1. Biochemistry and Chemical Biology
  2. Structural Biology and Molecular Biophysics
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The fluoride permeation pathway and anion recognition in Fluc family fluoride channels

  1. Benjamin C McIlwain
  2. Roja Gundepudi
  3. B Ben Koff
  4. Randy B Stockbridge  Is a corresponding author
  1. Department of Molecular, Cellular, and Developmental Biology, University of Michigan, United States
  2. Program in Biophysics, University of Michigan, United States
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Cite this article as: eLife 2021;10:e69482 doi: 10.7554/eLife.69482

Abstract

Fluc family fluoride channels protect microbes against ambient environmental fluoride by undermining the cytoplasmic accumulation of this toxic halide. These proteins are structurally idiosyncratic, and thus the permeation pathway and mechanism have no analogy in other known ion channels. Although fluoride-binding sites were identified in previous structural studies, it was not evident how these ions access aqueous solution, and the molecular determinants of anion recognition and selectivity have not been elucidated. Using x-ray crystallography, planar bilayer electrophysiology, and liposome-based assays, we identified additional binding sites along the permeation pathway. We used this information to develop an oriented system for planar lipid bilayer electrophysiology and observed anion block at one of these sites, revealing insights into the mechanism of anion recognition. We propose a permeation mechanism involving alternating occupancy of anion-binding sites that are fully assembled only as the substrate approaches.

Introduction

Microbes are protected from the cytoplasmic accumulation of environmental fluoride ion (F-) by export of the toxic anion via fluoride channels known as Flucs (Baker et al., 2012; Ji et al., 2014; McIlwain et al., 2021). These small, homodimeric ion channels are remarkable proteins in two regards: first, their unusual ‘dual-topology’ architecture, in which the two subunits of the homodimer are arranged antiparallel with respect to each other (Stockbridge et al., 2014; Stockbridge et al., 2013), yielding a double-barreled pair of pores related by twofold symmetry (Stockbridge et al., 2015; Last et al., 2016; Turman et al., 2015; Turman and Stockbridge, 2017); second, the Flucs stand out among anion channels for their extreme substrate selectivity (Stockbridge et al., 2013). In contrast to most characterized families of anion channels, which tend to be non-selective among anions and sometimes poorly discriminate against cations, the Flucs are arguably the most selective ion channels known, with >10,000-fold selectivity against the biologically abundant chloride (Stockbridge et al., 2013). This extreme selectivity prevents collapse of the membrane potential due to chloride or cation leak through the Fluc channels, which are constitutively open. Among anion channels, the stringent selectivity displayed by the Flucs is atypical. Most characterized anion channels handle the most abundant ion in their milieu, usually chloride ion (Cl-), and other halides and pseudohalides that might compete with the physiological ion are present at much lower concentrations.

Crystal structures of representative Fluc channels from Bordetella pertussis (Fluc-Bpe) and an Escherichia coli virulence plasmid (Fluc-Ec2) provide an opportunity to understand the molecular basis for anion permeation in the Flucs (Stockbridge et al., 2015; McIlwain et al., 2018). The protein possesses two deep, aqueous vestibules with an electropositive character due to an absolutely conserved arginine sidechain and a deeply buried sodium ion at the center of the protein (Turman and Stockbridge, 2017; McIlwain et al., 2020). The structures captured four electron densities assigned as fluoride ions, two in each pore, positioned near the center of the protein, at some distance from the vestibules. These ions are aligned along the polar face of TM4, referred to as the polar track. They are located 6–10 Å from the aqueous solution, with no clear aqueous pathway leading to the external solution. Mutation of the sidechains that coordinate the proposed fluoride ions inhibits fluoride throughput but does not alter the ion selectivity of these proteins (Stockbridge et al., 2015; Last et al., 2017). Thus, characterizing the rest of the fluoride permeation route is the first step toward identifying the residues responsible for fluoride ion recognition.

Here we combine x-ray crystallography, planar lipid bilayer electrophysiology, and liposome flux assays to identify access points to the polar track, including a non-specific anion-binding site at the bottom of the aqueous vestibule. We propose that fluoride ions accumulate in this electropositive vestibule before entering the fluoride-selective region of the pore, reprising a familiar feature of many ion channels. After traversing the polar track, the fluoride ions then emerge at another point in the opposite vestibule on the opposite side of the membrane, near a conserved glutamate that plays a role in discriminating against Cl-.

Results

Anions enter the fluoride pathway through the electropositive vestibule

The electropositive vestibule, lined with conserved, polar sidechains, is an obvious candidate for fluoride entry into the channel. Spherical, non-protein electron densities were observed in this region, but without additional evidence of anionic character, they were assigned as water molecules (Stockbridge et al., 2015). To test whether any of these densities might better be assigned as anions, we endeavored to crystallize Fluc channels with bromide (Br-), an anomalous scatterer. We were unable to generate diffracting Fluc-Bpe crystals in the presence of Br-, but we were successful in solving the structure of Fluc-Ec2 in the presence of 100 mM Br- (Table 1).

Table 1
Crystallography data collection and refinement statistics.
Ec2-WTEc2-S81AEc2-S81CEc2-S81A/T82AEc2-S81T
Data collection
Space groupP41P41P41P41P41
Cell dimensions
a, b, c (Å)87.6, 87.6, 14487.4, 87.4, 141.987.2, 87.2, 142.787.5, 87.5, 147.487.1, 87.1, 145.2
α, β, γ (°)90, 90, 9090, 90, 9090, 90, 9090, 90, 9090, 90, 90
Resolution (Å)34.4–3.11 (3.3–3.11)39.1–2.5 (2.6–2.5)46.7–2.9 (3.0–2.9)41.9–3.1 (3.3–3.1)28.4–2.7 (2.8–2.7)
Rmerge0.491 (2.31)0.140 (1.846)0.363 (3.437)0.723 (6.147)0.217 (2.104)
Rpim0.203 (0.938)0.057 (0.742)0.156 (1.434)0.290 (2.446)0.088 (0.833)
Mn I/σI7.2 (2.0)11.9 (1.7)9.8 (2.5)8.5 (2.1)10.3 (2.0)
CC1/20.996 (0.61)0.998 (0.61)0.98 (0.59)0.998 (0.73)0.998 (0.71)
Completeness (%)99.85 (100)99.5 (100)99.83 (100)99.85 (99.95)99.8 (100)
Multiplicity13.3 (13.9)13.7 (14.1)12.5 (13.0)13.6 (14.0)13.8 (14.4)
Refinement
Resolution33.3–3.1137.68–2.546.65–2.939.11–3.128.28–2.7
No. of reflections19,50036,59123,58020,05529,192
Rwork/Rfree23.7/27.624.0/25.222.3/25.823.0/25.221.9/25.6
Ramachandran favored93.396.595.994.796.1
Ramachandran outliers0.230.460.230.230.46
r.m.s. deviations
Bond length (Å)0.0050.0020.0020.0080.008
Bond angle (°)0.6530.5320.4890.7820.934
PDB code7KKR7KKA7KKB7KK97KK8
  1. Statistics for the highest-resolution shell are shown in parentheses. r.m.s., root-mean-square.

Anomalous difference maps show two prominent peaks, located in equivalent, non-crystallographic symmetry-related positions at the bottom of the aqueous vestibules (Figure 1A). These densities are coordinated by a sidechain that is invariant among Fluc channels, Ser81, along with the highly conserved Thr82 (Figure 1B, upper panel). In maps from previous Fluc-Bpe structures (Stockbridge et al., 2015), a positive density occupies this same position between the homologous hydroxyl sidechains (Figure 1B, lower panel). This site is exposed to bulk water in the vestibule but is likely to be partially dehydrated, with aliphatic sidechains, including Ile48 in close proximity to the bound bromide ion (Figure 1C).

Figure 1 with 2 supplements see all
An anion-binding site in the Fluc channel vestibule.

(a) Structure of Fluc-Ec2 with Br-. Monomers are shown in maize and blue, with fluoride ions as pink spheres, sodium as a gray sphere, and anomalous difference map shown as an orange mesh, contoured at 5σ. Zoomed-in views depict Br- as orange spheres, with the aqueous vestibule indicated by a blue mesh and vestibule arginines shown as sticks. (b) Comparison of vestibule anion-binding site for Fluc-Ec2 (top) and Fluc-Bpe (bottom; PDB: 5NKQ). For Fluc-Ec2, the Br- anomalous difference map is displayed as an orange mesh and contoured at 5σ. For Fluc-Bpe, the Fo-Fc map is displayed in green and contoured at 3σ. 2Fo-Fc electron density is shown for sidechains and displayed as a gray mesh, contoured at 2σ. (c) Additional views of the Br--binding site in Fluc-Ec2, with Ile48, Ser81, and Thr82 shown as sticks. (d) Electrical recordings for multichannel bilayers of Fluc-Ec2 I48C and wild-type (WT) Fluc-Ec2. Dashed line indicates the zero-current level. Saturating (2-sulfonatoethyl)methanethiosulfonate (MTSES) was added at the indicated time. Regions of the recording with electrical noise from mixing are colored light gray to assist with figure interpretation. Traces are representative of data collected from five independent bilayers. Right panel, normalized current after MTSES addition. Replicates from two independent preps are shown in black or white. Average current change for Ec2 I48C upon MTSES addition (mean ± SEM from five bilayers): 56 ± 3% decrease. Current change for Ec2 WT upon MTSES addition (mean ± SEM from five bilayers): 0.7 ± 1.7% increase.

Figure 1—source data 1

Measurements of current decrease upon MTSES addition.

https://cdn.elifesciences.org/articles/69482/elife-69482-fig1-data1-v2.xlsx
Figure 1—source data 2

Fluoride efflux measurements for Bpe-I50W.

https://cdn.elifesciences.org/articles/69482/elife-69482-fig1-data2-v2.xlsx

In order to test whether this anion-binding site is part of the fluoride permeation pathway, we introduced an I48C mutation to Fluc-Ec2 and assessed the effect of modification by a bulky, anionic, thiol-reactive reagent, (2-sulfonatoethyl)methanethiosulfonate (MTSES), on fluoride conduction. We performed these experiments on a C74A background, which behaves like the wild-type (WT) protein in F- efflux assays (Figure 1—figure supplement 1). A second cysteine in the native Ec2 sequence, C16, cannot be altered without destabilizing the protein. However, this residue is buried at the interface of helices 1 and 1’, and does not react with thiol reagents in the folded protein. In planar lipid bilayers, I48C mediates robust fluoride currents, which rapidly diminish by ~50% upon addition of saturating MTSES to the cis chamber (Figure 1D), consistent with full modification of a cis-facing thiol in a population of channels with oppositely oriented pores. In contrast, MTSES addition to channels with WT I48 (C74A background) does not alter the fluoride currents (Figure 1D). We sought to recreate the MTSES block experiment in Fluc-Bpe channels, but we did not observe efficient labeling of a cysteine introduced at the corresponding position, Ile50. However, mutation of Ile50 in Bpe to a bulkier tryptophan sidechain diminished the rate of fluoride transport by ~400-fold in liposome efflux experiments, possibly by sterically hindering fluoride access to the bottom of the vestibule (Figure 1—figure supplement 2).

In order to probe the role of the anion-coordinating sidechains in more detail, we mutated Fluc-Ec2’s bromide-coordinating Ser81 to alanine, threonine, or cysteine, and also constructed a S81A/T82A double mutant. For all four mutants, we measured fluoride channel activity using either single-channel electrophysiology or bulk liposome efflux experiments (Figure 2A and B), and we solved x-ray crystal structures of the mutants together with Br- (Figure 2C). No other halides or pseudohalides were present in crystallization solutions.

Figure 2 with 2 supplements see all
Mutagenesis of vestibule anion-binding site.

(a) Fluoride efflux from liposomes monitored with a fluoride-selective electrode: wild-type (WT) Fluc-Ec2 (gray), S81A (green), S81A/T82A (blue), S81T (red), and S81C (purple). Efflux initiated by the addition of valinomycin (black triangle). After reaching steady state, the remaining encapsulated fluoride was released by detergent addition (open triangles). Each trace is normalized against total encapsulated fluoride. Traces are representative of results from at least two independent biochemical purifications. Results from all replicates are tabulated in Table 5. (b) Representative single-channel recording of S81A in the presence of a blocking monobody to identify the zero-current level (dashed line). (c) Bromine anomalous difference maps for S81A, S81A/T82A, S81T, and S81C contoured at 5σ. The frame around each panel is colored as in panel (a). (d) Comparison of the position of Br- density in S81C (orange sphere) and WT Ec2 (dashed orange circle). (e) Fluoride currents mediated by Ec2-S81C, WT Ec2, and Bpe-S83C channels. pH was adjusted during the experiment as indicated. Regions of the recording with electrical noise from mixing are colored light gray to assist with figure interpretation. Traces are representative of recordings from three to six independent bilayers. Additional replicate traces can be found in Figure 2—figure supplement 1. (f) Summary of all replicates of experiments shown in panel (e). Values are normalized against the maximum steady-state current (5 s average) measured at pH 5.5 for that trace. Black and white points indicate different protein preparations.

The functional experiments showed that fluoride throughput is inhibited in these mutants. S81A had a mild effect, with a ~50% decrease in conductance to 3.7 ± 0.4 pS (seven single-channel measurements), compared to 7 pS for the WT protein (Stockbridge et al., 2013Figure 2B). The S81A/T82A double mutant had a more severe effect, with fluoride throughput diminished to 8530 ± 30 s−1, a ~100-fold reduction in the rate (Figure 2A). In accord with the fluoride transport experiments, a strong anomalous peak persisted in the S81A structure but was weaker in the S81A/T82A double mutant (Figure 2C). In both cases, the densities shifted away from the channel center toward the external solution, moving about 2 Å closer to the vestibule Arg22s.

The S81C and S81T phenotypes were more extreme: for both mutants, fluoride efflux from liposomes was completely abolished (Figure 2A). From the structural data, it is not readily apparent why S81T does not transport fluoride. A bromide density is observed in a similar position, coordinated by the threonine’s hydroxyl, and with similar intensity as wild-type, and the surrounding residues are not perturbed by this mutation.

In contrast, the structure of S81C provides a possible explanation for the lack of fluoride transport observed in the liposome flux assays. An anomalous density is present in the vestibule, but has moved ~2 Å farther up into the aqueous vestibule, relative to the Br- position in the WT protein (Figure 2D). We posit that the electropositive environment of the vestibule perturbs the cysteine pKa such that it is deprotonated at the pH of these experiments (pH 9 in the crystal structure and pH 7.5 in the liposome flux experiments). The pKa prediction software PropKa reinforces this possibility, calculating an approximate pKa value of 6 for S81C in the crystal structure of this mutant (Bas et al., 2008). To test this idea explicitly, we monitored currents mediated by Ec2 S81C in planar lipid bilayers as a function of changing pH. Whereas fluoride currents were near zero at pH 7.4, currents increased dramatically when the pH was decreased to 5.5 (Figure 2E, Figure 2F, Figure 2—figure supplement 1). The increase in F- currents was fully reversible with pH, and WT activity was not altered by changing pH over this range. The analogous mutation in Fluc-Bpe channels, S83C, exhibits similar pH sensitivity (Figure 2E, Figure 2F, Figure 2—figure supplement 1).

Taken together, these experiments show that the anion-binding site at the bottom of the vestibule is on the fluoride permeation pathway . This anion-binding site is located immediately adjacent to one of the fluoride ions in the polar track, and we imagine fluoride ions enter the vestibule, become dehydrated, before eventually being stripped of water entirely as the ion is translocated from the bottom of the vestibule to the polar track. Translocation between the vestibule and the polar track must contribute to anion selectivity since the bromide anomalous density is observed in the former location, but never in the latter. However, we could not detect any change in chloride transport by these mutants (Figure 2—figure supplement 2), motivating us to search for additional pore-lining sidechains on the opposite end of the pore.

A trio of sidechains defines the opposite end of the pore

To identify additional pore-lining sequences, we began by analyzing the sequences of the eukaryotic relatives of the homodimeric bacterial Flucs, known as Fluoride Export proteins (Li et al., 2013). Whereas the bacterial Flucs assemble as dual-topology homodimers with a pair of symmetry-related pores, the eukaryotic fluoride channels are expressed as a two-domain single polypeptide with a linker helix that enforces antiparallel topology of the domains (Smith et al., 2015). In the FEX proteins, this ancient fusion event has permitted drift of redundant sequences, including degradation of one of the two pores (Berbasova et al., 2017). A clear pattern has been identified in which residues that line one pore (mostly, but not entirely, from the C-terminal domain) are highly conserved, whereas the corresponding residues from the second, vestigial pore (mostly, but not entirely, from the N-terminal domain) have drifted (Stockbridge et al., 2015; Smith et al., 2015; Berbasova et al., 2017). We reasoned that the other amino acids that follow this pattern of conservation and degradation might be expected to also contribute to the pore.

We selected representative eukaryotic FEX proteins from yeasts and plants, and aligned the N- and C-terminal domains with the sequence of Fluc-Bpe in order to identify residues that follow eukaryotic pore conservation patterns (Figure 3A). We chose Fluc-Bpe for this analysis rather than Fluc-Ec2, because Fluc-Bpe has higher sequence homology to the eukaryotic FEX domains. We identified three additional residues that follow the same pattern of conservation as other pore-lining sequences: a threonine, a tyrosine, and a glutamate (blue highlighting in alignment). In the Fluc-Bpe structure, the three homologous sidechains (Thr39, Glu88, and Tyr104) associate within hydrogen-bonding distance of each other, one contributed by each pore-lining helix, TM2, TM3, and TM4. They are positioned near the protein’s aqueous vestibules, and Tyr104 is also within hydrogen-bonding distance of a fluoride ion within the pore (Figure 3B).

Figure 3 with 3 supplements see all
Identification and characterization of triad residues.

(a) Sequence alignment of Fluc-Bpe with N- and C-terminal domains of representative eukaryotic fluoride channels (transmembrane helices only). Invariant pore-lining residues are shown in yellow. Pore-lining residues that are conserved in only one pore of the eukaryotic FEX channels are highlighted in blue. Residue numbering from Fluc-Bpe is shown (note that S83 in Fluc-Bpe is equivalent to S81 in Fluc-Ec2). (b) Structure of Fluc-Bpe (PDB: 5NKQ) with triad residues indicated as sticks, aqueous vestibule as a mesh, and fluoride ions as pink spheres. (c) Single-channel currents for wild-type (WT) Fluc-Bpe and indicated mutants measured at a holding voltage of 200 mV. Error bars represent the mean and SEM. Black and white points indicate different protein preparations. (d) Representative single-channel electrophysiological recordings for WT Fluc-Bpe, Bpe T39S, and Bpe-Y104F.

These residues are well-conserved among Flucs more generally (Macdonald and Stockbridge, 2017). From an alignment of all homodimeric Fluc sequences in the PFAM database (Mistry et al., 2021), we found that Thr39 is conserved in ~95% of the sequences we studied, Glu88 is conserved in >85% of sequences (~10% Asp and ~ 5% Gln), and Tyr104 is conserved in 55% of sequences (~35% Asn and ~15% Ser). The strong conservation of these residues across multiple kingdoms, the asymmetric distribution among eukaryotic domains that is consistent with other pore-lining sequences, and their close spatial relationship with one another motivated further functional analysis of this molecular triad.

T39 and Y104 proved sensitive to mutagenesis, and only conservative mutations were permitted at these positions. Using bulk liposome efflux assays as a binary measurement of F- transport, we found that we obtained transport-competent mutants when Thr39 was replaced by Ser, but not Val, Asn, Ala, or Cys (Figure 3—figure supplement 1). When Tyr104 was replaced by Phe, robust fluoride efflux activity was observed, but mutants with Ser, His, or Ile in this position all had anemic fluxes in the range of 100 ions/s (Figure 3—figure supplement 1). Glu88 was somewhat more permissive: Ala, Asp, and Gln were all tolerated, but not Lys (Figure 3—figure supplement 1). To experimentally probe whether Glu88 is in the anionic carboxylate form or protonated at pH 7, we performed bilayer experiments in which we recorded currents at pH 7 and then raised the pH in a stepwise fashion to 8.7. We observed reduced fluoride currents as pH was increased, but the difference in these effects between channels bearing Glu and Gln at position 88 was minimal (Figure 3—figure supplement 2). Since changing the protonation state of an acidic sidechain along the permeation pathway would be expected to have substantial ramifications for fluoride currents, these experiments suggest that the protonation state of Glu88 does not change as the pH is increased from 7 to 8.7 and, therefore, that the pKa of E88 falls below ~6.5 or above ~9. A pKa perturbation of a glutamate to >9 would be quite unusual, and we therefore argue that it is more likely that Glu88 is not protonated at physiological pH. In agreement with this interpretation, Propka calculates an approximate pKa for Glu88 of 5.7 (Bas et al., 2008).

Those triad mutants that permitted fluoride transport in efflux assays were also assessed using single-channel electrophysiology (Figure 3C,D). T39S, E88D, and Y104F retained F- conductance to at least 75% of WT levels, and we do not interpret these differences as mechanistically important. In contrast, E88Q exhibited currents one-fifth of the WT levels, a more substantial difference that is also statistically significant at p<0.0001 (unpaired t-test). T39S and Y104F both showed differences in dynamic behavior compared to WT Fluc-Bpe proteins, which are constitutively open and show no closures or sub-conductance states. T39S undergoes long periods of robust throughput (τo = 9.2 ± 0.2 s), punctuated by brief channel closures (τc = 35.3 ± 0.4 ms) (Figure 3D). Y104F was more dynamic, with shorter open intervals (τo = 1.9 ± 0.2 s and τc = 33.2 ± 2.5 ms) (Figure 3D, inset). Thus, single-channel recordings suggest that one role of the triad is to stabilize the three pore-lining helices in an open, fluoride-conducting conformation. Upon addition of channel-binding monobodies (Stockbridge et al., 2014; Turman and Stockbridge, 2017), familiar current block is observed, indicating that despite the increased conformational flexibility, the structure of the channel is not perturbed to a significant extent (Figure 3—figure supplement 3).

Anion recognition at the triad

None of the fluoride-conductive mutants constructed thus far transport chloride ion, as probed using our most sensitive metric of chloride transport, liposome efflux assays (Figure 4—figure supplement 1). However, we noticed that halides and pseudohalides inhibit fluoride currents with a wide range of potencies (Figure 4A, Table 2). The recognition series (OCN->SCN->NO3->Cl-) deviates from common determinants of anion selectivity, such as anion radius, ΔGhydration, ΔGBorn, or the lyotropic (Hofmeister) series (Table 3, Figure 4—figure supplement 2). In these titrations, full inhibition of the fluoride currents is not achieved. The inhibitory effects are best fit by a two-site-binding isotherm, with weak binding to a second site (Table 2). Because the Fluc channel possesses a pair of antiparallel pores, the observed behavior might reflect anion interactions at both the vestibule and triad sides of the channel. In order to separate the effects of anion block at these two positions, and to better quantify the affinity, we exploited the S83C vestibule mutant described in Figure 2E by recording channels under asymmetric pH conditions. The cis side of the bilayer was maintained at pH 7.5, silencing any pore with a cis-facing vestibular S83C. The trans side of the bilayer was adjusted to pH 5.5, so that pores with a trans-facing vestibular S83C retained WT-like function (Figure 4B).

Figure 4 with 6 supplements see all
Inhibition of Fluc-Bpe and Fluc-Bpe E88Q currents by halides and pseudohalides.

(a) Fraction of blocked current as a function of anion addition. The solid lines represent fits to a two-site-binding isotherm, constrained so that the maximum PB for each site is 0.5. In this model, anions bind to single sites that are located on opposite sides of the dual-topology pores. Ki values for fits are reported in Table 2. Data are collected from three independent bilayers. Where present, error bars represent SEM of independent replicates. (b) Cartoon of strategy for orienting Bpe channels for anion block experiments. Gray area indicates aqueous vestibules. Sidechains E88 and S83C are shown as sticks. (c) Representative electrical recording showing OCN- addition to fluoride currents mediated by oriented Bpe-S83C channels. The zero-current level is indicated with a dashed line. Cyanate additions are indicated by the arrows. Regions of the recording with electrical noise from cyanate addition and mixing are colored light gray to assist with figure interpretation. (d) Lineweaver-Burke analysis of OCN- block as a function of F- concentration. Dashed lines represent linear fits to the data. All measurements were performed in triplicate from independent bilayers; where not visible, error bars are smaller than the diameter of the point. (e, f) Fraction of blocked current in S83C (e) or S83C/E88Q (f) oriented channels as a function of anion addition. Points and error bars represent the mean and SEM from three independent bilayers. Where not visible, error bars are smaller than the diameter of the point. Solid lines represent fits to a single-site-binding isotherm with PB,max = 1. Ki values from fits are reported in Table 2. Comparison of replicate measurements from independent preps are shown in Figure 4—figure supplement 3.

Table 2
Fit parameters for anion block experiments.
Dual-topology channelsOriented channels
WT/OCN-WT/SCN-WT/NO3-WT/Cl-WT/OCN-WT/Cl-E88Q/OCN-E88Q/Cl-
Ki,16.8 mM9.0 mM45 mM137 mM7.9 mM480 mM48.9 mM213 mM
Bmax10.50.50.50.51.01.01.01.0
Ki,2100 mM190 mM530 mM1.1 M--------
Bmax20.50.50.50.5--------
Dual-topology channels
E88Q/SCN-E88Q/Cl-
Ki,1398 mM542 mM
Bmax10.50.5
Ki,21.2 M5.8 M
Bmax20.50.5
Table 3
Fluc-Bpe inhibition and physical properties of halides and pseudohalides.
Ki,1 (mM)Ki,2 (mM)Ki (oriented system, mM)Radius (Å)pKaΔGhyd (kcal/mol)ΔGBorn (kcal/mol)Log KCl-X*
F-------1.333.2−112−114−1.5
Cl-13711004801.81−7−83−860
NO3-45530--1.99−1.3−73−721.9
SCN-9.0190--2.491−69−633.23
OCN-6.81077.92.163.7−89−720.82
  1. *Relative anion partition coefficient between water and PVC membrane, a measurement that reflects the lyotropic (Hofmeister) series, described in Smith et al., 1999.

With this oriented system, we tested the effect of OCN- and Cl- addition to the cis (pH 7.5) side of the bilayer, isolating anion interactions at the side of the pore defined by the T-E-Y triad. In OCN- titration experiments, currents were reduced almost to the zero-current level at 30 mM OCN-, showing that the higher affinity OCN--binding site is on the triad side of the pore (Figure 4C). Using the oriented system, we performed OCN- addition experiments in the presence of 30–300 mM F-. The apparent affinity of OCN- increased as F- concentration decreased, showing that binding and inhibition at the triad site is competitive with fluoride (Figure 4D). For both OCN- and Cl-, block of the fluoride currents was well approximated by a single-site-binding isotherm that saturates at full inhibition, although we did not perform experiments at the ~molar Cl- concentrations that would be required to fully block currents (Figure 4E, Figure 4—figure supplement 3, Table 2). In contrast, fits to the data with the two-site-binding model used for the dual-topology WT channels were poor. Under our usual experimental conditions with 300 mM F-, a fit to a single-site-binding isotherm yielded Ki values of ~400 mM for chloride, the most abundant biological halide, and 8 mM for OCN-, in very good agreement with the value estimated from the dual-topology WT channels (Table 2). Although OCN- blocked Fluc-Bpe with relatively high affinity, liposome flux experiments showed that OCN- is not permeant (Figure 4—figure supplement 4).

It is notable that one of the participants in the triad, E88, is itself an anion. In order to understand the interplay between the E88 carboxylate and the blocking anions, we mutated E88 to glutamine on the S83C background and measured fluoride current inhibition by Cl- and OCN-. The appreciable ~60-fold difference in Cl- and OCN- block characteristic of WT channels is almost eliminated for E88Q channels, which display only ~fourfold difference in Cl- and OCN- affinity (Figure 4F, Figure 4—figure supplement 5, Table 2). This effect is almost entirely due to the 10-fold less potent block of E88Q by OCN-. Qualitatively similar results were obtained for SCN- block of randomly oriented WT and E88Q channels (Figure 4—figure supplement 6). Even if we are cautious in quantifying the effect because Cl- block cannot be measured to saturation, a qualitative reading of these experiments suggests that Glu88 contributes to anion recognition at the end of the pore defined by the T-E-Y triad.

Discussion

The vestibule end of the pore

In this work, we fused electrophysiology, X-ray crystallography, and liposome flux assays to identify the routes by which fluoride ions access the previously identified fluoride-binding sites along the polar track of Fluc homologs Fluc-Bpe and Fluc-Ec2. One anion-binding site, identified by the anomalous diffraction of Br- in the Fluc-Ec2 homolog, is located at the bottom of the electropositive vestibule and is sensitive to mutagenesis as well as modification of a nearby sidechain with the bulky thiol-reactive anion MTSES. Moreover, conversion of a serine from this anion-binding site to a cysteine introduces a strong pH-dependence to the fluoride channel activity, demonstrating that this position comprises part of the permeation pathway. Ion accumulation in aqueous entryways is a well-characterized feature of many ion channels, serving to increase the rate at which ions process to the constricted selectivity filter (Doyle et al., 1998; Latorre and Miller, 1983; Payandeh et al., 2011).

We speculate that the vestibule serine (S81 in Fluc-Ec2/S83 in Fluc-Bpe), which is absolutely invariant in Fluc channels, plays a central role in fluoride access to the dehydrated polar track. It is worth noting that a rotamerization of the vestibule serine would bring this sidechain within hydrogen-bonding distance of one such polar track fluoride position, F1 (Figure 5, right panel). A mechanism involving translocation of fluoride ions by rotamerization of amino acid sidechains lining the pore has been proposed for the Fluc channels previously and would be consistent with the measured conductance of these proteins (Stockbridge et al., 2015; Last et al., 2017). Since threonine enjoys less conformational flexibility than serine, such a mechanism might explain why S81T is non-functional in Fluc-Ec2 and why the Ser to Thr substitution has not arisen over evolutionary time in any Fluc channel. The hydrogen bond between the fluoride and the vestibule serine seems to be dispensable, and mutant channels with an alanine at the position retain robust fluoride currents. Similarly, conversion of polar track residues to alanine also had mild consequences for Fluc-Ec2 (Last et al., 2017). We note that, in experiments to monitor fluoride currents, especially single channels, saturating fluoride concentrations and high potentials are required due to the channels’ relatively low conductance. We speculate that these mutants might have more drastic consequences at the low mM fluoride concentrations typical in the biological context.

Figure 5 with 1 supplement see all
Proposed multi-ion permeation mechanism for Fluc-Bpe.

For clarity, only one pore is shown. Cartoon structure is shown in transparent gray, aqueous vestibules are shown as pale cyan surfaces, and residues that have been shown to contribute to the pore (this work and references Stockbridge et al., 2015; Last et al., 2016; McIlwain et al., 2020; Last et al., 2017) are shown as yellow sticks. The five pore-lining residues identified in this work are labeled. Asterisks indicate that the rotamer shown is hypothetical and has not been observed crystallographically. Occupied fluoride ion sites are shown in pink, unoccupied fluoride-binding sites are shown as dashed circles, and the proposed movement of ions between binding sites is indicated by arrows.

The T-E-Y triad end of the pore

Based on sequence analysis and site-directed mutagenesis, we have also identified the opposite end of the pore, which, in Fluc-Bpe, is defined by a hydrogen-bonded trio of conserved sidechains, T39, Y104, and E88, contributed by each of the three pore-lining helices. We propose that, in the resting state of the channel, the E88 carboxylate resides in the position observed in the crystal structures (Figure 5, right panel) (in structures, this position is additionally enforced by monobody binding). E88 is stabilized in this position by the positive dipole of helix 3b and hydrogen bond donors T39 and Y104, where it helps compensate the otherwise positive electrostatics of the unoccupied channel. We suggest that when F- is present, the permeant anion electrostatically repels the E88 carboxylate, perhaps competing for the same binding site at the top of helix 3b (Figure 5, left panel).

Other anions are also able to compete for this site in the permeation pathway, competitively inhibiting fluoride currents when bound. We observed that, for a series of halides and pseudohalides, the selectivity series is correlated to the pKa of the conjugate acid (Table 3, Figure 4—figure supplement 2); we propose that pKa is actually a proxy for the anion’s strength as a hydrogen bond acceptor (basicity). Although pKa and basicity are not strictly correlated across anion types, the properties are relatively well correlated within a single anionic series, such as the halide/pseudohalide series tested here (Pike et al., 2017; Gilli et al., 2009). Thus, we suggest that an anion’s propensity to serve as a hydrogen bond acceptor contributes to its recognition by the Flucs, helping to explain the channel’s remarkable indifference to Cl-, the fluoride ion’s most biologically relevant competitor. In contrast to Cl-, and like OCN-, F- is a famously strong hydrogen bond acceptor.

Proposed mechanism of fluoride permeation

For all of Fluc’s idiosyncrasies, we propose a mechanism with much in common with other well-characterized ion channels (Figure 5). The negative charge of the fluoride ions is counterbalanced by the protein’s few positive charges, the vestibule arginines and the structural central Na+. Experiments have shown that both pores are functional for F- permeation (Last et al., 2016), but it seems highly unlikely that all six anion positions (three anions in each of two pores) are simultaneously occupied. Rather, we imagine a scenario of alternating occupancy, as proposed for other multi-ion pores, in which a fluoride moving into one binding site electrostatically hastens its neighbor into the next position in the sequence. We propose either that the densities observed in the crystal structure represent partially occupied fluoride sites or that the monobodies used as crystallization chaperones alter the electrostatic landscape in the pore, increasing ion occupancy. Indeed, in crystal structures of Fluc-Bpe with a monobody occupying only one side of the channel, each pore contained only one fluoride density, rather than two, in the polar track (McIlwain et al., 2018).

In Figure 5, the starting configuration (left panel) shows an F- bound in the site identified by anomalous scattering, at the bottom of the vestibule, labeled F0. We propose that as additional fluoride ions enter the electropositive vestibule, the fluoride ion at F0 is electrostatically repelled, providing energy for desolvation and translocation into the narrowest part of the channel at position F1 (Figure 5, right panel). But the F1 binding site is not pre-assembled: rotamerization of the vestibule serine (S83 in Fluc-Bpe), which is possible with serine but not threonine, accompanies the lateral movement of the anion. Other sidechains have also been proposed to adopt new rotameric conformations in order to ligand the anion at F1, including N43 (Stockbridge et al., 2015) and S84 (Last et al., 2017). Thus, we propose that the F1 binding site is assembled simultaneously with its occupation by fluoride. The rotamerization of channel sidechains to accompany ions through the pore has been proposed for other channels as well, including the Orai and voltage-gated calcium channels (Hou et al., 2020; Sather and McCleskey, 2003).

We imagine that this configuration is short-lived: a new fluoride ion settles into the deep vestibule F0 site, the fluoride at F1 moves farther down the channel to F2, and the S83 sidechain returns to its position facing the vestibule. The binding site at F2 is in close proximity to the E88 carboxylate; the electrostatic conflict could be resolved if E88 swings out into solution, allowing the fluoride at F2 to exit the channel, having now traversed the bilayer (whereby E88 could then resume its position at the pore exit without conflict). We have shown that the E88Q mutant reduces both fluoride currents and block of fluoride currents by OCN-. We propose that both behaviors arise because the mutant sidechain, which does not bear a negative charge, is not easily dislodged from the binding site via electrostatic conflicts with the permeant fluoride or the cyanate blocker.

This proposed mechanism introduces several previously unrecognized amino acids involved in fluoride permeation and extends the pathway to the aqueous solutions on both sides of the bilayer. It also explains the evolutionary conservation and physiological consequences of mutation described for conserved sidechains, including the invariant serine (position S81 in Ec2 or S83 in Bpe) and triad glutamate (position E88 in Bpe) (Smith et al., 2015; Berbasova et al., 2017). Also, while these experiments provide the first hints of a molecular mechanism for anion recognition by the Flucs, they also emphasize how robust the channel’s anion selectivity is. Despite dozens of point mutations to two homologs, alone and in combination (summarized in Figure 5—figure supplement 1 and Table 4), no mutant that permits the permeation of any other anion has been reported yet. It may be that there is no unique selectivity filter but that several regions of the channel work together to achieve selectivity, so that abolishing anion selectivity requires destruction of the channel itself. Alternatively, channel selectivity might be achieved by matching the number of available ligands in the pore to the preferred coordination number of the anion, as has been proposed for K+ channels (Bostick and Brooks, 2007; Bostick and Brooks, 2009). F- is a superlative in this regard, requiring fewer ligands than any other anion. If this is the case, relaxing the selectivity might require adding coordinating ligands along the pore, which would be difficult to accomplish with site-directed mutagenesis alone. Indeed, even accounting for the addition of coordinating ligands via sidechain rotamerization, the F1 and F2 sites have relatively small coordination numbers (~four including the phenylalanine ring edges). Chloride, in contrast, prefers at least six ligands in its coordination sphere (Bostick and Brooks, 2009; Ohtaki and Radnai, 1993; Cametti and Rissanen, 2009; Merchant and Asthagiri, 2009).

Table 4
Compiled results of anion transport experiments for Fluc-Bpe and Fluc-Ec2.

Results from Fluc-Ec2 are shown in italics, with numbering according to Fluc-Bpe for reference to the structure in Figure 5—figure supplement 1.

ReferenceMutant (no F- permeation)Mutant
(F- permeation retained, no Cl- permeation)
Stockbridge et al., 2015F82I, F85I, N43D
Last et al., 2016F82I, F85I
Last et al., 2017F82Y, F82S, F82A, F82L, F82I, F82T, F85Y, F85S, F85AS112A, T116V, T116I, S83A, F82M
McIlwain et al., 2020N43S, R22K
This workS83T, S83C, T39A, T39V, T39C, T39N, E88K, Y104S, Y014H, Y104W, Y104IS83A, S83A/S84A, Y104F, T39S, E88Q, E88D, E88A

As a rare example of an anion channel required to select against the biologically dominant anion, the Fluc channels present an excellent case study of biochemical anion recognition. But the Fluc channel’s stringent anion recognition, as quantified here, is physiologically essential, too. In electrophysiology experiments, in the presence of saturating 300 mM F-, the apparent Ki values for block by Cl- and other anions are correspondingly low. But in the bacterial cytoplasm, during an F- challenge, with F- ion between 100 μM and 10 mM (Ji et al., 2014), and Cl- ion between 10 and 100 mM (Schultz et al., 1962), even a small increase in the inhibitory effects of Cl- would represent a serious challenge to the efficacy of these channels and the survival of the bacteria.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (Bordetella pertussis)Fluc-BpeNCBIWP_003818609.1Bears mutation R28K to increase yield
(PMID:26344196)
Gene (Escherichia coli virulence plasmid)Fluc-Ec2NCBIWP_001318207.1Bears mutation R25K to increase yield (PMID:26344196). For cysteine modification experiments, C74A (this paper — see Figure 1—figure supplement 1).
Recombinant DNA reagentFluc-Bpe in pET21a (plasmid)PMID:26344196Expression vector for Fluc-Bpe. Available upon request.
Recombinant DNA reagentFluc-Ec2 in pET21a (plasmid)PMID:26344196Expression vector for Fluc-Ec2. Available upon request.
Chemical compound, drugIsethionic acidWako Chemicals, Richmond VA107-36-8
Chemical compound, drugMTSESToronto Research ChemicalsS672000
Chemical compound, drugE. coli polar lipidsAvanti, Alabaster, AL#100600C
Chemical compound, drugn-decyl-β -D-maltopyranosideAnatrace, Maumee, OHD322
OtherMonobodies S9 and S12PMID:25290819Purified from E. coli according to the protocol described in the reference.
PMID:25290819

Chemicals and reagents

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Potassium isethionate was prepared from isethionic acid (Wako Chemicals, Richmond, VA). Detergents were from Anatrace and lipids from Avanti Polar Lipids. MTSES ((2-sulfonatoethyl)methanethiosulfonate) was from Toronto Research Chemicals.

Protein expression, purification, and reconstitution

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Mutant channels were constructed using standard molecular biology techniques and verified by sequencing. All constructs bore functionally neutral mutations, R25K (Fluc-Ec2) or R28K (Fluc-Bpe), which increase protein yield (Stockbridge et al., 2015). Constructs that introduced a cysteine (Ec2-I48C and Ec2-S81C) also bore the mutation C74A. WT Fluc-Bpe is cysteine-free. Histidine-tagged Fluc-Bpe and Fluc-Ec2 were expressed in E. coli and purified via cobalt affinity chromatography according to published protocols (Stockbridge et al., 2014; Stockbridge et al., 2015; McIlwain et al., 2020). The buffer for the final size-exclusion step was 100 mM NaBr, 10 mM 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES), pH 7, for crystallography applications, or 100 mM NaCl, 10 mM HEPES, pH 7, for functional reconstitution. For reconstitution, proteins were mixed with detergent-solubilized E. coli polar lipids (Avanti Polar Lipids; 10 mg/ml) at a ratio of 0.1 μg protein/mg lipid for single-channel bilayer electrophysiology, 0.2 μg protein/mg lipid for liposome flux experiments, or 5 μg protein/mg lipid for macroscopic bilayer experiments. The protein/detergent/lipid mixture was dialyzed for 36 hr (6 l buffer per 50 mg lipid over three buffer changes). Proteoliposomes were stored at −80°C until use, at which point the suspension was freeze-thawed three times and extruded 21 times through a 400 nm filter to form liposomes.

X-ray crystallography

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After purification, monobody S9 and Fluc-Ec2 were mixed in a 1:1 molar ratio as described in Stockbridge et al., 2015. For Ec2-S81C, the protein mixture was used to set up sitting drop vapor diffusion crystal trays with a 1:1 mixture of protein solution and mother liquor. Crystals were formed in either 0.1 M glycine, pH 8.7–9.2, 31–36% polyethylene glycol (PEG) 600 or 0.1 M ammonium sulfate, 0.1 M N-(2-acetamido)iminoacetate (ADA), pH 6–6.5, 31–36% PEG 600 over 3–7 days and were frozen in liquid nitrogen prior to data collection at 13.5 keV at the Life Sciences Collaborative Access Team beamline 21-ID-D at the Advanced Photon Source, Argonne National Laboratory. Phases were calculated by molecular replacement with Phaser (McCoy et al., 2007) using Fluc-Ec2 and the monobody S9 as search models (pdb:5A43), followed by refinement with Refmac (Murshudov et al., 2011) and Phenix (Liebschner et al., 2019) and model building in real space with Coot (Emsley et al., 2010).

Planar lipid bilayer electrophysiology

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Experiments were performed as described previously (Stockbridge et al., 2013). Electrophysiological recordings were acquired at a holding voltage of −200 mV, electronically filtered at 1 kHz during acquisition, and digitally filtered to 500 Hz for analysis. Solutions in the cis and trans chambers varied as described in the text. Typical solutions contained 300 mM NaF with 10 mM 3-morpholinopropane-1-sulfonic acid (MOPS), pH 7. For MTSES and anion block experiments, the sodium salt of each anion was prepared as a concentrated solution in 300 mM NaF and 10 mM MOPS, pH 7, and added to the cis chamber with thorough manual mixing. The final MTSES concentration was 1 mM. For experiments in which the pH was varied, recording buffers additionally contained 10 mM 2-(N-morpholino)ethanesulfonic acid (MES, for pH 5.5 experiments) or 10 mM glycine (for pH 9 experiments). A pre-determined aliquot of dilute isethionic acid or NaOH was added to adjust the pH in the cis chamber, and the final pH value was confirmed after each experiment. Because hydrofluoric acid has a pKa of 3.4 and is extremely hazardous, we avoided lowering the pH of fluoride solutions below 5.5. Macroscopic bilayer recordings shown are representative of three to seven independent bilayer experiments, and single-channel experiments are from 9 to 17 independent channel fusions for each mutant. All constructs used for electrophysiology experiments were purified from at least two independent protein preparations, and no prep-to-prep variation was observed.

Fluoride efflux from liposomes

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Fluoride efflux from liposomes was monitored using a fluoride-selective electrode as described previously (Brammer et al., 2014). Intraliposomal solution contained 300 mM KF, 10 mM Na+ isethionate, 10 mM HEPES-KOH, pH 7. The external solution was exchanged by passing liposomes over a Sephadex G-50 spin column equilibrated in 300 mM K+ isethionate, 10 mM Na isethionate, 10 mM HEPES-KOH, pH 7. Proteoliposomes were diluted 20-fold in matching buffer and fluoride efflux initiated by addition of 1 μM valinomycin. At the end of the experiment, remaining encapsulated fluoride was released from the liposomes by addition of 50 mM n-octyl-β-D-glucoside. Fluoride efflux was normalized against total encapsulated fluoride. In most cases, the result of this assay was binary: either the mutant had no activity relative to background leak (<100 ions/s) or the rate of fluoride efflux exceeded the response time of the electrode (>104 ions/sec). Efflux experiments were performed three to six independent times, with replicates derived from at least two independent protein preparations. In all cases of a binary result (no activity or >104 ions/sec), all replicates were in agreement (Table 5). Light-scattering experiments (Figure 4—figure supplement 3) were performed as previously described (Stockbridge et al., 2012). Proteoliposomes containing 300 mM KF, KCl, or KOCN and 10 mM HEPES, pH 7, were diluted in assay buffer (300 mM K+ isethionate, 10 HEPES, pH 7). 90° light scattering was monitored at 550 nm upon addition of valinomycin (0.1 μg/ml final concentration).

Table 5
Liposome efflux experiments: compiled results from all replicates.
ConstructAnionFigureRate (ions/s): Prep 1Rate (ions/s): Prep 2Mean ± SEM
Ec2 WTF-1-S1>104, >104>104, >104>104
Ec2 C74AF-1-S1>104, >104>104, >104>104
Ec2 WTF-2a>104, >104>104, >104>104
Ec2 S81AF-2a>104, >104, >104>104, >104, >104>104
Ec2 S81TF-2a<100, <100<100, <100<100
Ec2 S81CF-2a<100, <100<100, <100<100
Ec2 S81A/S82AF-2a8860, 64009640, 7840, 88608320 ± 560 ions/sec
Ec2 S81A/S82ACl-2-S2<50, <50<50, <50<50
Bpe S83A/T84ACl-2-S2<50, <50<50, <50<50
Bpe T39VF-3-S1<100, <100<100, <100<100
Bpe T39SF-3-S1>104, >104, >104>104, >104>104
Bpe T39CF-3-S1<100, <100, <100<100, <100<100
Bpe T39AF-3-S1<100, <100<100, <100<100
Bpe T39NF-3-S1<100<100, <100<100
Bpe E88AF-3-S1>104, >104>104, >104, >104>104
Bpe E88QF-3-S1>104, >104>104, >104, >104>104
Bpe E88DF-3-S1>104, >104>104, >104, >104>104
Bpe E88KF-3-S1<100<100, <100<100
Bpe Y104FF-3-S1>104, >104, >104>104, >104>104
Bpe Y104SF-3-S1<100, <100, <100<100, <100<100
Bpe Y104HF-3-S1<100, <100<100, <100<100
Bpe Y104IF-3-S1<100, <100<100<100
Bpe I50WF-1-S1600, 720, 960650, 550, 720700 ± 60 ions/sec
Bpe Y104FCl-4-S1<50, <50<50, <50<50
Bpe T39SCl-4-S1<50, <50<50, <50<50
Bpe E88QCl-4-S1<50, <50<50, <50<50
Bpe E88ACl-4-S1<50, <50<50, <50<50

Data availability

Atomic coordinates for the Fluc-Ec2 and mutants in the presence of Br- have been deposited in the Protein Data Bank under accession numbers 7KKR (WT); 7KKA (S81A); 7KKB (S81C); 7KK8 (S81T); 7KK9 (S81A/T81A). Source data files have been provided for all figures. No custom code was used.

The following data sets were generated
    1. McIlwain BC
    2. Stockbridge RB
    (2021) RCSB Protein Data Bank
    ID 7KKR. Fluoride channel Fluc-Ec2 wild-type with bromide.
    1. McIlwain BC
    2. Stockbridge RB
    (2021) RCSB Protein Data Bank
    ID 7KKA. Fluoride channel Fluc-Ec2 mutant S81A with bromide.
    1. McIlwain BC
    2. Stockbridge RB
    (2021) RCSB Protein Data Bank
    ID 7KKB. Fluoride channel Fluc-Ec2 mutant S81C with bromide.
    1. McIlwain BC
    2. Stockbridge RB
    (2021) RCSB Protein Data Bank
    ID 7KK8. Fluoride channel Fluc-Ec2 mutant S81T with bromide.
    1. McIlwain BC
    2. Stockbridge RB
    (2021) RCSB Protein Data Bank
    ID 7KK9. Fluoride channel Fluc-Ec2 mutant S81A/T82A with bromide.

References

    1. Emsley P
    2. Lohkamp B
    3. Scott WG
    4. Cowtan K
    (2010) Features and development of Coot
    Acta Crystallographica Section D Biological Crystallography 66:486–501.
    https://doi.org/10.1107/S0907444910007493

Decision letter

  1. Merritt Maduke
    Reviewing Editor; Stanford University School of Medicine, United States
  2. Kenton J Swartz
    Senior Editor; National Institute of Neurological Disorders and Stroke, National Institutes of Health, United States
  3. Merritt Maduke
    Reviewer; Stanford University School of Medicine, United States
  4. Rachelle Gaudet
    Reviewer; Harvard University, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This manuscript will be of interest to scientists in physiology and membrane biophysics. Using functional tests guided by new structures that reveal ion-binding sites, the authors propose an elegant permeation mechanism that helps explain the unusually high fluoride selectivity of the microbial "Fluc" ion channels.

Decision letter after peer review:

Thank you for submitting your article "The fluoride permeation pathway and anion recognition in Fluc family fluoride channels" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, including Merritt Maduke as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Kenton Swartz as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Rachelle Gaudet (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

1) The electrophysiology experiments with MTS reagents and the I48C mutation are lacking a control experiment with a positively charged MTS reagent such as MTSET. For the experiment to confirm that the MTS is inhibiting through the electropositive vestibule, it should be shown that a positively charged MTS reagent either doesn't inhibit or inhibits with significantly slower kinetics.

2) Related to the previous point, Figure 1d shows only "before and after" from a single

experiment with current before MTS addition, and a lower current after, with the actual reaction kinetics not shown. We suspect this part of the trace may have been removed due to stirring artifacts, though this is not stated. The interpretation of the result will be much more compelling if you could show the kinetics of the MTS reagent. This should be possible using a lower concentration of MTS reagent (in the μM range, prepared just before use). We note that Figure 2e shows a reasonable timecourse when changing pH.

3) The summary data for experiments of Figure 1d would be best shown in the Figure (perhaps in a bar graph) and not just in the legend.

4) In the Methods section, it is stated that all electrophysiology experiments were performed on at least two independent protein preparations. However, the data as presented do not reveal the prep-to-prep variability. We recommend that the summary data (Figure 1d, Supp Figure 1, Supp Figure 2, etc etc) include tabulated data that indicate the results specifically for different preparations.

5) For Supplementary Figure 2, we would prefer to see the actual data, and not just single summary points, as the current traces are wobbly and therefore it is not immediately clear how one picks a value for each pH condition. For Figure 2e, a bar graph or some other method of presenting the summary data should be included.

6) For the experiments in Figure 2a, summary data should be shown in addition to the representative data.

7) A minor shortcoming regards the effects of pH on the mutations of Glu88 (p6, bottom). It is noted that the difference between Glu and Gln at position 88 are minimal, yet the lack of pH sensitivity between pH 7 and 8.7 is interpreted as reflecting a deprotonated Glu88. We would suggest instead that the glutamate is likely protonated, leading to the lack of difference between Glu and Gln, but that it's pKa is sufficiently shifted that it remains nearly fully protonated through the tested pH range. Such shifts in pKa are not unusual, especially in membrane proteins.

8) Which of the changes in Figure 3 are statistically significant?

9) The methods mention that the "Ec2-S81C construct also bore a mutation C74A", but this is not mentioned elsewhere and not data are presented to evaluate the function of the C74A background. The authors should provide information about how mutation of this residue, which contacts the bound Br-, does or does not impact ion channel function.

10) The pH dependence of the S81C and S83C mutations are used to propose that the cysteine is likely to be readily deprotonated. It could strengthen your arguments to generate predicted pKa values based on their structures. (The same suggestion applies to the arguments about E88.) One way to do this is to use PROPKA.

11) In the validation reports for Ec2-S81T, one of the Br- ion (residue # 2) has a very high B-factor and poor fit to the map (RSCC and RSR). This prompted a closer look at the models and maps. Looking at the Ec2-S81T map, it seems like Br- ion 2 is not positioned at the peak of density, and would likely benefit from more careful refinement, especially because it is a crucial part of this model. There is also a very strong unmodeled spherical density (>5 σ in the 2Fo-Fc map and >10 σ in the Fo-Fc map for chain C) right next to S81 of each monobody. (Ec2-S81C and WTBr have similar unmodeled blobs.)

Figures:

1) The arrows and arrowheads in all figures should be carefully checked to make sure that they line up with the actual time of addition (valinomycin and detergent) and buffer switches.

2) Figure 2d: using purple for the Br- ion is very confusing with the F- ions almost the same color. They should be consistently represented in the same orange color as in the other figures in the manuscript.

3) Figure 3a: It is unclear how residues were chosen for yellow highlighting, as there are plenty of other residues in the figure that seem just as or more conserved as the highlighted residues. Also, it would be helpful if residue numbers for the Bpe sequence were included in the figure. Finally, the "blue" highlight is dark enough that the black letters are difficult to make out.

4) Figure 3d: can the region of the recordings corresponding to the insets be indicated in the corresponding top traces?

5) Supp Figure 12: A line pointing to F82 would be useful.

Reviewers Combined Review:

This manuscript describes experiments aimed at understanding the ion-permeation pathway in the unusual and interesting Fluc family of microbial fluoride-selective ion channels. Previously, the channel pathway had been postulated based on locations of fluoride ions seen in crystal structures, but the tightly packed nature of the protein precluded certainty on this point. Here, the authors present four new crystal structures of the E. coli Fluc channel in the presence of the anomalous scattering ion, bromide, providing provide strong evidence for previously uncharacterized ion binding site one of the channel entry vestibules.

The authors follow up this structural evidence with mutagenesis of nearby residues. They test the role of a residue adjacent to the density by mutating it to Cysteine and reacting with a negatively charged MTS reagent, which reduces ion conduction as measured electrophysiologically. Other mutations in that and nearby residues also reduce ion conduction. Particularly compelling results are that the low ion conduction can be overcome for the E. coli S81C (and B. pertussis S83C) variants by lowering the pH, suggesting that the channel permeation pathway is very sensitive to local changes at that site. To identify potential anion entryways at the other end of the channel, the authors perform a sequence alignment of relatively distant family members and find several candidate residues whose roles in conductance are analyzed using a set of elegant selectivity and block assays.

Overall, the work presented here is very nice and the interpretations are solidly based on the data. However, some aspects of the experiments need to be clarified and extended. The experiments with MTS reagents are missing a critical control; characterization of the C74A background mutation is missing; and some experimental details of reproducibility and are not fully reported.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "The fluoride permeation pathway and anion recognition in Fluc family fluoride channels" for further consideration by eLife. Your revised article has been reviewed by 2 peer reviewers and the evaluation has been overseen by Kenton Swartz as the Senior Editor, and a Reviewing Editor.

The Reviewers agree that you have substantially improved the manuscript, especially with respect to reproducibility and the presentation of the data in most cases. The Reviewers are not completely satisfied with your argument against the need for the MTSET control: since there are two other cysteines in the protein, the control should be performed. Nevertheless, given that WT-like construct is not affected by MTS, which provides some reasonable support for the idea that the introduced cysteine is indeed the target, the additional control is not required. Similarly, though the reviewers agree that the MTS experiment would have been better with lower concentrations (so that the reaction is not over by the end of the stirring), such experiments are not essential to your conclusions and therefore will not be required. The inclusion of the currents during stirring is helpful, as is the coloring scheme to deemphasize that part of the data. Since the records are now continuous, we request that you remove the "break" marks in the figure.

https://doi.org/10.7554/eLife.69482.sa1

Author response

Essential revisions:

1) The electrophysiology experiments with MTS reagents and the I48C mutation are lacking a control experiment with a positively charged MTS reagent such as MTSET. For the experiment to confirm that the MTS is inhibiting through the electropositive vestibule, it should be shown that a positively charged MTS reagent either doesn't inhibit or inhibits with significantly slower kinetics.

We do not agree with the reviewers that MTSET modification is an essential control. In this construct, there is a single modifiable cysteine, I48. When the cysteine is present, currents are blocked by MTSES addition. When the cysteine is removed, currents are completely insensitive to MTSES addition. We have a high-resolution structure that shows that residue 48 is located in the vestibule, and that the vestibule is electropositive. We can’t imagine any other interpretation of the data except that MTSES inhibits by modifying the cysteine introduced at position 48 in the vestibule. It’s a reasonable prediction that MTSET would have slower kinetics of modification, but we do not think this experiment would serve as a control for any plausible alternative possibility. In addition, the interpretation that fluoride ions enter the Fluc channels through the vestibule is bolstered by additional lines of evidence, including crystallography data, I50W mutant in the Bpe homologue, mutagenesis experiments, and S81C pH sensitivity experiments.

2) Related to the previous point, Figure 1d shows only "before and after" from a single

experiment with current before MTS addition, and a lower current after, with the actual reaction kinetics not shown. We suspect this part of the trace may have been removed due to stirring artifacts, though this is not stated. The interpretation of the result will be much more compelling if you could show the kinetics of the MTS reagent. This should be possible using a lower concentration of MTS reagent (in the μM range, prepared just before use). We note that Figure 2e shows a reasonable timecourse when changing pH.

We have updated our figures throughout the manuscript so that the electrical noise during stirring is shown in a light gray color. The kinetic response after MTSES addition is visible through the noise.

3) The summary data for experiments of Figure 1d would be best shown in the Figure (perhaps in a bar graph) and not just in the legend.

We have added a bar graph to Figure 1d.

4) In the Methods section, it is stated that all electrophysiology experiments were performed on at least two independent protein preparations. However, the data as presented do not reveal the prep-to-prep variability. We recommend that the summary data (Figure 1d, Supp Figure 1, Supp Figure 2, etc etc) include tabulated data that indicate the results specifically for different preparations.

We have color-coded the points in the bar graphs reported in Figure 1d, 2e, and 3c to indicate which measurements come from which biochemical prep. For liposome efflux experiments of point mutants, we thought that the most efficient way to convey this information was in a Table summarizing each result. We have added a new table (Table 5) that summarizes all replicates, including information about biochemical prep, for Figures 2a, 2b, 1-S1, 1-S2, 2-S2, 3-S1, 4-S1. For the experiments reported in Figure 4, we have included a supplementary figure (4-S3) that shows titrations of both the E88Q and WT constructs with Cl- and OCN- with datapoints color-coded by biochemical prep. For the traces shown in 2-S1, the prep is indicated on the graph. Source data is also color coded according to biochemical prep.

None of the constructs reported in this manuscript show prep-to-prep variability.

5) For Supplementary Figure 2, we would prefer to see the actual data, and not just single summary points, as the current traces are wobbly and therefore it is not immediately clear how one picks a value for each pH condition. For Figure 2e, a bar graph or some other method of presenting the summary data should be included.

We have included full traces replicates of this experiment in 2-S1 and added a panel with single summary points to Figure 2e.

6) For the experiments in Figure 2a, summary data should be shown in addition to the representative data.

For most of the mutants, the fluoride transport rate is either above or below the limit of detection (>104 ions per second and <100 ions/second, respectively). This is described in the Methods. We have updated this text slightly to include the lower limit of measurement:

“Fluoride efflux was normalized against total encapsulated fluoride. In most cases, the result of this assay is binary: either the mutant has no activity relative to background leak (<100 ions/s) or the rate of fluoride efflux exceeds the response time of the electrode (>104 ions/sec). In all cases of a binary result (no activity or >104 ions/sec) all replicates were in agreement.”

Of 27 different constructs tested in fluoride or chloride efflux experiments, 25 are either above or below the limit of detection. For these constructs, there is no way to calculate a mean or standard error for replicates – in other words, no way to summarize. (Ec2-S81A/S82A and Bpe-I50W are exceptions that fall within the dynamic range of the electrode.) The representative traces shown in Figure 2a (and 1-S1, 1-S2, 2-S2, 3-S1, 4-S1) are sufficient to convey this qualitative point (does a particular mutant have F- channel activity, yes or no?). We have added table in the supplement (Table 5) that shows the individual results from the replicates for each mutant. These are denoted “>104” for measurements that exceed the response time of the electrode, “<100” for measurements below the limit of detection, or, for S81A and I50W, individual measurements of the rate, and are broken down by protein prep.

7) A minor shortcoming regards the effects of pH on the mutations of Glu88 (p6, bottom). It is noted that the difference between Glu and Gln at position 88 are minimal, yet the lack of pH sensitivity between pH 7 and 8.7 is interpreted as reflecting a deprotonated Glu88. We would suggest instead that the glutamate is likely protonated, leading to the lack of difference between Glu and Gln, but that it's pKa is sufficiently shifted that it remains nearly fully protonated through the tested pH range. Such shifts in pKa are not unusual, especially in membrane proteins.

We agree that shifted pKa values are not unusual. However, a glutamate with a pKa shifted by almost 6 units to >9 would be fairly unusual. We performed the propKa analysis as suggested in point #10, which predicts a pKa for this glutamate of 5.7.

We would also like to point out that the single channel currents for E88 and E88Q are quite different (Figure 3C) – a difference we argue is both statistically and mechanistically significant. What doesn’t differ between E88 and E88Q is the modulation of those currents by pH. We interpret this to mean that E88 (like E88Q) is not protonatable between pH 7 and 9. Of the two possibilities (a pKa above 9, or a pKa below 7), the pKa below 7 is the most likely. The propka analysis nicely backs up that inference.

We have updated the text around E88 to read:

“Since changing the protonation state of an acidic sidechain along the permeation pathway would be expected to have substantial ramifications for fluoride currents, these experiments suggest that the protonation state of Glu88 does not change as the pH is increased from 7 to 8.7, and therefore that the pKa of E88 falls below ~6.5 or above ~9. […] In agreement with this interpretation, Propka calculates an approximate pKa for Glu88 of 5.7 [12].”

8) Which of the changes in Figure 3 are statistically significant?

Although we have recorded enough single channels that the 10-15% differences in conductance between WT, Y104F, T39S, and E88D are statistically significant, we do not wish to make the argument that these differences are mechanistically important.

In the text, we do argue that the difference between WT and E88Q is mechanistically significant, and, accordingly, the difference in single channel conductance is statistically significant with p<0.0001.

We have updated the text as follows:

“T39S, E88D, Y104F retained F- conductance at least 75% of WT levels, and we do not interpret these differences as mechanistically important. In contrast, E88Q exhibited currents one fifth of the wildtype levels, a more substantial difference that is also statistically significant at p<.0001 (unpaired t-test).”

9) The methods mention that the "Ec2-S81C construct also bore a mutation C74A", but this is not mentioned elsewhere and not data are presented to evaluate the function of the C74A background. The authors should provide information about how mutation of this residue, which contacts the bound Br-, does or does not impact ion channel function.

Thank you for pointing out our oversight in describing this construct. We use the C74A background as a “minimal cys” construct for all Ec2 cysteine labelling experiments. (The construct is not truly cysless because C16 cannot be altered without destabilizing the protein. However, C16 is buried at the dimer interface and does not react with thiol reagents.) In F- efflux experiments, it behaves like WT. We have added data to show this (Figure 1-S1 and Table 5). This same minimal-cys construct was used as the background for both the I48C MTSES addition experiments and the S81C experiments.

We have added language to the describe this construct when we introduce the I48C MTSES addition experiments, where we use it as the background for the “WT” control:

“We performed these experiments on a C74A background, which has fluoride transport properties similar to the WT protein. […] However, this residue is buried at the interface of helices 1 and 1’, and does not react with thiol reagents in the folded protein.”

We do not think that the structural evidence supports the suggestion that C74 contacts the Br-. Especially in our higher resolution structures, it is evident from the electron density that the C74 sidechain is pointed away from the Br- and packed against helix 2. As an example, electron density from the S81T structure is shown in Author response image 1. The closest atom of C74 is 4.4 Å from the Br-, and the SH group is 5.5 Å from the Br-. Even accounting for the possibility that C74 could adopt a different rotamer outside the crystal, none of the probable rotamers bring it within coordination distance of the Br-.

Author response image 1

10) The pH dependence of the S81C and S83C mutations are used to propose that the cysteine is likely to be readily deprotonated. It could strengthen your arguments to generate predicted pKa values based on their structures. (The same suggestion applies to the arguments about E88.) One way to do this is to use PROPKA.

Thank you for this suggestion. We have updated the text around S81C to read:

“We posit that the electropositive environment of the vestibule perturbs the cysteine pKa such that it is deprotonated at the pH of these experiments (pH 9 in the crystal structure, and pH 7.5 in the liposome flux experiments). The pKa prediction software PropKa reinforces this possibility, calculating an approximate pKa value of ~6 for S81C in the crystal structure of this mutant [12]. To test this idea explicitly…”

We have updated the text around E88 to read:

“Since changing the protonation state of an acidic sidechain along the permeation pathway would be expected to have substantial ramifications for fluoride currents, these experiments suggest that the protonation state of Glu88 does not change as the pH is increased from 7 to 8.7, and therefore that the pKa of E88 falls below ~6.5 or above ~9. […] In agreement with this interpretation, Propka calculates an approximate pKa for Glu88 of 5.7 [12].”

11) In the validation reports for Ec2-S81T, one of the Br- ion (residue # 2) has a very high B-factor and poor fit to the map (RSCC and RSR). This prompted a closer look at the models and maps. Looking at the Ec2-S81T map, it seems like Br- ion 2 is not positioned at the peak of density, and would likely benefit from more careful refinement, especially because it is a crucial part of this model. There is also a very strong unmodeled spherical density (>5 σ in the 2Fo-Fc map and >10 σ in the Fo-Fc map for chain C) right next to S81 of each monobody. (Ec2-S81C and WTBr have similar unmodeled blobs.)

We have re-refined Ec2-S81T and improved the fit to the density. A new PDB validation report is attached.

We are aware of the unmodelled blob near the monobody in the maps. We re-sequenced the monobody to ensure that the residue at position 81 is indeed a serine, and it is. The unmodelled blob is more electron dense than a water, and it is not a bromide. It may be a sulfate anion from the crystallization buffer. However, since this blob is associated with the monobody rather than the channel, is well removed from the bromide-binding site in the vestibule, and because it is difficult to assign it with certainty in these 2.5-3 Å-resolution structures, we have elected not to assign the density.

Figures:

1) The arrows and arrowheads in all figures should be carefully checked to make sure that they line up with the actual time of addition (valinomycin and detergent) and buffer switches.

2) Figure 2d: using purple for the Br- ion is very confusing with the F- ions almost the same color. They should be consistently represented in the same orange color as in the other figures in the manuscript.

3) Figure 3a: It is unclear how residues were chosen for yellow highlighting, as there are plenty of other residues in the figure that seem just as or more conserved as the highlighted residues. Also, it would be helpful if residue numbers for the Bpe sequence were included in the figure. Finally, the "blue" highlight is dark enough that the black letters are difficult to make out.

4) Figure 3d: can the region of the recordings corresponding to the insets be indicated in the corresponding top traces?

5) Supp Figure 12: A line pointing to F82 would be useful.

We have made all of the requested changes to the figures. With regards to point #3, we have clarified the color-coding in the legend:

“Invariant pore-lining residues are shown in yellow. Pore-lining residues that are conserved in only one pore of the eukaryotic FEX channels are highlighted in blue.”

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The Reviewers agree that you have substantially improved the manuscript, especially with respect to reproducibility and the presentation of the data in most cases. The Reviewers are not completely satisfied with your argument against the need for the MTSET control: since there are two other cysteines in the protein, the control should be performed. Nevertheless, given that WT-like construct is not affected by MTS, which provides some reasonable support for the idea that the introduced cysteine is indeed the target, the additional control is not required.

We thank the editors for recognizing that the proposed experiment is not essential. Although we appreciate the suggestion from the reviewers, we would like to further elaborate on our rationale for not performing the MTSET modification experiment. We hope that this explanation is satisfactory.

For reasons described below, we think that (1) the control that we have already performed accomplishes the goal of distinguishing between the two potential cysteines and (2) the outcome of the MTSET experiment would be difficult to interpret and would be less likely to distinguish between the cysteines.

For the trace in the left panel of Figure 1d (labelled Ec2-I48C), the protein has cysteines at positions 16 (buried in the structure) and I48C. For the trace in the right panel (labelled Ec2-WT), the protein has cysteines at position 16 only (as described in the text, the C16 sidechains are buried and cannot be removed without destabilizing the protein).

The structure-guided introduction of a cysteine at position I48 provides a strong prediction that the MTS-mediated inhibition occurs through this site in the vestibule. However, we absolutely agree with the reviewers that, by itself, the electrophysiological experiment with the I48C construct (left panel) does not establish that MTSES reacts with the cysteine at position I48C. Modification of the other cysteine, C16, is an unlikely but valid possibility that must be controlled for. We can also imagine that MTSES interacts with the protein as a non-specific anionic blocker, blocking currents without forming any covalent linkage.

The experiment shown in the right panel of Figure 1d controls for both of these possibilities. If the current reduction in the left panel was due to modification of C16, then we would see a similar inhibitory effect in the right panel, since C16 is present in both constructs. We don’t see this. The same logic also rules out the possibility that MTSES blocks non-specifically. Any other interpretation would need to invoke the idea that the Ec2-WT and Ec2-I48C constructs adopt substantially different conformations. We think this possibility is quite unlikely, since the I48C’s biochemical and functional properties resemble WT, and extensive structural and electrophysiological experiments targeted at identifying conformational changes for the Flucs have uniformly suggested that these constitutively open channels do not undergo significant structural rearrangements (Turman and Stockbridge, 2017, JGP 149: 511-522; McIlwain, Newstead and Stockbridge, 2018, Structure 26: 635-639).

The reviewers propose that modification with MTSET would provide an additional control to distinguish between modification at C16 and I48C. Although this experiment might provide additional evidence beyond the structure that the modified cysteine is in an electropositive environment, we do not think that this experiment would be a superior control to confirm that MTS-mediated inhibition occurs at I48C. Both residues are located near the center of the protein, and the environment is likely to be electropositive for both (see Author response image 2). Thus, changing the electrostatic properties of the MTS reagent would be unlikely to distinguish between modification of C16 and I48C in a clearly interpretable way. Although we thank the reviewers for the suggestion, and we would be eager to perform any experiment that would rule out likely alternative explanations for the results in Figure 1, we do not think that the proposed MTSET modification experiment would do this.

Thank you for considering our work.

Author response image 2
The structure shown here is of the WT channel.

The Br- ions are shown as brick red spheres. Positions that are cysteines in the MTS experiments are colored magenta (I48C and C16). The features that contribute to the electropositive character of the channel are the central sodium ion (purple sphere) and R22 (cyan). These positive charges are in close proximity to both I48C and C16, and are likely to electrostatically influence both positions.

Similarly, though the reviewers agree that the MTS experiment would have been better with lower concentrations (so that the reaction is not over by the end of the stirring), such experiments are not essential to your conclusions and therefore will not be required. The inclusion of the currents during stirring is helpful, as is the coloring scheme to deemphasize that part of the data. Since the records are now continuous, we request that you remove the "break" marks in the figure.

We’ve removed the “break” marks in Figure 1 and 2. Thanks for catching this.

https://doi.org/10.7554/eLife.69482.sa2

Article and author information

Author details

  1. Benjamin C McIlwain

    Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, United States
    Contribution
    Conceptualization, Formal analysis, Investigation, Visualization, Methodology, Writing - original draft
    Competing interests
    No competing interests declared
  2. Roja Gundepudi

    Program in Biophysics, University of Michigan, Ann Arbor, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  3. B Ben Koff

    Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  4. Randy B Stockbridge

    1. Department of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann Arbor, United States
    2. Program in Biophysics, University of Michigan, Ann Arbor, United States
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    stockbr@umich.edu
    Competing interests
    Reviewing editor, eLife
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8848-3032

Funding

National Institutes of Health (R35-GM128768)

  • Randy B Stockbridge

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank the LS-CAT beamline staff for technical assistance, Christian Macdonald for assistance with sequence analysis, and members of the Stockbridge lab for comments on the manuscript and project. We are grateful to José Faraldo-Gómez and Robyn Stix (NIH/NHLBI) for insightful conversations about channel electrostatics.

Senior Editor

  1. Kenton J Swartz, National Institute of Neurological Disorders and Stroke, National Institutes of Health, United States

Reviewing Editor

  1. Merritt Maduke, Stanford University School of Medicine, United States

Reviewers

  1. Merritt Maduke, Stanford University School of Medicine, United States
  2. Rachelle Gaudet, Harvard University, United States

Publication history

  1. Preprint posted: April 4, 2021 (view preprint)
  2. Received: April 16, 2021
  3. Accepted: July 9, 2021
  4. Accepted Manuscript published: July 12, 2021 (version 1)
  5. Version of Record published: July 27, 2021 (version 2)

Copyright

© 2021, McIlwain et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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