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Department of Microbiology & Molecular Genetics, University of California, Irvine, California 92697-4025
Correspondence: Address reprint requests to A. L. Goldin, Tel.: 949-824-5334; Fax: 949-824-8504; E-mail: agoldin{at}uci.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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; Goldin, 2001). In mammals, there are nine sodium channel 
-subunit isoforms named Nav1.1–Nav1.9 (Goldin, 2002; Goldin et al., 2000). Three of these (Nav1.1, Nav1.2, and Nav1.6) are expressed at high levels in the adult central nervous system (CNS). The genes encoding Nav1.1 (SCN1A) and Nav1.2 (SCN2A) are clustered together on chromosome 2 in both mouse and human, whereas the gene encoding Nav1.6 (SCN8A) is located on chromosome 15 in mouse and chromosome 12 in human (Litt et al., 1989; Malo et al., 1991, 1994; Plummer et al., 1998). Although the amino acid sequence of Nav1.6 is >84% identical to the sequences of Nav1.1 and Nav1.2, it is more distant from those two than they are to each other (Goldin et al., 2000).
The three adult CNS isoforms are selectively expressed in different cell types and localized in different neuronal regions, which may indicate that they serve different purposes as determined by their electrophysiological properties. Nodes of Ranvier are the axonal regions without myelin and glial cell wrapping, which are essential for the efficient conduction of action potentials. Nav1.6 is the predominant isoform at the nodes of Ranvier in both sensory and motor neurons of the adult CNS and peripheral nervous system (Caldwell et al., 2000). Studies of developing retinal ganglion cells have shown that Nav1.2 and ß2 are clustered at immature nodes of Ranvier. As myelination proceeds, Nav1.6 replaces Nav1.2 (Boiko et al., 2001; Kaplan et al., 2001). Little Nav1.6 and much more of Nav1.2 are detected on myelin-deficient axons of shiverer mice (Boiko et al., 2001). Nav1.6 disappears from the nodes after nerve injury (Novakovic, 1999). These data suggest that Nav1.6 may play a critical role in faithfully transmitting high-frequency action potentials, which could reflect unique electrophysiological properties of Nav1.6 compared to Nav1.2.
In this study, we compared the properties of Nav1.6 and Nav1.2 in response to a rapid series of repetitive depolarizations. Previous studies have shown that currents through Nav1.1 and Nav1.2 with or without the auxiliary ß-subunits decrease in response to high-frequency depolarizations (Pugsley and Goldin, 1998; Spampanato et al., 2001). In contrast, currents through Nav1.6 coexpressed with ß1 increased in response to high-frequency depolarizations. The use-dependent potentiation still occurred when fast inactivation was removed by mutation, suggesting that it resulted from changes in channel activation. Consistent with this interpretation, high-frequency depolarizations accelerated a slow phase of activation in the Nav1.6 channel that had fast inactivation removed. The activation kinetics of Nav1.2 with ß1 were faster than those of Nav1.6, so that it might already be close to the maximal rate. Repetitive depolarizations only slightly accelerated the activation kinetics of Nav1.2 with ß1 and the current decreased due to its faster slow inactivation. These results suggest that Nav1.6 might be specialized to more faithfully propagate high-frequency firing at nodes of Ranvier compared to Nav1.2.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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The mutation was confirmed by sequencing. The MfeI to BstEII fragment was then ligated back into the full-length clone to make Nav1.6Q3. The Nav1.2Q3 mutant was previously described (West et al., 1992).
Expression of the sodium channels - and ß1-subunits in Xenopus oocytes
Stage V oocytes were removed from adult female Xenopus laevis frogs and prepared as previously described (Goldin, 1991). Oocytes were incubated in ND-96 media, which consists of 96 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, and 5 mM HEPES, pH 7.5, supplemented with 0.1 mg/ml gentamicin, 0.55 mg/ml pyruvate, and 0.5 mM theophylline. Plasmids containing the sodium channel wild-type and mutant -subunit or ß1-subunit cDNA were linearized with NotI, and capped full-length transcripts were synthesized in vitro using T7 RNA polymerase (mMESSAGE mMACHINE kit; Ambion., Austin, TX). RNA was dissolved in 1 mM Tris-HCl, pH 7.5, and injected into oocytes. The ratio of - to ß1-subunits was 1:10. Oocytes were incubated in ND96 at 20°C for 1–2 days before recording.
Electrophysiological recording
Electrophysiological recording used either the two-electrode voltage clamp or the cut-open oocyte voltage clamp. The two-electrode voltage clamp experiments were performed using the oocyte clamp OC-725B (Warner Instruments., Hamden, CT), DigiData 1322A interface (DAGAN, Minneapolis, MN), and pCLAMP software (version 8.1, Axon Instruments, Burlingame, CA). The recording solution was ND-96 without supplements. Tetrodotoxin (TTX) subtraction was used to eliminate all non-TTX sensitive currents by subtracting those currents recorded in the presence of 400 nM TTX from those recorded in the absence of TTX.
The cut-open oocyte voltage-clamp experiments were performed using the CA-1 high-performance oocyte clamp (DAGAN), DigiData 1321A interface, and pClamp software (Smith and Goldin, 1998). All recordings were obtained after stable baseline and ionic current levels were achieved. The experiments were performed with an external solution containing (in mM) 120 NaOH-methyl sulfonate, 1.8 Ca(OH)2-methyl sulfonate, and 10 HEPES, pH 7.5. The internal solution consisted of 44 K2SO4, 5 Na2SO4, 10 EGTA-Cs, and 10 HEPES-Cs, pH 7.5.
Data analysis
Data analysis was performed using pCLAMP software (version 8.1, Axon Instruments, Burlingame, CA). Peak conductance was calculated using the following equation:
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The kinetics of fast inactivation were analyzed by fitting the current traces between 0.4 ms after the peak current and the end of the 20-ms depolarization with a double exponential equation:
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s is the slow time constant of inactivation, f is the fast time constant of inactivation, As and Af are the percentages of current inactivating with the slow and fast time constants, respectively, and C is the noninactivating current.
The kinetics of activation were analyzed by fitting the current traces between the start of activation and the peak current with either a single or double exponential equation of the form:
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s is the slow time constant of activation, f is the fast time constant of activation, and As and Af are the percentages of current activating with the slow and fast time constants, respectively. The choice of a single exponential equation for Nav1.2 and a double exponential equation for Nav1.6 was made empirically by fitting activation for both channels with both types of equations and visually inspecting the quality of the fits.
The kinetics of slow inactivation were analyzed by fitting the current traces between the peak current and the end of the 10-s depolarization with a double exponential equation:
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s is the slow time constant of inactivation, f is the fast time constant of inactivation, and As and Af are the percentages of current inactivating with the slow and fast time constants, respectively. |
RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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). Two of these isoforms (Nav1.1 and Nav1.2) demonstrate a decrease in current with repetitive depolarizations (Pugsley and Goldin, 1998; Spampanato et al., 2001), which might limit their ability to propagate high-frequency action potentials. Because Nav1.6 is the primary channel at nodes of Ranvier (Caldwell et al., 2000), that channel might have different characteristics that enable it to sustain high-frequency firing in myelinated axons. To test this possibility, we examined the current through Nav1.6 compared to Nav1.2 during a 10-Hz train of 20 depolarizations, each 20-ms long from –100 mV to –10 mV. The peak current during each depolarization was normalized to the peak current during the first depolarization and plotted against the depolarization number (Fig. 1, A and B). The first and the last traces of the 20 depolarizations are shown in Fig. 1, C–F. When the -subunit of Nav1.6 or Nav1.2 was expressed alone, the current decreased and reached a steady-state level (panels A, C, and D). The use-dependent decrease in current for Nav1.6 was less pronounced than that for Nav1.2. In contrast, when Nav1.6 was coexpressed with the ß1-subunit, the channel demonstrated an increase in current with successive depolarizations (panels B and F). Nav1.2 with ß1 demonstrated no significant change in current amplitude under the same set of conditions (panels B and E).
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with ß1 during this protocol. The current level was stable during the baseline depolarizations at 1-min intervals (open circles), suggesting that 1 min provides sufficient time for all of the channels to return to the original state. The current decreased during the period of rapid stimulation (open squares), consistent with previous results for Nav1.2 (Pugsley and Goldin, 1998). The current returned to the original level after only three depolarizations at 10-s intervals (open triangles). The fact that complete recovery required >10 s was surprising because complete recovery was obtained within 300 ms when current through Nav1.2 with ß1 was fully inactivated with a single 50-ms depolarization (Smith et al., 1998). This difference could be due to the fact that the total depolarization time in this experiment was 4 s (200 depolarizations of 20 ms each), which would have caused some of the channels to enter the slow inactivated state. Depolarization to –10 mV for 4 s results in slow inactivation of 
80% of current through Nav1.1 with ß1, and it takes significantly longer to recover from slow inactivation than from fast inactivation (Spampanato et al., 2001).
In contrast to the results with Nav1.2, current through Nav1.6 with ß1 increased with successive depolarizations and didn't stabilize until after 50 depolarizations (Fig. 2 C, open squares). The current gradually returned to the baseline level during the recovery period (open triangles), although it required seven depolarizations (60 s) for complete recovery. This recovery time was approximately three times as long as the time required for Nav1.2 with ß1 to recover from the current decrease, suggesting that the two changes are not the result of comparable processes. The current remained steady during the 1-min intervals at the end of the protocol (solid circles), demonstrating that the potentiation was due to the high-frequency depolarizations.
To examine the effect of rapid stimulation on the kinetics of fast inactivation, the current traces during the first (solid line) and last (dotted line) depolarizations of rapid stimulation were plotted together for Nav1.2 with ß1 (Fig. 3 A) and Nav1.6 with ß1 (Fig. 3 B). Rapid stimulation accelerated the fast inactivation of both Nav1.2 and Nav1.6. The current traces were fit with a double exponential equation to quantitatively compare the kinetics of fast inactivation, and the results are shown in Table 1. For Nav1.2, rapid stimulation accelerated all aspects of fast inactivation, whereas for Nav1.6 the primary effects were a large increase in the percentage of current inactivating with the fast time constant and a decrease in the amount of persistent current.
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Nav1.6 potentiation depends on the extent of activation and inactivation
To examine the relationship between the use-dependent potentiation of Nav1.6 and channel activation, we compared the voltage dependence of both processes. The voltage dependence of activation for the Nav1.6 -subunit alone (Fig. 4 A, open squares) was 3.7 mV more positive than that for the Nav1.2 -subunit alone (Fig. 4 A, open circles; Table 2). Coexpression of the ß1-subunit shifted the voltage dependence of activation for both channels to more negative values that were similar (Fig. 4 B; Table 2). To determine the voltage dependence of potentiation for Nav1.6 with ß1, the current increase during a series of depolarizations to different voltages was measured by comparing the current amplitude during the first depolarization of the recovery interval with that of the last depolarization of the baseline interval. The increase was compared to the current change observed during repetitive depolarizations to –50 mV, at which potential there was no potentiation with rapid stimulation. The amount of increase was normalized to the maximal increase in current, and the data from different oocytes were averaged and plotted against the depolarization voltage (Fig. 4 C). The potentiation increased from –40 mV to 0 mV, at which voltage the increase stabilized. The relationship is similar to that observed for the voltage dependence of activation, with a V1/2 of –17 mV. These results suggest that the increased current of Nav1.6 with ß1 correlates with the extent of channel activation during rapid stimulation.
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Because rapid stimulation with depolarization times shorter than 30 ms resulted in smaller increases in current, it is possible that very short depolarizations that would be more similar to those during an action potential might not result in any potentiation. To test this possibility, a rapid-stimulation protocol was performed using 2-ms depolarizations from –120 to –10 mV. The normalized current amplitude during each depolarization was plotted against the depolarization number for Nav1.2 or Nav1.6 with ß1 (Fig. 5 B). With 2-ms depolarizations, current through Nav1.6 with ß1 still increased slightly in response to rapid stimulation, whereas current through Nav1.2 with ß1 decreased very slightly.
Potentiation of Nav1.6 is unstable at hyperpolarized potentials
The return of current to the baseline level after repetitive depolarizations suggests that the increase was unstable at negative potentials. To examine the stability of the current increase, we varied the time between depolarizations from 100 to 1000 ms. The current increase for each time interval between depolarizations was compared to the change in current with a 2-s interval, for which time there was no significant potentiation. The amount of increase was normalized to the maximal increase in current, and the data from different oocytes were averaged and plotted against the time interval (Fig. 5 C). The increase was larger with smaller time intervals, corresponding to a higher frequency of depolarization. This result demonstrates that the channels recover from the current increase during the intervals of hyperpolarization between depolarizations during rapid stimulation, similar to the recovery from inactivation at negative potentials.
Potentiation of Nav1.6 results from faster activation
The data thus far suggest that the increase of Nav1.6 with ß1 during rapid stimulation might be caused by increasing channel activation. To examine the kinetics of activation, we removed fast inactivation from both Nav1.2 and Nav1.6 by replacing the critical isoleucine, phenylalanine, and methionine (IFM) residues in the III-IV linker inactivation particles with three glutamine residues (West et al., 1992). The noninactivating channels were termed Nav1.2Q3 and Nav1.6Q3. Activation was elicited from a holding potential of –120 mV by 200-ms depolarizations to potentials ranging between –80 and 50 mV (Fig. 6). Sample traces from oocytes expressing Nav1.2 and Nav1.2Q3 -subunits are shown in Fig. 6, A and B. The currents through Nav1.2 were fully inactivated within 100 ms, whereas there was no significant inactivation of Nav1.2Q3 current during the 200-ms depolarization. Similarly, currents through Nav1.6 were fully inactivated within 100 ms and there was no significant inactivation of Nav1.6Q3 current during the 200-ms depolarization (Fig. 6, C and D). Nav1.2Q3 and Nav1.6Q3 demonstrated similar voltage dependences of activation that were more negative than those for Nav1.2 and Nav1.6 (Fig. 4 A and Table 2). Coexpression of ß1 did not affect the voltage dependence of activation for Nav1.2Q3 nor that of Nav1.6Q3, whereas the voltage dependences of activation for Nav1.2 and Nav1.6 were both shifted toward the values of the noninactivating mutants in the presence of ß1 (Fig. 4 B and Table 2). These data suggest that both the differences in the voltage dependence of activation between wild-type Nav1.2 and Nav1.6 and the effects of the ß1-subunit on the voltage dependence of activation are due to fast inactivation.
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-subunit channel (Fig. 6 E, dotted line) activated significantly faster than the Nav1.6Q3 -subunit channel (solid line). Coexpression of the ß1-subunit speeded up the activation of both channels so that activation appeared equivalent on this timescale (Fig. 6 F). There was no significant inactivation of Nav1.6Q3 with ß1 (Fig. 6 F, solid line), but the Nav1.2Q3 with ß1 channel showed a decrease in current of 20% by the end of the 200-ms depolarization (dotted line), presumably due to slow inactivation. These data suggest that Nav1.2 enters the slow inactivated state more rapidly than Nav1.6. The responses of the noninactivating channels to rapid stimulation were examined using the protocol shown in Fig. 2 A. Nav1.2Q3 with ß1 showed a slight increase in current during the first 10 depolarizations of rapid stimulation, after which the current decreased with successive depolarizations (Fig. 7 A, open squares). Currents through Nav1.6Q3 with ß1 also showed an increase during repetitive stimulation, but in this case the increase was larger and it continued for 30 depolarizations (Fig. 7 B). The increase was followed by a decrease in current, similar to the results with Nav1.2Q3. The decrease in current for both Nav1.2Q3 and Nav1.6Q3 with ß1 did not appear to stabilize, in contrast to the decrease for Nav1.2 with ß1 (Fig. 2 B). Because fast inactivation had been eliminated in Nav1.2Q3 and Nav1.6Q3, the continuing decrease in current was most likely due to accumulation of slow inactivation.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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; Spampanato et al., 2001). The extent of Nav1.6 potentiation increased with increasing depolarization times up to 30 ms, and it decreased with increasing times between depolarizations. The voltage dependence of potentiation correlated with the voltage dependence of activation, and the kinetics of activation were accelerated after rapid stimulation. One mechanism to explain the potentiation of Nav1.6 with ß1 is that repetitive depolarizations cause faster channel activation. Four characteristics of the use-dependent potentiation are consistent with this hypothesis. First, the voltage dependence of potentiation correlated with the voltage dependence of activation (Fig. 4). Second, potentiation occurred with sodium channels in which fast inactivation had been removed (Fig. 7 B). Third, the potentiation of Nav1.6 with ß1 increased with increasing depolarization times up to 30 ms (Fig. 4 A), and activation of Nav1.6Q3 with ß1 continued to increase until at least 20 ms (Fig. 7 D). Finally, the kinetics of activation for Nav1.6Q3 plus ß1 were faster when potentiation was maximal compared to the kinetics during the first depolarization (Fig. 7 D and Table 3).
Rapid stimulation resulted in both faster activation and faster inactivation of Nav1.6 sodium channels. A single mechanism that could account for acceleration of both activation and inactivation is increased synchronization, because inactivation is coupled to activation in neuronal sodium channels (Aldrich et al., 1983). Sodium channels pass through multiple closed states before opening, which causes a delay in channel opening after depolarization (Armstrong and Gilly, 1979; Keynes and Elinder, 1998; Vandenberg and Bezanilla, 1991). Rapid stimulation might drive the channels into a later closed state and they don't revert to the most extreme closed state between depolarizations, which could result in more synchronous opening and inactivation.
The fact that potentiation occurred with only Nav1.6 and not with Nav1.2 probably reflects differences in both the activation and slow-inactivation kinetics of these two channels. Although both Nav1.2Q3 and Nav1.6Q3 coexpressed with ß1 demonstrated faster activation after repetitive depolarization, the acceleration was more pronounced for Nav1.6Q3. This difference might reflect the fact that activation of Nav1.6Q3 was slower than that of Nav1.2Q3. In addition, only Nav1.6Q3 demonstrated a slow component of activation with ß1, and it was this component that was accelerated most dramatically by rapid stimulation (Table 3). The kinetics of activation for Nav1.2 with ß1 may already be close to the maximal rate, so that repetitive depolarizations did not have much of an effect. In addition to the differences with respect to activation, there was a significant difference between the two channels with respect to slow inactivation. Slow inactivation of Nav1.2Q3 was much faster than that of Nav1.6Q3, and ß1 accelerated the slow inactivation of Nav1.2Q3 while slowing the slow inactivation of Nav1.6Q3. Therefore, the fact that potentiation of current through Nav1.2Q3 with rapid stimulation was transient could be due to channels entering the slow inactivated state. Nav1.6Q3 showed a similar but delayed decrease in current, most likely because slow inactivation in Nav1.6Q3 was less rapid than slow inactivation in Nav1.2Q3. The fact that potentiation continued for 200 depolarizations for Nav1.6 but was transient for Nav1.6Q3 may indicate that fast inactivation affects the kinetics of slow inactivation, either by slowing entry into the slow inactivated state or accelerating recovery from that state.
The differences between Nav1.6 and Nav1.2 in their responses to repetitive depolarizations may be important for the physiological roles of the different channels. Nodes of Ranvier develop at sites that contain clusters of sodium channels. The initial clusters contain Nav1.2 and the ß2-subunit (Kaplan et al., 2001), but Nav1.2 is then replaced by Nav1.6 when myelination occurs (Boiko et al., 2001). In the mature neurons, Nav1.6 is localized at nodes of Ranvier whereas Nav1.2 is localized in the unmyelinated regions (Boiko et al., 2001). It may be advantageous to have sodium channels that can faithfully propagate high frequencies of action potentials without loss of amplitude. Slow inactivation could decrease the amplitude or frequency of the spike train and could also result in fewer channels being available for the next wave of stimulation, which might cause a loss of information transmission. The difference in response of the two channels to repetitive depolarizations suggests that Nav1.6 with ß1 would be more able to keep up with multiple rapid stimulations compared to Nav1.2.
Colbert et al. (1997) have shown that sodium current in the soma and dendrites of CA1 pyramidal neurons decreased to 37 and 73% of their original levels during 10 2-ms depolarizations to –10 mV at 20 Hz. They attributed the current decrease to slow inactivation of the channels. Compared to these results, the potentiation that we observed for Nav1.6 with ß1 during 2-ms depolarizations at 10 Hz was quite small, and the current decrease for Nav1.2 with ß1 was <1%. These differences probably reflect differences in the two sets of experimental conditions. Colbert et al. (1997) recorded from neurons at 37°C. Gating is much faster in those conditions, with full activation and inactivation within 2 ms compared to 20 ms in oocytes at 20°C. In addition, neurons contain multiple sodium channel isoforms including Nav1.1, Nav1.2, and Nav1.6 and associated ß-subunits, whereas we recorded from oocytes expressing a single sodium channel -subunit isoform. Despite the quantitative differences between our results and those obtained from neurons, it is probable that the qualitative differences that we observed between the responses to repetitive depolarizations of Nav1.6 and Nav1.2 sodium channels will be observed in vivo. However, a true test of the physiological significance of Nav1.6 use-dependent potentiation requires recording from neurons expressing isolated populations of either Nav1.6 or Nav1.2.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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This work was supported by the National Institutes of Health (grants NS26729 and NS48336). Wei Zhou was supported by a fellowship from the American Heart Association.
Submitted on May 12, 2004; accepted for publication September 22, 2004.
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REFERENCES |
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Spampanato, J., A. Escayg, M. H. Meisler, and A. L. Goldin. 2001. Functional effects of two voltage-gated sodium channel mutations that cause generalized epilepsy with febrile seizures plus type 2. J. Neurosci. 21:7481–7490.
Vandenberg, C. A., and F. Bezanilla. 1991. A sodium channel gating model based on single channel, macroscopic ionic, and gating currents in the squid giant axon. Biophys. J. 60:1511–1533.
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; n = 8) 
; n = 5), Nav1.2Q3 (•; n = 4), and Nav1.6Q3 (
; n = 3) 
) during 200 depolarizations from –100 to –10 mV, each lasting 2 ms. The current amplitudes were normalized to the peak current amplitude during the initial depolarization. The data were recorded using the cut-open oocyte voltage clamp as described in Materials and Methods. (C) Average normalized current increase levels with different time intervals between depolarizations during rapid stimulation. The current increase for different time intervals between depolarization was compared to the change in current with a 2-s interval, because there was no significant potentiation in current with a 2-s interval. The amount of increase was normalized to the maximal increase in current, and the data from different oocytes were averaged and plotted against the time interval (n = 5). Recordings were obtained in ND96 using a two-electrode voltage clamp. The symbols represent the means and the error bars indicate the standard deviations.
