S165F mutation-induced Junctophilin-2 autoinhibition impairs Ca2+ signaling in hypertrophic cardiomyopathy

Li, Zhiwei; Wang, Jingxin; Fang, Xiao; Liu, Haiyang; Liu, Wei; Zhang, Wenqing; Li, Jianchao

doi:10.1038/s42003-025-09244-9

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Published: 25 November 2025

S165F mutation-induced Junctophilin-2 autoinhibition impairs Ca²⁺ signaling in hypertrophic cardiomyopathy

Communications Biology volume 8, Article number: 1846 (2025) Cite this article

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Abstract

Hypertrophic cardiomyopathy (HCM) is a common inherited cardiac disorder often linked to mutations in various genes, including junctophilin-2 (JPH2), a key non-sarcomeric protein essential for forming junctional membrane complexes between the plasma membrane (PM) and endoplasmic/sarcoplasmic reticulum (ER/SR). Deciphering the molecular mechanisms behind these mutations is crucial for the development of effective therapies. In this study, we investigate the pathological effects of the HCM-associated S165F mutation in JPH2. Through integrated structural and biochemical analyses, we demonstrate that the S165F mutation induces an unintended intramolecular interaction in JPH2, disrupting its normal interaction with CaV1.2. This mutation compromises ER-PM junctions and disrupts Ca²⁺ signaling, leading to cellular hypertrophy in COS7 and H9c2 cell models. Furthermore, zebrafish overexpressing JPH2_S165F exhibit cardiac dysfunction, including pericardial edema and reduced heart rate. Notably, treatment with the ryanodine receptor agonist PCB-95 ameliorates these phenotypes, underscoring its potential clinical relevance. These findings offer new insights into the molecular mechanisms of JPH2-related HCM and provide a foundation for exploring novel therapeutic interventions.

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Introduction

Hypertrophic cardiomyopathy (HCM) is the most common inherited cardiac disorder, affecting approximately 1 in 500 individuals globally^1,2. Characterized by abnormal thickening of the heart muscle, HCM can lead to severe clinical complications, including heart failure and sudden cardiac death³. Mutations in over 15 genes have been implicated in HCM, with most pathogenic variants found in genes encoding sarcomeric proteins^4,5. However, mutations in genes encoding non-sarcomeric proteins, such as JPH2, also contribute to the disease, indicating multiple pathways in HCM pathogenesis^6,7. Understanding these mechanisms is crucial for developing novel therapeutic strategies⁸.

JPH2 encodes the junctophilin-2 (JPH2) protein, a member of the junctophilin (JPH) family⁹. JPH proteins are key structural components of the junctional membrane complex (JMC), which anchors the plasma membrane (PM) to the endoplasmic/sarcoplasmic reticulum (ER/SR)^10,11,12. The JMC facilitates bidirectional communication between these membranes, crucial for various cellular processes^13,14. In cardiomyocytes, the JMC organizes Ca²⁺ signaling pathways. Ca²⁺ influx through L-type Ca²⁺ channels (LTCCs) at the transverse (T-) tubule triggers Ca²⁺ release from the SR via ryanodine receptor (RyR) channels, a process known as Ca²⁺-induced Ca²⁺ release (CICR)^15,16. This process is essential for excitation–contraction (E-C) coupling, linking membrane depolarization to myofilament contraction^17,18. JPH proteins mediate the precise positioning of LTCC and RyR channels within the JMC, ensuring effective E-C coupling^19,20.

In mammals, the JPH family consists of four members: JPH1, JPH2, JPH3, and JPH4, all sharing a similar domain architecture²¹. Each JPH protein contains eight MORN (membrane occupation and recognition nexus) motifs at its N-terminal, which can bind to ion channels in the PM and phospholipids^9,22. These motifs are arranged in a 6 + 2 pattern, separated by a serine-rich joining region (JR). The JR might mediate the homo- or heterodimer formation of JPH1 or JPH1/2²³. Following the MORN repeats is a long alpha-helix that, along with the MORN motifs, forms an integral structural unit, as shown in recent structural studies²². A transmembrane helix (TM) at the C-terminal anchors JPH proteins to the ER/SR membrane²⁴. Between the alpha-helix and TM are proline-rich divergent domains, essential for differential RyR recruitment by JPH3 and JPH4²⁵.

JPH2 is the predominant isoform in cardiomyocytes, playing a critical role in Ca²⁺ signaling and cardiac muscle function^20,26. In mouse models, JPH2 knockout leads to embryonic lethality due to heart failure⁹, and shRNA-mediated knockdown in adult mice causes acute heart failure and eventually death within a week^27,28. Human genetic studies link JPH2 mutations to HCM^29,30,31, dilated cardiomyopathy (DCM)^32,33, atrial fibrillation (AF)^34,35, and other cardiac conditions¹⁰. The ClinVar database (https://www.ncbi.nlm.nih.gov/clinvar/) lists hundreds of missense mutations in patients with cardiovascular diseases, including the S165F variant, one of the first HCM-associated mutations identified²⁹. S165F leads to cardiomyocyte hyperplasia and altered Ca²⁺ signaling in cellular models and impairs PKC-mediated phosphorylation and TRPC3 interaction in skeletal myocytes³⁶. However, the exact mechanisms by which S165F causes disease in cardiomyocytes require further investigation.

In this work, we explored the effects of the S165F mutation on JPH2 function. Through biochemical characterization and Alphafold-based structure prediction, we demonstrated that S165F impairs the interaction between JPH2 and CaV1.2 by autoinhibiting the inner groove of the N-terminal MORN repeats, the binding site for the CaV1.2 cytoplasmic tail. We showed that S165F disrupts CaV1.2-JPH2-ER coupling, leading to impaired Ca²⁺ signaling in COS7 and H9c2 cell lines. Additionally, zebrafish overexpressing JPH2_S165F exhibit pericardial edema and a slowed heart rate, a phenotype attenuated by a RyR agonist PCB-95 but exacerbated by a late sodium current inhibitor GS-967. Our study provides mechanistic insights into the HCM-related S165F mutation of JPH2, potentially informing personalized HCM treatments.

Results

Characterization of JPH2-CaV1.2 interactions and the impact of pathogenic mutations

Previous studies have demonstrated that JPH1 and JPH2 can directly interact with a specific stretch of residues at the C-terminal cytoplasmic tails of CaV1.1 and CaV1.2 (Fig. 1a)^22,37. To further validate these findings, we used purified recombinant proteins. Full-length JPH2 is a transmembrane protein that is notoriously difficult to express and purify in a soluble, stable form, often leading to aggregation. Therefore, following the combination of sequence conservation analysis and optimization of constructs, we used the N-terminal cytoplasmic domain (denoted as JPH2 NT, aa 1-440) of mouse JPH2. The JPH2 NT recombinant protein was found to be stable and monomeric in a buffer containing 100 mM MgSO₄. Consistent with earlier reports, our size-exclusion chromatography coupled with multiangle light scattering (SEC-MALS) and isothermal titration calorimetry (ITC) results confirmed that JPH2 NT interacts with residues 1647–1777 (referred to as the JPH binding motif, JBM) at the C-terminal tail of CaV1.2, with a dissociation constant (K_d) of approximately 6.04 μM (n = 3) and a 1:1 stoichiometry (Fig. 1b, c and Supplementary Fig. 1a). Deletion of the JR (Δ176-273) from JPH2 NT (termed JPH2 MORN-Helix) had minimal impact on its interaction with CaV1.2 JBM (Fig. 1b–d). To determine if the interaction with CaV1.2 is a conserved feature across the junctophilin family, we performed both in vitro ITC and cell lysate-based strep pull-down assays using purified recombinant N-terminal domains of mouse JPH1 (JPH1 NT, aa 1-436) and JPH3 (JPH3 NT, aa 1-446). ITC results showed that JPH1 NT and JPH3 NT bind to the CaV1.2 JBM with affinities of 5.54 ± 0.71 µM and 5.82 ± 0.36 µM, respectively (Fig. 1d), which are comparable to that of JPH2. The pull-down results also revealed that a similar amount of GFP-tagged full-length CaV1.2 was pulled down by Strep-tagged JPH1, JPH2, and JPH3 NT fragments (Fig. 1e), further confirming this conserved interaction.

**Fig. 1: Biochemical characterizations of the CaV1.2-JPH interaction.**

A recent structural study indicated that the HCM-related mutation E47A weakens this interaction by disrupting a salt bridge with R1599 of CaV1.1²². Similarly, we found that the DCM-related E85K (corresponding to E85K in mouse JPH2) mutation and the HCM-related A399S (corresponding to A405S in mouse JPH2) mutation in human JPH2 led to a threefold reduction in binding affinity (Fig. 1f). Structurally, E85 forms hydrogen bonds with the backbone amides of β8 and is proximal to K391 on the alpha-helix. Mutating to lysine likely destabilizes the structure. Likewise, A399, located on the alpha-helix, when mutated to serine, affects the packing between the MORN repeats and the alpha-helix (Fig. 1g). Interestingly, we discovered that the HCM-related S165F mutation decreased binding affinity by more than fivefold (Fig. 1f). However, S165 is located in the JR between β13 and β14, far from the CaV1.2 JBM binding site, and was not resolved in previous structural analyses (Fig. 1g). This raises an intriguing question: how does the S165F mutation impact the interaction?

S165F mutation in JPH2 impairs the JPH2-CaV1.2 interaction

We focused on verifying the impact of the S165F mutation on the JPH2-CaV1.2 interaction. To ensure that our observations were not species-specific, we obtained the human version of JPH2 NT and its S165F mutant (Fig. 2a, b). ITC results indicated that wild-type (WT) human/zebrafish JPH2 NT binds to CaV1.2 with an affinity comparable to that of the mouse protein, while the S165F mutation significantly weakened this binding (Fig. 2c and Supplementary Fig. 2). To further confirm the effect of the S165F mutation, we employed various approaches. SEC-MALS results demonstrated that JPH2 NT_S165F and CaV1.2 JBM did not co-elute when mixed at a 1:1 ratio, in sharp contrast to the WT JPH2 NT (Fig. 2d). A Strep pull-down assay showed that Strep-tagged full-length JPH2 WT, but not its S165F mutant, could efficiently pull down full-length CaV1.2 (Fig. 2e). Consistently, purified Strep-tagged CaV1.2 JBM could pull down WT Human/Mouse/Zebrafish JPH2 NT but not the S165F mutant (Fig. 2f, g). Additionally, co-transfection in COS7 cells revealed that WT JPH2, rather than the S165F counterpart, could effectively co-localize with CaV1.2 (Fig. 2h–j). Sequence analysis indicated that the S165 residue is conserved across the JPH family (Fig. 2b). ITC and pull-down results showed that mutating this serine to phenylalanine (S165F for JPH1 and S166F for JPH3) similarly impaired the binding between CaV1.2 and JPH1/JPH3 (Fig. 2c, f, g). Collectively, these findings confirm that the S165F mutation significantly disrupts the JPH2-CaV1.2 interaction.

A potential autoinhibitory mechanism by which the S165F mutation in JPH2 impairs CaV1.2 interaction

An earlier study indicated that the S165F mutation in JPH2 disrupts the JPH2-TRPC3 interaction due to defects in PKC-mediated phosphorylation in skeletal myotubes³⁶. However, our in vitro binding assays clearly demonstrated that S165F impairs the JPH2-CaV1.2 interaction independent of phosphorylation, suggesting alternative mechanisms contributing to this binding defect. To elucidate the underlying mechanism, we tested the binding affinities between CaV1.2 and JPH2 with S165 substituted by each of the other 19 amino acids (Fig. 3a and Supplementary Table 1). Interestingly, mutating S165 to Phe resulted in the largest decrease in binding affinity. S165Y also weakened the binding by about threefold, while substitutions to the hydrophobic residues Leu and Ile led to a two-fold decrease in affinity (Fig. 3a). These observations were further validated by pull-down assays, which showed significantly less of the S165F or S165Y mutant forms of JPH2 NT being pulled down by Strep-tagged CaV1.2 JBM (Fig. 3b, c). These results implied that the Ser at position 165 is not inherently crucial for binding; rather, mutations to aromatic residues Phe or Tyr disrupt the interaction.

**Fig. 3: The S165F mutation-induced specific autoinhibition in JPH2 MORN-Helix.**

By carefully analyzing the crystal structure of the JPH2-CaV1.1 complex, we found that F1605 of CaV1.1 (corresponding to F1703 in mouse CaV1.2) makes extensive hydrophobic interactions with several aromatic residues from JPH2_MORN. Additionally, R1600, located five residues ahead of F1605, forms a salt bridge with D10 (Fig. 3d, g). Intriguingly, R160 in JPH2_JR is also five residues ahead of S165, and the S165F mutation generates an RXXXXF motif (Fig. 3d). We hypothesized that this putative RXXXXF motif in JPH2_JR, created by the S165F mutation, might occupy the CaV1.1/1.2 JBM binding site through an intramolecular interaction. To gain further structural insight, we used AlphaFold3³⁸ to predict the potential intramolecular interaction between JR_S165F (aa 154-173) and the MORN motifs of JPH2. In one of the top five scored predictions, the JR containing S165F binds to the inner groove of MORN1-4 in an antiparallel manner (Fig. 3e, f). In this predicted structure, F165 inserts into the hydrophobic pocket created by the aromatic residues F6, F8, and Y14. R160 forms a charge-charge interaction with the negatively charged D10. Additionally, L163 occupies a position similar to that of L1604 of CaV1.1 in the JPH2-CaV1.1 structure, making hydrophobic interactions with F8 and Y14 (Fig. 3f, g). Consistent with this prediction, mutating R160 to Glu together with S165F (denoted as double mutations, DM) restored the interaction with CaV1.2 to levels comparable to WT, as revealed by both ITC and Co-immunoprecipitation (Co-IP) experiments (Fig. 3h–j). The L163E/S165F mutant form of JPH2 NT bound to CaV1.2 JBM more strongly than the S165F mutant alone but weaker than the WT counterpart, further supporting our hypothesis (Fig. 3h).

ITC measurements failed to detect a direct interaction between JR S165F and the JPH2 MORN-Helix domain (Supplementary Fig. 3a). To test if the S165F mutation creates a novel intramolecular binding site within JPH2, we engineered a “CaV-swap” variant replacing mouse JPH2 residues 156–167 (RSPLRTSLSFLR) with the homologous CaV1.2 sequence surrounding F1703 (DIFRRAGGLFGN, JR-JBM chimera, Supplementary Fig. 3b), preserving the backbone while introducing the critical hydrophobic motif. SEC-MALS confirmed both WT/S165F and JR-JBM chimera remain monomeric with no dimerization (Supplementary Fig. 3c–e). SEC assays demonstrated no co-elution of JR-JBM chimera with CaV1.2 JBM, similar to S165F (Supplementary Fig. 3f, g). ITC showed JR-JBM chimera weakened the binding significantly. The double mutant (R160E/F165S) on the JR-JBM chimera recovers affinity (K_d = 42 µM, Supplementary Fig. 3h), indicating intramolecular sequestration of the binding site.

We also examined the impact of the S165F mutation on known binding partners of JPH2. We performed Co-IP assays in HEK293T cells expressing full-length GFP-tagged RyR2 or TRPC3 with either WT, S165F, or DM JPH2. The results showed that the amount of RyR2 or TRPC3 co-precipitated by S165F and DM JPH2 was not significantly different from that of WT JPH2 (Supplementary Fig. 4a, b), indicating that S165F does not alter JPH2–RyR2 or JPH2-TRPC3 interaction. Likewise, we also observed robust co-localization between TRPC3 and both WT and S165F JPH2 in COS7 cells, further indicating the mutation does not prevent their association in a cellular context (Supplementary Fig. 4c, d).

Taken together, these findings suggested that the S165F mutation impairs binding to the CaV1.2 JBM by creating an unintended autoinhibitory interaction.

The S165F mutation in JPH2 hinders ER-PM junction assembly and Ca²⁺ signaling in COS7 cells

JPHs play a critical role in the assembly of the JMC and the promotion of ER-PM junctions^10,11,12. We previously demonstrated that the S165F mutation in JPH2 disrupts the JPH2-CaV1.2 interaction, while the DM does not. This disruption is expected to impair the formation of ER-PM junctions, leading to reduced ER-PM junction integrity and slow Ca²⁺ mobilization within cells.

To investigate this, we first utilized heterologous COS7 cells. When co-transfected with its β subunit CaVβ1b, CaV1.2 predominantly localized to the PM, with reduced localization to the ER, as shown by immunostaining for the ER marker GRP94 (Supplementary Fig. 5a1, d). However, co-transfection with mCherry-JPH2, but not mCherry alone, resulted in JPH2 co-localizing with CaV1.2 in punctate structures, suggesting an interaction between the two proteins within cells (Supplementary Fig. 5a2, a3, b). These puncta also co-localized with the ER, likely drawing the ER close to the PM and indicating the formation of ER-PM junctions (Supplementary Figs. 5a3, 6a). Consistent with our biochemical findings, co-transfection with JPH2_S165F resulted in the mutant JPH2 co-localizing with the ER but not with CaV1.2, leaving the ER primarily in the cytosol and indicating a failure to form ER-PM junctions due to the disrupted interaction between JPH2_S165F and CaV1.2 (Supplementary Figs. 5a4, b, e–g and 6a). Conversely, the JPH2_DM restored ER-PM junction formation to levels observed with the wild type, as evidenced by the co-localization of JPH2_DM with CaV1.2 and the ER, along with the presence of membrane-localized ER signals (Supplementary Fig. 5a3–a5, e−g, and 6a, b). To sensitively measure JPH2-CaV1.2 proximity, we performed seFRET using GFP-CaV1.2 (donor) and mCherry-JPH2 (acceptor) in COS7 cells. FRET efficiency significantly decreased in the S165F mutant compared to WT, while the DM construct restored it (Supplementary Fig. 6c, d).

To further evaluate the impact of the S165F mutation on Ca²⁺ mobilization in COS7 cells, we utilized the CYB5 ER-TM domain-tethered GCaMP6m biosensor (denoted as GCaMP6m-CYB5), which detects Ca²⁺ changes in a narrow spatial domain on the ER outer surface^39,40. As a control, we demonstrated that co-transfected GCaMP6m-CYB5 co-localized well with the ER and did not affect the localization of CaV1.2 or JPH2 and its mutant forms (Supplementary Fig. 7a). In live-cell imaging experiments, co-transfection with JPH2_WT revealed a significant and prolonged ER Ca²⁺ transient following KCl treatment, characterized by an average amplitude of approximately 2.5 and a full duration at half maximum (FDHM) of around 100 s. In contrast, cells co-transfected with mCherry alone exhibited only small Ca²⁺ changes with an amplitude of ~1 (Supplementary Fig. 7c–e). However, co-transfection with JPH2_S165F resulted in impaired ER Ca²⁺ transients, evidenced by substantial reductions in both amplitude and FDHM. The JPH2_DM, on the other hand, nearly restored the Ca²⁺ transients to levels comparable to the WT, with only a slight decrease in amplitude (Supplementary Fig. 7b–e).

Functional implications of the S165F mutation in JPH2 on ER-PM junction formation and Ca²⁺ signaling in H9c2 cells

To enhance the functional relevance of our study, we conducted similar experiments on H9c2 cells, a rat myoblast cell line frequently used in cardiovascular disease research⁴¹. Western blot analysis confirmed that H9c2 cells do not express detectable levels of endogenous JPH2 under our culture conditions (Supplementary Fig. 8a). This provides an effective JPH2-null background, allowing direct assessment of WT and mutant JPH2 effects without interference from endogenous protein. Consistent with our findings in COS7 cells, JPH2_WT and CaV1.2 co-localized within puncta near the PM and also co-localized effectively with the ER, as indicated by the co-transfected GCaMP6m-CYB5 (Fig. 4a2, b–d). In contrast, JPH2_S165F failed to promote the formation of ER-PM junctions, demonstrated by reduced co-localization of JPH2_S165F/CaV1.2 and CaV1.2/GCaMP6m-CYB5, accompanied by increased cytosolic localization of GCaMP6m-CYB5. However, JPH2_S165F still co-localized with GCaMP6m-CYB5, reinforcing that the S165F mutation specifically impairs the JPH2-CaV1.2 interaction (Fig. 4a3, b–f). Furthermore, H9c2 cells overexpressing JPH2_S165F were significantly larger than those expressing WT JPH2 or mCherry alone. This phenomenon of increased cell size, previously reported²⁹, suggests a potential association with myocyte hypertrophy. Notably, JPH2_DM restored the formation of ER-PM junctions as well as normalized cell size (Fig. 4a, g). Furthermore, we used Structured Illumination Microscopy (SIM) to distinguish closely located but non-co-localized proteins. We observed a clear loss of co-localization between CaV1.2 and JPH2 in the S165F mutant, showing disrupted association at ER-PM junctions (Supplementary Fig. 8b, c). We performed seFRET assays in H9c2 cells. FRET efficiency significantly decreased in the S165F mutant compared to WT (Supplementary Fig. 8d, e).

**Fig. 4: S165F mutation in JPH2 induces defects in H9c2 cells.**

We also examined the impact on Ca²⁺ signaling by recording Ca²⁺ transients using live-cell imaging. Upon treatment with KCl, H9c2 cells expressing JPH2_WT exhibited large and prolonged ER Ca²⁺ transients, marked by high amplitude and FDHM (Fig. 4h, i). In contrast, cells expressing JPH2_S165F showed significantly reduced amplitude and shorter FDHM of the Ca²⁺ signals near the ER, indicating defects in Ca²⁺ signaling. However, cells expressing JPH2_DM exhibited Ca²⁺ transients with amplitude and FDHM values comparable to those of JPH2_WT, suggesting that the DM mutation likely restores normal Ca²⁺ signaling function by re-establishing the interaction with CaV1.2 (Fig. 4h–k).

In summary, these findings demonstrate that the S165F mutation in JPH2 disrupts ER-PM junction formation and impairs ER-PM junction integrity and slow Ca²⁺ mobilization in both heterologous COS7 cells and H9c2 cells, highlighting the potential functional impact of this mutation.

JPH2_S165F induces cardiac dysfunction in zebrafish embryos, which can be relieved by a RyR agonist

The preceding cell biology results prompted us to further investigate the functional impact of the S165F mutation using zebrafish. To assess the effects on cardiac function, we overexpressed EGFP-tagged mouse JPH2_WT or its mutants (S165F or DM) by microinjecting their corresponding mRNAs into one-cell stage zebrafish embryos (Fig. 5a). Successful expression of the mRNAs was confirmed by the presence of EGFP signals throughout the bodies of 2-day postfertilization (dpf) embryos (Supplementary Fig. 9). At 3 dpf, zebrafish overexpressing JPH2_WT did not exhibit any obvious abnormalities compared to non-injected siblings or the EGFP-only injected group, consistent with previous studies showing that overexpression of JPH2 in the mammalian heart affects neither cardiac function nor viability (Fig. 5b1–b3)⁴². On the contrary, zebrafish overexpressing JPH2_S165F displayed significant pericardial edema, as indicated by an average enlarged pericardium (Fig. 5c). Additionally, these zebrafish exhibited slightly shorter body lengths (Fig. 5d). And the average heart rate or blood flow velocity (Fig. 5e, f) was significantly slower in the JPH2_S165F group compared to the non-injected, EGFP-injected, and EGFP-JPH2_WT-injected groups. Moreover, we performed whole-mount in situ hybridization (WISH) at 3 dpf using nkx2.5 as a probe to visualize the developing heart. Our results revealed a striking and statistically significant ~2-fold increase in ventricular area in zebrafish overexpressing JPH2_S165F compared to those expressing WT JPH2 or GFP controls (Fig. 5g, h). To validate our findings, we cloned the zebrafish versions of JPH2 and overexpressed their mRNAs in zebrafish embryos. The phenotypes observed were identical to those obtained with the murine constructs (Supplementary Fig. 10). All these indicated that overexpression of JPH2_S165F may lead to cardiac dysfunction. As anticipated, these abnormalities were not observed in zebrafish overexpressing JPH2_DM, which suggests that the DM mutation mitigates the adverse effects associated with the S165F mutation (Fig. 5b5, c–h).

**Fig. 5: Heart failure in JPH2_S165F-overexpressing zebrafish embryos.**

These observations, showing that zebrafish overexpressing JPH2_S165F exhibited cardiac dysfunction, suggest that it could serve as a potential disease model for HCM in the assessment or discovery of therapeutic drugs (Fig. 6a). We first treated these zebrafish models with two experimentally tested drugs (Fig. 6b2, b3 and Supplementary Fig. 11a2, a3)^43,44: GS-967, a selective late sodium current inhibitor⁴⁵, and MYK-461 (Mavacamten), a small molecule that directly binds to β-cardiac myosin and decreases its ATPase activity^46,47. Unexpectedly, GS-967 treatment worsened the phenotype in JPH2_S165F-overexpressing fish, leading to enlarged pericardia, increased whole-heart area, pronounced developmental defects with markedly shorter body lengths, and a progressively reduced survival rate (Fig. 6c–e and Supplementary Fig. 11), compared with both DMSO-treated counterparts and GS-967-treated siblings or the JPH2_WT group. Conversely, MYK-461 treatment produced no measurable benefit (Fig. 6c–e and Supplementary Fig. 11). Based on our earlier findings that the S165F mutation impairs ER-PM junction integrity in cells, we next treated the fish with PCB-95, a RyR channel agonist, an effect demonstrated in multiple RyR isoforms and mapped to highly conserved inter-domain interfaces near the pore region (Supplementary Fig. 12)^48,49,50. Remarkably, PCB-95 treatment restored the phenotype in JPH2_S165F-overexpressing zebrafish embryos, as evidenced by a smaller average pericardium and an increased heart rate (Fig. 6c, d). As controls, PCB-95 had no effect on the siblings and the JPH2_WT group (Fig. 5c–e and Supplementary Fig. 11). These results underscore the potential of JPH2_S165F zebrafish as a model for studying HCM and for identifying effective therapeutic interventions.

**Fig. 6: PCB-95 treatment rescues the phenotype of JPH2_S165F-overexpressing zebrafish embryos.**

Discussion

Missense mutations in protein-coding regions have been linked to numerous inherited diseases^51,52. Protein structures, either experimentally determined or computationally predicted, offer valuable insights into the impact of these mutations: when the mutation site is located within the folding core, it destabilizes the protein; when it is situated at the interaction interface, it disrupts complex formation. For instance, HCM-related E47A, A399S, and DCM-related E85K mutations in JPH2 can be explained by these structural principles. However, the S165F mutation in JPH2, which is neither in the folding core nor at any known interaction interface, impairs its interaction with CaV1.1/1.2. Our biochemical assays demonstrated that S165F does not affect JPH2 binding to RyR2 or TRPC3, but selectively disrupts its interaction with CaV1.2. This specificity is explained by our autoinhibitory model, in which the S165F-containing joining region folds back to occlude the inner groove of the N-terminal MORN repeats, thereby blocking CaV1.2 binding without interfering with other partners likely located on distinct and spatially separate surfaces of JPH2. This novel finding broadens our understanding of how missense mutations can cause disease beyond the classic mechanisms of protein destabilization and disrupted complex formation. By revealing that mutations can create novel intramolecular interactions leading to functional impairment, our study emphasizes the complexity of genetic disorders. This insight, together with the diverse mechanisms discussed elsewhere⁵¹, underscores the importance of considering these alternative mechanisms in the understanding of genetic diseases.

We also provided cellular analyses to reveal a functional link between the JPH2 S165F mutation, disruption of ER-PM junctional integrity, and altered intracellular Ca²⁺ handling. These results highlight the critical role of JPH2 in maintaining the structural and functional organization of JMC and in safeguarding Ca²⁺ signaling homeostasis. The observed alterations are consistent with a potential contribution of the S165F variant to the development of HCM, supporting the concept that impaired junctional architecture can predispose to maladaptive calcium dynamics underlying disease pathology. However, the present conclusions are based on cellular models that, while useful for delineating mechanistic insights, may not fully capture the physiological complexity of cardiomyocytes. It is therefore necessary to extend these findings to primary cardiomyocytes to assess the direct impact of the S165F mutation on excitation–contraction coupling and contractile performance.

Following this, we generated a zebrafish model exhibiting cardiac dysfunction by overexpressing JPH2_S165F. This model enabled us to demonstrate that certain drugs, such as PCB-95, might potentially attenuate the phenotype, while others, like GS-967, even exacerbated it. Our finding that GS-967 worsened cardiac dysfunction serves as a critical preclinical observation, suggesting that therapies targeting the late sodium current may be contraindicated in HCM patients with specific mutations that impair, rather than enhance, Ca²⁺ signaling at the dyad. This underscores the vital importance of genotype-specific therapeutic strategies. These results highlight the critical importance of understanding detailed disease-causing mechanisms for personalized treatment, as inappropriate therapies can be detrimental. HCM is a complex genetic disorder influenced by multiple pathways^5,44. Our zebrafish model provides a robust platform for screening potential therapeutics and studying their effects in a living organism, thereby facilitating the identification of drugs that can specifically target the molecular mechanisms underlying HCM. This paves the way for more effective and individualized treatments. It is important to note that at present, we have only characterized relatively simple phenotypic readouts. To establish this system as a true disease model, more detailed functional studies will be required, including analyses of contractility defects, arrhythmias, and CICR in the zebrafish heart. Moreover, as overexpression may not accurately mimic the heterozygous state typical of HCM patients, our current model has inherent limitations. Despite this limitation, our current model and future optimized versions based on it offer valuable preliminary insights that will guide further investigation in more complex and clinically relevant systems.

Mutations in JPH2 are commonly implicated in HCM, with many of these mutations located in the MORN repeats^11,12,22. MORN repeats were originally identified in JPHs and were initially thought to bind directly to lipids or membranes^9,53. Our earlier studies suggested that, similar to other repeat-containing modules such as ankyrin repeats or armadillo repeats, MORN repeats can also function as protein–protein interaction modules. For instance, MORN4 utilizes its MORN repeats to bind to unconventional myosin IIIa, serving as its cargo⁵⁴. Similarly, SETD7 employs its MORN repeats to interact with various positively charged transcription factors, acting as a docking site for efficient methylation⁵⁵. As demonstrated in this study and corroborated by others²², JPHs use their MORN repeats to directly interact with CaV1.1/1.2. Notably, the JBM of CaV1.1/1.2 only occupies the inner groove of the N-terminal four repeats. Considering that residues in the inner groove of the entire MORN repeat region are highly conserved through evolution and among the JPH family, and given the spread of mutations along the entire MORN repeat region, it is highly plausible that other constitutive or regulatory components of the JMC might also bind to the MORN repeats of JPHs.

Methods

Constructs and mutagenesis

The cDNA encoding the human JPH2 fragment (residues 1-446) was PCR amplified using the N-terminal Homo_JPH2 cDNA template (purchased from Sangon Biotech, residues 1-446, NCBI accession code: NM_020433.5). The coding sequences of mouse JPH2 full-length (residues 1-696, NCBI accession code: NM_021566.2), mouse JPH1/JPH3 N-terminal fragments (residues 1-436/1-455, NCBI accession code: NM_020604.2/NM_020605.3), mouse CaV1.2 full-length (residues 1-2139, NCBI accession Number: NM_009781.4), mouse CaV1.1 fragment (residues 1552-1672, NCBI accession code: NM_014193.4) and mouse CaVβ1b full-length (residues 1-597, NCBI accession code: NM_145121.4) were PCR amplified using Q5 High-Fidelity polymerase (NEB Lyo Sciences) from a mouse cDNA library. All point mutations (including S165F, R160E/S165F, etc.) were generated by QuikChange-style PCR using Q5 High-Fidelity polymerase (NEB Lyo Sciences, M0492L). All constructs were fully sequence-verified (Sangon Biotech, China) to ensure accuracy. All of these constructs were cloned into a modified pETMBP.3 C/pET32M.3 C vector for protein expression in bacteria or a modified pTTagBFP/pTGFP/pTmCherry vector for expression in COS7/H9c2 cells. All primers used in this study are listed in Supplementary Table 2.

Fusion-protein expression & purification

All recombinant proteins were expressed in Escherichia coli Rosetta (DE3) cells. Bacterial cultures were grown at 37 °C in LB medium supplemented with 50 µg/mL chloramphenicol and 100 µg/mL Ampicillin to an OD₆₀₀ of 0.6–0.8. Protein expression was then induced with 0.2 mM IPTG at 16 °C overnight. Cells were harvested by centrifugation, and pellets were resuspended in lysis buffer containing 20 mM Tris-HCl (pH 7.5), 500 mM NaCl, 5 mM imidazole, and 1 mM PMSF. Lysis was performed by high-pressure homogenization, followed by centrifugation to clarify the lysate. Clarified lysates were applied to a Ni²⁺-NTA Sepharose™ 6 Fast Flow column (Cytiva). After washing, His₆-tagged fusion proteins were eluted with an imidazole gradient. The purification tag was subsequently cleaved with 3 C protease. The proteins were further purified by size-exclusion chromatography (Superdex-200, Cytiva) and, where necessary, an additional step of ion-exchange chromatography (Mono Q or Mono S, Cytiva). Final polishing was performed in a buffer containing 50 mM Tris-HCl (pH 7.5), 1 mM DTT, and either 100 mM NaCl, 1 mM EDTA, or 100 mM MgSO₄, depending on the downstream assay requirements.

Co-immunoprecipitation (Co-IP) assay and immunoblotting

Human HEK293T cells (from ATCC) were cultured in Dulbecco’s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FBS, Gibco). Cells were transiently transfected with 10–20 μg of GFP-tagged CaV1.2 full-length using jetOPTIMUS^® DNA Transfection Reagent (Polyplus). Cells were harvested 36 h post-transfection and lysed in the RIPA or a buffer containing 50 mM Tris (pH 7.4), 150 mM NaCl, 1% Triton X-100, PMSF, and a protease inhibitor cocktail (Roche). Each lysate was incubated with Strep-tagged proteins at 4 °C for 1–2 h.

After extensive washing of the beads with the lysis buffer, bound proteins were boiled in SDS-loading buffer and subjected to SDS-PAGE. Proteins were transferred to a 0.45 μm NC/PVDF membrane (Millipore), which was blocked using 3% bovine serum albumin in TBST (20 mM Tris-HCl (pH 7.4), 137 mM NaCl, and 0.1% Tween-20) buffer at room temperature for 1 h, followed by incubation with the anti-GFP antibody (Abcam, ab290) or anti-HA antibody (Abcam, ab9110) at a 1/10,000 dilution at 4 °C overnight. Membranes were washed three times with TBST buffer, incubated with Goat Anti-Rabbit IgG H&L (HRP) antibody (Abcam, ab205718) at a 1/1000 dilution, at room temperature for 1–2 h, and visualized by an immunohistochemistry kit (DAB detection, Elabscience).

Size-exclusion chromatography coupled with multiangle light scattering (SEC-MALS) assay

Protein samples (typically 200 μL at a concentration of 50–100 μM) were injected into an ÄKTA-SEC system with a Superose^TM 10/300 GL column (Cytiva) using the column buffer of 50 mM Tris-HCl pH 7.5, 100 mM MgSO_4, and 1 mM DTT. The chromatography system was coupled to a static multiangle light scattering system equipped with a 3-angle static light scattering detector (miniDAWN, Wyatt) and a differential refractive index detector (Optilab, Wyatt). The molecular weights were analyzed using the ASTRA 7 software (Wyatt).

Isothermal titration calorimetry (ITC) assay

ITC measurements were carried out on a MicroCal PEAQ-ITC (Malvern) at 25 °C. All proteins were dissolved in 50 mM Tris-HCl (pH 7.5) buffer containing 100 mM MgSO₄ and 1 mM DTT. The concentrations of the protein in the syringe were typically 300 μM, while the concentrations of the protein in the cell were typically 30 μM. Each titration point was performed by injecting a 2 µL aliquot of the syringe sample into the cell sample at a time interval of 120–150 s to ensure that the titration curve returned to the baseline. The titration data were analyzed by MicroCal PEAQ-ITC analysis software (Malvern) and fitted by the one-site binding model.

Strep pull-down assay

Cell lysate-based Strep pull-down: Strep-tagged proteins were bound to 40 µL Strep-Tactin XT beads (IBA Lifesciences) with 1 mg HEK293T lysate in RIPA buffer (50 mM Tris pH 7.4, 150 mM NaCl, 1% Triton X-100) for 2 h at 4 °C, washed three times, and eluted in SDS-loading buffer. After centrifugation, supernatants containing soluble proteins were resolved on a polyacrylamide gel with 10% SDS and analyzed by Western blotting with anti-GFP antibody (Abcam, ab290) or anti-HA antibody (Abcam, ab9110) at a 1/10,000 dilution.

Purified protein-based Strep pull-down: 1 µM MBP-JPH2 NT and 1 µM Trx-Strep–CaV1.2 JBM were mixed, incubated with 40 µL Strep-Tactin beads in binding buffer (50 mM Tris pH 7.5, 100 mM MgSO₄, 1 mM DTT) for 1 h, washed twice, and eluted by boiling in SDS sample buffer. Samples from the pellet fraction were analyzed by SDS-PAGE with Coomassie blue R250 staining.

Band intensities were quantified by densitometry using ImageJ, performed exclusively on the intact protein band, ignoring any lower molecular weight bands. All groups were compared with the JPH WT group by the two-tailed independent-sample t-test in the IBM SPSS Statistics 21 software.

AlphaFold3-based predictions

The structures of JR_S165F (amino acids 154-173) and the MORN motifs of JPH2. (amino acids 1-153, 275-421) were generated with AlphaFold3³⁸ on the AlphaFold Server (https://alphafoldserver.com). All structure figures were prepared using PyMOL (https://pymol.org/).

Cell culture, cell imaging, and data analysis

COS7/H9c2 cells (from ATCC) were cultured in DMEM supplemented with 10% FBS and 50–100 units of penicillin-streptomycin (P/S). Cultured COS7/H9c2 cells were maintained at 37 °C with 5% CO₂. The cell lines were not further authenticated. Cells were tested negative for mycoplasma contamination by cytoplasmic DAPI staining.

Before transfection, 3–4 × 10⁴ cells were plated on 6-well cell culture plates and allowed to adhere to 18–22 mm² 1.5–1.8# glass coverslips overnight. After 8–16 h, cells were transiently transfected using jetOPTIMUS^® DNA Transfection Reagent (Polyplus) following the manufacturer’s instructions. Images were taken at 30–36 h. For immunostaining, ERs were stained with an Anti-GRP94 antibody (Abcam, ab108606). Endogenous and overexpressed levels of JPH2 were assessed by western blot using anti-JPH2 antibody (Abcam, ab79071).

All the images were acquired using a Zeiss LSM 800 laser-scanning confocal microscope, with a 40 × 1 or 63 × 2 NA oil-immersion objective and a pinhole setting of 1 Airy unit. Data were collected from three independent batches of cultures. At least 80–100 fluorescence-positive cells were counted for each group of experiments. All experiments were conducted in a double-blinded fashion. The data from each cell were quantified by Zeiss ZEN 3.3 (blue edition) and analyzed using ImageJ and GraphPad Prism 9.

Z-stack image

For Z-stack collection, the region of interest (ROI) was first identified in the XY plane using fluorescent channels. The Z-stack range was defined to encompass the entire vertical depth of the sample, from the bottom to the top focal plane, typically covering 10 µm in total thickness depending on the sample. Images were acquired at uniform Z-intervals of 0.5 µm to satisfy Nyquist sampling criteria. All channels were acquired sequentially to minimize spectral overlap. Maximum intensity projections were generated from raw stacks using Zeiss ZEN 3.3 (blue edition) for further analysis. Identical settings were used for all samples within a given experiment to ensure comparability.

Membrane vs cytoplasmic localization analysis

Confocal images were analyzed by line-scan, Manders’ coefficients (tM1 and tM2), and Pearson’s co-localization coefficient (>50 cells/condition from three independent experiments, Zen 3.3 Blue). Manders’ coefficients were calculated using Fiji. Pearson’s correlation coefficients were calculated in parallel to assess overall signal correlation. Membrane/cytosolic (M/C) intensity ratios for TagBFP-CaV1.2, mCherry-JPH2, and the ER marker (GRP94 or GCaMP6m-CYB5) were calculated by segmenting “membrane” pixels (≤1.5 µm from the cell edge) and “cytosolic” pixels (>1.5 µm inward) from Zen-detected cell boundaries. Mean fluorescence intensities were measured per region, M/C values computed, and statistical analysis performed by one-way ANOVA (GraphPad Prism 9). n values are reported in figure legends.

The sensitized-emission FRET (seFRET) assay

The seFRET assay was performed to evaluate protein–protein interactions in cells using GFP-CaV1.2 as donor and mCherry-JPH2 WT/S165F/DM constructs as acceptor. Donor-only, acceptor-only, and donor–acceptor co-expressing samples were prepared to enable correction for spectral crosstalk. Fluorescence images were acquired using identical microscope settings across all samples in a Zeiss LSM 800 laser-scanning confocal microscope, including channels for donor emission under donor excitation (I_DD), acceptor emission under donor excitation (I_DA), and acceptor emission under acceptor excitation (I_AA). Background fluorescence was subtracted from each image. Crosstalk correction coefficients were calculated from the single-labeled controls: donor bleed-through ratio (DER = I_DA/I_DD from donor-only samples) and acceptor excitation ratio (AER = I_DA/I_AA from acceptor-only samples). Corrected FRET signals (cFRET) were then calculated using the equation: cFRET = I_DA – (AER × I_AA) – (DER × I_DD). The resulting cFRET values were further normalized as appropriate to compare FRET efficiency across samples.

Super-resolution imaging and data analysis

Super-resolution imaging was performed using a commercial Hessian-SIM system (HIS-SIM, High Intelligent and Sensitive SIM; CSR Biotech, Guangzhou, China) as described previously^56,57. Images were acquired using a 100×/1.5 NA oil-immersion objective (Olympus). For image reconstruction, SIM Pattern Denoise (notch filter) was used to remove periodic artifacts based on the previous description (equation 17 in supplementary information of Huang et al.⁵⁶), with the following parameters: v = 2, a₀ = 1/(2 × σ²), σ = 5. Additionally, a 3D Pattern Denoise filter (equation 11 in Gustafsson et al.⁵⁸, parameter w = 0.3) was used to suppress reconstruction artifacts. Sparse deconvolution was performed according to the method and parameters reported by Zhao et al.⁵⁹. All raw images were reconstructed with identical settings, and the processed images were subsequently analyzed in Fiji software.

KCl stimulation

COS7/H9c2 cells were transfected with TagBFP-CaV1.2-P2A-CaVβ1b, GCaMP6m-CYB5, and either mCherry-JPH2 WT or mCherry-JPH2 Mut as described above. Transfected cells, cultured on glass-bottomed dishes, were then superfused with rodent Ringer solution (10 mM HEPES pH 7.4, 146 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂). Calcium transients were triggered by 150 mM KCl rodent Ringer solution (K⁺ replacing Na⁺, 10 mM HEPES pH 7.4, 150 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂) for 2.5 s. Cells exhibiting clear mCherry-JPH2 and TagBFP-CaV1.2 signals at the periphery were chosen for analysis.

Live-cell calcium imaging

The live-cell imaging setup was stable and well-controlled in order to maintain the cells at 37 °C for GCaMP6m-CYB5 imaging. For confocal Ca²⁺ imaging, time-lapse imaging was performed on an inverted confocal microscope (Zeiss LSM 800) with a 20 × 1 objective and a pinhole setting of 1 Airy unit. The excitation was 488 nm, and the emission was 490 to 660 nm. Images were acquired at 1024 × 1024 pixels, 2 s/frame, and 30 frames/min for a total duration of 600 s.

For data analysis, regions with intensity transitions were cropped manually. The intensity-time curves were normalized by the intensity of the first image. The GCaMP6m-CYB5 signal trace was normalized by the averaged initial baseline fluorescence of the whole focal plane. For analyzing Ca²⁺ transient amplitude, baseline fluorescence (F0) was subtracted from the peak fluorescence (F), then F-F0 was normalized by the baseline value (F0) to give the transient amplitude ΔF/F0 = (F-F0)/F0.

For parametric quantification, we measured the GCaMP6m-CYB5 dynamic change (F/F0), amplitude (ΔF/F0), and full duration at half maximum (FDHM) of each cell. Scatter plots of the ER Ca²⁺ transient amplitude, peak time, and FDHM were generated by GraphPad Prism 9.

Ethical statement and zebrafish maintenance

All experimental procedures involving zebrafish were approved by the Institutional Animal Care and Use Committee of the South China University of Technology (approval number SCUT-2019-079). Wild-type AB strain zebrafish were maintained under a 14-h light/10-h dark cycle at 28.5 °C. Embryos were collected and staged according to Kimmel et al.⁶⁰. Zebrafish embryos were euthanized by immersion in 0.8 mM Tricaine (MS-222, Sigma-Aldrich). All procedures conformed to the guidelines outlined in Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes, as well as the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

Generation of JPH2-overexpression lines and validation

The pT-Sp6-EGFP-JPH2 plasmid is composed of a Sp6 promoter, a full-length JPH2 cDNA fragment obtained via PCR from mouse samples, and the SV40 poly A sequence. GFP-Mus/Danio_JPH2 WT/Mut-pA mRNAs were synthesized using mMESSAGE mMACHINE mRNA transcription-synthesis kits (Ambion). GFP-Mus/Danio_JPH2 WT/Mut-pA mRNA (0.1 ng/embryo) was injected into single-cell wild-type embryos. Overexpression embryos were identified through GFP at 24–48 h. The body length, area of pericardium, heart rate, and blood flow velocity were quantified in 3 dpf embryos.

Whole-mount in situ hybridization in zebrafish embryos

Whole-mount in situ hybridization (WISH) was performed to assess gene expression in zebrafish embryos. Embryos were fixed in 4% paraformaldehyde at 4 °C overnight, dehydrated in methanol, and stored at −20 °C. After rehydration, embryos were permeabilized with proteinase K and hybridized overnight at 65 °C with digoxigenin-labeled antisense nkx2.5 RNA probes. Post-hybridization washes were followed by incubation with alkaline phosphatase-conjugated anti-DIG antibody at 4 °C overnight. Color development was carried out using NBT/BCIP or BM Purple substrates. Embryos were then post-fixed, cleared, and imaged nkx2.5 signal using a stereomicroscope.

Drugs or inhibitors treatment

Embryos at 24 h postfertilization (hpf) were soaked in egg water containing DMSO (Sigma-Aldrich #D8418), GS-967, MYK-461 (selected from TargetMol’s Bioactive Compound Library), or PCB-95 (Macklin P863826). The concentration used was 1 ppm. During the treatment, body length, area of pericardium, and survival rate were quantified. To control for variability, we randomized embryos from the same clutch into treatment and control groups immediately after injection, ensuring matched genetic background and any potential maternal effects.

Statistics and reproducibility

Data from the in vitro Strep Pull down assay and Co-IP assay were expressed as mean ± SD. Data from cell imaging and zebrafish experiments were expressed as a scatter plot.

All datasets were first tested for normality using the Shapiro–Wilk test (α = 0.05). The statistical analysis was performed using a two-tailed independent-sample T-test in the IBM SPSS Statistics 23 software or one-way ANOVA in the GraphPad Prism 9, n.s. not significant, *p < 0.05, **p < 0.01, ***p < 0.001. All experiments related to cell cultures, COS7/H9c2 imaging, and zebrafish experiments were performed in a double-blinded fashion.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All the data relevant to this study can be found within the main text or the Supplementary Materials. Uncropped blot and gel images are provided in Supplementary Figs. 13–16. All underlying source data for the main and Supplementary Fig. graphs are available in Supplementary Data 1.

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Acknowledgements

This work was supported by grants from the National Natural Science Foundation of China (No. 32271270), the Guangdong Basic and Applied Basic Research Foundation (2024B1515040019), and the Key-Area Research and Development Program of Guangdong Province (2023B0303010001) to J.L.

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Authors and Affiliations

The Innovation Centre of Ministry of Education for Development and Diseases, the Second Affiliated Hospital, School of Medicine, South China University of Technology, Guangzhou, China
Zhiwei Li, Jingxin Wang, Wenqing Zhang & Jianchao Li
Department of Hematology, Guangzhou First People’s Hospital, School of Medicine, South China University of Technology, Guangzhou, China
Xiao Fang
Greater Bay Biomedical Innocenter, Shenzhen Bay Laboratory, Shenzhen, China
Haiyang Liu & Wenqing Zhang
Shenzhen Key Laboratory for Neuronal Structural Biology, Biomedical Research Institute, Shenzhen Peking University-The Hong Kong University of Science and Technology Medical Center, Shenzhen, China
Wei Liu

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Zhiwei Li
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Jingxin Wang
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Xiao Fang
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Haiyang Liu
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Wei Liu
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Wenqing Zhang
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Jianchao Li
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Contributions

Z.L. and J.L. designed the study. Z.L., J.W. and X.F. performed the experiments. W.L. and H.L initiated this project. All authors analyzed the results. Z.L., J.W. and J.L. wrote the manuscript with inputs from other authors. W.Z. and J.L. supervised the research. J.L. coordinated the project.

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Correspondence to Jianchao Li.

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Li, Z., Wang, J., Fang, X. et al. S165F mutation-induced Junctophilin-2 autoinhibition impairs Ca²⁺ signaling in hypertrophic cardiomyopathy. Commun Biol 8, 1846 (2025). https://doi.org/10.1038/s42003-025-09244-9

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Received: 12 December 2024
Accepted: 12 November 2025
Published: 25 November 2025
Version of record: 30 December 2025
DOI: https://doi.org/10.1038/s42003-025-09244-9