ClC-1 protein channels facilitate rapid passage of chloride ions across cellular membranes, thereby orchestrating skeletal muscle excitability. Malfunction of ClC-1 is associated with myotonia congenita, a disease impairing muscle relaxation. Here, we present the cryo-electron microscopy (cryo-EM) structure of human ClC-1, uncovering an architecture reminiscent of that of bovine ClC-K and CLC transporters. The chloride conducting pathway exhibits distinct features, including a central glutamate residue (“fast gate”) known to confer voltage-dependence (a mechanistic feature not present in ClC-K), linked to a somewhat rearranged central tyrosine and a narrower aperture of the pore toward the extracellular vestibule. These characteristics agree with the lower chloride flux of ClC-1 compared with ClC-K and enable us to propose a model for chloride passage in voltage-dependent CLC channels. Comparison of structures derived from protein studied in different experimental conditions supports the notion that pH and adenine nucleotides regulate ClC-1 through interactions between the so-called cystathionine-β-synthase (CBS) domains and the intracellular vestibule (“slow gating”). The structure also provides a framework for analysis of mutations causing myotonia congenita and reveals a striking correlation between mutated residues and the phenotypic effect on voltage gating, opening avenues for rational design of therapies against ClC-1–related diseases.
CLC proteins comprise a large family of chloride (Cl−) -transporting integral membrane proteins with diverse physiological functions [1–3]. The first identified human member, ClC-1, is essential for maintaining the permeability of Cl− across the plasma membrane of skeletal muscle fibers, gCl, accounting for approximately 80% of the resting membrane conductance and assuring precise neuronal control of muscle contraction [3]. Mutations of the ClC-1 gene cause myotonia congenita, a disease that allows a single nerve action potential to trigger a series of muscle action potentials (myotonic runs), leading to prolonged muscle contraction [4–7]. Despite distinct roles as passively conducting Cl− channels and stoichiometrically coupled secondary active Cl−/H+ antiporters [2,3], members of the CLC family share a common homodimeric core architecture, with each subunit harboring an independent ion translocation pathway [8,9]. The molecular mechanisms of ion transport in CLC antiporters have been extensively studied functionally and structurally [8,10–15]. Yet it is poorly understood how the antiporters and channels establish their separate functions. In addition, the complex gating processes that regulate CLC channel activity remain elusive, with only a single available structure of a channel member, namely, that of bovine ClC-K [9]. Each CLC monomer has a gate that operates independently from the other (also known as “protopore” or “fast gate”), structurally attributed to a specific glutamate, “GluGATE” [10]. A slower gate controls both conducting pathways simultaneously (“common” or “slow gate”) [16], but the principles and determinants of this regulation are enigmatic. Furthermore, activity of ClC-1 is modulated by cellular cues such as phosphorylation [17], pH, and nucleotides [18,19] in an unknown manner. Such regulation is, however, physiologically essential because intense muscle exercise leads to acidosis, resulting in an increased nucleotide sensitivity of ClC-1 and consequent reduction of gCl, thereby assisting in preventing muscle fatigue [20,21]. The recent ClC-K structure provided the first insights into the differences between CLC channels and transporters; in particular, it revealed a pore widening on the intracellular side. Yet there are surprisingly few known structural differences between the CLC channels and transporters. However, ClC-K channels exhibit only limited gating as GluGATE is missing [2,3], and their activity has not been reported to depend on nucleotide binding [22]. Therefore key questions concerning CLC channel function and regulation remain unanswered. Furthermore, a deeper understanding on structure–phenotype relationships of myotonia-causing mutations in ClC-1 is required to shed further light on how the muscle disease is manifested at a molecular level.
Here, we have determined structures of full-length human ClC-1 using single-particle cryo-electron microscopy (cryo-EM), exploiting a purified protein sample that displays Cl−-dependent single-channel–derived ion conductance (S1 Fig and S1 Data). For structural characterization, sample in the presence of 100 mM Cl− at pH 7. 5 and in the absence of nucleotides or antibodies was initially employed (Fig 1). Three-dimensional (3D) classification of particles resulted in several different groups, of which one yielded a 3. 6 Å overall resolution density map for the transmembrane domain, allowing confident model building (S2–S4 Figs). The final model represents the membrane-spanning portion (note that the N terminus and intracellular αA helix are lacking) as well as parts of two C terminal’s so-called cystathionine-β-synthase (CBS) domains present per monomer (for which some cryo-EM density is left unmodeled) and includes several features that were not observed in the ClC-K structure (S5 Fig). The homodimeric architecture of ClC-1 is reminiscent of that of bovine ClC-K and available structures of CLC proteins from lower organisms (Fig 2A). The monomers consist of membrane-spanning helices and half-helices (αB to αR) with connecting loops (e. g. , αB–C, between αB and αC) as well as the CBS domains (Fig 1). Each protomer holds a separate chloride conducting pathway across the membrane, established by a vestibule on either side of the membrane, and an interconnecting narrow and short pore. In CLC transporters, the Cl− conducting pore (Fig 2B) is marked by distinct Cl− binding sites (denoted sext, scen, and sint, respectively, but no Cl− ions are resolved in the current structure), and the constricting Glu232 (of αF, also known as GluGATE; ClC-1 numbering throughout) and Tyr578 (of αR, TyrC) [9]. Furthermore, Ser189 (of αC-D, SerC) is located in the vicinity of the pore (Fig 2B, 2C and 2E and S5A Fig). In ClC-1, voltage-dependent gating is established by GluGATE, which is perhaps being displaced by competing Cl− ions and/or protonation. In contrast, in voltage-independent ClC-K channels, GluGATE is replaced by a valine, and, indeed, substitutions of GluGATE with uncharged residues render ClC-1 similarly voltage independent [24]. Unfortunately, the GluGATE side chain is not visible in our cryo-EM density maps (S4 Fig), but carboxylate groups of interacting acidic residues are known to be frequently undetectable using cryo-EM due to radiation damage. A similar orientation of the side chain as observed in ClC-K would be in agreement with Cl− passage through a maintained scen, as a concomitant adaptation of αR significantly shifts the position of TyrC and thus maintains the GluGATE-TyrC distance (Fig 2B–2F and S5G Fig). However, we cannot exclude that the side-chain of GluGATE is buried deep into the hydrophobic pocket established by Phe279, Phe288, and Phe484 (S5H Fig). The pore aperture of the extracellular vestibule is constricted by a hydrophobic barrier with Met485 (Met427 in ClC-K), but in contrast to ClC-K, the gate opening is also controlled by Lys231 (of αE–F) and Arg421 (of αL) (Fig 2B–2F) that may orchestrate Cl− permeation to or from the extracellular environment [25–27]. This difference can be attributed to αE–F, with its GluGATE and Lys231 adopting a more CLC-transporter–like configuration because this loop is considerably shorter than in ClC-K, alongside a side-chain reorientation of Arg421 (Fig 2C–2F). We also observe a structural adjustment on the intracellular side of the pore, with αC–D being displaced as compared to the corresponding loop in ClC-K. This rearrangement opens the vestibule even deeper toward GluGATE (Fig 2C–2F), providing intracellular access beyond the sint site present in CLC antiporters and suggesting that no tight Cl− binding occurs on the intracellular side, in agreement with electrophysiological data [28]. The wider intracellular vestibule of the CLC channels, as compared to the transporters, has been proposed to allow for the higher Cl− conductance in channels, lowering the kinetic barrier between scen and the cytosol [9]. We note that the vestibule width of ClC-1 is similar to that of ClC-K at SerC, with the side chain of this residue being positioned away from the Cl− permeation pathway in both channels, establishing the SerC location as another of the distinguishing features between CLC channels and transporters. It remains obscure whether the channel has been captured in the open configuration, a priori induced by the experimental conditions (0 mV, 100 mM Cl−). Molecular dynamics simulations of the ClC-1 structure suggest that Cl− from the intracellular side spontaneously interacts with GluGATE upon protonation of its side chain but that free energy is required to complete the passage across the membrane (S6 Fig). We anticipate that GluGATE and the Lys231–Arg421 constricting interactions attenuate chloride flux, in agreement with the smaller conductance of ClC-1 versus ClC-K [2,3], and we cannot exclude that Cl− shuttling occurs directly between protonated GluGATE and Lys231 across the Met485 barrier (GluGATE overlays sext in some CLC transporters [8,14,29]); chloride interaction with the latter may be unfavorable, however. The molecular mechanisms that govern slow gating in CLC proteins remain elusive. It is known that CBS nucleotide binding and low pH inhibit ClC-1 activity by favoring closure of the common gate [19,29]. Assessment of the 3 major cryo-EM maps obtained in our structural classification (see also S2 Fig and Methods) reveals different arrangements of the CBS domains, suggesting intrinsic domain flexibility at pH 7. 5 (Fig 3A and 3B and S7 and S8 Figs). To test this, we determined the structure of ClC-1 also at lower pH (6. 2) in the presence of 0. 3 mM of the nucleotide nicotinamide adenine dinucleotide (NAD) to unravel the regulation mechanism (S2, S3 and S8 Figs). In these conditions, the CBS domains appear significantly more rigid (in comparison to pH 7. 5; Fig 3A and 3B and S7 and S8 Figs). This observation is also supported by ClC-1 size-exclusion chromatography profiles (S9 Fig), with samples at low pH being shifted toward lower molecular weight (more compact). Therefore, the CBS arrangements seem to correlate with slow gating, being rigid at low pH in the presence of nucleotides and more flexible at higher pH in the absence of nucleotides, bringing to mind a mechanism that has been proposed based on electrophysiological data [29]. The complete effects of such putative rearrangements are, however, not demonstrated experimentally by our structures, because they remain closed also at the higher pH (determined from particles in detergent environment). How then can the Cl− conductance of 2 separate pores be affected by structural shifts of the CBS domains? Examination of the interface between the CBS and the transmembrane domain suggests that CBS2 interacts with αD–E, a loop previously shown to affect slow gating (Fig 3C and 3D) [25,31]. Nucleotides may also interact directly with the transmembrane domain when bound in the cleft between CBS1 and CBS2 (the latter observed in structures of isolated CBS domains [13]; Fig 3E). It is conceivable that these structural arrangements and the direct physical connection between CBS and αR—all structural elements leading to the GluGATE constrictions site—allow structural adjustment of the transport pathway and thus chloride conductance regulation (Fig 3C). Such structural effects will be propagated between the monomers via the CBS domains, in agreement with concurrent modulation of the 2 conducting pathways in the dimer [16]. We note that the CBS portions that interact with the transmembrane and the CBS domain of the adjacent monomer are structurally (and at interaction sites also sequencewise; S10 Fig) conserved (Fig 3E), and therefore this may represent a unifying mechanism of slow gating for CLC proteins. ClC-1 defects cause recessive (Becker type) or dominant (Thomsen type) myotonia congenita, typically associated with complete disruption of channel function or with a dominant negative effect in heterodimeric wild-type (WT) -mutant complexes [7], respectively. Our structure now allows mapping of such (or other experimental) ClC-1 substitutions for evaluation of structure–function–disease and -phenotype relationships (Fig 4). Several dominant and recessive mutations induce an alteration of the overall gating from depolarization to hyperpolarization activated, yielding a similar intracellular Cl−-sensitive gating as described for ClC-2 [32]. Therefore, the different gating profiles of ClC-1 and ClC-2 likely do not necessitate major structural differences. These residues are generally surface exposed and localized to the extracellular half, including the vestibule and the pore-constricting residues Lys231 and Arg421 (Fig 4B) [26,27,32–35]. In contrast, many dominant mutations exert a “shift” of the common gate to open probability to positive voltages, leading to significant reduction of gCl at the physiological membrane potential [36]. Such mutations cluster primarily at the dimer interface and in the intracellular vestibule and pore region (Fig 4C, and 4D and S5D Fig). One is located in CBS2, close to the membrane domain, in agreement with the above-mentioned mechanism of slow-gating regulation exerted via CBS2. Residues that affect binding of one of the most commonly used ClC-1 inhibitors, the lipophilic 9-anthracene-carboxylic acid (9-AC), are all buried into a CAVER [37]-computed membrane-embedded cavity on the intracellular side that stretches to GluGATE, in agreement with the intracellular mechanism of action proposed for this compound (Fig 4E and 4F and S11 Fig) [24]. Because this pocket is lined by multiple hydrophobic and a few negatively charged residues, it is unlikely to allow chloride conductance (proton access is possible) but rather 9-AC–induced interference of flux across GluGATE and may thus represent a suitable site for future drug-discovery efforts.
In summary, we report the molecular structure of Cl−-conducting human ClC-1, sharing an overall fold similar to other CLC proteins, with a narrow connecting pore and positively charged vestibules attracting Cl- ions similar to CFTR [38]. The structure exhibits several unique features, including shifts in the central GluGATE-TyrC pair, a more closed extracellular vestibule, and a wider penetration profile from the intracellular side, the latter representing a distinct feature of CLC channels separating them from transporters. We propose a model for adenine nucleotide and pH regulation of the common gate via CBS2 and the intracellular loops congruent with previous functional data. Overall, these findings significantly increase our understanding of Cl− conductance in physiology and open new opportunities for biomedicine. For example, the positively charged constriction of the extracellular vestibule and the putative 9-AC pocket may serve as favorable target sites for stimulators or inhibitors from outside or inside the cell, respectively. During the course of the preparation of this manuscript, the structure of human ClC-1 was reported by another group [39]. The ClC-1 structures display only limited differences despite that different overproduction hosts were exploited. The authors detected a similar putative 9-AC binding pocket (the alternative pathway) and conformational flexibility in the CBS region (determined at pH 7. 4), in agreement with our findings. We anticipate that the pH-dependent conformational changes reported here—in conjunction with mutational efforts using, e. g. , single-channel recordings, as for the first time demonstrated in this work, will allow for more refined studies to further resolve the mechanism of slow-gating in CLC proteins.
Yeast codon-optimized cDNA encoding human ClC-1 (UniProt accession P35523) was purchased from Genscript (Genscript, USA). cDNA was inserted into pEMBLyex4 [40] along with yeast-enhanced GFP by homologous recombination to encode ClC-1, followed by a Tobacco Etch Virus (TEV) cleavage site, GFP, and a His10 tag. The correct nucleotide sequence of the expression construct was verified by DNA sequencing (Eurofins MWG Operon, Germany). Human ClC-1 was produced in the PAP1500 strain [41] grown in computer controlled 15-L bioreactors as previously reported but without addition of any chloride salts (such as NaCl) [42]. Yeast cells were harvested approximately 90 hours after induction of ClC-1 expression. For crude membrane preparations, approximately 25 g of yeast cells were resuspended in 25 mL lysis buffer (25 mM imidazole [pH 7. 5], 1 mM EGTA, 1 mM EDTA, 10% glycerol, 5 mM β-mercaptoethanol) supplemented with protease inhibitors (1 μg/mL leupeptin, pepstatin, and chymostatin, and 1 mM PMFS). Cells were disrupted by addition of glass beads (0. 4–0. 8 mm) and vortexed in 50-mL Falcon tubes 8 times for 1 minute. The supernatant was collected, and glass beads were washed several times in ice-cold lysis buffer. The cell lysate was centrifuged at 1,000g for 10 minutes to remove cell debris. Crude membranes were pelleted from the supernatant by ultracentrifugation at 160,000g for 90 minutes; resuspended in a buffer containing 50 mM Tris (pH 7. 5), 300 mM NaCl, 10% glycerol, 1 mM PMSF, and EDTA-free protease inhibitors (Sigma); and homogenized in a Potter-Elvehjem homogenizer. Subsequently, membranes were solubilized by adding dodecyl-β-maltoside (DDM) and cholesteryl semi succinate (CHS; from Anatrace) at final concentrations of 1% and 0. 33%, respectively, and incubated at 4°C for 3 hours under gentle stirring. Nonsolubilized material was removed by ultracentrifugation at 30,000 rpm for 30 minutes in a Beckman Ti 60 rotor. Ni-beads from 5 mL of slurry (Thermofisher) were incubated with the supernatant for 2 hours under gentle stirring. To prevent unspecific binding, 30 mM imidazole was added. Resin was transferred to a 5-mL Econo column (Bio-Rad) and washed with 10 column volumes of high-salt buffer (50 mM Tris [pH 7. 5], 800 mM NaCl, 5% glycerol, 0. 4 mg/mL DDM, and 0. 04 mg/mL CHS) followed by 10 column volumes of low-salt buffer (50 mM Tris [pH 7. 5], 300 mM NaCl, 5% glycerol, 0. 4 mg/mL DDM, and 0. 04 mg/mL CHS). ClC-1 protein was liberated from the beads by overnight incubating in 10 mL low-salt buffer containing 0. 2 mg of TEV protease. Ni-beads were washed twice with 5 mL of low-salt buffer, and all collections were pooled and concentrated to approximately 1 mL using a 100,000 kDa cutoff concentrator device (Sartorius). Amphipol PMAL-C8 (Anatrace) was added to the purified protein at a mass ratio of 1: 5 and incubated overnight. To remove DDM, protein was dialyzed overnight against final buffer (20 mM Tris [pH 7. 5], 100 mM NaCl, 0. 2 mM TCEP) supplemented with 100 mg of SM-2 Bio-Beads (Bio-Rad). The protein-amphipol complex was applied to a Superdex-200 column equilibrated with final buffer. Peak fractions were collected and concentrated to approximately 0. 5 mg/mL. For the low pH samples, the purification procedure was identical except for using 20 mM BisTris (pH 6. 2) (instead of Tris [pH 7. 5]) in the final buffer (final protein concentration only reached approximately 0. 3 mg/mL due to precipitation). Single-channel ion current was recorded using 2 separate methods, as follows: Cryo-EM grids were prepared with the Vitrobot Mark IV (FEI) operated at 100% humidity at 4°C. Immediately prior to sample vitrification, Quantifoil 1. 2/1. 3-μm holy carbon grids were glow-discharged with Easyglow (TedPella), and fluorinated fos-choline-8 (Anatrace) was added to the protein sample to a final concentration of 3 mM, which was an essential step for producing good quality thin ice. For each grid, an aliquot of 3. 5 μL was applied and incubated for 20 seconds inside the Vitrobot. Blotting time was set to 2. 5 seconds with 2 seconds of drain time. The low pH sample was treated identically, except for incubation with 0. 3 mM NAD before freezing (and that no fluorinated fos-choline-8 was added to obtain one of the pH 6. 2 data sets). Cryo-EM data sets were collected on a Titan Krios electron microscope (FEI) operating at 300 keV with a Gatan K2 Summit direct electron detector attached to a Gatan imaging filter (GIF). Movies were recorded under super-resolution counting mode at a pixel size of 0. 535 Å and a dose rate of 0. 876 e/pixel/frame for a total of 60 frames. The total electron dose was 45 electrons per Å2 per movie for 9 seconds. Cryo-EM movies were first gain-corrected and 2× binned to a final pixel size of 1. 07 Å. Dose-weighted and nondose-weighted summed micrographs were generated with MotionCorr2 [43] using all frames except the first one. Defocus values were calculated with the nondose-weighted micrographs using Gctf [44]. Next, image processing was conducted using dose-weighted micrographs with the predetermined defocus. Template-free particle picking was done using Kai Zhang’s Gautomatch software (https: //www. mrc-lmb. cam. ac. uk/kzhang/Gautomatch). All following processing steps were done in Relion 2. 0 [45] using a box size of 288 pixels. For the pH 7. 5 data set, a total of 594,609 auto-picked particles from 4,475 micrographs with a defocus range of −1. 0 to −3. 0 μm were subjected to several rounds of reference-free 2D classification to remove defective particles. The selected 477,729 particles were sorted using 3D classification. Selected classes were refined using masks, either with the complete protein excluding the amphipol belt or with the membrane domain only. Multiple cryo-EM density maps were calculated demonstrating structural heterogeneity of the protein. 3D classification of particles into 5 classes provided the best class consisting of 176,871 particles (representing more than 37% of all particles). A soft mask covering the entire protein without amphipol belt yielded a map with an overall resolution of 4. 00 Å, and a tighter mask only containing the membrane domain resulted in map with resolution of 3. 63 Å. To further investigate the structure heterogeneity in the cytoplasmic domain, the 2D selected particles were first refined, and then the refined per-particle parameters were applied for 3D classification, only performing local angular searches within ±10 degrees. This local 3D classification resulted in 9 classes, and the 2 major classes differed primarily in the cytoplasmic domain. Refinement of these 2 classes, each representing approximately 15% of all selected particles, yielded overall map resolutions of 4. 34 Å and 4. 28 Å, respectively. For the pH 6. 2 data set collected with fluorinated fos-choline-8,552,914 particles were autoselected from 4,119 motion-corrected micrographs, and 300,572 particles were selected after 2D classification for further processing; 3D classification into 5 classes generated the best class, which eventually was refined to a final resolution of 4. 47 Å. Combination of the data collected at pH 6. 2 with and without fluorinated fos-choline-8, and a similar local angular search strategy as for the pH 7. 5 data set, generated a final map of 4. 2 Å of the best class (based on approximately 30% of the total particles). C2 symmetry was applied for all classification procedures, and all maps were sharpened with a B-factor of −100 Å2. Local resolution was calculated using the postprocessed map, and the map was filtered according to the local resolution and used for model building. The initial model was generated using the SWISS-MODEL online server and the ClC-K structure [9] (PDB-ID 5TQQ) as a template. The model was first fitted into the cryo-EM density map and later manually built in COOT [46]. The 3. 6 Å membrane domain density map was sufficient for building the entire membrane domain (residues 115 to 589) with only 1 loop missing (residues 254–261). The built model was refined using phenix. real_space_refine of the Phenix software package [47]. C2 symmetry was imposed during the refinement by using strong non-crystallographic symmetry (NCS) restraints. Secondary structure restraints and Ramachandran restraints were also imposed during refinement. The resolution and connectivity of the cytoplasmic domain was insufficient for de novo model building. Instead, a homology model based on the available structure of the CBS domains of ClC-0 (PDB-ID 2D4Z [30]) was generated and docked into different maps. The refinement of the cytoplasmic domain was conducted by local grid minimization, model morphing, and simulated annealing implemented in the phenix. real_space_refine software [47]. To prevent overfitting, the map resolution was restricted to 5 Å, the local resolution of the cytoplasmic domains as determined by Relion postprocessing. After model building, the models were trimmed to only include the minimal CBS architecture, consisting of 2 helices and a β-sheet. The quality of the models were validated assessed using Molprobity [48] (see S1 Table for statistics). All figures except for Fig 3A and 3B were generated using the model based on the 4. 0 Å (Map 1). The ClC-1 dimer with Glu232 either protonated or deprotonated was inserted into a palmitoyloleoylphosphocholine (POPC) membrane, and CHARMM36 force field parameters [49,50] were generated using CHARMM-GUI [51]. The simulations were performed using the GROMACS 2016. 4 simulation software [52]. Each system was energy minimized and equilibrated in a stepwise manner using 25-ps NVT simulations with decreasing restraints on the protein and lipid heavy atoms. In these simulations, a 1-fs time step was used and the temperature was maintained at 310 K with a Berendsen temperature-coupling scheme [53]. The following set of NPT simulations further released heavy-atom restraints for 0. 1 ns, 10 ns, and 10 ns, respectively. Here, a 2-fs time step was used and the pressure was kept constant at 1 bar using a Berendsen pressure barostat [53]. In a 100 ns production simulation, all atoms were unrestrained, and the temperature and pressure coupling schemes were Nose-Hoover [54,55] and Parrinello-Rahman [56,57], respectively. The GROMACS pull code with a force constant of 1,000 kJ mol−1 nm−2 was applied for 300 ps to the Cl− ion in closest vicinity of Glu232 in 1 monomer. The pull rate was 0. 1 Å per ps, and the pull force was directed along the vertical axis of the membrane. The potential of mean force (PMF) was calculated using umbrella sampling from 1 Å windows along the ion path. The figures were generated using VMD software[58].
Chloride transporting CLC proteins are expressed in a wide range of organisms, and the family encompasses several members with numerous roles in human health and disease by allowing movement of chloride ions across the membranes that encapsulate cells and cellular organelles. Structurally, CLCs form dimers possessing a separate ion translocation pathway in each monomer, and they can operate as either channels or transporters that exchange chloride for protons. The CLC channel ClC-1 is critical to skeletal muscle excitability and has been proposed as a target to alleviate neuromuscular disorders. Here, we have analyzed the structure of human ClC-1 and revealed the high similarity of its ion conducting pathway to those observed in other CLC members, including prokaryotic and algal transporters. Our data suggest how ClC-1 is regulated by environmental cues to allow opening and closure, thereby permitting attenuation of muscle function. Our results help with understanding the principal determinants that govern CLC proteins and may guide downstream translational applications to combat muscle pathologies.
Structure of the human ClC-1 chloride channel