Introduction

Sodium-calcium exchangers (NCX) are transporters that control the flux of Ca²⁺ across the plasma membrane and play a vital role in maintaining cellular calcium homeostasis for cell signaling^1–6. NCXs facilitate the exchange of 3 Na⁺ for 1 Ca²⁺ in an electrogenic manner, primarily responsible for extruding Ca²⁺ from the cytoplasm. However, this process can be reversed to permit Ca²⁺ entry, depending on the chemical gradients of Na⁺ and Ca²⁺, as well as the membrane potential^3,7–13. Three NCX isoforms (NCX1-3) are present in mammals, with each isoform bearing multiple splice variants expressed in distinct tissues, thereby modulating numerous fundamental physiological events^4,14–19. Dysfunctions of NCXs are associated with a plethora of human pathologies including cardiac hypertrophy, arrhythmia, and postischemic brain damage^3,20–22. The cardiac variant NCX1.1 has been extensively studied, with its function playing a central role in cardiac excitation and contractile activity^23–27.

The eukaryotic NCX consists of a transmembrane (TM) domain with 10 TM helices and a large intracellular regulatory domain between TMs 5 and 6^4,28–33. The TM domain is responsible for the ion exchange function in NCX. It consists of two homologous halves (TMs 1-5 and TMs 6-10) with TMs 2-3 and TMs 7-8 forming the core of the TM domain and hosting the residues that coordinate the transported Na⁺ and Ca^2+34,35. The large regulatory domain contains two calcium-binding domains (CBD1 and CBD2). Ca²⁺ binding at CBDs enhances NCX activity and rescues it from the Na⁺-dependent inactivation, a process that manifests as slow decay of the exchange current due to elevated levels of cytosolic Na⁺ ions^{12,30,31,36–41}. A stretch of residues known as XIP region (eXchanger Inhibitory Peptide) at the N terminus of the regulatory domain plays a pivotal role in the Na⁺ inactivation process^42,43.

Several other cellular cues can also modulate NCX1 activity, including phosphatidylinositol 4,5-bisphosphate (PIP₂) in the membrane. PIP₂ has been shown to stimulate the NCX1 activity by reducing the Na⁺-dependent inactivation and the XIP region was suggested to participate in the PIP₂ activation^39,44–46. In addition, several small molecule NCX inhibitors have been developed to provide valuable tools for studying the physiological and pharmacological properties of NCXs, among which compound SEA0400 is a highly potent and selective NCX1 inhibitor that promotes the Na⁺-dependent inactivation of the exchanger^20,47–52.

We previously determined the human cardiac NCX1 structure in an inward-facing inactivated state in which XIP and the β-hairpin between TMs 1 and 2 form a TM-associated four-stranded β-hub and mediate a tight packing between the TM and cytosolic domains, resulting in the formation of a stable inactivation assembly that blocks the TM movement required for ion exchange function³². The study also provides a mechanistic insight into how cytosolic Ca²⁺ binding at CBD2 destabilizes the inactivation assembly and activates the exchanger³². To expand our understanding of NCX modulation by PIP₂ lipid and small molecule inhibitors, we present the cryo-EM structures of human cardiac NCX1 in complex with PIP₂ or SEA0400, revealing the structural basis underlying PIP₂ activation and SEA0400 inhibition of NCX1.

Results

PIP₂ activation of NCX1

Phosphatidylinositol 4,5-bisphosphate (PIP₂) has been shown to modulate NCX1 activity by reducing the Na⁺-dependent inactivation of the exchanger^39,44–46. To characterize the effect of PIP₂ on HsNCX1, we expressed the exchanger in Xenopus laevis oocytes and recorded the outward exchanger currents using the giant patch technique in the inside-out configuration with or without applying porcine brain PIP₂ (Fig. 1, and Methods). The recording was performed with 12 µM free cytosolic [Ca²⁺]_i (bath) and the outward NCX1 current was elicited by the rapid replacement of 100 mM Cs⁺ with 100 mM Na⁺ in the bath solution. In the patches without applying PIP₂, the outward exchanger currents quickly decay and reach a steady state due to Na⁺-dependent inactivation (Fig. 1A). Introducing 10 µM PIP₂ at the steady state progressively increases the current that plateaus at about two-fold of the peak current measured before PIP₂ addition (Fig. 1A). After PIP₂ washout, the outward current remains at the pre-removal level without obvious inactivation, indicating high-affinity PIP₂ binding and its positive modulation of NCX1 by both potentiating the peak current and reducing the Na⁺-dependent inactivation. Intriguingly, the commonly used shorter chain PIP₂ substitute (PIP₂ diC8) does not have the equivalent activation effect on NCX1 and the exchanger remains susceptible to Na⁺-dependent inactivation when recorded in the presence of 10 µM PIP₂ diC8 (Fig. 1B). However, PIP₂ diC8 still binds and stimulates both the peak and steady currents of the exchanger. This stimulation effect is abolished after PIP₂ diC8 washout, indicating a lower affinity of short-chain PIP₂ than that of the long-chain native PIP₂.

Fig. 1.

To verify that the short-chain PIP₂ diC8 and the long-chain brain PIP₂ share the same binding site, we performed a competition assay. As shown in Fig. 1C, introducing high-affinity brain PIP₂ at the steady state yields a long-lasting potentiation of NCX1 current that is irreversible even after a 5-minute washout. Applying PIP₂ diC8 can steadily decrease the brain PIP₂-potentiated NCX1 current, suggesting that both lipids compete for the same binding site.

Structural insight into PIP₂ binding in NCX1

To reveal the structural mechanism of PIP₂ activation, we tried to obtain the EM structure of HsNCX1 in the presence of the long-chain porcine brain PIP₂. However, the exchanger becomes highly dynamic, yielding a low-resolution EM map with an overall shape similar to a cytosolic Ca²⁺-activated NCX1 whose β-hub-mediated inactivation assembly is destabilized and cytosolic domain (CBD1 and CBD2) is detached from the TM domain³² (Fig. S1). We suspect the long-chain PIP₂ exerts the same activation effect on NCX1 as high cytosolic Ca²⁺ by destabilizing the inactivation assembly. As NCX1 retains its Na⁺-dependent inactivation property in the presence of the short-chain PIP₂, we reasoned that the PIP₂ diC8-bound NCX1 likely remain in an inward-facing inactivated state in high Na⁺ low Ca²⁺ condition and its structure would still reveal how PIP₂ binds in NCX1. We therefore determined the EM structure of NCX1 in complex with PIP₂ diC8 at 3.5Å (Fig. 2, Fig. S2, Table S1, and Methods), which indeed adopts an inward-facing conformation with intact inactivation assembly. Due to the relative dynamic movement between the TM and cytosolic domains, we also performed local refinement to improve the map quality for each domain (Fig. S2). The density of the IP₃ head group from the bound PIP₂ diC8 is well-defined in the local-refined EM map focused on the TM domain (Fig. 2A and Fig. S2B). This density is not present in the apo NCX1 structure (Fig. S3). The acyl chains, however, are flexible and could not be resolved in the structure (Fig. S2). The lipid is attached to the cytosolic sides of TMs 4 and 5 with its head group positioned at the C-terminal end of TM5 (Fig. 2A). Four positively charged residues including K164 and R167 from the N-terminus of TM4 and R220 and K225 from the C-terminus of TM5 are positioned in the vicinity of the PIP₂ head group and likely participate in the electrostatic interactions with the head group (Fig. 2A).

Fig. 2.

PIP₂ diC8-induced conformational changes in NCX1

Two major conformational changes occur in NCX1 upon PIP₂ diC8 binding (Fig. 2B-D). The first change occurs at the C-terminus of TM5 which ends at R220 and is connected to the 2-stranded XIP β-sheet (β3 and β4) via a 6-residue loop in the apo structure. When PIP₂ binds, the TM5 helix extends by one helical turn. This loop-to-helix transition significantly changes the locations of these connecting loop residues and their side-chain orientations to accommodate PIP₂, enabling K225 to reorient and interact with the IP₃ head group (Fig. 2D, top panel). The second conformational change is the partial detachment of the CBD2 domain from XIP upon PIP₂ binding, resulting in a downward swing of cytosolic CBD domains (CBD1 and CBD2) as a rigid body (Fig. 2B&C). This CBD2 detachment is a result of the first PIP₂-induced conformational change at TM5 that disrupts part of the interactions between CBD2 and XIP as follows: In the apo inactivated state, Y226 and R247, the two termini residues of the 2-stranded XIP β-sheet, form H-bonds with several CBD2 residues including E554 sidechain and backbone carbonyls of I518 and I520 (Fig. 2D, bottom panel); The loop-to-helix transition of TM5 upon PIP₂ binding leads to a dramatic rotation of Y226 that allows it to move closer to and directly interact with R247, resulting in the loss of H-bonding interactions between XIP and CBD2 (Fig. 2D, bottom panel); In addition, the rotation of Y226 also causes a direct collision with CBD2 if it remains closely attached to XIP. Thus, the PIP₂-induced TM5 movement, particularly the reorientation of Y226, abolishes some local interactions between CBD2 and XIP and also pushes CBD2 away from XIP, causing the partial detachment of CBD2. However, the short-chain PIP₂ only partially destabilizes rather than completely disassembles the inactivation assembly as the CH2 helix of CBD2 still engages in extensive interactions with the C-shaped β-hub.

To test if the PIP₂-interacting residues play a critical role in the native long-chain PIP₂ activation, we performed mutagenesis to those positively charged residues, including K164A, R167A, R220A, and K225A single mutants. We also generated an R220A/K225A double mutant as these two residues undergo PIP₂-induced conformational change at TM5. While all mutants remain susceptible to current potentiation upon brain PIP₂ application as seen in the wild-type NCX1, the PIP₂-potentiation effects on peak and steady-state currents are weakened in some mutants, most notably in R220A and R220A/K225A (Fig. 3A-C). Interestingly, these two mutants also have reduced Na⁺-dependent inactivation as demonstrated by their higher fractional activity (ratio between stead-state and peak currents) before applying PIP₂ (Fig. 3D). The retained PIP₂ activation on these mutants implies that the longer acyl chain from native PIP₂ plays an important role in its interaction and activation of NCX1. Furthermore, as the interactions between PIP₂ and NCX1 are both electrostatic involving multiple charged residues and hydrophobic involving the long lipid acyl chain, those single or double amino acid substitutions may only decrease the affinity of PIP₂ rather than abolish its binding. To test that, we also mutated all four positively charged residues to alanine. The K164A/R167A/R220A/K225A mutant is no longer sensitive to PIP₂ activation. The currents from this quadruple mutant are small in most recordings and show no Na⁺-dependent inactivation. The unresponsiveness to PIP₂ and lack of Na⁺-dependent inactivation in this mutant is consistent with previous studies demonstrating that PIP₂ activates NCX by tuning the amount of Na⁺-dependent inactivation and any mutation that decreases NCX sensitivity to PIP₂ will also affect the extent of Na⁺-dependent inactivation⁴⁴. This quadruple NCX1 mutant likely abolishes the potentiation effect from both endogenous and externally applied PIP₂ and thereby functions in a Na⁺-inactivated steady state.

Fig. 3.

Structure of NCX1 in complex with SEA0400 inhibitor

SEA0400 is known to potently inhibit cardiac NCX1 by facilitating the inactivation of the exchanger^50–52. To reveal the structural mechanism of its inhibition, we determined the structure of HsNCX1 in complex with SEA0400 at 2.9 Å (Fig. 4, Fig. S4, Table S1, and Methods). The SEA0400-bound NCX1 structure adopts an inward-facing, inactivated state identical to the apo NCX1 structure obtained at high Na⁺, nominal Ca²⁺-free condition ³² (Fig. 4A and Fig. S5). SEA0400 binds at the TM domain in a pocket enclosed by the internal halves of TMs 8 (8a segment), 2 (2ab segments), 4, and 5 (Fig. 4B). The pocket has a lateral fenestration in the middle of the membrane (Fig. 4B), which provides a portal for the SEA0400 entrance. The pocket is sealed off at the cytosolic side by E244 from the XIP β-sheet. As demonstrated in our previous study³², XIP and the linker β-hairpin (β1 and β2) between TMs 1 and 2ab form a β-hub that stabilizes the exchanger in an inactivated state. This β-hub has to be dissembled in an activated NCX1 and XIP is expected to be detached from the TM domain, which would lead to the opening of the SEA400-binding pocket to the cytosol and provide a cytosolic portal for the release of the inhibitor release as further discussed below. Fig. 4C summarizes the interactions between SEA0400 and NCX1 and demonstrates that SEA0400 fits perfectly in the pocket, making extensive contact with surrounding residues. Mutations of some key interacting residues, such as F213 and G833, have been shown to compromise the inhibitor binding⁵². The same SEA0400 binding was also demonstrated in the recent study of human NCX1.3 and mutations at some pocket-forming residues mitigate the SEA400 inhibition of the exchanger³³.

Fig. 4.

Inhibition mechanism of SEA0400

The TM domain of NCX1 shares a similar overall architecture to the archaea exchanger NCX_Mj³⁴. The structural comparison between the TM domains of the inward-facing NCX1 and the outward-facing NCX_Mj reveals the conformational changes that occur during ion exchange, providing structural insight into the SEA0400 inhibition mechanism (Fig. 5A). The inward-outward transition mainly involves the sliding motion of TMs 1 and 6 and the bending movement of TMs 2ab and 7ab^32,34,35,53. As TM2ab directly interacts with SEA0400 in the inward-facing state, its bending movement towards the outward conformation would cause a direct collision with the inhibitor (Fig. 5A). Thus, the binding of SEA0400 stabilizes the exchanger in the inward-facing state and blocks the conformational change from the inward to the outward state. The Na⁺-dependent NCX1 inactivation occurs when the exchanger is in a Na⁺-loaded, inward-facing state with low cytosolic [Ca²⁺]^40,41,54. As demonstrated in our previous study, only in this state, the β-hub can form and readily interact with the cytosolic CBD domains, generating the inactivation assembly that locks TMs 1 and 6 and prevents the TM module from transporting ions³². Thus, SEA0400 promotes NCX1 inactivation by stabilizing NCX1 in the inward-facing conformation which facilitates the formation of the inactivation assembly. The formation of the inactivation assembly also reciprocally stabilizes SEA0400 binding as the XIP of the assembly interacts with the TM domain and seals off the inhibitor binding pocket from the cytosolic side (Fig. 5B). Under conditions in which the inactivation assembly cannot form in NCX1, such as chymotrypsin treatment or forward exchange mode (Na⁺ influx/Ca²⁺ efflux), the removal or detachment of XIP from the TM domain would generate a cytosolic open portal for SEA0400 release and effectively reduce its binding affinity (Fig. 5C)^50,51. Indeed, cysteine scanning mutagenesis studies have shown that the pocket-forming G833 residue is accessible to intracellular sulfhydryl reagents in the chymotrypsin-treated inward-facing NCX1, confirming the cytosolic exposure of the SEA400-binding pocket upon XIP removal⁵⁵. This cytosolic opening of the SEA0400 pocket explains the low efficacy of SEA0400 inhibition in NCX1 when the Na⁺-dependent inactivation is absent.

Fig. 5.

Discussion

In this study, we provide mechanistic underpinnings of cellular regulation and pharmacology of NCX family proteins by revealing the structural basis of PIP₂ activation and SEA0400 inhibition in HsNCX1. Both compounds modulate NCX1 activity by reducing or enhancing the physiologically relevant Na⁺-dependent inactivation process that occurs when the exchanger is in an inward-facing, Na⁺-loaded state with high [Na⁺] and low [Ca²⁺] on the cytosolic side^27,40. In this inactivation state, XIP and β-hairpin can assemble into a TM-associated four-stranded β-hub that mediates a tight packing between the TM and cytosolic domains, resulting in the formation of an inactivation assembly that blocks the TM conformational changes required for ion exchange function^32,33. PIP₂ or SEA0400 binding changes the stability of the inactivation assembly in NCX1, resulting in a reduction or potentiation of inactivation.

Although short-chain PIP₂ cannot fully recapitulate the NCX1 activation effect from long-chain native PIP₂, the structure of PIP₂ diC8 NCX1 allows us to define the lipid-binding site and the lipid-induced conformational changes at the interface between CBD2 and XIP that destabilize the inactivation assembly. The SEA0400-bound NCX1 structure presented here, along with the recent study by Dong et al.³³, suggests that the drug can directly inhibit NCX1 by blocking the inward-outward conformational change at TM2ab. Our structural analysis also explains the strong connection between SEA0400 binding and the Na⁺-dependent inactivation - SEA0400 is ineffective in an exchanger lacking Na⁺-dependent inactivation whereas enhancing the extent of Na⁺-dependent inactivation increases the affinity for SEA0400. As SEA0400 binding traps NCX1 in an inward-facing state that facilitates the formation of the inactivation assembly, the interaction between XIP and the TM domain in the assembly can in turn stabilize SEA0400 binding by sealing the drug-binding pocket from the cytosol. When XIP is removed as in the chymotrypsin-treated NCX1 or when the inactivation assembly cannot form as in the NCX1 at forward exchange mode, the SEA0400-binding pocket becomes exposed to the cytosol, mitigating the inhibition efficacy of SEA0400^50,51.

While we expect the long-chain PIP₂ to bind in the same location as PIP₂ diC8, it is unclear how it exacerbates the destabilization effect on the inactivation assembly. The long acyl chain of native PIP₂ may engage in some interactions with the TM module of NCX1 that are not present in the short-chain lipid, rendering a higher affinity binding and more profound destabilization effect on the inactivation assembly than PIP₂ diC8. The acyl-chain length-dependent PIP₂ activation is consistent with some previous studies. Before PIP₂ was demonstrated to regulate NCX, some earlier studies showed that negatively charged long-chain lipids such as phosphatidylserine (PS) or phosphatidic acid (PA) could have the same potentiation effects on NCX1 as PIP₂^56,57. Furthermore, long-chain acyl-CoAs could also have the same potentiation effects on NCX as PIP₂⁵⁸. All these studies demonstrated that activation of NCX by the anionic lipids depends on their chain length with the short chain being ineffective or less effective. These findings have two implications. First, it is the negative surface charge rather than the specific IP₃ head group of the lipid that is important for stimulating NCX1 activity, suggesting non-specific electrostatic interactions between the negatively charged lipids and those positively charged residues at the binding site. Second, a longer acyl chain is required for the high-affinity binding of PIP₂ or negatively charged lipids. In the PIP₂ diC8-bound structure, the tail of the acyl chain is positioned right at the fenestration of NCX1 that serves as the portal for the SEA0400 binding. Two pieces of evidence lead us to suggest that the tail of the long acyl chain from a native lipid can enter the same binding pocket for SEA0400 and thereby rendering its higher affinity binding than a shorter chain lipid. Firstly, a docking analysis^59,60 of both long-chain and short-chain PIP₂ at the binding site showed that the tail portion of the acyl chain from the native brain PIP₂ inserts into the SEA0400-binding pocket through the fenestration of NCX1 in all docking models with highest calculated affinity (Fig. S6A). This pocket is not reachable for PIP₂ diC8 due to its shorter chain length (Fig. S6B). Second, a stretch of density likely from a native lipid acyl chain is observed in the apo NCX1 structure inside the SEA0400-binding pocket (Fig. S6C), indicating its accessibility to a long-chain lipid. Thus, the accommodation of the long acyl chain in the open pocket of NCX1 through the fenestration in the middle of the membrane likely contributes to the high affinity binding of native lipids.

Methods

Protein expression and purification

The expression and purification of the HsNCX1 (cardiac isoform NCX1.1, indicated as HsNCX1 or NCX1 throughout the manuscript) were carried out as described previously³². Truncated HsNCX1 (Δ341-365aa) containing a C-terminal Strep-tag was cloned into a pEZT-BM vector and baculoviruses were produced in Sf9 cells⁶¹. For protein expression, cultured Expi293F GnTI-cells were infected with the baculoviruses at a ratio of 1:20 (virus: GnTI-, v/v) for 10 hours. 10 mM sodium butyrate was then introduced to boost protein expression level and cells were cultured in suspension at 30°C for another 60 hours and harvested by centrifugation at 4,000 g for 15 mins. All purification procedures were carried out at 4°C. The cell pellet was resuspended in lysis buffer (25 mM HEPES pH 7.4, 300 mM NaCl, 2 μg/ml DNase I, 0.5 μg/ml pepstatin, 2 μg/ml leupeptin, 1 μg/ml aprotinin, and 0.1 mM PMSF) and homogenized by sonication. NCX1 was extracted with 2% (w/v) N-dodecyl-β-D-maltopyranoside (DDM, Anatrace) supplemented with 0.2% (w/v) cholesteryl hemisuccinate (CHS, Sigma Aldrich) by gentle agitation for 2 hours and supernatant collected by centrifugation at 40,000 g for 30 mins was incubated with Strep-Tactin affinity resin (IBA) for 1 hour. The resin was then collected on a disposable gravity column (Bio-Rad) and washed with 30 column volumes of buffer A (25 mM HEPES pH 7.4, 200 mM NaCl, 0.06% digitonin). NCX1 was eluted in buffer A supplemented with 50 mM biotin and further purified by size-exclusion chromatography on a Superdex200 10/300 GL column (GE Healthcare). For the generation of NCX1-Fab 2E4 complex, NCX1 was incubated with purified Fab in a molar ratio of 1:1.2 (NCX1: Fab 2E4) for 2 hours and further purified by size-exclusion chromatography in buffer A. The peak fractions were collected and concentrated to ∼5-6 mg/ml for cryo-EM analysis. To prepare the protein samples in complex with various compounds, 0.9 mM SEA0400, 0.47 mM PI(4,5)P₂ diC8 or 0.42 mM brain PI(4,5)P₂ were added to the protein samples 2 hours before grid preparation.

Expi293F GnTI-cells were purchased from and authenticated by Thermo Fisher Scientific. The cell lines were tested negative for mycoplasma contamination.

Cryo-EM sample preparation and data acquisition

HsNCX1-Fab 2E4 samples (∼5-6 mg/ml) in various conditions were applied to a glow-discharged Quantifoil R1.2/1.3 300-mesh gold holey carbon grid (Quantifoil, Micro Tools GmbH, Germany), blotted for 4.0 s under 100% humidity at 4 °C and plunged into liquid ethane using a Mark IV Vitrobot (FEI). For the SEA0400-bound NCX1-Fab 2E4, raw movies were acquired on a Titan Krios microscope (FEI) operated at 300 kV with a K3 camera (Gatan) at 0.83 Å per pixel and a nominal defocus range of −0.9 to −2.2 μm. Each movie was recorded for about 5 s in 60 subframes with a total dose of 60 e^-/Ų. For other samples, raw movies were acquired on a Titan Krios microscope operated at 300 kV with a Falcon 4i (Thermo Fisher Scientific) at 0.738 Å per pixel and a nominal defocus range of −0.8 to −1.8 μm. Each movie was recorded for 4 s with a total dose of 60 e^-/Ų.

Image processing

Cryo-EM data were processed following the general scheme described below with some modifications to different datasets (Figs. S1, S2, and S4). First, movie frames were motion-corrected and dose-weighted using MotionCor2⁶². The CTF parameters of the micrographs were estimated using the GCTF program⁶³. After CTF estimation, micrographs were manually inspected to remove images with bad defocus values and ice contamination. Particles were picked using program Gautomatch (Kai Zhang, https://www.mrc-lmb.cam.ac.uk/kzhang/) or crYOLO⁶⁴ and extracted with a binning factor of 3 in RELION^65,66. Extracted particles were subjected to 2D classification, ab initio modeling, and 3D classification. The particles from the best-resolving 3D class were then re-extracted with the original pixel size and subjected to heterogenous 3D refinement, non-uniform refinement, CTF refinement, and local refinement in cryoSPARC⁶⁷. The quality of the EM density maps for the transmembrane (TM) and cytosolic domains was further improved through focused refinement, allowing for accurate model building for a major part of the protein. For the dataset of NCX1 in complex with brain PI(4,5)P₂, the maps of apo inactive NCX1 (PDB 8SGJ) and Ca²⁺-bound active NCX1 (PDB 8SGT) are used as references for heterogeneous refinement. Due to the highly dynamic nature of the protein samples, the particles sorted into the active state produce a map with very low resolution. All resolution was reported according to the gold-standard Fourier shell correlation (FSC) using the 0.143 criterion⁶⁸. Local resolution was estimated using cryoSPARC.

Model building, refinement, and validation

The EM maps of HsNCX1 in the SEA0400-bound and PI(4,5)P₂ diC8-bound states show high-quality density and model building is facilitated by previous apo NCX1 structure (PDB 8SGJ)³². Models were manually adjusted in Coot ⁶⁹ and refined against maps using the phenix.real_space_refine with secondary structure restraints applied⁷⁰. The final NCX1structural model contains residues 17-248, 370-467, 482-644, 652-698, 707-718, and 738-935. The EM map of HsNCX1 in complex with brain PI(4,5)P₂ is relatively poor. The Ca²⁺-bound activated NCX1 structure (PDB 8SGT) is directly docked into the EM map without adjustment.

The statistics of the geometries of the models were generated using MolProbity⁷¹. All the figures were prepared in PyMol (Schrödinger, LLC.), Chimera⁷², and ChimeraX⁷³.

Electrophysiological experiments

The wild-type HsNCX1 and its mutants were cloned into a pGEMHE vector and expressed in oocytes for electrophysiological recordings. RNA was synthesized using mMessage mMachine (Ambion) and injected into Xenopus laevis oocytes as described in⁷⁴. Oocytes were isolated from at least 3 different frogs and kept at 18°C for 4-7 days. Outward HsNCX1 currents were recorded using the giant patch technique in the inside-out configuration. Each data point shown in this study represents a recording obtained from a single oocyte. The external solution (pipette solution) contained the following (mM): 100 CsOH (cesium hydroxide), 10 HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 20 TEAOH (tetraethyl-ammonium hydroxide), 0.2 niflumic acid, 0.2 ouabain, 8 Ca(OH)₂ (calcium hydroxide), pH = 7 (using MES, (2-(N-morpholino) ethanesulfonic acid)); bath solution (mM): 100 CsOH or 100 NaOH (sodium hydroxide), 20 TEAOH, 10 HEPES, 10 EGTA(ethylene glycol-bis(β-aminoethyl ether)-N,N,N’,N’-tetra acetic acid) or HEDTA (N-(2-Hydroxyethyl) ethylenediamine-N,N’,N’-triacetic acid) and different Ca(OH)₂ concentrations to obtain the desired final free Ca²⁺ concentrations, pH = 7 (using MES). Free Ca²⁺ concentrations were calculated according to WEBMAXc program and confirmed with a Ca²⁺ electrode.

HsNCX1 currents were evoked by the rapid replacement of 100 mM Cs⁺ with 100 mM Na⁺, using a computer-controlled 20-channel solution switcher. As HsNCX1 does not transport Cs⁺, there is no current and only upon application of Na⁺ does the exchange cycle initiate. Data were acquired at 4 ms/point and filtered at 50 Hz using an 8-pole Bessel filter. Experiments were performed at 35°C and at a holding potential of 0 mV. The effects of the Na⁺-dependent inactivation were analyzed by measuring fractional currents calculated as the ratio of the steady-state current to the peak current (fractional activity). All P values were calculated using an unpaired, two-sided Welch’s t-test.

PI(4,5)P₂ diC8 (Phosphatidylinositol 4,5-bisphosphate diC8, Echelon Bioscience) and brain PI(4,5)P₂ (L-α-phosphatidylinositol-4,5-bisphosphate, Brain, Porcine, Avanti Polar Lipids) were dissolved in water and kept as stock at −20 °C. Immediately prior to recordings, PI(4,5)P₂ was diluted in the bath solution to 10 µM final concentration and perfused cytosolically for the indicated time.

Acknowledgements

Single particle cryo-EM data were collected at the University of Texas Southwestern Medical Center Cryo-EM Facility which is funded by the CPRIT Core Facility Support Award RP170644. Cryo-EM sample grids were prepared at the Structural Biology Laboratory at UT Southwestern Medical Center which is partially supported by grant RP170644 from CPRIT. This work was supported in part by the Howard Hughes Medical Institute (to Y.J.) and by grants from the National Institute of Health (R35GM140892 to Y.J. and R01HL152296 to M.O.), and the Welch Foundation (Grant I-1578 to Y.J.).

Additional information

Data availability

The cryo-EM density maps of the human NCX1 have been deposited in the Electron Microscopy Data Bank under accession numbers EMD-40456 for the SEA0400-bound state and EMD-60921 for the PI(4,5)P₂ diC8-bound state, respectively. Atomic coordinates have been deposited in the Protein Data Bank under accession numbers 8SGI for SEA0400-bound structure and 9IV8 for the PI(4,5)P₂ diC8-bound structure.

Author contributions

J.X. prepared the samples and performed data acquisition, image processing, and structure determination; M.O., S.J., N.A., and W.Z. performed electrophysiology recording; Y.J. supervised the work; all authors participated in research design, data analysis, discussion, and manuscript preparation.