Voltage-gated proton channels, HV1, were first reported in Helix aspersa snail neurons. These H+ channels open very rapidly, two to three orders of magnitude faster than mammalian HV1. Here we identify an HV1 gene in the snail Helisoma trivolvis and verify protein level expression by Western blotting of H. trivolvis brain lysate. Expressed in mammalian cells, HtHV1 currents in most respects resemble those described in other snails, including rapid activation, 476 times faster than hHV1 (human) at pHo 7, between 50 and 90 mV. In contrast to most HV1, activation of HtHV1 is exponential, suggesting first-order kinetics. However, the large gating charge of ∼5.5 e0 suggests that HtHV1 functions as a dimer, evidently with highly cooperative gating. HtHV1 opening is exquisitely sensitive to pHo, whereas closing is nearly independent of pHo. Zn2+ and Cd2+ inhibit HtHV1 currents in the micromolar range, slowing activation, shifting the proton conductance–voltage (gH-V) relationship to more positive potentials, and lowering the maximum conductance. This is consistent with HtHV1 possessing three of the four amino acids that coordinate Zn2+ in mammalian HV1. All known HV1 exhibit ΔpH-dependent gating that results in a 40-mV shift of the gH-V relationship for a unit change in either pHo or pHi. This property is crucial for all the functions of HV1 in many species and numerous human cells. The HtHV1 channel exhibits normal or supernormal pHo dependence, but weak pHi dependence. Under favorable conditions, this might result in the HtHV1 channel conducting inward currents and perhaps mediating a proton action potential. The anomalous ΔpH-dependent gating of HtHV1 channels suggests a structural basis for this important property, which is further explored in this issue (Cherny et al. 2018. J. Gen. Physiol. https://doi.org/10.1085/jgp.201711968).
Voltage-gated proton channels, HV1, remain relative newcomers to the ion channel family. Although the idea of a depolarization-activated proton-selective ion channel was proposed in 1972 by J. Woodland Hastings and colleagues (Fogel and Hastings, 1972), the first voltage-clamp study that established the existence of this channel type occurred a decade later in the snail Helix aspersa (Thomas and Meech, 1982). An HV1 gene was not identified until 2006 (Ramsey et al., 2006; Sasaki et al., 2006). Strong interest in this channel has arisen for two main reasons. First, its structure, with just four transmembrane helices, closely resembles the voltage-sensing domain of other voltage-gated ion channels, making it a unique model for voltage-gating mechanisms. By combining voltage sensing, gating, and conduction into a single module, HV1 uniquely provides a direct readout of its gating state. Second, exceedingly diverse functions have been identified for HV1 in many species and in many human tissues (DeCoursey, 2013).
The first systematic voltage-clamp characterization of voltage-gated proton currents was in Lymnaea stagnalis snail neurons (Byerly et al., 1984). When mammalian proton currents were identified a decade later (DeCoursey, 1991), the most obvious difference was that HV1 in snails activated two to three orders of magnitude faster. Here, we investigate the properties of the Helisoma trivolvis snail HV1 gene product. We searched a transcriptome of H. trivolvis and found a putative HtHV1; we then cloned the gene from a cDNA pool constructed from H. trivolvis brain tissue. We find many similarities to native proton currents studied in situ in other snail species, including rapid gating kinetics and other significant differences from mammalian HV1. HtHV1 currents differ from mammalian HV1 in having exponential (vs. sigmoid) activation, similarity of τact and τtail at overlapping voltages, and maximal time constants near the midpoint of the proton conductance–voltage (gH-V) relationship, all features suggestive of simple first-order gating kinetics expected of a monomeric protein. However, the existence of an extensive coiled-coil motif in the C terminus together with steep voltage dependence suggests “cooperative” gating of a dimeric protein. Potent inhibition of HtHV1 by Zn2+ and Cd2+ is explained by conservation of three of four members of the Zn2+-binding site (Takeshita et al., 2014). The most remarkable property of the HtHV1 channel is that its sensitivity to pHi is anomalously weak. The voltage-gating mechanism of all HV1 identified to date is unique in being nearly equally responsive to pHo and pHi, such that a one-unit change in either shifts the gH-V relationship by 40 mV. This “rule of forty” (DeCoursey, 2013) has the biologically crucial effect of ensuring that HV1 channels open only when there is an outward electrochemical gradient for H+. In other words, HV1 channels open only when doing so will result in acid extrusion from cells. Extensive mutation of hHV1 has failed to produce any significant violation of the rule of forty (Ramsey et al., 2010; DeCoursey, 2016). In this issue, Cherny et al. identify a single amino acid difference between HtHV1 and hHV1 that appears to be responsible for the anomalous ΔpH dependence of the snail channel.
Materials and methods
H. trivolvis, a pulmonate snail (order: Basommatophora; family: Planorbidae) from an albino stock maintained and continuously bred in aquaria at Georgia State University, was used for experiments. Snails were originally caught in the wild and introduced as an experimental model animal by S.B. Kater (Kater, 1974).
Gene cloning, mutagenesis, antibody synthesis, and Western blotting
Basic Local Alignment Search Tool searches of a transcriptome from H. trivolvis (unpublished data) yielded a hit that matched the criteria for an HV1 sequence (Smith et al., 2011). Brains were dissected from H. trivolvis (Cohan et al., 2003), RNA was extracted from brain tissue using the RNeasy kit (Qiagen), and a cDNA pool was constructed using the SuperScript III kit (Life Technologies) according to the manufacturer's instructions. Primers designed against the transcriptome hit were used to clone the putative HtHV1 coding sequence; the sequence was confirmed by commercial sequencing (SourceBio Science). This coding sequence was subcloned into eukaryotic expression vector pCA-IRES-eGFP. Site-directed mutagenesis of HtHV1 was performed and sequence verified commercially (Genewiz). Antibody was raised in rabbit to a synthetic peptide (RSPSDHGEGFEEPLC) based on the predicted HtHV1 epitope and affinity purified (Genscript) with a final concentration of 0.904 mg/ml. Total lysate was prepared from H. trivolvis brains that had been stored whole in Qiagen RLT buffer at −80°C for 12 mo. Brains were thawed and triturated briefly on ice; the lysate was cleared by centrifugation at 10,000 × g for 5 min. Proteins from H. trivolvis brain lysate were separated by SDS-PAGE, Western blotted, and probed with anti-HtHV1 antibody (diluted 1:10,000 in blocking buffer) either alone or preincubated with 1,000-fold molar excess of synthetic peptide corresponding to the epitope.
HEK-293 cells were grown to ∼80% confluence in 35-mm culture dishes. HEK-293 cells were transfected with 0.4–0.5 µg cDNA using Lipofectamine 2000 (Invitrogen) or polyethylenimine (Sigma). Plasmids that did not include GFP were cotransfected with GFP. After 24 h at 37°C in 5% CO2, cells were trypsinized and replated onto glass coverslips at low density for patch-clamp recording. We selected green cells under fluorescence for recording. Because HEK-293 cells often have small endogenous HV1 currents (Musset et al., 2011), cells that exhibited small currents suspected to be native were exposed to 1 µM Zn2+, which has generally weaker effects on HtHV1 (20% slowing of τact, ∼5 mV shift of the gH-V relationship, and a 24% decrease in gH,max in three to four cells at pHo 7) than on hHV1 (more than a twofold slowing of τact, ∼20 mV shift of the gH-V relationship; Musset et al., 2010b). Cells determined on this basis to exhibit native currents were excluded from the study.
Micropipettes were pulled using a Flaming Brown automatic pipette puller (Sutter Instruments) from Custom 8520 Patch Glass (equivalent to Corning 7052 glass; Harvard Apparatus), coated with Sylgard 184 (Dow Corning Corp.), and heat polished to a tip resistance range of typically 3–10 MΩ with highly buffered TMA+ pipette solutions. Electrical contact with the pipette solution was achieved by a thin sintered Ag-AgCl pellet (In Vivo Metric Systems) attached to a Teflon-encased silver wire, or simply a chlorided silver wire. A reference electrode made from a Ag-AgCl pellet was connected to the bath through an agar bridge made with Ringer’s solution. The current signal from the patch clamp (EPC-9 from HEKA Instruments or Axopatch 200B from Axon Instruments) was recorded and analyzed using Pulse and PulseFit software (HEKA Instruments), or P-CLAMP software supplemented by Sigmaplot (SPSS). Seals were formed with Ringer’s solution (in mM: 160 NaCl, 4.5 KCl, 2 CaCl2, 1 MgCl2, 5 HEPES, pH 7.4) in the bath, and the potential was zeroed after the pipette was in contact with the cell. Current records are displayed without correction for liquid junction potentials.
Whole-cell or excised inside-out patch configurations of the patch-clamp technique were performed. Bath and pipette solutions were used interchangeably. They contained (in mM) 2 MgCl2, 1 EGTA, 80–100 buffer, and 75–120 TMA+ CH3SO3–, adjusted to bring the osmolality to ∼300 mOsm, and were titrated using TMAOH. Buffers with pKa near the desired pH were used: homo-PIPES for pH 4.5–5.0, Mes for pH 5.5–6.0, BisTris for pH 6.5, N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid for pH 7.0, HEPES for pH 7.5, Tricine for pH 8.0, and N-cyclohexyl-2-aminoethanesulfonic acid for pH 9.0. Experiments were done at room temperature (∼20–25°C). Current records are shown without leak correction.
Reversal potentials (Vrev) in most cases were determined from the direction and amplitude of tail current relaxation over a range of voltages after a prepulse that activated the proton conductance, gH. When the gH was activated negative to Vrev, the latter could be determined directly from families of currents. Currents were fitted with a single exponential to obtain the activation time constant (τact), and the fitted curve was extrapolated to infinite time to obtain the steady-state current amplitude (IH), from which the gH was calculated as gH = IH/(V − Vrev). Thus we assume that the time-dependent component reflects H+ current, and time-independent current represents leak. Because of the strong voltage dependence of activation kinetics, we frequently applied longer pulses near threshold voltages and shorter pulses for large depolarizations to resolve kinetics and avoid proton depletion associated with large H+ flux. The voltage at which gH was 10% of gH,max (VgH,max/10) was determined after defining gH,max as the largest gH measured.
HtHV1 is a voltage-gated proton-selective channel
The gene coding for a putative voltage-gated proton channel was identified based on criteria established previously, namely the presence of four transmembrane helices homologous to S1–S4 of voltage sensor domains with an Asp in the middle of the S1 transmembrane helix and the RxWRxxR motif in S4 (Musset et al., 2011; Smith et al., 2011). We cloned the putative HtHV1 gene from a cDNA pool of brain tissue, verifying that this gene is expressed at the RNA level. Protein level expression was verified by Western blotting of H. trivolvis brain lysate probed with a commercially raised antibody to a synthetic peptide based on a HtHV1 epitope (Fig. 1, inset). The single protein detected ran at ∼50 kD, somewhat larger than the predicted size of 40 kD. Glycosylation at the five putative N-glycosylation sites in the S1–S2 linker could account for this discrepancy, given that N-linked oligosaccharides range from 1,884 to 2,851 D (Imperiali and O’Connor, 1999). Excess synthetic peptide abolished the binding of antibody to brain lysate, establishing the specificity of the antibody. The antibody did not significantly bind to two proteins that do not contain the epitope: human glutathione S-transferase and luciferin binding protein from Lingulodinium polyedrum.
The HtHV1 channel protein (Fig. 1) is substantially larger than the human hHV1, with 360 amino acids (hHV1 has 273). Much of this excess resides in the S1–S2 extracellular linker with 73 residues (vs. eight in hHV1), which contains five potential N-glycosylation sites (vs. 0 in hHV1). Focusing on the transmembrane regions, HtHV1 has charged amino acids nearly identical to those of hHV1. One exception is at the outer end of the S1 helix, where hHV1 has basic Lys125 but snail HtHV1 has acidic Glu120. There is extensive predicted coiled-coil in the C terminus: 36 residues (positions 289–324) with 90% stringency, and 28 residues (294–321) with 99% stringency according to MARCOIL (Delorenzi and Speed, 2002). HV1 in several species have been shown to exist as dimers, largely because of coiled-coil interactions in the C terminus (Koch et al., 2008; Lee et al., 2008; Tombola et al., 2008).
The HtHV1 gene was transfected into HEK-293 cells. Under voltage clamp, transfected cells displayed depolarization-activated currents. The selectivity of these currents was established by measuring the reversal potential, Vrev, over a range of pHo and pHi values (Fig. 2). The measured values of Vrev are close to the Nernst potential for H+, EH, shown as a dashed line. Clearly, the HtHV1 channel is highly proton selective over the pH range studied.
HtHV1 gating is rapid with unusual voltage dependence
A family of currents generated by HtHV1 at symmetrical pH 6.0 is illustrated in Fig. 3 A. The currents activate rapidly with depolarization, and activation becomes much faster at higher voltages. Although HV1 currents in all species activate more rapidly at more positive voltages, the τact of HtHV1 currents (solid and open red squares in Fig. 3 C) exhibits noticeably steeper voltage dependence. The maximum slope of the τact-V relationship in seven cells at pHo 6 was 13.0 ± 3.4 mV/e-fold change in τact (mean ± SD). In several mammalian HV1, τact changes e-fold in 40–72 mV (DeCoursey, 2003). Channel closing in HtHV1 was also steeply voltage dependent (Fig. 3 B and blue diamonds in Fig. 3 C), with τtail changing e-fold in 14.2 ± 1.9 mV in six cells. In mammalian cells, the slope is typically much flatter, 26–44 mV/e-fold change in τtail (DeCoursey, 2003).
A remarkable feature of HtHV1 is that at intermediate voltages where the measurements overlap, the time constants of H+ current turn-on (τact) and deactivation (τtail) essentially superimpose (Fig. 3 C). This behavior is suggestive of simple first-order kinetics, such as a two-state system:
in which α is the rate of channel opening and β is the rate of channel closing, and the time constant τ is (α + β)−1 (Hodgkin and Huxley, 1952). Another feature suggestive of first-order kinetics is evident in the gH-V relationship from this cell (Fig. 3 D). The voltage at which the gH is half-maximal is ∼40 mV, where the time constants are maximal (Fig. 3 C). However, the limiting slope of the gH-V relationship in Fig. 3 D, i.e., the slope of the most negative values obtained, indicates a gating charge of ∼6 e0. The mean gating charge in 18 limiting slope measurements was 5.5 ± 0.9 e0 (mean ± SD). Because the range of gH values resolved did not exceed three orders of magnitude, these gating charge estimates should be considered lower limits. In most species, cooperative gating of the dimeric HV1 channel doubles the gating charge from 2–3 to 4–6 e0 (Gonzalez et al., 2010, 2013; Fujiwara et al., 2012).
Mean gating kinetics determined at symmetrical pH 7.0 is shown in Fig. 4 A. As was also seen at pH 6.0 (Fig. 3 C), at voltages where τact and τtail overlap, they have similar values, suggestive of first-order gating kinetics. In the first description of proton currents in snail neurons, the activation time to half-peak current was 25 ms or less at pHo 7.4 (Byerly et al., 1984). With this in mind, the activation kinetics of HtHV1 is quite similar to that reported in neurons from L. stagnalis. When proton currents were first identified in mammalian species, they were found to be radically slower (DeCoursey, 1991; Bernheim et al., 1993; DeCoursey and Cherny, 1993; Demaurex et al., 1993; Kapus et al., 1993). The activation kinetics of HtHV1 is two to three orders of magnitude faster than that of hHV1 (Fig. 4 B), averaging 476 times faster between 50 and 90 mV at pHo 7.
HtHV1 is sensitive to inhibition by external Zn2+ and Cd2+
The polyvalent metal cations Zn2+ and Cd2+ were among the first HV1 inhibitors identified (Thomas and Meech, 1982; Mahaut-Smith, 1989b). Zn2+ in particular has been used widely on HV1 identified in new species and remains the most potent inhibitor (Cherny and DeCoursey, 1999). Fig. 5 illustrates the effects of 100 µM Zn2+ or Cd2+ on HtHV1 currents. Three main effects are evident: the current amplitude is reduced, the current activates more slowly (scaled currents in Fig. 5 D), and the gH-V relationship is shifted positively along the voltage axis. These three parameters are interrelated in that a positive shift of the gH-V relationship will in itself decrease the current and slow τact at any given voltage. The mean changes in these three parameters produced by 10 or 100 µM of the two metals are summarized in Fig. 5 E.
These three effects of polyvalent metal cations have been observed for HV1 from many species. As in rat HV1 (Cherny and DeCoursey, 1999), Zn2+ is more potent than Cd2+ in HtHV1. Focusing on the three main effects, HtHV1 was more sensitive, similar to, or less sensitive than human HV1 (hHV1). The reduction of gH,max is glaringly obvious for HtHV1, whereas in mammalian HV1 this effect is small and difficult to detect because of the interrelatedness of the three effects (Cherny and DeCoursey, 1999). Zn2+ at 10 µM slows τact by four- to fivefold in both hHV1 (Musset et al., 2010b) and HtHV1 (Fig. 5). In contrast, the shift of the gH-V relationship by Zn2+ is far more profound in human hHV1, with a 20-mV shift produced by 1 µM Zn2+ (Musset et al., 2010b) compared with a 12-mV shift by 10 µM Zn2+ in HtHV1 (Fig. 5).
Unique ΔpH dependence of HtHV1 gating
Families of proton currents generated by the H. trivolvis proton channel gene product, HtHV1, in a cell studied at four pHo values with pHi 6 are illustrated in Fig. 6 (A–D). The currents activate with depolarization, and activation becomes much faster at higher voltages. Both voltage dependence and kinetics were exquisitely sensitive to pHo. At higher pHo, the proton conductance, gH, turned on at more negative voltages and turned on much more rapidly (note the different time bases). Fig. 6 (E–H) shows deactivation kinetics at each pHo during tail current measurements in this cell. Channel closing becomes much more rapid at more negative voltages.
In Fig. 6 I, time constants of H+ current turn-on (activation, τact) and turn-off (deactivation, τtail) from the same cell are plotted. Several intriguing features emerge. Unlike HV1 in other species, at intermediate voltages where τact and τtail overlapped, they were of similar magnitude (as was seen in Figs. 3 and 4). Because this behavior suggests first-order kinetics (Scheme 1), the data in Fig. 6 I were analyzed in this way, and the expressions for each rate constant are given on the graph in the following form:
The voltage dependence of τact is steep (small kα) and appears to become steeper at lower pHo. The voltage dependence of channel closing, τtail, is also steep (small kβ) but appears to be independent of pHo. To a first approximation, β is independent of pHo, whereas α is markedly influenced by pHo. Mean values for the rate constants are plotted in Fig. 7. Confirming the impression from Fig. 6 I (where kα was 11.4, 18, 26, and 30 mV at pHo 5, 6, 7, and 8), kα increased with pHo, and kβ was pH independent. But by far the strongest effect of pH is that increasing pHo massively increases the opening rate constant α, which increased more than an order of magnitude per unit increase in pHo. Stated differently, protonation at the external face of the HtHV1 channel strongly inhibits channel opening. More subtly, it is evident that for any given pHo, α is higher and β is lower at pHi 6 than at pHi 7; hence, lower pHi both promotes opening and slows closing.
The gH-V relationships from the cell in Fig. 6 (A–I) are plotted in Fig. 6 J. Like other HV1, HtHV1 exhibits robust pHo-dependent shifts with increasing pHo shifting the gH-V relationship negatively. The shifts for pHo 5 → 6 and 6 → 7 are closer to 50 than 40 mV, indicating that HtHV1 exceeds the rule of forty for changes in pHo. To reconstruct gH-V relationships using the simple first-order assumption (Scheme 1 and Eqs. 1 and 2), which predicts that Popen = α/(α + β), the solid curves in Fig. 6 J were drawn from the rate constant equations in Fig. 6 I scaled by gH,max. Their limiting slope at negative voltages is shallower than observed. Squaring the Popen-V relationships, as in the classic Hodgkin–Huxley n2 approach (Hodgkin and Huxley, 1952), produces the steeper dashed curves, which better approximate the data. Without pushing the model too far, we conclude that it is probable that, like several other HV1 (Gonzalez et al., 2010; Musset et al., 2010b; Tombola et al., 2010; Fujiwara et al., 2012), HtHV1 functions as a dimer in which both protomers must activate before either one conducts.
The effects of changes in pHi were explored in inside-out patches, as illustrated in Fig. 8. The resolution was limited somewhat by the typically small current amplitude combined with rapid activation kinetics at some pH. Activation kinetics could be resolved at low but not at high pHi. For example, at pHi 8 (Fig. 8 C), inward current is clearly activated, but the kinetics is ambiguous. Nevertheless, it is evident in Fig. 8 D that activation kinetics depends only weakly on pHi, in stark contrast to the strong dependence seen for pHo (Figs. 6 and 7), and in contrast to mammalian HV1, in which lowering pHi speeds activation fivefold/unit (DeCoursey and Cherny, 1995; Villalba-Galea, 2014). Deactivation kinetics was poorly resolved in most patches. The most surprising feature (Fig. 8 E) is that the heretofore universal rule of forty governing ΔpH-dependent gating is violated by HtHV1. Changing pHi shifts the gH-V relationship of HtHV1 by just 20 mV/unit or less. The aberrant behavior of HtHV1 provides clues to the mechanism of ΔpH-dependent gating.
Fig. 9 summarizes the ΔpH dependence of HtHV1. For a variety of reasons discussed elsewhere (Cherny et al., 2015), we have adopted V(gH,max/10), the voltage at which the gH is 10% of its maximal value, as a parameter to define the position of the gH-V relationship. We find this preferable to other parameters that have been used for this purpose, such as the midpoint of a Boltzmann curve (which frequently does not fit the data well or is ill determined) or the threshold voltage at which current is first detectable (which is arbitrary, depends on the signal-to-noise ratio, and is particularly difficult to resolve when it occurs near Vrev, as frequently occurs in HtHV1). It is evident in Fig. 9 that when pHo < 7, changes in pHo shift V(gH,max/10) by more than 40 mV/unit (for reference, this slope is shown as a dashed green line in Fig. 9). HV1 in two other species (coccolithophore EhHV1 and insect NpHV1) also exhibit shifts with pHo greater than 40 mV/unit (Cherny et al., 2015; Chaves et al., 2016). At pHo higher than 7, the shift decreases, which may reflect saturation of the response caused by the ambient pH approaching the pKa of a critical titratable group. Saturation of ΔpH dependence has been observed previously in hHV1 at pH > 8 (Cherny et al., 2015).
The most striking result in Fig. 9 is the data for changes in pHi (dark red diamonds), which reveal that the position of the gH-V relationship depends only weakly on pHi. There is no clear indication of saturation, although the slope appears to increase at larger ΔpH (i.e., lower pHi). This is qualitatively like the whole-cell pHo response, which is steepest at low pHo and saturates at high pHo. Over the entire ΔpH range, the mean slope is only 15.3 mV/unit change in pHi. HtHV1 is the first HV1 in which such weak ΔpH dependence has been identified.
The rapid kinetics of HtHV1 resembles that of other snail proton channels but differs from mammalian HV1
The snail HV1, HtHV1, exhibits all of the major features of HV1 in all species studied thus far. It is highly proton selective and it is voltage gated, opening with depolarization, and opening more rapidly at more positive voltages. Furthermore, its voltage dependence is strongly modulated by pH, such that increasing pHo or decreasing pHi shifts the gH-V relationship negatively, in what has been called ΔpH-dependent gating (Cherny et al., 1995). Beyond these qualitative similarities, however, HtHV1 differs markedly from HV1 in humans and other mammalian species. The main differences include very rapid activation kinetics, steeply voltage-dependent activation kinetics, activation in a more negative voltage range, exponential rather than sigmoid activation, and distinctly aberrant ΔpH dependence. These properties are discussed below.
The first voltage-gated proton channels to be characterized by voltage clamp were in neurons from the snails L. stagnalis (Byerly et al., 1984), H. aspersa (Thomas and Meech, 1982; Mahaut-Smith, 1989b), and Helix pomatia (Doroshenko et al., 1986). All activated rapidly, with time constants, τact, of a few milliseconds. When mammalian proton currents were identified, the most obvious difference was much slower activation, with τact in the range of seconds (DeCoursey, 1991; Bernheim et al., 1993; Demaurex et al., 1993; Kapus et al., 1993) or even minutes (DeCoursey and Cherny, 1993). A more subtle difference was that mammalian HV1 activate with a distinct delay, whereas snail HV1 activate exponentially. We show here that the HtHV1 channel shares both properties with other snail HV1. Byerly et al. (1984) reported half-times for activation of less than 25 ms for proton currents in L. stagnalis neurons at pHo 7.4, as observed here at pHo 7 (Fig. 4 A).
Paradoxically, some aspects of gating in HtHV1 suggest a simple first-order transition between closed and open states
Activation and deactivation time constants in HtHV1 are of similar magnitude at voltages where they overlap. This property is typical of a simple first-order system (Scheme 1). In mammalian HV1 (DeCoursey, 1991; Cherny et al., 1995, 2001; DeCoursey and Cherny, 1996, 1997; Cherny and DeCoursey, 1999; Schilling et al., 2002), activation tends to be slower than deactivation. This asymmetrical behavior is typical of cooperatively gated multimeric channels (Hodgkin and Huxley, 1952; Hille, 2001), because all subunits must activate before the channel conducts, whereas only one subunit needs to deactivate to close the pore. In rat HV1, deactivation was rapid, pHo independent, and weakly voltage dependent at large negative voltages (Cherny et al., 1995). However, near the threshold voltage for gH activation, a second slower component of τtail appeared that was pHo dependent and of comparable magnitude to τact. Also suggestive of a first-order system in HtHV1 is that τact and τtail were slowest at the midpoint of the gH-V relationship (Figs. 3 and 6). Finally, activation kinetics was well described by a single exponential and could not be fitted reasonably with a higher-order function.
There is general agreement that the HV1 dimer in several species gates “cooperatively,” but it is less clear what this word means; in drug binding, cooperativity can be produced by quite different mechanisms (Colquhoun, 1973). One sense is that, like the Hodgkin–Huxley model, multiple subunits must move before the channel can conduct. Another sense is that, like oxygen binding to the four hemes of hemoglobin, the movement of one HV1 protomer promotes the movement of the other. The sigmoid activation kinetics of HV1 in several species (Gonzalez et al., 2010; Musset et al., 2010b; Tombola et al., 2010; Fujiwara et al., 2012) appears to reflect that both protomers must undergo a conformational change before either can conduct (first sense; Gonzalez et al., 2010) or highly cooperative gating (second sense; Tombola et al., 2010). When HV1 is forced to exist as a monomer, by splicing it with the N terminus of Ciona intestinalis voltage-sensing phosphatase or by truncating the C terminus, the current turns on exponentially and five to seven times faster than with the WT dimeric protein (Koch et al., 2008; Musset et al., 2010a,b; Tombola et al., 2010; Fujiwara et al., 2012). The dimerization of HV1 in several species appears strongly dependent on coiled-coil interactions in the C terminus (Koch et al., 2008; Lee et al., 2008; Tombola et al., 2008; Li et al., 2010). HtHV1 has extensive predicted coiled-coil in its C-terminal region. This complicates the interpretation for HtHV1, because the exponential activation and the apparently first-order kinetics suggest monomeric behavior. One explanation might be that HtHV1 exists in the membrane as a dimer because of the coiled-coil region, but the coupling between C terminus and S4 segment is dysfunctional, as can be achieved experimentally by introducing a flexible linker between S4 and the C terminus (Fujiwara et al., 2012). However, this appears unlikely to be the case, because the apparent gating charge of HtHV1 is nearly 6 e0 (5.5 ± 0.9, mean ± SD in a sample of 18 gH-V curves), based on the limiting slope of the gH-V relationship. Monomeric HV1 typically exhibit gating charge roughly half that of the dimer, 2–3 versus 4–6 e0. When the coupling between the C terminus and the S4 helix was disrupted by a flexible linker, the gating charge was halved (Fujiwara et al., 2012). Given that HtHV1 has charged amino acids in its transmembrane regions similar to those of other HV1, we assume that its gating charge has analogous origins. The gH-V relationships in Fig. 6 J also are compatible with Hodgkin–Huxley-type gating. One possibility is that a concerted rate-limiting step in opening occurs late, presumably after the conformational changes in each monomer (Gonzalez et al., 2010; Musset et al., 2010b; Villalba-Galea, 2014). The voltage-dependent movement of monomers may be so rapid in HtHV1 that the concerted opening step becomes rate limiting. Another speculative explanation for its exponential activation is that HtHV1 enjoys tighter coupling between protomers than HV1 in other species; in essence, both S4 helices move together.
The gating of HtHV1 depends steeply on voltage
Perceptibly different from mammalian HV1, the activation kinetics of snail HV1, HtHV1, is more steeply voltage dependent, giving a family of currents a distinctive gestalt (Fig. 3 A). In HtHV1, τact decreased e-fold in 13.8 mV, in contrast to several mammalian HV1, where τact changes e-fold in 40–72 mV (DeCoursey, 2003). In addition, τtail increased e-fold in 14.0 mV in HtHV1, compared with a slope typically 26–44 mV/e-fold change in τtail in mammalian cells (DeCoursey, 2003). The steeply voltage-dependent gating kinetics of HtHV1 is strikingly reminiscent of voltage-gated K+ channel behavior (Cahalan et al., 1985).
Gating kinetics in HtHV1 is strongly dependent on pHo
Byerly et al. (1984) noted that activation kinetics in snail LsHV1 slowed at lower pHo more than could be accounted for by the shift of the gH-V relationship. This is clearly true of HtHV1 as well. The τact-V relationship shifts positively with lower pHo (Fig. 6 I), but its maximum increases by roughly an order of magnitude per unit decrease in pHo. In stark contrast, in rat HV1, the τact-V relationship mainly shifted along the voltage axis with changes in pHo, with little change in kinetics. However, the τact-V relationship in rat was strongly affected by pHi, slowing fivefold per unit increase in pHi (DeCoursey and Cherny, 1995). In one study of human hHV1, gating was described by three exponentials with activation generally faster at lower pHi and deactivation faster at higher pHi (Villalba-Galea, 2014). Qualitatively similar results were reported in mouse macrophages, but the largest change was a two- to threefold slowing of τact for a 1.5-unit increase in pHi or a 2.1-unit decrease in pHo (Kapus et al., 1993). An insect HV1, NpHV1, however, exhibited nearly as strong pHo dependence of kinetics as found here in HtHV1 (Chaves et al., 2016). In contrast to the strong dependence of activation kinetics on pHi in rat (DeCoursey and Cherny, 1995), in HtHV1, τact was in a similar range at all pHi values from 5 to 8. These differences in gating kinetics among species may make it challenging to produce a single universal model that describes the voltage and pH dependence of gating in all species.
Metal binding site in HtHV1
HtHV1 was moderately sensitive to inhibition by Zn2+, the classic (Thomas and Meech, 1982; Mahaut-Smith, 1989a) and still most potent (Cherny and DeCoursey, 1999) HV1 inhibitor. Somewhat weaker effects were observed for Cd2+ (Fig. 5). The principal effects of Zn2+ on mammalian HV1 are a slowing of activation, a positive shift of the gH-V relationship, and possibly a reduction of the maximum H+ conductance, gH,max (Cherny and DeCoursey, 1999). These effects are also observed in HtHV1, but the decrease in gH,max is much more obvious, whereas the shift of the gH-V relationship is substantially weaker in HtHV1 than in hHV1. Thus, the gH-V relationship in human hHV1 is shifted more by 1 µM Zn2+ (Musset et al., 2010b) than HtHV1 is shifted by 10 µM Zn2+ (Fig. 5).
In mammalian HV1, Zn2+ binds mainly to two His: His140 and His193 in hHV1 (Ramsey et al., 2006; Musset et al., 2010b). Surprisingly, when mHV1 was crystallized, it contained a Zn2+ atom, coordinated by the corresponding two His with contributions from two acids, Glu115 and Asp119, given in Table 1 (Takeshita et al., 2014). Mutation of both acids simultaneously decreases Zn2+ affinity of mHV1, but neutralizing either alone does not (Takeshita et al., 2014). As indicated in Table 1, three of these four corresponding residues are conserved in HtHV1: Glu114, Glu118 (conservatively replacing Asp), and His201, with Val254 replacing the second His. Consistent with the partial conservation of the mammalian Zn2+ site, Zn2+ was generally less potent in HtHV1 but still quite effective. The main difference in the presumed Zn2+ binding residues in HtHV1 is the lack of His193. One might therefore speculate that His193 in human hHV1 is important in Zn2+ shifting the gH-V relationship positively. Evidently, when the binding site includes His193 located in the external S2–S3 linker, Zn2+ binding biases the membrane potential more effectively. Coordination by four amino acids is more typical of a structural Zn2+ binding site, whereas catalytic Zn2+ binding sites usually have three amino acids and one water as a ligand (Auld, 2001). Of interest is a study showing that the metal transport site of ZnT transporters is selective for Zn2+ over Cd2+ when the four ligands are 2 His + 2 acids, but cannot discriminate the two metals with 1 His + 3 acids (Hoch et al., 2012). The HtHV1 channel has 1 His + 2 acids and is moderately selective for Zn2+ over Cd2+. As shown in Table 1, the NpHV1, CiHV1, CpHV1, SpHV1, and DrHV1 channels share a 1 His + 3 acids scheme and are much less sensitive to Zn2+ than mammalian HV1 (Cd2+ was not tested) and generally less sensitive than HtHV1. Intriguingly, the D145H mutation in NpHV1 results in 2 His + 2 acids, which markedly increases its Zn2+ sensitivity (Chaves et al., 2018). Empirically, Table 1 indicates that the configurations of HV1 for Zn2+ binding to HV1 in order of decreasing efficacy are: 2 His + 2 acids > 1 His + 2 acids > 1 His + 3 acids. It appears that the 1 His + 3 acids motif is somewhat less favorable for Zn2+ binding than 1 His + 2 acids as found in HtHV1, which seems paradoxical, because the 1 His + 3 acids motif has four ligands instead of three, possibly plus water. Perhaps geometrical factors can be more important than the number of ligands.
The gH-V relationship of HtHV1 depends more on pHo and less on pHi than HV1 in other species
A unique property of HV1 is that its voltage-dependent gating is strongly modulated by pH in a manner called ΔpH dependence (Cherny et al., 1995). The gH-V relationship is shifted equally by increasing pHo or decreasing pHi, by −40 mV/unit change, thus responding to the pH gradient (ΔpH) rather than to the absolute pH (Cherny et al., 1995). The practical consequence is that HV1 opens only when the electrochemical gradient for H+ is outward, such that when the channel opens it will always extrude acid from the cell (Doroshenko et al., 1986; DeCoursey and Cherny, 1994). To a rough approximation, all HV1 appear to shift by 40 mV/unit at all pH values (DeCoursey, 2003). Until recently, the rare exceptions to this rule of forty were ignored as anomalies, perhaps reflecting difficulties of the measurements, in particular with control over pH (DeCoursey and Cherny, 1997). However, measurements explicitly addressing this point revealed that the ΔpH-dependent gating of hHV1, kHV1, and EhHV1 does indeed deviate by saturating at high pH, namely above pHo 8 or pHi 8 (Cherny et al., 2015). Byerly et al. (1984) reported little shift between pHo 7.4 and 8.4 in L. stagnalis, and this observation is consistent with the saturation at high pHo observed here for HtHV1 (Fig. 9). The slope in HtHV1 begins to decrease above pHo 7 (Fig. 9), suggesting that saturation begins at lower pHo than in hHV1. Saturation of ΔpH-dependent gating suggests that pH is approaching the effective pKa of one or more titratable groups that sense pHo. Given this interpretation, the effective pKa is roughly 1 unit lower in HtHV1 than in hHV1.
Another deviation from the rule of forty is that the gH-V relationship in HtHV1 shifted ∼60 mV/unit change in pHo between pHo 5 and 7 (Fig. 9), well above the classic value of 40 mV/unit change in pHo (Cherny et al., 1995). This unusual property is shared by several disparate species, including other snails. In H. pomatia, the shift was 63 mV from pHo 7.5 to 6.6 (Doroshenko et al., 1986). In L. stagnalis, the shift was 46 mV from pHo 7.4 to 6.4 (Byerly et al., 1984). Changes in pHo in a coccolithophore EhHV1 produced shifts of ∼50 mV/unit (Cherny et al., 2015). An insect HV1 (NpHV1) shifts 54 mV/unit change in pHo (Chaves et al., 2016).
More dramatically, changes in pHi produced much smaller shifts of the gH-V relationship in HtHV1 than the 40 mV in mammalian species (Cherny et al., 1995). The mean shift in HtHV1 between pHi 5 and 9 was only 15.3 mV/unit (Fig. 9). This is in remarkable agreement with the 15 mV/unit reported in the snail LsHV1 between pHi 5.9 and 7.3 (Byerly et al., 1984). Meech (2012) recently emphasized the stronger effects of pHo over pHi after reanalyzing old data. However, in another snail, H. pomatia, HpHV1 apparently shifted normally, roughly 30–50 mV/unit change in pHi (Doroshenko et al., 1986), so on this point it is not possible to generalize about molluscan HV1.
The ΔpH dependence of mammalian HV1 results in only outward H+ currents under most circumstances, which is crucial to many if not all of its functions (DeCoursey, 2003). One striking consequence of the anomalous ΔpH dependence of HtHV1 and perhaps of other snail HV1 is that inward currents are readily observed at certain ΔpH. Even at symmetrical pH, there are often inward currents. More conspicuously, because of the weak dependence on pHi, an inward pH gradient (ΔpH < 0) produces inward currents over an extensive voltage range (e.g., Fig. 8 C). Inward currents would affect neuronal excitability by providing a depolarizing current. They at first appear incompatible with an early proposal that proton currents in snail neurons function to extrude protons that enter via Ca2+/H+ exchange after each Ca2+-mediated action potential (Ahmed and Connor, 1980; Thomas and Meech, 1982; Byerly et al., 1984), but under normal conditions of an outward H+ gradient, inward currents would likely not be activated. Nevertheless, the possibility arises that HV1 might mediate action potentials in molluscan neurons under certain conditions, although Ca2+ channels are thought to be primarily responsible (Hagiwara and Byerly, 1981). HV1 appears to mediate action potentials in bioluminescent dinoflagellates (Fogel and Hastings, 1972; Smith et al., 2011; Rodriguez et al., 2017).
As L. stagnalis and H. trivolvis snails live in similar habitats, we expect that their proton channels should function similarly. This view is supported by the similarity of the LsHV1 sequence to that of HtHV1, especially in the S2/S3 region that we have identified for its importance in pHi sensing in the accompanying paper (Cherny et al., 2018). In that paper, we identify a single amino acid difference between hHV1 and HtHV1 that appears largely responsible for the difference in pHi sensing.
We thank Kristie Bishop (Albany State University) for technical assistance with Western blotting experiments. We gratefully acknowledge heroic efforts by David Colquhoun to disambiguate our fuzzy thoughts on cooperativity.
This work was supported by the National Institutes of Health (grants GM121462 to T.E. DeCoursey and GM102336 to T.E. DeCoursey and S.M.E. Smith) and the National Science Foundation (grant MCB-1242985 to T.E. DeCoursey and S.M.E. Smith and Neuroscience Cluster award 0843173 to V. Rehder). Ms. Bishop was a participant in the Kennesaw State Research Experience for Undergraduates, Chemistry and Biochemistry Summer Undergraduate Research Experience (National Science Foundation CHE-1560329).
The authors declare no competing financial interests.
Author contributions: Conceptualization: S.M.E. Smith and T.E. DeCoursey; data curation: S.M.E. Smith and T.E. DeCoursey; formal analysis: V.V. Cherny, D. Morgan, L.R. Artinian, V. Rehder, and T.E. DeCoursey; funding acquisition: V. Rehder, S.M.E. Smith, and T.E. DeCoursey; investigation: S. Thomas, V.V. Cherny, and D. Morgan; project administration: S.M.E. Smith and T.E. DeCoursey; resources: S. Thomas, L.R. Artinian, V. Rehder, and S.M.E. Smith; visualization: S.M.E. Smith and T.E. DeCoursey; writing (original draft): T.E. DeCoursey; writing (review and editing): V.V. Cherny and S.M.E. Smith.
Richard W. Aldrich served as editor.