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Physiologisches Institut, University of Kiel, Kiel, Germany
ABSTRACT I. INTRODUCTION A. Scope B. Terminology II. PHENOMENA A. Time Course of (Fast) Inactivation B. Differences Between Channel Isoforms C. Slow Inactivation D. Single-Channel Results E. Temperature Effects III. INTERPRETATIONS A. Kinetic Models B. Gating Currents and Inactivation IV. MOLECULAR MECHANISMS A. Channel Structure and How to Relate It to Function 1. Structure 2. Methods B. Localization of the Gate Mediating Fast Inactivation C. Localization of Site(s) Responsible for Slow Inactivation D. Importance of the {beta}-Subunits V. CHEMICAL MODULATION A. Modulation by Toxins and Local Anesthetics 1. Site 2 toxins: veratridine, batrachotoxin, grayanotoxin, and aconitine 2. Site 3 toxins: scorpion, sea anemone, and spider toxins 3. Site 5 toxins and persistent sodium current 4. Site 6 toxins: {delta}-conotoxins 5. Other toxins and agents affecting inactivation A) OTHER POLYPEPTIDE TOXINS. B) INSECTICIDES. C) CHLORAMINE-T. D) N-BROMOACETAMIDE. E) GLUTARALDEHYDE. F) IODATE. G) POSITIVE INOTROPIC AGENTS. H) PROTEOLYTIC ENZYMES. I) FREE FATTY ACIDS AND PHOSPHOLIPIDS. 6. Local anesthetics preferentially bind to inactivated channels B. Toxins and Receptor Sites 1. Molecular determinants of toxin binding site 2 2. Molecular determinants of toxin binding site 3 3. Molecular determinants of the binding sites of local anesthetics 4. Binding sites for agents of low molecular weight A) CT. B) NBA. VI. GENETIC MODULATION: CHANNELOPATHIES A. Heart Muscle B. Skeletal Muscle C. Central Nervous System VII. CONCLUSION ACKNOWLEDGMENTS REFERENCES
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ABSTRACT |
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-subunits of the channel molecule. Systematic studies of these modulating factors and of the effects of point mutations (experimental and in hereditary diseases) in the channel molecule have yielded a fairly consistent picture of the molecular background of fast inactivation, which for the slow inactivation is still lacking. |
I. INTRODUCTION |
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The subject of this article is the voltage-gated sodium channel that plays a key role in membrane excitation and is dealt with in an enormous number of papers. Inactivation (defined in sect. IIA) of this channel appears to be its most vulnerable kinetic feature as it is influenced, mostly slowed or abolished, by all kinds of chemical agents such as drugs, toxins, or mutations, often of only a single amino acid residue of the channel molecule. The amino acid sequence of the sodium channel molecule and its transmembrane topology have been known for quite some time. Nevertheless, relating function to structural details is still unsatisfactory, but site-directed mutations are increasingly employed to solve such questions. The molecular exploration of hereditary diseases has helped much to identify relevant regions.
Papers dealing with sodium channels, even those that are limited to inactivation, are so numerous that often only summaries, in particular of older work, can be quoted. Reviews of more recent papers on structure and function of sodium channels have been written by Catterall (67, 68), Denac et al. (106), Fozzard and Hanck (130), Marban et al. (272), and Ogata and Ohishi (331); extensive reviews of kinetic aspects of inactivation by Patlak (341) and of mechanisms by Goldin (146) have appeared. Highly readable accounts are found in books on ion channels in general (5, 181). Other reviews will be quoted in several other sections.
In this paper inactivation is first described as observed in electrophysiological experiments including patch-clamp studies. These are followed by interpretations by kinetic models and gating current experiments. Then the structure will be dealt with on a molecular level as derived from experimental point mutations. Next chemical modulation will be described and finally genetic modulation, observed in hereditary diseases (channelopathies).
There exist many isoforms of the sodium channel (144) that have been differently termed. To eliminate the confusion a panel of leading researchers has agreed on a uniform nomenclature following that for potassium channels (147). These terms will be given, at least once, in parentheses to those used in the original papers.
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II. PHENOMENA |
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The typical voltage-gated sodium channel opens on depolarization and closes rapidly on repolarization or, more slowly, on sustained depolarization. The latter process is termed inactivation and leaves the channel refractory for some time after repolarization. In the classical study of the squid giant axon, Hodgkin and Huxley (184, 185) described the sodium conductance (channel gating) being proportional to m3 · h, where m and h are variables of time and potential that can assume values between 0 and 1: m increases and h decreases on depolarization. This leads to a nonmonotonic time course with 1 – h expressing inactivation whereby h develops exponentially as h(t) = h
– (h – ho) exp (-t/
h) with ho and h the starting and final value of h, respectively, and h the inactivation time constant. For large depolarizations h becomes 0, meaning that inactivation reaches completion.
Changes in methods led to results that necessitated different formalisms. Thus in squid axons internally perfused with NaF solution, Chandler and Meves (75) found inactivation to be incomplete (h = 
0.1) and at very positive membrane potentials h increasing to 0.3 or more. These authors explained their results by assuming that h is the sum of two components, h = h1 + h2, with the components connected through an inactive (closed) state. Only during very long depolarizing pulses lasting for several seconds did the "maintained" current inactivate, which Chandler and Meves (76) described by an additional variable s so that h = (h1 + h2)s. Such slow inactivation of varying time constants is observed in many preparations as will be shown below; in KF-perfused squid axons, it gives rise to action potentials lasting for seconds. On blocking IK, a noninactivating fraction of INa was later found also in nonperfused squid axons (412). It was further studied at very positive membrane potentials and with an inverse Na+ gradient (92). The voltage dependence of this fraction has again been examined in more detail recently (359), but the conclusions have been questioned (88).
In amphibian myelinated nerve fibers, inactivation was originally studied with short impulses and could be described by a monoexponential process (131). When in this preparation the potassium current was eliminated with tetraethylammonium ions (TEA), or by other means, the sodium current could be followed for any length of time. Inactivation then turned out to be diphasic (see Fig. 4, trace "toxin free") and so was recovery from inactivation (84) which was interpreted by a second inactivating state in series. Several authors observed diphasic inactivation (see Ref. 230), but some interpreted it differently, recognizing that recovery from inactivation posed the more difficult part of description. Ochs et al. (330) tested several three-state models and found their results to be best fit with two open states connected through a closed one. To account for biphasic tail currents on repolarization, Elinder and 
rhem (119) suggested another model with two open states leading to two different inactivated states. Schmidtmayer (394) formulated a cyclic three-state model with one open and two closed inactivated states. Whereas these models were based on the idea of a single population of sodium channels, Benoit et al. (35) assumed that there are two, however interconvertible, types of sodium channels that differ not only in their inactivation kinetics but also in their susceptibility to blocking agents.
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(V) curve was clearly shifted to more negative potentials for weaker test pulses (151). Such shift had been predicted by Hoyt (197) for activation-inactivation coupling. Another point in favor of coupling was seen in a delayed onset of inactivation as determined with a constant test pulse following a conditioning pulse of constant amplitude but varying duration. Such delay was reported for Myxicola axons (151) and squid axons (45), but in the latter preparation, the delay could be minimized by leaving a gap between conditioning and test pulse to allow activation to subside (140). This does not seem to apply to Myxicola axons in which introduction of a gap did not eliminate the delay (148, 149, 150) nor did it in neuronal and cardiac channels (42). Kniffki et al. (230) showed that in toad nodes of Ranvier the initial delay is due to the activation during the conditioning pulse and implicit in the classical m · h description. Thus the situation is complicated, moreover since the kinetic experiments are prone to flaws in method, some of which were reviewed by Meves (282). The question of whether or not inactivation is coupled to activation is of course essential for understanding the mechanism of inactivation. A great number of kinetic models have been proposed that attempt to reconcile results on bulk sodium currents with so-called gating currents (see sect. IIIB) or single-channel recordings (see sect. IID).
B. Differences Between Channel Isoforms
Activation and inactivation characteristics like Vm, the potential at which activation reaches half-maximal values, and Vh, the potential at which h = 0.5, differ in different isoforms, but also among species and, with cloned channels, depend on the cells in which they are expressed (269; see below). Thus, in Na+ channels of human heart (hH1 = Nav1.5), skeletal muscle (hSkM1 = Nav1.4), and rat brain (rIIA = Nav1.2), expressed in mammalian tsA201 cells, Vm = –48, –28, and –22 mV and Vh = –92, –72, and –61 mV, respectively. The kinetics of activation and inactivation are also quite different. Thus t1/2, the time to reach half of their maximal current amplitude, values are 0.82, 0.48, and 0.40 ms for the three isoforms at 20°C (Ref. 333, containing ample references). The time constants of the rapid component of inactivation decrease with depolarization and reach asymptotic values at V >20 mV of 0.41 ms (hH1), 0.26 ms (hSkM1), and 0.27 ms (rIIA). Clearly at positive potentials, where channels inactivate mostly from the open state, heart channels do so more slowly. Recovery from inactivation is also quite different: rec, measured at V = –100 mV, was 44.7 ms (hH1), 4.7 ms (hSkM1), and 7.6 ms (rIIA). Extensive descriptions of these isoforms (70) and their evolution (142) have recently been compiled.
After prolonged depolarization, often achieved by changes of holding potential in the voltage clamp, recovery from inactivation may proceed very slowly with time constants in the second to minute range and in several phases. Such changes have been termed "slow" or "ultraslow" inactivation and have been reported for amphibian myelinated nerve fibers (52, 129, 238, 318, 458), frog muscle (12), frog ventricular myocytes (132), and rat muscle (382, 415). Slow changes in INa following changes in membrane potential have also been observed in lobster, crayfish (378), Myxicola (392), and squid (1, 75, 76, 287) axons, in the latter preparation even after block of fast inactivation by intra-axonal application of pronase (379).
Slow inactivation was also observed in mammalian neuronal cells, such as rat hippocampal neurons (288), tetrodotoxin-resistant cells of dorsal root ganglia of rat (332), or cultivated neuroblastoma cells (353). In human cardiac muscle (471), slow inactivation is present but is only 40% complete compared with 80% observed in channels of human skeletal muscle (367).
Fast inactivation is important for action potential repolarization, and in mammalian nodes of Ranvier, which almost lack phasic potassium channels, it is the only repolarizing force besides the leakage current (85). Slow inactivation, on the other hand, may play a role in regulating excitability (383), such as by modulating burst discharges as can be demonstrated by computations (120). In reality, however, this modulation appears to be complicated since slow inactivation not only depends on resting potential but also on previous history of action potential firing (288, 448). Also, persistent INa of tetrodotoxin (TTX)-resistant channels (100) may affect the resting potential as suggested by computer simulations (175).
Single-channel records of normal sodium channels were preferentially obtained from cell-attached neuroblastoma and myocardial cells. They show one or a few short openings at the first 10–20 ms (at room temperature) after the start of a depolarizing impulse which, if kept on, leads the channels in an inactivated state. An early thorough analysis of single neuroblastoma channels revealed a fast "microscopic" inactivation from the open state even in the case of slow "macroscopic" inactivation as expressed by h at smaller depolarizations where a slower activation is rate limiting (6, 7). The underlying scheme predicts that removal of inactivation would unmask the slow activation reaching its peak much later than INa of untreated cells. This was indeed observed in neuroblastoma cells intracellularly treated with papain (153) but seems to be the exception rather than the rule, since other channel subtypes show a fast activation combined with relatively slow inactivation.
In some heart cells, openings lasting clearly longer than in neuronal cells were observed that correspond to a slow inactivating state as illustrated by ensemble-averaged records; even a transition from fast to slowly inactivating state was observed as well as a persistent state (48, 49). In rat skeletal muscle, slow inactivation manifests itself as a decreasing number of channels of unchanged open time and conductance (380).
An extensive single-channel study has also been done on cells of the rat entorhinal cortex which generate a persistent current caused by prolonged and often delayed bursts (267). Open times within these bursts (but not interburst closed times) were strongly voltage dependent. In neuroblastoma cells, multiple, longer, and late openings were seen only in channels modified by batrachotoxin (354), sea anemone toxin, scorpion toxin, or chloramine-T (226, 303, 308, 323). Sea anemone toxin prolonged the open state (including repeated openings) in cardiomyocytes (121, 402) as well as in cloned myocardial channels (hH1 = Nav1.5; Ref. 74). In a comparative study on single cardiac and neuronal channels, modified by sea anemone toxin, initial clusters of multiple openings and periodic subsequent reopenings were observed in either preparation. However, toxin induced a larger persistent current in neuronal than in cardiac channels (42).
Similar single-channel phenomena have been observed in mutant channels with slowed inactivation (280) whereby particularly delayed inactivation is accompanied with very long mean open times (244, 417). Mutant channels completely deficient of inactivation (IFM/QQQ; see sect. IVB) showed repeated openings and closings throughout long depolarizing pulses (159). In almost all these experiments, single-channel conductance was unaffected by modifications of kinetics. Analysis, however, may be complicated by the existence of conductance substrates and changes following excisions of membrane patches (see Ref. 309).
In their classical description of ion channel kinetics in squid axons, Hodgkin and Huxley (185) assumed a temperature coefficient, Q10, of inactivation of 3. Similar results have since been obtained with many other preparations (181). Hence, in comparing time constants it is necessary to consider the temperature at which they were measured. Usually the Q10 values of the time constant(s) increase at lower temperatures as one would expect if kinetics are described by a simple Arrhenius plot of log h vs. T–1. However, in some preparations such as skeletal muscle, a break in this plot is observed which may point to a transition in the lipid membrane phase (85, 405), complicating the interpretation in terms of activation enthalpy. In rat nerve, the steady-state inactivation h(V) was found to be shifted in the hyperpolarizing direction on cooling (404), so was h(V) in Xenopus nerve (207), whereas in rat muscle only the steady-state curve of slow inactivation s(V) was similarly shifted; this may explain the reduced availability of mammalian sodium channels if tested at room rather than at body temperature (381).
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III. INTERPRETATIONS |
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As already mentioned in section IIA, the classical description of sodium channel gating by Hodgkin and Huxley (184, 185) implied activation and inactivation to be independent processes, linked only by their dependence on membrane potential. However, later results such as the existence of more than one inactivation component, single-channel results, and the discovery of four nonidentical domains to form the sodium channel 
-subunit required extended models, competently reviewed by Hille (181). Most of these models propose that inactivation derives its voltage dependence from that of activation. However, often amino acid substitutions in the voltage sensors shift h(V) in the opposite direction compared with shifts in m(V), arguing against a strict activation-inactivation coupling (236).
The general concept is that on depolarization a channel moves from (several) closed (C) resting state(s) through an open (O) state to one (or several) inactivated (I) state(s). Gating is assumed to be a Markov process meaning that the rate constants of transitions between these states are "oblivious" of how a given state is reached. A critical feature to model is the recovery from inactivation that does not seem to pass through the open state as no ionic current is usually seen during this period. An exception is observed in cerebellar neurons that produce a "resurgent" current on repolarization (2, 360). This current is assumed to reflect unblocking of an open-channel block in a portion of channels by a hypothetical particle during strong depolarization, constituting an additional mechanism of inactivation. It thus enhances recovery from inactivation, which enables these neurons to fire at high frequency (360). In most other channels the details of recovery require C
I and IC steps leading to cyclic connections of states. Such cycles underlie the restraint of microscopic reversibility so that the product of rate constants going clockwise must equal the product going counterclockwise. In the following scheme 1 this would mean k34kOIkIC = k43kCIkIO.
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It should be noted that the classical Hodgkin-Huxley formulation m3 · h can be restated as a consecutive movement through eight states as in scheme 3 with k12 = 3m, k21 = m, k23 = 2m, k32 = 2m, k34 = m and k43 = 3m (180); to describe a number of single-channel results, it appears not to be inferior to a model of the type of scheme 1 (78).
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While the kinetic models described so far are based on voltage-clamp experiments that employ square voltage pulses, a different method called "nonequilibrium response spectroscopy" uses rapidly fluctuating (up to 14 kHz) potentials to extract more kinetic details (290).
B. Gating Currents and Inactivation
The potential-dependent gating of channels requires a voltage sensor bearing charges that move within the membrane as its potential changes, generating a tiny "gating current." This became measurable on channel opening and closing after suppression of the much larger ionic current (16, 221), whereas movement of the inactivation gate did not seem to contribute a specific fraction of gating current. Instead, inactivation causes a substantial fraction (2/3) to become immobilized, and immobilization proceeds and recovers with the same time course as does inactivation (17, 286, 287; for reviews, see Refs. 11, 15). This indicates that inactivation is coupled to activation from which it derives most of its voltage dependence (181). Nevertheless, some genuine voltage dependence of inactivation exists (194, 219, 220). The advantage of gating current measurements is to reveal kinetic steps that are not accompanied by changes of ionic currents.
A great number of papers have been published dealing, one way or the other, with the relationship between inactivation and gating current, many of them in K+ channels. As for Na+ channels, more recent studies tried to reconcile gating currents with details of channel structure, in particular the existence of four domains, D1–D4, each with six segments of which S4 acts as voltage sensor (see sect. IVA). Although segments S4 of all domains seem to be involved in activation (237), D4S4 clearly has the largest effect on inactivation (236), and mutation of central arginines has specific effects on inactivation and gating charge immobilization in rNav1.2 (239). S4 immobilization in D3 and D4 (but not in D1 or D2) has also been observed by site-directed fluorescence labeling (73).
Another approach is to compare effects of modulating agents on ionic currents with those on gating currents (223). As one would expect, irreversibly removing inactivation in squid axons by chloramine-T (CT) or pronase also stops charge immobilization but with other similarly acting agents like batrachotoxin (BTX) immobilization remains (439). In amphibian nodes of Ranvier, comparable results were obtained with CT (111) but not with BTX, which eliminated immobilization (114) as did internally applied iodate (113). Comparative studies on nodes of Ranvier, also with site 3 toxins, have been summarized by Meves (284). In mammalian muscle channels (rNav1.4) and cardiac channels (hNav1.5), site 3 toxins of Anthopleura, ApA and ApB, reduce the maximal gating charge by about one-third, although muscle channels inactivate clearly faster than heart channels (407). Studies of ApA effects on gating charge in mutant hNav1.5 revealed the importance of the outermost arginine in D4S4 for charge immobilization (409) Charge immobilization has also been achieved in squid axons by intracellular treatment with the positively charged sulfhydryl reagents MTSET (2-trimethylammonioethylmethane thiosulfonate) and MTS-PTrEA {[3(triethylammonium)propyl]methanesulfonate}. These reagents irreversibly modify native cysteine(s) in a potential-dependent manner and promote inactivation from closed states, whereas they do not affect activation; MTS-PTrEA shifts the steady-state inactivation curve to more negative potential values and renders it much shallower (222).
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IV. MOLECULAR MECHANISMS |
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During the last 20 years the molecular analysis of the sodium channel has made enormous progress through experiments most recently described by Catterall (68). Here it may suffice to summarize our present knowledge. The channel purified from mammalian brain consists of the large -subunit (260 kDa) with the pore, the 1- (36 kDa) and 2-subunits (33 kDa) that contain extracellular immunoglobulin-like folds as illustrated by Figure 1 in section IVB. The -subunit consists of four domains (D1-D4) each with six transmembrane -helical segments (S1-S6) of which S4 bears several positive charges originating from arginine or lysine residues. The proteins of the domains wrap around a central pore such that the P-loops (SS1-SS2) between S5 and S6 form part of the pore lining.
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-helix. The "classical" view has it that negatively charged residues in adjacent segments form ion pairs that stabilize S4 against the pull exerted at rest by the (inside) negative membrane potential. Depolarization releases S4, the voltage sensor, to move outward initiating the structural changes that open the pore (66, 166). This general concept has been varied and adapted to comply with new evidence, in particular gained from the three-dimensional structure of the bacterial potassium channel KcsA as reviewed by Bezanilla (43). Comparable results of sodium channels are soon to be expected as a recently described bacterial channel, NaChBac, may provide the protein in quantities needed for structural studies (365). A mutation, G219P, in S6 of this channel, dramatically reduces the rate of the already very slow deactivation and inactivation and shifts membrane potential (Vm) by –51 mV (521). It seems that this mutation strongly favors bending of the S6 helix and that G219 serves as a gating hinge. A low-resolution three-dimensional structure study of a sodium channel (employing cryo-electron microscopy and image reconstruction) revealed, in addition to the ion-conducting central pore, four peripherally located transmembrane "gating pores" in which the S4 movement is thought to take place (69, 391).
More recently, X-ray crystallography of another bacterial channel, KvAP, yielded results that led to a quite different interpretation of voltage sensor movement. S4 and part of S3 are supposed to form a "paddle" at the periphery of the channel which, on depolarization, moves like a lever through the surrounding membrane lipid from the intracellular to the extracellular side, thereby opening the pore (205, 206). This unconventional view, summarized by MacKinnon (265), is at present controversially discussed on the basis of new experimental results (4, 29, 53, 133, 247; see also comment in Ref. 190). Whatever the outcome, it certainly will affect our interpretation of sodium channel gating.
To derive function from structure, many methods have been employed. Early experiments revealed the sidedness of certain modulating agents or procedures giving valuable information as to where certain functions may be located. Thus, for instance, scorpion toxins act only from outside, pronase treatment only from inside the excitable cell to slow or delete inactivation. Comparable information was obtained with specific antibodies against portions of the channel (see sect. IVB). Decisive progress, however, was achieved by molecular genetics enabling site-directed mutagenesis. Mutants are expressed in Xenopus oocytes but most frequently in mammalian cells such as HEK293 (human embryonic kidney cells) and their derivative tsA201, also in Chinese hamster ovary cells (CHO) or in the Escherichia coli strain BL21. Very successful was replacing single amino acid residues by cysteine to which access could then be tested with sulfhydryl reagents ("substituted-cysteine-accessibility method", SCAM; Ref. 213), whereby access could change with the functional state. Experimental point mutations were employed in most of the papers cited in section IV, B–D, whereas natural mutants found in hereditary diseases ("channelopathies") are described in section VI. In Figure 1, the localization of the most important mutations is marked by black numbers on white, those of channelopathies as white numbers on black background.
B. Localization of the Gate Mediating Fast Inactivation
The intracellular linker between domains D3 and D4 plays an important role as derived from studies with antibodies directed against the linker which completely blocks fast inactivation (39, 461, 462); also, inactivation is lost or slowed if this linker is cut (432) or bears deletions or mutations (217, 344). In the following description, mutants of Nav1.2 are given if not otherwise mentioned. In these Nav1.2 channels, the critical motif is the hydrophobic triad I1488, F1489, M1490 (IFM; "h" in Fig. 1). Inactivation is completely blocked in I1488Q-F1489Q-M1490Q (IFM/QQQ, corresponds to I1303Q-F1304Q-M1305Q in Nav1.4), but mutant F1489Q alone considerably slows inactivation (502) as does the equivalent F1304Q in muscle channels (Nav1.4) for which kinetic modeling led to the assumption of three inactivated states (328). Inactivation is restored by adding short peptides containing IFM (KIFMK-amide) to the intracellular side (117, 118) or in channels in which inactivation is slowed by external application of -toxin of the scorpion Leiurus quinquestriatus (116). SCAM experiments with the mutant F1489C show that F becomes inaccessible to MTSET on inactivation, suggesting that IFM serves as a hydrophobic latch for a hinged lid formed by the D3-D4 linker (216). The NMR solution structure of the isolated inactivation gate (the linker peptide) has been identified as a stably folded core consisting of a -helix capped by an NH2-terminal turn (373). It is supposed that on gate closing the core (the latch) pivots on a more flexible hinge region. Molecular dynamic simulation predicts an additional helical segment in the D3-D4 linker, on the other side of IFM, which possibly helps to guide the inactivation particle towards its receptor (416). Another solution structure study has been done in a medium of low dielectric constant (293).
More recent experiments on mutant muscle channels, Nav1.4, revealed that the charge of residues beyond the IFM motif also plays an important role: reduction of positive charge (K1317N,K1318N) accelerates inactivation kinetics in the absence of the -subunit (289), whereas charge reversal in the presence of the -subunit (E1314R,E1315R) slows open-state inactivation but accelerates closed-state inactivation (165). It is concluded that clusters of negatively and positively charged residues in the D3-D4 linker differentially regulate the kinetics of fast inactivation.
The D3-D4 linker of Nav1.2 (brain) channels also contains a serine, S1506, distal of IFM, whose phosphorylation by protein kinase C (PKC) slows inactivation without an effect on the steady-state curve h(V) (500, 501). Phosphorylation of other sites in the D1-D2 linker reduces INa, an effect that is also observed on phosphorylation by protein kinase A (99, 401). In cardiac Na+ channels (Nav1.5) phosphorylation by PKC at the equivalent site S1505 causes a strong negative shift of h(V) pointing to stabilized inactivation from closed states; the S1505A mutant is almost unaffected by PKC (352). In muscle µ1-channels (Nav1.4), the equivalent serine S1321 does not seem to play a key role in shifting h(V) by PKC, since it is also observed in a S1321A mutant (30).
Fast inactivation of Nav1.2 is also interrupted in mutants F1764A/I1765A/L1766A of D4S6 (corresponding to F1579/80/81 in Nav1.4, 
in Fig. 1), originally thought to form part of the hydrophobic latch receptor (278). Later experiments with F1764A/V1774A excluded a direct interaction with the IFM motif, since application of KIFMK-amide nevertheless restored inactivation (279). KIMFK restores fast inactivation of open but not of closed channels in the F1651A/L1660A mutant in the D4S4-S5 intracellular loop (approximately at 
in Fig. 1). This suggests that the IFM motif interacts with the loop during inactivation of closed channels (280). In Nav1.4 cysteine substitution, F1579C, inhibits both fast and slow inactivation, whereas Y1586C and I1575C enhance them (21). Interestingly, these residues belong to the LA binding site, which hence appears to be involved in both types of inactivation.
The D3-D4 linker, however, is not the only determinant of fast inactivation. Thus if in the more slowly inactivating heart channel (Nav1.5) the linker is replaced by that of the faster brain channel (Nav1.2), its properties are not transferred (171). On the other hand, replacing the COOH terminals accelerates inactivation but also magnifies the differences in voltage dependence of the steady-state inactivation (270). Similar results have been obtained with Nav1.4/Nav1.5 chimeras (108, 109). In a more detailed study of the heart channel COOH terminal, it was shown that truncation of its distal part only reduces current density. Truncation of the proximal part, consisting of six helices, additionally shifts the inactivation curve and clearly increases the fraction of noninactivating channels (91). Further experiments with mutants, in particular truncation of the highly charged sixth helix which increases the persistent INa, suggest that the COOH terminal stabilizes inactivation and minimizes channel reopening (301). The COOH terminal contains a binding site of calmodulin, which is important for the functional expression of channels. In Nav1.6 calmodulin also slows inactivation in a calcium-dependent manner (176).
Important insights have also been gained from experimental changes of the voltage sensor segment, S4. Point mutations of charged amino acids in S4 to cysteine changed the kinetics of deactivation from open and inactivated states in a domain-specific fashion (164). In heart channels (Nav1.5), charge-neutralizing or -reversing substitutions shifted the activation curve m(V) and decreased its slope as had already been demonstrated for D1S4 in the pioneering work of Stühmer et al. (432). Inactivation time constants were markedly decreased only in D4S4 mutations (Ref. 82; see also sect. IVB). In rat brain channels (rNav1.2), addition of a positive charge just external of the outermost positively charged residue of D4S4 (F1625R or F1625K) led to a split of h(V) into two components and a large shift towards hyperpolarization but only to a shift in neutral mutants. These and other findings add to the idea that the D4S4 movement controls the inactivation gate, whereby the countercharges may play an important role in the D4S4 position (512).
The still poorly understood connection between sensor movement and gating has recently been subject of several "perspective" papers (44, 134, 189, 243). It should be mentioned that the "paddle" hypothesis leaves the problem of the sensor-gate coupling unresolved.
C. Localization of Site(s) Responsible for Slow Inactivation
The mechanism of slow inactivation is more complicated, located in structures distinct from those of fast inactivation, and it is less well understood (472). Since it bears similarities to the C-type inactivation of potassium channels, which is connected to the external pore lining, corresponding residues in sodium channels have been the target of site-directed mutagenesis. Thus, in the absence of 1, mutant W402C (P-region of D1, 
; Ref. 22) eliminates slow inactivation, whereas mutation of the adjacent residue, E403C or E403R, seems to favor entry into the slow inactivation state (520) as does W434A (492). E403 together with E758, D1214, and D1532 at comparable positions in domains 2, 3, and 4, respectively, form an outer ring of charges whose structural rearrangement was found to be associated with slow inactivation (507). Also, slow inactivation alters the accessibility by the positively charged MTSEA (2-aminoethylmethanethiosulfonate) to the outer pore cysteine of mutant F1236C (P-region of D3,
), pointing to a structural rearrangement which, incidentally, may be linked to use-dependent LA action (334). A mutant of the adjacent K1237S or E leads to a very slow ("ultra-slow") type of inactivation (447). Such very slow inactivation is also enhanced by mutant A1529D in the P-loop of D4, part of the putative selectivity filter (177).
As already mentioned, slow inactivation of human cardiac sodium channels (hNav1.5) develops more slowly and is only 40% complete versus 80% in skeletal muscle channels (Nav1.4) (367). Chimeras of domains of these two channels show that slow inactivation can be modulated by all four domains, with D1 and D2 being more prominent (335). Interestingly, a single residue in D2S5-S6 (P-region), V754 of Nav1.4 and the corresponding I891 of Nav1.5, confers their parental properties to the chimera; however, considering the other experimental evidence, it seems unlikely that the P-region is directly involved in slow inactivation (470). Also, mutant V787K (D2S6; 
) markedly enhanced slow inactivation, whereas with V787C it was further slowed, incomplete, and less voltage dependent than in wild type (336). Mutants Y401C and G1530C in the P-region of D1 and D4, respectively, were modified by MTSET at the same rate during slow inactivation as in the noninactivated state, which suggests that the outer mouth of the pore remains open in either state (430).
In mutant channels devoid of fast inactivation, slow inactivation remains intact (127, 463). In such mutants, substitution of positive charges in S4 of D1 or D2 shifts the slow inactivation to more positive potentials, whereas with intact fast inactivation, S4 mutations in D2 and D3 cause a negative shift of slow inactivation; obviously fast and slow inactivation interact (236), but the extent is differently interpreted (127, 450, 463). Coupling between fast and slow inactivation has also been proposed from analysis of point mutation F1403Q in Nav1.4 (328). Also, mutation L1482C in the D4S4-S5 loop of human muscle Na+ channels disrupts fast but enhances slow inactivation (8). Mutants of charged residues around the fast-inactivation IFM particle, D1309Q or R and EE1314,15RR, right-shift the steady-state relation between slow inactivation and membrane potential, suggesting that these residues may interact with the structures that control slow inactivation (275).
Further studies of the interaction between fast and slow inactivation were done on mutations in segment D4S4 (which is likely to be involved in the coupling) with the method of cysteine modification. Fast inactivation of mutant R1454C (near 
) is similar to that of wild type but becomes much slower on modification by MTSET and MTSES (2-sulfonatoethylmethanethiosulfonate) (510). However, only treatment with the latter (negatively charged) reagent also affects slow inactivation, whereby peak current is reduced ("use dependent") during trains of depolarizing pulses (291). Interestingly, this effect is partially counteracted by a point mutation in the P-region of D1 (W408A) which, in the presence of the 1-subunit, shifts activation to more negative potentials, hastens fast inactivation (449), and attenuates recovery from slow inactivation (211).
D. Importance of the -Subunits
In Xenopus oocytes, coexpression of the -subunit of Nav1.2 with 1-subunit of neuronal and skeletal muscle channels increases the current density, accelerates inactivation, and shifts the steady-state inactivation curve in the hyperpolarizing direction (202, 343) as illustrated by Figure 2. Comparable results have been obtained with Nav1.8 and, to a lesser extent, with Nav1.7, channels found in dorsal root ganglia (466). Coexpression also accelerates recovery from inactivation (32, 62). It seems that the -subunit has a slow and a fast gating mode leading to diphasic inactivation. The latter mode is favored on binding of 1 whereby the inactivation kinetics of either component does not change (296). An opposite effect, slowing of inactivation and inducing a persistent INa, is observed by coexpression of G protein 
-subunits with Nav1.2 (264). -Subunits expressed alone in mammalian cells inactivate almost as fast as the native preparation (203), which has been attributed to an endogenous splice variant, 1A, present in these cells (299). This hypothesis was subsequently rejected on the grounds of experiments with antisense oligonucleotides (297). An additional subunit, 3, predominant during development (406), has been found that accelerates inactivation but less than 1 with which it is closely related (300). The distribution of 3 in human tissues and comparative amino acid sequences of 1, 2, and 3 is described in Stevens et al. (426). Most recently, a novel disulfide-linked subunit, 4, has been identified which shows similarities with 2; on coexpression it shifts the activation curve in the hyperpolarizing direction without affecting inactivation (516).
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2-subunit does not modulate hH1 (=Nav1.5) or IIA (=Nav1.2) and 1/2 chimeras, expressed in Xenopus oocytes, have served to determine the regions of 1 necessary for modulation: hH1 channels via the transmembrane portion ("membrane anchor") plus additional regions, IIA channels via the extracellular region only (276, 277, 522) as has been suggested before (79).
These effects are not yet fully understood, but the functional domains for the interaction seem to be highly conserved (62, 343). Further experiments revealed that only the extracellular domain of 1 is essential for the interaction (79, 277), whereby segment D4SS2-S6 of the -subunit plays an important role (see broken-line connection in Fig. 1; Ref. 351). Earlier studies pointed to this region: inactivation of human heart channels (hH1 = Nav1.5), in contrast to skeletal muscle channels (hSkM1 = Nav1.4), is not much accelerated on coexpression (in Xenopus oocytes, Ref. 329) with 1, but chimeras containing only D4S5-S6 of hSkM1 show the typical acceleration (268). As for the increased channel density, it is interesting to note that in HEK293 cells the hH1-1 complex forms already in the endoplasmic reticulum which may facilitate trafficking to the plasma membrane (523).
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V. CHEMICAL MODULATION |
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Various toxins and other chemical agents slow or even abolish inactivation. Several groups of toxins have been characterized by binding studies that originally led to the definition of five binding sites (47, 67, 495) that were eventually extended to nine sites (524). Experiments on mutant channels (see sect. IV) reveal that toxins within one binding group often do not occupy identical but rather overlapping sites. Also, there exists considerable allosteric interaction between the sites. In the following section, toxin binding to sites 2, 3, 5, and 6 will be discussed as well as other chemicals that affect Na+ channel inactivation.
1. Site 2 toxins: veratridine, batrachotoxin, grayanotoxin, and aconitine
The toxins of this group are lipid soluble, which enables them to access binding sites embedded in the membrane. The most important of these compounds are veratridine (VT), an alkaloid from lilaceous plants, BTX, which is secreted by the skin of Colombian arrow-poison frogs, aconitine (AC) from plants of the buttercup family, and grayanotoxin (GTX) from plants of the heather family. Although these toxins differ widely in their structure, their common effect is to keep sodium channels open, hence they are termed "agonists." The underlying mechanism is a large shift of activation in the hyperpolarizing direction and a slowed or even abolished inactivation. Also, the toxins clearly bind to open channels. Details are found in reviews (64, 106, 224, 294, 455) and in the book of Hille (181).
A typical VT (60 µM) effect is illustrated by Figure 3, which was obtained with long depolarizing impulses on a frog node of Ranvier. It not only shows the slowly developing Na+ inward current followed by a large slowly decaying current tail (cut off) at the end of the pulse but also the slow current reduction on adding benzocaine during the impulse. In contrast, if benzocaine was applied to Na+ channels of a Ranvier node kept open by CT (see sect. VA5), block was very fast (half-time 60 ms). From this and other results it is hypothesized that channels kept open by VT cannot be directly blocked, and the current reduction shown in trace 2 in Figure 3 is determined by the rate with which VT-modified channels close during the pulse and become susceptible to the local anesthetic (457).
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Obviously these toxins considerably distort the channel since also selectivity for Na+ is reduced. Thus permeability of ammonium ions increases considerably in the presence of VT (251) and in the presence of AC ammonium ions may even become more permeable than Na+ and the relative permeabilities of Cs+, Rb+, and K+ increase (161, 304). Comparable results have been reported under the influence of BTX and VT (135) and AC (362). On the other hand, Li+ that passes native sodium channels readily becomes clearly less permeable under VT treatment (135). Hence, in Li+ Ringer solution, the typical VT-induced afterpotentials are no longer observed (453). Details of altered selectivity caused by VT including ion flux measurements are found in a review (455), that of other site 2 toxins also elsewhere (181).
2. Site 3 toxins: scorpion, sea anemone, and spider toxins
Another binding site has been characterized and termed site 3 to which -scorpion toxins and sea anemone toxins bind (for a review, see Ref. 67). Site 3 toxins have been the subject of many studies so that the reader is referred to a selection of reviews treating toxin structure (156, 218, 326, 347, 370, 498) and/or electrophysiological effects (245, 246, 283, 414, 454). These toxins consist of a polypeptide chain held together by several disulfide bonds; they act from outside the membrane and inhibit inactivation and render it incomplete, but their binding sites seem to be overlapping rather than identical (372). An example of the effect of ATX II, a toxin of the sea anemone Anemonia sulcata, is given in Figure 4, which was obtained in a voltage-clamped frog node of Ranvier at a saturating (with respect to the late current) concentration of 5 µM.
There exists a large sequence homology among the many -scorpion toxins (347) and sea anemone toxins (326) but little between the two. Not only do the toxins of various scorpion species differ with respect to their effects, but also to the target channel isoforms as in amphibian nodes of Ranvier (285, 305, 485), rat skeletal muscle (80) versus heart muscle (81). TTX-resistant sodium channels of dorsal root ganglion cells are also resistant to sea anemone toxins (and other similarly acting toxins) in contrast to TTX-sensitive channels of this preparation (Ref. 389; see below and sect. V, A3, A5, and B).
As for the toxin molecule, subtle changes may yield largely different effects. Thus in neuroblastoma cells, sea anemone toxins APE 1–1 and 1–2 of Anthopleura elegantissima, differing by four amino acids, cause noninactivating currents of 20 and 40%, respectively (54). Likewise, two toxins from another sea anemone (Anthopleura xantogrammatica), ApA and ApB, show great differences in affinity for mammalian (tsA201 cells) channels despite a strong sequence homology, arising solely from different rates, koff, of toxin-channel dissociation (41). In frog nerve ATX I, which differs from ATX II by several amino acids, is ineffective and does not antagonize ATX II (395). Affinity may also be affected: a 20-fold change in EC50 of slowing inactivation of cloned hH1 (=Nav1.5) by Bunodosoma granulifera toxins II and III which differ by only one amino acid (157). The exclusive targets of some -scorpion toxins are insect sodium channels, which makes them potential selective insecticides (155, 524).
Another type of scorpion toxins, termed -like, is toxic to both mammals and insects. Typical representatives are LqhIII of Leiurus quinquestriatus hebraeus and BomIII and BomIV of Buthus occitanus mardochei (72, 80, 137, 138); LqhIII also affects frog axons where, however, it acts like the classical -toxin (37). The -like toxins seem to bind to a site that is differentially related to site 3 (72). The venom of the scorpion Tityus serrulatus contains an interesting toxin, TiTx, which binds to muscle surface channels with a very high affinity (26). In neuroblastoma cells, TiTx reduces peak INa, causes an inward current to flow near the resting potential, but also slows inactivation (468). The toxin thus shows effects of both - and -toxins. Similar results have been observed in Xenopus nodes of Ranvier (208).
Binding of the scorpion -toxins is weaker on depolarization (36, 71, 285, 403, 429, 496), but the authors do not completely agree as to whether the open and/or inactivated state confers the reduced affinity. Toxin II of the sea anemone Anemonia sulcata (ATX II) binding is not reduced on depolarization (285, 473), whereas actions of ATX III and IV on crayfish axons were clearly reduced (497). More recently, voltage dependence of scorpion toxin binding was found in a voltage range where activation and inactivation saturates so that it may originate from other sources (80, 81). Moreover, in steady-state binding experiments to rat brain synaptosomes, depolarization, achieved by high K+ concentration, yielded different kinetic results than short applications.
The toxins not only slow inactivation but render it incomplete (reviewed in Refs. 106, 454), inducing a persistent INa component (36) which is more prominent in neuronal than in cardiac Na+ channels. Also, the toxins enhance the rate of recovery from inactivation through closed states (42).
Slowed and incomplete inactivation is also induced by funnel-web spider toxins that have been shown to bind to site 3 and compete with scorpion -toxins (258, 259), some of them also act on insect Na+ channels (163). These toxins are without effect on the inactivation of TTX-resistant channels of rat dorsal root ganglia (321, 322, 434). The spider toxins 
-atracotoxins form a new family of polypeptides with no similarity to scorpion -toxins (128). -ACTX-Hv1a (formerly versutoxin) is contained in the venom of the funnel-web spider Hadronyche versuta, which also produces the less effective -atracotoxin-Hv1b of no insecticidal effect (434). -ACTX-Ar1 (formerly robustoxin) from Atrax robustus is dangerous to humans; it exhibits a 83% amino acid sequence homology with -ACTX-Hv1a (321). Another spider, Paracoelotes luctuosus, produces the insecticidal -palutoxins with effects similar to those of -scorpion toxins (95). Reviews of spider toxins have been published (94, 162).
3. Site 5 toxins and persistent sodium current
In many preparations a small persistent INa is observed already in normal saline; although small, it may be important in regulating excitability (97, 436, 443). Different channel isoforms with comparable inactivation kinetics nevertheless have persistent components of distinctly different size (83, 104, 267, 525). Some such results have been interpreted as "window" currents flowing in the potential range where steady-state activation and inactivation curves overlap (19, 317, 384), but this interpretation does not fit results in ventricular myocytes (388) and especially in mammalian neurons (13, 97, 214, 340).
In addition to site 3 toxins, another group of toxins, binding to site 5, causes persistent currents including ciguatoxin, which is responsible for the ciguatera fish poisoning. Site 5 toxins are lipid-soluble polyethers produced by dinoflagellates (20, 136). At least in frog nodes of Ranvier, ciguatoxin merely induces a late (persistent) component without affecting inactivation kinetics. Activation of this component is shifted by 30 mV towards hyperpolarization. The size of this component depends on the holding potential (VH) being three times larger at VH = –70 than –120 mV (38). In TTX-sensitive channels of rat dorsal root ganglia neurons, pacific ciguatoxin-1 shifts the activation curve and the steady-state inactivation curve in the hyperpolarizing direction. In TTX-resistant channels, toxin mainly increases the rate of recovery from inactivation (428).
Another group of dinoflagellate toxins that bind to site 5 are the brevetoxins (also polyether molecules), the cause of paralytic or neurotoxic shellfish poisoning. Brevetoxin-3 (PbTx-3), for example, produces a hyperpolarizing shift of the activation curve in TTX-sensitive channels of rat sensory neurons accompanied by an inhibition of inactivation (204). In TTX-sensitive Na+ channels of rat brain, brevetoxin PbTx-3 causes a similar shift of the activation curve which, together with a slowing of inactivation, leads to hyperexcitability (349). Actions of site 5 toxins on ion channels have been reviewed (20, 102, 274, 505). Studies of brevetoxin derivatives to elucidate active centers of the toxin have appeared (136, 204); some of these derivatives act as antagonists (350).
Marine snails of the genus Conus produce a great variety of toxins that act on different ion channels including sodium channels (reviewed in Refs. 125, 445). Many block these channels, but a group of polypeptides, consisting of 26–32 amino acids with 3 disulfide bridges, termed -conotoxins, slows inactivation like -toxins of scorpions but does not bind to site 3. Hence, a new site 6, located at the extracellular side of the membrane, was defined (122; summarized in Ref. 524). The main target of the toxins TxVIA (C. textile), NgVIA (C. nigropunctatis), and GmVIA (C. gloriamaris) is the sodium channel of mollusk neurons (122, 123, 172, 173, 411). Mammalian cells, as contained in rat brain synaptosomes, generally do not respond (but see below), but 22Na influx stimulated by veratridine is further increased (123). Also, TxVIA, lacking electrophysiologically detectable effects on insect axons or frog muscle, nevertheless bind to "silent" receptors on these preparations (410).
Toxins of the fish-hunting Conus striatus retard inactivation in mouse neuroblastoma cells (154), in frog sympathetic neurons (55), and in frog nodes of Ranvier (as does -EVIA) and shift
denotes position of glycosylation. Loops between segments are not to scale. Numbers in white circles give positions of experimental point mutations mentioned in text; numbers on black background are those observed in hereditary diseases. For details see text.
