In excitable cells, the initiation of the action potential results from the opening of voltage-gated sodium channels. These channels undergo a series of conformational changes between open, closed, and inactivated states. Many models have been proposed for the structural transitions that result in these different functional states. Here, we compare the crystal structures of prokaryotic sodium channels captured in the different conformational forms and use them as the basis for examining molecular models for the activation, slow inactivation, and recovery processes. We compare structural similarities and differences in the pore domains, specifically in the transmembrane helices, the constrictions within the pore cavity, the activation gate at the cytoplasmic end of the last transmembrane helix, the C-terminal domain, and the selectivity filter. We discuss the observed differences in the context of previous models for opening, closing, and inactivation, and present a new structure-based model for the functional transitions. Our proposed prokaryotic channel activation mechanism is then compared with the activation transition in eukaryotic sodium channels.

Eukaryotic sodium channels are complex, multidomain proteins that assemble into pseudotetrameric structures. Sodium channels are also present in some prokaryotes (Ren et al., 2001), where they are homologous but simpler, single-domain proteins that assemble to form functional homotetramers (Nurani et al., 2008). Both eukaryotic and prokaryotic sodium channels consist of voltage-sensor (VS) and pore regions that are physically separated in three dimensions. Pore-only channels can be constructed, which are capable of undergoing opening and closing transitions and support ion flux in a manner similar to that of intact channels (McCusker et al., 2011; Shaya et al., 2011). Although the kinetics of their processes of conversion between states are different, both eukaryotic and prokaryotic channels exhibit activation, recovery, and slow inactivation transitions (Koishi et al., 2004; Charalambous and Wallace, 2011); in addition, eukaryotic channels undergo a fast inactivation process that is not observed in prokaryotic channels (Pavlov et al., 2005). Prokaryotic sodium channels have also been shown to be blocked by eukaryotic sodium channel antagonists (Bagnéris et al., 2014).

Many models have been proposed for the structural changes that result in the open, closed, and slow inactivated states of prokaryotic sodium channels (e.g., Kuzmenkin et al., 2004; Zhao et al., 2004; Pavlov et al., 2005; Shafrir et al., 2008). Although there are, as yet, no crystal structures of eukaryotic sodium channels, the structures of several prokaryotic sodium channel orthologues have recently been determined by x-ray crystallography (Payandeh et al., 2011, 2012; McCusker et al., 2012; Zhang et al., 2012; Bagnéris et al., 2013; Shaya et al., 2014). These include the voltage-gated sodium channels from Magnetococcus marinus, formerly known as Magnetococcus spirillium (NavMs); from Arcobacter butzler (NavAb); from Rickettsiales sp. HIMB114 (NavRh); and from Alkalilimnicola ehrlichei (NavAe). These structures now make it possible to observe various states of prokaryotic sodium channels because they have apparently captured the pore regions in partially and fully open (NavMs), closed (NavAb and NavAe), and inactivated (NavAb and NavRh) conformations, thereby enabling structural comparisons that provide new insights into the transition processes.

Because the transmembrane pore regions of the NavMs and NavAb pores have very high sequence identities (∼69%; McCusker et al., 2012; Fig. 1), our comparisons will primarily be confined to these structures because with this level of similarity, differences seen are likely to be state dependent rather than a consequence of sequence dissimilarities. They enable comparisons of structures that we designate as the following: partially (pOpenMs; Protein Data Bank [PDB] accession no. 4F4L; 3.5-Å resolution; McCusker et al., 2012) and fully open (OpenMs; PDB accession no. 3ZJZ, 2.9-Å resolution; Bagnéris et al., 2013), closed (ClosedAb; PBD accession no. 3RVY, an I217C mutant; 2.7-Å resolution; Payandeh et al., 2011), and inactivated (InactivatedAb; PDB accession no. 4EKW, 3.2-Å resolution; Payandeh et al., 2012) forms. The other inactivated form for which there is a crystal structure, NavRh (PDB accession no. 4DXW; 3.0-Å resolution; Zhang et al., 2012), and the other closed form, NavAe (PDB accession no. 4LTO; 3.5-Å resolution; Shaya et al., 2014), are more distant homologues (only 42 and 45% identity, respectively, in the pore region; Fig. 1). In addition, although both the NavAb and NavMs homologues have been shown to support sodium flux (Payandeh et al., 2011; D’Avanzo et al., 2013), neither NavRh nor NavAe has yet been shown to exhibit functional activity; therefore, we considered them to be less suitable for detailed comparisons. Although the resolutions of the structures are modest, all are sufficient to define the types and magnitudes of the conformational differences described in the comparisons made in this Review. In addition to these crystal structures, however, two structures of another orthologue, NavCt (PDB accession no. 4BGN, ∼9-Å resolution; Tsai et al., 2013), have been determined by electron crystallography, but those structures are at too low a resolution to make viable comparisons.

This Review focuses on comparisons of the ClosedAb–pOpenMs–OpenMs–InactivatedAb crystal structures, developing a structure-based model for the functional transitions. Although these crystal structures provide an important new structural context for understanding the nature of the transitions, it must be remembered that any models developed from them will ultimately require verification by functional studies under voltage-clamp conditions that probe native structures in real time and in the context of cell membranes.

Defining the conformational states represented in the crystal structures

The full-length NavAb channel has its VS in the “up” conformation (with residues E59 and D81 forming salt-bridged pairs to arginines in S4; Fig. 1) (Payandeh et al., 2011). This conformation of the VS is considered to be associated with the activated or fully open state as defined by disulfide cross-linking experiments (DeCaen et al., 2008, 2009). In addition, because in crystals there is no transmembrane potential, we expected that the structure would include an open form of the pore, as would be seen when the channel gate opens in response to membrane depolarization. However, in these crystals, the activation gate, and thus the pore region, appears to be in the closed conformation: the pore has no continuous transmembrane pathway that would enable the passage of sodium ions from the extracellular to the intracellular surface (McCusker et al., 2012; Fig. 2, A and B). Although the selectivity filter (SF) is sufficiently open to enable ions to enter the pore, the bottom of its cavity (at the activation gate) is blocked (Payandeh et al., 2011) so ions cannot exit. This structure has thus been described as being in a “pre-open” state, in other words, with a VS conformation that has primed the gate for opening but in which the gate is actually closed. This demonstrates that the structures of the VS and pore regions can be uncoupled, and may indicate the presence of an additional conformation intermediate between the “open” and “closed” states. In the comparisons in this Review, which focuses on the pore region responsible for the opening and closing of the transmembrane pathway, we have designated this structure as “closed” (ClosedAb) based on the state of the pore region. It is very similar in dimensions and features to the NavAe closed pore (Shaya et al., 2014), a structure that does not contain a VS region, another indication that the pore region state is not solely determined by the VS state.

One of the two NavMs pore structures (Bagnéris et al., 2013) has been designated to be in the fully open state (OpenMs) by virtue of its transmembrane pathway being of sufficient diameter along the full length of the pore, from the extracellular surface to the intracellular surface (Fig. 2, A and B), to enable the translocation of sodium ions across the cell membrane (Ulmschneider et al., 2013). It has all four monomers with a splayed conformation at their intracellular ends. In contrast, in the partially open (pOpenMs) structure (McCusker et al., 2012) (which lacks the last 37 residues of its C-terminal domain [CTD]), only one of the four monomers of the tetramer adopts an open splayed conformation, with the result that the activation gate is only partially open; it is not sufficient to permit ion translocation but is more widely open than the closed structure (Fig. 2 A). The other three monomers in this structure are effectively equivalent to the closed-state pore conformations. A model “open” structure was constructed (McCusker et al., 2012) based on using the most open of the four monomers to produce a symmetric tetramer, which was very similar to the structure later determined of the fully open state (Bagnéris et al., 2013). In the partially open structure, the distal end of the CTD, which has been proposed to form a stabilizing four-helix coiled-coil (Powl et al., 2010; Irie et al., 2012; Bagnéris et al., 2013; Shaya et al., 2014), is absent, although it is present in the fully open structure; this may have enabled the uncoupled opening of the four S6 helices, and this asymmetry suggests there may be an asynchrony in the functional transition between states.

The two inactivated forms of NavAb (InactivatedAb; Payandeh et al., 2012) and NavRh (InactivatedRh; Zhang et al., 2012) have been so designated based on their closed pores and collapsed SFs (although the authors of the NavAb structure were careful to describe it as being in a “potentially” inactivated state, largely because there was no confirming functional data). Both of these structures have closed activation gates, which are very similar to the ClosedAb structure, but their SFs are unlike any of the other structures, and would be too narrow to enable ions to enter the pore (Figs. 2 A, middle, and 4, C and D, and Table S1).

Structural transitions associated with opening and closing

The extracellular turret and vestibule surfaces (comprised of the S5–S6 linker, including the P1, P2, and SF regions; Fig. 1) are very similar in the OpenMs and ClosedAb pores, suggesting that these are not substantially involved in the gating transition. In addition, the N-terminal ends of their S6 helices (Fig. 3 B) are virtually superimposable, as are the C-terminal ends of the S5 helices. However, the intracellular surface features differ substantially (Fig. 3 A) as a result of differences in the relative orientations and splaying of the C-terminal ends of their S6 helices away from the pore axis. Also, the N-terminal ends of the S5 helices appear to have moved slightly with respect to each other. Had the open pore structure had its VS attached, this latter movement would have impinged on the S4–S5 linker (Fig. 3 A, circle) of an adjacent monomer. We speculate that the reason the NavAb pore is closed is that the linker region is in the closed conformation, and that if the linker had been more closely coupled to the VS in its open conformation in the crystal, the NavAb pore structure would also have been of an open conformation.

When the structures of the S6 pore helices of all the conformations are examined in detail (Fig. 3 B), the differences at the quaternary structural level between the open and closed (and inactivated) structures appear to arise from a twist in the middle of the S6 helix structures beginning at residue T206. The extracellular ends of all of the S6 helices in all of the closed and inactivated structures superimpose closely (Fig. 3 B), but the OpenMs structure is an outlier. This results from a change in the backbone Ramachandran angles between the open and closed forms, which perpetrates along the C-terminal end of S6 starting at T206 (Fig. 3 C). The consequence of this deviation is that the C-terminal end of the S6 helix in the OpenMs structure moves away from the axis of the top part of the helix, thereby opening the activation gate. In the lower resolution pOpenMs structure, residue T209 appeared to be the focal point of the bend, but in the higher resolution fully OpenMs structure, it is clear that the beginning of the bend occurs one turn earlier in S6, at T206. Notably, the changes between open and closed structures are subtle enough so that they do not change the secondary structure (Fig. 3 D) of the S6 helix. What starts as a relatively small change in the middle of the helix is translated to a motion of >5 Å at the end of S6 that forms the activation gate. This suggests a simple (energetically inexpensive) mechanism for opening and closing: a twist in the backbone that does not disrupt the hydrogen bonding pattern is sufficient to open the gate. It does not require major rearrangements of the rest of the structure and is thus compatible with a rapid opening, such as that which gives rise to the initial phase of the action potential in excitable cells. The asynchrony of the motions of the four monomers, as suggested by the partially open structure, suggests that there may also be an asynchrony of the process of opening.

The open and closed structures have very similar cavity regions (Fig. 2 A), with a minor hydrophobic constriction (designated HC1) near the bottom of the cavity (Fig. 2 A) near residue I215 in NavMs (equivalent to V213 in NavAb) that is not sufficient to prevent ions continuing their passage. However, the ClosedAb form has two additional major hydrophobic constrictions (HC2 and HC3) further down toward the extracellular surface (Fig. 2 A) at approximately the level of residues M221 and I217 (numbering according to NavAb sequence; notably, the latter corresponds to the site of the mutation to a cysteine in the NavAb construct). Neither of these constrictions is seen in the open NavMs structure. HC3 (which obscures the exit) would exclude passage of even totally dehydrated sodium ions in the closed structure.

Because the SFs of NavMs and NavAb have identical sequences (TLESWS; Fig. 1), they can be directly compared. The SFs of the OpenMs (Fig. 4 A) and ClosedAb (Fig. 4 B) structures are superimposable (Fig. 4 B, right), thus indicating that the closing of the activation gate, which prevents ion translocation, does not affect the potential entry of ions into the pore. In contrast, the SFs of the inactivated forms (Fig. 4, C and D) are very different from those of both the open and closed states, suggesting that they are altered during the inactivation process.

The structure and role of the CTD in opening and closing

All of the initial crystal structures determined (NavAb, NavMs, and NavRh) have elucidated the nature of the transmembrane domains (effectively to the end of the S6 helix) of the channels, but the structures of their CTDs (beyond the end of helix S6) were not interpretable, although these regions were present in the proteins used to produce the crystals. The structure of the CTD of the open conformation was first determined not by crystallography but by a combination of spectroscopic methods (Powl et al., 2010; Bagnéris et al., 2013), and shown to have a disordered region adjacent to the activation gate and a distal coiled-coil region (Figs. 1 and 5); molecular dynamics calculations (Bagnéris et al., 2013) suggested that flexibility of the region adjacent to the activation gate in the open state could account for the lack of defined structure in the crystal. The CTD in the closed NavAe pore (Shaya et al., 2014) is the only other such domain that has been defined structurally, this time by crystallographic means. Like the open CTD, its distal end formed a coiled-coil, but unlike the open CTD, the proximal end of the structure, nearest to the activation gate, was helical and relatively well ordered. These observations suggested that although the C-terminal end of the CTD remains unchanged in the two states, the end adjacent to the activation gate undergoes an ordered-to-disordered transition when the gate is opened (Fig. 5), thereby accommodating the separation movement at the end of the S6 helix that forms the gate, without uncoiling the distal coiled-coil that has been proposed to have a structural role in stabilizing the tetrameric structure in the membrane (Mio et al., 2010; Powl et al., 2010). Functional studies on the NavMs channel (Bagnéris et al., 2013), as well as on another orthologue (NavSulP from Sulfitobacer pontiacus; Irie et al., 2012), have indicated that this region (especially the EEE motif near the top of the CTD) plays a role in inactivation, recovery, and closing, perhaps by promotion of the conformational change of the S6 helix. In addition, in the NavAe pore structure, there is some density in the extracellular region beyond the end of the transmembrane segments that could be a calcium or other cation. Domain DIV of the human Nav1.4 sodium channel also possesses a similar negatively charged region just after its activation gate (Fig. 1), which appears to be a common feature in the prokaryotic sodium channels.

What produces the open and closed states in the crystals?

An obvious suggestion as to what causes NavMs pore structure to be open is that it is missing the constraints provided by the VS and S4–S5 linker regions present in the full-length channel structures. However, this cannot be the only reason, as the two structures of pore-only constructs, NavMs and NavAe, are in different states (NavMs is open, whereas NavAe is closed). It is also not because of the presence of CTD structures because, again, the CTDs are present in both of these two structures, albeit in different conformations. Furthermore, the partially open NavMs construct is missing the distal end of its CTD, yet one of its monomers is in the open state. The CTDs are present but not visible in any of the other structures. It is possible that the different CTD structures have a role in driving the transition (for example because of the different conformations at their N-terminal ends), but this cannot be determined from the structures in hand. It could also be because of the differences in crystal packing that enable the NavMs pores to open: in the NavMs crystals, the transmembrane regions of different pores are aligned next to each other, as if in a membrane bilayer, with a large enough gap to fit the CTD between two separate bilayers and thus allow sufficient room for the bottom of the pore to open. In the NavAe crystals, the CTDs are sandwiched between other CTDs and pore regions of symmetrically related molecules, rigidifying the whole structure and potentially preventing the opening.

The different structures in the crystals may be caused by (or alternatively, result in) the presence or absences of ions within the pore, as sodium ions are visible in the SF of only the NavMs open-state crystal forms. However, in a version of the NavAb orthologue engineered to be calcium selective (Tang et al., 2014), the presence of calcium ions in that pore did not produce an open state, so the mere presence of ions appears not to be sufficient to produce an open pore. The most likely candidate in vivo is the VS-pore linker between helices S4 and S5, which could act as a lever, either pushing or pulling the pore domain when the VS is activated to its “up” position. But the linker is not present in either the NavMs open or partially open structures, nor in the NavAe closed structures. And although it is present in the NavAb structures, it is in the “wrong” position to influence the opening of the pore, so it may be a contributing, but not the only, driving force. This is an important issue and will need to be resolved in the future to enable a full understanding of the opening and closing processes.

Structural transitions associated with inactivation

Comparisons of the structure of the InactivatedAb channel with both the ClosedAb and OpenMs structures indicate areas of the protein primarily involved in the inactivation transition. The InactivatedAb structure has a very similar upper cavity region to both the open and closed structures, and an asymmetric inactivation gate conformation that, like the symmetrical gate of the closed state, is occluded. The activation gate structures of the S6 helices of both the InactivatedAb and InactivatedRh channels closely overlay the ClosedAb (Fig. 3 B) but not the OpenMs structures, suggesting that the hinge necessary to open the gate has not been activated. Although their S6 conformations appear to be the same, the structures of the two types of InactivatedAb tetramers (“AB” and “CD”) and the ClosedAb form do exhibit differences at their C-terminal ends: the inactivated forms have between two and eight fewer ordered residues visible in the crystals, suggesting that they exhibit more mobility or flexibility in this region. Because the CTD has not been resolved in any of the inactivated form crystals, we cannot make any conclusions about whether the regions adjacent to the transmembrane helices are disordered (as in the open state) or helical (as in the closed state). Nor can we discern even if they exhibit the same coiled-coil structures at their distal ends that are present in both the open and closed forms.

In contrast, the SF region of the InactivatedAb structure (as well as that of the inactivated NavRh structure) is very different from either the open or closed states (Fig. 4). It has been described as “collapsed,” and indeed it is of insufficient diameter to enable a full dehydrated sodium ion to enter into the pore, much less exit it. Because in this case comparisons can be made for two crystal structures of the same orthologue (NavAb), the differences in the inactivated and closed structures cannot be attributed to SF sequence differences. The changes involve both the P1 and P2 helices as well as the SF itself, and involve modest differences in the secondary structures of the turret and top of S5 (Fig. 3 D). The observation that structural changes associated with the inactivated structure primarily involve the SF region corresponds well with earlier functional studies suggesting that the region close to the SF is responsible for inactivation (Pavlov et al., 2005). Interestingly, too, is the correspondence with studies done many years ago using circular dichroism spectroscopy (Cronin et al., 2003) on eukaryotic (electric eel) sodium channels that were induced to adopt open, closed, and inactivated states through the use of toxins and drugs. Those functional studies indicated that the secondary structures of the open and closed channels were surprisingly very similar but that the secondary structures of the inactivated form was considerably different. This also corresponds with the structural observations for the prokaryotic channels, where there are no apparent changes in secondary structure between open and closed states but significant differences between the open and the inactivated state.

The structural comparisons thus suggest that inactivation closes not only the extracellular but also the intracellular ends of the transmembrane passageway, although the extracellular changes appear to be the defining ones for the inactivation state.

Structural models for sodium channel opening–closing–inactivation mechanisms

Early models (e.g., Armstrong and Bezanilla, 1973; Hille, 1975; Lehmann-Horn and Jurkat-Rott, 1999; see also Hille, 2001, and Armstrong, 2007) proposed for eukaryotic sodium channel charge movement and gating were based primarily on functional observations and were produced, for the most part, before either any sodium channel sequences were determined or any crystal structures of any members of the voltage-gated cation channel superfamily were available. Structural models included suggestions such as the constriction of the SF or the shutting of two flaps (Lehmann-Horn and Jurkat-Rott, 1999), one at the extracellular surface and one at the intracellular surface, as being responsible for inactivation and closing, respectively. With the sequencing of the first eukaryotic sodium channel (Noda et al., 1984), three-dimensional models were developed that proposed the involvement of specific regions for the activation and inactivation processes (Guy and Seetharamulu, 1986; Yu et al., 2005). Many other approaches, including mutational effects, interactions with toxins, drug and ligand binding, and various spectroscopic studies, have since contributed to structure–function models of sodium channels (Catterall, 2012). More recently, the crystal structures of potassium channels (Doyle et al., 1998; Long et al., 2007) provided structural templates and information for understanding the electromechanical coupling and mechanism of their opening and closing. The sequence homology of sodium channels to these other members of the voltage-gated ion channel superfamily suggested that they would share a common architecture, albeit with very different SF regions, and led to more detailed models for sodium channel structures and functions (Yarov-Yarovoy et al., 2001; Shafrir et al., 2008; Zarrabi et al., 2010).

Soon after the prokaryotic sodium channels were identified as simpler target molecules for biophysical studies (Ren et al., 2001), and well before any of the prokaryotic orthologue crystal structures were determined, a hinge model for NaChBac (the first sodium channel orthologue whose sequence was determined) was proposed. This model for opening of the activation gate, like the models for the MthK potassium channel (Jiang et al., 2002), was based on the presence of a glycine residue in the middle of the S6 helix. Because glycines lack side chains, this type of residue could act as a flexible pivot for the top and bottom of the S6 helix, which could then move to open the gate. This model was supported by functional studies that showed that mutations of the glycine altered the rate of inactivation and recovery (Ito et al., 2004; Zhao et al., 2004), and spectroscopic studies that showed that changing the glycine to a conformationally less flexible serine produced a more thermally stable protein (O’Reilly et al., 2008); later molecular dynamics calculations (Barber et al., 2012) also supported a hinge-bending model involving this glycine. This model for the mechanism became less favored, however, after the determination of the sequences of other orthologues, as they did not have a glycine in the equivalent position.

The solution of the crystal structure of the prokaryotic sodium channel NavAb, with sequence and structural homology (especially in the VS regions) to single-domain tetrameric prokaryotic and eukaryotic potassium channel structures, prompted the suggestion (Payandeh et al., 2011) that the mechanism behind the coupling and pore opening might be similar to another model proposed for K+ channels (Long et al., 2007). Indeed, the VS of the ClosedAb structure aligns well with the VS of the open Kv1.2 structure, although they diverge in the pore, specifically at the beginning of the base of the S5 helix, and their SFs are completely different both in sequence and in structure (one using side chains and backbone carbonyls, and the other narrower one using only backbone carbonyls to create the ion-binding sites). That model had the VS and pore regions moving as modular units, with the S4–S5 linker movement causing the conformational change. In the sodium channel case, the proposed change was a rotation-like motion of the S5 and S6 helices relative to each other, opening the pore in an iris-like dilation motion. This bears semblance to the twisting mechanism proposed for the opening and closing of the nonspecific ion channel in nicotinic acetylcholine receptors (Unwin, 2003), where the transmembrane helices of adjacent subunits change positions relative to each other, primarily on one surface, producing a rotational squeezing and release mechanism that changes the size of the pore along its length.

A new structure-based model for the opening–closing–inactivation mechanisms

The observed structural differences described above between OpenMs and ClosedAb do not support a model based on a modular S5–S6 iris-like motion. In the prokaryotic sodium channel crystal structures, the C-terminal end of the S5 helix exhibits very little change between the two different states, and nothing suggests its movement is coupled with that of the S6 helix in either the same or an adjacent subunit within the tetramer. Instead, the opening and closing appears to involve a twist/bend in the middle of the S6 helix, which moves the activation gate to occlude or open the intracellular surface, enabling ion egress from the cavity (McCusker et al., 2012; Bagnéris et al., 2013; Video 1). This is remarkably similar to the earlier hinge model, except that there is no glycine at the hinge position. Instead, a threonine (T206 in the NavMs numbering scheme) is located at the juncture, in an equivalent position in the sequence to the glycine found in the NaChBac orthologue. The structure of NavMs shows that there would be sufficient space to enable the backbone motion even with the threonine side chain, as the Cβ substituent does not impinge on the region that moves. However, the enhanced flexibility of a glycine residue in NaChBac could be responsible for its faster rate of opening (Ren et al., 2001) than NavMs (Ulmschneider et al., 2013).

Because the S1–S4 helices of the VS structure are already in the open conformation in the ClosedAb and InactivatedAb structures, it initially seemed likely that the S4–S5 linker in these structures would be in a conformation similar to that which it would adopt in the open pore state. However, comparisons of the OpenMs and ClosedAb structures indicate that the positions of especially the S5 helices in the open pore would mitigate against the linker in the ClosedAb structure being completely in the activated state because the end of the open state S5 helix would impinge on the linker. It is therefore suggested that the linker is in a “primed” state but that it moves in concert with the S5 and S6 helices as the activation gate is opened.

The structure-based model for the slow inactivation process, like several of the early models (Lehmann-Horn and Jurkat-Rott, 1999), appears to primarily involve a closing of the SF to ion entry on the extracellular surface (Fig. 5). Rather than a flap folding over the entry, however, it appears that the SF itself folds in toward the center of the ion pathway, blocking entry of any sodium ions into the pore. It also appears, contrary to the double-flap mechanism, that the activation gate in the inactivated form is closed rather than open, although this could be a consequence of the crystallization and packing conditions (Video 2).

Flexibility at the C-terminal region of S6 thus appears to be the defining feature of the prokaryotic voltage-gated sodium channel. It may be at least partially responsible for channel opening and for the cascade of events that occur at the SF and P loops during inactivation. The presence of a flexible linking region to the CTD would enable the opening and closing to occur without uncoiling the distal coiled-coil region, which may play an important role in stabilizing the tetrameric quaternary structure of the prokaryotic sodium channels (Video 3). This feature is not present (nor necessary) in eukaryotic sodium channels, which are single polypeptide chains. Instead, they seem to have a CTD that plays an important role in fast inactivation, a function that prokaryotic sodium channels do not exhibit.

Similarities and differences in the activation mechanisms of prokaryotic and eukaryotic sodium channels

Prokaryotic and eukaryotic sodium channels exhibit considerable homologies in the sequences of their transmembrane regions (Fig. 1), especially in the C-terminal ends of their S6 helices, although their SFs and CTDs are entirely different. Likewise, both exhibit transitions between activation, recovery, and slow inactivation functional states, albeit with different kinetics (as well as the absence of fast inactivation in prokaryotic channels). Therefore, it is of interest to consider if they might involve similar structural features in a common mechanism for opening and closing.

New studies (Bagnéris et al., 2014) on the structure and electrophysiological consequences of binding channel blocker drugs to prokaryotic and human Nav1.1 channels have shown the parallel nature of their block by a range of ligands that bind in the hydrophobic cavity and prevent passage of ions from the extracellular to intracellular surfaces. This apparently occurs not by closing the activation gate but by the hydrophobic ligands creating a barrier to exit at the end of the SF region, and is entirely consistent with the mechanisms described above.

Furthermore, a recent study was done to test whether a similar mechanism of opening and closing could be operating in human Nav1.4 and prokaryotic sodium channels (Oelstrom et al., 2014). Channels with their fast inactivation mechanism disabled could be functionally held in either the open or closed state. An accessibility labeling technique was used to probe sites in S6 of domain IV (Fig. 1). Residues C terminal to the proposed activation gate were accessible to labeling from the intracellular side in both the open and closed states, whereas residues above this were only accessible in the open state. In the open state, the channel could be labeled up to the ring of hydrophobic residues immediately preceding the equivalent to residue T206 in the prokaryotic channels, again consistent with a mechanism involving a hinge mechanism in the middle of S6. These results strongly support the type of mechanism for opening and closing presented above, with a flexible hinge region that opens the pore up to the intracellular compartment in the open state, but not in the closed state.

However, it is important to note that significant differences in the activation/inactivation and selectivity mechanisms do exist between prokaryotic and eukaryotic channels (see, for example, Ahern, 2013; Goldschen-Ohm et al., 2013; Finol-Urdaneta et al., 2014, for detailed discussions). These arise in large part because of the asymmetry of the pseudotetrameric eukaryotic structures as opposed to the simpler homotetramers that comprise the prokaryotic channels. They result in asynchronous movements of VSs, asymmetric ion-binding sites, and novel loops between the different domains that have specialized roles, for example in fast inactivation, which are not seen in the prokaryotic channels. Nevertheless, the availability of several crystal forms of prokaryotic voltage-gated sodium channels in different conformational states has vastly increased our knowledge of the structure–function relationships for these channels, and provides new insight into their mechanisms of opening, closing, and inactivation (Fig. 5). These, in turn, may inform our understanding of both the structure and function of sodium channels in general. The new model proposed, based on the detailed structures of prokaryotic channels in crystal environments, will, however, await confirmation by further functional experiments on channels in their biological context, as well as structural studies of eukaryotic channels.

Acknowledgments

This work was supported by grants from the UK Biotechnology and Biological Sciences Research Council (to B.A. Wallace).

The authors declare no competing financial interests.

Elizabeth M. Adler served as editor.

References

Ahern
,
C.A.
2013
.
What activates inactivation?
J. Gen. Physiol.
142
:
97
100
.
Armstrong
,
C.M.
2007
.
Life among the axons
.
Annu. Rev. Physiol.
69
:
1
18
.
Armstrong
,
C.M.
, and
F.
Bezanilla
.
1973
.
Currents related to movement of the gating particles of the sodium channels
.
Nature.
242
:
459
461
.
Bagnéris
,
C.
,
P.G.
DeCaen
,
B.A.
Hall
,
C.E.
Naylor
,
D.E.
Clapham
,
C.W.M.
Kay
, and
B.A.
Wallace
.
2013
.
Role of the C-terminal domain in the structure and function of tetrameric sodium channels
.
Nat. Commun.
4
:
2465
.
Bagnéris
,
C.
,
P.G.
DeCaen
,
C.E.
Naylor
,
D.C.
Pryde
,
I.
Nobeli
,
D.E.
Clapham
, and
B.A.
Wallace
.
2014
.
Prokaryotic NavMs channel as a structural and functional model for eukaryotic sodium channel antagonism
.
Proc. Natl. Acad. Sci. USA.
111
:
8428
8433
.
Barber
,
A.F.
,
V.
Carnevale
,
S.G.
Raju
,
C.
Amaral
,
W.
Treptow
, and
M.L.
Klein
.
2012
.
Hinge-bending motions in the pore domain of a bacterial voltage-gated sodium channel
.
Biochim. Biophys. Acta.
1818
:
2120
2125
.
Catterall
,
W.A.
2012
.
Voltage-gated sodium channels at 60: structure, function and pathophysiology
.
J. Physiol.
590
:
2577
2589
.
Charalambous
,
K.
, and
B.A.
Wallace
.
2011
.
NaChBac: The long lost sodium channel ancestor
.
Biochemistry.
50
:
6742
6752
.
Chovancova
,
E.
,
A.
Pavelka
,
P.
Benes
,
O.
Strnad
,
J.
Brezovsky
,
B.
Kozlikova
,
A.
Gora
,
V.
Sustr
,
M.
Klvana
,
P.
Medek
, et al
.
2012
.
CAVER 3.0: A tool for the analysis of transport pathways in dynamic protein structures
.
PLOS Comput. Biol.
8
:
e1002708
.
Cronin
,
N.B.
,
A.
O’Reilly
,
H.
Duclohier
, and
B.A.
Wallace
.
2003
.
Binding of the anticonvulsant drug lamotrigine and the neurotoxin batrachotoxin to voltage-gated sodium channels induces conformational changes associated with block and steady-state activation
.
J. Biol. Chem.
278
:
10675
10682
.
D’Avanzo
,
N.
,
E.C.
McCusker
,
A.M.
Powl
,
A.J.
Miles
,
C.G.
Nichols
, and
B.A.
Wallace
.
2013
.
Differential lipid dependence of the function of bacterial sodium channels
.
PLoS ONE.
8
:
e61216
.
DeCaen
,
P.G.
,
V.
Yarov-Yarovoy
,
Y.
Zhao
,
T.
Scheuer
, and
W.A.
Catterall
.
2008
.
Disulfide locking a sodium channel voltage sensor reveals ion pair formation during activation
.
Proc. Natl. Acad. Sci. USA.
105
:
15142
15147
.
DeCaen
,
P.G.
,
V.
Yarov-Yarovoy
,
E.M.
Sharp
,
T.
Scheuer
, and
W.A.
Catterall
.
2009
.
Sequential formation of ion pairs during activation of a sodium channel voltage sensor
.
Proc. Natl. Acad. Sci. USA.
106
:
22498
22503
.
Doyle
,
D.A.
,
J.
Morais Cabral
,
R.A.
Pfeutzner
,
A.
Kuo
,
J.M.
Gulbis
,
S.L.
Cohen
,
B.T.
Chait
, and
R.
MacKinnon
.
1998
.
The structure of the potassium channel: Molecular basis of K+ conduction and selectivity
.
Science.
280
:
69
77
.
Finol-Urdaneta
,
R.K.
,
Y.
Wang
,
A.
Al-Sabi
,
C.
Zhao
,
S.Y.
Noskov
, and
R.J.
French
.
2014
.
Sodium channel selectivity and conduction: Prokaryotes have devised their own molecular strategy
.
J. Gen. Physiol.
143
:
157
171
.
Goldschen-Ohm
,
M.P.
,
D.L.
Capes
,
K.M.
Oelstrom
, and
B.
Chanda
.
2013
.
Multiple pore conformations driven by asynchronous movements of voltage sensors in a eukaryotic sodium channel
.
Nat. Commun.
4
:
1350
.
Guy
,
H.R.
, and
P.
Seetharamulu
.
1986
.
Molecular model of the action potential sodium channel
.
Proc. Natl. Acad. Sci. USA.
83
:
508
512
.
Hille
,
B.
1975
.
Ionic selectivity, saturation, and block in sodium channels. A four-barrier model
.
J. Gen. Physiol.
66
:
535
560
.
Hille
,
B.
2001
.
Ion Channels of Excitable Membranes
.
Sinauer Associates
,
Sunderland, MA
.
814 pp
.
Irie
,
K.
,
T.
Shimomura
, and
Y.
Fujiyoshi
.
2012
.
The C-terminal helical bundle of the tetrameric prokaryotic sodium channel accelerates the inactivation rate
.
Nat. Commun.
3
:
793
.
Ito
,
M.
,
H.
Xu
,
A.A.
Guffanti
,
Y.
Wei
,
L.
Zvi
,
D.E.
Clapham
, and
T.A.
Krulwich
.
2004
.
The voltage-gated Na+ channel NaVBP has a role in motility, chemotaxis, and pH homeostasis of an alkaliphilic Bacillus
.
Proc. Natl. Acad. Sci. USA.
101
:
10566
10571
.
Jiang
,
Y.
,
A.
Lee
,
J.
Chen
,
M.
Cadene
,
B.T.
Chait
, and
R.
MacKinnon
.
2002
.
The open pore conformation of potassium channels
.
Nature.
417
:
523
526
.
Klose
,
D.P.
,
B.A.
Wallace
, and
R.W.
Janes
.
2010
.
2Struc: the secondary structure server
.
Bioinformatics.
26
:
2624
2625
.
Koishi
,
R.
,
H.
Xu
,
D.
Ren
,
B.
Navarro
,
B.W.
Spiller
,
Q.
Shi
, and
D.E.
Clapham
.
2004
.
A superfamily of voltage-gated sodium channels in bacteria
.
J. Biol. Chem.
279
:
9532
9538
.
Kuzmenkin
,
A.
,
F.
Bezanilla
, and
A.M.
Correa
.
2004
.
Gating of the bacterial sodium channel, NaChBac: Voltage-dependent charge movement and gating currents
.
J. Gen. Physiol.
124
:
349
356
.
Lehmann-Horn
,
F.
, and
K.
Jurkat-Rott
.
1999
.
Voltage-gated ion channels and hereditary disease
.
Physiol. Rev.
79
:
1317
1372
.
Long
,
S.B.
,
X.
Tao
,
E.B.
Campbell
, and
R.
MacKinnon
.
2007
.
Atomic structure of a voltage-dependent K+ channel in a lipid membrane-like environment
.
Nature.
450
:
376
382
.
McCusker
,
E.C.
,
N.
D’Avanzo
,
C.G.
Nichols
, and
B.A.
Wallace
.
2011
.
Simplified bacterial “pore” channel provides insight into the assembly, stability, and structure of sodium channels
.
J. Biol. Chem.
286
:
16386
16391
.
McCusker
,
E.C.
,
C.
Bagnéris
,
C.E.
Naylor
,
A.R.
Cole
,
N.
D’Avanzo
,
C.G.
Nichols
, and
B.A.
Wallace
.
2012
.
Structure of a bacterial voltage-gated sodium channel pore reveals mechanisms of opening and closing
.
Nat. Commun.
3
:
1102
.
Mio
,
K.
,
M.
Mio
,
F.
Arisaka
,
M.
Sato
, and
C.
Sato
.
2010
.
The C-terminal coiled-coil of the bacterial voltage-gated sodium channel NaChBac is not essential for tetramer formation, but stabilizes subunit-to-subunit interactions
.
Prog. Biophys. Mol. Biol.
103
:
111
121
.
Noda
,
M.
,
S.
Shimizu
,
T.
Tanabe
,
T.
Takai
,
T.
Kayano
,
T.
Ikeda
,
H.
Takahashi
,
H.
Nakayama
,
Y.
Kanaoka
,
N.
Minamino
, et al
.
1984
.
Primary structure of Electrophorus electricus sodium channel deduced from cDNA sequence
.
Nature.
312
:
121
127
.
Nurani
,
G.
,
M.
Radford
,
K.
Charalambous
,
A.O.
O’Reilly
,
N.B.
Cronin
,
S.
Haque
, and
B.A.
Wallace
.
2008
.
Tetrameric bacterial sodium channels: Characterization of structure, stability, and drug binding
.
Biochemistry.
47
:
8114
8121
.
O’Reilly
,
A.O.
,
K.
Charalambous
,
G.
Nurani
,
A.M.
Powl
, and
B.A.
Wallace
.
2008
.
G219S mutagenesis as a means of stabilizing conformational flexibility in the bacterial sodium channel NaChBac
.
Mol. Membr. Biol.
25
:
670
676
.
Oelstrom
,
K.
,
M.P.
Goldschen-Ohm
,
M.
Holmgren
, and
B.
Chanda
.
2014
.
Evolutionarily conserved intracellular gate of voltage-dependent sodium channels
.
Nat. Commun.
5
:
3420
.
Pavlov
,
E.
,
C.
Bladen
,
R.
Winkfein
,
C.
Diao
,
P.
Dhaliwal
, and
R.J.
French
.
2005
.
The pore, not cytoplasmic domains, underlies inactivation in a prokaryotic sodium channel
.
Biophys. J.
89
:
232
242
.
Payandeh
,
J.
,
T.
Scheuer
,
N.
Zheng
, and
W.A.
Catterall
.
2011
.
The crystal structure of a voltage-gated sodium channel
.
Nature.
475
:
353
358
.
Payandeh
,
J.
,
T.M.
Gamal El-Din
,
T.
Scheuer
,
N.
Zheng
, and
W.A.
Catterall
.
2012
.
Crystal structure of a voltage-gated sodium channel in two potentially inactivated states
.
Nature.
486
:
135
139
.
Powl
,
A.M.
,
A.O.
O’Reilly
,
A.J.
Miles
, and
B.A.
Wallace
.
2010
.
Synchrotron radiation circular dichroism spectroscopy-defined structure of the C-terminal domain of NaChBac and its role in channel assembly
.
Proc. Natl. Acad. Sci. USA.
107
:
14064
14069
.
Ren
,
D.
,
B.
Navarro
,
H.
Xu
,
L.
Yue
,
Q.
Shi
, and
D.E.
Clapham
.
2001
.
A prokaryotic voltage-gated sodium channel
.
Science.
294
:
2372
2375
.
Shafrir
,
Y.
,
S.R.
Durell
, and
H.R.
Guy
.
2008
.
Models of voltage-dependent conformational changes in NaChBac channels
.
Biophys. J.
95
:
3663
3676
.
Shaya
,
D.
,
M.
Kreir
,
R.A.
Robbins
,
S.
Wong
,
J.
Hammon
,
A.
Brüggemann
, and
D.L.
Minor
Jr
.
2011
.
Voltage-gated sodium channel (NaV) protein dissection creates a set of functional pore-only proteins
.
Proc. Natl. Acad. Sci. USA.
108
:
12313
12318
.
Shaya
,
D.
,
F.
Findeisen
,
F.
Abderemane-Ali
,
C.
Arrigoni
,
S.
Wong
,
S.R.
Nurva
,
G.
Loussouarn
, and
D.L.
Minor
Jr
.
2014
.
Structure of a prokaryotic sodium channel pore reveals essential gating elements and an outer ion binding site common to eukaryotic channels
.
J. Mol. Biol.
426
:
467
483
.
Sievers
,
F.
,
A.
Wilm
,
D.
Dineen
,
T.J.
Gibson
,
K.
Karplus
,
W.
Li
,
R.
Lopez
,
H.
McWilliam
,
M.
Remmert
,
J.
Söding
, et al
.
2011
.
Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega
.
Mol. Syst. Biol.
7
:
539
.
Tang
,
L.
,
T.M.
Gamal El-Din
,
J.
Payandeh
,
G.Q.
Martinez
,
T.M.
Heard
,
T.
Scheuer
,
N.
Zheng
, and
W.A.
Catterall
.
2014
.
Structural basis for Ca2+ selectivity of a voltage-gated calcium channel
.
Nature.
505
:
56
61
.
Tsai
,
C.-J.
,
K.
Tani
,
K.
Irie
,
Y.
Hiroaki
,
T.
Shimomura
,
D.G.
McMillan
,
G.M.
Cook
,
G.F.X.
Schertler
,
Y.
Fujiyoshi
, and
X.-D.
Li
.
2013
.
Two alternative conformations of a voltage-gated sodium channel
.
J. Mol. Biol.
425
:
4074
4088
.
Ulmschneider
,
M.B.
,
C.
Bagnéris
,
E.C.
McCusker
,
P.G.
DeCaen
,
M.
Delling
,
D.E.
Clapham
,
J.P.
Ulmschneider
, and
B.A.
Wallace
.
2013
.
Molecular dynamics of ion transport through the open conformation of a bacterial voltage-gated sodium channel
.
Proc. Natl. Acad. Sci. USA.
110
:
6364
6369
.
Unwin
,
N.
2003
.
Structure and action of the nicotinic acetylcholine receptor explored by electron microscopy
.
FEBS Lett.
555
:
91
95
.
Yarov-Yarovoy
,
V.
,
J.
Brown
,
E.M.
Sharp
,
J.J.
Clare
,
T.
Scheuer
, and
W.A.
Catterall
.
2001
.
Molecular determinants of voltage-dependent gating and binding of pore-blocking drugs in transmembrane segment IIIS6 of the Na+ channel alpha subunit
.
J. Biol. Chem.
276
:
20
27
.
Yu
,
F.H.
,
V.
Yarov-Yarovoy
,
G.A.
Gutman
, and
W.A.
Catterall
.
2005
.
Overview of molecular relationships in the voltage-gated ion channel superfamily
.
Pharmacol. Rev.
57
:
387
395
.
Zarrabi
,
T.
,
R.
Cervenka
,
W.
Sandtner
,
P.
Lukacs
,
X.
Koenig
,
K.
Hilber
,
M.
Mille
,
G.M.
Lipkind
,
H.A.
Fozzard
, and
H.
Todt
.
2010
.
A molecular switch between the outer and the inner vestibules of the voltage-gated Na+ channel
.
J. Biol. Chem.
285
:
39458
39470
.
Zhang
,
X.
,
W.
Ren
,
P.
DeCaen
,
C.
Yan
,
X.
Tao
,
L.
Tang
,
J.
Wang
,
K.
Hasegawa
,
T.
Kumasaka
,
J.
He
, et al
.
2012
.
Crystal structure of an orthologue of the NaChBac voltage-gated sodium channel
.
Nature.
486
:
130
134
.
Zhao
,
Y.
,
V.
Yarov-Yarovoy
,
T.
Scheuer
, and
W.A.
Catterall
.
2004
.
A gating hinge in Na+ channels: A molecular switch for electrical signaling
.
Neuron.
41
:
859
865
.

    Abbreviations used in this paper:
     
  • CTD

    C-terminal domain

  •  
  • NavAb

    voltage-gated sodium channel from Arcobacter butzleri

  •  
  • NavAe

    voltage-gated sodium channel from Alkalilimnicola ehrlichei

  •  
  • NavMs

    voltage-gated sodium channel from Magnetococcus marinus

  •  
  • NavRh

    voltage-gated sodium channel from Rickettsiales sp. HIMB114

  •  
  • SF

    selectivity filter

  •  
  • VS

    voltage sensor

Author notes

E.C. McCusker’s present address is Teva Pharmaceuticals, Chicago, IL 60622

This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.rupress.org/terms). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).

Supplementary data