Na⁺ channel chimaeras exhibit an intermediate slow inactivation development phenotype

Development of slow inactivation was studied in the three Na⁺ channel chimaeras constructed with domains from hH1 and µ1: µ1₍₁₎hH1_(2,3,4), µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎. All three chimaeras exhibited slow inactivation development phenotypes that were intermediate relative to hH1 and µ1. However, development of slow inactivation in µ1₍₁₎hH1_(2,3,4) (n = 8) was more similar to that in hH1 than to that in µ1 (Fig. 3A and B). Although the onset of slow inactivation in the chimaera µ1₍₁₎hH1_(2,3,4) appeared similar to that in hH1, ₁ was less in µ1₍₁₎hH1_(2,3,4) (6·0 vs. 6·9 s) and ₂ was greater (87·5 vs. 52·0 s) than in hH1 (Table 2). In addition, depolarization to 0 mV for 5 s or more produced significantly more slow inactivation in µ1₍₁₎hH1_(2,3,4) (to 66 % at 5 s and to 26 % at 300 s) than in hH1 (to 79 % at 5 s and to 36 % at 300 s; P < 0·01; Fig. 3B).

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Figure 3. Development of slow inactivation in Na+ channel chimaeras is intermediate relative to that in hH1 and in µ1 Data were collected as described in Fig. 2. A, development of slow inactivation in µ1(1)hH1(2,3,4) is more similar to that in hH1 than to that in µ1. Normalized representative traces for hH1 (continuous lines) and µ1(1)hH1(2,3,4) (dashed lines) before (larger traces) and after (smaller traces) depolarization to 0 mV for 120 s are shown. B, mean ± S.E.M. values for hH1 (, continuous line; n = 11) and µ1(1)hH1(2,3,4) (, dashed line; n = 8) were fitted with a double exponential function. * P < 0·05 vs. hH1. C, normalized traces from µ1 (continuous lines), µ1(1,2)hH1(3,4) (dotted lines), and µ1(1,2,3)hH1(4) (dashed lines) before and after depolarization to 0 mV for 120 s to induce slow inactivation. D, mean ± S.E.M. values for µ1 (, continuous line; n = 10), µ1(1,2)hH1(3,4) (, dotted line; n = 8) and µ1(1,2,3)hH1(4) (, dashed line; n = 10) were fitted with a double exponential function. * P < 0·05 vs. µ1.

The other two chimaeras, µ1_(1,2)hH1_(3,4) (n = 8) and µ1_(1,2,3)hH1₍₄₎ (n = 10), expressed a slow inactivation phenotype more similar to that of µ1 than to that of hH1 (Fig. 3C and D). The time constants ₁ and ₂ did not differ significantly among µ1_(1,2)hH1_(3,4), µ1_(1,2,3)hH1₍₄₎ and µ1 (Table 2). However, both µ1_(1,2)hH1_(3,4) (to 26 %) and µ1_(1,2,3)hH1₍₄₎ (to 25 %) showed less slow inactivation than µ1 (to 18 %; P < 0·01) after 10 s depolarization (Fig. 3D). With longer depolarization (>120 s), slow inactivation was less in µ1_(1,2)hH1_(3,4) (to 16-19 %) than in µ1_(1,2,3)hH1₍₄₎ or µ1 (to 11-14 %; P < 0·05; Fig. 3D).

Steady-state slow inactivation is less voltage dependent in hH1 than in µ1

Steady-state slow inactivation (s) was determined with a modification of the h protocol. Cells were held at the prepulse potential for 120 s, stepped to -140 mV for 50 ms to allow recovery from fast inactivation, and then stepped to +30 mV to measure I_Na (see Fig. 4A, inset). As with the development of slow inactivation, steady-state slow inactivation differed between hH1 (n = 6) and µ1 (n = 8). A Boltzmann fit of the mean data for steady-state inactivation showed that the V½ for hH1 was more negative than that for µ1 (-78·3 vs. -73·1; Fig. 4A and Table 2). In addition, the s curve for µ1 was much steeper than that for hH1 (k = 10·3 vs. 23·1; P < 0·001; Fig. 4A and Table 2).

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Figure 4. Voltage dependence of steady-state slow inactivation (s) differs among wild-type and chimaeric Na+ channels Peak available current was obtained with a test pulse to +30 mV after a 120 s prepulse at the voltages shown on the abscissa. An intervening step to -140 mV for 50 ms allowed recovery from fast inactivation. Mean ± S.E.M. values were fitted with a Boltzmann function. A, voltage dependence of steady-state slow inactivation is much less steep for hH1 (; n = 6) than for µ1 (; n = 8). B, the s curve for µ1(1)hH1(2,3,4) (, dashed line; n = 10) is similar to that for hH1 (continuous line) although the slope is steeper in µ1(1)hH1(2,3,4). C, the s curves for µ1(1,2)hH1(3,4) (, dotted line; n = 6) and µ1(1,2,3)hH1(4) (, dashed line; n = 8) are similar to that for µ1 (continuous line), although V½ is more positive in µ1(1,2)hH1(3,4) and more negative in µ1(1,2,3)hH1(4) than in µ1.

Steady-state slow inactivation in Na⁺ channel chimaeras is different from that in hH1 and µ1

s in the chimaera µ1₍₁₎hH1_(2,3,4) (n = 10) was more similar to that in hH1 than µ1 (Fig. 4B). However, V½ of the s curve was significantly more positive for µ1₍₁₎hH1_(2,3,4) than for hH1 (Table 2). In addition, the slope of the s curve was steeper for µ1₍₁₎hH1_(2,3,4) than for hH1 (Table 2 and Fig. 4B).

The s curves for the chimaeras µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎ were more like that for µ1 than that for hH1. However, V½ of the s curves for the chimaeras was shifted compared with µ1 (Fig. 4C). The s curve for µ1_(1,2)hH1_(3,4) (n = 6) was shifted > 10 mV in a positive direction relative to that for µ1 (P < 0·05), while the s curve for µ1_(1,2,3)hH1₍₄₎ (n = 8) was shifted > 10 mV in a negative direction relative to that for µ1 (Table 2). The slopes of the s curves were not different for µ1, µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎ (Table 2).

Steady-state slow inactivation shows little time-dependent shift

A well-known feature of steady-state fast inactivation is the time-dependent shift of V½ in the negative direction (Wang et al. 1996a). This shift is not readily apparent in steady-state slow inactivation. For example, during 45 min of whole-cell recording from one cell expressing µ1, the h curve showed a shift in V½ of -15 mV (Fig. 5A). In contrast, the V½ of s shifted < -3 mV in the same cell. The mean shift in s between 15 and 60 min of whole-cell recording for five cells expressing µ1 was -2·7 ± 1·0 mV (Fig. 5B). In three cells expressing hH1, the shift was -1·4 ± 1·3 mV (Fig. 5C). Similar negligible shifts in s over time (i.e. < -3 mV h^-1) were seen in the chimaeras (data not shown).

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Figure 5. Steady-state slow inactivation (s) changes little with time Slow inactivation data were obtained as described in Fig. 4. Steady-state fast inactivation (h) curves were obtained with a 300 ms prepulse to the voltage on the abscissa, then a test pulse to +30 mV. A, curves (s and h) from one cell expressing µ1. The s curve (continuous lines) changes little (V½ < -3 mV) between 15 min () and 60 min () of whole-cell recording, while V½ of the h curve shifts by ~-15 mV between 15 min () and 60 min (). B, mean ± S.E.M. values of s for five cells expressing µ1were fitted with a Boltzmann equation; curves shown are at 15 min () and 60 min () of whole-cell recording. C, mean ± S.E.M. values of s for three cells expressing hH1 were fitted with a Boltzmann equation; curves are at 15 min () and 60 min () of whole-cell recording.

Recovery from slow inactivation is faster in hH1 than in µ1

Na⁺ channels recover from the slow-inactivated state when the membrane is repolarized. Mean data for recovery from slow inactivation were fitted with a double exponential function. Following depolarization to 0 mV for 120 s to induce slow inactivation (Fig. 6A, inset), I_Na in hH1 (n = 6) at -140 mV recovered faster than that in µ1 (n = 10; Fig. 6A). The time constants of recovery from slow inactivation were significantly less for hH1 than for µ1 (Table 2). Also, recovery from slow inactivation (P > 0·05 compared with I_Na at 300 s) occurred much sooner in hH1 (by 30 s) than in µ1 (by 120 s; Fig. 6A).

Recovery from slow inactivation is intermediate in Na⁺ channel chimaeras

Recovery from slow inactivation appeared to be slower in the chimaera µ1₍₁₎hH1_(2,3,4) (n = 8) than in hH1 (Fig. 6B). However, the recovery time constants for µ1₍₁₎hH1_(2,3,4) and hH1 were not statistically different (Table 2). Recovery was complete for both hH1 and µ1₍₁₎hH1_(2,3,4) by 30 s at the holding potential of -140 mV following induction of slow inactivation (i.e. 120 s at 0 mV).

The recovery time constant ₁ was similar for µ1_(1,2)hH1_(3,4) (n = 6), µ1_(1,2,3)hH1₍₄₎ (n = 8), and µ1 (Table 2). However, ₂ for µ1 was greater than ₂ for the chimaeras (Table 2). The chimaera µ1_(1,2)hH1_(3,4) recovered sooner (by 60 s) than either µ1_(1,2,3)hH1₍₄₎ or µ1 (by 120 s; Fig. 6C). However, all wild-type and chimaeric Na⁺ channels recovered from slow inactivation (0 mV, 120 s) by 120 s at the holding potential of -140 mV (Fig. 6).

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Figure 6. hH1 recovers from slow inactivation faster than µ1, while recovery in the chimaeras falls between that in hH1 and in µ1 Cells were depolarized to 0 mV for 120 s, then stepped to -140 mV for various times before the test pulse to +30 mV (inset in A). Current was normalized to peak current recorded after 300 s at -140 mV. Mean ± S.E.M. values were fitted with a double exponential function. A, hH1 (; n = 6) recovers from slow inactivation much faster and sooner (by 30 s) than µ1 (by 120 s; ; n = 10). B, recovery in µ1(1)hH1(2,3,4) (, dashed line; n = 8) was slower but not statistically different from that in hH1 (continuous line). Both hH1 and µ1(1)hH1(2,3,4) recovered by 30 s. C, recovery in the chimaeras µ1(1,2)hH1(3,4) (, dotted line; n = 6) and µ1(1,2,3)hH1(4) (, dashed line; n = 6) is similar to that in µ1 (continuous line), although µ1(1,2)hH1(3,4) recovers sooner (by 60 s) than µ1(1,2,3)hH1(4) or µ1 (by 120 s).

	DISCUSSION

Top Abstract Introduction Methods Results Discussion References

This study compared slow inactivation in wild-type human heart Na⁺ channels (hH1), wild-type rat skeletal muscle Na⁺ channels (µ1) and Na⁺ channel chimaeras constructed with domains from hH1 and µ1 (µ1₍₁₎hH1_(2,3,4), µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎). In addition, activation and fast inactivation kinetics were also studied in wild-type and chimaeric Na⁺ channels. The results showed that development of slow inactivation in µ1 has a faster onset, a steeper voltage dependence and is more complete compared with hH1. In addition, recovery from slow inactivation was much slower for µ1 than for hH1. Slow inactivation in the chimaeras was intermediate to that of wild-type. However, the chimaera µ1₍₁₎hH1_(2,3,4) was more like hH1, while the chimaeras µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎ were more similar to µ1. Activation and fast inactivation of the wild-type Na⁺ channels (hH1 and µ1) also differed, while the chimaeras exhibited kinetics that were, in general, intermediate between those of the wild-type isoforms. Fast inactivation was slower, and V½ of activation and steady-state inactivation were more negative, in hH1 compared with µ1. The chimaeras were more similar to µ1 in activation, more similar to hH1 in fast inactivation, and intermediate to hH1 and µ1 in steady-state fast inactivation. The results demonstrate that Na⁺ channel kinetics can be modulated by all four domains of the Na⁺ channel protein. In particular, slow inactivation appears to be a process that may involve participation of all four domains of the -subunit of the Na⁺ channel molecule.

Activation and fast inactivation

Initial experiments determined the kinetics of activation and fast inactivation in wild-type and chimaeric Na⁺ channels. Some previous studies have presented data on these parameters in cloned hH1, µ1 and chimaeric Na⁺ channels (Chahine et al. 1996; Wang et al. 1996a; Wright et al. 1999). The data from our study are similar to those previously reported. For example, the activation and steady-state fast inactivation (h) curves (i.e. V½) for hH1 were more negative than those for µ1, and the macroscopic current decay (from +30 mV) was slower for hH1 than for µ1. In the chimaeras, activation was similar to that in µ1. Because all three chimaeras contained D1 from µ1, one could speculate that D1 is 'dominant' in determining the voltage dependence of activation in Na⁺ channels. However, the chimaera µ1_(2,3)hH1_(1,4) also exhibits a µ1 activation phenotype (Chahine et al. 1996), suggesting that the presence of any µ1 domain in a hH1-µ1 chimaera can produce a µ1 activation phenotype. In addition, point mutations in the S4 segment of all four domains affect the V½ of activation (Chen et al. 1996; Kontis et al. 1997). Taken together, these results suggest that activation is a complex process that probably involves all four domains of the Na⁺ channel molecule.

It should also be noted that the activation curve for the chimaeras was less steep than that for either hH1 or µ1. If it is assumed that hH1 and µ1 domains do not differ in net charge, a possible explanation for the difference in slope is a shift in the equilibrium of voltage-dependent activation steps (Sigworth, 1994). This shift might be due to disruption of interactions between domains that are important for stabilization of transitional conformation states (McCormack et al. 1991). Fast inactivation in the chimaeras (as measured by macroscopic current decay from +30 mV) resembled fast inactivation in hH1. All three chimaeras contained D4 and the D3-D4 linker from hH1. These results support the conclusion of other investigators that this region of the Na⁺ channel -subunit is involved in fast inactivation gating (Stuhmer et al. 1989; Chen et al. 1996; McPhee et al. 1998). Steady-state fast inactivation (h) for the chimaeras fell between that for hH1 and that for µ1. As proposed previously, the intermediate V½ of h for the chimaeras suggests that all four domains contribute to the voltage dependence of Na⁺ channel steady-state fast inactivation (Chen et al. 1996; Deschenes et al. 1998; Wright et al. 1999).

Development of slow inactivation

Having examined activation and fast inactivation, we determined the slow inactivation phenotype (development, steady state and recovery) in wild-type and chimaeric Na⁺ channels. Slow inactivation differs from fast inactivation in the time frame of onset and recovery (seconds vs. milliseconds) and in the amount of current that slow inactivates (slow inactivation is less complete than fast inactivation). In addition, slow inactivation appears to be structurally separate from fast inactivation (Rudy, 1978; Featherstone et al. 1996), although slow inactivation can be enhanced when fast inactivation is removed (Rudy, 1978; Featherstone et al. 1996). Comparison of hH1 and µ1 expressed in HEK cells showed a marked difference in slow inactivation phenotype between the two isoforms. Slow inactivation developed much faster and more completely in µ1 than in hH1. As only the -subunit of the respective Na⁺ channel was expressed in HEK cells and the same protocols were used to compare slow inactivation, the difference between hH1 and µ1 is probably due to molecular differences between the -subunits of the two isoforms (Trimmer et al. 1989; Gellens et al. 1992). In particular, several studies have shown that mutations in and around the pore region of Na⁺ channels can alter the slow inactivation phenotype (Balser et al. 1996; Hayward et al. 1997; Wang & Wang, 1997). Therefore, amino acid differences between hH1 and µ1 in and/or around the pore region may underlie the difference in slow inactivation phenotype of these two Na⁺ channel isoforms.

The results in this report complement several previous studies on slow inactivation in heart and muscle Na⁺ channels. Richmond et al. (1998) recently reported on slow inactivation in a human heart -subunit, hH1a, expressed in Xenopus oocytes and recorded with on-cell macropatches. The results are similar to the data in this study, although hH1a expressed in the oocyte system appears to reach a steady-state level of slow inactivation faster (60 s) than hH1 expressed in HEK cells in this study (120 s). In addition, onset of and recovery from slow inactivation in hH1a expressed in oocytes was fitted with a single exponential (Richmond et al. 1998). In hH1 expressed in HEK cells, a double exponential function best described both development of and recovery from slow inactivation (this study). In SkM1 (i.e. µ1) expressed with the ₁-subunit in oocytes, onset of and recovery from slow inactivation apparently also followed a single exponential (Featherstone et al. 1996), whereas slow inactivation in µ1 expressed in HEK cells is described by (at least) a double exponential function (this study; Hayward et al. 1997). Regardless of potential differences due to different expression systems, these studies demonstrate that there are dramatic differences in slow inactivation between hH1 and µ1.

To explore the molecular basis for these differences, we studied slow inactivation in Na⁺ channel chimaeras. These experiments tested the hypothesis that the molecular mechanism for slow inactivation is localized to one or more domains of the Na⁺ channel molecule. The results showed that slow inactivation in the Na⁺ channel chimaeras is intermediate relative to that in hH1 and in µ1. This result demonstrates that all four domains can influence or modulate the slow inactivation phenotype of the Na⁺ channel -subunit. However, slow inactivation of the chimaeras µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎, which have D1 and D2 from µ1, resemble the µ1 phenotype much more than the hH1 phenotype. Therefore, D1 and D2 (or the combination of D1 and D2) may play a more prominent role than D3 and D4 in the development of Na⁺ channel slow inactivation. This conclusion could be further tested in chimaeras with D1 and/or D2 from hH1, e.g. hH1₍₁₎µ1_(2,3,4), hH1_(1,2)µ1_(3,4) and hH1_(1,2,3)µ1₍₄₎. These chimaeras were constructed but, despite repeated transfections, expressed little to no Na⁺ current. It should be noted that while the data from the chimaera studies show that the slow inactivation phenotype can be altered by different combinations of domains in the chimaeras, the data do not determine the specific regions directly involved in the slow inactivation process.

Interaction between domains is found in other aspects of Na⁺ channel function. For example, all four domains appear to contribute to the voltage dependence of steady-state fast inactivation, although the respective contributions of each domain may not be equivalent (this study; Chen et al. 1996; Kontis & Goldin, 1997; Wright et al. 1999). In addition, activation gating is dependent on the S4 segment from all four domains of the Na⁺ channel, probably with unequal contributions (Kontis et al. 1997). The data from this study describe another kinetic process in Na⁺ channels, i.e. slow inactivation, that may be dependent on interactions involving all four domains.

Steady-state slow inactivation

The voltage dependence of steady-state slow inactivation (s) also appears to be dependent on all four domains of the Na⁺ channel. As with the development of slow inactivation, the chimaera µ1₍₁₎hH1_(2,3,4) was more similar to hH1, while the chimaeras µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎ were more similar to µ1 (at least in slope k). These results again suggest that D1 and D2 may play a more prominent role than D3 and D4 in determining Na⁺ channel slow inactivation phenotype. The reason for the differences in V½ between µ1_(1,2)hH1_(3,4), µ1_(1,2,3)hH1₍₄₎ and µ1 is unclear. While the s of µ1_(1,2)hH1_(3,4) could be interpreted as being intermediate relative to those of hH1 and µ1, the negative shift of µ1_(1,2,3)hH1₍₄₎ (relative to µ1) is more difficult to explain. One possible explanation is that an alteration in a co-operative interdomain mechanism in µ1_(1,2,3)hH1₍₄₎ promotes stabilization of the slow-inactivated state(s) (McCormack et al. 1991).

An interesting feature of the s curve is that it does not exhibit the same time-dependent negative shift as the h curve. Although the time-dependent shift in h has been described in detail in heart and muscle Na⁺ channels, the reason for the negative shift is unknown (Hanck & Sheets, 1992; Wang et al. 1996a). One possible explanation for the time-dependent difference between s and h is that the fast inactivation gate is accessible to a cellular process (e.g. phosphorylation) that produces the shift, while the slow inactivation gate is not modulated by or is not accessible to the same or similar processes.

Recovery from slow inactivation

After 120 s at 0 mV, recovery from slow inactivation appears to depend on the amount of current that is slow inactivated, i.e. the less slow-inactivated current, the faster the recovery. For example, hH1 and µ1₍₁₎hH1_(2,3,4) showed a similar amount of slow-inactivated current after 120 s at 0 mV, and the rate of recovery was essentially identical for these two Na⁺ channels. The chimaeras µ1_(1,2)hH1_(3,4) and µ1_(1,2,3)hH1₍₄₎ were more like µ1 in the amount of slow inactivation and rate of recovery. The greatest difference in recovery from slow inactivation was between µ1 and the other Na⁺ channels. µ1 had a much greater recovery time constant ₂ than either hH1 or the chimaeras. One possible explanation is that disruption of domain interaction by substitution of another domain into the µ1 background enhances the speed of recovery by destabilizing the slow inactivation state(s).

Relation of Na⁺ channel slow inactivation to C-type inactivation in K⁺ channels

Na⁺ channels are structurally similar and evolutionarily related to voltage-gated K⁺ channels (Hille, 1989) in which slow (C-type) inactivation has been described (Hoshi et al. 1991). K⁺ channels are homotetramers, and mutation studies have determined that C-type inactivation occurs by a co-operative mechanism involving all four identical subunits (Yellen et al. 1994; Ogielska et al. 1995; Panyi et al. 1995). The proposed mechanism for C-type inactivation is a constriction of the outer mouth of the channel pore with each subunit contributing equally (Panyi et al. 1995), perhaps altering the ion selectivity of the pore (Starkus et al. 1997). In contrast to K⁺ channels, Na⁺ channels are formed from a single protein with four homologous, but not identical, domains (Noda et al. 1986). However, studies show that Na⁺ channel slow inactivation is altered with point mutations in and around the pore region (Balser et al. 1996; Wang & Wang, 1997; Hayward et al. 1997). Therefore, slow inactivation in Na⁺ channels and C-type inactivation in K⁺ channels may occur via a similar molecular mechanism (Townsend & Horn, 1997; Wang & Wang, 1997).

Physiological and pathophysiological significance of slow inactivation

Slow inactivation is thought to play an important role in membrane excitability and firing properties (Ruff et al. 1988; Sawczuk et al. 1995). In brain, Na⁺ channel slow inactivation may contribute to spike frequency adaptation (Sawczuk et al. 1995; Fleidervish et al. 1996), and may encode information of previous neuronal activity (Toib et al. 1998). In skeletal muscle, Na⁺ channel slow inactivation may figure prominently in muscle fatigue (Ruff et al. 1988). Compared with brain and muscle Na⁺ channels, hH1 undergoes little slow inactivation (this study; Townsend & Horn, 1997; Richmond et al. 1998). The 'resistance' to slow inactivation of hH1 may avoid a potential reduction in Na⁺ channel availability during the relatively long cardiac action potential (300 ms). Therefore, Na⁺ channel slow inactivation may play only a minor role in normal cardiac function, while playing a more prominent and physiologically important role in brain and muscle.

Inheritable mutations that alter slow inactivation in Na⁺ channels have been identified in human skeletal muscle. These mutations underlie such diseases as hyperkalaemic periodic paralysis (Cannon, 1997). To date, no similar mutations have been identified in brain or heart Na⁺ channels. It is interesting to speculate that mutations in human brain or heart Na⁺ channels that alter or disrupt slow inactivation could have profound effects on normal physiological function in these tissues.

In summary, hH1 and µ1 exhibit dramatically different slow inactivation phenotypes. The difference is probably due to molecular differences between the -subunits of hH1 and µ1. Studies of Na⁺ channel chimaeras constructed with domains from hH1 and µ1 demonstrate that the slow inactivation phenotype can be modulated by all four Na⁺ channel domains. The data suggest that slow inactivation in Na⁺ channels may occur via a mechanism similar to that involved in C-type inactivation of voltage-gated K⁺ channels, i.e. a co-operative interaction involving all four domains. Future studies with site-directed mutagenesis (single, double or multiple mutations) in and around the pore region may provide further information on the molecular basis of Na⁺ channel slow inactivation.

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Acknowledgements

This work was supported by the National Institutes of Health (NIH) grants GM 35401 and GM 49090. Dr R. G. Kallen was supported by NIH grant AR 41762, the American Heart Association, Muscular Dystrophy Association and the University of Pennsylvania Research Foundation.

Corresponding author

J. P. O'Reilly: Department of Anesthesia, Brigham and Women's Hospital, Harvard Medical School, 75 Francis Street, Boston, MA 02115, USA.

Email: joreilly{at}zeus.bwh.harvard.edu

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