Articles

X-ray crystal structure of voltage-gated proton channel

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Kohei Takeshita1–3, Souhei Sakata2,3, Eiki Yamashita1, Yuichiro Fujiwara2, Akira Kawanabe2, Tatsuki Kurokawa2,4, Yoshifumi Okochi2, Makoto Matsuda1, Hirotaka Narita1, Yasushi Okamura2,3 & Atsushi Nakagawa1,3 The voltage-gated proton channel Hv1 (or VSOP) has a voltage-sensor domain (VSD) with dual roles of voltage sensing and proton permeation. Its gating is sensitive to pH and Zn2+. Here we present a crystal structure of mouse Hv1 in the resting state at 3.45-Å resolution. The structure showed a ‘closed umbrella’ shape with a long helix consisting of the cytoplasmic coiled coil and the voltage-sensing helix, S4, and featured a wide inner-accessible vestibule. Two out of three arginines in S4 were located below the phenylalanine constituting the gating charge–transfer center. The extracellular region of each protomer coordinated a Zn2+, thus suggesting that Zn2+ stabilizes the resting state of Hv1 by competing for acidic residues that otherwise form salt bridges with voltage-sensing positive charges on S4. These findings provide a platform for understanding the general principles of voltage sensing and proton permeation. Voltage-gated ion channels, such as the voltage-gated Na+, Ca2+ and K+ channels (Nav, Cav and Kv) are key proteins involved in membrane excitability. They have essential roles in neural function, muscle contraction and secretion. These proteins are composed of six transmembrane segments (S1 to S6); S1 to S4 constitute the VSD, and S5 and S6 form the ion-conduction pore1,2 Voltage-gated proton channels were first suggested to be involved in bioluminescence in dinoflagellates3 and were later discovered in snail neurons4, blood cells, airway epithelia, sperm, microglia and oocytes5,6. A voltage-sensor protein, Hv1 (or voltage-sensor domain–only protein (VSOP)), was discovered in Mus musculus and Homo sapiens and identified as the molecular correlate of the voltage-gated proton channel7,8. Hv1 is required for high-level superoxide production by phagocytes, through its tight functional coupling with NADPH oxidase, to eliminate pathogens9–11. Hv1 is also expressed in human sperm and has been suggested to regulate motility through activating alkalization-activated calcium channels12. The activities of Hv1 also have pathological implications, such as exacerbation of ischemic brain damage13 and progression of cancer14, and are reported to be suppressed by external Zn2+ (ref. 15). Hv1 has a VSD that is similar to the VSD of canonical voltagegated ion channels, but it lacks a pore domain (Fig. 1a), and protons permeate through the VSD. Hv1 is expressed as a dimer in biological membranes, with each monomer containing a separate proton pathway16–18. The fourth transmembrane segment (S4) of Hv1 has multiple basic residues, which are characteristic of the VSDs of voltage-gated ion channels and voltage-sensing phosphatases (Supplementary Fig. 1a). Electrophysiological studies19–21 indicate that S4 moves upward relative to other helices upon activation19, as it does in Kv and Nav channels. Voltage-dependent gating is suppressed by extracellular Zn2+ at micromolar levels15. An acidic residue,

Asp108 (Asp112 in human Hv1) in S1, is highly conserved among Hv1 orthologs and is critical for proton selectivity22. A recent study has suggested that Phe146 (Phe150 in hHv1) on S2 is located close to the ion-permeation pathway and is critical for inhibition by a guanidine compound, 2-guanidinobenzimidazole (2GBI), that acts as an open-channel blocker23. Although several models for molecular mechanisms of gating and permeation have been proposed21,24–27, it remains unclear how voltage sensing is coupled to proton permeation in the VSD because structural information on Hv1 is lacking. Here we present the first X-ray crystal structure of mouse Hv1 (mHv1). RESULTS Overall structure of a mouse Hv1 chimeric channel We prepared mHv1 by using various constructs including the wildtype (WT) sequence. We replaced the C-terminal coiled-coil region with a leucine-zipper motif of the transcriptional activator GCN4 from Saccharomyces cerevisiae to increase the thermostability of the molecule28. Then we replaced the region from the middle of S2 to the middle of S3 on the cytoplasmic side with the corresponding region from a Ciona intestinalis voltage-sensing phosphatase (Ci-VSP)29. In addition to these two chimerizations, we removed the N-terminal 74 residues. As a result, the chimeric protein (mHv1cc, Supplementary Fig. 1b) showed high thermostability with a melting temperature (Tm) of 70.6 °C, as measured with a thermal-shift assay using 7-diethylamino-3-(4′-maleimidylphenyl)4-methylcoumarin (CPM) (Supplementary Fig. 1c,d). We succeeded in crystallization with this construct. The proton current of the crystallization construct (mHv1cc) expressed in HEK293T cells showed voltage dependence with a threshold that shifted depending on the pH difference across the cell membrane (Fig. 1b and Supplementary Fig. 1e,f) and, with

1Institute

for Protein Research, Osaka University, Suita, Japan. 2Graduate School of Medicine, Osaka University, Suita, Japan. 3Institute for Academic Initiatives, Osaka University, Suita, Japan. 4Present address: Graduate School of Engineering, Kyoto University, Kyoto, Japan. Correspondence should be addressed to Y. Okamura ([email protected]) or A.N. ([email protected]). Received 22 October 2013; accepted 3 February 2014; published online 2 March 2014; doi:10.1038/nsmb.2783

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Figure 1  Electrophysiological properties and crystal structure of mHv1cc. (a) Schematic illustration of the Kv channel and Hv1. Hv1 consists of only a VSD (S1–S4) and lacks an ion-pore domain (S5 and S6). (b,c) Representative current-voltage (I-V) plots (b) and the reversal potentials (Erev; c) recorded with or without pH gradient. The open- and closed-circle symbols show pHout/pHin of 7.0/6.0 and 7.0/7.0, respectively. The closedtriangle symbols show the I-V curve of WT mHv1 in pHout/pHin of 7.0/7.0. Error bars, mean ± s.e.m. (n = 5 technical replicates). (d) Ribbon model of crystal structure of mHv1cc, viewed parallel to the membrane from three different angles rotated 90° along the vertical axis. The four transmembrane segments S1 (green), S2 (yellow), S3 (orange) and S4 (magenta) connected to the cytoplasmic coiled-coil region (pink) are shown. The N-terminal cytoplasmic helix (S0) is in cyan. Three arginine residues (Arg201, Arg204 and Arg207) in S4 are shown as stick models.

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reversal potentials, shifted under distinct pH conditions in accordance with the Nernst equation (Fig. 1c). In addition, we observed voltage-gated proton-channel activity with the protein reconstructed in proteoliposomes (Supplementary Fig. 1h). These results indicate that the crystallization construct exhibited the basic electrochemical properties of a voltage-gated proton channel. The initial electron density map, obtained by the multiple anomalous diffraction (MAD) method with anomalous signals of three seleno­ methionines (SeMet) in the S2 and the S4 transmembrane helices (Met138 and Met147 in S2 and Met217 downstream of S4) clearly showed a four-helix-bundle transmembrane structure. The register of sequences of the S2 and the S4 helices was easily assigned with the guide of Bijvoet anomalous difference signals of the three SeMet sites in WT mHv1cc (Supplementary Fig. 2a). We prepared two mutants, L107M L118M and L182M, to identify the topology of the transmembrane helices (Supplementary Fig. 2b,c). We modeled the four transmembrane helices (His95–Lys121, Val132–Gly158, Phe162–Phe186 and Phe191–Ser215), the cytoplasmic coiled-coil (Arg216–Leu241) and the N-terminal short helix (Phe84–Ser93) and refined the structure at 3.45-Å resolution (Fig. 1d and Supplementary Fig. 2e). In addition, we found the Bijvoet anomalous difference signal of zinc in the four transmembrane helices (Fig. 2a,b and Supplementary Fig. 2d). Crystallographic results are summarized in Table 1 and Supplementary Table 1. Hv1 consists of two functional domains: the VSD and the cytoplasmic coiled-coil domain. The crystal structure of mHv1cc showed that the entire shape looks like a ‘closed wagasa’ (traditional Japanese umbrella) with four helices opening wider toward the intracellular side (Fig. 1d). The fourth transmembrane helix (S4) was directly connected to the cytoplasmic coiled-coil region to form a slightly bent, long helical structure. The structure of mHv1cc in the crystal showed a trimer related by a crystallographic three-fold axis. The mHv1cc expressed in HEK293T cells formed a dimer similar to that of WT Hv1, as shown by a crosslinking assay (Supplementary Fig. 1g). The crystallographic trimer of mHv1cc might be induced by the trimerization property of GCN4 under crystallization conditions because GCN4 can be oligomerized either in dimeric or in trimeric forms, both in solution and in crystals30.

S0

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Although the trimeric structure might be an artifact caused by the interaction of the GCN4 coiled-coil region, the crystal structure of mHv1cc captures the molecular mechanism of the voltage-gated proton channel activity because artificially trimerized mHv1 channels with amino acid replacement in the coiled-coil motif exhibit properties indistinguishable from those of the WT voltage-gated proton channels, except for reduced cooperativity in channel activity31. The structure also showed a short helix (S0), which was located almost perpendicular to the transmembrane helices, at the N-terminal region upstream of S1. The position of S0 corresponded to the location of head groups of lipids, as predicted from the amino acids of other helices and loops (Fig. 1d and Supplementary Fig. 2e), thus suggesting that the S0 helix might be important for anchoring Hv1 to the membrane. Zn2+ binding of mHv1cc We identified the Zn2+ in mHv1cc by Bijvoet anomalous difference Fourier maps with the data collected below and above the absorption edge of zinc (Fig. 2a,b and Supplementary Fig. 2d). The binding of Zn2+ indicates that the crystal structure of mHv1cc represents the resting state because Zn2+ specifically inhibits activities of voltagegated proton channels. Two histidine residues, His136 and His189 of mHv1cc, corresponding to residues known to be critical for Zn2+ binding in hHv1 (ref. 32), exist at positions that would be able to coordinate Zn2+. The distance between the Zn2+ and the Cα atom of His136 was about 5 Å. His189 was disordered and could not be modeled in the electron density map, but it also would be positioned near the Zn2+ ion. In addition to these histidine residues, we found two negatively charged residues, Glu115 and Asp119, near the Zn2+. These two acidic residues were located too far from Zn2+ to make direct contacts but may be involved in placing water molecules around Zn2+

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Figure 2  Zn2+-binding site and Zn2+ sensitivity of mHv1 mutants. (a,b) Zn2+ anomalous difference map, drawn in blue mesh and contoured at 5.0σ. These maps were calculated from 20.0-Å to 5.0-Å resolution from a reflection data set collected at 1.275 Å. Views from different directions are shown in a and b. (c,d) Zn2+ sensitivity of WT (c) and E115S D119S (d) mHv1. Voltage-gated proton currents were elicited by test pulses to 100 mV (pHout/pHin = 7.0/7.0). The holding potentials was −60 mV. Black, 0 µM Zn2+; red, 1 µM Zn2+; blue, 10 µM Zn2+. (Additional data are in Supplementary Fig. 2g–i). (e) Normalized current amplitudes plotted against Zn 2+ concentration. Black, WT (n = 4); red, E115S D119S (n = 4); blue, E115S (n = 4); green, D119S (n = 5); yellow, ∆N∆C (n = 5). Error bars, mean ± s.d. (n values are for technical replicates).

through hydrogen bonds, as found in the NMDA receptor NR2B33. The Glu115 and Asp119 residues are highly conserved among the Hv1s from many species including humans (Supplementary Fig. 3). To examine whether Glu115 and Asp119 were actually essential residues for Zn2+ binding, we replaced the sites with serine in mHv1. Single mutants of either E115S or D119S exhibited a sensitivity to Zn2+ indistinguishable from that of the WT mHv1 channel, thus indicating that either Glu115 or Asp119 is dispensable for sensitivity to Zn2+ (Fig. 2c–e and Supplementary Fig. 2g–i). In contrast, the double mutant (E115S D119S) was clearly resistant to Zn2+. Thus, a Zn2+ ion is bound in the VSD structure, and four residues—His136, His189, Glu115 and Asp119—are likely to contribute to its coordination.

Position of sensor helix, S4, in VSD in mHv1cc A series of arginines on S4 function as voltage-sensing residues, and S4 is known to change its orientation relative to other helices in the VSD upon membrane-potential change1. Several VSD structures have been determined by X-ray crystallography, and all of these structures were considered to be in the activated state, as evidenced by their biophysical and electrophysiological properties34. All of the known S4 structures in VSDs show that the positively charged residues line the same side of S4. Despite the limited resolution of the electron density of mHv1cc, the electron density of the region from Arg201 (R1) to Arg204 (R2) was apparently narrower than that of the other regions of the S4 helix. This suggests that the Arg201–Arg204 region formed a 310 helix, whereas the other regions in S4, including around Arg207 (R3), formed an Table 1  Data collection and refinement statistics α-helix (Supplementary Fig. 2f). The 310 S76-mHv1cc SeMet1b helix can form a linear alignment of residues with three residues per helix turn. Data collection Consistent with this, the S4 helices of other Space group P 63 P 63 VSDs in the activated state show a 310-helix Cell dimensions structure in the region forming the gating   a, b, c (Å) 86.5, 86.5, 89.6 83.2, 83.2, 89.6 82.6, 82.6, 89.4 83.2, 83.2, 89.6 pore35–37. In the current structure of mHv1cc,   α, β, γ (°) 90.0, 90.0 120.0 90.0, 90.0, 120.0 90.0, 90.0, 120.0 90.0, 90.0, 120.0 it seemed that two of the sensor residues, Peak Inflection Remote Arg204 (R2) and Arg207 (R3), slid toward Wavelength 0.90000 0.97884 0.97930 0.96407 the inner-membrane side relative to the Resolution (Å) 100.0–3.45 50.00–4.30 50.00–4.30 50.00–4.20 conserved phenylalanine, Phe146, on S2 in (3.51–3.45)a (4.37–4.30)a (4.37–4.30)a (4.37–4.20)a a charge-transfer center (Fig. 3a). However, Rmerge 0.042 (0.747) 0.074 (0.0642) 0.051 (0.332) 0.057 (0.556) many positive residues on S4 in other VSDs I / σI 44.4 (1.98) 33.8 (2.44) 31.4 (3.63) 29.0 (1.79) in the activated state were located above the Completeness (%) 99.1 (100.0) 99.2 (100.0) 98.2 (89.3) 98.0 (91.7) corresponding phenylalanine34–39 (Fig. 3 and Redundancy 5.6 (4.4) 7.1 (5.7) 4.6 (5.0) 4.3 (5.0) Supplementary Fig. 2j,k). Taken together, the Zn2+ binding and location of S4 indicate Refinement that this is a resting state of the mHv1cc Resolution (Å) 28.3–3.45 VSD structure. No. reflections 5,008 Resting-state structures of Kv channels40–42 Rwork / Rfree 0.341 / 0.357 and bacterial Nav channels43 have been No. atoms extensively studied by several approaches   Protein 1,085 including molecular modeling based on B factors activated-state structures of VSDs. In the   Protein 196.0 deepest resting state of Kv channels, R1 of S4 r.m.s. deviations is located close to the highly conserved hydro  Bond lengths (Å) 0.011 phobic residue of the gating charge–transfer   Bond angles (°) 1.500 aValues in parentheses are for highest-resolution shell. bData sets were derived from the T57-mHv1cc construct (Cryst-A). center, phenylalanine on S2. Given that R1

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Figure 3  Comparison of S4 position between Kv1.2-2.1 NavAb mHv1cc and other VSDs. (a) The VSD structures mHv1cc (mHv1cc; Kv1.2-2.1, Shaker family voltagegated potassium channel Kv1.2-Kv2.1 paddle chimera channel; and NavAb, voltage-gated R1 R2 R3 sodium channel from Acrobacter butzleri), mHv1cc viewed parallel to the membrane. Positively Kv1.2-2.1 charged residues, which are known to act as NavAb effective charges in voltage sensing, are shown as stick models (magenta). The highly conserved hydrophobic residues (phenylalanine) on S2 are shown as yellow spheres. The N-terminal helix (S0, cyan) and transmembrane helices (S1, green; S2, yellow; S4, magenta) are shown by a Cα-traced model. Structures of VSDs were extracted from this structure (PDB 3WKV), Kv1.2-2.1 (2R9R37) and NavAb (3RVY35). (b) Sequence alignment of S4 segments among mHv1cc, Kv1.2-2.1 and NavAb by Clustal W54. The red letters show the residues acting as effective charges, which are involved in membrane-potential sensitivity.

of Hv1 corresponds to R2 of the Kv1.2-2.1 chimera and Shaker Kv channel (Supplementary Fig. 1a), S4 of Hv1 in our structure is positioned less deeply than in the deepest resting state of the Kv channel. As is well known in canonical voltage-gated ion channels, voltagegated proton currents exhibit a delay in activation because the holding potential is more negative, a phenomenon called the Cole-Moore effect, thus suggesting that there exist multiple resting states in Hv1 (ref. 5). It is possible that our structure represents an intermediate resting state41 and that S4 could be positioned more deeply in the completely deactivated state of Hv1. It should also be noted that the positions of arginine residues of S4 in our structure, as compared with the resting-state structure of native mHv1, may be slightly shifted because Phe146, which was used as the reference position, is adjacent to the inserted Ci-VSP sequence in mHv1cc. Double-hydrophobic-layer structure The crystal structure of mHv1cc showed two hydrophobic layers (Fig. 4). The lower layer at the cytoplasmic side (HL in) included Phe146 (Phe150 in hHv1) and Phe178 (Phe182 in hHv1). These residues are highly conserved among species (Supplementary Fig. 3). In particular, Phe146 (Phe150 in hHv1) is an important residue for binding a 2GBI23. We found an upper layer at the extracellular side (HLex), which consisted of four highly conserved hydrophobic residues from four helices (Val112 (S1), Leu143 (S2), Leu185 (S3) and Leu197 (S4)). Residues forming the HL ex are not conserved in the VSDs of other voltage-sensing proteins (Supplementary Fig. 1a). HLex and HLin might also prevent the penetration of water molecules, which can be proton carriers. The structure of mHv1cc showed a cavity that was shielded by the two hydrophobic layers. The cavity had enough space to accommodate several water molecules (Fig. 4 and Supplementary Fig. 4). In our model of the mHv1cc structure, Asp108 (Asp112 in hHv1), which is critical for selective proton permeation22, seems to be located in the hydrophobic layer. However, recent PEGylation experiments detecting the accessibility of maleimide reagents44 suggested that Asp108 faces the aqueous vestibule. The two hydrophobic layers probably have distinct roles in regulating the proton-conduction pathway. Previously, state-dependent motion of VSD was studied elec­ trophysiologically by examining the accessibility of an MTS reagent to a mutagenized cysteine on S4 of the sea-squirt Hv1 ortholog, CiHv1 (refs. 19,20), and the accessibility of Zn2+ to a mutagenized cysteine on S4 of hHv1 (ref. 21). The S4 position in our structure was more consistent with the results of those studies under the resting state19,20 (Supplementary Fig. 4). The mHv1cc showed a wide inner-accessible vestibule below HLin, and water molecules could access up to half of the molecule from the

cytoplasmic side (Fig. 4 and Supplementary Fig. 4). This structure of the water-accessible vestibule is also consistent with previous findings of electrophysiological and biochemical studies of the mHv1 S4 segment: mHv1 truncated at Ala206 (between Arg204 and Arg207) retained voltage-gated proton-channel activities, and cysteine introduced into individual sites ranging from Ile212 (Ile216 in hHv1) to Ala206 (Ala210 in hHv1) on S4 were accessible to a maleimidecontaining reagent, 4-acetamido-4′-maleimidylstilbene-2,2′disulfonic acid (AMS), as detected by a PEGylation protection assay45. The water-accessibility profile in this structure is also consistent with our recent results of comprehensive cysteine scanning on all sites of S1 to S4 helices44. Additional discussion is presented in the Supplementary Note. Proposed model of dimer structure The crystal structure of mHv1cc showed a trimer related by the crystallographic c axis. This trimer structure might be caused by the interaction of the coiled coil in the GCN4 region because the GCN4 coiled-coil structure exhibits either dimer or trimer structures, according to small changes in the environment46. We modeled the dimeric mHv1cc structure by superimposing the C-terminal GCN4 region of mHv1cc onto the WT dimeric coiled-coil structure of mHv1 from our previous observations28. In this dimer model, there was no region where the atoms clashed, including the N-terminal S0 helix

L197 L185

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Figure 4  Double hydrophobic layers (HLex and HLin), cavity and inner vestibule. The two hydrophobic layers consist of the hydrophobic residues, which are shown by a stick model around the constriction sites (HL ex and HLin). These water-accessible regions are represented by Connolly surface (probe radius = 1.4 Å, blue surface). The space size of the intermolecular cavity is ~8 Å. The black dashed lines show the putative surface of the lipid-bilayer membrane. The lower hydrophobic layer, HL in, is located distally from the inserted fragment of Ci-VSP.

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articles region. The S0 helix is located outside the dimer structure of mHv1cc as if the S0 helix anchored the Hv1 molecule to the membrane (Supplementary Fig. 5). The cytoplasmic coiled-coil motif connects with the helix of S4, thus forming the longest helix, as previously suggested28, and two S4s run in parallel next to each other between two protomers. This is also consistent with recent findings of a systematic study of cross-linking between two S4s47. Our dimer model differs from a previous model based on a study on Zn2+ sensitivity of hHv1, in which a single Zn2+ was claimed to sit at the interface of two monomers within dimers, binding to one histidine on the S1–S2 loop on one monomer and another histidine on the S3–S4 loop of the other monomer32. In our model, unlike the previous study on hHv1 (ref. 32), Zn2+ sensitivity of the N terminus– and C terminus–deleted monomeric channels (∆N∆C; Supplementary Fig. 2i) was not substantially distinct from that of the WT dimeric channels (Fig. 2e). Moreover, our mHv1cc structure suggests that Zn2+ is coordinated by the residues within an individual protomer. It has been proposed that an extracellular loop adjacent to S1 makes contacts with the corresponding loop in an adjacent subunit and that the charged residues in this region are critical for cooperative gating17,48. In our structure, the S1 is not close to the S1 in the adjacent protomer within dimers. This raises the possibility that the configuration of spatial arrangement of four helices of the VSD is more dynamic than was previously thought, and S1-S1 interactions within dimers may occur during the transition from the closed state to the open state. DISCUSSION The mHv1cc crystal structure is the first, to our knowledge, of a VSD structure in the resting state. Our judgment that this structure represents a resting state rather than an activated state is based on three findings: (i) the structure had bound Zn2+, which is known to inhibit gating of voltage-gated proton channels15, (ii) the position of the arginines in S4 was lower in our structure than in previously resolved VSD structures in the activated state and (iii) the position of S4 in our structure was consistent with the maps of MTS-reagent accessibility by electrophysiology and maleimide-reagent accessibility by PEGylation protection assay, both in the resting state19,20,44. Molecular dynamics simulations have predicted the structure of Hv1 in the activated state21,24–27. We propose that negative charges of Glu115 (Glu119 in hHv1) and Asp119 (Asp123 in hHv1) interact with Zn2+ in the inhibited resting state but with arginine on S4 in the activated state (Fig. 5). In other words, Zn2+ prevents Glu115 (Glu119 in hHv1) and Asp119 (Asp123 in hHv1) from interacting with arginine, thus leading to inhibition of the S4 upward motion. The coordinating geometry of Zn2+, specific to the resting state of VSD, is consistent with the previous finding that Zn2+ does not affect the kinetics of deactivation15, i.e., the transition from activated state to resting state. The suppression of motion of S4 by metal ions through binding to negatively charged residues on the VSD from the extracellular side observed in our study could represent a general regulatory mechanism shared by many voltage-dependent ion channels, including Shaker-type Kv channels49,50 and the ether-à-go-go (eag) Kv channel51, in which metal ions slow activation but do not affect deactivation. It is known that the intratesticular environment is high in Zn2+ concentration, and the channel activity of hHv1 is inhibited by binding of Zn2+ (ref. 12). When sperm is transferred from the testis into the epididymis or vagina, where the Zn2+ concentration is substantially lower, proton-channel activity is derepressed. This may lead to alkalization of intracellular pH in sperm, resulting in sperm motility through the activities of the CatSper channel, an alkalizationactivated cation channel52. Zn2+ inhibition of Hv1 may also have 356

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Figure 5  Gating mechanism of Hv1. (a) Schematic illustration of Zn2+bound resting state of mHv1cc. The red sphere represents Zn 2+. (b) Proposed schematic illustration of the activated state of mHv1cc.

physiological relev­ance in the brain. Hv1 is expressed in microglia11 and is involved in neuronal cell death in brain ischemia, through its regulatory role in the production of reactive oxygen species13. Zn2+ is known to be released from synapses53, and its concentration in the vicinity of microglia may fluctuate depending on neuronal activity. Future elucidation of crystal structures of Hv1 at higher resolution, and of other gating states, will reveal detailed operating mechanisms of Hv1 and general principles of the VSD that are shared among voltage-gated ion channels and voltage-sensing phosphatases29 upon membrane-potential changes. Structural information, combined with modeling, will also help to unravel mechanisms underlying the efficient conduction of protons, which are available only at concentrations lower than those of other cations by a factor of 10−4 to 10−5. Methods Methods and any associated references are available in the online version of the paper. Accession codes. Coordinates and structure factors have been deposited in the Protein Data Bank under accession code 3WKV. Note: Any Supplementary Information and Source Data files are available in the online version of the paper. Acknowledgments We wish to thank N. Nakamura, K. Nishiwaki, M. Kobayashi and W. Kumano for their support with experiments. We thank D.M. Standley for suggestions and comments on the manuscript. We are grateful to S. Ogasawara, S. Iwata and S. Yokoyama for advice on membrane-protein crystallization. We wish to thank T. Tsukihara and Y. Yoneda for encouragement throughout this project. We also thank all members of the Nakagawa and Okamura laboratories for their suggestions and comments. This work was supported by the Target Proteins Research Program (A.N. and Y. Okamura) and the Platform for Drug Discovery, Informatics, and Structural Life Science (A.N.) from the Ministry of Education, Culture, Sports, Science and Technology (MEXT) in Japan, by Grants-in-Aid for basic research S (no. 25253016) and A (no. 21229003) (Y. Okamura and A.N.), a Grant-in-Aid for Scientific Research on Innovative Areas (no. 24111529) (Y. Okamura) from the Japan Society for the Promotion of Science, by the JAXA-GCF (Japan Aerospace Exploration Agency–Granada Crystallization Facility) project “High quality protein crystallization project on the protein structure and function analysis for application,” conducted from the Japan Aerospace Exploration Agency (A.N.), and by the National Project on Protein Structural and Functional Analyses from the MEXT (A.N.). Diffraction data were collected at the Osaka University beamline BL44XU at SPring-8 (Harima, Japan) under proposal numbers 2010A6500, 2010B6500, 2011A6500, 2011B6500, 2012A6500, 2012B6500 and 2013A6500. The detector, MX225HE, is financially supported by Academia Sinica and the National Synchrotron Radiation Research Center in Taiwan, Republic of China. AUTHOR CONTRIBUTIONS K.T. expressed, purified and crystallized, collected and processed X-ray data, refined and analyzed the structure and wrote the paper. E.Y. assisted with collection

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articles and processing of X-ray data. S.S. performed physiological experiments. Y.F. assisted with physiological and structural studies. T.K., Y. Okochi and A.K. assisted with physiological studies. M.M. and H.N. assisted with protein expression. Y. Okamura and A.N. designed the study and wrote the paper. All authors commented on the manuscript. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests.

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26. Morgan, D. et al. Peregrination of the selectivity filter delineates the pore of the human voltage-gated proton channel hHV1. J. Gen. Physiol. 142, 625–640 (2013). 27. Chamberlin, A. et al. Hydrophobic plug functions as a gate in voltage-gated proton channels. Proc. Natl. Acad. Sci. USA 111, E273–E282 (2014). 28. Fujiwara, Y. et al. The cytoplasmic coiled-coil mediates cooperative gating temperature sensitivity in the voltage-gated H+ channel Hv1. Nat. Commun. 3, 816 (2012). 29. Murata, Y., Iwasaki, H., Sasaki, M., Inaba, K. & Okamura, Y. Phosphoinositide phosphatase activity coupled to an intrinsic voltage sensor. Nature 435, 1239–1243 (2005). 30. Gonzalez, L., Woolfson, D.N. & Alber, T. Buried polar residues and structural specificity in the GCN4 leucine zipper. Nat. Struct. Biol. 3, 1011–1018 (1996). 31. Fujiwara, Y. et al. Gating of the designed trimeric/tetrameric voltage-gated H+ channel. J. Physiol. 591, 627–640 (2013). 32. Musset, B. et al. Zinc inhibition of monomeric and dimeric proton channels suggests cooperative gating. J. Physiol. 588, 1435–1449 (2010). 33. Karakas, E., Simorowski, N. & Furukawa, H. Structure of the zinc-bound aminoterminal domain of the NMDA receptor NR2B subunit. EMBO J. 28, 3910–3920 (2009). 34. Yu, F.H., Yarov–Yarovoy, V., Gutman, G.A. & Catterall, W.A. Overview of molecular relationships in the voltage-gated ion channel superfamily. Pharmacol. Rev. 57, 387–395 (2005). 35. Payandeh, J., Scheuer, T., Zheng, N. & Catterall, W.A. The crystal structure of a voltage-gated sodium channel. Nature 475, 353–358 (2011). 36. Zhang, X. et al. Crystal structure of an orthologue of the NaChBac voltage-gated sodium channel. Nature 486, 130–134 (2012). 37. Long, S.B., Tao, X., Campbell, E.B. & MacKinnon, R. Atomic structure of a voltagedependent K+ channel in a lipid membrane-like environment. Nature 450, 376–382 (2007). 38. Tao, X., Lee, A., Limapichat, W., Dougherty, D.A. & MacKinnon, R. A gating charge transfer center in voltage sensors. Science 328, 67–73 (2010). 39. Jiang, Y. et al. X-ray structure of a voltage-dependent K+ channel. Nature 423, 33–41 (2003). 40. Tombola, F., Pathak, M.M., Gorostiza, P. & Isacoff, E.Y. The twisted ion-permeation pathway of a resting voltage-sensing domain. Nature 445, 546–549 (2007). 41. Delemotte, L., Tarek, M., Klein, M.L., Amaral, C. & Treptow, W. Intermediate states of the Kv1.2 voltage sensor from atomistic molecular dynamics simulations. Proc. Natl. Acad. Sci. USA 108, 6109–6114 (2011). 42. Henrion, U. et al. Tracking a complete voltage-sensor cycle with metal-ion bridges. Proc. Natl. Acad. Sci. USA 109, 8552–8557 (2012). 43. Yarov–Yarovoy, V. et al. Structural basis for gating charge movement in the voltage sensor of a sodium channel. Proc. Natl. Acad. Sci. USA 109, E93–E102 (2012). 44. Kurokawa, T. & Okamura, Y. Mapping of sites facing aqueous environment of voltagegated proton channel at resting state: a study with PEGylation protection. Biochim. Biophys. Acta 1838, 382–387 (2014). 45. Sakata, S. et al. Functionality of the voltage-gated proton channel truncated in S4. Proc. Natl. Acad. Sci. USA 107, 2313–2318 (2010). 46. Gonzalez, L., Brown, R.A., Richardson, D. & Alber, T. Crystal structures of a single coiled-coil peptide in two oligomeric states reveal the basis for structural polymorphism. Nat. Struct. Biol. 3, 1002–1009 (1996). 47. Fujiwara, Y., Kurokawa, T. & Okamura, Y. Long α-helices projecting from the membrane as the dimer interface in the voltage-gated H+ channel. J. Gen. Physiol. (in the press). 48. Qiu, F., Rebolledo, S., Gonzalez, C. & Larsson, H.P. Subunit interactions during cooperative opening of voltage-gated proton channels. Neuron 77, 288–298 (2013). 49. Gilly, W.F. & Armstrong, C.M. Divalent cations and the activation kinetics of potassium channels in squid giant axons. J. Gen. Physiol. 79, 965–996 (1982). 50. Hoshi, T. & Armstrong, C.M. Initial steps in the opening of a Shaker potassium channel. Proc. Natl. Acad. Sci. USA 109, 12800–12804 (2012). 51. Silverman, W.R., Tang, C.Y., Mock, A.F., Huh, K.B. & Papazian, D.M. Mg2+ modulates voltage-dependent activation in ether-à-go-go potassium channels by binding between transmembrane segments S2 and S3. J. Gen. Physiol. 116, 663–678 (2000). 52. Kirichok, Y., Navarro, B. & Clapham, D.E. Whole-cell patch-clamp measurements of spermatozoa reveal an alkaline-activated Ca2+ channel. Nature 439, 737–740 (2006). 53. Qian, J. & Noebels, J.L. Visualization of transmitter release with zinc fluorescence detection at the mouse hippocampal mossy fibre synapse. J. Physiol. 566, 747–758 (2005). 54. Larkin, M.A. et al. Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947–2948 (2007).

nature structural & molecular biology  VOLUME 21  NUMBER 4  APRIL 2014

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Plasmids. mHv1cc is a chimeric channel constructed for X-ray crystallography and other biochemistry studies, in which the cytoplasmic coiled coil (Val216– Asn269) and S2–S3 half intracellular side (Glu149–Phe171) were replaced with the leucine-zipper transcriptional activator GCN4 from S. cerevisiae (Arg249–Arg281) and the intracellular portion of Ci-VSP (Asp164–Leu188), respectively. cDNA encoding mHv1cc was amplified by PCR and ligated into a pFastBac1 vector (Life Technologies). For determination of the location of S1 and S3, two methionine mutants, L107M L118M and L182M, were prepared by PCR and subcloned into the pFastBac1 vector. An N-terminal 56-residue deletion mutant (T57-mHv1cc) and N-terminal 75-residue deletion mutant (S76-mHv1cc) was used for X-ray crystallography. For electrophysiological analysis of mHv1cc, a cDNA fragment of mHv1cc was subcloned into pIRESEGFP (Clontech). Genes of the other mHv1 mutants (E115S, D119S and E115S D119S) were amplified by PCR with the mHv1 gene as a template and subcloned into pcDNA3 (Life Technologies). ∆N∆C (N terminus- and C terminus-truncated mHv1) was prepared as described elsewhere16. High-level expression and purification of mHv1cc. Baculoviruses encoding mHv1cc with an eight-polyhistidine tag (8× His tag) and Tobacco Etch Virus (TEV) protease cleavage site at the N terminus were used to infect Sf9 cells with Cellfectin II (Life Technologies). Baculovirus-infected Sf9 cells were cultured at 28 °C and harvested 3 d after infection. The collected cells were resuspended in 20 mM HEPES-Na, pH 7.0, 350 mM NaCl, 5mM MgCl2, Benzonase endo­ nuclease (Merck) and Complete protease inhibitor cocktail tablets without ETDA (Roche) and disrupted with an ultrasonic disruptor UD-201 (TOMY Seiko). The membrane fraction–expressed mHv1cc was collected by ultracentrifugation at 100,000g. The membrane fraction was washed with wash buffer (20 mM HEPESNa, pH 7.0, and 350 mM NaCl). The membrane fraction was solubilized with wash buffer containing 40 mM imidazole and 1% (w/v) dodecyl-β-d-maltoside (Anatrace). The solubilized mHv1cc protein was purified by nickel-chelating resin (Ni-NTA, Qiagen), and the His tag was removed by TEV protease–fused His tag. The protease reaction mixture was applied to cobalt-chelate resin (Talon, Clontech). We collected the flow-through fraction containing mHv1cc. mHv1cc was further purified by HiTrap Q HP (GE Healthcare). Finally, mHv1cc was purified by size-exclusion chromatography with HiLoad 16/60 Superdex 200 prep grade (GE Healthcare) in 0.2% (w/v) CYMAL-5 (Anatrace), 200 mM NaCl and 20 mM HEPES-Na, pH 7.0. For crystallization, purified mHv1cc was concentrated to 10 mg ml−1 by ultrafiltration in VIVASPIN 6 concentrators with a 30-kDa MWCO (Sartorius Stedim Biotech). Before crystallization, mHv1cc protein solution was dialyzed in 0.2% (w/v) CYMAL-5, 200 mM NaCl and 20 mM HEPES-Na, pH 7.0, to remove excess detergent, with an Xpress Micro Dialyzer with a MWCO of 12–14 kDa (Scienova). To prepare selenomethionine (SeMet)-labeled mHv1cc, we used serum-free culture medium without methionine and cysteine (Life Technologies). Sf-9 cells were collected 24 h after infection, washed with sterilized PBS and transferred into medium including 20 mg L−1 SeMet (Wako) and 150 mg L−1 l-cysteine (Wako). After 4 h incubation, the cells were transferred into the medium supplemented with 50 mg L−1 SeMet and 150 mg L−1 l-cysteine. The cells were collected after 48 h, and SeMet-labeled mHv1cc was purified with the same protocol as for native protein. Thermal stability assay. Thermal stability of mHv1cc was assessed by 7-diethylamino-3-(4′-maleimidylphenyl)-4-methylcoumarin (CPM) (SigmaAldrich) assay55. Thermal denaturation of the protein was monitored by chemical reactivity of the interior cysteine residues in the folded state as a sensor for the overall structural integrity. 6 µg of the protein was dissolved in 20 mM HEPESNa, pH 7.0, 200 mM NaCl, 0.2% (w/v) CYMAL-5 and 0.1 mg ml−1 CPM. The reaction mixture was transferred to a 0.2-ml Hi-8 tube (TaKaRa). CPM reactions were performed under an elevated temperature range starting from 25 °C and ending at 98.5 °C, with a 0.5 °C/min temperature gradient and a quantitative PCR system (Stratagene Mx3005p, Agilent Technologies). The excitation wavelength was set at 387 nm, and the emission wavelength was 463 nm. Data processing and calculation of the melting temperature (Tm) was performed with MxPro QPCR software (Agilent Technologies). Crystallization and X-ray diffraction data collection. We obtained crystals by the vapor-diffusion method. The reservoir solution for crystallization consisted

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of 29–31% (v/v) PEG 400, 100 mM KCl and 100 mM Tris-HCl, pH 8.0. Equal volumes of the protein solution of mHv1cc (10 mg ml−1) and reservoir solution were mixed and incubated at 20 °C. All reflection data sets were collected on BL44XU at SPring-8 (Harima, Japan) with an MX225-HE charge-coupled device detector (Raynonix). All X-ray experiments were performed under a cryostream at 90 K. Diffraction data of SeMet derivative crystals were collected with X-ray wavelength of 0.90000 Å (remote), 0.97894 Å (peak) and 0.97954 Å (edge). Diffraction data for the identification of the zinc site were collected with X-ray wavelengths at 1.275 Å and 1.285 Å. All diffraction data were pro­ cessed and scaled with HKL2000 (ref. 56) (HKL Research). All crystals belonged to space group P63. One mHv1cc molecule was present in the crystallographic asymmetric unit. All crystallographic data are summarized in Table 1 and Supplementary Table 1. Structure determination. The initial electron density map was obtained by the multiple anomalous diffraction (MAD) method with SHELXC/D/E57 integrated in CCP4i58. The anomalous signals of three selenomethionines in the S2 and S4 transmembrane helices (Met138 and Met147 in S2 and Met217 downstream of S4) were used for assignment of the register of the S2 and the S4 transmembrane helices. Moreover, we prepared L107M L118M and L182M mutants to identify the topology of S1 and S3 transmembrane helices, using the SeMet substitutions. Beside the peak corresponding to the anomalous signal of Se, weak anomalous signal was observed in these anomalous maps (Supplementary Fig. 2b,c). As a result, we identified the unknown anomalous signal as Zn2+ bound to mHv1cc by using data from two wavelengths, collected below and above the absorption edge of zinc (1.290 Å and 1.275 Å). The highest resolution of T57-mHv1cc (native crystal; Cryst-A) was 3.55 Å. Higher-resolution data were obtained from S76-mHv1cc, and the structure was refined at 3.45-Å resolution (Fig. 1d). In addition, the Bijvoet anomalous difference signal of Zn2+ was found in S76mHv1cc at the same position as that in T57-mHv1cc. Model building was performed with Coot59, and structural refinement was carried out with PHENIX60. Crystallographic R and Rfree for 5% of the reflections excluded from the refinement were calculated to monitor the structural refinement procedures. The results of the structural analysis are summarized in Table 1. The main chain dihedral angles for 86.92% were in preferred regions of the Ramachandran plot; 8.46% were in the allowed regions; and 4.62% were in the outliers. The structure was validated with MolProbity61. All molecular graphics were produced with PyMOL (http://www.pymol.org/). Electrophysiological analysis of mHv1cc and mutants. For electrophysiological analysis, plasmids were transfected into HEK293T cells. Macroscopic currents were recorded in the whole-cell patch-clamp configuration with an Axopatch200B amplifier (Molecular Devices) or an EPC9 (HEKA Electronik). The pipette resistance in the solution was 3–11 MΩ. 60–80% of the voltage error due to the series resistance was compensated by a circuit in the amplifier. The recorded currents were low-pass filtered at 1 kHz with a quadrupole Bessel filter circuit built into the amplifier. The external solution contained 75 mM N-methyl-d-glucamine (NMDG), 1 mM CaCl2, 1 mM MgCl2, 10 mM glucose and 180 mM HEPES-Na, pH 7.0. The pipette solution contained 65 mM NMDG, 3 mM MgCl2, 1 mM EGTA and 183 mM HEPES-Na, pH 7.0. Reversal potentials were calculated as the ratio between the current amplitude at +80 mV pulse end and the tail current amplitude at −60mV. Data were analyzed with Clampfit (Axon Instruments, Inc) or PatchMaster (HEKA Electronik) and Igor Pro (WaveMetrics, Inc). In the analysis of Zn2+ inhibition, the current amplitudes at 100 mV were measured in the extracellular solutions containing 0 or 1 or 10 µM of ZnCl2 under the DAD rapid perfusion system (ALA Scientific). Proton flux by liposome containing mHv1cc. The liposome was prepared as a 1:1 mixture of l-α-phosphatidylcholine (PC) from chicken egg (Avanti) and E. coli lipid (Avanti). Protein was added to the liposome at 0.1% (w/w) protein to lipid. Proton flux was performed by fluorescence-based proton flux assay62. The proteoliposome with high-concentration K+ was diluted into low-concentration K+ buffer containing the fluorescence dye 9-amino-6-chloro-2-methoxyacridine (ACMA) (Life Technologies). Addition of valinomycin generates an electrochemical driving force for proton flux. The decrease of pH inside the liposome was monitored by fluorescence quenching of ACMA. Data were collected on an F-2500 fluorescence spectrophotometer (Hitachi) with excitation at 410 nm, emission at

doi:10.1038/nsmb.2783

490 nm, bandwidth 5 nm and an integration time of 2 s. A baseline was collected for 100 s before the addition of valinomycin (20 nM). After the fluorescence stabilized, carbonyl cyanide m-chlorophenyl hydrazine (CCCP) (Sigma-Aldrich) was added to 2.0 µM, thus rendering all vesicles proton permeable. Cross-linking analysis of mHv1cc. In the amine-reactive cross-linking, HEK293T cells expressing mHv1cc containing an HA tag were treated by disuccinimidyl suberate (DSS), and the reaction was quenched with 100 mM Tris-HCl, pH 8.0. Proteins were separated on 12.5% SDS-PAGE under reducing conditions and electrophoretically transferred to Immobilon-P (Merck, Millipore). A polyclonal rabbit anti–HA tag antibody63 (Life Technologies, cat. no. 71-5500) was used (1:500) for detection. Horseradish peroxidase– conjugated donkey anti-rabbit IgG64 (GE Healthcare, cat. no. NA9340V) was used (1:2,000) as the secondary antibody.

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55. Alexandrov, A.I., Mileni, M., Chien, E.Y., Hanson, M.A. & Stevens, R.C. Microscale fluorescent thermal stability assay for membrane proteins. Structure 16, 351–359 (2008).

56. Otwinowski, Z. & Minor, W. Processing of X-ray diffraction data collected in oscillation mode. Methods Enzymol. 276, 307–326 (1997). 57. Sheldrick, G.M. A short history of SHELX. Acta Crystallogr. A 64, 112–122 (2008). 58. Potterton, E., Briggs, P., Turkenburg, M. & Dodson, E. A graphical user interface to the CCP4 program suite. Acta Crystallogr. D Biol. Crystallogr. 59, 1131–1137 (2003). 59. Emsley, P., Lohkamp, B., Scott, W.G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010). 60. Adams, P.D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 (2010). 61. Chen, V.B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D Biol. Crystallogr. 66, 12–21 (2010). 62. Lee, S.Y., Letts, J.A. & MacKinnon, R. Functional reconstitution of purified human Hv1 H+ channels. J. Mol. Biol. 387, 1055–1060 (2009). 63. Tu, Y., Li, F. & Wu, C. Nck-2, a novel Src homology 2/3-containing adaptor protein that interacts with the LIM-only protein PINCH and components of growth factor receptor kinase-signaling pathways. Mol. Biol. Cell 9, 3367–3382 (1998). 64. Nakane, P.K. & Kawaoi, A. Peroxidase-labeled antibody. A new method of conjugation. J. Histochem. Cytochem. 22, 1084–1091 (1974).

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