Essential role of NONO-HOXA1-Wnt axis in cardiomyocyte differentiation

Domain: neuroscience
Published: 2026-01-24
Editorial outcome: major_revision

Abstract
NONO is recognized as a critical molecular scaffold involved in both transcriptional and posttranscriptional regulation. Mutations in NONO are frequently linked to congenital heart diseases (CHDs) in humans. However, the mechanisms by which NONO regulates cardiac development remain elusive. Here, we identified NONO as a pivotal dual-function regulator of cardiomyocyte differentiation in human induced pluripotent stem cells (hiPSCs). NONO deficiency in hiPSCs results in a distinct defect in early cardiomyocyte differentiation. Mechanistically, NONO interacts with HOXA1 and regulates the dynamic expression of key genes during early cardiomyocyte differentiation. ChIP-seq analysis reveals that NONO loss reduces HOXA1 occupancy at target genes, compromising its transcriptional regulation. Additionally, NONO and HOXA1 cooperatively activate the Wnt signaling. Taken together, these findings establish the NONO-HOXA1-Wnt axis as a key molecular mechanism in cardiomyocyte differentiation and provide insights into the etiology of CHDs associated with NONO mutations.

Introduction
Congenital heart disease (CHD) is the most common birth defect, affecting ~1% of live births and leading to significant morbidity and mortality1,2. Heart development requires the precise regulation of gene expression, primarily via cardiac transcription networks governing early cellular differentiation and later cellular maturation3. These networks cooperate to finely tune the regulatory machinery for gene expression, which is essential for the proper spatiotemporal control of cardiac morphogenesis and provides the foundation for proper cardiac physiology4,5.

The non-POU domain-containing octamer-binding protein (NONO) is characterized as an important molecular chaperone that is best known for its association with molecular machinery involved in both transcriptional and posttranscriptional regulation. NONO is widely expressed and is encoded by an X-linked gene that contains two RNA recognition motifs, a NonA/paraspeckle domain, and coiled-coil and intrinsically disordered regions at the N- and C-termini6,7. This unique structure enables NONO to bind both DNA and RNA, facilitating its roles in transcription regulation, mRNA splicing, and DNA double-strand break repair8–12. Our previous work demonstrated that NONO recruits 10‒11 translocation 1 (TET1) to specific genomic loci in mouse embryonic stem cells (mESCs), thereby modulating 5-hydroxymethylcytosine (5hmC) levels and playing a critical role in neuronal differentiation13. More importantly, in the clinic, dysfunctional variants of NONO have been identified in a broad spectrum of CHDs and cardiomyopathies, including atrial septal defect (ASD), ventricular septal defect (VSD), pulmonary stenosis (PS), right aortic arch (RAA), Ebstein anomaly, and left ventricular noncompaction (LVNC)14–17, particularly in male patients. The birth rate of Nono-KO mice is significantly lower than that of wild-type littermates, and for those that survive, Nono-KO mice exhibit impaired diastolic cardiac function, leading to heart failure and early lethality18,19. Despite these previous observations, the biological role and underlying molecular mechanisms by which NONO regulates cardiogenesis and cardiomyocyte (CM) differentiation are poorly understood.

In this study, we tested the hypothesis that NONO is critical for heart development by employing hiPSC-directed CM differentiation as a model system. Our findings revealed that NONO is essential for proper CM differentiation. We demonstrated that the DNA-binding domain of NONO directly interacts with the homeobox domain of homeobox A1 (HOXA1), a master transcription factor for establishing the anterior‒posterior axis during embryonic development20. This NONO-HOXA1 interaction facilitates the association of HOXA1 with chromatin and promotes the expression of genes in the cardiac and Wnt pathways, such as MESP1, PDGFRA, MIXL1, and WNT5A. NONO deficiency impairs this interaction, leading to the disrupted regulation of the Wnt signaling pathway and abnormal CM differentiation. Overall, our study reveals the NONO-HOXA1-Wnt axis as a critical mechanism driving CM differentiation and provides insights into the potential pathogenesis of CHDs and cardiomyopathies caused by NONO dysfunctional variants in humans.

Results
NONO depletion impairs CM differentiation
Previous studies have established that the mutations in NONO are frequently associated with various types of CHDs17. To elucidate the mechanisms underlying how NONO mutations cause CHDs, we employed an episomal vector-based CRISPR/Cas9 (Epi-CRISPR) gene-editing approach to generate NONO knockout human induced pluripotent stem cells (hiPSCs), and two NONO-KO hiPSC lines were obtained (Supplementary Fig. 1A, B). Karyotype analysis of WT-hiPSCs and NONO-KO clones (3 and 11) confirmed stable, normal chromosomal integrity (Supplementary Fig. 1C). As all NONO-KO hiPSCs presented similar phenotypes, NONO-KO clone-3 cells were used in subsequent analyses. These NONO-KO hiPSCs retained normal pluripotency (Supplementary Fig. 1D, E). We then differentiated these cells into CMs following previously established protocols (Supplementary Fig. 2A)21. First, we analyzed the temporal expression patterns of key cardiac differentiation factors, including OCT4, T, EOMES, MESP1, ISL1, GATA4, PDGFRA, NKX2-5, MYH6, MYH7, and TNNT2 within our in vitro cardiac differentiation system using qRT‒PCR (Supplementary Fig. 2B). Day 15 scRNA-seq revealed that TNNT2+ CMs exhibited high MYL7 (73.9%) and low MYL2 (0.4%) expression (Supplementary Fig. 2C), consistent with an immature, atrial-like phenotype as previously reported21,22. Expression analysis of cardiac lineage markers from Day 2 to Day 7 revealed that NONO-KO hiPSCs exhibited substantially reduced induction of T, EOMES, MIXL1, MESP1, PDGFRA, GATA4, NKX2-5, and MYOCD compared with WT cells, highlighting the essential role of NONO in early CM differentiation (Supplementary Fig. 2D). Flow cytometry demonstrated reduced proportions of PDGFRA+ (Day 3), GATA4+ (Day 5), and NKX2-5+ (Day 6) cells in NONO-KO cultures compared with WT (67.0% vs. 83.5%; 18.3% vs. 35.0%; 41.2% vs. 77.2%) (Supplementary Fig. 2E–J). At Day 15, the percentage of cTNT+ CMs was diminished in NONO-KO cells (11.6%) relative to WT (70.9%) (Fig. 1A), about a 59.3% reduction in CM differentiation. Consistent with the notion, qRT‒PCR analysis further revealed largely reduced expression levels of major myofibril genes, such as ACTN2, TNNT2, MYL2, MYL7, MYH6, MYH7, TITIN, TNNI1, and TNNI3, in NONO-KO CMs (Fig. 1B).

Immunofluorescence analysis revealed that ~80% of NONO-KO CMs on Day 15 presented disorganized sarcomere structures, in contrast with the well-organized sarcomeres observed in WT CMs (Fig. 1C, D). Transmission electron microscopy (TEM) confirmed those findings, revealing a lack of Z-disc structures in NONO-KO CMs, whereas WT CMs exhibited well-defined sarcomeres (Fig. 1E). Compared with WT CMs, NONO-KO CMs presented irregular contraction patterns and a reduced beating rate (Fig. 1F). To investigate these irregularities, we assessed spontaneous Ca2+ transients, a critical physiological marker closely linked to CM-beating rates and myocardial functionality. NONO-KO CMs displayed irregular calcium transients (Fig. 1G right). We developed an R-based script for analyzing and visualizing calcium transient signals (see Methods for details) (Fig. 1H), which clearly highlighted aberrant patterns in NONO-KO CMs. These abnormalities were characterized by a significantly prolonged decay time, an increased time to reach peak and an extended peak-to-peak duration (Fig. 1I). Consistent with these observations, qRT‒PCR analysis revealed reduced expression levels of key calcium-handling genes, including RYR2 (calcium release), SERCA2a (calcium reuptake), and CACNA1C (calcium influx)23 (Fig. 1J).

Differentiation of hPSCs into CMs generates cultures containing multiple lineage intermediates alongside the predominant CM population22. To examine how the loss of NONO affects these developmental trajectories, we performed time-course single-cell RNA-seq on WT and NONO-KO cells collected at Days 2, 5, and 15 of differentiation. UMAP embedding resolved major developmental populations—including mesendoderm, cardiac progenitors, CMs, endoderm, epithelial, endothelial, and neural progenitor cells—which were consistently identified across samples based on canonical marker expression (Supplementary Fig. 3A, B). Cells from all replicates were well integrated, indicating minimal batch effects (Supplementary Fig. 3C). Monocle 224 trajectory analysis further demonstrated a continuous differentiation path from mesendoderm to cardiac progenitors and ultimately CMs in WT cells. However, NONO-KO cells showed hindered progression along the cardiac trajectory, with a subset retaining mesendoderm-like features at Day 5. Cardiac progenitor markers such as ISL1, GATA4, HAND1, and TBX20 exhibited attenuated induction, accompanied by inappropriate persistence of POU5F1, resulting in a markedly lower CM yield at Day 15 (Fig. 1K, L, Supplementary Fig. 3D). CytoTRACE25 scores showed that NONO-KO cells remained less differentiated than WT cells at both CPC and CM stages (Fig. 1M). Collectively, our results highlight that NONO is essential for CM differentiation. NONO deletion reduces differentiation efficiency, which is characterized by significantly fewer cTNT-positive cells, and disrupts CM function, as demonstrated by disorganized sarcomeres and impaired calcium handling in the low percentage of remaining differentiated CMs. Single-cell analyses show that NONO is required for the timely exit from the mesendoderm state. In the absence of NONO, Day 5 progenitors retain mesendoderm signatures and exhibit attenuated expression of key cardiac regulators, thereby failing to acquire the transcriptional competence necessary for efficient CM production.

Reactivation of NONO expression in NONO-KO hiPSCs rescues CM differentiation
To determine whether reactivating NONO can rescue CM differentiation defects in NONO-KO hiPSC, we utilized a PiggyBac (PB) transposon-based doxycycline-inducible system to reactivate NONO expression in NONO-KO hiPSC lines26. To mimic the natural expression pattern of NONO during differentiation (Supplementary Fig. 4A), doxycycline was administered from Days 1–2 of differentiation (Fig. 2A). The generated NONO-RE (NONO-KO NONO OE with Dox) clones exhibited inducible NONO expression upon doxycycline treatment. Compared with wild-type cells, NONO-RE cells showed only a partial rescue at Day 2, as confirmed by Western blot analysis (Supplementary Fig. 4B, C). To assess whether NONO reactivation could restore early cardiac specification in NONO-KO cells, we examined the expression of lineage markers from Day 2 to Day 7 by qRT-PCR. NONO reactivation reinstated the induction of T, EOMES, MIXL1, MESP1, PDGFRA, GATA4, NKX2-5, and MYOCD (Fig. 2B, C). By Day 15, flow cytometry analysis of differentiated cells revealed a significant improvement in the proportion of cTNT-positive cells among NONO-RE (29.5%) compared with that observed among NONO-KO NONO OE without Dox (4.07%), ~25.43% enhancement in CM differentiation (Fig. 2D, E). Immunofluorescence analysis revealed a significant improvement in sarcomere formation in NONO-RE CMs, and TEM revealed the restoration of Z-line structures (Fig. 2F–H). qRT‒PCR analysis of the expression levels of major sarcomeric genes also demonstrated marked recovery (Fig. 2I).

To confirm those observations, we performed bulk RNA-sequencing using WT, NONO-KO and NONO-RE CMs on Day 15 of differentiation. A 9-square plot was used to determine potential genes that are directly associated with NONO-mediated regulation (see Methods for details). This in silico analysis, in which genes were categorized by a log2 (1.5-fold changes) cutoff, revealed that the reactivation of NONO expression restored the expression of a significant number of NONO-modulated genes (Groups G and C) (Supplementary Fig. 4D), including essential cardiac genes such as ACTN2 and TNNT2 (Supplementary Fig. 4E). Gene Ontology (GO) enrichment analysis of these NONO-modulated genes highlighted significant enrichment in biological processes related to muscle contraction and cardiac muscle development (Supplementary Fig. 4F). Corresponding heatmaps illustrated the expression distribution of these genes, emphasizing the role of NONO in modulating genes essential for CM differentiation and function (Supplementary Fig. 4G, H). These findings reinforce the notion that NONO plays a critical role in CM differentiation.

Pathogenic NONO variants fail to rescue CM differentiation defects
To evaluate the functional impact of clinically identified NONO variants, we reconstituted NONO-KO hiPSCs with doxycycline-inducible wild-type NONO (NONO-KO WT-NONO OE) or four patient-specific truncation mutants (p.Arg73*, p.Arg184*, p.Arg337*, and p.Arg365*). Immunofluorescence staining for α-ACTININ and cTNT at Day 15 showed that re-expression of WT-NONO restored well-organized sarcomeric structures. In contrast, cells expressing the mutant variants displayed highly disorganized myofibrils and remained comparable to uninduced NONO-KO controls (Supplementary Fig. 5A, B).

Consistent with these morphological observations, flow cytometry analysis showed that WT-NONO robustly rescued CM differentiation, increasing the cTNT+ population to ~33.5% compared to ~4.0% in the −DOX control. In contrast, none of the four mutants improved differentiation, as cTNT+ fractions remained low (8.8%–11.9%) (Supplementary Fig. 5C, D). qRT-PCR further confirmed that WT-NONO, but not the mutant variants, restored expression of TNNT2, MYH6, and MYH7 (Supplementary Fig. 5E). Collectively, these findings confirm that these clinical truncating mutations result in a loss of NONO function, thereby impairing CM differentiation.

NONO dynamically regulates the expression of genes essential for CM differentiation
To investigate the molecular mechanisms underlying the effects of NONO depletion and subsequent rescue during CM differentiation, we assessed NONO expression levels via qRT‒PCR. The results revealed that NONO expression peaked on Day 2, suggesting a critical role in the early stages of differentiation (Supplementary Fig. 4A). On the basis of these findings, we further analyzed mRNA-seq data of WT, NONO-KO, and NONO-RE hiPSCs at Days 2, 5, 9, and 15 of differentiation. Based on the 9-square plot to categorize genes on the basis of log2 (1.5-fold change) in expression from Day 2 to Day 15 in WT (x-axis) and NONO-KO (y-axis) cells, the genes in Group I (n=332) were upregulated during differentiation in the WT cells but downregulated in the NONO-KO cells, suggesting that these genes, which are normally activated during CM differentiation, were suppressed by NONO loss. In contrast, Group A genes (n=565) presented the opposite pattern, indicating that dysregulation was caused by NONO depletion (Fig. 3A).

We then compared the gene expression levels in NONO-RE (x axis) and NONO-KO (y axis) cells between Day 2 and Day 15 of differentiation (Fig. 3B and Supplementary Fig. 6A). Among the 332 genes in Group I (Fig. 3A), 60 genes (18%) in Group I’ (Fig. 3B) presented restored expression in NONO-RE cells. This suggests that the dynamic expression of these genes during differentiation is dependent on NONO. To validate this dynamic regulation, we analyzed the expression trajectories of Group I’ genes from Day 2 to Day 15. The results revealed that the expression of these genes was consistently downregulated in NONO-KO cells but upregulated by Day 15 in NONO-RE cells, a finding that aligns closely with the expression patterns observed in WT cells (Fig. 3C). Gene Ontology biological process (GO-BP) enrichment analysis revealed that Group I’ genes were significantly associated with cardiac muscle cell contraction processes (Fig. 3D). Notably, the expression of key cardiac genes, such as GATA4, RYR2, KCNE2, and HCN4, was restored in NONO-RE cells (Fig. 3E).

We then employed a 9-square plot to determine potential genes directly associated with NONO-mediated regulation during the early differentiation stages (Days 2 and 5). This analysis revealed that the reactivation of NONO expression restored the expression of a significant number of NONO-modulated genes (Groups G and C), including essential cardiac mesodermal and Wnt canonical genes such as PDGFRB, MESP1, HAND1, WNT3A, and WNT11 (Supplementary Fig. 6B, C). The same analysis of upregulated and downregulated genes measured at different time points during CM differentiation (Days 5 and 2) revealed significant enrichment in cardiac ventricle morphogenesis (Supplementary Fig. 6D–G), emphasizing the pivotal role of NONO in regulating genes crucial for CM function. These findings demonstrate that NONO dynamically modulates the expression of genes critical for CM development.

NONO regulates transcriptional activity independently of its role in pre-mRNA splicing function
Previous studies have identified NONO as a key regulator of both transcription and alternative splicing during retinal development27. In our experimental system, NONO deletion reduced cTNT-positive cell numbers and disrupted sarcomere structure and calcium handling, leading to impaired function. To determine whether these phenotypes are driven by transcriptional or splicing defects, we analyzed differentially expressed genes (DEGs) and alternative splicing events (ASEs) between WT and NONO-KO cells during differentiation (Supplementary Fig. 7A, B). We then examined the overlap between DEGs (4267), ASEs genes (242), and NONO-targeted genes (2212) on Day 2 of differentiation. NONO-targeted genes identified through NONO ChIP-seq were intersected with NONO-KO RNA-seq data, yielding 556 transcriptionally regulated genes. Additionally, 62 alternatively spliced genes were identified by intersecting NONO-KO RNA-seq data with splicing gene datasets. Among these genes, 14 were dually regulated by NONO through transcription and alternative splicing (Fig. 4A). Notably, the 542 transcriptionally regulated genes were enriched in heart development pathways, whereas the 48 genes exclusive to alternative splicing showed no significant functional enrichment (Supplementary Fig. 7C–F). Our results suggest that NONO regulates transcriptional activity independently of its splicing function, with these two roles likely influencing distinct cellular processes during early differentiation.

NONO directly interacts with HOXA1
To investigate how NONO regulates the expression of genes involved in CM development, we examined its potential interactions with transcription factors critical for cardiac differentiation. Transcription factor networks play essential roles in the precise regulation of gene expression during cardiac development4,28. Previous studies have demonstrated that NONO participates in neuronal differentiation and hormone regulation as part of a complex13,29. We performed immunoprecipitation‒mass spectrometry (IP‒MS) analysis of NONO using Day 2 differentiated cells, corresponding to peak NONO mRNA expression. The results led to the identification of core components of the NONO complex, including PSPC1, NONO, and SFPQ. Notably, the transcription factor homeobox A1 (HOXA1), known to be involved in cardiac development, was also identified as part of the NONO complex (Supplementary Fig. 8A). HOXA1 is an essential member of the Hox gene family and is pivotal for establishing the anterior‒posterior axis during embryonic development20. Hox proteins typically function in conjunction with cofactors to precisely regulate gene expression30, and HOXA1 genetic mutations are associated with heart defects31,32. HOXA1 genes are not expressed in ES cells but are activated during differentiation33,34. Consistently, both NONO and HOXA1 exhibited synchronous increases in expression levels on Day 2, suggesting a coordinated expression pattern (Supplementary Fig. 4A, 8B). We validated the interaction between HOXA1 and NONO via co-immunoprecipitation (Co-IP) using HEK293T cells in which NONO-FLAG and HOXA1-HA were overexpressed. Co-IP results confirmed that HOXA1 coprecipitated with NONO and vice versa (Fig. 4B). To investigate the relevance of this interaction during differentiation, we generated a doxycycline-inducible HOXA1-HA overexpression hiPSC line (WT HOXA1-HA OE), in which HOXA1-HA was induced during Day 1 to Day 2, using the PiggyBac transposon system (Supplementary Fig. 8C). Co-IP of Day 2 differentiated cells with doxycycline-induced HOXA1-HA expression confirmed that endogenous NONO coprecipitated with HOXA1-HA, reinforcing the interaction between NONO and HOXA1 in the context of differentiation (Fig. 4C). Immunofluorescence analysis with anti-NONO and anti-HA antibodies demonstrated the nuclear colocalization of NONO and HOXA1 in HEK293T cells (Fig. 4D) and Day 2 differentiated cells (Fig. 4E). Using a GST-pull-down assay, we confirmed that the GST-NONO fusion protein specifically binds to HOXA1-HA from HEK293T cell lysates (Fig. 4F and Supplementary Fig. 8D). These results indicate that NONO interacts with HOXA1 in the nucleus.

To identify the structural domains mediating the interaction between HOXA1 and NONO, we generated HOXA1 deletion mutants targeting three key regions: ΔPH (Poly-Histidine region deletion), ΔHP (Hexapeptide motif deletion), and ΔHD (Homeodomain deletion). In parallel, we constructed NONO deletion mutants lacking major functional domains: ΔRRM1 (RNA Recognition Motif 1 deletion), ΔRRM2 (RNA Recognition Motif 2 deletion), and ΔCoil (Coiled-coil domain deletion) (Fig. 4G). Co-IP analysis in HEK293T cells co-expressing NONO and HA-tagged HOXA1 fragments showed that ΔPH and ΔHP retained NONO binding, whereas deletion of the homeodomain (ΔHD) markedly impaired the interaction (Fig. 4H). Co-IP assays using FLAG-tagged NONO fragments further revealed that deletion of either RRM2 or the coiled-coil domain substantially reduced binding to HOXA1-HD (Fig. 4I). Together, these results demonstrate that the HOXA1 homeodomain and the NONO RRM2 and coiled-coil domains are essential for mediating the NONO-HOXA1 interaction. To explore the structural basis of this interaction, we performed molecular modeling via the AlphaFold2 multimer35. The prediction indicated that residues N230, I275, and K286 within the HOXA1, together with E278, Q281, and R287 in NONO, form key hydrogen-bonding contacts that are critical for the NONO-HOXA1 interaction (Fig. 4J). To validate these predictions, we mutated these residues to alanine (HOXA1-HD: N230A, I275A, K286A; NONO: E278A, Q281A, R287A) to disrupt hydrogen bond formation. Co-IP assays in HEK293T cells co-expressing NONO and GFP-tagged HOXA1-HD mutants showed that the K286A substitution markedly reduced NONO binding, indicating that K286 in HOXA1 is essential for the interaction (Fig. 4K). Similarly, Co-IP analyses using GFP-tagged HOXA1-HD and FLAG-tagged NONO mutants demonstrated that the E278A mutation substantially diminished binding to HOXA1-HD, establishing E278 in NONO as a critical determinant of the NONO-HOXA1 interaction (Fig. 4L). These findings demonstrate that the interaction between HOXA1 and NONO requires key residues within both proteins: K286 in the HOXA1 and E278 within the NONO coiled-coil region. Together, these results establish a direct and residue-specific interaction between HOXA1 and NONO.

Given that NONO has been proposed to function as a protein chaperone, we examined whether it regulates HOXA1 protein stability. We generated NONO-KO HEK293T cells using the Epi-CRISPR system (Supplementary Fig. 1H). To examine whether NONO regulates HOXA1 protein stability, we overexpressed HOXA1-HA in WT and NONO-KO HEK293T cells and conducted cycloheximide (CHX) chase assays. We observed that HOXA1-HA decayed significantly faster in NONO-KO cells than in WT controls, indicating that NONO is required to maintain HOXA1 stability (Fig. 4M). To further elucidate the underlying mechanism, we co-expressed HOXA1-HA with either WT-NONO or the E278A mutant in WT HEK293T cells. In contrast to WT-NONO, the E278A variant failed to stabilize HOXA1 (Fig. 4N).

These results demonstrated that NONO binds to the HOXA1 homeodomain via its RRM2 and coiled-coil domains, with residues E278 of NONO and K286 of HOXA1 being critical for this interaction. Functionally, NONO acts as a chaperone to stabilize HOXA1 protein, a process strictly dependent on their physical association.

NONO and HOXA1 co-occupy genes that are critical for precardiac mesoderm differentiation
To explore the genomic landscape regulated by the NONO-HOXA1 axis during CM differentiation, we performed ChIP-seq in Day 2 differentiated cells. NONO profiling identified 4350 peaks, which were markedly enriched in promoter regions (53.55%) (Supplementary Fig. 8E) and targeted key early cardiac and mesoderm genes such as EOMES, TBXT, DVL2, and MECOM36,37 (Fig. 5A). Gene Ontology (GO) analysis revealed the significant enrichment of NONO binding to genes involved in Wnt signaling, embryonic development regulation, and cardiac septum formation (Fig. 5B). Parallel HOXA1 ChIP-seq identified 26,143 peaks, predominantly located in intergenic and intronic regions (Fig. 5A, Supplementary Fig. 8F). GO analysis of HOXA1 target genes revealed enrichment in pathways related to Wnt signaling and cardiac development (Supplementary Fig. 8G). Bioinformatics analysis revealed 1169 overlapping peaks between the NONO and HOXA1 ChIP-seq datasets (Fig. 5C and Supplementary Fig. 8H), suggesting a shared chromatin association. GO analysis of these shared peaks revealed enrichment in processes such as the Wnt signaling pathway, gastrulation, and heart morphogenesis, suggesting a cooperative regulatory role for NONO and HOXA1 (Fig. 5D).

To further determine whether NONO and HOXA1 co-occupy the same genomic loci, we performed sequential ChIP (Re-ChIP) assays using a doxycycline-inducible HOXA1-HA line. HA-ChIP material was competitively eluted with HA peptide to preserve protein–DNA complexes before a second immunoprecipitation with anti-NONO. As visualized in the genome browser tracks, distinct Re-ChIP signals were successfully detected at the regulatory regions of key early cardiac and mesoderm drivers (Fig. 5A). These Re-ChIP peaks align precisely with the overlapping regions observed in the individual ChIP-seq profiles. Taken together, these findings suggest that NONO and HOXA1 co-occupy genomic regions associated with critical biological processes, including Wnt signaling, gastrulation, and heart morphogenesis, underscoring their cooperative role in CM differentiation.

NONO is indispensable for the HOXA1-mediated regulation of precardiac mesoderm genes
Given that HOX transcription factors require cofactors for stable and specific DNA binding38, we hypothesized that NONO and HOXA1 cooperate to regulate precardiac mesoderm genes. To test this hypothesis, we induced HOXA1-HA overexpression in WT HOXA1-HA OE and NONO-KO HOXA1-HA OE cells using Dox, and western blotting confirmed consistent overexpression efficiency (Supplementary Fig. 9A). HOXA1 ChIP-seq with an anti-HA antibody subsequently revealed 26,143 binding peaks in WT cells and 19,293 peaks in NONO-KO cells, with NONO-KO resulting in the loss of 12,483 HOXA1 binding peaks (Fig. 5E and Supplementary Fig. 9B, C). The loss of HOXA1 binding to target genes, including MESP1, PDGFRA, HAS2 and MIXL1 (Fig. 5F, G and Supplementary Fig. 9D, E), was closely linked to cardiac lineage development (Supplementary Fig. 9F). To determine whether the loss of these 12,483 HOXA1 binding peaks affects gene expression, we generated HOXA1-KO hiPSCs using the Epi-CRISPR system (Supplementary Fig. 1F, G), without an obvious effect on pluripotency (Supplementary Fig. 1D, E). We then performed a cross-analysis between the target genes associated with the 12,483 HOXA1 binding peaks lost upon NONO-KO and the DEGs identified in HOXA1-KO cells versus WT cells on Day 2 of differentiation. This analysis revealed 155 upregulated and 304 downregulated genes, indicating that these HOXA1-regulated genes are associated with HOXA1 promoter binding, which is dependent on NONO (Supplementary Fig. 9G). These DEGs were significantly enriched in cardiac development-related terms and the Wnt signaling pathway (Fig. 5H). Notably, 13 genes associated with cardiac development-related terms were also downregulated in NONO-KO cells (Supplementary Fig. 9H). qRT‒PCR validation further confirmed the downregulation of these key precardiac mesoderm genes (MESP1, PDGFRA, HAS2, and MIXL1) in both HOXA1-KO and NONO-KO cells on Day 2 of differentiation (Fig. 5I and Supplementary Fig. 9I).

To elucidate the molecular mechanism underlying the impaired HOXA1 genomic occupancy in NONO-deficient cells, we examined the oligomeric states of these proteins. Using doxycycline-inducible systems, Co-IP assays first confirmed that NONO forms homodimers in differentiated cells (Fig. 5J). Crucially, we investigated whether NONO influences HOXA1 self-association. Co-IP analysis revealed that HOXA1 homodimer formation was markedly compromised in NONO-KO cells compared to WT controls (Fig. 5K, L). This suggests that NONO facilitates or stabilizes HOXA1 homodimerization, a conformational state often requisite for robust DNA binding.

Finally, to confirm the functional significance of this cooperation, we generated NONO-HOXA1 double-knockout (DKO) cells. Phenotypic analysis demonstrated that DKO cells exhibited significantly more severe defects in cardiac differentiation compared to either single KO line (Fig. 5M–P, Supplementary Fig. 1I, J). Together, these findings support a model where NONO promotes HOXA1 homodimerization to enable stable DNA binding and synergistic activation of the precardiac mesoderm program.

The NONO-HOXA1 interaction is critical for Wnt/β-catenin activation
As demonstrated above, HOXA1-regulated genes dependent on NONO are significantly enriched in the Wnt signaling pathway. Among these genes, three were upregulated, and six were downregulated in both HOXA1-KO and NONO-KO cells (Fig. 6A, B). The expression patterns of key canonical Wnt signaling pathway genes, such as WNT5A, WIF1, and GRB10, were validated in cells on Day 2 of differentiation via qRT‒PCR and ChIP‒qPCR (Supplementary Fig. 10A, B). The findings suggested that NONO is critical for the HOXA1-mediated activation of key Wnt signaling components.

The transcription factor β-catenin is an important downstream effector of the canonical Wnt signaling pathway. Upon Wnt receptor activation, β-catenin stabilizes, enters the nucleus, and activates Wnt target genes together with TCF439. To investigate the role of HOXA1 and NONO in regulating β-catenin, we assessed β-catenin expression in both the cytoplasmic and nuclear fractions of WT, HOXA1-KO, and NONO-KO cells on Day 2 of differentiation. Western blot analysis revealed that, compared with those in WT cells, cytoplasmic β-catenin levels in HOXA1-KO cells remained unchanged, but nuclear β-catenin levels were significantly lower, suggesting that HOXA1 plays a critical role in facilitating the nuclear accumulation of β-catenin (Fig. 6C and Supplementary Fig. 10C). Similarly, in NONO-KO cells, the cytoplasmic β-catenin level was unaffected, but the nuclear β-catenin level was significantly reduced. Partial recovery of nuclear β-catenin was observed in NONO-RE cells (Fig. 6D and Supplementary Fig. 10D). To determine whether this nuclear depletion stems from protein destabilization, we generated HOXA1-KO (Supplementary Fig. 1K, L) and NONO-KO HEK293T cell lines, and conducted CHX chase assays on nuclear fractions. The results demonstrated a marked acceleration in the turnover of nuclear β-catenin in both knockout lines compared to WT controls, despite stable cytoplasmic levels (Fig. 6E, F).

To evaluate the functional roles of HOXA1 and NONO in Wnt/β-catenin signaling in HEK293T cells, we used the TOP/FOP-Flash reporter assay stimulated with the Wnt activator CHIR99021 (CHIR). HOXA1 knockdown significantly reduced CHIR-induced TOP/FOP Flash reporter activity, particularly at higher siRNA concentrations (Fig. 6G and Supplementary Fig. 10E). Conversely, HOXA1 overexpression enhanced reporter activity in a dose-dependent manner, confirming that HOXA1 positively regulates Wnt/β-catenin signaling (Fig. 6H). Similarly, NONO knockdown via siRNA reduced CHIR-induced reporter activity, indicating that NONO is essential for optimal Wnt signaling activation (Fig. 6I and Supplementary Fig. 10F). NONO overexpression also increased TOP/FOP-Flash reporter activity in a dose-dependent manner, further validating its role as a positive regulator of Wnt signaling (Fig. 6J).

Given that NONO depletion impairs the regulation of genes involved in the Wnt signaling pathway, a TOP/FOP luciferase assay was used to investigate the role of NONO in HOXA1-mediated Wnt activation. NONO overexpression enhanced HOXA1-driven Wnt signaling, suggesting that NONO facilitates the HOXA1-mediated activation of the pathway (Supplementary Fig. 10G). Further analysis revealed that the HOXA1-mediated activation of Wnt signaling is largely dependent on its HD, as mutating this domain significantly reduced luciferase activity (Fig. 6K). Additionally, the disruption of the HOXA1-NONO interaction resulted in decreased Wnt activity (Fig. 6L), emphasizing the importance of this interaction for maintaining Wnt signaling. To evaluate whether stronger Wnt activation could rescue the differentiation defect in NONO-KO hiPSCs, we performed a CHIR99021 dosage–response assay. Increasing the Wnt stimulus to 10–12µM at Day 0–2 restored CM differentiation, as reflected by higher TNNT2+ cell proportions (Fig. 6M, N) and increased expression of TNNT2, MYH6, and MYH7 (Fig. 6O). In contrast, excessive CHIR exposure (14µM) diminished differentiation efficiency, consistent with previously reported cytotoxic or lineage-disruptive effects of supraphysiological Wnt activation. These findings indicate that NONO deficiency elevates the threshold for Wnt-mediated mesoderm induction and that this defect can be mitigated by strengthening early Wnt activation.

Collectively, these findings indicate that NONO is indispensable for the HOXA1-mediated activation of the Wnt/β-catenin signaling pathway, emphasizing its cooperative role in regulating the Wnt pathway during CM differentiation (Fig. 6P).

Methods
hiPSC culture and CM differentiation
Research involving human induced pluripotent stem cells (hiPSCs) was performed in strict accordance with the 2021 ISSCR Guidelines for Stem Cell Research and Clinical Translation. Human Induced pluripotent stem cells (WT-hiPSCs) derived from normal individuals were provided by Professor Ning Sun from the Department of Pathophysiology, Basic Medical College, Fudan University. This established cell line was generated from a skin punch biopsy of a healthy donor under ethical approval from the Human Research Ethical Review Board of Zhongshan Hospital of Fudan University, and informed consent was obtained from the donor as previously described63. hiPSCs were cultured on Matrigel-coated plates using mTeSRTM plus medium (Stemcell Technologies, 100-0276). Upon reaching ~80% confluence, the cells were split at a ratio of 1:5 using Accutase (Sigma, A6964). We utilized a previously established protocol involving WNT activation and inhibition to differentiate hiPSCs into CMs21. Briefly, hiPSCs were seeded at 80% confluence, treated with B27 minus insulin (Thermo, A1895601)/RPMI1640 (Thermo, 11875119) plus 7μM CHIR99021 (Selleck, S1263). Following two Days of CHIR99021 treatment, the medium was switched to B27 minus insulin/RPMI1640. After 24 hours, the medium was replaced with B27 minus insulin/RPMI1640 containing 5μM IWR1 (Selleck, S7086) for 2 Days. Subsequently, IWR1 was removed, and differentiation continued in CDM3 medium containing RPMI 640, 50 mg/ml BSA, and 213 mg/ml ascorbic acid. Beating CMs was observed by Day 8.

HEK293T cell culture and transient transfection
HEK293T cell line was obtained from ATCC (ATCC: ACS-4500) and cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, 11995-065) supplemented with 10% fetal bovine serum (FBS) (Gibco, 10099-141C) and 1% Penicillin-Streptomycin (Gibco, 15070-063). Transient transfection was performed using jetPRIME® (Polyplus, 10100046), following the manufacturer’s protocol.

siRNAs were obtained from QianMo Biotechnology Co., Ltd. (Shanghai, China): NONO siRNA target sequence, AGGTCATGCTAATGAGACA and HOXA1 siRNA target sequence, CTACGCGTTAAATCAGGAA. Transfection of these siRNAs was carried out using jetPRIME® (Polyplus, 10100046) according to the manufacturer’s protocol.

Generation of CRISPR-mediated gene knockout hiPSC and HEK293T cell lines
Single-guide RNAs (sgRNAs; sequences provided in Supplementary Data 1) were designed using DeepHF (https://www.deephf.com/64) and subsequently cloned into the EpiCRISPR vector, kindly provided by Dr. Yongming Wang. All sgRNA oligonucleotides were synthesized by Tsingke Biotechnology Co., Ltd. (Beijing, China). The EpiCRISPR-gRNA construct was transfected into hiPSCs or HEK293T cell lines using the Lonza 4D Nucleofector system (Lonza, AAF-1002X). Gene knockout (KO) clones were selected with puromycin (Thermo, A1113803) for one week, followed by screening through genomic sequencing, qRT-PCR and western blot analysis.

Generation of doxycycline-inducible NONO and HOXA1 expression in hiPSC lines
NONO-FLAG (pcDNA3.1) and HOXA1-HA (pcDNA3.1) plasmids were already developed in our laboratory. A PiggyBac transposon system kindly provided by Professor Ning Sun was used to establish doxycycline-inducible NONO expression in NONO-KO hiPSC lines and doxycycline-inducible HOXA1 overexpression hiPSC lines. The cDNA encoding the FLAG-NONO/HOXA1-HA tag fusion protein was subcloned downstream of the TRE3G promoter using the ClonExpress Ultra One Step Cloning Kit V2 (Vazyme, C116-01), resulting in the construction of the PB-TRE-NONO-FLAG, PB-TRE-NONO-HA, PB-TRE-HOXA1-FLAG, and PB-TRE-HOXA1-HA expression vectors. These PiggyBac expression plasmids were co-transfected together with the rtTA and PiggyBac transposase vectors (2:2:1) into WT and NONO-KO hiPSCs using the Lonza 4D Nucleofector system. The electroporated cells were subjected to 1 week of combined puromycin (Selleck, S7471) and G418 (Gibco, 10131) selection. Positive clones expressing NONO and HOXA1-HA in response to doxycycline induction were jointly screened using western blot analysis. This strategy enabled doxycycline-inducible reactivation of NONO in NONO-KO hiPSCs, overexpression of NONO-FLAG and NONO-HA in WT-hiPSCs, and inducible expression of HOXA1-FLAG and HOXA1-HA in both WT and NONO-KO backgrounds.

Immunofluorescence staining
Individual hiPSC clones and single beating CMs, dissociated using TrypLE Select (Thermo, 12563029), along with HEK293T cells dissociated with 0.25% Trypsin-EDTA (Thermo, 25200056), were plated on 35 mm imaging dishes (ibidi, 81156) pre-coated with Matrigel. The cells were fixed with 4% paraformaldehyde (Solaribio, P1110) for 15 minutes (min), permeabilized with 0.02% Triton-X-100 for another 15 min, and blocked with 5% BSA for 1 hour prior to staining. For pluripotency assessment, hiPSC colonies were stained with antibodies against OCT4 (Abcam, ab181557) at a 1:500 dilution and SSEA4 (Abcam, ab16287) at a 1:200 dilution. CM differentiation was assessed using antibodies against TNNT2 (Thermo, MA5-12960) at a 1:200 dilution and ACTN2 (Proteintech, 14221-1-AP) at a 1:100 dilution. For colocalization studies of NONO and HOXA1, staining involved antibodies against NONO (Santa Cruz, sc-376865) at a 1:200 dilution and HA tag (Abcam, ab9110) at a 1:500 dilution. Following primary antibody application, all cells were incubated with Alexa Fluor-conjugated secondary antibodies (Thermo, A32731 and A32723) at a 1:1000 dilution at room temperature for 1 hour. Cells were then counterstained with DAPI for 15 min at room temperature. The labeled cells were visualized using a Leica DMi8 fluorescence microscope.

Alkaline phosphatase staining (AP)
hiPSCs were seeded on a six-well plate coated with Matrigel and cultured for 2–3 Days. AP staining was performed using the Quantitative Alkaline Phosphatase ES Characterization Kit (Millipore, SCR004), following the manufacturer’s protocol.

Calcium transient imaging
Day 15 differentiated CMs were loaded with 5μM Cal-520® AM (AAT Bioquest, 21130) and 0.02% Pluronic F-127 (AAT Bioquest, 20053), and then incubated at 37°C for 15 min in Tyrode’s solution (Solaribio, T1420). After that, CMs were washed three times using Tyrode’s solution. The calcium transients of CMs were recorded using a Leica DMi8 fluorescence microscope (10 ms per scan, 30 s). The LAS X Science Microscope Software (Leica) and Excel were used for the transition of the calcium image into data. We developed an R script (https://doi.org/10.5281/zenodo.18059608) to detect inflection points for the data of calcium transition, followed by calculation and statistical analysis based on the calcium transient parameters schematic shown in Fig. 1H.

Flow cytometry assay
Single beating CMs, dissociated using TrypLE Select, Dissociated cells were washed with PBS and Perm/Wash Buffer (BD Biosciences, 554715), then fixed and permeabilized using Fixation/Permeabilization Solution for 60 min. Cells were incubated for 1 hour with either anti-cardiac troponin T (cTnT) primary antibodies or isotype-matched controls. For additional lineage analyses, dissociated cells were separately stained with antibodies against PDGFRA (R&D Systems, AF1062), NKX2-5 (CST, 8792), or GATA4 (Santa Cruz, sc-25310) using the same fixation and permeabilization conditions. After washing, cells were stained with secondary antibodies (Proteintech, SA00003-1, SA00008-9 and SA00008-2) for 30 min, then washed, resuspended, and analyzed using a FACSCalibur flow cytometer (BD Biosciences). Data were processed with FlowJo software v10. All data were analyzed using FlowJo software (BD). The gating strategies used for data analysis are shown in Supplementary Fig. 11.

Transmission electron microscopy
CMs were immediately fixed with 2.5% glutaraldehyde solution for 15 min at room temperature, CMs underwent dehydration in escalating ethanol concentrations: 60%, 70%, 80%, 90%, and 95%, each for 24 hours, followed by two treatments in 100% ethanol, each lasting 12–24 hours. The samples were then embedded in a mold with T7200 resin and cured under light for 12 hours to polymerize. Sectioning involved adhering the embedded blocks to slides using T4000 glue, smoothing with 1200-grit sandpaper, and affixing to a top slide with T7210 precision adhesive using a parallel sectioning device. This configuration was exposed to blue light for 10 min before disassembly and then vacuum-mounted on a hard tissue microtome to cut 200 μm thick sections. Finally, these sections were polished down to 30 μm thickness using a hard tissue grinder and analyzed using a TEM (HT7800; HITACHI).

Quantitative reverse transcription real-time polymerase chain reaction (qRT-PCR)
Total RNA was extracted using Trizol (Thermo, 15596018) according to the manufacturer’s protocol. Complementary DNA (cDNA) was prepared using the PrimeScript™ RT reagent Kit (Takara, RR036A) with 1 μg RNA. qRT-PCR was performed using the TB Green® Premix Ex Taq™ (Takara, RR420A) on a QuantStudio 3 system (Applied Biosystems). Relative mRNA expression was normalized to the GAPDH expression level. Primer sequences are provided in Supplementary Data 1, and all primers were synthesized by Tsingke Biotechnology Co., Ltd. (Beijing, China).

Bulk RNA-seq and alternative splicing analysis
RNA-seq library preparation was performed according to the manufacturer’s instructions (Vazyme, TR503-01). The libraries underwent high-throughput sequencing conducted by Haplox company (Jiangxi, China) using the Illumina NovaSeq 6000 platform, ensuring three biological replicates.

For RNA-seq data analysis, sequencing reads in FASTQ format were first processed using TrimGalore (v0.6.4) with the parameters ‘--paired --illumina’ to remove low-quality reads and adapter sequences. The trimmed reads were subsequently aligned to the human reference genome (UCSC hg38) using TopHat (v2.1.1) under default settings. Gene expression quantification was performed using Cufflinks (v2.2.1) with default parameters, which assigned the mapped reads to human gene annotations and generated expression abundances in FPKM (Fragments Per Kilobase of transcript per Million mapped reads). The FPKM values were then normalized to TPM (Transcripts Per Kilobase Million) to enable cross-sample comparison of gene expression levels. Differential expression analysis was carried out using Cuffdiff within the Cufflinks suite, which computed fold-changes and P values for each gene. The analysis included three biological replicates for Days 2, 5, 9, and 15, with differentially expressed genes defined as those exhibiting a fold-change greater than 1.5 and a P value less than 0.05. To identify alternative splicing events, aligned reads were analyzed using rMATS, with significant splicing changes defined by P value < 0.05 and junction read counts ≥5. Functional enrichment analysis, including GO term and KEGG pathway analysis, was conducted on the differentially expressed genes using the R package clusterProfiler (v3.14.0) from Bioconductor.

9-square plot analysis
To compare the changes in differentially expressed genes (DEGs) between KO vs WT and KO vs Rescue, we first identified DEGs in both comparisons with a cutoff of 1.5-fold change. To visually represent these DEG changes in a scatter plot, we categorized the genes into a 9-square matrix based on their regulation patterns. Each point in the matrix represents a gene, and genes were assigned to 9 groups (A–I) based on their up- or downregulation according to the 1.5-fold change threshold. Log2 (fold-change) values greater than 4 or less than −4 were scaled to 4 or −4, respectively. Group C contains genes that are upregulated in both KO vs WT and KO vs Rescue, while group G contains genes that are downregulated in both comparisons. This matrix allows for a direct comparison of DEG changes between KO vs WT and KO vs Rescue, as shown in the scatter plot, where the different groups are visually distinguished by their respective fold-change patterns.

Western blot (WB)
Whole cell lysates were prepared using RIPA buffer (Thermo, 89901) containing protease inhibitor cocktail. Lysates were boiled with 1× Sample buffer (GenScript, M01015) for 5 min and separated by 4–20% SurePAGE™ Bis-Tris Gels (GenScript, M00655), followed by transfer to PVDF membranes (Millipore, IPVH00010). The Membranes were blocked for 1 hour at room temperature in 1× Tris Buffered Saline containing Tween-20 (TBST) (Beyotime, ST673) with 5% non-fat dry milk (BD biosciences, 232100), then incubated overnight at 4°C with primary antibodies against NONO at a dilution 1:1000, HA at a dilution 1:5000, GAPDH (Proteintech, 10494-1-AP) at a dilution 1:10000, GFP (Abcam, ab290) at a dilution 1:2000, GST (Proteintech, 10000-0-AP), β-catenin (Abcam, ab32572) at a dilution 1:3000, and Lamin A/C (CST, 4777) at a dilution 1:2000. The next Day, membranes were washed three times of 10 min each with TBST, and incubated 2 hours at room temperature with HRP-conjugated secondary antibodies (CST, 7074/7076) at 1:3000 dilution. The bands were incubated with SuperSignal West Pico PLUS (Thermo, 34580) for 2 min and detected using ChemiDoc Imaging System (Biorad XRS+). The intensity of each band was quantified using Image Lab software.

CHX chase assays
HEK293T cells were seeded in six-well plates and transfected with 1 μg of plasmids using JetPrime reagent. To assess protein stability, cells were treated with 200 μg/mL CHX (Sigma-Aldrich, 01810) 24 hours post-transfection to inhibit protein synthesis. ImageJ software was used to estimate protein abundance.

Subcellular fractionation
Subcellular fractionation was prepared using NU-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo, 78835) according to the manufacturer’s protocol. The entire process was performed at 4°C. The cells were harvested with TryPLE Select and then centrifuged for 5 min. Cells were washed with PBS 2 times and incubated with CER I buffer containing protease inhibitor cocktail for 10 min, then incubated with CER II for 1 min. Centrifuge the tube to get the cytoplasmic extraction, then suspend the insoluble fraction with NER containing protease inhibitor cocktail. Vortex for 15 seconds every 10 min, for a total of 40 min, then sonicate in low mode (15 s on, 15 s off, 15 cycles) using Biorupter® Plus sonication device (Diagenode, B01020001). Centrifuge the tube to get the nuclear extraction. The Nuclear and Cytoplasmic proteins were then detected by WB.

Co-immunoprecipitation
For 293T cells, cells overexpressing HOXA1 and NONO were lysed in a 10 cm dish using 1 ml IP lysis buffer containing protease inhibitor cocktail and incubated on ice for 20 min. Similarly, Day 2 differentiated cells with DOX-induced HOXA1-HA overexpression (7–8×106 cells) were treated with the same lysis protocol. The lysates were then centrifuged at 12,000×g for 20 min. Supernatants were precleared with 15 μl each of Dynabeads Protein G (Thermofisher, 10004D) and Protein A (Thermofisher, 10002D), rotated at 4°C for 1 hour to reduce non-specific binding, and subsequently cleared beads were removed with a DynaMag Magnet. 50 μL of each supernatant was saved as input. The remaining supernatant was incubated either with NONO and rabbit control IgG or HA and rabbit control IgG (4 μg of each antibody), rotated at 4°C overnight (16 hours). Following antibody binding, 15 μl each of Dynabeads Protein G and Protein A were added and incubated for an additional 2 hours at 4°C. The beads were washed three times with IP lysis buffer and then boiled at 95°C for 10 min in 50 μl of 2× SDS sample buffer. The supernatant from this final boiling step was collected for further analysis.

LC–MS/MS analysis and NONO protein complex identification
Wild-type and NONO-KO hiPSCs were differentiated for two Days, followed by immunoprecipitation with a NONO antibody. Protein complexes were eluted and analyzed by mass spectrometry. NONO-KO Day 2 differentiated cells served as a negative control. (n=1). Proteins bound to the IP beads were eluted using SDT lysis buffer (4% SDS, 100 mM DTT, 100 mM Tris-HCl, pH 8.0). Samples were boiled for 3 min and further disrupted by ultrasonication. Insoluble material was removed by centrifugation at 16,000×g for 15 min, and the supernatant containing solubilized proteins was collected. Protein digestion was performed using the filter-aided sample preparation (FASP) method as previously described65. Briefly, samples were processed in urea buffer containing iodoacetamide to block reduced cysteines. Proteins were then digested with 2 μg trypsin (Promega) overnight at 37°C. Peptides were collected by centrifugation at 16,000×g for 15 min and desalted using C18 StageTips prior to LC–MS/MS analysis.

LC–MS experiments were performed on a Q Exactive HF-X mass spectrometer that was coupled to Easy nLC 1200 (Thermo Scientific). The peptide was first loaded onto a trap column (100 μm × 20 mm, 5 μm, C18, Dr. Maisch GmbH, Ammerbuch, Germany) in buffer A (0.1% Formic acid in water). Reverse-phase high-performance liquid chromatography (RP-HPLC) separation was performed using a self-packed column (75 μm × 150 mm; 3 μm ReproSil-Pur C18 beads, 120 Å, Dr. Maisch GmbH, Ammerbuch, Germany) at a flow rate of 300 nL/min. The RP-HPLC mobile phase A was 0.1% formic acid in water, and B was 0.1% formic acid in 95% acetonitrile. The gradient was set as follows: 2%–4% buffer B from 0 min to 2 min, 4% to 30% buffer B from 2 min to 47 min, 30% to 45% buffer B from 47 min to 52 min, 45% to 90% buffer B from 52 min to 54 min, 90% buffer B kept till to 60 min. MS data were acquired using a data-dependent top 20 method dynamically choosing the most abundant precursor ions from the survey scan (350–1800 m/z) for HCD fragmentation. A lock mass of 445.120025 Da was used as an internal standard for mass calibration. The full MS scans were acquired at a resolution of 60,000 at m/z 200, and 15,000 at m/z 200 for the MS/MS scan. The maximum injection time was set to 50 ms for MS and 25 ms for MS/MS. Normalized collision energy was 28, and the isolation window was set to 1.6 Th. Dynamic exclusion duration was 30 s.

The MS data were analyzed using MaxQuant software version 2.0.1.066. MS data were searched based on the UniProt-Reference proteome-Homo sapiens (Human) [96060-81791-20230317.fasta (https://www.uniprot.org/proteomes/UP000005640, 81791 total entries). Settings were set as below: Enzyme-trypsin; Max Missed cleavages-2; Precursor Tolerance (Main search)−4.5 ppm; Precursor Tolerance (First search)−20 ppm; Variable modifications-Oxidation (M), Acetyl (Protein-N-term); Database pattern-Target-Reverse; PSM FDR-0.01; Protein FDR-0.01; Site FDR-0.01.

GST Pulldown
NONO GST-fusion constructs were expressed in BL21 Escherichia coli, which were grown to an appropriate optical density and induced with 0.5 mM IPTG (Selleck, S6826) at 22°C for 4 hours. Cell pellets were resuspended in 10 mL of lysis buffer (10 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 0.5% NP-40, 0.09% sodium azide) and sonicated for 30 min (10 seconds on, 10 seconds off) at 4°C. The lysates were centrifuged at 12,000×g for 20 min. The resulting supernatants were mixed with supernatants from HOXA1-HA overexpressing HEK293T cell lysates and incubated with rotation at 4°C for 16 hours. To prepare for affinity purification, 25 μL of GST Trap Agarose (Proteintech, sta) was added to a microcentrifuge tube along with 500 μL of dilution buffer (10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 0.018% sodium azide), centrifuged at 2500×g for 5 min at 4°C, and the supernatant was discarded. The equilibrated agarose was then introduced to the mixed proteins and incubated with rotation at 4°C for 2 hours. The agarose was washed three times with 500 μL of wash buffer (10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5 mM EDTA, 0.05% NP-40, 0.018% sodium azide), each time centrifuging at 2500×g for 5 min at 4°C and discarding the supernatant. Finally, the agarose was resuspended in 50 μL of 2× SDS sample buffer and boiled at 95°C for 10 min. The proteins were then detected by WB.

Structure modeling with AlphaFold2
The interaction between NONO (Uniprot accession, Q15233) and HOXA1 homeodomain (HOXA1-HD, amino acid 229-288 of Uniprot accession P49639) was modeled utilizing AlphaFold-multimer (v2.3.2) with default parameters. The complex formed by NONO and HOXA1-HD was then visualized and analyzed using Chimera (v1.18).

Chromatin immunoprecipitation (ChIP) and ChIP-seq
7–8×106 Day 2 differentiated cells were crosslinked with 1% formaldehyde for 10 min at room temperature, then quenched with 0.125 M glycine for 5 min. Cells were washed three times with cold PBS and harvested in DNA LoBind Tubes (Eppendorf, 0030108035/0030108051). Sonication was performed using a Bioruptor (Diagenode, B01020001) on high setting (30 s on, 30 s off) for 30 min in high salt lysis buffer (20 mM Tris-HCl pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.05% SDS, 1% Triton-X-100) containing a protease inhibitor cocktail. Chromatin samples were precleared with 15 μL each of Dynabeads Protein G and Protein A, and rotated at 4°C for 1 hour. Precleared samples were then incubated with 4 μg of specific antibody at 4°C for 16 hours. After antibody binding, 15 μL each of Dynabeads Protein G and Protein A were added and incubated for an additional 2 hours at 4°C. The chromatin-antibody complexes underwent four washes with high salt buffer, two washes with low salt buffer (10 mM Tris-HCl pH 8.0, 250 mM LiCl, 1 mM EDTA, 0.5% NP-40, 0.5% sodium deoxycholate), and one wash with TE buffer (50 mM Tris-HCl pH 8.0, 10 mM EDTA, 50 mM NaCl). Complexes were eluted in elution buffer (50 mM Tris-HCl, pH 8.0, 10 mM EDTA, 1% SDS) on a Thermal Mixer (Thermo, 13687712) at 65°C for 4 hours, followed by treatment with 10 μg RNase and 10 μg Proteinase K at 55°C for 1 hour on the Thermal Mixer. DNA was purified using a QIAquick PCR Purification Kit (QIAGEN, 28006). For Re-ChIP assays, the starting material was scaled to 2×107 cells. Chromatin was first immunoprecipitated with 12 μg of anti-HA antibody and competitively eluted under native conditions using HA peptide (1 mg/mL) (MCE, HY-P0239). Subsequently, 30% of the eluate was retained as the primary ChIP fraction, while the remaining 70% was diluted and subjected to incubation with 12 μg of anti-NONO antibody overnight, and then captured with Dynabeads. The purified DNA library preparation was constructed using VAHTS Universal Plus DNA Library Prep Kit for Illumina (Vazyme, ND617) in accordance with the manufacturer’s instructions. Libraries were sequenced by the Haplox company (Jiangxi, China) using Illumina NovaSeq 6000.

ChIP-seq data analysis
Sequencing reads underwent trimming with Trim_galore (v0.6.4_dev) and were subsequently aligned to human genomes using Bowtie2 (v2.2.5)67. PCR duplicate reads were eliminated using the rmdup command of samtools (v1.7)68. To visualize ChIP signals in the UCSC Genome Browser, genomic coverage bedGraph files were generated using the bamCoverage in deeptools (v3.0.2)69. Peak calling was performed using MACS270 with the following parameters: -q 0.1 -f BAMPE --extsize=500 --bw 300 for NONO, and -f BAMPE --extsize=300 --bw 300 for HOXA1. Heatmaps and aggregate plots were generated using deeptools computeMatrix for data organization, followed by plotHeatmap and plotProfile for visualization, respectively. The genomic distribution of ChIP-seq peaks was assessed using the R package ChIPseeker (v3.10)71, with promoter regions defined from TSS-2 kb to TSS+2 kb.

Single-cell RNA-sequencing and preprocessing
iPSCs were differentiated into CMs for scRNA-seq. Differentiation efficiency was evaluated by flow cytometry. Differentiated cells collected on Days 2, 5, and 15 were dissociated using Accutase and neutralized with FBS in DMEM. For each time point, three biological replicates were obtained from both WT and KO groups. After centrifugation at 1000×g for 5 min, cell pellets were resuspended in DPBS containing 0.04% bovine serum albumin and immediately submitted for scRNA-seq. Single-cell libraries were generated using the Chromium GEM-X Single-Cell 3′ Kit v4 (10x Genomics) according to the manufacturer’s instructions, and sequenced on an Illumina Novaseq X to a mean depth of at least 20 K reads per cell.

Sequencing reads were aligned to the human reference genome (GRCh38), and gene-by-cell count matrices were generated. Downstream analysis was performed using Seurat (v5.3.0)72 and the Pegasus pipeline (v1.9.0)73. Quality control included removing cells with low gene counts (<500) or elevated mitochondrial transcript proportions (>10%) to exclude low-quality or dying cells. Gene expression matrices were then normalized using SCTransform, and the top 3000 highly variable genes were selected. The Seurat object was subsequently converted to an AnnData object for further processing with Pegasus. Principal component analysis was performed, and the top 50 principal components were used to construct a shared nearest-neighbor graph. Batch effects across samples were mitigated using scVI74 implemented in the Pegasus framework, followed by dimensionality reduction with UMAP. Cell type identities were assigned based on cluster-specific marker genes, supported by previously published annotations.

Cell differentiation potential was estimated using the CytoTRACE kernel implemented in the CellRank package, applying the CytoTRACEKernel().compute_cytotrace() function with default parameters75. Pseudotime trajectories for mesendoderm, cardiac progenitor, and CM populations were reconstructed using Monocle 224. Highly variable genes were used for dimensionality reduction with DDRTree, after which cells were ordered along the inferred trajectories to assign pseudotime values.

TOP/FOP reporter assay
HEK293T cells were plated in 12-well plates and transfected with the indicated plasmids or siRNAs for 24 hours, followed by a second transfection with either TOP-FLASH or FOP-FLASH reporter plasmids together with pRL-TK. After another 24 hours, cells were dissociated and re-plated into 96-well plates, distributing the contents of each well from the 12-well plate into six wells of the 96-well plate. The cells were then treated with either DMSO or 7 μM CHIR99021, a WNT/β-catenin activator, for 24 hours. Luciferase activity was measured using the Dual-Luciferase Reporter Assay System (Promega, E1960) according to the manufacturer’s protocol, and the results were detected using a Varioskan LUX Multimode Microplate Reader (Thermo, VL0000D0). The average luciferase activity from the six wells was calculated, normalized to Renilla luciferase activity, and expressed as the TOP/FOP ratio to determine relative Wnt activity. All transfection experiments were conducted with at least three biological replicates.

Statistical analysis
Data are presented as mean ± standard deviation (SD) or mean ± standard error of the mean (SEM), as indicated in the figure legends. All quantitative experiments were performed with at least three independent biological replicates. An unpaired two-tailed Student’s t test and Fisher’s exact test were used to calculate significant differences between the two groups. P values < 0.05 were considered to indicate a statistically significant result. All analyses were performed using GraphPad Prism version 9.0.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript by Feng et al. presents a comprehensive investigation into the role of the non-POU domain-containing octamer-binding protein (NONO) in heart development, using human iPSC-directed cardiomyocyte (CM) differentiation as a model system. The authors generated a series of hiPSC lines to address this question. Specifically, they employed CRSIPR/Cas9 to compare the CM differentiation in NONO-KO hiPSC to that ub WT hiPSC. RNAseq analysis revealed that genes involved in early heart specification were downregulated in NONO-KO hiPSC, resulting in impaired CM maturation in NONO-KO cells, accompanied by structural and functional abnormalities. These defects were rescued upon reactivation of NONO (NONO-RE) in NONO-KO hiPSC-derived CMs. Transcriptomic analysis of WT, NONO-KO and NONO-RE hiPSCs at differentiation days 2,5,9 and 15 showed that NONO modulates the expression of genes critical for heart development. Given that NONO regulates both transcription and alternative splicing, the authors further analyzed transcriptomic data at day 2 and compared these findings with NONO-ChIP-seq data. Based on this comparison, they suggest that NONO regulates gene expression through transcriptional activity, independently of its splicing function. To explore how NONO influences gene expression in CMs, the authors performed immunoprecipitation-mass spectrometry and identified HOXA1 as a NONO interactor, biding through its homeodomain. ChIP-seq analysis at day 2 revealed 1,169 overlapping peaks between NONO and HOXA1, with an enrichment in genes involved in process such as Wnt signaling pathway, gastrulation, and heart morphogenesis. These data led the authors to propose that NONO is indispensable for HOXA1-mediated regulation of precardiac mesoderm genes. Finally, focusing on canonical Wnt signaling, they show that the NONO-HOXA1 interaction is critical for Wnt/beta-Catenin activation.\n\nThis study provides a rich dataset combining mRNA-seq and ChIP-seq in hiPSC-derived CMs, offering mechanistic insights into the role of NONO in heart development. The use of bioinformatic analyses enhances our understanding of how NONO regulates CM specification/differentiation. The study identifies NONO-HOXA1 interaction as a key regulator in cardiac development. However, a major limitation of the study is the lack of experimental validation for key conclusions, particularly the proposed role of the NONO-HOXA1 interaction in regulating early cardiac genes. The manuscript would also benefit from a clearer discussion of how these results advance beyond previous works on the role of NONO during early development.\n\nOverall, the study is thoughtfully conducted and offers valuable observations that contribute to our understanding of early heart development. Nonetheless, the central conclusion regarding the NONO-HOAX1-Wnt signaling axis in cardiac specification is not sufficiently substantiated and would be strengthened by additional experimental validation.\n\nMajor points:\n- The author state that NONO plays a critical role in CM differentiation. However, based on the presented data, it appears more accurate to conclude that NONO is required for early specification, and that disruption at this stage subsequently impairs CM differentiation. To clarify this point, it is important to quantify the proportion of progenitor cells expressing MESP1, GATA4, and NKX2-5. It remains unclear whether the NONO-KO cells fail to specify into cardiac progenitors or adopt non-cardiac lineages instead.\n- The authors should specify which subtype of CMs are generated using their differentiation protocol. It is well established that in vitro differentiation of hiPSC can give rise to heterogeneous CM populations. Clarifying the major type of CM subtype would help also to understand the phenotype during CM maturation.\n- The authors demonstrate that reactivation of NONO expression in NONO-KO hiPSCs rescues CM differentiation. However, to fully support their model, it is essential to also show that early cardiac specification is restored upon this reactivation.\n- NONO expression peaks at day 2, which is consistent with a potential role in early cardiac specification. To support this, the authors should indicate the timing if CM differentiation by presenting the temporal expression profile of key differentiation markers.\n- The comparison of gene expression levels between NONO-RE and NONO-KO cells from day 2 to day 15 identified 332 genes in Group I and 60 genes in Group I’, which are potentially regulated by NONO. However, it remains unclear whether these genes are direct or indirect targets of NONO. To address this point, the authors should integrate and compare the NONO ChIP-seq data with these gene sets to determine which are directly bound by NONO. This analysis would help distinguish primary regulator targets from downstream effects.\n- The authors should address the temporal differences between their in vitro differentiation model and in vivo cardiac development during embryogenesis. Notably, several studies have established that MESP1 as a critical factor for early cardiac specification, acting upstream of key transcription factors in both the FHF and SHF (e.g., Lescroart et al. 2014, 2018). In this context, it is unexpected that MESP1 expression peaks at day 4 in the present model, as MESP1 expression in vivo occurs much earlier during mesodermal specification. This point should be more detailed and included in the discussion.\n- The analysis of DEG and ASE between WT and NONO-KO cells, in conjunction of NONO-targeted genes, identified 542 transcriptionally regulated genes enriched in heart development pathways. However, as shown in Supplementary Fig 4C, there is an additional category of genes involved in the negative regulation of stem cell differentiation, which is also highly relevant and should discussed. Pointed to this category would provide a more comprehensive view of how NONO may influence both lineage commitment and the suppression of alternative fates during early cardiac differentiation.\n- The immunoprecipitation-mass spectrometry analysis performed at day 2 identified NONO and HOXA1 as potential interacting partners. The authors should clarify whether NONO forms homodimers, as this may influence its mode of interaction with other proteins. Additionaly, since HOXA1 is known to form homodimers, it is important to address the possibility of protein competition or cooperation between HOXA1 homodimers and HOXA1-NONO complexes in regulating target gene expression.\n- To strengthen the analysis of overlapping HOXA1 and NONO binging peaks, it would be essential to include additional epigenetic data, such as histone modification marks or ATACseq profiles. These data would help to identify active enhancer regions.\n- Figure 5F: The peak enrichment at the PDGFRA and MESP1 loci shown in Fig. 5F is not convincing. The authors should either provide higher-resolution tracks, quantitative peak calling data or additional replicates to support their claims.\n\nMinor Points:\n-Figure 6F: SiCon must be changed to NC.", "reviewer_2": "In this manuscript, the authors showed interesting findings about molecular mechanism of NONO in cardiomyocyte differentiation. Key contribution of NONO in cardiomyocyte differentiation is confirmed, and it is also demonstrated to be independent of NONO’s known splicing function. Furthermore, the authors identify HOXA1 as a key effector of NONO, and establish that the domain HD and residue K286 of HOXA1 play essential role in the direct interaction. The HOXA1-NONO complex regulates precardiac mesoderm gene expression by active downstream Wnt signaling. This manuscript is written fluently and is convinced by detailed data. However, there are still some major problems to be solved.\n\nMajor Concerns:\n1. The authors did not properly distinguish the concepts between differentiation and maturation. Maturation means the developmental process from a naïve cardiomyocyte to a better functional cardiomyocyte. The authors differentiated the cells for only 15 days and did not use methods to boost hiPSC-CM maturation (T3, fatty acid, electric stimulation or EHT formation). Therefore, all the conclusions about cardiomyocyte maturation are inappropriate and are indeed about differentiation. Here I recommend the authors to replace all maturation by differentiation and to avoid mentioning maturation, which the authors did not really examined.\n2. The cell composition of the NONO-KO versus control hiPSC-CM populations are apparently different due to the impaired differentiation efficiency in the mutant group. Consequently, bulk RNA-seq would largely reflect cell composition changes rather than transcription changes in CMs or CPCs. To address this concern, the authors should add scRNA-seq validation and look at the cardiac progenitor cell population to see if the observed gene expression changes are still real.\n3. The authors mentioned that NONO is likely a chaperone, thus the authors are recommended to look at HOXA1 stability to check this point. In addition, although the authors mapped the binding sites in HOXA1, they did not map the protein interaction sites in NONO and determine if this binding is necessary for HOXA1 stability and function.\n4. HD domain is responsible for HOXA1-DNA binding. Thus it is critical to test how NONO affects HOXA1 affinity to DNA. In addition, the authors emphasized that HOXA1 ChIP seq peak decreased in the NONO-KO cells. This could still be due to changes in cell composition rather than NONO-KO.\n5. The authors are recommended to validate key findings in an animal model, since the hiPSC-CM differentiation process is still quite artificial and may not truly reflect in vivo development.\n6. The mechanism how HOXA1 and NONO are required for nuclear localization of β-catenin lacks further explanation. If the cytoplasmic b-catenin level does not change, how could it be a nuclear translocation phenotype? Alternatively, could these factors affect nuclear b-actinin stability?\n7. The authors used a classic two-small molecule differentiation methods that involve WNT activation. If NONO-KO truly impair WNT activation and thereby CM differentiation, then one would expect rescued cardiomyocyte differentiation simply by increasing the concentration of CHIR activating Wnt signaling at the first step of differentiation? This might be a stronger support for Wnt signaling mechanism.\n8. NONO genetic variants are found in CHD patients. The results in this study should be validated in a hiPSC line with patient-related genetic variants to make sure the results in this study is disease related.\n9. The conclusion that NONO cooperates with HOXA1 is weak. The authors could do a reChIP experiment to see if the two molecules indeed act on the same loci. In addition, NONO-HOXA1 double silencing cells should be compared to each single KO lines to show cooperativity, rather than a upstream-downstream relationship in the same pathway.\n\nMinor Comments:\n1. A karyotype detection of the new hiPSC lines should be supplied. It is ideal to test the results in multiple hiPSC clones, even though the authors claimed that they are similar.\n2. Fig 1F: the images do not show reduced peak intensity, but rather increased.\n3. Fig2A: in the rescue experiment, the restored NONO protein level should be compared to endogenous level to determine why there is only partial rescue.\n4. Fig3B should label clearly that the genes were all from group I. In addition, the authors should display how the rescue changes genes in each group, not just group I.\n5. In line 569 and 618, the word “hESC” is misused. Human embryonic stem cell (hESC) is different from human induced pluripotent stem cell (hiPSC). The authors should be clear what cells are used for each cell line."}, "author_response": "We sincerely thank both Reviewer #1 and Reviewer #2 for their thoughtful and constructive comments. Their insightful suggestions have significantly improved the clarity, rigor, and overall quality of our manuscript. Below, we provide a point-by-point response to each comment. Reviewer comments are shown in italic. All corresponding revisions have been incorporated into the main text, Materials and Methods, Supplementary Information, and figures.\n\nReviewer #1 (Remarks to the Author):\n[...]\nWe thank the reviewer for the constructive assessment and for recognizing the value of our genetic models and the proposed NONO–HOXA1–Wnt mechanistic framework. This feedback has significantly improved the clarity and rigor of our manuscript.\nWe have carefully addressed the reviewer’s suggestions regarding experimental validation and the differentiation of our work from prior studies. Accordingly, we have incorporated new data to bolster our conclusions and expanded the Discussion to better define the novelty of our findings in the context of early cardiac development. A point-by-point response detailing these revisions follows below.\n\nReviewer #1 – Major points 1:\n[...]\nResponse:\nWe thank the reviewer for this important clarification. We agree that our data more accurately support the conclusion that NONO is required for early cardiac specification, and that impaired cardiomyocyte differentiation in NONO-KO cells arises as a downstream consequence of defective lineage induction.\nTo directly address the reviewer’s concern, we quantified the proportions of early cardiac progenitor populations at key differentiation stages. Because MESP1 is not reliably detected by flow cytometry, we used PDGFRA surface expression at day 3 as an established proxy for MESP1⁺ mesodermal progenitors1. Consistent with defective specification, NONO-KO cells showed a marked reduction in PDGFRA⁺ cells compared with WT (67.0% vs. 83.5%). This reduction persisted at subsequent specification checkpoints, with fewer GATA4⁺ progenitors at day 5 (18.3% vs. 35.0%) and fewer NKX2-5⁺ cardiac progenitors at day 6 (41.2% vs. 77.2%) (Panels A–C).\nOur newly added single-cell RNA-seq analysis further supports this interpretation. Although the CPC-like cluster is present in both genotypes, NONO-KO progenitors exhibit significantly reduced expression of core cardiac transcription factors such as GATA4 and ISL1, and an increased bias toward neural-associated lineages at day 5 (Panels D–E). These data demonstrate that NONO-KO cells do not undergo a complete loss of mesoderm formation but instead fail to properly activate the cardiac specification program, leading to diminished downstream differentiation potential.\nTogether, these results clarify that NONO functions at the specification stage and that mis-specification, rather than an intrinsic block in cardiomyocyte differentiation, underlies the observed differentiation defects.\nThe results shown here in Panels A–C have been integrated into the revised manuscript as Supplementary Fig. 2E–J. Similarly, the data in Panel D and Panel E are now presented as Supplementary Fig. 3D and Fig. 1L, respectively. Corresponding descriptions have been updated in the revised text (Page 4, lines 133–136; Page 6, lines 197–200).\n\nReviewer #1 - Major points 2:\n[...]\nResponse:\nWe thank the reviewer for this insightful comment. We agree that defining the cardiomyocyte (CM) subtype composition is important for interpreting the differentiation phenotype observed in our system. To address this point, we analyzed our day 15 single-cell RNA-seq dataset and examined the expression of canonical atrial and ventricular markers within TNNT2⁺ cardiomyocytes.\nThis analysis showed that TNNT2⁺ cells exhibit predominant MYL7 expression (73.9%) and minimal MYL2 expression (0.4%) (Panel A), indicating that the majority of CMs generated at this stage resemble atrial-like or immature ventricular-like cardiomyocytes. This is consistent with the known developmental timeline in hiPSC-derived cardiomyocytes, where MYL7 (MLC2a) is expressed early, whereas MYL2 (MLC2v)—a hallmark of ventricular maturation—typically appears only at substantially later stages of in vitro differentiation (often > day 40). 2,3\nTogether, these data clarify that our differentiation protocol predominantly produces immature, MYL7-enriched cardiomyocytes at day 15, which is consistent with the early-stage differentiation phenotypes characterized in our study.\nThe results shown here in Panels A have been integrated into the revised manuscript as Supplementary Fig. 2C. Corresponding descriptions have been updated in the revised text (Page 5, lines 126–129).\n\nReviewer #1 - Major points 3:\n[...]\nResponse:\nWe thank the reviewer for this important comment. We fully agree that demonstrating the rescue of early cardiac specification is essential to support our model that NONO acts upstream of lineage commitment and that its reactivation functionally restores the differentiation trajectory.\nTo directly address this point, we examined the temporal expression of key mesodermal and early cardiac transcription factors from day 2 to day 7 in NONO-RE cells. Quantitative RT–PCR analysis showed that NONO reactivation robustly restored the induction of T, EOMES, MIXL1, MESP1 and PDGFRA at early mesoderm stages, and subsequently rescued the activation of GATA4, NKX2-5 and MYOCD during cardiac progenitor formation (Panels A–B). The restored expression levels closely mirrored those observed in WT cells.\nThese results demonstrate that NONO reactivation effectively reinstates the early cardiac specification program, thereby providing functional evidence that the impaired differentiation phenotype in NONO-KO cells arises from defective lineage induction rather than an inability to undergo later maturation. This rescue further strengthens our conclusion that NONO is required at the specification stage to initiate the transcriptional cascade necessary for proper cardiomyocyte development.\nThe results shown here in Panels A, B have been integrated into the revised manuscript as Fig. 2B, C. Corresponding descriptions have been updated in the revised text (Page 9, lines 249–253).\n\nReviewer #1 - Major points 4:\n[...]\nResponse:\nWe thank the reviewer for this insightful suggestion. We agree that defining the temporal dynamics of NONO relative to established lineage markers is essential for clarifying its developmental role.\nTo address this point, we performed a detailed RT–qPCR time-course analysis across cardiac differentiation (days 0, 1, 2, 3, 4, 5, 6, 7, 9, and 15), examining the expression trajectories of pluripotency, mesoderm, cardiac progenitor, and cardiomyocyte markers, including OCT4, T, EOMES, MESP1, ISL1, GATA4, PDGFRA, NKX2-5, MYH6, MYH7, and TNNT2 (Panel A).\nThis analysis revealed that NONO expression is transiently upregulated at day 2, coinciding with the induction of early mesodermal regulators T and EOMES, and immediately preceding the peak of the cardiac mesoderm marker MESP1 at day 3. The subsequent activation of ISL1, GATA4, and NKX2-5 further delineates the normal progression toward cardiac progenitor formation.\nThe alignment of the NONO peak with these early lineage transitions strongly supports the conclusion that NONO acts at a pre-specification to early-specification window, functioning upstream of the transcriptional network that initiates cardiac lineage commitment.\nThe results shown here in Panels A have been integrated into the revised manuscript as Supplementary Fig. 2B. Corresponding descriptions have been updated in the revised text (Page 4 and 5, lines 114–126).\n\nReviewer #1 - Major points 5:\n[...]\nResponse:\nThank you for this important suggestion. In response, we performed an integrative analysis combining our NONO ChIP-seq dataset with the 332 Group I genes and 60 Group I′ genes identified from the NONO-RE versus NONO-KO comparisons. Using the day 2 NONO ChIP-seq dataset, we identified 4,350 NONO peaks associated with 2,212 gene promoters. Intersecting these promoter-associated peaks with our DEG groups revealed that 23 of the 332 Group I genes and 3 of the 60 Group I′ genes exhibit direct NONO promoter binding at day 2. These genes therefore represent the most likely primary transcriptional targets of NONO.\nWe note, however, that the expression differences defining Group I and I′ genes were derived from comparisons spanning day 2 to day 15, a developmental interval during which transcriptional regulatory networks undergo substantial and stage-specific rewiring. Because the available NONO ChIP-seq data capture binding only at day 2, many DEGs emerging at later stages are expected to be regulated indirectly, through downstream transcription factors, signaling changes, or lineage-specific cascades initiated earlier in differentiation.\nTogether, these integrative analyses distinguish a small subset of genes with direct NONO promoter occupancy from a broader set of indirectly regulated genes, thereby providing a clearer interpretation of NONO’s primary versus downstream regulatory effects.\n\nReviewer #1 - Major points 6:\n[...]\nResponse:\nWe thank the reviewer for this insightful comment. We apologize for the confusion caused by the missing day-3 label in the initial figure, which may have led to the impression that MESP1 peaks at day 4. After correcting the X-axis and presenting the full time course, our data clearly show that MESP1 is induced at day 2 and peaks at day 3, which is consistent with previously reported PSC-based cardiac differentiation systems³ (as also addressed in our response to Major points 4).\nWe agree with the reviewer that timing differences between in vitro hiPSC differentiation and in vivo embryogenesis are well recognized. We have now expanded the Discussion to acknowledge these temporal differences and to clarify how our results align with established in vivo findings. We also note that future studies using mouse embryonic models will be necessary to examine the precise in vivo temporal relationship between NONO, MESP1, and early mesodermal specification.\nDiscussion part:\nRegarding the temporal dynamics of this regulation, we noted that MESP1 induction peaks at day 3 in our hiPSC differentiation model (Supplementary Fig. 2B). While this timing is slightly delayed compared to mesodermal specification in vivo, it is consistent with established kinetics in pluripotent stem cell-based differentiation systems4-6. We acknowledge that in vitro models cannot fully recapitulate the precise chronometry of embryonic development, nor do they possess the complex spatial architecture of native organogenesis. However, the hiPSC system offers an ideal, controlled platform for dissecting molecular mechanisms. Therefore, validating our findings in vivo is a critical next step. Future studies using Nono-deficient mice will determine whether the molecular defects observed here translate into anatomical malformations during heart development.\nCorresponding descriptions have been updated in the revised text (Page 28, lines 1084–1094).\n\nReviewer #1 - Major points 7:\n[...]\nResponse:\nWe appreciate the reviewer for highlighting the biological significance of the \"negative regulation of stem cell differentiation\" category. We agree that discussing this category provides a more comprehensive view of NONO’s role in balancing lineage commitment and the suppression of alternative fates.\nIn the revised Discussion, we have added a dedicated section to address this. We now explain that NONO likely functions to dampen the expression of these \"gatekeeper\" genes (such as EZH2, REST, and HES1). This discussion clarifies that NONO is required not only to activate cardiac pathways but also to overcome the molecular barriers that restrict differentiation, thereby ensuring a robust exit from the progenitor state.\nDiscussion part:\nBeyond activating cardiac developmental pathways, our analysis revealed that NONO actively suppresses lineage-restricting factors. Established repressors such as EZH27, REST8, and HES19 were aberrantly upregulated in NONO-KO cells (Supplementary Fig 7D). This suggests a dual mechanism wherein NONO facilitates lineage commitment by synergistically activating cardiac identity genes while simultaneously dismantling transcriptional barriers imposed by progenitor-maintenance programs.\nThe results shown here in Panel have been integrated into the revised manuscript as Supplementary Fig. 7D. Corresponding descriptions have been updated in the revised text (Page 28, lines 1079–1084).\n\nReviewer #1 - Major points 8:\n[...]\nResponse:\nWe thank the reviewer for these insightful comments, which have significantly strengthened our mechanistic understanding of the HOXA1–NONO interaction. In particular, the questions regarding the interplay between HOXA1 homodimers and HOXA1–NONO complexes prompted us to further examine how NONO affects HOXA1 dimerization and DNA binding.\nIn response, we performed additional experiments to directly address the dimerization properties of NONO and HOXA1 and the potential regulatory interplay between HOXA1 homodimers and HOXA1–NONO complexes.\n1. NONO homodimerization\nTo experimentally validate whether NONO forms homodimers in our differentiation system, we generated doxycycline-inducible NONO-FLAG and NONO-HA hiPSC lines. Doxycycline was added at day 1 of differentiation, and co-expression of NONO-FLAG and NONO-HA was induced.\nConsistent with the reviewer’s comment, co-immunoprecipitation (co-IP) experiments demonstrated that NONO forms homodimers in day-2 differentiated cells (Panel A). This property is likely to influence its mode of interaction with other transcriptional regulators, including HOXA1.\n2. Competition or cooperation between HOXA1 homodimers and HOXA1-NONO complexes\nTo further investigate whether NONO influences HOXA1 dimerization, we established doxycycline-inducible HOXA1-FLAG and HOXA1-HA hiPSC lines in both WT and NONO-KO backgrounds. Doxycycline was added at day 1 of differentiation, and HOXA1 overexpression was induced.\nThe co-IP analyses revealed that HOXA1 homodimer formation was markedly reduced in NONO-KO cells compared with WT controls (Panel B and C). Based on our current data, we favor a cooperative rather than competitive relationship between the NONO–HOXA1 complex and HOXA1 homodimers.\nThe results shown here in Panel A-C have been integrated into the revised manuscript as Fig. 5J-L. Corresponding descriptions have been updated in the revised text (Page 20, lines 776–783).\n\nReviewer #1 - Major points 9:\n[...]\nResponse:\nWe thank the reviewer for this constructive suggestion. To characterize the epigenetic landscape of the NONO-HOXA1 co-occupied regions, we performed ATAC-seq and H3K27ac ChIP-seq analyses in Day-2 differentiated cells. By intersecting these profiles with our NONO and HOXA1 ChIP-seq data, we found that the vast majority of NONO-HOXA1 co-occupied peaks reside within accessible chromatin regions (~91% overlap with ATAC-seq peaks, 1110 / (1110+104) x 100 = 91%) and are marked by active histone modifications (~72% overlap with H3K27ac peaks, 901 / (901+349) x 100 = 72%) (Panel A). these data support a model where NONO and HOXA1 cooperate at open, active chromatin regions to regulate target gene expression.\nThe processed peak calling data are available at: https://drive.google.com/drive/folders/1wzqwnzLNZB0LX3Ik0ojFa1qiOeZmrRnf?usp=sharing\n\nReviewer #1 - Major points 10:\n[...]\nResponse:\nWe appreciate the reviewer’s rigorous scrutiny regarding the ChIP-seq enrichment. To provide more quantitative support beyond the browser tracks, we performed sequential ChIP (Re-ChIP) assays using a doxycycline-inducible HOXA1-HA hiPSC line differentiated to day 2. Chromatin was first immunoprecipitated with an anti-HA antibody (targeting HOXA1), eluted under native conditions, and subsequently re-immunoprecipitated with an anti-NONO antibody. As shown in Panel A, we observed robust enrichment at the MESP1 locus in the primary HA-ChIP, and PDGFRA demonstrated significant enrichment in the NONO Re-ChIP fraction, indicating that both loci are co-occupied by HOXA1 and NONO.\nTogether with the updated higher-resolution browser tracks, these Re-ChIP results provide strong and direct evidence supporting NONO–HOXA1 co-occupancy at key precardiac mesoderm regulatory loci, including MESP1 and PDGFRA.\nThe results shown here in Panel A have been integrated into the revised manuscript as Fig. 5F. Corresponding descriptions have been updated in the revised text (Page 19 and 20, lines 730–761).\n\nMinor Points:\n-Figure 6F: SiCon must be changed to NC.\nResponse:\nWe thank the reviewer for noting this oversight. Fig.6 “SiCon” has been corrected to “NC” as suggested.\n\nSummary and final appreciation for Reviewer #1\nWe are deeply grateful to Reviewer #1 for the thorough and constructive evaluation of our work. In response to these comments, we have (i) refined our conceptual framework to emphasize NONO’s role in early cardiac specification rather than late-stage maturation, (ii) added new flow cytometry and single-cell RNA-seq analyses to quantify cardiac progenitor populations and cell-state transitions, (iii) integrated NONO ChIP-seq data with time-resolved transcriptomic profiles to distinguish direct from indirect targets, (iv) clarified the temporal relationship between NONO, MESP1, and key developmental markers, and (v) expanded the Discussion to incorporate the “negative regulation of stem cell differentiation” gene set and to more carefully compare our findings with in vivo and previous studies.\nIn addition, we have strengthened the mechanistic model of NONO–HOXA1 cooperation by characterizing their dimerization properties, mapping interaction domains, performing Re-ChIP and epigenomic profiling (ATAC-seq and H3K27ac ChIP-seq), and examining the impact of NONO loss on HOXA1 chromatin binding and downstream Wnt signaling. Together, these revisions have, in our view, substantially improved the clarity, depth, and robustness of the manuscript. We sincerely thank Reviewer #1 again for the insightful guidance that has been instrumental in shaping the revised version of this study.\n\nReviewer #2 (Remarks to the Author):\n[...]\nWe thank Reviewer #2 for the positive overall evaluation of our study and for recognizing the mechanistic insights we provide into NONO function during cardiomyocyte differentiation, including its interaction with HOXA1 and downstream effects on Wnt signaling. We also appreciate the reviewer’s careful reading of the manuscript and the constructive identification of several important issues that require further clarification or experimental support. In the responses that follow, we address each of these points in detail and describe the revisions, additional analyses, and new experiments incorporated into the revised manuscript.\n\nReviewer #2 – Major concerns 1:\n[...]\nResponse:\nWe thank the reviewer for this critical distinction. We agree that given our experimental endpoint of 15 days and the absence of specific maturation stimuli (such as T3, fatty acid supplementation, or electromechanical stimulation), our study reflects the process of cardiac differentiation and the generation of naïve cardiomyocytes, rather than functional maturation.\nFollowing the reviewer’s recommendation, we have revised the manuscript to replace the term \"maturation\" with \"differentiation\" throughout the text, figure legends, and title.\n\nReviewer #2 - Major concerns 2:\n[...]\nResponse:\nWe thank the reviewer for raising this important concern. To determine whether the transcriptional differences observed in bulk RNA-seq reflect true regulatory changes within cardiac progenitors or cardiomyocytes rather than altered cell-type composition, we performed time-course single-cell RNA-seq on WT and NONO-KO cells collected at days 2, 5, and 15 of differentiation.\nUMAP analysis resolved major developmental populations—including mesendoderm, cardiac progenitors (CPCs), cardiomyocytes (CMs), endoderm, epithelial, endothelial, and neural progenitor cells—consistently across samples and defined by canonical lineage markers (Panels A–B). Cells from the three biological replicates were well integrated, indicating minimal batch effects (Panel C).\nMonocle210 trajectory analysis further demonstrated a continuous progression from mesendoderm to cardiac progenitors and ultimately cardiomyocytes in WT cells. However, NONO-KO cells showed reduced progression along the cardiac trajectory, with a subset retaining mesendoderm-like features at Day 5 and cardiac progenitor markers such as ISL1, GATA4, HAND1, and TBX20 exhibiting attenuated induction, accompanied by inappropriate persistence of POU5F1 (Panel D and E).\nCytoTRACE11 analysis further confirmed that NONO-KO cells remained transcriptionally less differentiated than WT counterparts within both CPC and CM clusters (Panel F).\nTogether, these single-cell datasets demonstrate that the transcriptional defects in NONO-KO cells are intrinsic to cardiac progenitor and cardiomyocyte populations, and are not merely a consequence of altered cell composition in bulk RNA-seq.\nThe results shown here in Panels A–C have been integrated into the revised manuscript as Supplementary Fig. 3A–C. Similarly, the data in Panel D-F are now presented as Fig. 1K-L, respectively. Corresponding descriptions have been updated in the revised text (Page 6, lines 185–193; Page 6, lines 197–200).\n\nReviewer #2 - Major concerns 3:\n[...]\nResponse:\nWe thank the reviewer for this insightful comment. To determine whether NONO functions as a molecular chaperone for HOXA1 and to define the interaction interface between the two proteins, we performed a combination of protein stability assays and domain-mapping experiments.\nFirst, we generated NONO-KO HEK293T cells and conducted cycloheximide (CHX) chase assays using nuclear lysates. HOXA1-HA protein exhibited markedly accelerated decay in NONO-KO cells compared with WT controls (Panels A–B), indicating that NONO is required to maintain HOXA1 protein stability.\nTo map the NONO regions responsible for binding HOXA1, we constructed deletion mutants lacking RRM1, RRM2, or the coiled-coil domain (Panel C). Co-immunoprecipitation revealed that removal of RRM2 or the coiled-coil region substantially reduced HOXA1 binding (Panel D), identifying these domains as critical for the interaction.\nGuided by our structural model of the HOXA1-HD–NONO-Coil interface (Figure 4J), we further introduced a point mutation (E278A) within the coiled-coil region at a predicted contact site. This single-residue substitution dramatically weakened the NONO–HOXA1 interaction (Panel E). Functionally, CHX chase assays demonstrated that the E278A mutant failed to stabilize HOXA1, whereas WT NONO preserved HOXA1 protein levels over time (Panels F, G).\nTogether, these results show that NONO stabilizes HOXA1 in the nucleus and that this stabilizing function depends on the integrity of NONO’s RRM2 and coiled-coil domains. These findings provide strong mechanistic support for the reviewer’s suggestion that NONO plays a chaperone-like role in regulating HOXA1 function.\nThe results shown here in Panels A–G have been integrated into the revised manuscript as Fig. 4. Corresponding descriptions have been updated in the revised text (Page 15 and 16, lines 549-609).\n\nReviewer #2 - Major concerns 4:\n[...]\nResponse:\nWe thank the reviewer for emphasizing the importance of evaluating HOXA1–DNA binding, particularly given that the HOXA1 homeodomain (HD) mediates its interaction with chromatin. To clarify whether NONO directly affects HOXA1’s chromatin affinity, we performed additional mechanistic analyses.\nFirst, we examined whether NONO influences HOXA1 homodimerization, a property known to enhance HOXA1 DNA-binding capacity. Co-immunoprecipitation in day-2 differentiated hiPSCs expressing HOXA1-HA and HOXA1-FLAG revealed that HOXA1 homodimer formation is markedly reduced in NONO-KO cells (Panels A–B). We further showed that HOXA1 nuclear protein stability is substantially decreased in NONO-KO cells (Panels C–D). Together, these findings support a model in which NONO promotes HOXA1 chromatin engagement by stabilizing the protein and supporting its dimeric state.\nThe results shown here in Panel A, B have been integrated into the revised manuscript as Fig. 5K, L. Panel C, D have been integrated into the revised manuscript as Fig. 4M. Corresponding descriptions have been updated in the revised text (Page 20, lines 779-783; Page 16, lines 598-602).\nRegarding the reviewer’s concern that reduced HOXA1 ChIP-seq enrichment in NONO-KO hiPSCs might be attributable to altered cell composition rather than loss of NONO itself, we performed an additional experiment in a controlled, lineage-independent system. HOXA1-HA was overexpressed at comparable levels in WT and NONO-KO HEK293T cells, and HOXA1 chromatin occupancy was quantified by ChIP–qPCR using an anti-HA antibody. Even in this uniform cellular context, HOXA1 binding was significantly diminished at multiple target promoters, including MESP1, PDGFRA, HAS2, and MIXL1, in NONO-KO cells (Panel A).\nThese results confirm that the reduction in HOXA1 ChIP signal is not solely due to cell-type composition differences but reflects an intrinsic requirement for NONO in enabling HOXA1 chromatin binding.\n\nReviewer #2 - Major concerns 5:\n[...]\nResponse:\nWe appreciate the reviewer’s valid point regarding the importance of validating mechanistic findings in an in vivo context. We fully agree that hiPSC-derived cardiomyocyte systems, while powerful, do not fully recapitulate embryonic development.\nHowever, the primary aim of this study was to delineate the molecular mechanism by which NONO cooperates with HOXA1 during early cardiac differentiation. The hiPSC model provides a highly controlled environment that allows us to isolate and interrogate these interactions without the confounding systemic effects and compensatory pathways present in whole-animal systems. Establishing such mechanistic foundations in vitro is an essential step before moving to more complex in vivo models.\nIn accordance with the reviewer’s suggestion, we have expanded the Discussion to explicitly acknowledge this limitation and to emphasize that testing the NONO–HOXA1 regulatory axis in an appropriate animal model represents an important direction for future research.\nDiscussion part:\nWe acknowledge that in vitro models cannot fully recapitulate the precise chronometry of embryonic development, nor do they possess the complex spatial architecture of native organogenesis. However, the hiPSC system offers an ideal, controlled platform for dissecting molecular mechanisms. Therefore, validating our findings in vivo is a critical next step. Future studies using Nono-deficient mice will determine whether the molecular defects observed here translate into anatomical malformations during heart development.\nCorresponding descriptions have been updated in the revised text (Page 28, lines 1088-1094)\n\nReviewer #2 - Major concerns 6:\n[...]\nResponse:\nWe thank the reviewer for this insightful comment. As noted, the observed reduction in nuclear β-catenin could arise either from impaired nuclear translocation or from decreased stability of the protein once in the nucleus. To distinguish between these possibilities, we examined the stability of nuclear β-catenin in the absence of HOXA1 or NONO.\nWe generated HOXA1-KO and NONO-KO HEK293T cell lines and performed cycloheximide (CHX) chase assays, collecting nuclear lysates at 0, 4, 8, and 12 hours after CHX treatment. These experiments revealed that loss of either HOXA1 or NONO markedly accelerates the decay of nuclear β-catenin, whereas cytoplasmic β-catenin levels remained unchanged (Panel A-D).\nThese findings indicate that HOXA1 and NONO are required not primarily for β-catenin nuclear import but rather for maintaining the stability of β-catenin within the nucleus. This result provides a mechanistic explanation for the reduced nuclear β-catenin observed in NONO-KO and HOXA1-KO cells and supports the conclusion that both factors promote canonical Wnt signaling by stabilizing β-catenin in the nuclear compartment.\nThe results shown here in Panels A–D have been integrated into the revised manuscript as Fig. 6E, F. Corresponding descriptions have been updated in the revised text (Page 23, lines 886-891).\n\nReviewer #2 - Major concerns 7:\n[...]\nResponse:\nWe thank the reviewer for this insightful suggestion. To test whether enhanced WNT activation could rescue the differentiation defect in NONO-KO cells, we performed a CHIR99021 dose–response experiment during the early induction phase. Increasing the CHIR concentration from the standard condition to 10 μM and 12 μM produced a dose-dependent improvement in TNNT2⁺ cardiomyocyte differentiation. This effect was further confirmed by RT-qPCR, which showed significantly increased expression of TNNT2, MYH6, and MYH7 in the 10–12 μM groups compared with the standard condition. In contrast, increasing the CHIR concentration to 14 μM markedly reduced differentiation efficiency, consistent with previous studies showing that supraphysiological CHIR levels can induce off-target cytotoxicity and disrupt proper lineage specification12.\nCollectively, these data indicate that NONO deficiency reduces cellular sensitivity to WNT signaling, thereby raising the activation threshold required for mesoderm induction, and that this defect can be partially overcome by appropriately increasing the dosage of a WNT agonist.\nThe results shown here in Panels A–C have been integrated into the revised manuscript as Fig. 6M-O. Corresponding descriptions have been updated in the revised text (Page 23 and 24, lines 911-940).\n\nReviewer #2 - Major concerns 8:\n[...]\nResponse:\nWe thank the reviewer for this insightful suggestion. To validate the pathogenic effects of NONO variants identified in CHD patients, and to further evaluate their functional impact during human cardiomyocyte differentiation, we performed a DOX-inducible overexpression assay in a NONO-knockout hiPSC line. Four nonsense mutations (c.217C>T [p.Arg73*], c.550C>T [p.Arg183*], c.1009C>T [p.Arg337*], and c.1093C>T [p.Arg365*])13-15,which have been clinically associated with congenital cardiac defects including NCC, VSD, PDA, and aortic arch anomalies, were selected for functional validation. DOX-induced expression of WT NONO or the mutant variants was initiated during the early differentiation window (days 1–2).\nImmunofluorescence staining revealed that NONO-KO cardiomyocytes exhibited severely disorganized sarcomeres at day 15, whereas DOX-induced WT NONO expression restored well-organized α-ACTININ/cTNT structures. In contrast, none of the four pathogenic variants were able to rescue sarcomere organization, with the majority of cells retaining disorganized sarcomere structures (Panel A). Consistent with this structural defect, flow cytometry showed a robust increase in TNNT2⁺ cardiomyocytes upon WT NONO re-expression (33.5% vs. 4.0% without DOX), whereas all four mutant variants produced only minimal TNNT2⁺ populations (8.8–11.9%) (Panel B, C). qRT-PCR revealed a robust restoration of TNNT2, MYH6, and MYH7 expression by WT NONO, while all pathogenic variants failed to activate these sarcomeric genes (Panel D).\nTaken together, these results demonstrate that NONO variants associated with CHD are functionally pathogenic and incapable of supporting cardiomyocyte lineage commitment, thereby providing direct evidence that NONO dysfunction contributes to disrupted cardiac development in humans.\nThe results shown here in Panels A–D have been integrated into the revised manuscript as Supplementary Fig. 5. Corresponding descriptions have been updated in the revised text (Page 11, lines 363-378).\n\nReviewer #2 - Major concerns 9:\n[...]\nResponse:\nWe sincerely thank the reviewer for this constructive suggestion, which has significantly strengthened our mechanistic conclusions regarding the cooperative interaction between NONO and HOXA1.\nTo validate whether NONO physically co-occupies target loci with HOXA1, we performed the suggested sequential ChIP (Re-ChIP) assays using a doxycycline-inducible HOXA1-HA-overexpressing hiPSC line (WTHOXA1-HA OE) differentiated to Day 2. To ensure the preservation of protein-protein-DNA complexes during the sequential immunoprecipitation, we employed a rigorous native elution strategy: chromatin was first immunoprecipitated with an anti-HA antibody and eluted competitively using HA peptide, to maintain complex integrity. The eluate was then subjected to a second immunoprecipitation using an anti-NONO antibody. As visualized in the newly added genome browser tracks (Panel A), distinct Re-ChIP signals were successfully recovered at the regulatory regions of key cardiac development genes, including EOMES, TBXT, DVL2, and MECOM6,16. Crucially, these Re-ChIP peaks precisely align with the overlapping regions observed in the individual NONO and HOXA1 ChIP-seq tracks, providing direct evidence that NONO and HOXA1 form a physical transcriptional complex on chromatin to co-regulate these key developmental genes.\nThe results shown here in Panels A have been integrated into the revised manuscript as Fig. 5A. Corresponding descriptions have been updated in the revised text (Page 19, lines 710-717).\nTo examine the relationship between NONO and HOXA1, we generated a double-knockout (Double KO) hiPSC line by deleting HOXA1 exon 1 in the NONO-KO background (Panel A). qRT–PCR confirmed efficient loss of HOXA1 expression (Panel B). Upon cardiac differentiation, the Double KO cells exhibited a markedly stronger defect than the NONO-KO line alone. At Day 15, the proportion of cTNT⁺ cardiomyocytes decreased from ~12.7% in NONO-KO cultures to ~4.9% in the Double KO line (Panel C). Immunofluorescence further showed that ~96% of Double KO cells displayed disorganized sarcomeres, compared with ~75% in NONO-KO cardiomyocytes (Panel D), indicating that loss of HOXA1 accentuates the structural abnormalities caused by NONO deficiency.\nThe results shown here in Panel A, B have been integrated into the revised manuscript as Supplementary Fig. 1K, L. Panel C, D have been integrated into the revised manuscript as Fig. 5M, N. Corresponding descriptions have been updated in the revised text (Page 20, lines 784-789).\n\nReviewer #2 - Minor Comments 1:\n[...]\nResponse:\nWe thank the reviewer for this suggestion. In response, we performed karyotype analysis on WT hiPSCs and two independent NONO-KO clones (clone 3 and clone 11).\nThe results shown here in Panel A have been integrated into the revised manuscript as Supplementary Fig. 1C. Corresponding descriptions have been updated in the revised text (Page 4, lines 109-110).\n\nReviewer #2 - Minor Comments 2:\n[...]\nResponse:\nWe acknowledge that the specific representative trace shown in Fig 1F for the NONO-KO group appears to have a robust amplitude, which visually contradicts a \"reduced intensity\" phenotype. When selecting this representative image, our primary intention was to highlight the most striking and statistically significant phenotypes: the drastically reduced beating frequency and the prolonged calcium decay. Notably, this specific pattern of calcium handling is consistent with previous findings; a similar trace morphology was observed in Figure 3F6.\n\nReviewer #2 - Minor Comments 3:\n[...]\nResponse:\nWe thank reviewer for this suggestion. In response, we performed Western blot on day 2 differentiated WT, NONO-KO (clone3) and NONO-RE cells with or without DOX induction. The results show that NONO re-expression provides only a partial rescue at day 2. The results, now shown in Fig. 2A, confirm that NONO re-expression reaches only a partial level compared to endogenous NONO, consistent with the partial rescue phenotype.\nThe results shown here in Panel A have been integrated into the revised manuscript as Supplementary Fig. 4B. Corresponding descriptions have been updated in the revised text (Page 9, lines 248-249).\n\nReviewer #2 - Minor Comments 4\n[...]\nResponse:\nWe thank the reviewer for the suggestion. We have now clearly labeled that the heatmaps show rescue-associated changes for genes in each group (A–I).\nThe results shown here in Panel have been integrated into the revised manuscript as Supplementary Fig. 6A. Corresponding descriptions have been updated in the revised text (Page 11, lines 393-394).\n\nReviewer #2 - Minor Comments 5\n[...]\nResponse:\nWe thank the reviewer for pointing out this important correction. We apologize for the misuse of the term “hESC” and have revised the manuscript to correctly refer to the cells as “hiPSCs” in all relevant sections.\n\nSummary and final appreciation for Reviewer #2\nWe are very grateful to Reviewer #2 for the careful and constructive evaluation of our work. In response to these comments, we have (i) refined our terminology to focus on cardiomyocyte differentiation rather than maturation, (ii) added time-course single-cell RNA-seq analyses to disentangle true transcriptional changes from shifts in cell composition, (iii) defined the structural basis and functional consequences of the NONO–HOXA1 interaction using domain mapping and protein stability assays, (iv) demonstrated that NONO is intrinsically required for HOXA1 chromatin binding and for maintaining nuclear β-catenin, thereby strengthening the link between the NONO–HOXA1 complex and canonical Wnt signaling, (v) tested the sensitivity of NONO-KO cells to graded Wnt activation, and (vi) functionally validated CHD-associated NONO variants in a human hiPSC-based model. We have also addressed all minor points, including karyotype analysis of edited hiPSC lines, clarification of figure labeling and grouping, and correction of cell-type nomenclature. We believe that these additions and revisions have substantially strengthened the mechanistic depth, internal consistency, and translational relevance of the manuscript. We sincerely thank Reviewer #2 again for the insightful guidance that has been instrumental in improving this study."}, {"round": 2, "reviewer_comments": {"reviewer_1": "In this revised manuscript, the authors have thoroughly addressed all my comments. The overall quality of the study has been subsequently improved, and the data are now clearly presented and well interpreted.\n\nIn response to my comments, the authors have refined their conclusions regarding the role of NONO in early cardiac specification rather than late-stage maturation. Additional experiments, including new flow cytometry and single-cell RNA-seq analyses to quantify cardiac progenitor populations and cell-state transitions, have helped them better assess the role of NONO during precardiac mesoderm development. Cardiomyocytes (CM) are now better characterized using markers such as Myl2 and Myl7, which indicate that the CMs generated have predominantly atrial identity. The rescue experiments are also better characterized thanks to the inclusion of early markers. A temporal expression analysis, shown in Supplementary Figure 2, highlights the progression of CM differentiation in their model. Integration of the NONO ChIP-seq data with transcriptomic profiles has enabled the authors to identify direct targets of NONO. Clarifying the temporal relationships between NONO, MESP1, and key developmental markers has strengthened the understanding of their model and the role of NONO in CM differentiation. The discussion now includes a section on the “negative regulation of stem cell differentiation” gene set and provides a more careful comparison of their findings with in vivo and previous studies.\n\nAddressing the homodimerization of NONO and HOXA1 has further strengthened the mechanistic model of NONO–HOXA1 cooperation by characterizing their dimerization properties. Through sophisticated approaches such as Re-ChIP, they map interaction domains and assess how loss of NONO affects HOXA1 chromatin binding and downstream Wnt signaling.\n\nOverall, the revised manuscript demonstrates significant improvements in clarity, and mechanistic insight.", "reviewer_2": "The authors did a commendable work at addressing my comments. Most of my questions were well answered by supplemental experiments, including scRNA-seq (for the major comments 2), protein stability assays(for the major comments 3 and 6), domain-mapping experiments (for the major comments 3), mechanistic analyses (for the major comments 4), CHIR99021 dose–response experiment (for the major comments 7), validation in hiPSC lines with patient-related genetic variants (for the major comments 8), reCHIP (for the major comments 9), double knockout hiPSC lines generation (for the major comments 9) and karyotype detection (for the minor comments 1). For the major comments 5, although the authors didn’t provide additional experiments, they gave a reasonable explanation in the discussion. The authors also corrected their misuse of “maturation” and “hiPSC”. The only imperfection is that replicates and a quantification are needed for Supplementary Fig. 4B. I have no further requests."}, "author_response": "We sincerely thank both Reviewer #1 and Reviewer #2 for their thoughtful and constructive comments. Their insightful suggestions have significantly improved the quality of our manuscript. Below, we provide a point-by-point response to each comment. Reviewer comments are shown in italic. \n\nReviewer #1 (Remarks to the Author):\n[...]\nWe sincerely thank the reviewer for their positive assessment of our revised manuscript and for their constructive feedback throughout the review process. We are pleased to learn that the additional experiments—including the detailed characterization of cardiac progenitor populations and the mechanistic insights gained from the homodimer and heterodimer protein interaction assays—have satisfactorily addressed your concerns. We agree that these revisions have significantly strengthened the conclusions of our study.\n\nReviewer #1 (Remarks on code availability):\nThe GitHub link provides access to the bioinformatic datasets and analysis scripts used in this study.\nWe thank the reviewer for confirming the accessibility of the code. we have archived the code on Zenodo as indicated in the revised manuscript. (https://doi.org/10.5281/zenodo.18059608.)\n\nReviewer #2 (Remarks to the Author):\n[...]\nWe are grateful for the reviewer’s encouraging comments and are pleased that the extensive additional experiments—ranging from scRNA-seq analyses to the generation of double-knockout lines—have resolved the major concerns.\nRegarding the specific request for Supplementary Fig. 4B, we have performed the necessary biological replicates as suggested. We have also performed the corresponding quantification analysis (~60% efficiency), which is now presented in the newly added Supplementary Fig. 4C.\nWe thank the reviewer for their thorough assessment, which has helped ensure the robustness of our data."}]

Decision letter