D identical to those observed for D1309R, suggesting that charge content at this residue does not play an important role in open-state deactivation in hNa_v1.4.

Are the structural determinants of fast inactivation and deactivation interactive or independent?

Tail current decay was slowed in a charge-dependent manner by mutations at EE1314,15, and allosterically by mutations at D1309. Both open-state deactivation and open-state fast inactivation were slowed by charge reversal and neutralization at EE1314,15 and by charge reversal at D1309R. We used the IFM/QQQ mutation to isolate the voltage dependence of deactivation from fast inactivation in tail current decay (Featherstone et al. 1998). The finding that mutagenic removal of fast inactivation increases _D in hNa_v1.4, as it does in rNa_v1.4 (Groome et al. 2002), suggests that fast inactivation may contribute to the rate of tail current decay. An increase in _D following removal of fast inactivation by mutagenesis is also observed with enzymatic removal of fast inactivation (Cota & Armstrong, 1989). Since tail current decay was increased for EE1314,15RR and D1309R following conditioning depolarizations which increase _h in these mutants, we cannot preclude the possibility that slowed deactivation with reduction of negative charge at either of these loci is the result of slowed fast inactivation.

Do negative linker charges interact with the inactivation particle?

D1309 and EE1314,15 flank the IFM(T) motif in hNa_v1.4, whose solution structure in rNa_v1.4 suggests that isoleucine and phenylalanine residues of the putative inactivation gate are solvent-exposed and thus available for interaction (Rohl et al. 1999). The proximity of the IFM(T) motif to the residues studied here raises the possibility that D1309 and EE1314,15 influence the kinetics of fast inactivation and deactivation through interactions with one or more of the residues in the inactivation particle. Allosteric effects of D1309R might be an indirect consequence of an alteration in the structural interactions of D1309 with I1310 or F1311. In rNa_v1.4, each of the glutamate residues in the cluster analogous to EE1314,15 in hNa_v1.4 are located in the proximity of the proposed inactivation surface (Rohl et al. 1999). Both the structure and polarity of the threonine residue of the IFM(T) motif in rNa_v1.4 (T1491) are important determinants in the stability of fast inactivation (Kellenberger et al. 1997). In rNa_v1.4, the paramyotonia congenita mutation T1313M produces effects on fast inactivation and deactivation similar to those reported here for mutations at EE1314,15 (Hayward et al. 1996). Thus, it is interesting to speculate that effects of EE1314,15RR reported here might be in part a consequence of an alteration in electrostatic interactions of E1314 or E1315 with T1313 in hNa_v1.4.

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Acknowledgements

We thank J. Repscher for help with experiments, and J. Abbruzzese for comments on a draft of this manuscript. This work was supported by a Harvey Mudd College faculty research grant to J.R.G., and by Public Health Service grant R01-NS29204 and a Research Grant from the Muscular Dystrophy Association to P.C.R.

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