1. Structural Biology and Molecular Biophysics

Cryo-EM structures of an LRRC8 chimera with native functional properties reveal heptameric assembly

  1. Hirohide Takahashi
  2. Toshiki Yamada
  3. Jerod S Denton
  4. Kevin Strange
  5. Erkan Karakas  Is a corresponding author
  1. Department of Molecular Physiology and Biophysics, School of Medicine, Vanderbilt University, United States
  2. Center for Structural Biology, Vanderbilt University, United States
  3. Department of Anesthesiology, Vanderbilt University Medical Center, United States
  4. Department of Pharmacology, School of Medicine, Vanderbilt University, United States
https://doi.org/10.7554/eLife.82431
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Cite this article
  1. Hirohide Takahashi
  2. Toshiki Yamada
  3. Jerod S Denton
  4. Kevin Strange
  5. Erkan Karakas
(2023)
Cryo-EM structures of an LRRC8 chimera with native functional properties reveal heptameric assembly
eLife 12:e82431.
https://doi.org/10.7554/eLife.82431
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Abstract

Volume-regulated anion channels (VRACs) mediate volume regulatory Cl- and organic solute efflux from vertebrate cells. VRACs are heteromeric assemblies of LRRC8A-E proteins with unknown stoichiometries. Homomeric LRRC8A and LRRC8D channels have a small pore, hexameric structure. However, these channels are either non-functional or exhibit abnormal regulation and pharmacology, limiting their utility for structure-function analyses. We circumvented these limitations by developing novel homomeric LRRC8 chimeric channels with functional properties consistent with those of native VRAC/LRRC8 channels. We demonstrate here that the LRRC8C-LRRC8A(IL125) chimera comprising LRRC8C and 25 amino acids unique to the first intracellular loop (IL1) of LRRC8A has a heptameric structure like that of homologous pannexin channels. Unlike homomeric LRRC8A and LRRC8D channels, heptameric LRRC8C-LRRC8A(IL125) channels have a large-diameter pore similar to that estimated for native VRACs, exhibit normal DCPIB pharmacology, and have higher permeability to large organic anions. Lipid-like densities are located between LRRC8C-LRRC8A(IL125) subunits and occlude the channel pore. Our findings provide new insights into VRAC/LRRC8 channel structure and suggest that lipids may play important roles in channel gating and regulation.

Editor's evaluation

This paper is valuable to the field of ion channels, as it provides useful information about the assembly of the volume-regulated anions channels formed by LRRC8 proteins. The evidence that the LRRC8C-LRRC8A chimera has native functional properties and adopts a heptameric assembly is convincing. The evidence supporting the physiological relevance of the heptameric assembly and the hypothesized role of lipids is incomplete and will be addressed in future studies.

https://doi.org/10.7554/eLife.82431.sa0

Introduction

Volume-regulated anion channels (VRACs) are expressed widely in vertebrate cell types where they mediate the efflux of Cl- and organic solutes required for cell volume regulation (Jentsch, 2016; Strange et al., 2019). VRACs are activated by increases in cell volume and by large reductions in intracellular ionic strength (Strange et al., 2019).

VRACs are encoded by the Lrrc8 gene family (Qiu et al., 2014; Voss et al., 2014), which comprises five paralogs termed Lrrc8a-e (Abascal and Zardoya, 2012; Voss et al., 2014). Native VRAC/LRRC8 channels are heteromers with unknown stoichiometry. LRRC8A is an essential VRAC/LRRC8 subunit and must be co-expressed with at least one other paralog to reconstitute volume-regulated channel activity (Syeda et al., 2016; Voss et al., 2014). High-resolution cryo-electron microscopy (cryo-EM) structures have been determined for homomeric LRRC8A (Deneka et al., 2021; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019) and LRRC8D (Nakamura et al., 2020) channels demonstrating that they have a hexameric configuration.

Defining the molecular basis by which VRAC and other volume-sensitive channels detect cell volume changes is a fundamental and longstanding physiological problem. Detailed molecular understanding of this important problem requires accurate channel structural information. Homomeric LRRC8A and LRRC8D channels have abnormal functional properties or are not expressed in the plasma membrane (Voss et al., 2014; Yamada et al., 2021; Yamada and Strange, 2018; Zhou et al., 2018). Structure-function studies of VRAC/LRRC8 heteromeric channels are complicated by their undefined, likely variable, and experimentally uncontrollable stoichiometry. To circumvent these problems, we developed a series of novel homomeric LRRC8 chimeras that exhibit functional properties similar to the native heteromeric VRAC/LRRC8 channels (Yamada and Strange, 2018).

We describe here the cryo-EM structures of the LRRC8C-LRRC8A(IL125) chimera, hereafter termed 8C-8A(IL125). 8C-8A(IL125) consists of a 25-amino acid sequence unique to the first intracellular loop, IL1, of LRRC8A inserted into the corresponding region of LRRC8C (Figure 1). Like native VRAC/LRRC8 channels, 8C-8A(IL125) chimeras are activated strongly by cell swelling and low intracellular ionic strength (Yamada and Strange, 2018). We demonstrate that the 8C-8A(IL125) chimeric channel is a large-pore seven-subunit heptamer, like homologous pannexin channels (Bhat and Sajjad, 2021; Deng et al., 2020; Jin et al., 2020; Kuzuya et al., 2022; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020). The pore diameter is similar to that estimated for native VRACs, and permeability to large organic anions is significantly higher than that of homohexameric LRRC8A channels. Lipid-like densities are located between 8C-8A(IL125) channel subunits and occlude the channel pore, as has been shown recently for human pannexin 1 (Kuzuya et al., 2022). Our results, together with the previous studies (Deneka et al., 2021; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019; Nakamura et al., 2020), demonstrate that LRRC8 proteins can form anion channels with different subunit numbers.

Figure 1 with 1 supplement see all
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Construct design of 8C-8A(IL125).

(A) Sequence alignment of LRRC8A and LRRC8C around the swapped IL125 region. The swapped region in the 8C-8A(IL125) construct is shown inside a black box. The key below amino acid sequences denotes identical (*), conservative (:), and semi-conservative (.) sequences. Space indicates the residues that are not conserved. (B) The amino acid sequence of the 8C-8A(IL125) construct around the swapped IL125 region. (C) Schematic diagram of the 8C-8A(IL125) protein highlighting the relative position of the swapped IL125 region.

Results

Structure determination

To facilitate structure-function understanding of VRAC/LRRC8 channel regulation and pore properties, we expressed the 8C-8A(IL125) chimera in Sf9 insect cells and purified the detergent-solubilized complexes by affinity and size exclusion chromatography (Figure 1—figure supplement 1). The purified 8C-8A(IL125) complex appeared larger than the LRRC8A hexamers in both native-PAGE and size exclusion chromatography analysis (Figure 1—figure supplement 1B–C). We performed single-particle cryo-EM analysis to determine the structure of the complex (Figure 2 and Figure 2—figure supplements 1 and 2). Following 2D classification and ab initio 3D reconstruction, we performed 3D classification and obtained five distinct 3D classes for 8C-8A(IL125) (Figure 2—figure supplement 1). The 8C-8A(IL125) maps showed no apparent symmetric arrangement. Therefore, the final reconstructions were done without enforcing any symmetry and resulted in five structures with resolutions in the range of 3.4–4.0 Å (Figure 2A and B, Figure 2—figure supplements 1 and 2, and Table 1).

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Cryo-electron microscopy (cryo-EM) structure of 8C-8A(IL125).

(A–B) Cryo-EM maps of 8C-8A(IL125) class 1 structure viewed through the membrane plane (A) and from the cytoplasm (B). (C) Ribbon representation of the 8C-8A(IL125) class 1 structure viewed through the membrane plane. (D–E) Unsharpened cryo-EM maps of 8C-8A(IL125) class 1 structure viewed through the membrane plane (D) and from the cytoplasm (E), emphasizing low-resolution features. (F) Structural comparison of the 8C-8A(IL125) (cyan) and LRRC8A (magenta, PDB ID: 5ZSU) subunits.

Table 1
Cryo-electron microscopy (cryo-EM) data collection, refinement, and validation statistics.
Data collection and processing
MicroscopeFEI Krios G3i microscope
DetectorGatan K3 direct electron camera
Nominal magnification×81,000
Voltage (kV)300
Electron exposure (e/Å2)54
Defocus range (µm)–0.8 to –1.5
Pixel size (Å)1.1
Number of Micrographs3198
Particles images (no.)846,122
Conformational stateClass 1Class 2Class 3Class 4Class 5
Symmetry imposedC1C1C1C1C1
Final particles images (no.)203,011132,722100,77293,17985,591
Map resolution (Å)
(FSC threshold = 0.143)		3.43.63.73.84.0
Refinement    
Model resolution (Å)
(original map, FSC threshold = 0.5)		3.63.93.94.24.4
B-factor used for map sharpening (Å2)–102.0–88.2–84.7–70.5–83
Model composition    
Non-hydrogen atoms17,49917,49917,46510,43510,435
Protein residues21012101209721012101
Mean B factors (Å2)    
Protein32.423.532.444.938.34
R.m.s. deviations    
Bond lengths (Å)0.0030.0020.0030.0020.004
Bond angles (°)0.5630.5250.5530.5160.807
Molprobity score1.721.681.751.371.49
Clash score5.044.846.021.141.54
Poor rotamers (%)0.00.00.00.00.0
Favored (%)92.993.593.689.9588.3
Allowed (%)7.16.56.410.011.6
Disallowed (%)0000.050.1

Similar to LRRC8A and LRRC8D structures, the 8C-8A(IL125) structure comprises four domains, the extracellular domain (ECD), transmembrane domain (TMD), intracellular domain (ICD), and leucine-rich repeat (LRR) motif-containing domain (LRRD) (Figure 2). The cryo-EM maps for the ECD and TMD revealed high-resolution features allowing us to build an atomic model comprising most of the ECD and TMD except residues 60–94 in the ECD and the first 15 residues of the N-terminus. The quality of the cryo-EM maps for the ICDs was improved by performing local refinements, and the resulting maps were used to build a model (Figure 2—figure supplement 2). However, the first intracellular loop containing the swapped IL125 region remained unresolved. Although the unsharpened maps revealed clear features for the entire protein, the local resolution for the LRRD was insufficient to build an atomic model, and the local refinement strategy we applied did not provide any meaningful improvement in LRRD resolution (Figure 2A–E and Figure 2—figure supplements 1 and 2). Therefore, we did not build an atomic model for the LRRD and used the maps without B-factor sharpening to assess their structure and overall arrangement relative to the rest of the protein complex (Figure 2D–E).

8C-8A(IL125) chimeras form heptameric channels

The overall structure of an individual 8C-8A(IL125) subunit is similar to that of LRRC8A (Figure 2F) and LRRC8D. However, unlike homomeric LRRC8A and LRRC8D channels, which are hexamers (Deneka et al., 2021; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019; Nakamura et al., 2020), the 8C-8A(IL125) chimeric channel is a seven-subunit heptamer, similar to the homologous pannexin channels (Figure 2B and E; Figure 2—figure supplement 3; Bhat and Sajjad, 2021; Deng et al., 2020; Jin et al., 2020; Kuzuya et al., 2022; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020).

The subunit arrangement of the 8C-8A(IL125) heptameric channel is asymmetric as opposed to LRRC8A and LRRC8D hexamers, which have two-, three-, or sixfold symmetric arrangements (Deneka et al., 2021; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019; Nakamura et al., 2020). When the ICD and TMD are viewed from the cytoplasm, 8C-8A(IL125) subunits are organized into two distinct groups, one with four subunits and the other with three subunits (Figure 3A–D). These two groups of subunits associate via a loose interface, while the subunits within each group associate via a tight interface (Figure 3A–D and Figure 3—figure supplements 1–3).

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Subunit arrangement of the 8C-8A(IL125) chimera.

(A–B) Surface representation of the 8C-8A(IL125) class 1 structure viewed from two sides, highlighting the ‘tight’ (A) and ‘loose’ (B) interfaces. (C–E) Intracellular domain (ICD), transmembrane domain (TMD), and extracellular domains (ECDs) are viewed from the cytoplasm with a depth of view, as shown in panel (B).

In contrast to the ICDs and TMDs, the ECDs are arranged symmetrically and form extensive contacts between the subunits in both tight and loose interfaces (Figure 3E and Figure 3—figure supplement 1). Most of the subunit interaction is between the residues on ELH1 and the linker that connects EL2β1 and EL2β2 in one subunit and the residues on the three β-strands of the neighboring subunit (Figure 3E and Figure 3—figure supplement 1). The contacts between each subunit pair bury about 1900 Å2 of solvent-accessible surface area at the ECDs (Table 2). The buried solvent-accessible surface areas between the subunit pairs in the TMDs and ICDs are considerably smaller and exhibit high variability between the tight and the loose interfaces. In the TMD, the buried areas are about 650 and 110 Å2 for the tight and loose interfaces, respectively (Table 2). The residues within the core of the TMD are loosely associated and the major interactions of the subunits within the TMD are mediated by the residues closer to the ECDs or ICDs at the tight interface (Figure 3—figure supplement 2). At the loose interface, only residues closer to the ECD are in close range for direct interaction (Figure 3—figure supplement 2). In the ICD, the buried areas in the tight interfaces range from 290 to 560 Å2, whereas there are no solvent-inaccessible contacts at the loose interfaces (Figure 3E, Figure 3—figure supplements 2–3, and Table 2).

Table 2
Total buried solvent-accessible surface area between subunits for each domain.
Buried surface area between the neighboring subunits (Å2)*
Subunits
A-B		Subunits B-CSubunits C-DSubunits D-ESubunits E-FSubunits F-GSubunits G-A
TightTightTightLooseTightTightLoose
ECD†1995189719641905190018961869
TMD†66462562012266067099
ICD†46229450005595020
  1. *

    Buried solvent-accessible surface area calculations were performed using the software NACCESS v2.1.1 (Hubbard and Thornton, 1993).

  2. †

    Domain definitions used for these calculations are as follows: ECD: residues 49–121 and 288–310; TMD: residues 20–48, 122–150, 260–287, and 311–342; ICD: residues 151–259 and 343–405.

The asymmetric arrangement observed for the TMDs and ICDs is likely due to their divergent orientation relative to the ECDs. Figure 4A shows the alignment of the seven subunits on their ECDs. TMDs do not align and exhibit up to an 8° difference in their orientation relative to the ECD, while the difference is larger for the ICDs (Figure 4A). Another notable difference between the subunits is in the linker that connects the first β-strand on extracellular loop 1, EL1β1, of the ECD to the transmembrane helix 1 (TM1) of the TMD (Figure 4A–B). In two of the subunits in the class 1 structure, K51 points toward the pore, while D50 points in the opposite direction. In the five other subunits, D50 points toward the pore, and K51 is oriented toward the subunit interface (Figure 4C). When K51 points toward the pore, it creates a groove between the subunits (Figure 4D). These grooves, located between the subunits forming the loose interface in the class 1 structure, connect the space within the ECD to the membrane-facing surface of the TMD (Figure 4E). When D50 points toward the pore, the side chain of K51 occupies this space, closing the groove (Figure 4F).

Figure 4
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Structural heterogeneity of 8C-8A(IL125) protomers.

(A) Structural comparison of the 8C-8A(IL125) subunits in the class 1 structure. The structures are aligned based on their extracellular domains (ECDs). (B) Close-up view of the box region in panel (A), highlighting the structural differences in the loop that connects transmembrane helix 1 (TM1) to EL1β1. Only two subunits are shown. (C) Close-up view of the pore around the residues D50 and K51, which are shown as sticks. The residues that adopt different conformations compared to others are labeled in red. The dashed circle indicates the pore-lining surface. (D) The same view as panel (C) but including the surface representation (transparent) to highlight the distinct interfaces between the subunits. (E–F) Close-up view of the interfaces from the points of view shown in panel (D).

Structural heterogeneity of 8C-8A(IL125) chimeras

We obtained five distinct 3D classes for the 8C-8A(IL125) chimera. The most apparent difference between the structures is in the arrangement of the LRRDs (Figure 5A–B). In class 1, all seven LRRDs are arranged circularly with roughly sevenfold symmetry. For class 5, one of the LRRDs is positioned outside of the quaternary assembly formed by six LRRDs arranged with pseudo-twofold symmetry. The density for the LRRD located outside the assembly is poorly visible, indicating high flexibility relative to the rest of the complex (Figure 5A–B). The arrangements of the LRRDs in the other three 3D classes exhibit diverse arrangements and subunit interactions. As a result, several different subunit interfaces exist between the neighboring LRRDs. However, a detailed analysis of these interfaces is not possible due to the limited resolution of the cryo-EM maps for these regions, prohibiting the building of the models with amino acid assignments.

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Conformational heterogeneity of 8C-8A(IL125) chimeras.

(A–B) Cryo-electron microscopy (cryo-EM) maps (before sharpening) of 8C-8A(IL125) 3D classes viewed through the membrane plane (A) and from the cytoplasm (B). Individual subunits are colored as in Figure 1. The detergent micelle is shown in white. Red arrows point to the weak density of the leucine-rich repeat motif-containing domain (LRRD), which is not located within the LRRD quaternary assembly. Lines represent the depth of the view for LRRD, intracellular domain (ICD), and transmembrane domain (TMD). (C–D) Ribbon representation of the ICD (C) and TMD (D) structures viewed from the cytoplasm with a depth of view indicated in panel (A). The Cα atoms for G163 in ICD and V33 in TMD are shown as spheres. The distances between these atoms in neighboring subunits are shown as lines (in black and red for the tight and loose interfaces, respectively) and labeled in Å.

Consistent with the conformational differences in the arrangements of the LRRDs, the ICDs also exhibit conformational heterogeneity, albeit less pronounced, among the five 3D classes we observed (Figure 5C and Figure 5—figure supplement 1). Figure 5C shows the arrangement of the ICDs and the distances between the Cα atoms of G163 of the neighboring subunits. The non-symmetric and diverse subunit arrangement is evident. The distances between the subunits forming the tight interfaces appear similar in all classes, whereas the distances between the subunits forming the loose interfaces are variable among different classes. A similar variation is also observed within the TMD (Figure 5D and Figure 5—figure supplement 1). However, there are no apparent structural differences at the ECD of the structures (Figure 5—figure supplement 1).

The distinct arrangement of the loops formed by D50 and K51 is also observed in other classes (Figure 5—figure supplement 2). However, the number of subunits with K51 pointing toward the pores differs. For example, the class 2 structure has only one subunit with K51 pointing toward the pore, compared to two subunits in the class 1 structure (Figure 5—figure supplement 2). The groove created by the distinct K51 arrangement is located between the subunits that form one of the two loose interfaces in the class 2 structure, despite the presence of two loose interfaces. It is plausible that this loop is flexible and adopts distinct conformations, possibly depending on the orientation of the TMD relative to the ECD. However, it is not clear if there is any direct correlation between the K51 orientation and the loose interfaces.

Pore structure

Figure 6 shows the pore domain of the 8C-8A(IL125) heptameric chimera compared to the pore domain of LRRC8A hexameric channels. The narrowest region of the heptameric 8C-8A(IL125) channel pore has an average solvent-accessible radius of 4.7 Å and is formed by L105 located on the extracellular side of the protein (Figure 6A). This is considerably larger than the pore radius of 2.0 Å determined from cryo-EM structures of hexameric LRRC8A channels (Figure 6B–C; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019).

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Comparison of channel pores.

(A–B) Pore openings of the 8C-8A(IL125) heptameric channel (class 1 structure) (A) and LRRC8A (PDB ID: 5ZSU) homohexameric channel (B) calculated using the software program HOLE (Smart et al., 1996). Only two opposing subunits are shown. Residues forming the constriction sites are shown as sticks. The first modeled residues at the N-termini are labeled as NT. (C) 1D graph of the average radius along the length of the 8C-8A(IL125) (cyan) and LRRC8A (magenta) channel pores. (D) Relative (Px/PCl) glutathione and lactobionate permeabilities calculated from reversal potential changes induced by replacing bath Cl- with the test anion. Values are means ± SEM (N=4–7). Statistical significance was determined using Student’s unpaired t-test. (E) Representative LRRC8A and 8C-8A(IL125) current traces in the presence of bath Cl- or after substitution with glutathione (Glut-) or lactobionate (Lac-). Currents were elicited by ramping membrane voltage from −100 to +100 mV.

The difference in pore diameters of hexameric LRRC8A channels and heptameric 8C-8A(IL125) channels implies that subunit number impacts channel solute permeability. To assess this directly, we quantified the relative permeabilities (i.e., Px/PCl) of LRRC8A and 8C-8A(IL125) to the large organic anions glutathione and lactobionate. Pglutathione/PCl and Plactobionate/PCl were both significantly (p<0.01) higher for the 8C-8A(IL125) heptameric channel (Figure 6D–E), consistent with its larger pore diameter.

Interaction of lipids with the 8C-8A(IL125)

As shown in Figure 7A, we observe non-protein densities penetrating through the openings between the subunits in the TMD. The shape and length of these densities are consistent with lipid molecules. Similar densities are observed in LRRC8A hexameric channels reconstituted in lipid nanodiscs (Kern et al., 2019). We observe these lipid-like densities on both the cytoplasmic and extracellular sides of the TMD. The lipid-like densities on the extracellular side do not show apparent differences between tight and loose interfaces. However, the lipid-like density closer to the intracellular side is considerably weaker at the loose interfaces compared to the ones at the tight interfaces (Figure 7A). This difference may be due to the larger separation of the subunits at the loose interface, creating more room for the lipid molecules to move.

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Interaction of lipids with the 8C-8A(IL125) chimera.

(A) Ribbon representation of the 8C-8A(IL125) class 1 structure along with the lipid-like cryo-electron microscopy (cryo-EM) densities (yellow) between the subunits viewed through the membrane plane. Loose and tight interfaces are indicated with arrows. (B) Close-up views of the boxed regions in panel (A). The cryo-EM map for the entire region visualized is shown as gray mesh. The phospholipid molecule shown as yellow sticks within the lipid-like density is used for illustration purposes only and not included in the deposited coordinate files. Select residues near the lipid-like densities are shown as sticks and labeled. (C–D) Surface representation of the 8C-8A(IL125) pore colored based on electrostatic charge (C) and hydrophobicity (D). The hypothetical lipid bilayer around the transmembrane domain (TMD) is built using CHARMM-GUI (Wu et al., 2014) and shown as sticks (cyan). Positioning of the protein within the bilayer is calculated using PPM 2.0 (Lomize et al., 2012). (E) A sliced view of the unsharpened cryo-EM map (gray, transparent) with the ribbon representation of the 8C-8A(IL125) class 1 structure. Arrows indicate densities corresponding to pore-blocking lipid-like densities and detergent micelles. (F) Hypothetical interaction of lipids with the inner surface of the pore is shown by placing phospholipids using CHARMM-GUI (Wu et al., 2014). Surface representation of the 8C-8A(IL125) colored based on hydrophobicity as in panel (D).

Although the quality of the maps does not allow us to identify the lipid molecules leading to the densities we observe, the ones with the strongest features suggest phospholipids as the most likely candidates (Figure 7B). As shown in Figure 7B, a phospholipid molecule can be reasonably fitted into the strongest density we observe. The polar phosphate moiety of the lipid molecule is coordinated by two lysine residues (K125 and K318) and a histidine residue (H314) of the neighboring subunits (Figure 7B). W122, Y126, and L130 are close to the acyl chains (Figure 7B). One of the acyl chains extends through the space between the subunit and reaches the interior surface of the channel. The second acyl chain is less defined. The area contains additional non-protein densities suggesting the presence of other ordered lipid molecules.

When we examined the surface properties of the pore-lining residues, we found high densities of charged amino acid residues that align with the phospholipid head groups of the membrane bilayer surrounding the TMD (Figure 7C). A hydrophobic patch of amino acid residues aligns with the hydrophobic core of the membrane bilayer (Figure 7D).

Intriguingly, we observed two layers of non-protein densities within the channel pore that align with the boundaries of the hydrophobic patch and the detergent micelle around the TMD in the cryo-EM maps for all classes (Figure 7E). Similar densities have been observed in the structures of other large-pore channels, including pannexins and innexin (Burendei et al., 2020; Kuzuya et al., 2022). It is plausible that these densities are of a lipid bilayer that occupies the pore as illustrated in Figure 7F. In this model, the hydrophobic acyl chains would occupy the space encircled by the hydrophobic patch, and the polar head groups would interact with the charged residues at the borders of the hydrophobic patch.

To confirm that these densities are not detergent-induced artifacts, we reconstituted the 8C-8A(IL125) chimera in lipid nanodiscs and performed cryo-EM analysis (Figure 7—figure supplement 1). Although the resolution of the cryo-EM maps could not be improved beyond 7 Å, the same bilayer-like density within the channel pore was visible (Figure 7—figure supplement 1). It is noteworthy that the lipid-like densities within the pores are similar in intensity and shape to the lipid densities observed around the protein within the nanodisc (Figure 7—figure supplement 1).

Discussion

Except for LRRC8A, homomeric LRRC8 channels do not traffic to the plasma membrane and cannot be functionally characterized (Voss et al., 2014; Yamada and Strange, 2018; Zhou et al., 2018). LRRC8A homomers form plasma membrane channels, but they exhibit abnormal functional properties. Most notably, LRRC8A channels are not activated by cell swelling under normal physiological conditions (Yamada et al., 2021; Yamada and Strange, 2018) and are only weakly activated by extreme reductions in cytoplasmic ionic strength (Figure 6—figure supplement 1; Yamada et al., 2021). LRRC8A channels also exhibit grossly abnormal DCPIB pharmacology (Yamada et al., 2021). The non-native functional properties of LRRC8A channels indicate that they have a non-native structure, which presents significant limitations for understanding how VRAC/LRRC8 channels are regulated and how pore structure determines solute permeability characteristics.

Homomeric LRRC8 channels with physiologically relevant functional properties are formed by chimeric proteins containing parts of the essential subunit LRRC8A and another LRRC8 paralog (Yamada and Strange, 2018). The 8C-8A(IL125) chimeric channel exhibits normal sensitivity to cell volume changes and intracellular ionic strength and normal DCPIB pharmacology (Figure 6—figure supplement 1; Yamada et al., 2021; Yamada and Strange, 2018).

Unlike homomeric LRRC8A and LRRC8D channels, which are hexamers, our studies demonstrate that 8C-8A(IL125) chimeric channels have a heptameric structure (Figures 2–3). The number of subunits required to form native VRAC/LRRC8 channels is unknown. However, native gel, crosslinking, and mass spectrometry studies (Syeda et al., 2016) and photobleaching experiments (Gaitán-Peñas et al., 2016) suggest that VRAC/LRRC8 channels are formed by ≥6 LRRC8 subunits.

The 8C-8A(IL125) has a limiting pore radius of 4.7 Å (Figure 6 and Figure 6—figure supplement 2A), which unlike the much narrower LRRC8A pore (Figure 6 and Figure 6—figure supplement 2B), is very consistent with the 6–7 Å pore radius estimated for native VRAC/LRRC8 channels (Droogmans et al., 1999; Ternovsky et al., 2004) and the role of these channels play in the transport of large anionic and uncharged organic solutes (Sabirov and Okada, 2005; Strange et al., 2019). A leucine residue located at position 105 forms the 8C-8A(IL125) pore constriction (Figure 6A). In LRRC8A and LRRC8B, L105 is replaced by arginine. LRRC8D and LRRC8E paralogs have either phenylalanine or leucine at the homologous position. Hexameric LRRC8A and LRRC8D channels have limiting pore radii of 2.0 and 3.5 Å, respectively (Figure 6B and Figure 6—figure supplement 2). When we create hypothetical heptamers by aligning LRRC8A and LRRC8D subunits onto individual 8C-8A(IL125) subunits, the pore radii increase to 4.7 and 7.1 Å, respectively, indicating that the oligomeric state has a direct impact in pore size and likely in channel transport properties (Figure 6—figure supplement 2D–F). Consistent with this conclusion, LRRC8A hexameric channels have lower relative permeability to the large organic anions glutathione and lactobionate compared to heptameric 8C-8A(IL125) channels (Figure 6D–E).

The narrow pore radius of 2 Å of the LRRC8A hexameric channel (Figure 6B) also likely accounts for its abnormal DCPIB pharmacology. Native VRAC/LRRC8 channels are inhibited >90% by 10 μM DCPIB. Inhibition is voltage-insensitive and exhibits a Hill coefficient of 2.9 (Friard et al., 2017), suggesting that multiple DCPIB molecules are required to inhibit the channel. The DCPIB pharmacology of 8C-8A(IL125) recapitulates that of native VRAC/LRRC8 channels (Yamada et al., 2021). In contrast, LRRC8A hexameric channels are weakly inhibited by 10 μM DCPIB, and inhibition is strongly voltage-dependent with a Hill coefficient close to 1 (Yamada et al., 2021). The low Hill coefficient is consistent with an LRRC8A cryo-EM structure described by Kern et al., 2019, showing a single DCPIB molecule blocking the channel pore at the narrowest constriction formed by R103 (Figure 6B).

Two recent studies have defined high-resolution cryo-EM structures of LRRC8A/LRRC8C heteromeric channels (Kern et al., 2022; Rutz et al., 2023). Both studies demonstrated that the heteromeric channels could adopt a hexameric conformation with a limiting pore radius of 2–3 Å, similar to that of LRRC8A hexamers. However, these channels either had 5:1 (Kern et al., 2022) or 4:2 LRRC8A:LRRC8C stoichiometries, indicating that LRRC8A:LRRC8C heteromers can adopt multiple oligomeric forms.

Taken together, existing cryo-EM structural data suggest that native VRAC/LRRC8 channels may exist in multiple oligomeric conformations, as has been reported for CALHM channels. It will be critical to relate the functional properties of various channel oligomeric conformations to those of native VRAC/LRRC8 channels. It will also be important to define the role of LRRC8A in channel assembly and conformation. Four groups have shown that LRRC8A has a dominant-negative function. Overexpression of LRRC8A suppresses endogenous (Qiu et al., 2014; Syeda et al., 2016; Voss et al., 2014) and heterologously expressed (Yamada et al., 2016) VRAC/LRRC8 currents. LRRC8A protein levels may therefore impact channel assembly and function.

Kern et al. identified lipids located between individual subunits in LRRC8A hexameric channels reconstituted in lipid nanodiscs (Kern et al., 2019). Intersubunit non-protein densities, which are most likely lipids, are also observed between 8C-8A(IL125) subunits in detergent reconstituted channels (Figure 7A). Importantly, we observed two layers of density resembling lipid bilayer blocking the pore of the 8C-8A(IL125) heptameric channel (Figure 7 and Figure 7—figure supplement 1). In agreement with our observations, in a recent pre-print reporting the hexameric structure of LRRC8A-LRRC8C (Kern et al., 2022), ordered lipid molecules are observed on the extracellular side of the pore opening aligning well with the densities we observed. Intrapore lipids have recently been proposed to be critical structural and regulatory elements of pannexins (Kuzuya et al., 2022) and closely related innexin (Burendei et al., 2020) and CALHM (Drożdżyk et al., 2020; Syrjanen et al., 2020) channels, as well as bacterial mechanosensitive channels (Rasmussen et al., 2019; Reddy et al., 2019; Zhang et al., 2021). Kuzuya et al. suggest that movement of lipids out of the pore through gaps between the subunits of the human pannexin 1 channel is required for channel opening (Kuzuya et al., 2022). A similar mechanism can be suggested for 8C-8A(IL125) heptameric channels. However, further experimental characterization is required to assess if lipids occupy the pore and play an essential role in channel gating.

Leucine-rich repeat motifs have been identified in over 14,000 proteins from viruses to eukaryotes and play essential roles in signal transduction and as sites for protein-protein interactions (Bella et al., 2008; Kobe and Kajava, 2001; Matsushima et al., 2019). The LRR motif can also function as a mechanosensor (Ju et al., 2016; Ju et al., 2015). VRAC/LRRC8 channels most likely arose by fusion of the transmembrane domain encoding portion of a pannexin channel gene with the LRRD portion of an unrelated gene type (Abascal and Zardoya, 2012). Conformational changes in the VRAC/LRRC8 channel LRR domain are correlated with changes in channel activity (Deneka et al., 2021; König et al., 2019), and multiple cryo-EM structures demonstrate that this region is conformationally flexible (Figure 5; Deneka et al., 2021; Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Kern et al., 2019; Nakamura et al., 2020). These data all point to the high likelihood that the VRAC/LRRC8 channel LRRD is a critical element of the channel’s cell volume sensing apparatus.

LRRC8 chimeras with normal cell volume and ionic strength sensitivity must contain all or part of the LRRC8A IL1 (Yamada and Strange, 2018), indicating that this protein region is also critical for channel regulation. The 25-amino acid region of the LRRC8A IL1 inserted into the 8C-8A(IL125) chimera is predicted to be intrinsically disordered (Yamada and Strange, 2018). Consistent with this prediction, this region was not visible in our cryo-EM structures. It is conceivable that the LRRC8A IL1 is required for correct channel assembly and conformation, that it plays a direct role in cell volume sensing and/or that it functions to transduce cell volume-induced conformational changes into changes in the gating.

The mechanisms by which eukaryotic cells sense cell volume changes and transduce those changes into regulatory responses remain mysterious. Understanding how VRAC/LRRC8 channels detect cell volume is greatly constrained by the abnormal functional properties of LRRC8A homomers and by the undefined heteromeric nature of native channels. The high-resolution cryo-EM structures of the 8C-8A(IL125) homomeric chimera, which has native physiological properties, have revealed several novel structural elements of VRAC/LRRC8 channels. These structures now provide the foundation for defining the molecular basis of channel cell volume sensing utilizing structure-guided mutagenesis combined with electrophysiological functional analysis of channels with defined stoichiometry and subunit arrangement.

Methods

Constructs

Human LRRC8A and LRRC8C cDNAs cloned into pCMV6 were purchased from OriGene Technologies. The 8C-8A(IL125) chimera cDNA construct was generated using the Phusion High-Fidelity PCR kit (New England BioLabs). All cDNAs were tagged on their carboxy terminus with Myc-DDK epitopes. For protein expression, the cDNAs encoding human LRRC8A and the 8C-8A(IL125) chimera were sub-cloned with a C-terminal FLAG tag into pAceBac1 vectors and incorporated into baculovirus using the Multibac expression system (Fitzgerald et al., 2006). All constructs were verified by DNA sequencing.

Cell lines

Patch-clamp experiments were performed using Lrrc8-/- HCT116 cell line in which the five Lrrc8 genes were disrupted by genome editing. Lrrc8-/- cells were prepared and authenticated in the Thomas Jentsch Lab (Ullrich et al., 2016). They were a generous gift from Thomas Jentsch. The absence of VRAC/LRRC8 channel activity in the Lrrc8-/- cell line was confirmed routinely by patch-clamp electrophysiology performed on cells transfected with GFP cDNA only. The cells were tested negative for Mycoplasma contamination.

The cells were grown in McCoy’s 5A media (HyClone) supplemented with 10% fetal bovine serum (R&D Systems), 50 units/ml of penicillin, and 50 µg/ml streptomycin in a humidified incubator at 37°C with 5% CO2.

Protein expression and purification

8C-8A(IL125) chimera was expressed in Sf9 cells (4×106 cells/ml) at 27°C for 48 hr. Cells were harvested by centrifugation (2000 × g) and resuspended in a lysis buffer composed of 150 mM NaCl and 50 mM Tris-HCl, pH 8.0, 1 mM phenylmethylsulfonyl fluoride. After cell lysis using Avastin EmulsiFlex-C3, the cell lysate was clarified from large debris by centrifugation at 6000 × g for 20 min. The cleared lysate was centrifuged at 185,000 × g (Type Ti45 rotor, Beckman) for 1 hr. Membrane pellets were resuspended and homogenized in ice-cold resuspension buffer (150 mM NaCl, 50 mM Tris-HCl, pH 8.0) and solubilized using 0.5% lauryl maltose neopentyl glycol (LMNG) at a membrane concentration of 100 mg/ml. The solubilized pellets were stirred gently for 4 hr at 4°C, and the insoluble material was separated by centrifugation at 185,000 × g (Type Ti45 rotor, Beckman) for 40 min. The supernatant was then mixed with anti-FLAG affinity gel resin (Sigma) at 4°C for 1 hr. After washing the resin with 10 column volume wash buffer composed of 150 mM NaCl, 50 mM Tris-HCl, pH 8.0, and 0.005% LMNG, the protein was eluted using the wash buffer supplemented with 100 µg/ml FLAG peptide. Protein was further purified by size exclusion chromatography using a Superose 6 Increase column (10/300 GL, GE Healthcare) equilibrated with 150 mM NaCl, 50 mM Tris-HCl, pH 8.0, 0.005% LMNG. The peak fraction corresponding to 8C-8A(IL125) was concentrated to 3.0 mg/ml, centrifuged at 220,000 × g using an S110-AT rotor (Thermo Fisher Scientific) for 10 min, and used immediately for cryo-EM imaging. Human LRRC8A with a C-terminal Flag tag was purified using the same protocol described above for 8C-8A(IL125).

For nanodisc incorporation, 8C-8A(IL125) chimera was purified as described above except n-dodecyl-β-D-maltopyranoside (DDM, 1% for solubilization of the membrane and 0.05% in the purification buffers) was used instead of LMNG. The membrane scaffold protein MSP1E3D1 was expressed using the p MSP1E3D1 plasmid, a gift from Stephen Sligar (Addgene plasmid # 20066; http://n2t.net/addgene:20066; RRID: Addgene_20066) (Denisov et al., 2007). MSP1E3D1 was expressed in Escherichia coli BL21(DE3) cells and purified as described previously using Ni2+ affinity resin (Ritchie et al., 2009). The histidine tag was cleaved off using TEV protease, and the protein was further purified by SEC using a HiLoad 16/600 Superdex 200 pg column equilibrated with 300 mM NaCl and 40 mM Tris-HCl, pH 8.0. Peak fractions containing MSP1E3D1 were collected and stored at –80°C for the nanodisc formation.

MSP1E3D1 nanodisc formation

The preparation of nanodiscs was performed by mixing 8C-8A(IL125) purified in the presence of DDM with POPC lipids (Avanti Polar Lipids, Inc) and MSP1E3D1 at a final molar ratio of 1:2.5:250 (8C-8A(IL125):MSP1E3D1:POPC). The mixture was incubated with Biobeads SM2 (Bio-Rad) overnight. After removing the biobeads by centrifugation, the protein sample was concentrated and purified by SEC using a Superose 6 10/300 Increase column equilibrated with 150 mM NaCl and 50 mM Tris-HCl, pH 8.0. Peak fractions corresponding to 8C-8A(IL125)-MSP1E3D1 nanodiscs were concentrated to 2.0 mg/ml and used for cryo-EM grid preparation immediately.

Cryo-EM sample preparation and data collection

Purified 8C-8A(IL125) was applied to 300 mesh UltrAuFoil holey gold 1.2/1.3 grids (Quantifoil Microtools) that were glow discharged for 10 s at 25 mA. The grids were blotted for 4 s at force 12 using double-layer Whatman filter papers (1442-005, GE Healthcare) before plunging into liquid ethane using an FEI MarkIV Vitrobot at 8°C and 100% humidity. Samples were imaged using a 300 kV FEI Krios G3i microscope equipped with a Gatan K3 direct electron camera. Movies containing 400 frames were collected in super-resolution mode at ×81,000 magnification with a physical pixel size of 1.1 Å/pixel and defocus values at a range of –0.8 to –1.5 µm using the automated imaging software SerialEM (Mastronarde, 2005).

8C-8A(IL125) nanodisc grids were prepared as described above. Samples were imaged using a 300 kV FEI Krios G4 microscope equipped with a Gatan K3 direct electron camera. Movies containing 50 frames were collected in super-resolution mode at ×103,000 magnification with a physical pixel size of 0.818 Å/pixel and defocus values at a range of –0.8 to –2.2 µm using the automated imaging software EPU (Thermo Fisher Scientific).

Cryo-EM data processing

All image processing was performed using CryoSparc2 (Punjani et al., 2017). Motion correction and CTF estimations were performed locally using Patch Motion Correction and Patch CTF Estimation procedures. Initial particle picking was performed by blob search. Particles were then binned 4× and extracted. After 2D classification, classes with clear structural features were selected and used for template-based particle picking. The new set of particles was binned four times and extracted. After 2D classification, a cleaned particle set was used for the ab initio 3D reconstruction. The resulting map was used as a model for 3D classification. After a series of 2D and 3D classification runs, particles were reextracted with a box size of 360×360 pixels. These particles were separated into six classes using 3D classification. Five out of six classes revealed interpretable density maps, and these particles were processed further using non-uniform refinement to obtain the final cryo-EM maps for each class.

To improve the quality of the cryo-EM maps around the ICDs, we performed local refinements using masks that cover the ICD of each subunit (Figure 2—figure supplement 2). We observed a noticeable improvement in class 1 structure and used these maps for model building. We applied the same strategy for the LRRDs. However, the quality of the maps did not improve to a level allowing us to build reliable models.

Data processing for 8C-8A(IL125) nanodiscs was performed as described above, with the following exceptions. Motion correction was performed using MotionCor2 (Zheng et al., 2017) in Relion 3.0 (Zivanov et al., 2018). The final set of particles was extracted with a box size of 480×480 pixels, and they were classified into three 3D classes.

Model building

Models were built using Coot (Emsley and Cowtan, 2004). We initially placed the human LRRC8C model from the AlphaFold protein structure prediction database (Jumper et al., 2021; Varadi et al., 2022) in the class 1 density. We manually fitted the individual residues into the density while removing the parts that did not have corresponding interpretable densities. Once we built a complete subunit, we copied the model into the other six protomers and manually fit individual residues into the density. The model was refined using Phenix real-space refinement (Afonine et al., 2018). We performed iterative build-refine cycles till a satisfactory model was obtained. The resulting model was fitted into the class 2 and 3 maps and fitted into the density using the same build-refine iterations as described above. Because of their limited resolution, most parts of class 5 and 6 structures were modeled without their side chains (i.e., as alanines) while maintaining their correct labeling for the amino acid type. The LRRDs were not modeled in all five structures. 8A-IL125 is longer by four amino acids than the corresponding region in LRRC8C (Figure 1). Therefore, the residue numbering after 8A(IL125) in the 8C-8A(IL125) chimera increases by four amino acids in the polypeptide. However, we kept the LRRC8C numberings in the models to facilitate easier comparison with other studies. Validations of the structural models were performed using MolProbity (Williams et al., 2018) implemented in Phenix (Afonine et al., 2018).

Some of the data processing and refinement software was supported by SBGrid (Morin et al., 2013).

Patch-clamp electrophysiology

LRRC8A and 8C-8A(IL125) cDNA constructs were expressed in Lrrc8-/- HCT116 cells transfected using Turbofectin 8.0 (OriGene Technologies) with 0.125 μg GFP cDNA and 0.25 μg of 8A and 8C-8A(IL125).

Transfected cells were identified by GFP fluorescence and patch clamped in the whole-cell mode at room temperature using patch electrodes pulled from 1.5 mm outer-diameter silanized borosilicate microhematocrit tubes. Recordings were not performed on cells where access resistance was >2-fold that of the pipette resistance.

The composition of the control bath and pipette solutions used in these studies is shown in Table 3. Intracellular ionic strength was reduced by the removal of CsCl or cesium methanesulfonate from the patch pipette solution and replacement with sucrose to maintain solution osmolality.

Table 3
Composition of patch pipette and solutions.
Patch pipette solutionsBath solutions
ControlControlControlHypotonic
CsCl126 mM26 mM75 mM75 mM
Cesium methanesulfonate100 mM
MgSO42 mM2 mM5 mM5 mM
Ca-gluconate21 mM1 mM
ATP-Na22 mM2 mM
GTP-Na20.5 mM0.5 mM
Glutamine2 mM2 mM
EGTA1 mM1 mM
HEPES20 mM20 mM12 mM12 mM
Tris8 mM8 mM
CsOH12 mM12 mM
HCl2 mM2 mM
Glucose5 mM5 mM
Sucrose16 mM16 mM115 mM70 mM
pH*7.27.27.47.4
Osmolality275 mOsm275 mOsm300 mOsm250 mOsm
Ionic strength0.162 M0.162 M
  1. *

    The pH of patch pipette and bath solutions was adjusted with CsOH and HCl, respectively.

Whole-cell currents were measured with an Axopatch 200A (Axon Instruments) patch-clamp amplifier. Electrical connections to the patch-clamp amplifier were made using Ag/AgCl wires and 3 M KCl/agar bridges. Series resistance was compensated by >85% to minimize voltage errors. Data acquisition and analysis were performed using pClamp 10 software (Axon Instruments).

Changes in current amplitude were quantified using a voltage-ramping protocol. Membrane voltage was held at –30 mV throughout all experiments. Ramps were initiated by stepping membrane voltage to –100 mV and then ramping membrane voltage over 1 s to +100 mV. This was followed by a step back to –30 mV for 4 s before the ramp was repeated.

Relative anion permeability (PX/PCl) was measured from Cl- substitute-induced changes in reversal potential using a modified Goldman-Hodgkin-Katz equation (Voss et al., 2014). LRRC8A and 8C-8A(IL125) currents were activated with low ionic strength, 0.062 M, patch pipette solution, and by cell swelling induced by reducing bath osmolality to 250 mOsm. After stable current activation was achieved, changes in reversal potential were induced by replacing bath CsCl with either cesium glutathione or cesium lactobionate. Reversal potentials were corrected for anion-induced changes in liquid junction potentials.

Electrophysiological data are presented as means ± SEM. All patch-clamps studies were performed on at least two independently transfected groups of cells. n represents the number of patch-clamped cells from which currents were recorded. Statistical significance was determined using Student’s t test for unpaired means.

Figure preparation

Figures were prepared using Chimera (Pettersen et al., 2004), ChimeraX (Pettersen et al., 2021), and The PyMOL Molecular Graphics System (Version 2.0, Schrödinger, LLC). Calculation of the pore radii was performed using the software HOLE (Smart et al., 1996).

Data availability

Cryo-EM maps and atomic coordinates are deposited to the Electron Microscopy Data Bank (EMDB) and Protein Data Bank (PDB) databases. The accession codes are EMD-27770 and 8DXN for class 1; EMD-27771 and 8DXO for class 2; EMD-27772 and 8DXP for class 3; EMD-27773 and 8DXQ for class 4; EMD-27774 and 8DXR for class 5.

References

    1. Emsley P
    2. Cowtan K
    (2004) Coot: model-building tools for molecular graphics
    Acta Crystallographica. Section D, Biological Crystallography 60:2126–2132.
    https://doi.org/10.1107/S0907444904019158
  17. Book
    1. Hubbard S
    2. Thornton J
    (1993)
    NACCESS, Computer Program Department of Biochemistry Molecular Biology
    University College London Press.

Decision letter

  1. Merritt Maduke
    Reviewing Editor; Stanford University School of Medicine, United States
  2. Richard W Aldrich
    Senior Editor; The University of Texas at Austin, United States

Our editorial process produces two outputs: (i) public reviews designed to be posted alongside the preprint for the benefit of readers; (ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "A LRRC8 chimera with native functional properties is a heptamer with a large lipid-blocked pore" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Richard Aldrich as the Senior Editor. The reviewers have opted to remain anonymous.

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

Overall, the reviewers agree that the structural and functional characterization of the chimera is interesting from a channel architecture and assembly point of view but that the physiological relevance of the heptamer and the proposed lipid gating are not sufficiently supported by the data. As you will see below, the reviewers have recommended potential experiments to strengthen your conclusions. However, the consensus is that fully addressing these points would require extensive additional work, and therefore our recommendation is that a revised version of this manuscript should focus on the characterization of the chimera and not extend conclusions to physiological channel assemblies.

Essential revisions:

1) Physiological relevance:

Given that other LRRC8 channels are hexamers, the relevance of the heptameric assembly of the chimeric construct analyzed here should be experimentally evaluated. For example, the authors should identify structural elements in the heptameric assembly that account for physiological properties of native channels. Are there any functional channel properties that are explained by the heptameric assembly that are not compatible with the hexameric one (or viceversa?). The only evidence presented thus far is the increase in the permeability ratios of gluthatione and lactobionate between LLRC8A and 8C-8A(IL125). However, these effects are quite small, <50% increase between the two channels. Further, the LLRC8A homomeric pore is permeable to these ions. This seems at odds with its reported ~2 A radius, suggesting that the open homomeric channel is wider than the reported close conformation. Additional pharmacological and/or mutagenesis data is needed to corroborate this.

Native LRRC8 channels consist of the obligatory LRRC8A subunit and other paralogs (LRRC8B-E). The chimera in this manuscript is constructed on the basis of LRRC8C, with a swapped intracellular loop from LRRC8A that consists of 25 amino acids. It is interesting that the chimera recapitulates some native channel properties and forms a distinct heptameric channel, but this chimera is drastically different from native channels because the lack of an intact LRRC8A subunit. For instance, the selectivity filter arginine R103 in LRRC8A corresponds to L105 in LRRC8C. Thus, the pore properties of the chimera with all Leucine residues at this position would be very different from native LRRC8 channels. Therefore, the physiological relevance is unclear.

The authors propose that native heteromeric LRRC8 channels form heptamers based on the heptameric structure of the chimera. However, previous studies (Deneka et al. 2018, Nature) have shown that the functional heteromeric LRRC8A/C channels also form hexamers. In addition, two recent preprints corroborated the hexameric assembly of LRRC8A/C heteromeric channels.

2) Lipid characterization and function:

As shown in Figure 6b, the cryoEM density appears to be shown at very low threshold (as assessed by the micelle density), the assigned lipid-like density is similar to the micelle density. We are concerned that these are just noisy density that does not represent ordered/occupied lipids in the channel.

Further, the evidence supporting pore occlusion by lipids is weak. Based on the hydrophobic profile of the pore, it is difficult to visualize how the lipids can rearrange into the pore in two layers. Even if lipids are blocking the pore, authors cannot rule out that this is an artifact because of the purification process. Although they performed channel reconstitution in nanodiscs, the lipid-densities inside the pore can still be artifactual due to significant amounts of detergents that are used during the purification process. In addition, even when lipids-like densities have been found inside the pore of other channels, there is not functional evidence for supporting a gating role in large-pore channels and current hypotheses are still speculative. Thus, it is hard to visualize a gating mechanism by lipids in VRAC channels; if pore occlusion by lipids occurs in native cells, how the lipids can move in and out from the pore during a gating event? Overall, without proper data, it is not suitable to conclude that lipids are essential for normal channel gating and regulation of VRAC.

Experiments aimed at supporting the idea that this assembly is physiologically relevant are essential for supporting the authors' conclusion that "associated lipids are essential for normal channel gating and regulation." For example, the authors could show that the 8C-8A(IL125) channels co-purify with many more lipids than the homomeric 8A ones using lipidomics and/or native mass spec. Alternatively, functional experiments showing that lipids affect not only channel gating, but also ion permeation and selectivity would greatly strengthen the authors' conclusion.

3) Pore properties:

The authors argued that chimera structures reveal that the narrower part of pore is wide enough (~ 5 Å radius) for the permeation of molecules like glutathione and lactobionate. Further evidence is needed to support this conclusion, such as molecular modeling. The functional data alone are not sufficient to support that the new structure corresponds to the molecule-permeable conformation. In addition, the electrophysiological data are poorly described/presented. What is the reversal potential of non-transfected cells under the same experimental conditions?

A very interesting finding of this work is the heterogeneity observed in different protomers; specifically, a switch between a positive charged residue (K51) and a negative charged residue (D50) at the entrance of the pore. Considering that these residues are conserved in the LRRC8A and LRRC8E, it would be very interesting to evaluate how these residues affect gating and permeability properties of the chimera and LRRC8A channels.

An intriguing question that arises after reading this manuscript is related the mechanism blockade of DCPIB in VRAC channels. Previous structural work proposed that this blocker occludes the pore by interacting with residue R103, but this residue is only present in LRRC8A and B channels. LRRC8 C, D and E do not have an arginine at this position. In a previous manuscript by coauthors (Yamada et al., Am J Physiol Cell Physiol. 2021), the apparent affinity for the DCPIB is higher in the LRRC8 chimeric suggesting a different blockade mechanism to that previously proposed. How could the authors explain these results?

Presentation Clarity:

1) Pg. 4 The authors should tone down some of their claims. For example, they claim "…five high resolution (3.3-4.0 A) structures…" Given current standards, I do not think that 3.3-4 Å is not 'high resolution'. Similarly, they state "…allowing us to build an atomic model comprising most of the protein, except residue 60-94 in the ECD and the first 15 residues of the N-terminus." This is not a proper description of the data as the very large LLRD domains are not resolved in their structures, so that most of the protein could not actually be resolved.

2) Please find a different nomenclature than 'wide' and 'narrow' interface. I find these adjectives to be confusing and somewhat misleading descriptors of the structural features of LRRC8 channels. A wide/narrow interface suggest that the interface is wide or narrow. Instead, it is the separation between the protomers that is wide or narrow. Thus, the interface surface in the 'wide' arrangement is smaller than in the 'narrow' conformation, further confusing a reader. Maybe 'tight' and 'loose' would be better descriptors of the interfaces observed?

3) Please expand on the differences in the TMD regions of the different classes. When the authors say "the clustered arrangement is preserved" do the authors mean the 4 and 3 subunit arrangement? if yes, this is not apparent is the data shown in Supp Figure 5.

4) What are the consequences of the relative changes in protomer positions in the 5 classes? How do the interfaces change? An analysis of interaction surfaces in the 5 classes would be helpful to clarify what the differences are.

5) Are the lipid-like densities between protomers the same in the narrow and wide interfaces? Are the densities the same in all 7 protomers and 5 classes, despite the differences in interaction surfaces? This should be documented and discussed.

6) All structural figures show the intracellular domains at the top and the extracellular domains at the bottom. This is opposite to the conventional orientation of the plasma membrane of a cell.

7) Based on their current data, the authors suggest that formation of native heteromeric VRAC channels are heptamers. It is possible, however, that different LRRC8 isoforms can have different oligomeric composition (hexamer and heptamer) and heptamers can be found for LRRC8C homomeric channels. An example of how different isoforms from the same family can different oligomer are the CALHM channels. This should be discussed.

8) The manuscript lacks proper data description/analysis of the structural basis for an heptameric assembly. For example, what are the interactions between subunits in the chimera? What key differences are observed with the already known LRRC8 hexameric structures?

9) Bands in the native page blot shown in Figure Sup. 1, are not clear (especially the chimera). Do the chimeric channels form exclusively heptamers or also can form another oligomeric state?

10) Figure Supplementary 2 and 7. FSC plot quality is quite low and almost unreadable. In addition, the scale of Y-axis in Figure Sup. 2 is apparently wrong.

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

Thank you for resubmitting your work entitled "A LRRC8 chimera with native functional properties is a heptamer with a large-diameter lipid-blocked pore" for further consideration by eLife. Your revised article has been evaluated by Richard Aldrich (Senior Editor) and a Reviewing Editor.

This new version of the manuscript has considerably toned down the conclusions from the resolved structures and incorporated new data, making it more focused on the description/characterization of this novel structure. There are only a few remaining issues that need to be addressed, as outlined below:

2) The evidence presented does not definitively demonstrate that the pore is completely occluded by lipid, as suggested in the title of this manuscript and the Result section (page 9, lines 209-211). We request that the title be changed and the Results section reworded in order to avoid potential confusion in the field.

3) Additional thoughts on the point-by-point letter concerning lipid densities:

Author statement: "To clarify the representation of the lipid-like densities, we modified Figure 6 (Figure 7 in the revised manuscript) as follows. The threshold of the cryo-EM map was increased. We aligned the panels showing the cryo-EM density, hydrophobicity plot, and electrostatic surface charges side by side to reveal how the lipid-like density aligns with the chemical properties of the inner surface of the pore. We used PyMol to prepare hydrophobicity and electrostatic surface potential figures in the revised manuscript."

From the figures, it is still unclear how the lipid-like densities align with the hydrophobicity/electrostatic profile of the pore. Surface pore properties are not correlative with the formation of two lipid layers. Also, a better representation of lipid-like densities between subunits is needed. A zoomed image could help to show this better. If authors can identify a conserved hydrophobic pocket for these lipid interactions, that could be highlighted. This point is not critical, but the manuscript may benefit by providing further information on the residues interacting with lipid-like densities between subunits.

4) You have included new figures highlighting the tight and loose interfaces in ECD, TMD, and ICD (Figure 3 —figure supplement 1-3). This is interesting, but an accurate description of the main interactions is not found in the main text.

5) You could elaborate on the description of D50/K51 rearrangements in the Result section. You state, "When K51 points toward the pore, it creates a groove between the subunits (Figure 4D)". Although this statement seems correct in the current context, the sentence confuses the overall interpretation of D50/K51 rearrangements in LRRC8C channels because the statement only applies for structures of class 1 and 3 but not for classes 2, 4, and 5. For example, the structure of class 2 only has one K51 side chain pointing toward the pore (Figure 5 – Figure sup2). Still, it keeps two loose interfaces anyway (Figure 5C), which suggests that D50/K51 rearrangement is not necessarily coupled with the loose interface as one could interpret from the text.

6) Page 4, Line 80-82 "The oligomeric state directly impacts pore size, pharmacology, and permeability to large anionic solutes. Our results also suggest that associated membrane lipids may be involved in channel gating and regulation".

This work did not evaluate if the oligomeric state affects the pharmacology of the channels. As discussed in the point-by-point letter, current evidence supports the hypothesis that pore size could impact pharmacology, but this could also be related to the intrinsic properties of LRRC8C. Also, in view that there is no evidence supporting the role of lipids in channel function, the above sentence is speculative. Thus, this entire sentence should be restricted to the Discussion section only.

7) Figure 1: Please Indicate in the figure legend what the symbols shown below the alignment in panel A (*. :) mean.

8) Please specify in the figure legends what class is shown in figures 2, 3, and 4. Are these the class 1 structure?

9) Figure 4: The arrow showing rotation between structures of panels D and E seems to be in the wrong direction. Please check.

10) Figure legend 7: "B) A sliced view of the unsharpened cryo-EM map (grey, transparent) with the ribbon representation of the 8C-8A(IL125) structure".

The ribbon representation is not shown in the new version of the manuscript. Please, check.

11) Table 3: The concentration unit for sucrose concentration is missing. Please check.

https://doi.org/10.7554/eLife.82431.sa1

Author response

Essential revisions:

1) Physiological relevance:

Given that other LRRC8 channels are hexamers, the relevance of the heptameric assembly of the chimeric construct analyzed here should be experimentally evaluated. For example, the authors should identify structural elements in the heptameric assembly that account for physiological properties of native channels. Are there any functional channel properties that are explained by the heptameric assembly that are not compatible with the hexameric one (or viceversa?). The only evidence presented thus far is the increase in the permeability ratios of gluthatione and lactobionate between LLRC8A and 8C-8A(IL125). However, these effects are quite small, <50% increase between the two channels. Further, the LLRC8A homomeric pore is permeable to these ions. This seems at odds with its reported ~2 A radius, suggesting that the open homomeric channel is wider than the reported close conformation. Additional pharmacological and/or mutagenesis data is needed to corroborate this.

Native LRRC8 channels consist of the obligatory LRRC8A subunit and other paralogs (LRRC8B-E). The chimera in this manuscript is constructed on the basis of LRRC8C, with a swapped intracellular loop from LRRC8A that consists of 25 amino acids. It is interesting that the chimera recapitulates some native channel properties and forms a distinct heptameric channel, but this chimera is drastically different from native channels because the lack of an intact LRRC8A subunit. For instance, the selectivity filter arginine R103 in LRRC8A corresponds to L105 in LRRC8C. Thus, the pore properties of the chimera with all Leucine residues at this position would be very different from native LRRC8 channels. Therefore, the physiological relevance is unclear.

The authors propose that native heteromeric LRRC8 channels form heptamers based on the heptameric structure of the chimera. However, previous studies (Deneka et al. 2018, Nature) have shown that the functional heteromeric LRRC8A/E channels also form hexamers. In addition, two recent preprints corroborated the hexameric assembly of LRRC8A/C heteromeric channels.

It is unclear at present whether native LRRC8 channels are hexamers, heptamers or whether they exist in multiple conformations including higher order assemblies. We have reduced the discussion of the physiological relevance of the heptameric assembly and instead point out the need for much additional study to understand the configuration of native channels.

We agree that additional pharmacological and mutagenesis data are needed to address this important problem. Some of these data will be presented at the upcoming Biophysical Society meeting and will be the focus of a forthcoming manuscript. However, as we discussed in the current manuscript, we have shown previously that the DCPIB pharmacology of hexameric LRRC8A channels is grossly abnormal compared to native channels and 8A+8C heteromers. DCPIB pharmacology of 8A hexamers is also dramatically altered by mutation of R103 to phenylalanine. R103 forms the narrow constriction of 8A hexameric channels. In contrast, 8A+8C heteromers are unaffected by the R103F mutation.

The two new manuscripts that have appeared on bioRxiv show clear hexameric structures of 8A+8C heteromers. However, the 8A:8C stoichiometries differ strikingly for the two labs. The Dutzler lab shows a 4A:2C stoichiometry, while the Brohawn lab shows a 5A:1C stoichiometry. It is important to note that four labs have shown that 8A functions as a dominant-negative. High levels of 8A expression completely block native LRRC8 currents and dramatically reduce heterologously expressed channel activity. Thus, one can well imagine that high levels of 8A expression drive the formation of non-native channel configurations. This is an important problem that must be addressed in order to understand the configurations and stoichiometries of native channels.

Finally, it is important to note that the Dutzler lab states in their bioRxiv manuscript that it is unclear how a hexameric configuration can give rise to a large pore channel permeable to diverse organic solutes.

2) Lipid characterization and function:

As shown in Figure 6b, the cryoEM density appears to be shown at very low threshold (as assessed by the micelle density), the assigned lipid-like density is similar to the micelle density. We are concerned that these are just noisy density that does not represent ordered/occupied lipids in the channel.

Further, the evidence supporting pore occlusion by lipids is weak. Based on the hydrophobic profile of the pore, it is difficult to visualize how the lipids can rearrange into the pore in two layers. Even if lipids are blocking the pore, authors cannot rule out that this is an artifact because of the purification process. Although they performed channel reconstitution in nanodiscs, the lipid-densities inside the pore can still be artifactual due to significant amounts of detergents that are used during the purification process. In addition, even when lipids-like densities have been found inside the pore of other channels, there is not functional evidence for supporting a gating role in large-pore channels and current hypotheses are still speculative. Thus, it is hard to visualize a gating mechanism by lipids in VRAC channels; if pore occlusion by lipids occurs in native cells, how the lipids can move in and out from the pore during a gating event? Overall, without proper data, it is not suitable to conclude that lipids are essential for normal channel gating and regulation of VRAC.

Experiments aimed at supporting the idea that this assembly is physiologically relevant are essential for supporting the authors' conclusion that "associated lipids are essential for normal channel gating and regulation." For example, the authors could show that the 8C-8A(IL125) channels co-purify with many more lipids than the homomeric 8A ones using lipidomics and/or native mass spec. Alternatively, functional experiments showing that lipids affect not only channel gating, but also ion permeation and selectivity would greatly strengthen the authors' conclusion.

We agree with the reviewers that additional experiments are required to conclude that these densities are lipid molecules and that lipids are essential for normal channel gating. Therefore, we have modified the text and presented our observation as a possible mechanism that requires further experimental validation.

As pointed out, the cryo-EM density we assign as lipid-like density is not as strong as the protein densities. However, we think this would be expected since most lipid molecules within the pore would not interact with the protein residues and be highly flexible, resulting in weaker cryo-EM densities similar to the ones for the detergent micelle. Moreover, the "lipid-like densities" in the nanodisc structures are highly similar, both in intensity and shape, to the lipid densities observed for the lipid bilayer around the protein within the nanodisc, as shown in Figure 7—figure supplement 1 (Supp. Figure 7 in the original submission). In agreement with our observations, in a recent preprint reporting the hexameric structure of LRRC8A-LRRC8C (Kern et al. 2022), ordered lipid molecules are observed on the extracellular side of the pore opening, aligning well with the densities we observed. It is plausible that the narrower opening in the hexameric structure compared to that of the heptameric structure led to the visualization of the ordered lipid molecules.

To clarify the representation of the lipid-like densities, we modified Figure 6 (Figure 7 in the revised manuscript) as follows. The threshold of the cryo-EM map was increased. We aligned the panels showing the cryo-EM density, hydrophobicity plot, and electrostatic surface charges side by side to reveal how the lipid-like density aligns with the chemical properties of the inner surface of the pore. We used PyMol to prepare hydrophobicity and electrostatic surface potential figures in the revised manuscript.

3) Pore properties:

The authors argued that chimera structures reveal that the narrower part of pore is wide enough (~ 5 Å radius) for the permeation of molecules like glutathione and lactobionate. Further evidence is needed to support this conclusion, such as molecular modeling. The functional data alone are not sufficient to support that the new structure corresponds to the molecule-permeable conformation. In addition, the electrophysiological data are poorly described/presented. What is the reversal potential of non-transfected cells under the same experimental conditions?

Our intent was to show that there are clear differences in the relative permeabilities of these two solutes in the 8C-8A(IL125) heptameric channel when compared directly to 8A hexameric channels. Our results demonstrate that the relative permeability to these solutes is significantly higher in the chimera. This is consistent with the larger pore diameter of the heptameric chimera versus the 8A hexamer. This section has now been rewritten to improve clarity.

Countless labs have measured relative permeabilites of native LRRC8/VRAC channels to a host of diverse solutes. The estimated pore diameter of native channels is similar to that of 8C-8A(IL125) heptameric channels. In contrast, the narrow pore diameters of 8A hexamer as well as the 8A+8C heteromers described in the recent manuscripts from the Brohawn and Dutzler labs are well below that estimated for native channels. As noted above, the Dutzler lab states in their bioRxiv manuscript that it is unclear how a hexameric configuration can give rise to a large pore channel permeable to diverse organic solutes.

A very interesting finding of this work is the heterogeneity observed in different protomers; specifically, a switch between a positive charged residue (K51) and a negative charged residue (D50) at the entrance of the pore. Considering that these residues are conserved in the LRRC8A and LRRC8E, it would be very interesting to evaluate how these residues affect gating and permeability properties of the chimera and LRRC8A channels.

To evaluate the effect of the switch between K51 and D50 at the entrance of the pore, we modified Figure 4 (Figure 3 in the original manuscript) by adding 3 new panels. These new panels show the surface representation of the protein, highlighting the differences in the spacing between the subunits due to this switch. We also modified the text accordingly.

An intriguing question that arises after reading this manuscript is related the mechanism blockade of DCPIB in VRAC channels. Previous structural work proposed that this blocker occludes the pore by interacting with residue R103, but this residue is only present in LRRC8A and B channels. LRRC8 C, D and E do not have an arginine at this position. In a previous manuscript by coauthors (Yamada et al., Am J Physiol Cell Physiol. 2021), the apparent affinity for the DCPIB is higher in the LRRC8 chimeric suggesting a different blockade mechanism to that previously proposed. How could the authors explain these results?

We believe that the larger pore diameter of native LRRC8 channels, heterologously expressed 8A+8C heteromers, and the 8C-8A(IL125) heptameric chimera is the simplest explanation for the difference. Notably, the Hill coefficient for DCPIB is close to 1 for narrow pore 8A hexameric channels suggesting that a single DCPIB molecule blocks the pore. This would be consistent with the cryo-EM structure published by Kern et al. showing that the pore is occluded by a single DCPIB molecule at its narrowest constriction formed by R103. In contrast, the Hill coefficients for the chimera and native channels are >2, suggesting multiple cooperative DCPIB binding sites.

Also of note is the effect of R103 mutations on DCPIB inhibition. Mutation of R103 to phenylalanine dramatically reduces DCPIB inhibition of narrow pore 8A hexamers. This is predicted due to the further narrowing of the already narrow pore by the phenylalanine. In contrast, R103F has no effect on DCPIB inhibition of 8A(R103F)+8C heteromers (Yamada et al.). Phenylalanine mutation of the equivalent residue, L105, has no effect on DCPIB inhibition of the 8C-8A(IL125) heptameric chimera or on 8A(R103F)+8C(L105F) heteromers. These latter data have been published in abstract form and are the subject of a forthcoming manuscript.

Finally, DCPIB inhibition is strongly voltage-dependent in 8A hexameric channels, whereas it is voltage insensitive in native channels, 8A+8C heteromers, and 8C-8A(IL125) heptameric chimeras. The voltage sensitivity of narrow pore 8A hexamers is most readily explained by the DCPIB block at the channel constriction site formed by R103.

Presentation Clarity:

1) Pg. 4 The authors should tone down some of their claims. For example, they claim "…five high resolution (3.3-4.0 A) structures…" Given current standards, I do not think that 3.3-4 Å is not 'high resolution'. Similarly, they state "…allowing us to build an atomic model comprising most of the protein, except residue 60-94 in the ECD and the first 15 residues of the N-terminus." This is not a proper description of the data as the very large LLRD domains are not resolved in their structures, so that most of the protein could not actually be resolved.

We have changed the text to tone down the claims as follows:

"….Therefore, the final reconstructions were done without enforcing any symmetry and resulted in five structures with resolutions in the range of 3.3 to 4.0 Å (Figures 1a,b; Supplementary Figure 2-3)."

"…The cryo-EM maps for the ECD and TMD revealed high resolution features allowing us to build an atomic model comprising most of the ECD and TMD except residues 60-94 in the ECD and the first 15 residues of the N-terminus…."

2) Please find a different nomenclature than 'wide' and 'narrow' interface. I find these adjectives to be confusing and somewhat misleading descriptors of the structural features of LRRC8 channels. A wide/narrow interface suggest that the interface is wide or narrow. Instead, it is the separation between the protomers that is wide or narrow. Thus, the interface surface in the 'wide' arrangement is smaller than in the 'narrow' conformation, further confusing a reader. Maybe 'tight' and 'loose' would be better descriptors of the interfaces observed?

We have changed the nomenclature of "wide" and "narrow" interfaces to "loose" and "tight" interfaces, respectively, on the text and the figures.

3) Please expand on the differences in the TMD regions of the different classes. When the authors say "the clustered arrangement is preserved" do the authors mean the 4 and 3 subunit arrangement? if yes, this is not apparent is the data shown in Supp Figure 5.

We added a panel to Figure 5 (Figure 4 in the original manuscript) to compare the TMD structures in all five classes. We used reference points to compare the separation between the neighboring subunits, along with the comparison of the ICD structures. We also added a new panel to Figure 5-supplement 1 (Supp. Figure 5 in the original manuscript), showing the alignment of all five structures at the TMD to compare the structures. We removed the phrase "the clustered arrangement is preserved" and emphasized the similarity to the differences in subunit separation at the loose and tight interfaces to that of ICD.

4) What are the consequences of the relative changes in protomer positions in the 5 classes? How do the interfaces change? An analysis of interaction surfaces in the 5 classes would be helpful to clarify what the differences are.

The main effect of the changes in the protomer positions is the separation between the subunits at the TMD and ICD. The changes are more robust at the loose interfaces and likely affect how the interior and outside of the pore are connected through the gaps between the subunits. The new Figure 5 (Figure 4 in the original manuscript) shows the changes in subunit separation at the ICD and TMD in the five classes.

5) Are the lipid-like densities between protomers the same in the narrow and wide interfaces? Are the densities the same in all 7 protomers and 5 classes, despite the differences in interaction surfaces? This should be documented and discussed.

The densities at the loose cytoplasmic side of the loose interfaces are considerably weaker than the ones at the tight interfaces. We changed Figure 7A (Figure 6A in the original manuscript) to show this difference and edited the text accordingly.

6) All structural figures show the intracellular domains at the top and the extracellular domains at the bottom. This is opposite to the conventional orientation of the plasma membrane of a cell.

All the figures are changed to place the extracellular domains at the top and the intracellular domains at the bottom.

7) Based on their current data, the authors suggest that formation of native heteromeric VRAC channels are heptamers. It is possible, however, that different LRRC8 isoforms can have different oligomeric composition (hexamer and heptamer) and heptamers can be found for LRRC8C homomeric channels. An example of how different isoforms from the same family can different oligomer are the CALHM channels. This should be discussed.

We included the following paragraphs in the manuscript to discuss the possibility of multiple oligomeric states for LRRC8 channels.

8) The manuscript lacks proper data description/analysis of the structural basis for an heptameric assembly. For example, what are the interactions between subunits in the chimera? What key differences are observed with the already known LRRC8 hexameric structures?

To expand on the description of the subunit interfaces, we included 3 additional supplementary figures (Figure 3-supplement 1-3) showing detailed view of the interfaces between the subunits at the loose and tight interfaces of the ECD, TMD, and ICD. We also calculated the buried surface areas between the subunits for each domain in the class 1 structure.

“Two recent studies have defined high-resolution cryo-EM structures of LRRC8A/LRRC8C heteromeric channels (Kern et al., 2022; Rutz et al., 2022). Both studies demonstrated that the heteromeric channels could adopt a hexameric conformation with a limiting pore radius of 2-3 Å, similar to that of LRRC8A hexamers. However, these channels either had 5:1 (Kern et al., 2022) or 4:2 (Rutz et al., 2022) LRRC8A:LRRC8C stoichiometries, indicating that LRRC8A:LRRC8C heteromers can adopt multiple oligomeric forms.

Taken together, existing cryo-EM structural data suggest that native VRAC/LRRC8 channels may exist in multiple oligomeric conformations, as has been reported for CALHM channels. It will be critical to relate the functional properties of various channel oligomeric conformations to those of native VRAC/LRRC8 channels. It will also be important to define the role of LRRC8A in channel assembly and conformation. Four groups have shown that LRRC8A has a dominant-negative function. Overexpression of LRRC8A suppresses endogenous (Qiu et al., 2014; Syeda et al., 2016; Voss et al., 2014) and heterologously expressed (Yamada et al., 2016) VRAC/LRRC8 currents. LRRC8A protein levels may therefore impact channel assembly and function.”

9) Bands in the native page blot shown in Figure Sup. 1, are not clear (especially the chimera). Do the chimeric channels form exclusively heptamers or also can form another oligomeric state?

The Flag tag on the constructs appear to cause smeary bands on the native PAGE blots, but 8A and the chimera run separately in all experiments we have done. We also do not observe any classes that suggested other possible stoichometries in our cryo-EM experiments and obtain single peaks in the size exclusion chromatography experiments suggesting that the chimeric channels are mainly heptameric.

10) Figure Supplementary 2 and 7. FSC plot quality is quite low and almost unreadable. In addition, the scale of Y-axis in Figure Sup. 2 is apparently wrong.

We enlarged the FSC plots so that they are more easily readable. We also corrected the labeling of the Y-axis in Supplemental Figure 2.

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

2) The evidence presented does not definitively demonstrate that the pore is completely occluded by lipid, as suggested in the title of this manuscript and the Result section (page 9, lines 209-211). We request that the title be changed and the Results section reworded in order to avoid potential confusion in the field.

We changed the title to "Cryo-EM structures of a LRRC8 chimera with native functional properties reveal heptameric assembly" and reworded the Results section. It should now be clear to the reader that further experiments are required to verify the presence of pore-blocking lipid molecules.

3) Additional thoughts on the point-by-point letter concerning lipid densities:

Author statement: "To clarify the representation of the lipid-like densities, we modified Figure 6 (Figure 7 in the revised manuscript) as follows. The threshold of the cryo-EM map was increased. We aligned the panels showing the cryo-EM density, hydrophobicity plot, and electrostatic surface charges side by side to reveal how the lipid-like density aligns with the chemical properties of the inner surface of the pore. We used PyMol to prepare hydrophobicity and electrostatic surface potential figures in the revised manuscript."

From the figures, it is still unclear how the lipid-like densities align with the hydrophobicity/electrostatic profile of the pore. Surface pore properties are not correlative with the formation of two lipid layers. Also, a better representation of lipid-like densities between subunits is needed. A zoomed image could help to show this better. If authors can identify a conserved hydrophobic pocket for these lipid interactions, that could be highlighted. This point is not critical, but the manuscript may benefit by providing further information on the residues interacting with lipid-like densities between subunits.

We have made several changes to Figure 7 and the text titled "Interaction of lipids with the 8C-8A(IL125)" on page 9. Specifically;

– We changed the color of lipid-like densities between the subunits to yellow from red to avoid potential confusion with the red color used to show the negative charge.

– We added a new panel (panel B) showing the zoomed view of one of the lipid-like densities between the subunits. In this figure, the cryo-EM map was shown as a grey mesh. The density was not trimmed to a particular region in this figure so that the readers can judge the quality of the lipid density relative to the surrounding region. Although we did not model any ligands to the deposited structures, we placed a phospholipid into the density for illustrative purposes. Select residues in proximity to the phospholipid molecule were shown.

– In the panel showing the surface properties of the pore lining residues (panels C and D in the revised manuscript), we included hypothetical lipid molecules that would form the bilayer around the TMD to emphasize how surface properties align with the membrane. The positioning of the membrane boundaries around the protein is calculated using PPM 2.0 (https://opm.phar.umich.edu/ppm_server) incorporated into CHARMM-GUI (https://www.charmm-gui.org/?doc=input/membrane.bilayer).

– We changed the coloring of the hydrophobicity from red to orange to avoid potential confusion with the red color used to show electronegative surface charge.

– We added a new panel showing a hypothetical representation of the pore with lipid molecules to help with the visualization of how the lipid-like densities align with the surface properties of the pore.

– We modified the text to clarify the correlation between surface properties and bilayer formation and to incorporate the description of new information included in the figures.

4) You have included new figures highlighting the tight and loose interfaces in ECD, TMD, and ICD (Figure 3 —figure supplement 1-3). This is interesting, but an accurate description of the main interactions is not found in the main text.

We expanded on the description of the intersubunit interactions refering to Figure 3—figure supplement 1-3 in the "8C-8A(IL125) chimeras form heptameric channels" section in page 6.

5) You could elaborate on the description of D50/K51 rearrangements in the Result section. You state, "When K51 points toward the pore, it creates a groove between the subunits (Figure 4D)". Although this statement seems correct in the current context, the sentence confuses the overall interpretation of D50/K51 rearrangements in LRRC8C channels because the statement only applies for structures of class 1 and 3 but not for classes 2, 4, and 5. For example, the structure of class 2 only has one K51 side chain pointing toward the pore (Figure 5 – Figure sup2). Still, it keeps two loose interfaces anyway (Figure 5C), which suggests that D50/K51 rearrangement is not necessarily coupled with the loose interface as one could interpret from the text.

We modified the last paragraphs of "8C-8A(IL125) chimeras form heptameric channels (page 7)" and "Structural heterogeneity of 8C-8A(IL125) chimeras (page 8)" sections to clarify that the D50/K51 orientation and the presence of loose interface do exhibit variability in different classes and are not necessarily correlated.

6) Page 4, Line 80-82 "The oligomeric state directly impacts pore size, pharmacology, and permeability to large anionic solutes. Our results also suggest that associated membrane lipids may be involved in channel gating and regulation".

This work did not evaluate if the oligomeric state affects the pharmacology of the channels. As discussed in the point-by-point letter, current evidence supports the hypothesis that pore size could impact pharmacology, but this could also be related to the intrinsic properties of LRRC8C. Also, in view that there is no evidence supporting the role of lipids in channel function, the above sentence is speculative. Thus, this entire sentence should be restricted to the Discussion section only.

We removed these sentences from the introduction.

7) Figure 1: Please Indicate in the figure legend what the symbols shown below the alignment in panel A (*. :) mean.

We added the description of the symbols to the figure legend.

8) Please specify in the figure legends what class is shown in figures 2, 3, and 4. Are these the class 1 structure?

We used the class 1 structure in these figures and clarified this in the figure legends.

9) Figure 4: The arrow showing rotation between structures of panels D and E seems to be in the wrong direction. Please check.

We checked the figure to make sure the arrow direction was correct.

10) Figure legend 7: "B) A sliced view of the unsharpened cryo-EM map (grey, transparent) with the ribbon representation of the 8C-8A(IL125) structure".

The ribbon representation is not shown in the new version of the manuscript. Please, check.

We checked the figure and confirmed that the legend is accurate. The ribbon representation is present in the figure.

11) Table 3: The concentration unit for sucrose concentration is missing. Please check.

We added the concentration unit for sucrose.

https://doi.org/10.7554/eLife.82431.sa2

Article and author information

Author details

  1. Hirohide Takahashi

    1. Department of Molecular Physiology and Biophysics, School of Medicine, Vanderbilt University, Nashville, United States
    2. Center for Structural Biology, Vanderbilt University, Nashville, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2553-8806

  2. Toshiki Yamada

    Department of Anesthesiology, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared

  3. Jerod S Denton

    1. Department of Anesthesiology, Vanderbilt University Medical Center, Nashville, United States
    2. Department of Pharmacology, School of Medicine, Vanderbilt University, Nashville, United States
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Project administration, Writing – review and editing
    Competing interests
    No competing interests declared

  4. Kevin Strange

    Department of Anesthesiology, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Writing – original draft, Project administration, Writing – review and editing
    Competing interests
    is cofounder and principal scientist of Revidia Therapeutics, Inc

  5. Erkan Karakas

    1. Department of Molecular Physiology and Biophysics, School of Medicine, Vanderbilt University, Nashville, United States
    2. Center for Structural Biology, Vanderbilt University, Nashville, United States
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Visualization, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    erkan.karakas@vanderbilt.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6552-3185

Funding

National Institute of Diabetes and Digestive and Kidney Diseases (DK51610)

  • Jerod S Denton

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

Acknowledgements

We thank Dr Kunpeng Lee at Case Western Reserve University and Melissa Chambers and Scott Collier at the Cryo-EM facility at Vanderbilt University for cryo-EM data collection. This work was conducted in part using the CPU and GPU resources of the Advanced Computing Center for Research and Education (ACCRE) at Vanderbilt University. We used the DORS storage system supported by the National Institute of Health (NIH) (S10 RR031634 to Jarrod Smith). This work was supported by the National Institute of Diabetes, Digestive, and Kidney Diseases Grant R01 DK51610 to JSD.

Senior Editor

  1. Richard W Aldrich, The University of Texas at Austin, United States

Reviewing Editor

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

Version history

  1. Preprint posted: July 28, 2022 (view preprint)
  2. Received: August 3, 2022
  3. Accepted: March 9, 2023
  4. Accepted Manuscript published: March 10, 2023 (version 1)
  5. Version of Record published: March 28, 2023 (version 2)

Copyright

© 2023, Takahashi 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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  1. Hirohide Takahashi
  2. Toshiki Yamada
  3. Jerod S Denton
  4. Kevin Strange
  5. Erkan Karakas
(2023)
Cryo-EM structures of an LRRC8 chimera with native functional properties reveal heptameric assembly
eLife 12:e82431.
https://doi.org/10.7554/eLife.82431

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    Elliott L Paine, Jack J Skalicky ... Wesley I Sundquist
    Research Advance

    The Endosomal Sorting Complexes Required for Transport (ESCRT) machinery mediates the membrane fission step that completes cytokinetic abscission and separates dividing cells. Filaments composed of ESCRT-III subunits constrict membranes of the intercellular bridge midbody to the abscission point. These filaments also bind and recruit cofactors whose activities help execute abscission and/or delay abscission timing in response to mitotic errors via the NoCut/Abscission checkpoint. We previously showed that the ESCRT-III subunit IST1 binds the cysteine protease CAPN7 (Calpain-7) and that CAPN7 is required for both efficient abscission and NoCut checkpoint maintenance (Wenzel et al., 2022). Here, we report biochemical and crystallographic studies showing that the tandem MIT domains of CAPN7 bind simultaneously to two distinct IST1 MIT interaction motifs. Structure-guided point mutations in either CAPN7 MIT domain disrupted IST1 binding in vitro and in cells, and depletion/rescue experiments showed that the CAPN7-IST1 interaction is required for: 1) CAPN7 recruitment to midbodies, 2) efficient abscission, and 3) NoCut checkpoint arrest. CAPN7 proteolytic activity is also required for abscission and checkpoint maintenance. Hence, IST1 recruits CAPN7 to midbodies, where its proteolytic activity is required to regulate and complete abscission.

    1. Biochemistry and Chemical Biology
    2. Structural Biology and Molecular Biophysics

    Vibrio cholerae's ToxRS bile sensing system

    Nina Gubensäk, Theo Sagmeister ... Tea Pavkov-Keller
    Research Article

    The seventh pandemic of the diarrheal cholera disease, which began in 1960, is caused by the Gram-negative bacterium Vibrio cholerae. Its environmental persistence provoking recurring sudden outbreaks is enabled by V. cholerae's rapid adaption to changing environments involving sensory proteins like ToxR and ToxS. Located at the inner membrane, ToxR and ToxS react to environmental stimuli like bile acid, thereby inducing survival strategies e.g. bile resistance and virulence regulation. The presented crystal structure of the sensory domains of ToxR and ToxS in combination with multiple bile acid interaction studies, reveals that a bile binding pocket of ToxS is only properly folded upon binding to ToxR. Our data proposes an interdependent functionality between ToxR transcriptional activity and ToxS sensory function. These findings support the previously suggested link between ToxRS and VtrAC-like co-component systems. Besides VtrAC, ToxRS is now the only experimentally determined structure within this recently defined superfamily, further emphasizing its significance. In-depth analysis of the ToxRS complex reveals its remarkable conservation across various Vibrio species, underlining the significance of conserved residues in the ToxS barrel and the more diverse ToxR sensory domain. Unravelling the intricate mechanisms governing ToxRS's environmental sensing capabilities, provides a promising tool for disruption of this vital interaction, ultimately inhibiting Vibrio's survival and virulence. Our findings hold far-reaching implications for all Vibrio strains that rely on the ToxRS system as a shared sensory cornerstone for adapting to their surroundings.

    1. Microbiology and Infectious Disease
    2. Structural Biology and Molecular Biophysics

    A pH-sensitive switch activates virulence in Salmonella

    Dasvit Shetty, Linda J Kenney
    Research Article Updated

    The transcriptional regulator SsrB acts as a switch between virulent and biofilm lifestyles of non-typhoidal Salmonella enterica serovar Typhimurium. During infection, phosphorylated SsrB activates genes on Salmonella Pathogenicity Island-2 (SPI-2) essential for survival and replication within the macrophage. Low pH inside the vacuole is a key inducer of expression and SsrB activation. Previous studies demonstrated an increase in SsrB protein levels and DNA-binding affinity at low pH; the molecular basis was unknown (Liew et al., 2019). This study elucidates its underlying mechanism and in vivo significance. Employing single-molecule and transcriptional assays, we report that the SsrB DNA-binding domain alone (SsrBc) is insufficient to induce acid pH-sensitivity. Instead, His12, a conserved residue in the receiver domain confers pH sensitivity to SsrB allosterically. Acid-dependent DNA binding was highly cooperative, suggesting a new configuration of SsrB oligomers at SPI-2-dependent promoters. His12 also plays a role in SsrB phosphorylation; substituting His12 reduced phosphorylation at neutral pH and abolished pH-dependent differences. Failure to flip the switch in SsrB renders Salmonella avirulent and represents a potential means of controlling virulence.