Abstract
Bestrophin (BEST1–4 in humans) channels are ligand gated chloride (Cl−) channels that are activated by calcium (Ca2+). Mutations in BEST1 cause retinal degenerative diseases. Partly because these channels have no sequence or structural similarity to other ion channels, the molecular mechanisms underlying gating are unknown. Here, we present a series of cryo-electron microscopy (cryo-EM) structures of chicken BEST1, determined at 3.1 Å resolution or better, that represent the principal gating states of the channel. Unlike other channels, opening of the pore is due to the repositioning of tethered pore-lining helices within a surrounding protein shell that dramatically widens a “neck” of the pore through a concertina of amino acid rearrangements within the protein core. The neck serves as both the activation and the inactivation gate. The binding of Ca2+ to a cytosolic domain instigates pore opening and the structures reveal that, unlike voltage-gated Na+ and K+ channels, similar molecular rearrangements are responsible for inactivation and deactivation. A single aperture within the 95 Å-long opened pore separates the cytosol from the extracellular milieu and controls anion permeability. The studies define the basis for Ca2+-activated Cl− channel function and reveal a new molecular paradigm for gating in ligand-gated ion channels.
The family of bestrophin proteins (BEST1–4) was identified by linkage analysis to hereditary macular degenerations caused by mutations in BEST1 (1, 2); to date more than 200 mutations in BEST1 are linked with eye disease (3, 4). BEST1–4 proteins are expressed in the plasma membrane and form Ca2+-activated Cl− channels by assembling as pentamers (5–9). Data suggest that BEST1 mediates a Ca2+-activated Cl− current that is integral to human retinal pigment epithelial function (10). The broad tissue distribution of the family suggests additional physiological functions that are not fully realized (5), and these may include processes as diverse as cell volume regulation (11), pH homeostasis (12), and neurotransmitter release (13). An X-ray structure of chicken BEST1, which shares 74 % sequence identity with human BEST1 and possesses analogous Ca2+-activation and anion-selectivity properties, revealed an architecture that is distinct from other ion channel families (6). A prokaryotic ion channel (KpBest) has discernable sequence (14 % identity) and structural homology with BEST1, but the Ca2+-activated Cl− channel function of bestrophin proteins appears specific to metazoan organisms; KpBest is cation-selective and not activated by Ca2+ (14). The pore of BEST1 contains two hydrophobic constrictions, the “neck” and the “aperture”, and ostensibly either of these could function as a gate. Ionic currents through BEST1 have recently been found to decrease over time due to the inactivation of the channel, which is caused by the binding of a C-terminal peptide to a receptor on the channel’s cytosolic surface (15). Because the inactivation peptide is bound to its receptor in the structure and because the antibody that was used as a crystallization chaperone also promotes inactivation, we now realize that the previously determined structure of BEST1 likely represents an inactivated state (6, 15). How the channel opens is not known.
To begin to address the conformational changes associated with channel opening, wedetermined a 3.1 Å resolution cryo-EM structure of BEST1 without an antibody (the same construct used for X-ray studies, comprising residues 1–405, and termed BEST1405). From single particle analysis we obtained a single conformation, which is indistinguishable from the X-ray structure, including a bound inactivation peptide, and therefore this presumably also represents a Ca2+-bound inactivated state (fig. S1, fig. S3; RMSD for Cα atoms =0.5 Å). This result suggests that antibody binding does not distort the channel from a native conformation and that differences we might observe in channel structures are not the result of differences in methodologies between cryo-EM and X-ray crystallography.
In an aim to obtain structural information on an open conformation of the channel, we removed the C-terminal inactivation peptide (by using a construct spanning amino acids 1–345, termed BEST1345) and used this for cryo-EM studies. Importantly, whilst BEST1345 does not inactivate, it possesses normal Cl− selectivity and Ca2+-dependent activation (15). Single particle cryo-EM analysis of Ca2+-bound BEST1345 (fig. S2, fig. S3, fig. S5) revealed two distinct conformations of the channel (Fig. 1). The first, determined at 3.0 Å resolution, represents 86% of the particles and is essentially indistinguishable from the structure of BEST1405 (Fig. 1a, fig. S1c, RMSD for Cα atoms =0.5 Å), except for the absence of the inactivation peptide. Because BEST1345 does not inactivate, the structure presumably represents a Ca2+-bound closed conformation. In this and in the inactivated structure, the neck adopts an indistinguishable closed conformation (Fig. 2c). The second structure, which represents 14% of the particles and is determined to 2.9 Å resolution, contains a dramatically widened pore within the neck (Fig. 1b). Based on discussions presented herein we conclude that this represents the open conformation of the channel. The relative abundance of the closed conformation suggests that it is energetically favorable.
In the closed conformation, the neck is less than 3.5 Å in diameter and approximately 15 Å long; three hydrophobic amino acids on the neck helix (S2b) from each of the channel’s five subunits, I76, F80 and F84, form its walls (Fig. 1a, c, d) (6). Its narrowness, length and hydrophobicity create an energetic barrier to ion permeation that seals the channel shut by hydrophobic block (16). In the open conformation, the neck has dilated to approximately 13 Å in diameter, which is more than sufficient to allow permeation of hydrated Cl− ions (Fig. 1c). No appreciable conformational difference is present in the cytosolic region of the channel, and in particular, the aperture constriction of the pore retains its dimensions.
Comparison of the open and closed conformations of the neck highlights an unusual structural element of the pore that distinguishes the mechanism of gating in BEST1 from most other channels. The neck helix (S2b) is flanked on both of its ends by disruptions of α-helical secondary structure. These disruptions provide impressive flexibility on the one hand and tethering on the other that allow the helix to “float” between closed and open conformations (Fig. 2). In the closed conformation, hydrophobic packing at the center of the neck among the I76, F80 and F84 residues themselves stabilize this conformation. In the open conformation, the tendency of the phenylalanine residues to seclude their hydrophobicity from an aqueous environment is satisfied by their interactions with other hydrophobic amino acids (Y236 and W287) on the S3b and S4a helices located behind the neck helix (Fig. 2 b). The conformational change moves F80 and F84 away from the center of the pore and involves a slight rotation along the helical axis of S2b (~ 10 ° clockwise viewed from the extracellular side), outward displacement of S2b (~ 2.5 Å at F80), a slight expansion of the entire transmembrane region (~ 1 Å increase in radius), and a coordinated set of side chain rotamer changes (Fig. 2a, b and video). Both F80 and F84 move from the most commonly observed rotamer for phenylalanine (observed 44% of the time in the pdb) in the closed conformation to the second most commonly-observed rotamer conformation (observed 33% of the time) in the open conformation (17). By these conformational changes, F80 and F84 rotate away from the axis of the pore by 80° and 105°, respectively (Fig. 2, fig. S5). In a domino effect, side chain rotamer changes of Y236, F282, F283, and W287 allow for the movements of F80 and F84 (Fig. 2a, b and movie S1). The conformational change in I76 is also dramatic. When the channel opens, the first α-helical turn of the neck helix unravels such that I76 packs with F247, F276, L279 and F283 in the open conformation and has shifted by approximately 10 Å (fig. S5a, d). The unraveling is facilitated by P77, which is perfectly conserved among BEST channels and is part of the neck helix in its closed conformation, but marks its N-terminal end in the open conformation (Fig. 2c, d). The repositioning within the neck also exposes S79 and G83 on the neck helix, which are secluded behind the F80 and F84 in the closed conformation, to the pore in the open conformation (Fig. 2a, b and video). Thus, through a concertina of coordinated conformational changes in and around the neck helix, amino acids that formed the hydrophobic barrier that prevented ion permeation have dispersed and reveal a wide aqueous vestibule.
To address how Ca2+ binding contributes to BEST1 channel gating, we determined the cryo-EM structure of BEST1345 in the absence of Ca2+ to 3.0 Å resolution (fig. S6, fig. S7; a cryo-EM structure of BEST1405 without Ca2+ was also determined, at 3.6 Å resolution, and is indistinguishable). The Ca2+-free structure is very similar to the inactivated and Ca2+-bound closed conformations with the only notable differences near the Ca2+ clasp (fig. S8). Of particular note, the neck shares the same closed conformation as in the inactivated and Ca2+- bound closed structures (Fig. 2c). In structures with Ca2+ bound, the five Ca2+ clasps, one from each subunit, resemble a belt that wraps around the midsection of BEST1 (Fig. 1a, b). Without Ca2+, the majority of the Ca2+ clasp becomes disordered (fig. S8b). 3D classification of the Ca2+-free dataset yielded only closed conformations of the neck but did indicate a degree of flexibility in the channel between the transmembrane and cytosolic regions that was manifested as a ~ 5° rotation along the symmetry axis (fig. S8c). This conformational flexibility was not observed in the Ca2+-bound datasets, which suggests that Ca2+ binding rigidifies the channel and we hypothesize that this may be necessary for stabilization of the open conformation.
To investigate how the coupling of the amino acid side chain movements are involved in the transition between the open and closed conformations of the neck and how Ca2+ binding might bias these, we studied the effects of mutating W287 to phenylalanine. W287 is highly conserved among BEST channels. We chose to study this residue because it adopts one side chain rotamer and packs with both F80 and F84 in the open conformation of the neck and another side chain rotamer in the closed conformation that buttresses the space between adjacent S2b helices (Figs. 2a, b and 3d, e), and thus it might govern conformational changes in the neck. We find that the W287F mutation produces channels with dramatically altered gating. Whilst the W287F mutant retained normal Cl− versus K+ selectivity (Fig. 3a), the mutation makes the channel nearly insensitive to Ca2+; approximately 80% of the Cl− current level was maintained when Ca2+ was chelated with EGTA (Fig. 3a, b). To understand the molecular basis of this behavior, we determined cryo-EM structures of the W287F mutant in the presence and absence of Ca2+ at 2.7 Å and 3.0 Å resolutions, respectively (fig. S7, 9). In Ca2+, the channel adopts an open conformation that is essentially indistinguishable from the open conformation observed for BEST1345 (fig. S9, Cα RMSD = 0.2 Å). Unlike BEST1345, 3D classification did not reveal a closed conformation within the cryo-EM dataset, which indicates that essentially all of the particles are in an open conformation and that the open state is preferential for this mutant.
In the absence of Ca2+, in spite of missing density for Ca2+ and a disordered Ca2+-clasp region, the neck also adopts an open conformation (Fig 3c and fig. S9). Thus, in accord with the electrophysiological recordings, the W287F mutation decouples Ca2+ binding from the conformational changes in the activation gate. Modeling of the W287F mutation on a closed conformation of the channel introduces a void behind the neck (fig. S9h), which we hypothesize energetically disfavors the closed conformation. The effects of the relatively conservative mutation of tryptophan to phenylalanine give context to a myriad of disease-causing mutations in and around the neck (fig. S5e).
The structures reveal that the open pore of BEST1 comprises a 90 Å-long water-filled vestibule and a short constriction at the cytosolic aperture (Fig. 1b). The aperture constriction is only 3 Å long (measured where the pore diameter is < 4 Å); its walls are formed solely by the side chains V205 of the five subunits (Fig. 4a). Retinitis pigmentosa can be caused by mutation of the corresponding residue of human BEST1 (I205T mutation)(18), which suggests that the aperture has an important role in channel function. The structures reveal that the aperture has the same conformation in the open and closed states (Fig. 1a, b); accordingly the V205A mutation of chicken BEST1, which would be expected to widen the aperture markedly, has no effect on Ca2+-dependent activation or inactivation (7, 15). We conclude that the aperture does not function as the activation or inactivation gate. However, mutations of V205 have dramatic effects on ion permeability(7) (Fig 4c, d). Both human and chicken BEST1 have a lyotropic permeability sequence in which small anions that are more easily dehydrated than Cl−, such as Br−, I− and SCN−, are more permeable(7, 19–21) (Fig. 4c). We find that mutation of V205 to a smaller or more hydrophilic residue (e.g. glycine, alanine, or serine) abolishes the lyotropic sequence whereas mutation to isoleucine, a bulkier hydrophobic amino acid, makes the permeability differences between Cl−, Br−, I− and SCN− more dramatic (Fig. 4c). Thus, the aperture controls permeability among anions. Based on the narrow diameter of the aperture, anions would shed at least some of their water molecules as they pass through it; this would give rise to the channel’s lyotropic permeability sequence and may contribute to its low single channel conductance (reported at ~2 μS for Cl− for drosophila BEST1(22)). The permeability to large anions such as acetate, propionate and butyrate increases when V205 is substituted by alanine or glycine (Fig. 4d), and thus, as has been suggested previously(7), the aperture functions as a size-selective filter that would tend to prevent permeation of large cellular constituents such as proteins or nucleic acids. Notably, the amino acid sequence at and around the aperture varies among BEST channels (Fig. 4b), and this may endow these channels with distinct permeabilities related to their specific physiological functions. Data suggest that the inhibitory neurotransmitter GABA permeates through BEST1 to underlie a tonic form of synaptic inhibition in glia (13). While this possibility seemed incongruous with the narrowness of the neck observed in the initial structure, the widened neck of the open conformation and the presence of a single constriction that controls permeability make the possibility of slow conductance of GABA and/or other solutes of similar size more plausible. Although the aperture adopts an indistinguishable conformation in all of the structures, we suspect that “breathing” (e.g. thermal motions) of the protein could allow larger ions to move through the aperture than might otherwise fit. It is also conceivable that the binding of cellular ligands near the aperture, as has been suggested for ATP (23), could influence channel behavior by changing its dimensions somewhat. The structure of the open pore hints at a rich diversity of potential physiological functions for BEST channels that are largely unexplored.
The structures presented herein represent the major gating transitions in the channel (Fig. 4e). Unlike voltage-dependent K+ and Na+ channels, in which ions are prevented from flowing by different mechanisms in the inactivated and deactivated states (21, 24), the same closed conformation of the neck is responsible for the deactivated (Ca2+-free) and inactivated states of BEST1. While localized twisting or domain motions often constitute the activation mechanism of ion channels, dramatic molecular choreography within the protein core of BEST1 underlies opening and represents a new paradigm for ion channel gating.
Materials and Methods
Cloning, expression and purification of BEST1
Chicken BEST1 (UniProt E1C3A0) constructs (amino acids 1–405 or 1–345 followed by a Glu-Gly-Glu-Glu-Phe tag) were expressed in Pichia Pastoris as described previously(6). Mutations were made using standard molecular biology techniques.
In preparation for cryo-EM analysis, purification of BEST1 proteins were performed as described previously with modification (6). BEST1 protein was purified by size-exclusion chromatography (SEC; Superose 6 increase 10/300 GL; GE Healthcare) in buffer containing 20 mM Tris, pH 7.5, 50 mM NaCl, 1 mM n-dodecyl-β-D-maltopyranoside (DDM; Anatrace) and 0.1 mM cholesteryl hemisuccinate (CHS; Anatrace). Purified BEST1 was concentrated to 5 mg ml−1 using a 100 kDa concentrator (Amicon Ultra-4, Millipore) and divided into aliquots. For structures with Ca2+, 1 μM CaCl2 was added to the freshly purified protein. For Ca2+-free structures, and 5 mM EGTA, pH 7.5 was added to the freshly purified protein. These samples were immediately used for cryo-EM grid preparation.
EM sample preparation and data acquisition
5 μl of sample was pipetted onto Quantifoil R1.2/R1.3 holy carbon grids (Au 400, Electron Microscopy Sciences), which had been glow discharged for 10 s using a PELCO easiGlow glow discharge cleaning system (Ted Pella). A vitrobot Mark IV cryo-EM sample plunger (FEI) (operated at room temperature with a 1–2 s blotting time under a blot force of 0 and 100% humidity) was used to plunge-freeze the sample into liquid nitrogen-cooled liquid ethane. For Ca2+-free conditions, the blotting paper used for grid freezing was pre-treated with 2 mM EGTA solution (4x), rinsed with ddH2O (4x) and dried under vacuum. Grids were clipped and loaded into a 300 keV Titan Krios microscope (FEI) equipped with a K2 Summit direct electron detector (Gatan). Images were recorded with SerialEM(25) in super-resolution mode at a magnification of 22,500x, which corresponds to a super-resolution pixel size of 0.544 Å, and a defocus range of −0.7 to −2.15 μm. The dose rate was 9 electrons per physical pixel per second, and images were recorded for 10 seconds with 0.25 s subframes (40 total frames), corresponding to a total dose of 76 electrons per Å2.
Image processing
Figures S1, S2, S6 and S9 show the cryo-EM workflow for Ca2+-bound BEST1405, Ca2+-bound BEST1345, Ca2+-free BEST1345 and BEST1345 W287F with and without Ca2+, respectively. Movie stacks were gain-corrected, two-fold Fourier cropped to a calibrated pixel size of 1.088 Å, motion corrected and dose weighted using MotionCor2(26). Contrast Transfer Function (CTF) estimates for motion-corrected micrographs were performed in CTFFIND4 using all frames (27).
Ca2+-bound BEST1405, BEST1345and BEST1345W287F datasets
All subsequent image processing was carried out with RELION2.1(28), using a particle box size of 384 pixels and a spherical mask with a diameter of 140–160 Å. A total of 1740, 1644 and 1597 micrographs were collected for Ca2+-bound BEST1405, BEST1345 and BEST1345 W287F, respectively, and all micrographs were inspected manually; poor quality micrographs and those having CTF estimation fits lower than 5 Å were discarded. Approximately 1000 particles were selected manually for reference-free 2D classification to generate templates that were then used for automatic particle picking. Auto-picking yielded ~312,000, ~309,000 and ~308,000 particles for BEST1405, BEST1345 and BEST1345 W287F, respectively. One round of 2D classification, using 100 classes, was used to remove outlier particles (e.g. ice contaminants), and this yielded ~290,000 particles for BEST1405 and BEST1345 datasets and ~265,000 particles for BEST1345 W287F. 3D refinement, using C5 symmetry, was performed for each dataset using an initial model (generated from a previously collected, lower resolution cryo-EM dataset of Ca2+-free BEST1 using EMAN2(29)) that was low-pass filtered at 60 Å resolution. This yielded consensus reconstructions at 3.1 Å (BEST1405)and 2.9 Å (BEST1345) overall resolutions that have the closed conformation of the neck and 2.8 Å for BEST1345 W287F that an open conformation at the neck. (Refinement using C1 symmetry also yielded reconstructions with 5fold symmetry.) All overall resolution estimates are based on gold-standard Fourier shell correlations (FSC).
To identify the distinct conformational states within the Ca2+-bound BEST1345 dataset, we performed 3D classification using the consensus reconstruction as an initial model (low-pass filtered at 5 Å resolution) and sorting the particles into 9 classes. One class with a widened neck (BEST1345 open) was isolated, containing ~ 30,000 particles. To identify additional open particles from the dataset, this reconstruction was low-pass filtered at 5 Å resolution and used as an initial model for 3D classification (with 4 classes) on the entire dataset. This procedure yielded one class in the open conformation (containing approximately 40,000 particles) and three classes in the Ca2+-bound closed conformation (containing the remainder of the particles). One class for the closed conformation was chosen (containing approximately 44,000 particles) because it contained better-resolved density for the residues lining the neck (I76, F80, F84). 3D Refinement of these two classes yielded reconstructions at 3.0 Å overall resolution. Particles from these two classes were “polished” using aligned movie frames generated from MotionCor2(26). 3D refinement using the polished particles and a global angular sampling threshold of 1.75° yielded final reconstructions at 3.0 Å and 2.9 Å overall resolutions for the Ca2+-bound closed and open reconstructions of BEST1345, respectively. The same polishing strategy for the BEST1345 W287F dataset yielded a final reconstruction of 2.7 Å. Several analogous 3D classification procedures were performed to try to identify an open conformation in the Ca2+-bound BEST1405 dataset but none were found. Conversely, 3D classification approaches with the Ca2+-bound BEST1w287f dataset to identify multiple conformations yielded only reconstructions with an open neck.
Ca2+-free BEST1345and Ca2+-free BEST1345W287F datasets
Initial image processing was carried out with RELION2.1(28), using a particle box size of 384 pixels and a mask diameter of 140. A total of ~1000 or ~1600 micrographs were collected for Ca2+-free BEST1345 and Ca2+-free BEST1345 W287F, respectively, and manually pruned as described for the Ca2+-bound dataset. Auto-picking templates were generated as described and the selected particles (~150,000 for the Ca2+-free BEST1345 and ~185,000 particles for the BEST1345 W287F datasets) were subjected to one round of 2D classification with 100 classes. 3D refinement was performed using the selected particles from 2D classification (~130,000 for the Ca2+-free BEST1345 and ~150,000 particles for the Ca2+-free BEST1345 W287F datasets), C5 symmetry, and the EMAN2-generated initial model. This yielded reconstructions of 3.4 Å and 3.2 Å overall resolutions, respectively. Particle polishing was performed on each dataset and the polished particles were imported into the cisTEM cryo-EM software package for further refinement and classificatio(30) 3D refinement was performed in cisTEM using a mask to apply a 15 Å low-pass filter to the micelle region. This yielded final reconstructions to 3.0 Å overall resolution for both datasets. The ~ 5 ° relative rotation of the cytosolic region with respect to the transmembrane region that was observed under Ca2+-free conditions was identified using 3D classification (using 6 or 8 classes for Ca2+-free BEST1345 and Ca2+-free BEST1345 W287F, respectively) using spatial frequencies up to 6 Å for refinement. Refinement of 3D classes with the most extreme rotation (e.g. approximately ± 2.5 ° rotations relative to the consensus reconstruction) in cisTEM yielded overall resolutions of 3.6 Å (Ca2+-free BEST1345 conformation A, ~11,000 particles), 3.4 Å (Ca2+-free BEST1345 conformation B, ~21,000 particles), 3.4 Å (Ca2+-free BEST1345 W287F conformation A, ~21,000 particles) and 3.5 Å (Ca2+- free BEST1345 W287F conformation B, ~17,000 particles). These reconstructions for Ca2+-free BEST1345 are depicted in Supplementary Figure 8.
RELION2.1(28) was used to estimate of the local resolution all of the final maps. The maps shown in figures are combined maps, were sharpened (using a 5-factor of −50–75 Å2), and low-pass filtered at the final overall resolution of each map.
Model building and refinement
The atomic models were manually built into one of the half-maps (which had been sharpened using a 5-factor of −50–75 Å2 and low-pass filtered at the final overall resolution) using the X-ray structure of BEST1 as a starting point (PDB ID: 4RDQ) and were refined in real space using the COOT software(31). The atomic models were further refined in real space against the same half-map using PHENIX (32). The final models have good stereochemistry and good Fourier shell correlation with the other half-map as well as the combined map (Supplementary Figures 3 and 7). Structural figures were prepared with Pymol (pymol.org), Chimera (33), and HOLE (34).
Liposome reconstitution
SEC-purified protein [in SEC buffer: 150 mM NaCl, 20 mM Tris-HCl, pH7.5, 3 mM n-decyl-β-D-maltoside (DM; Anatrace)] was reconstituted into liposomes. A 3:1 (wt/wt) mixture of POPE (Avanti) and POPG (Avanti) lipids was prepared at 20 mg ml−1 in reconstitution buffer (10 mM Hepes-NaOH, pH 7.6, 450 mM NaCl, 0.2 mM EGTA, 0.19 mM CaCl2). 8% (wt/vol) n-octyl-β-D-maltopyranoside (Anatrace) was added to solubilize the lipids and the mixture was incubated with rotation for 30 min at room temperature. Purified protein was mixed with an equal volume of the solubilized lipids to give a final protein concentration of 0.2–1 mg ml−1 and a lipid concentration of 10 mg ml−1. Proteoliposomes were formed by dialysis (using a 8000 Da molecular mass cutoff) for 1–2 days at 4 °C against 2–4 L of reconstitution buffer and were flash frozen in liquid nitrogen and stored at −80 °C until use.
Electrophysiological recordings
Proteoliposomes were thawed and sonicated for approximately 10 s using an Ultrasonic Cleaner (Laboratory Supplies Company). All data are from recordings made using the Warner planar lipid bilayer workstation (Warner Instruments). Two aqueous chambers (4 mL) were filled with bath solutions. Chlorided silver (Ag/AgCl) wires were used as electrodes, submerged in 3 M KCl, and connected to the bath solutions via agar-KCl salt bridges [2% (wt/vol) agar, 3 M KCl]. The bath solutions were separated by a polystyrene partition with a ~200μM hole across which a bilayer was painted using POPE:POPG in n-decane [3:1 (wt/wt) ratio at 20 mg ml−1]. Proteoliposomes were applied to the bilayer with an osmotic gradient across the bilayer with solutions consisting of: 30 mM KCl or NaCl (cis side) and 10 mM KCl or NaCl (trans side), 20 mM Hepes-NaOH, pH 7.6 and 0.21 mM EGTA/0.19 mM CaCl2 ([Ca2+]free ~ 300 nM) or 1 μM CaCl2. Proteoliposomes were added, 1 μL at a time, to the cis chamber to a preformed bilayer until ionic currents were observed. Solutions were stirred using a stir plate (Warner Instruments stir plate) to aid vesicle fusion. After fusion, the solutions were made symmetric by adding 3M KCl or 5M NaCl, depending on the starting solutions, to the trans side. Unless noted, all reagents were purchased from Sigma-Aldrich. All electrical recordings were taken at room temperature (22–24°C).
Measurements of relative permeabilities among anions were performed as described previously(7). Briefly, after establishing symmetric (30/30 mM KCl or NaCl) conditions, the bath solution in the trans chamber was replaced by perfusion with solutions in which KCl or NaCl was replaced by various potassium salts (Br, I, SCN, acetate, propionate) or sodium salts (butyrate).
Currents were recorded using the Clampex 10.4 program (Axon Instruments) with an Axopatch 200B amplifier (Axon Instruments) and were sampled at 200 μs and filtered at 1 kHz. Data were analyzed using Clampfit 10.4 (Axon Instruments). Graphical display and statistical analyses were carried out using GraphPad Prism 6.0 software. In all cases, currents from bilayers without channels were subtracted. Error bars represent the SEM of at least three separate experiments, each in a separate bilayer. We define the side to which the vesicles are added as the cis side and the opposite trans side as electrical ground, so that transmembrane voltage is reported as Vcis-Vtrans. Ion channels are inserted in both orientations in the bilayer.
Funding
This work was supported, in part, by NIH Grant R01 GM110396 (to S.B.L) and a core facilities support grant to Memorial Sloan Kettering Cancer Center (P30 CA008748).
Author contributions
A.N.M, G.V and S.B.L conceived of and designed the project. A.N.M determined structures of BEST1405, the W287F mutant, and structures in the absence of Ca2+. G.V determined structures of the Ca2+-bound closed state and the Ca2+-bound open state. A.N.M. and G.V performed electrophysiology experiments. All authors contributed to data analysis and the preparation of the manuscript.
Competing interests
The authors declare no competing financial interests.
Data and materials availability
Atomic coordinates and cryo-EM density maps of have been deposited with the PDB and Electron Microscopy Data Bank with the deposition numbers: D_1000236792 (BEST1405, inactivated), D_1000236798 (W287F mutant, Ca2+-free), D_1000236800 (W287F mutant, Ca2+-bound), D_1000236801 (Ca2+-free closed state), D_1000236802 (Ca2+-bound closed state), and D_1000236804 (Ca2+-bound open state). Correspondence and requests for materials should be addressed to S.B.L (Longs{at}mskcc.org).
Acknowledgements
We thank N. Grigorieff, members of his laboratory, and the staff at the Howard Hughes Medical Institute Cryo-EM facility for training and initial advice on cryo-EM. We thank M.J. de la Cruz of the Memorial Sloan Kettering Cancer Center Cryo-EM facility, M. Ebrahim, and the staff of the New York Structural Biology Center Simons Electron Microscopy Center for help with data collection. G.V received funding and mentorship from the Boehringer Ingelheim Fonds Predoctoral Fellowship Program.