Wang J et al. (1998), Regulation of deactivation by an amino terminal...

XB-ART-14002

J Gen Physiol 1998 Nov 01;1125:637-47. doi: 10.1085/jgp.112.5.637.

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Regulation of deactivation by an amino terminal domain in human ether-à-go-go-related gene potassium channels.

Wang J , Trudeau MC , Zappia AM , Robertson GA .

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Abnormalities in repolarization of the cardiac ventricular action potential can lead to life-threatening arrhythmias associated with long QT syndrome. The repolarization process depends upon the gating properties of potassium channels encoded by the human ether-à-go-go-related gene (HERG), especially those governing the rate of recovery from inactivation and the rate of deactivation. Previous studies have demonstrated that deletion of the NH2 terminus increases the deactivation rate, but the mechanism by which the NH2 terminus regulates deactivation in wild-type channels has not been elucidated. We tested the hypothesis that the HERG NH2 terminus slows deactivation by a mechanism similar to N-type inactivation in Shaker channels, where it binds to the internal mouth of the pore and prevents channel closure. We found that the regulation of deactivation by the HERG NH2 terminus bears similarity to Shaker N-type inactivation in three respects: (a) deletion of the NH2 terminus slows C-type inactivation; (b) the action of the NH2 terminus is sensitive to elevated concentrations of external K+, as if its binding along the permeation pathway is disrupted by K+ influx; and (c) N-ethylmaleimide, covalently linked to an aphenotypic cysteine introduced within the S4-S5 linker, mimics the N deletion phenotype, as if the binding of the NH2 terminus to its receptor site were hindered. In contrast to N-type inactivation in Shaker, however, there was no indication that the NH2 terminus blocks the HERG pore. In addition, we discovered that separate domains within the NH2 terminus mediate the slowing of deactivation and the promotion of C-type inactivation. These results suggest that the NH2 terminus stabilizes the open state and, by a separate mechanism, promotes C-type inactivation.

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Species referenced: Xenopus laevis
Genes referenced: gnao1 gnas kcnh2 sh2b3 tbx2

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	Figure 4. Deactivation rate in HERG channels increases with elevated external K+. (A) Scaled tail currents from WT channels in 5, 150, and 200 mM external K+. Currents were evoked at −100 mV subsequent to a 3-s step to 60 mV. (B) Deactivation time constants at various external K+ concentrations for both WT (n = 6) and Δ2-354 (n = 5) channels for tail currents evoked as described in A. Tail currents were fit with a single exponential function in this case because, at −100 mV, the fast component accounts for >90% of the current. (C) The change in deactivation rate (1/τ) with increasing external K+ exhibits a saturating dose–response behavior with a Hill coefficient of 5.36 ± 0.69. At −100 mV, the forward (activation) rate is negligible and most of the deactivating current is described by the fast deactivation τ (Table I) so the deactivation rate ≅ 1/τfast. As shown, deactivation rate increased in a dose-dependent manner with increased external K+ ions. These data were fitted with the Hill equation: ln[(1 − S′)/S′] = −n(lnK − lnL), where S′= S/S0 is the fraction of sites occupied by the ligand, S is the number of sites occupied, So is the total number of sites available, K is the dissociation constant of the ligand, L is the ligand concentration, and n is the Hill coefficient. We determine S′ using the formula (k − k5)/(k300 − k5), where k is the deactivation rate (1/τ at −100 mV) at any given external potassium concentration, k5 is the rate at 5 mM [K]o, and k300 the rate at 300 mM [K]o. The S′ values at different [K]o (i.e., L) were plotted as fractional increase of deactivation rate in C. The resulting sigmoidal dose–response curve was then fitted with the Hill equation indicated above to obtain the Hill coefficient.
	Figure 1. The HERG NH2 terminus slows deactivation but does not alter activation kinetics. (A) Families of tail currents from HERG wild-type (WT) and N deletion (Δ2-354) channels. Channels were allowed to fully activate and inactivate during a 3-s depolarizing pulse to 60 mV, only part of which is shown (Δ); during a subsequent step to potentials ranging from −120 to −40 mV (⋄), currents recovered from inactivation, and then deactivated. The holding potential was −80 mV. The zero current level is indicated by the dashed line. (B) Scaled tail currents evoked by a −100-mV repolarizing pulse from the experiment in A show that Δ2-354 channels deactivate faster than WT channels. (C) Fast (top) and slow (bottom) deactivation time constants obtained from double exponential fits to the closing phase of the tail currents plotted vs. repolarizing voltage values (see methods) for wild-type and Δ2-354 channels (n ≥ 10). Each point represents the mean, and the error bars are SEM for this and all subsequent figures. (D) Activation time constants for WT and Δ2-354 channels are plotted for different voltages. There is no significant difference between these values for WT and Δ2-354 channels (P ≥ 0.58 at confidence interval of 0.05 in one-way analysis of variance test). Top inset shows a series of currents evoked by the envelope of tails protocol to measure time-dependent increases in conductance, as described in methods, with a step to 40 mV (Trudeau et al., 1995; Wang et al., 1997); bottom inset shows exponential fit to resulting conductance vs. time plot (n = 6).
	Figure 2. The HERG NH2 terminus promotes C-type inactivation. (A) Inactivating current traces scaled to the peak current from WT and Δ2-354 channels generated using the three-pulse protocol: channels are allowed to fully activate and inactivate during a 3-s prepulse to 60 mV; a subsequent, brief repolarizing pulse to −100 mV allows channels to recover; near the peak of this tail current before deactivation becomes apparent, a third pulse to one of a range of voltages is given to drive channels from the open into the inactivated state (40 mV for the traces shown). The repolarizing pulse is 35 ms in duration for WT, 15 ms for Δ2-354. (Based on our fits, we calculate that recovery is 99.02 ± 0.72%, n = 14, complete for WT and 99.50 ± 0.29%, n = 8 complete for Δ2-354 at the peak of the tail current; i.e., end of recovery phase. This was determined by fitting the recovery phase of the tail currents with a single exponential of the form y = A0 + Ae−t/τ. The fit, which in most cases followed very closely to the peak currents, was extended beyond the peak to a plateau. The current level of the plateau was considered to represent the full recovery from inactivation. The amplitude of the peak tail current was normalized to this value to give the percentage of recovery, the mean values for which are reported above.) Thus, an inactivating current is measured in relative isolation from other kinetic components. Arrowheads point to the steady state inactivation level of WT and Δ2-354 channels. The dashed line indicates the zero current level. The capacitance artifact (∼2 ms) ends at the arrow. Capacitance artifacts are also shown at the beginning of the tail currents upon repolarization. (B) Currents from A scaled to peak and minimum to show the difference in time course. Time constants extracted from single exponential fits are shown along with the error of the fit. The arrow corresponds to the settling of the clamp as shown in A. (C) Inactivation time constants extracted from single exponential fits to the inactivating current illustrated in A are plotted at different voltages (n ≥ 10). (D) Change in free energy as a function of voltage shows that deletion of the NH2 terminus increases the free energy of the inactivated state relative to the open state (n = 6). The free energy change associated with open to inactivated transition can be calculated as ΔGO ↔ I = −RTlnKO ↔ I = −RTln(I/O) = RTln(O/I), where ΔG is the difference in free energy between the open and closed states. I and O are the fraction of channels in the inactivated and open states, respectively, R is the gas constant and T is the temperature. The fraction of open and inactivated channels are measured from the inactivation current as illustrated in the inset: the instantaneous outward current level, indicated as b, reflects the total number of channels that are open and ready to undergo the open to inactivated transition. The value for b is determined by fitting the inactivating current with a single exponential function and extrapolating back to the moment of the voltage change; the fit departs from the recorded current during the capacitive artifact, which has a duration of ∼2 ms. The steady state current level (the channels remaining in O) is indicated as a. Thus, the number of inactivated channels can be calculated as (b − a). Then, at any given voltage, ΔGO ↔ I = RTln(O/I) = RTln[b/(b − a)]. The example shown is a Δ2-354 current. (E) Normalized steady state outward current–voltage relationships for WT and Δ2-354 channels (n = 6). All outward currents are normalized to the maximal tail current level evoked at −100 mV subsequent to a 3-s step to 60 mV, which represents the channel expression level for each oocyte assuming the same single-channel conductance for the two constructs.
	Figure 3. NH2 terminus regulation of deactivation is independent of the inactivation process. (A) Scaled outward currents from S620T and S620T Δ2-354 channels show that deleting the NH2 terminus increases deactivation rate even in the absence of C-type inactivation. (B) Fast and slow deactivation time constants for WT (n ≥ 10), Δ2-354 (n ≥ 10), S620T (n = 6), and S620TΔ2-354 (n = 6) channels showing that the change in deactivation rate due to deletion of the NH2 terminus is the same in the absence of C-type inactivation as in the wild-type background.
	Figure 5. NEM modification of the S4–S5 linker mimics the NH2 terminus deletion. (A) Deactivation rates for cysteine substitutions at each position within the S4–S5 linker region. The open bars with the asterisks indicate the positions where cysteine substitution significantly speeds deactivation rate (n ≥ 3 for each mutant). Note that deactivation rate is not significantly altered by G546C. (B) Fractional change of deactivation rate by NEM modification for all the cysteine substitutions. Positive numbers indicate increased rate. *Dramatic effect on the G546C mutant. (C) Families of tail currents from G546C channels before and after NEM modification. (D) Summary of effects of NEM modification on WT, G546C, and G546C Δ2-354 channels. Time constants were determined by fitting tail currents to a single exponential function because, at −100 mV, the fast component accounts for >90% of the current. The first pair of bars indicates that NEM modification does not significantly affect deactivation time constants for wild-type channels. The second set of bars shows marked effect of NEM modification on G546C mutant. The third set of bars shows that modification of G546C Δ2-354 does not further accelerate the deactivation rate, indicating that NEM and deletion of the NH2 terminus disrupt the same process (n = 6 for each construct). (E) Inactivation time constants measured from G546C channels show that modification of the S4–S5 linker results in slower C-type inactivation (n = 7).
	Figure 6. Domains regulating deactivation and inactivation are spatially distinct within the NH2 terminus. (A) Scaled tail currents from Δ2-12 and WT channels show that the small deletion increases deactivation rate. (B) Fast and slow deactivation time constants for WT (n > 10), Δ2-354 (n > 10), and Δ2-12 (n = 6) channels indicate that the two deletions have similar effects on deactivation kinetics. (C) Inactivation time constants for HERG WT (n > 10), Δ2-354 (n > 10), and Δ2-12 (n = 6) show that the small deletion does not significantly alter the inactivation rate (P > 0.6 at 0.05 confidence interval for each voltage).
	Scheme II.

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