Structural insights into the voltage and phospholipid activation of the mammalian TPC1 channel


The organellar two-pore channel (TPC) functions as a homodimer, in which each subunit contains two homologous Shaker-like six-transmembrane (6-TM)-domain repeats1. TPCs belong to the voltage-gated ion channel superfamily2 and are ubiquitously expressed in animals and plants3,4. Mammalian TPC1 and TPC2 are localized at the endolysosomal membrane, and have critical roles in regulating the physiological functions of these acidic organelles5,6,7. Here we present electron cryo-microscopy structures of mouse TPC1 (MmTPC1)—a voltage-dependent, phosphatidylinositol 3,5-bisphosphate (PtdIns(3,5)P2)-activated Na+-selective channel—in both the apo closed state and ligand-bound open state. Combined with functional analysis, these structures provide comprehensive structural insights into the selectivity and gating mechanisms of mammalian TPC channels. The channel has a coin-slot-shaped ion pathway in the filter that defines the selectivity of mammalian TPCs. Only the voltage-sensing domain from the second 6-TM domain confers voltage dependence on MmTPC1. Endolysosome-specific PtdIns(3,5)P2 binds to the first 6-TM domain and activates the channel under conditions of depolarizing membrane potential. Structural comparisons between the apo and PtdIns(3,5)P2-bound structures show the interplay between voltage and ligand in channel activation. These MmTPC1 structures reveal lipid binding and regulation in a 6-TM voltage-gated channel, which is of interest in light of the emerging recognition of the importance of phosphoinositide regulation of ion channels.


TPC1 and TPC2 represent two major subfamilies of mammalian TPC channels and their functions are associated with various physiological processes, including hair pigmentation8,9,10, autophagy regulation11,12, blood vessel formation13, acrosome reaction in sperm14, mTOR-dependent nutrient sensing15, lipid metabolism16 and Ebola virus infection17, to name a few. Mammalian TPCs were initially suggested to mediate nicotinic acid adenine dinucleotide phosphate (NAADP)-dependent calcium release from endolysosomes18,19,20. However, several recent studies have demonstrated that mammalian TPCs are Na+-selective channels activated by endolysosome-specific PtdIns(3,5)P2 rather than NAADP15,21. The dual regulation of TPC2 by both PtdIns(3,5)P2 and NAADP has also been reported22. Distinct from TPC2, mammalian TPC1 activation is voltage-dependent, conferring electrical excitability to the endolysosome23,24. The atomic structure of a plant TPC1 from Arabidopsis thaliana (AtTPC1) was recently determined by X-ray crystallography, revealing the overall architecture of the TPC family25,26. However, mammalian TPCs share low sequence identity with their plant counterparts (Extended Data Fig. 1) and exhibit different gating and selectivity properties. Here we present the structural and functional analysis of MmTPC1.

When overexpressed in HEK293 cells, some MmTPC1 channels are trafficked to the plasma membrane, enabling us to directly measure channel activity by patching the plasma membrane (Extended Data Fig. 2 and Methods). In brief, MmTPC1 activation requires both membrane depolarization and the PtdIns(3,5)P2 ligand (Extended Data Fig. 2b, c). The voltage activation of MmTPC1 is modulated by endolysosomal luminal pH23, and a lower pH shifts voltage activation towards a more positive potential (Extended Data Fig. 2d, e). In our recordings, MmTPC1 exhibits higher selectivity for Na+ than K+ and Ca2+ (Extended Data Fig. 2f, g); this is different from plant TPC1, which is non-selective27,28.

MmTPC1 structures were determined in the presence and absence of PtdIns(3,5)P2 to a resolution of 3.2 and 3.4 Å, respectively, using single particle electron cryo-microscopy (cryo-EM) (Fig. 1, Extended Data Figs 3, 4 and Extended Data Table 1). The cryo-EM density maps of both structures are of sufficient quality for de novo model building of major parts of the protein (Extended Data Fig. 5). Here we use the higher-resolution PtdIns(3,5)P2-bound structure for the initial description of the overall structural features. Similar to AtTPC1, each MmTPC1 subunit contains two homologous 6-TM domains (6-TMI and 6-TMII) and two subunits that assemble into a rectangle-shaped functional channel, which is equivalent to a tetrameric Shaker-like channel (Fig. 1a, b and Extended Data Fig. 6). Following the same nomenclature as other voltage-gated channels, we labelled the six transmembrane helices within each 6-TM domain as IS1–IS6 and IIS1–IIS6, respectively (Fig. 1c). The transmembrane region of MmTPC1 is domain-swapped; the S1–S4 voltage-sensing domain (VSD) from one 6-TM interacts with the S5–S6 pore domain from the neighbouring 6-TM (Fig. 1b). The pore domain of the second 6-TM contains a luminal loop between IIS5 and pore helix 1 (IIP1) that forms an upright antenna-like β-hairpin; Asn600 and Asn612 on this luminal loop are glycosylated with visible density for the covalently linked N-acetylglucosamine moiety of the sugar29 (Fig. 1b, e and Extended Data Fig. 5c).

Figure 1: Overall structure of MmTPC1.

a, A 3D reconstruction of PtdIns(3,5)P2-bound (purple density) MmTPC1 dimer with each subunit in individual colour. b, Cartoon representation of MmTPC1 in the same orientations as the electron microscopy maps in a. N-acetylglucosamine (NAG) molecules and PtdIns(3,5)P2 (purple) are shown as sticks. c, Topology and domain arrangement of MmTPC1 subunit. d, Structure of the 6-TMI and the soluble domain, with individual elements coloured as in c. Inset, zoomed-in view of the cytosolic soluble domain. e, Structure of the 6-TMII.

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Multiple cytosolic components within each TPC1 subunit—including the N-terminal H1 helix, the linker between the two 6-TMs and the C-terminal post IIS6 region—assemble into a tightly packed cytosolic domain (Fig. 1d). Despite low sequence homology, the linker between the two 6-TMs adopts the EF-hand domain structure with two EF-hand motifs (EF-1 and EF-2), similar to plant TPC1, and the C-terminal portion of the exceptionally long IS6 serves as the E1 helix (Fig. 1d and Extended Data Fig. 6d). Ca2+ is unlikely to bind to the EF motifs of MmTPC1 as these motifs lack essential Ca2+-chelating acidic residues (Extended Data Fig. 1). The N-terminal H1 helix is tightly packed with the EF-1 motif and becomes an integral part of the EF-hand domain (Fig. 1d). Compared with plant TPC1 and mammalian TPC2, MmTPC1 has a much longer C-terminal region, which adopts a horseshoe-shaped structure with four α-helices and two β-strands and tightly wraps around the EF-hand domain (Fig. 1d and Extended Data Figs 1, 6d).

The MmTPC1 ion conduction pore, which consists of S5, S6 and two pore helices, adopts a closed conformation in the apo structure and an open conformation in the PtdIns(3,5)P2-bound structure (Fig. 2a–d). In the apo structure, the four pore-lining S6 helices form a bundle-crossing at the cytosolic side, with two layers of hydrophobic residues—the Leu317 and Phe321 from each IS6 helix, and the Val684 and Leu688 from each IIS6 helix—that form the constriction points that prevent the passage of hydrated cations (Fig. 2b, c). In the PtdIns(3,5)P2-bound state, the S6 helices undergo outward movement along with rotational motion (Fig. 2d). Consequently, the constriction-forming residues dilate and rotate away from the central axis, resulting in a much wider opening at the intracellular gate (Fig. 2b–d). In addition, the side chains of four acidic residues (Asp322 from each IS6 and Glu689 from each IIS6) that point tangentially to the pore in the closed structure undergo inward rotation and face the ion conduction pathway in the open state (Fig. 2b–d), generating a ring of negative charges at the gate that could facilitate channel conductance. The molecular mechanism of PtdIns(3,5)P2-induced channel opening will be discussed later.

Figure 2: Ion conduction pore of MmTPC1.

a, Ion conduction pore, comprising IS5–S6 (pore 1) and IIS5–S6 (pore 2). b, c, Side view of the bundle-crossing formed by IS6 (b) and IIS6 (c) in the apo closed (salmon) and PtdIns(3,5)P2-bound open (blue) states. Numbers are cross distances (in Å) at the constriction points. d, Structural comparison of the cytosolic gate between the closed and open states viewed from the cytosolic side in three sections: Leu317/Val684 (left), Phe321/Leu688 (middle) and Asp322/Glu689 (right). e, Side view of the selectivity filter formed by filter I and filter II with the front subunit removed for clarity. Numbers are cross distances (in Å) at the constriction points. f, Top view of the selectivity filter. Inset, zoomed-in view of the filter with the stabilization H-bonds for Asn648 (dotted line) and electron microscopy density (grey) shown. g, Sample IV curves of the filter mutations recorded with high Na+ or K+ in the bath solution. PNa/K, permeability ratio of Na+ to K+. Original traces are shown in Extended Data Fig. 7. The experiments were repeated five times independently with similar results.

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The selectivity filter region remains identical in both structures. Two sets of filter residues, Thr280–Ala281–Asn282 (filter I) in 6-TMI and Val647–Asn648–Asn649 (filter II) in 6-TMII, enclose the central ion pathway with different dimensions (Fig. 2e, f). The residues of filter I line the pathway with predominantly main-chain backbone carbonyls and have atom-to-atom cross distances of about 8 Å (Fig. 2e). The residues of filter II use side chains to generate a much narrower pathway, with two constriction points formed by the Asn648 and Asn649 residues (Fig. 2e). Positioned at the centre of the filter and stabilized by hydrogen-bonding interactions with the filter I backbone carbonyls of Thr280 and Ala281, the Asn648 side chain forms the narrowest point along the filter pathway; it has a cross distance of about 3.7 Å and has the central role in defining the Na+ selectivity of MmTPC1 (Fig. 2e, f). Asn648Ala mutation results in a complete loss of Na+ selectivity (Fig. 2g and Extended Data Fig. 7). The Asn649 residues are positioned at the luminal entrance of the channel at a wider distance from one another, and Asn649Ala mutation reduces but does not abolish Na+ selectivity (Fig. 2g and Extended Data Fig. 7). With an elongated coin-slot-like ion pathway at the filter, Na+ ions probably pass through the MmTPC1 filter in a partially hydrated form. The two Asn648 side chains are positioned to provide optimal coordination to stabilize the permeating Na+ ion, but are too close to permit the passage of K+. Thus, Asn648 forms a simple size sieve to exclude K+ or larger ions.

The two VSDs (VSD1 and VSD2) have virtually the same structures as their respective counterparts between the apo and ligand-bound states and, therefore, the higher-resolution PtdIns(3,5)P2-bound structure will be used in the discussion (Extended Data Fig. 8a, b). Figure 3a provides the numbering of the S4 gating charge residues (R1–R5) from TPCs and other canonical voltage-gated channels for comparison. Although VSD1 contains three arginine residues in IS4 (Arg200, Arg203 and Arg206, at positions R2, R3 and R4, respectively) (Fig. 3b), it lacks some key features of canonical voltage sensors (see Extended Data Fig. 8a legend) and does not contribute to voltage-dependent gating—similar to the VSD1 of plant TPC125—as shown by the fact that replacing these arginines individually with a neutral residue does not affect the voltage activation of MmTPC1 (Fig. 3c and Extended Data Fig. 8c).

Figure 3: The voltage-sensing domains.

a, Partial S4 sequence alignment and arginine registry. NavAb, Nav channel from Arcobacter butzleri. b, Side view of VSD1 with IS1 omitted for clarity. c, G/GmaxV curves of wild-type MmTPC1 and IS4 arginine mutations. Sample traces are shown in Extended Data Fig. 8. All data points are mean ± s.e.m. (n = 5 independent experiments). d, Side view of VSD2 with IIS1 omitted for clarity. e, Sample IV curves of wild-type MmTPC1 (obtained from the peak currents at various activation potentials) and Arg540Gln mutant (obtained by applying a voltage pulse ramp from −100 to 100 mV). Currents were recorded with 2 μM PtdIns(3,5)P2 in the pipette and repeated five times independently with similar results. f, Structural comparison of VSD2 between the PtdIns(3,5)P2-bound MmTPC1 (orange) and AtTPC1 (cyan), with S1 helices omitted for clarity. g, Cartoon representation of VSD2 conformational change from the activated to resting state. Red arrows indicate the concurrent movements of S4 and S4–S5 linker.

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Source data

VSD2 contains only two S4 arginines (Arg540 at position R3 and Arg546 at position R5), preserves the key features of a canonical voltage sensor—including the 310-helix in IIS4 and the conserved gating-charge transfer centre (Fig. 3d)—and is responsible for the voltage gating of MmTPC1. Mutations of Arg540 and Arg546 have a profound but opposite effect on the voltage dependence of the channel. The Arg540Gln mutation stabilizes VSD2 in an activated state and yields a voltage-independent channel that has a linear current–voltage relationship between −100 and 50 mV and can be activated by PtdIns(3,5)P2 even at hyperpolarization (Fig. 3e). The Arg546Gln mutant, by contrast, can barely be activated by voltage even at high concentrations of PtdIns(3,5)P2, as if the voltage sensor is trapped in the resting state (Extended Data Fig. 8d).

The MmTPC1 VSD2 adopts an activated conformation with its final voltage-sensing arginine (Arg546) positioned in the gating-charge transfer centre formed by Tyr487 and Glu490 from IIS2 and Asp512 from IIS3, and the other voltage-sensing residues (Arg540 and Gln543) facing the external, luminal side (Fig. 3d). The VSD2 of AtTPC1, with its S4 arginines residing at positions R3–R5, is also responsible for the voltage-gating of the channel, and its structure is in the resting state25. We can, therefore, extrapolate the conformational change of MmTPC1 VSD2 from the activated to the resting state by comparing its structure with that of AtTPC1 (Fig. 3f, g). Except for the S4 helix, the two structures superimpose well. This suggests that upon hyperpolarization the IIS4 of MmTPC1 would slide down by about two helical turns without undergoing structural change in the rest of VSD2, and position its R3 arginine (Arg540) at the gating-charge transfer centre (Fig. 3g). Concurrent with the IIS4 sliding, the IIS4–S5 linker would swing downward and move closer to IIS6. Notably, VSD2 is in the activated state in both the apo and PtdIns(3,5)P2-bound structures, indicating that the voltage sensor can be activated without opening the channel in MmTPC1.

The bound PtdIns(3,5)P2 can be unambiguously identified from the electron microscopy density map of the ligand-bound structure (Extended Data Figs 5d, 9a). PtdIns(3,5)P2 is situated at the junction formed by IS3, IS4 and the IS4–S5 linker of 6-TMI; its inositol 1,3,5-trisphosphate head group is positioned on the cytosolic side and its acyl chains are inserted upright into the membrane (Fig. 4a and Extended Data Fig. 9a). Figure 4b summarizes the protein–ligand interactions, which involve predominantly basic residues from the C terminus of H1, the N terminus of IS3, the IS4–S5 linker and the C-terminal part of IS6. Buried deep in the protein, the two phosphate groups on the C1 and C3 positions of the inositol muster the majority of protein–ligand interactions and probably define the ligand specificity. The C5 phosphate protrudes outwardly away from the ligand-binding pocket, and forms salt bridges with Lys87 and Lys331; the interaction with Lys331 participates in the coupling between the ligand and IIS6, and has an important role in the ligand activation of the channel. Among all the ligand-interacting residues at the PtdIns(3,5)P2-binding site, mutations of the residues that predominantly interact with the C3 phosphate—including the three arginines (Arg220, Arg221 and Arg224) on the IS4–S5 linker and Lys331 on IS6—appear to have the most profound effect on PtdIns(3,5)P2 activation, which illustrates the central role of the C3 phosphate (Extended Data Fig. 9b). In a recent study, the three linker arginines have also been reported to be important for NAADP-mediated Ca2+ release30.

Figure 4: PtdIns(3,5)P2 binding in MmTPC1.

a, PtdIns(3,5)P2 binding in 6-TMI of MmTPC1. Inset: zoomed-in view of the PtdIns(3,5)P2 site. b, Schematic of the protein–ligand interactions. c, Concentration-dependent PtdIns(3,5)P2 activation of Arg540Gln mutant at −100 mV. Curve is least square fit to the Hill equation. Data points are mean ± s.e.m. (n = 5 independent experiments). Sample IV curves are shown in Extended Data Fig. 9c. d, Ligand specificity of MmTPC1 measured using the Arg540Gln mutant. Sample IV curves were recorded on the same patch with different PtdInsP2 isoforms. The experiments were repeated five times independently with similar results. e, Close proximity between the C4 hydroxyl of PtdIns(3,5)P2 and the surrounding residues. f, Structural comparison at the region around Lys331 between the apo (green) and PtdIns(3,5)P2-bound MmTPC1 (salmon).

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To investigate the affinity and specificity of the ligand, we measured the activity of the ligand-dependent channel in excised patches by using the voltage-independent Arg540Gln mutant, which simplifies ligand-dependent gating by eliminating the voltage effect. The mutant also elicits much larger currents, which makes it suitable for inside-out patches. The PtdIns(3,5)P2-dependent activation of the mutant yielded a half-maximal effective concentration (EC50) of about 145 nM (Fig. 4c and Extended Data Fig. 9c), similar to that of human TPC1 measured in whole lysosome patch23. The PtdIns(4,5)P2 isoform cannot activate the channel or inhibit PtdIns(3,5)P2 activation (Fig. 4d), indicating high lipid specificity of MmTPC1. The lack of PtdIns(4,5)P2-binding can be explained by the missing C3 phosphate and the close proximity of Asn85 and Lys87 to the C4 hydroxyl group, which sterically excludes the C4 phosphate and thereby prevents the binding of the lipid (Fig. 4e).

Compared to the apo structure, PtdIns(3,5)P2-binding does not introduce major structural changes around the ligand-binding pocket (Extended Data Fig. 9d), except for one key conformational change on IS6 mediated by Lys331 (Fig. 4f). In the apo state, the Lys331 side chain points away from the ligand-binding pocket. In the presence of PtdIns(3,5)P2, the Lys331 side chain adopts an extended configuration to form salt bridges with both the C3 and C5 phosphates as well as a hydrogen bond with the C4 hydroxyl, pulling IS6 towards the ligand-binding pocket (Fig. 4f). This movement propagates to the other part of IS6, as well as IIS6, and opens the gate. Lys331 appears to be the only residue that couples IS6 to the bound PtdIns(3,5)P2 and its mutation to Ala completely abolishes PtdIns(3,5)P2 activation (Extended Data Fig. 9b).

Our structures demonstrate that PtdIns(3,5)P2 only binds to the first 6-TM domain and directly introduces conformational changes in IS6 helix, whereas voltage influences only the VSD2 in the second 6-TM domain, the conformational change of which is likely to affect the movement of IIS6 helix (Figs 3, 4). A global structural comparison between the apo and PtdIns(3,5)P2-bound structures explains the interplay between the two stimuli (Fig. 5). Despite having an activated voltage sensor, the MmTPC1 pore remains closed in the apo structure, implying that PtdIns(3,5)P2-binding is required to trigger the opening of the gate. Upon PtdIns(3,5)P2-binding, the ensuing tethering interaction between Lys331 and PtdIns(3,5)P2 straightens the IS6 helices that are initially bent at the π-helix just below the filter region in the closed state, resulting in the outward dilation and rotation at the bundle crossing (Figs 2d, 5a). The five-residue π-helix is present only in IS6 and may facilitate the helix bending. To open the pore, the IIS6 helices also have to undergo concurrent outward and rotational movements to accommodate the PtdIns(3,5)P2-induced conformational change in IS6 helices, particularly the rotation of the two IS6 gating residues with large hydrophobic side chains (Leu317 and Phe321). Consequently, the two IIS6 gating residues (Val684 and L688) also rotate away from the central axis and open the gate (Figs 2d, 5b). The IIS6 motion is hinged around the residue immediately below the filter region, and is propagated to a much larger movement at the C-terminal end of IIS6, which swings upward and makes direct contact with the IIS4–S5 linker. Such motion is permitted only when IIS4 of VSD2 is in the activated, up state. Under hyperpolarized membrane potential, IIS4 is expected to slide downward and push the IIS4–IIS5 linker along with it, occluding the space necessary for upward IIS6 movement upon PtdIns(3,5)P2 activation (Fig. 3f, g). PtdIns(3,5)P2 can probably still bind MmTPC1 under hyperpolarization, but the resting VSD2 prevents channel opening by blocking the movement of IIS6. Thus, membrane potential modulates the TPC1 channel activity by imposing a voltage-dependent constraint on PtdIns(3,5)P2 activation and the upward movement of VSD2 under depolarization is a prerequisite for the PtdIns(3,5)P2-induced gate opening (Fig. 5c).

Figure 5: Gating mechanism of MmTPC1.

a, b, Structural comparison between the apo closed (grey) and PtdIns(3,5)P2-bound open (green) MmTPC1 with zoomed-in views of the IS3–S6 (a) and IIS3–S6 (b) regions. Arrows indicate the S6 movements. Key gating residues are shown as sticks. IS6 contains a five-residue π-helix (coloured red). c, Working model for voltage-dependent PtdIns(3,5)P2 activation of MmTPC1. Red arrows mark the direction of the driving force.

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No statistical methods were used to predetermine sample size. The experiments were not randomized and investigators were not blinded to allocation during experiments and outcome assessment.

Protein expression and purification

Mouse TPC1 (MmTPC1, NCBI accession: NM_145853.2) containing a C-terminal thrombin cleavage site followed by a GFP tag and a 10× His tag was cloned into a pEZT-BM vector31 and heterologously expressed in HEK293F cells (Life Technologies) using the BacMam system (Thermo Fisher Scientific). The baculovirus was generated in Sf9 cells (Life Technologies) following the standard protocol and used to infect HEK293F cells at a ratio of 1:40 (virus:HEK293F, v/v) and supplemented with 10 mM sodium butyrate to boost protein expression. Cells were cultured in suspension at 37 °C for 48 h and collected by centrifugation at 3,000g. All purification procedures were carried out at 4 °C. The cell pellet was re-suspended in buffer A (20 mM Tris, pH 8.0, 150 mM NaCl) supplemented with a protease inhibitor cocktail (containing 2 μg/ml DNase, 0.5 μg/ml pepstatin, 2 μg/ml leupeptin, and 1 μg/ml aprotinin and 0.1 mM PMSF) and homogenized by sonication on ice. MmTPC1 was extracted with 1% (w/v) n-dodecyl-β-d-maltopyranoside (Anatrace) supplemented with 0.2% (w/v) cholesteryl hemisuccinate (Sigma Aldrich) by gentle agitation for 2 h. After extraction, the supernatant was collected after a 60-min centrifugation at 20,000g and incubated with Ni-NTA resin (Qiagen) using gentle agitation. After 2 h, the resin was collected on a disposable gravity column (Bio-Rad). The resin was washed with buffer B (20mM Tris, pH 8.0, 150 mM NaCl and 0.06% glycol-diosgenin (Anatrace) supplemented with 20 mM imidazole. The washed resin was left on column in buffer B and digested with thrombin (Roche) overnight. After thrombin digestion, the flow-through containing untagged MmTPC1 was collected, concentrated and purified by size exclusion chromatography on a Superdex 200 column (GE Heathcare) pre-equilibrated with buffer B. The peak fraction was pooled and concentrated to 4.7 mg/ml for cryo-EM analysis. To obtain PtdIns(3,5)P2-bound structure, the protein sample was supplemented with 0.5 mM PtdIns(3,5)P2 diC8 (Echelon Biosciences) for 30 min on ice before electron microscopy grid preparation.

Electron microscopy data acquisition

The cryo-EM grids were prepared by applying MmTPC1 (4.7 mg/ml, with or without 0.5 mM PtdIns(3,5)P2) to a glow-discharged Quantifoil R1.2/1.3 300-mesh gold holey carbon grid. Grids were blotted for 4.0 s under 100% humidity at 4 °C before being plunged into liquid ethane using a Mark IV Vitrobot (FEI). Micrographs were acquired on a Titan Krios microscope (FEI) operated at 300 kV with a K2 Summit direct electron detector (Gatan), using a slit width of 20 eV on a GIF-Quantum energy filter. Images were recorded with EPU software (FEI) in super-resolution counting mode with a super resolution pixel size of 0.535 Å. The defocus range was set from −1.5 μm to −3 μm. Each micrograph was dose-fractionated to 30 frames under a dose rate of 4 e per pixel per s, with a total exposure time of 15 s, resulting in a total dose of about 50 e per Å2.

Image processing

Micrographs were motion corrected and binned twofold (yielding a pixel size of 1.07 Å per pixel) with MotionCor232. The CTF parameters of the micrographs were estimated using the GCTF program33. All other steps of image processing were performed using RELION v.2.034,35. Initially, ~1,000 particles were manually picked from a few micrographs. Class averages representing projections of MmTPC1 in different orientations were selected from the 2D classification of the manually picked particles, and used as templates for automatic particle picking from the full dataset. For the Apo MmTPC1 dataset, 1,411,763 particles were picked from 2,937 micrographs. The particles were extracted and binned 3 times (3.21 Å per pixel). After 2D classification, a total of 1,117,348 particles were finally selected for 3D classification using the AtTPC1 structure as the initial mode. Three of the 3D classes showed good secondary structure features and were selected and re-extracted into the original pixel size of 1.07 Å. After 3D refinement with C2 symmetry imposed, and particle polishing, the resulting 3D reconstructions from ~536,000 particles showed a clear two-fold symmetry with a resolution of 3.5 Å. We then performed a focused 3D classification with density subtraction to improve the density of transmembrane domain36. In this approach, only the density corresponding to the transmembrane domain was kept in each particle image, by subtracting the density for all other parts including the belt-like detergent density from the original particles. The subsequent 3D classification on the modified particles was carried out by applying a mask around the transmembrane domain with all the particle orientations fixed at the value determined in the initial 3D refinement. After this round of classification, one class (~43,000 particles) showed better density in the transmembrane domain. The corresponding particles before density subtraction from this class were selected and 3D-refined, yielding an electron microscopy map of 3.4 Å for the entire channel.

The data for MmTPC1 in the presence of PtdIns(3,5)P2 were processed similarly to that of apo MmTPC1. In brief, 941,754 particles were picked from a total of 2,348 micrographs. After 2D classification, 620,307 particles were selected for 3D classification. Three classes with a total of ~245,000 particles were selected and combined for 3D auto-refinement, which resulted in a map with an overall resolution of 3.3 Å. One round of 3D classification was then performed by focusing on the transmembrane domain. One class (~83,000 particles) showed better density in the transmembrane domain and was selected for final 3D refinement, yielding an electron microscopy map of 3.2 Å. All resolutions were estimated by applying a soft mask around the protein density and the gold-standard Fourier shell correlation (FSC) = 0.143 criterion. ResMap was used to calculate the local resolution map37.

Model building, refinement and validation

De novo atomic model buildings were conducted in Coot38. Amino acid assignment was achieved based mainly on the clearly defined densities for bulky residues (Phe, Trp, Tyr and Arg). Real-space model refinement was performed in Phenix39. Models were validated using previously described methods, to avoid overfitting40,41. The final structure models for both apo and PtdIns(3,5)P2-bound states include residues 66–701 and residues 709–795. Residues 1–65, 702–708 and 796–817 are disordered and not modelled. The statistics of the geometries of the models were generated using MolProbity42. All the figures were prepared in PyMol43 or Chimera44. Programs used for model building, refinement and validation are compiled by SBGrid45.


In human TPC2, the Leu11Ala and Leu12Ala mutations at the N-terminal targeting sequence have previously been shown to promote channel expression and trafficking to the plasma membrane of the HEK293 cell, enabling channel activity measurement using patch clamp22,46. We therefore also introduced the equivalent mutations (Leu11Ala and Ile12Ala) to MmTPC1. HEK293 cells overexpressed with the Leu11Ala/Ile12Ala mutant of MmTPC1 elicited much larger whole-cell currents than those expressed with wild-type MmTPC1 (Extended Data Fig. 2a). Therefore, the Leu11Ala/Ile12Ala mutant was used and considered as the wild-type channel in all our recordings. All other mutations in our experiments were generated on the background of this plasma-membrane-targeting MmTPC1. With the channels targeted to the plasma membrane, the extracellular side is equivalent to the luminal side of TPC1 in endosomes or lysosomes. MmTPC1 and its mutants were cloned into pCGFP-EU vector47. About 2 μg of the plasmid containing the C-terminal GFP-tagged MmTPC1 or its mutant was transfected into HEK293 cells grown in a six-well tissue culture dish using Lipofectamine 2000 (Life Technology). Forty-eight hours after transfection, cells were dissociated by trypsin treatment and kept in complete serum-containing medium and re-plated on 35-mm tissue culture dishes in a tissue culture incubator until recording.

Patch clamp in whole-cell configuration was used to measure channel activity in most of the experiments except the measurements of ligand affinity and specificity, which were recorded in excised patches (inside-out patches) using the voltage-independent Arg540Gln mutant. This mutant channel can be activated solely by PtdIns(3,5)P2 and also yields much larger plasma membrane currents, which makes it more amenable for inside-out patches. The standard intracellular solution contained (in mM): 145 sodium methanesulfonate (Na-MS), 5 NaCl, 4 MgCl2, 1 EGTA, 10 HEPES buffered with Tris, pH = 7.4. The extracellular solution contained (in mM): 145 Na-MS, 5 NaCl, 1 MgCl2, 1 CaCl2, 10 HEPES buffered with Tris, pH = 7.4. Various concentrations of PtdIns(3,5)P2 as specified in each experiment were added to the intracellular solutions to activate the channel. For patches in whole-cell configuration, the intracellular solution was in the pipette and the extracellular solution was in the bath; the solution arrangement was reversed for the inside-out patches. The lipid ligands used in our studies are phosphatidylinositol-3,5-bisphosphate diC8 (PtdIns(3,5)P2 diC8, Echelon) and phosphatidylinositol-4,5-bisphosphate diC8 (PtdIns(4,5)P2 diC8, Echelon).

The data were acquired using an AxoPatch 200B amplifier (Molecular Devices) and a low-pass analogue filter set to 1 kHz. The current signal was sampled at a rate of 20 kHz using a Digidata 1322A digitizer (Molecular Devices) and further analysed with pClamp 9 software (Molecular Devices). Patch pipettes were pulled from borosilicate glass (Harvard Apparatus) and heat-polished to a resistance of 3–5 MΩ. After the patch pipette attached to the cell membrane, a giga-seal (>10GΩ) was formed by gentle suction. The whole-cell configuration was formed by short zap or suction to rupture the patch. The inside-out configuration was formed by pulling the pipette away from the cell, and the pipette tip was exposed to the air for a short period in some cases. The holding potential was set to −70 mV. To generate G/Gmax versus V curves (G = I/V), the membrane was stepped from the holding potential (−70 mV) to various testing potentials (−100 mV to 100 mV) for 1 s and then stepped to −70 mV (Extended Data Fig. 2b). The peak tail currents were used to plot the GV curve. Gmax was obtained from the peak tail current at 100 mV testing potential. V1/2 and Z values were obtained from the fits of data with Boltzmann equation, in which V1/2 is the voltage at which the channels have reached half of their maximum fraction open and Z is the apparent valence of voltage dependence. The same protocol was used to obtain current and voltage relationships (IV curve) of the wild-type MmTPC1 (Fig. 3e, top trace), except that the peak current at each testing potential was used to generate the IV curve. For voltage-independent Arg540Gln mutant, the holding potential was set to 0 mV, and the current and voltage relationship (IV curve, Fig. 3e, bottom trace) was obtained directly by using voltage pulses ramp from −100 to 100 mV over 800-ms duration.

For measuring ion selectivity of MmTPC1 and its mutants in whole-cell patches, 10 μM PtdIns(3,5)P2 was included in intracellular (pipette) solution to fully activate the channel. The membrane potential was stepped from the holding potential (−70 mV) to 100 mV for 1 s to activate the channels, and then stepped to various testing potentials (−120 mV to 4 mV) for 1 s (Extended Data Fig. 2f). The peak tail currents at various testing potentials were plotted to determine the reversal potential (Vrev). To measure the relative permeability between Na+ and K+, the extracellular (bath) solution (in mM) was changed to 145 K-MS, 5 NaCl, 1 MgCl2, 1 CaCl2, 10 HEPES buffered with Tris, pH 7.4. To measure the relative permeability between Na+ and Ca2+, the extracellular solution (in mM) was changed to 95 Ca-(MS)2, 5 CaCl2, 10 HEPES buffered with Tris, pH 7.4. The ion permeability ratios were calculated with the equations: PNa/PK = [K]o/([Na]iexp(VrevF/RT) – [Na]o) and PNa/PCa = 4[Ca]o/([Na]iexp(VrevF/RT)(1 + exp(VrevF/RT))), in which Vrev is the reverse potential, F is Faraday’s constant, R is the gas constant, T is the absolute temperature, o is extracellular and i is intracellular.

All electrophysiological recording were repeated at least five times using different patches. Most data points shown are mean ± s.e.m. (n = 5 independent experiments).

Data availability

The cryo-EM density maps of the MmTPC1 have been deposited in the Electron Microscopy Data Bank under accession number EMD-7434 for the apo state, and accession number EMD-7435 for the PtdIns(3,5)P2-bound state. Atomic coordinates have been deposited in the RCSB Protein Data Bank under accession number 6C96 for the apo state, and accession number 6C9A for the PtdIns(3,5)P2-bound state. Source Data for Fig. 3c and Extended Data Fig. 2c, e are available in the online version of the paper.

Accession codes

Primary accessions

Electron Microscopy Data Bank

Protein Data Bank


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We thank N. Nguyen for manuscript preparation and M. X. Zhu for providing clones of animal TPC genes. Single particle cryo-EM data were collected at the University of Texas Southwestern Medical Center (UTSW) Cryo-Electron Microscopy Facility that is funded by the CPRIT Core Facility Support Award RP170644. We thank D. Nicastro and Z. Chen for facility access and data acquisition. Negatively stained sample screening was performed at UTSW Electron Microscopy core. This work was supported in part by the Howard Hughes Medical Institute (Y.J.) and by grants from the National Institute of Health (GM079179 to Y.J.) and the Welch Foundation (Grant I-1578 to Y.J.). X.B. is supported by the Cancer Prevention and Research Initiative of Texas and Virginia Murchison Linthicum Scholar in Medical Research fund.

Author information




J.S., J.G. and Q.C. prepared the samples; J.S., J.G., Q.C. and X.B. performed data acquisition, image processing and structure determination; W.Z. performed electrophysiology; Y.J. supervised the project and revised the manuscript; all authors participated in research design, data analysis and manuscript preparation.

Corresponding authors

Correspondence to Youxing Jiang or Xiao-chen Bai.

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Extended data figures and tables

Extended Data Figure 1 Sequence alignment of MmTPC1, HsTPC1, AtTPC1, MmTPC2 and HsTPC2.

Secondary structure assignments are based on the structure of PtdIns(3,5)P2-bound MmTPC1. Red dots mark the ligand-binding residues; black dots mark the S4 arginine residues and residues at the gating-charge transfer centre; cyan dots mark the key S6 gating residues; green dots mark the residues predicted to participate in Ca2+ coordination in EF-hand domains of AtTPC1. MmTPC1 and AtTPC1 share about 25% sequence identity.

Extended Data Figure 2 Gating and selectivity of MmTPC1.

a, Sample traces and current density (current/capacitance) of wild-type MmTPC1 and the L11A/I12A mutant of MmTPC1, recorded in the whole-cell configuration with 100 μM PtdIns(3,5)P2 in the pipette (cytosolic). The experiments were repeated five times independently with similar results. Data points for current density are mean ± s.e.m. (n = 5 independent experiments). The L11A/I12A mutant elicited much larger whole-cell currents and was therefore used as the wild-type channel in all recordings. The extracellular side of MmTPC1 in plasma membrane is equivalent to the luminal side of MmTPC1 in lysosomes. b, Sample traces of PtdIns(3,5)P2-dependent voltage activation of MmTPC1. Whole-cell currents were recorded with varying PtdIns(3,5)P2 concentrations in the pipette (cytosolic) at pH 7.4. The experiments were repeated five times independently with similar results. c, G/GmaxV curves of MmTPC1 at various PtdIns(3,5)P2 concentrations. Boltzmann fit yields V1/2 (mV) = 21.6 ± 1.2, 15.2 ± 1.0, 16.1 ± 0.9 and −2.0 ± 1.0, and Z = 0.78 ± 0.04, 0.82 ± 0.03, 0.89 ± 0.02 and 0.84 ± 0.05 for voltage activation in 0.05, 0.2, 2.0 and 10 μM cytosolic PtdIns(3,5)P2, respectively, in which V1/2 is the membrane potential for half maximum activation and Z is apparent valence. All data points are mean ± s.e.m. (n = 5 independent experiments). d, Luminal pH modulates the voltage activation of MmTPC1. Whole-cell currents of MmTPC1 recorded in the presence of 2 μM cytosolic PtdIns(3,5)P2 with a varying luminal (bath) pH of 7.4, 6.0 or 4.6. Sample traces were obtained from the same patch. The experiments were repeated five times independently with similar results. e, G/GmaxV curves of MmTPC1 at various luminal pH values. Boltzmann fit yields V1/2 = 16.2 ± 0.8 mV, Z = 0.91 ± 0.02 at pH 7.4, V1/2 = 38.2 ± 1.2 mV, Z = 0.95 ± 0.02 at pH 6.0. All data points were normalized against Gmax obtained at 100 mV activation voltage and pH 7.4. All data points are mean ± s.e.m. (n = 5 independent experiments). f, Sample traces of whole-cell currents with 150 mM Na+ in the pipette solution, and either 150 mM Na+ or 145 mM K+ and 5 mM Na+ in the bath solution, and the IV curves generated from the tail currents of the sample traces. g, Sample traces of whole-cell currents with 150 mM Na+ in the pipette solution and 150 mM Na+ or 100 mM Ca2+ in the bath solution, and the IV curves generated from the tail currents of the sample traces. Data in f and g were recorded with 10 μM PtdIns(3,5)P2 in the pipette at pH 7.4 and both experiments were repeated five times independently with similar results. Source data

Extended Data Figure 3 Structure determination of PtdIns(3,5)P2-bound MmTPC1.

a, Representative electron micrograph of PtdIns(3,5)P2-bound MmTPC1; 2,348 micrographs were used for structure determination. b, 2D class averages. c, Euler angle distribution of particles used in the final 3D reconstruction, with the heights of the cylinders corresponding to the number of particles. d, Final density maps coloured by local resolution. e, Gold-standard FSC curves of the final 3D reconstructions. f, FSC curves for cross-validation between the models and the maps. Curves for model versus summed map in black (sum), for model versus half map in blue (work) and for model versus half map not used for refinement in red (free). g, Flowchart of electron microscopy data processing for PtdIns(3,5)P2-bound MmTPC1 particles.

Extended Data Figure 4 Structure determination of apo MmTPC1.

a, Representative electron micrograph of apo MmTPC1; 2,937 micrographs were used for structure determination. b, 2D class averages. c, Euler angle distribution of particles used in the final 3D reconstruction, with the heights of the cylinders corresponding to the number of particles. d, Final density maps coloured by local resolution. e, Gold-standard FSC curves of the final 3D reconstructions. f, FSC curves for cross-validation between the models and the maps. Curves for model versus summed map in black (sum), for model versus half map in blue (work) and for model versus half map not used for refinement in red (free). g, Flowchart of electron microscopy data processing for apo MmTPC1 particles.

Extended Data Figure 5 Sample electron microscopy density maps (blue mesh) for MmTPC1.

a-d, Sample electron microscopy density maps for various parts of PtdIns(3,5)P2-bound MmTPC1: IS1–IS6 and filter I (a), IIS1–IS6 and filter II (b), NAGs of Asn600 and Asn612 (c), and PtdIns(3,5)P2-binding site (d). The maps are low-pass filtered to 3.2 Å and sharpened with a temperature factor of −105 Å2. e, f, Sample electron microscopy density maps for the key parts of apo MmTPC1: ligand binding site (e) and S6 helices (f). The maps are low-pass filtered to 3.4 Å and sharpened with a temperature factor of −98.5 Å2. Residues discussed in main text are labelled in red.

Extended Data Figure 6 Structure comparison between MmTPC1 and AtTPC1.

a, Superposition of the overall structures of MmTPC1 (blue) and AtTPC1(salmon). b, Superposition of the pore regions. c, Superposition of VSD1 domains. The comparison of the VSD2 domains is shown in Fig. 3f. d, Superposition of cytosolic soluble domains.

Extended Data Figure 7 Sample traces of whole-cell currents for Asn648Ala and Asn649Ala filter mutants.

The pipette solution contained 150 mM Na+ and the bath solution contained 150 mM Na+, or 145 mM K+ and 5 mM Na+. The tail currents were used to generate the IV curves shown in Fig. 2g. The experiments were repeated five times independently with similar results.

Extended Data Figure 8 Voltage-sensing domains.

a, Superimposition of MmTPC1 VSD1 structures in the PtdIns(3,5)P2-bound (green) and apo (pink) states with S1 helices removed for clarity. The MmTPC1 VSD1 lacks some key features of canonical voltage sensors: the conserved aromatic residue on S2 and acidic residue on S3 that form the gating-charge transfer centre become Val152 and Lys177, respectively, in MmTPC1; the conserved basic residue at the R5 position becomes Phe209 in MmTPC1; no arginine from IS4 is positioned in the gating-charge transfer centre. b, Superimposition of MmTPC1 VSD2 structures in the PtdIns(3,5)P2-bound (orange) and apo (cyan) states. c, Sample traces of voltage activation of MmTPC1 and its IS4 arginine mutations, recorded in whole-cell configuration with 2 μM PtdIns(3,5)P2 in the pipette. Peak tail currents were used to generate the G/GmaxV curves shown in Fig. 3c. The experiments were repeated five times independently with similar results. d, Sample traces of voltage activation of Arg546Gln mutation, recorded in whole-cell configuration with 2 μM and 100 μM PtdIns(3,5)P2 in the pipette. The experiments were repeated five times independently with similar results.

Extended Data Figure 9 PtdIns(3,5)P2-binding in MmTPC1.

a, Model of bound PtdIns(3,5)P2 (left) and its electron microscopy density (right). Density of two other membrane lipid molecules (blue mesh in left panel) was also observed near PtdIns(3,5)P2 in the structure. b, Current density of mutations at the PtdIns(3,5)P2-binding site measured at −100 mV in whole-cell recordings. All mutants were generated on the background of Arg540Gln mutant, which was used as control. All data points are mean ± s.e.m. with the number of independent experiments for each mutant shown in parentheses. c, Sample IV curves of Arg540Gln mutant recorded in excised patches with varying concentrations of PtdIns(3,5)P2 in the bath (cytosolic). The experiments were repeated five times independently with similar results. Currents at −100 mV were used to generate the concentration-dependent PtdIns(3,5)P2 activation curve shown in Fig. 4c. Imax is the current recorded at −100 mV with 10 μM PtdIns(3,5)P2 in the bath. d, Structural comparison at the ligand-binding site between the PtdIns(3,5)P2-bound (green) and apo (salmon) states.

Extended Data Table 1 Cryo-EM data collection and model statistics

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She, J., Guo, J., Chen, Q. et al. Structural insights into the voltage and phospholipid activation of the mammalian TPC1 channel. Nature 556, 130–134 (2018).

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