p16INK4a promotes myocardial ischemia-reperfusion injury by regulating bile acid transport via Slco1a4
Tingting Yang, Qiulian Zhou, Yihua Bei, Danni Meng, Songwei Ai, Yuhui Zhang, Jian Zhang, Li Liu, Hongjian Chen, Xue Pan, Xiaohang Yin, Michail Spanos, Guoping Li, Dragos Cretoiu, Joost P G Sluijter, Anthony Rosenzweig, Junjie Xiao
TL;DR
The protein p16INK4a worsens heart damage after blood flow is restored by affecting bile acid transport in heart cells.
Contribution
This study reveals a new regulatory pathway involving p16INK4a, CUGBP1, Npas2, and Slco1a4 in cardiac ischemia-reperfusion injury.
Findings
p16INK4a is upregulated in I/R injury and promotes cardiomyocyte apoptosis.
p16INK4a inhibition reduces cell death in I/R models.
p16INK4a regulates Npas2 via CUGBP1, leading to increased Slco1a4 and bile acid transport.
Abstract
Myocardial ischemia-reperfusion (I/R) injury remains a significant challenge in cardiovascular medicine, with its molecular mechanisms still not fully understood. Screening the GEO and Comparative Toxicogenomics Database as well as spatial multi-omics data, we identify Cdkn2a, encoding p16INK4a, as a determinant in I/R injury. Cdkn2a expression is elevated in the myocardium of ischemic cardiomyopathy patients and p16INK4a protein is enriched in cardiomyocytes within ischemic zones of myocardial infarction tissues. We find that p16INK4a is consistently upregulated in both in vivo and in vitro I/R models, promoting apoptosis in neonatal rat cardiomyocytes (NRCMs) and human embryonic stem cell-derived cardiomyocytes (hESC-CMs) exposed to oxygen-glucose deprivation/reperfusion (OGD/R). p16INK4a inhibition confers cellular protection, an effect also observed in in vivo I/R injury models.…
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Figure 15- —National Natural Science Foundation of China
- —National Natural Science Foundation of China
- —http://dx.doi.org/10.13039/501100003399Science and Technology Commission of Shanghai Municipality (STCSM)
- —Dawn Program of Shanghai Education Commission
- —Oriental Scholars of Shanghai Universities
- —http://dx.doi.org/10.13039/501100002858China Postdoctoral Science Foundation (中国博士后科学基金)
- —Postdoctoral Fellowship Program of CPSF
- —Shanghai Sailing Program
- —National Institutes of Health
- —The American Heart Association
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Taxonomy
TopicsDrug Transport and Resistance Mechanisms · Cardiac Ischemia and Reperfusion · Kruppel-like factors research
Introduction
Ischemic heart disease remains the leading cause of cardiovascular mortality worldwide (Ezzatvar et al, 2021; Hausenloy and Yellon, 2013), with reperfusion strategies such as percutaneous coronary intervention (PCI), thrombolysis, and surgical revascularization being the cornerstone of restoring function within the ischemic myocardium (Eefting et al, 2004; Takemura et al, 2009). However, reperfusion may paradoxically provoke additional cardiac insults, including calcium overload, oxidative stress, and mitochondrial dysfunction, culminating in apoptosis, necrosis, and adverse remodeling of the heart (Zhou et al, 2017; Zweier and Talukder, 2006). Myocardial infarction (MI) and ischemic injury exhibit a higher prevalence among older patients, and elderly patients appear to have greater cardiac dysfunction and worse injury despite successful reperfusion therapy, commonly developing heart failure in the ensuing five years (Ezekowitz and Kaul, 2010; Lesnefsky et al, 1996). The molecular intricacies driving myocardial ischemia/reperfusion (I/R) injury are complex, and despite significant research efforts, the full spectrum of molecular events that follow I/R is not completely understood. Unraveling these mechanisms is critical for identifying new therapeutic avenues.
The protein p16^INK4a^, encoded by the Cdkn2a gene, regulates cell cycle arrest and cellular senescence (Koh et al, 1995; Lukas et al, 1995; Mas-Bargues et al, 2017; Qiu et al, 2023), and its expression increases with age in animals (Baker et al, 2016) and humans (Li et al, 2011). In the context of cardiovascular pathology, such as atherosclerosis, p16^INK4a^ has been implicated in endothelial dysfunction and proliferative changes, often observed at sites prone to plaque formation (Gimbrone and Garcia-Cardena, 2016; Zhu et al, 2021). Conflicting evidence has emerged regarding the role of p16^INK4a^ after I/R, with prior reports suggesting either deleterious (Khatiwala et al, 2018) or beneficial effects (Shi et al, 2022).
To identify key modulators of cardiac I/R injury, we used the Comparative Toxicogenomics Database (CTD; http://ctdbase.org/), which provides manually curated information on chemical–gene/protein interactions and integrates these data to infer gene–disease associations (Huang et al, 2013). We identified p16^INK4a^ protein coding gene Cdkn2a as one of a small number of genes common to all searched conditions. In vitro and in vivo gain- and loss-of-function experiments demonstrated an important role for p16^INK4a^ in myocardial I/R injury, through regulation of Npas2/Slco1a4 axis. These studies provide a mechanistic link between aging biology and increased myocardial I/R injury.
Results
Upregulation of p16INK4a in response to cardiac I/R injury
To identify key mediators of myocardial ischemia/reperfusion (I/R) injury, we analyzed a published RNA-seq dataset from the Gene Expression Omnibus (GEO; GSE:161323) associated with myocardial I/R injury (P < 0.01, |logFC| > 2). By cross-referencing this finding with the Comparative Toxicogenomics Database (CTD; http://ctdbase.org/) for associations with myocardial ischemia (CTD_D017202), myocardial infarction (CTD_D009203) and myocardial reperfusion injury (CTD_D015428), we identified seven factors common to all searched conditions (Fig. 1A). Subsequent qPCR analysis confirmed elevated expression of Myc, Cxcl1, S100a9, Serpine1 and p16^INK4a^ transcript variant of Cdkn2a, as well as Thbs1 in both the infarct and border zones of hearts from mice after acute I/R, as well as hearts assessed for remodeling 3 weeks post I/R injury (Fig. 1B). Consistent upregulation of Myc, Cxcl1, Cdkn2a, and Thbs1 was also evident in neonatal rat cardiomyocytes (NRCMs) subjected to an in vitro oxygen-glucose deprivation/reperfusion (OGD/R) model (Fig. 1C). In both rats and mice, the Cdkn2a gene gives rise to two distinct proteins, p16^INK4a^ and p19^ARF^. According to current public database annotations, these proteins in mice are encoded by separate transcript variants and are therefore analyzed independently as Cdkn2a (p16^INK4a^) and Cdkn2a (p19^ARF^). By contrast, only a single Cdkn2a transcript is currently annotated in rats. In humans, Cdkn2a primarily gives rise to two distinct proteins, p16^INK4a^ and p14^ARF^. Notably, in vitro hypoxic-glucose deprivation/reperfusion (OGD/R) human embryonic stem cell-derived cardiomyocytes (hESC-CMs) revealed consistent upregulation solely of the p16^INK4a^ variants of Cdkn2a, alongside Thbs1 (Fig. 1D). Moreover, the expression of the Cdkn2a was also elevated in neonatal rat cardiac fibroblasts (NRCFs) under TGF-β-induced fibrotic activation (Fig. 1E). Considering the consistent elevation of the Cdkn2a transcript variant p16^INK4a^ across various species and multiple models, and the previously established role of Thbs1 in myocardial I/R injury (Kelm et al, 2020), we focused our investigation primarily on p16^INK4a^. We then confirmed the upregulation of p16^INK4a^ protein expression in both in vivo I/R and in vitro OGD/R models via immunoblotting. In contrast, p19^ARF^ exhibited increased expression specifically in the remodeling model of I/R injury at 3 weeks, while p14^ARF^ showed unchanged expression in the hESC-CMs OGD/R model (Fig. EV1A–D). Additionally, increased p16^INK4a^ protein expression was also observed in NRCFs subjected to TGF-β-induced fibrotic activation (Fig. EV1E). Moreover, in the canine myocardial I/R injury model, we observed elevated levels of p16^INK4a^ mRNA and protein expression (Figs. 1F and EV1F). Through the analysis of clinical samples, we subsequently identified a significant increase in the expression of the p16^INK4a^ protein-coding gene Cdkn2a in the myocardium of patients with ischemic cardiomyopathy (ICM) compared to control samples (NF, non-failing hearts) (Sweet et al, 2018; Data ref: Sweet et al, 2018) (Fig. 1G). ICM is the consequence of cumulative ischemic and reperfusion injuries over time, reflecting chronic pathological changes and manifesting as chronic inflammation, fibrosis, and myocardial remodeling. Taking a step further, our analysis of the spatial multi-omic map of human myocardial infarction data revealed that the expression of the p16^INK4a^ protein-coding gene Cdkn2a was markedly elevated in cardiomyocytes and fibroblasts within the ischemic zone (IZ) of myocardial infarction tissues (Kuppe et al, 2022; Data ref: Kuppe et al, 2022) (Fig. 1H,I). Collectively, these findings strongly suggest that p16^INK4a^ is likely a critical driver of cardiac I/R injury.Figure 1. Upregulation of p16^INK4a^ in response to cardiac I/R injury.(A) The Venn diagrams demonstrate the overlap among aberrant expression of genes in cardiac ischemia-reperfusion (GSE161323, identified using the limma statistical method; p < 0.01, |logFC| > 2) and myocardial ischemia (CTD_D017202), myocardial infarction (CTD_D009203), myocardial reperfusion injury (CTD_D015428). (B) qRT-PCR analysis of Agt, Myc, Cxcl1, Fosb, S100a9, Serpine1, Cdkn1a, Cdkn2a (p16^INK4a^), Cdkn2a (p19^ARF^), Col3a1 and Thbs1 expression in both acute myocardial I/R injury model and myocardial I/R injury remodeling model (n = 6 per group, biological replicates). Statistical analysis was performed by one-way ANOVA followed by Bonferroni or Dunnett T3 post hoc tests for (B) (Sham vs. Acute I/R (Infarct): Agt, **p = 0.004; Myc, *p = 0.014; Cxcl1, ****p < 0.0001; Fosb, **p = 0.001; S100a9, **p = 0.002; Serpine1, **p = 0.009; Cdkn2a (p16^INK4a^), **p = 0.008. Sham vs. Acute I/R (Border): Agt, **p = 0.001; Myc, **p = 0.003; Cxcl1, *p = 0.011; Fosb, **p = 0.001; S100a9, ****p < 0.0001; Serpine1, *p = 0.019; Cdkn2a (p16^INK4a^), **p = 0.001), by Kruskal-Wallis test with the original FDR method of Benjamini and Hochberg for (B) (Sham vs. Acute I/R (Infarct): Cdkn1a, *p = 0.0386; Thbs1, **p = 0.0074. Sham vs. Acute I/R (Border): Thbs1, **p = 0.0073) and by unpaired Student’s t-test for (B) (Sham vs. I/R (3 W): Myc, ****p < 0.0001; Cxcl1, ***p = 0.0001; S100a9, **p = 0.008; Serpine1, **p = 0.001; Cdkn2a (p16^INK4a^), **p = 0.001; Cdkn2a (p19^ARF^), ***p = 0.0002; Col3a1, ****p < 0.0001; Thbs1, ****p < 0.0001). (C) qRT-PCR analysis of Agt, Myc, Cxcl1, Fosb, S100a9, Serpine1, Cdkn1a, Cdkn2a (p16^INK4a^, p19^ARF^), Col3a1 and Thbs1 expression in NRCMs treated with OGD/R (n = 6 per group, biological replicates). Statistical analysis was performed by unpaired Student’s t-test. Agt, **** p < 0.0001; Myc, **p = 0.002; Cxcl1, **p = 0.001; Fosb, *p = 0.028; S100a9, *p = 0.028; Cdkn2a (p16^INK4a^, p19^ARF^), **p = 0.002, Col3a1, **** p < 0.0001; Thbs1, **** p < 0.0001). (D) qRT-PCR analysis of Agt, Myc, Cxcl1, Fosb, S100a9, Serpine1, Cdkn1a, Cdkn2a (p16^INK4a^), Cdkn2a (p14^ARF^), Col3a1 and Thbs1 expression in hESC-CMs treated with OGD/R (n = 6 per group, biological replicates). Statistical analysis was performed by unpaired Student’s t-test. Agt, ***p = 0.0001; Myc, ****p < 0.0001; Fosb, *p = 0.027; Cdkn2a (p16^INK4a^), **p = 0.001; Col3a1, ***p = 0.0001; Thbs1, **** p < 0.0001) and by Mann-Whitney U test for S100a9. (E) qRT-PCR analysis of Cdkn2a (p16^INK4a^, p19^ARF^) expression in NRCFs treated with TGF-β (n = 6 per group, biological replicates). Statistical analysis was performed by unpaired Student’s t-test, *p = 0.011. (F) qRT-PCR analysis of p16^INK4a^ expression in a canine myocardial I/R injury model (n = 4 per group, biological replicates). Statistical analysis was performed by unpaired Student’s t-test, **p = 0.004. (G) Analyzing of p16^INK4a^ protein coding gene Cdkn2a mRNA expression in the myocardium of patients with ischemic cardiomyopathy (ICM) (n = 14 for NF, n = 13 for ICM; biological replicates). Statistical analysis was performed by unpaired Student’s t-test, **p = 0.004. (H) Analysis of Cdkn2a gene expression in cardiomyocytes within the ischemic zone of human cardiac after myocardial infarction. (I) Analysis of Cdkn2a gene expression in fibroblasts within the ischemic zone of human cardiac after myocardial infarction. Data represent mean ± SD. Source data are available online for this figure.
Inhibition of p16INK4a expression reduces OGD/R-induced apoptosis in vitro
To elucidate the function of p16^INK4a^ in this context, we employed shRNA to suppress its expression (sh-p16^INK4a^) in NRCMs and hESC-CMs and subsequently validated the lentiviral knockdown effects via qPCR and immunoblotting (Appendix Fig. S1A,B). We found that p16^INK4a^ knockdown substantially reduced NRCMs and hESC-CMs apoptosis induced by OGD/R, with fewer TUNEL-positive cells (Fig. 2A,B) and decreased ratios of Bax/Bcl2 and Cleaved-caspase3/Caspase3 protein expression in NRCMs within the OGD/R model (Fig. 2C). In addition, we also employed lentivirus-mediated overexpression of p16^INK4a^ (p16^INK4a^ OE) in both NRCMs and hESC-CMs, followed by validation of the lentivirus overexpression effects via qPCR and immunoblotting (Appendix Fig. S1C,D). We found that overexpression of p16^INK4a^ in NRCMs and hESC-CMs did not affect apoptosis at baseline but exacerbated OGD/R-induced apoptosis (Fig. 2D–F).Figure 2. Inhibition of p16^INK4a^ expression reduces OGD/R-induced apoptosis in vitro.(A) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with sh-p16^INK4a^ or shScr under OGD/R or Control condition (n = 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. **p = 0.008; ****p < 0.0001. (B) Representative images of immunofluorescence staining and quantification of the TUNEL positive hESC-CMs treated with sh-p16^INK4a^ or shScr under OGD/R or Control condition (n = 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. ****p < 0.0001. (C) Immunoblotting analysis of NRCMs apoptosis by detection of Bax, Bcl2, caspase-3, and Cleaved-caspase3 in Control or OGD/R-induced apoptosis model treated with sh-p16^INK4a^ or shScr (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. Bax/Bcl2 ratio: ****p < 0.0001; Cleaved-caspase3/Caspase3 ratio: Control: shScr vs. OGD/R: shScr, ***p = 0.0005; OGD/R: shScr vs. OGD/R: sh-p16^INK4a^, ***p = 0.0004. (D) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with p16^INK4a^ OE or Fugw under OGD/R or Control condition (n = 4 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. ****p < 0.0001. (E) Representative images of immunofluorescence staining and quantification of the TUNEL positive hESC-CMs treated with p16^INK4a^ OE or Fugw under OGD/R or Control condition (n = 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. ****p < 0.0001. (F) Immunoblotting analysis of NRCMs apoptosis by detection of Bax, Bcl2, caspase-3, and Cleaved-caspase-3 in Control or OGD/R-induced apoptosis model treated with p16^INK4a^ OE or Fugw lentivirus (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. *p = 0.0111; **p = 0.0052; ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
To further evaluate the role of p16^INK4a^ in cardiac fibroblasts, we examined its effect in NRCFs following TGF-β-induced fibrotic activation. Neither knockdown nor overexpression of p16^INK4a^ significantly altered fibrotic responses, as indicated by comparable α-SMA immunofluorescence intensity, EdU incorporation, and mRNA levels of fibrotic markers such as Col1a1, Col3a1, and α-SMA (Appendix Fig. S2A–D). Collectively, these findings indicate that p16^INK4a^ does not significantly influence fibrotic activation in cardiac fibroblasts, prompting us to focus on its role in cardiomyocyte.
Cardiac-specific p16INK4a deletion protects against cardiac I/R in vivo
To examine the role of p16^INK4a^ in I/R injury in vivo, we generated cardiac myocyte-specific p16^INK4a^ conditional knockout mice (p16^INK4a^ cKO) by crossing p16^INK4a flox/flox^ mice with α-MHC-Cre transgenic mice. Notably, α-MHC-Cre mice exhibited no apparent morphological abnormalities in cardiac tissue compared to p16^INK4a flox/flox^ mice (Fig. EV2A). p16^INK4a flox/flox^ littermates lacking Cre expression were used as controls (Control). Both groups were subjected to cardiac I/R injury (Fig. 3A). Firstly, qPCR analysis revealed a reduction in p16^INK4a^ expression in the myocardial tissue of p16^INK4a^ cKO mice, whereas the expression of another Cdkn2a transcript, p19^ARF^, was not affected (Fig. 3B). After cardiac I/R injury, the infarct area was dramatically reduced in p16^INK4a^ cKO mice compared to littermate non-cKO control mice, as indicated by TTC staining during I/R surgery (Fig. 3C). Additionally, following I/R injury, p16^INK4a^ cKO mice displayed reduced myocardial apoptosis, evidenced by fewer TUNEL-positive cells and decreased ratios of Bax/Bcl2 and Cleaved-caspase3/Caspase3 protein expression in the infarct and border zones, and further exhibited enhanced cardiomyocyte proliferation, as demonstrated by increased Ki67-positive nuclei and upregulated protein expression of PCNA and Cyclin D1 after acute myocardial I/R injury (Figs. 3D and EV2B–E). Finally, the levels of lactate dehydrogenase (LDH) in the serum of p16^INK4a^ cKO mice after I/R surgery were also markedly decreased (Fig. 3E). These results serve as in vivo evidence that p16^INK4a^ deletion reduces infarct size and alleviates cardiomyocytes apoptosis and necrosis and enhances cardiomyocyte proliferation after acute cardiac I/R injury.
Based on these promising results, we sought to investigate whether p16^INK4a^ deletion also mitigates I/R-induced pathological cardiac remodeling (Fig. 3F). Three weeks after myocardial I/R, echocardiographic assessment revealed that p16^INK4a^ cKO mice maintained superior cardiac function post-remodeling, with substantially better ejection fraction (EF) and fractional shortening (FS) (Fig. 3G). Moreover, myocardial fibrosis as measured by Masson’s trichrome staining was significantly lower in p16^INK4a^ cKO mice, where lower expression of fibrosis-associated genes Col1a1, Col3a1, and Ctgf were also found according to qPCR analysis (Fig. 3H,I). Finally, the expression of heart failure markers Anp, Bnp, and β-Mhc was also suppressed in p16^INK4a^ cKO mice (Fig. 3J), indicating protection from I/R-induced pathological cardiac remodeling, whereas cardiomyocyte proliferation, as assessed by Ki67-positive nuclei, showed no significant difference (Fig. EV2F).
To assess the in vivo effects of increased p16^INK4a^, we performed cardiomyocyte-specific induction of p16^INK4a^ expression by injecting mice with AAV9 that carries p16^INK4a^ under the cardiomyocyte-specific cardiac troponin T (cTnT) (cTnT-p16^INK4a^-AAV9) followed by acute I/R injury surgery (Fig. EV3A). qPCR and Western blot results showed that intravenous tail injection of cTnT-p16^INK4a^-AAV9 in mice led to increased expression of p16^INK4a^ mRNA and protein in cardiac tissue (Fig. EV3B,C). Subsequent functional assays revealed that mice with cTnT-p16^INK4a^-AAV9 exhibited significantly larger myocardial infarct areas, increased cardiomyocyte apoptosis, reduced the expression of the cardiomyocyte proliferation marker Ki67, and greater myocardial necrosis 24 h post-surgery compared to cTnT-CTL-AAV9 (Fig. EV3D–H). Furthermore, in another group of mice that underwent I/R remodeling surgery one week after tail vein injection of cTnT-p16^INK4a^-AAV9 or cTnT-CTL-AAV9, we also observed that cTnT-p16^INK4a^-AAV9 increased the expression of p16^INK4a^ mRNA and protein in mouse cardiac tissue (Fig. EV4A–C). Additionally, mice treated with cTnT-p16^INK4a^-AAV9 showed markedly deteriorated cardiac function, elevated myocardial fibrosis, and pathological cardiac hypertrophy after three weeks, compared to cTnT-CTL-AAV9 injected mice (Fig. EV4D–G), whereas cardiomyocyte proliferation, as assessed by Ki67-positive nuclei, was not significantly affected (Fig. EV4H). These results reinforce the hypothesis that cardiac-specific overexpression of p16^INK4a^ exacerbates both acute myocardial injury and pathological cardiac remodeling following I/R.Figure 3. Cardiac-specific p16^INK4a^ deletion protects against cardiac I/R in vivo.(A) The schedule of acute myocardial I/R injury model establishment in p16^INK4a flox/flox^ (control) and α-MHC-Cre; p16^INK4a flox/flox^ (cKO) mice. (B) qRT-PCR analysis of p16^INK4a^ and p19^ARF^ expression in the hearts of p16^INK4a^ cKO mice and control mice (n = 6 per group, biological replicates). Statistical analysis was performed by unpaired Student’s t-test, ****p < 0.0001. (C) Representative images of TTC staining and quantification of the area at risk/left ventricular weight (AAR/LV) ratio and the infarct size/area at risk (INF/AAR) ratio in the hearts of p16^INK4a^ cKO mice and control mice treated with acute myocardial I/R injury (n = 6 per group, biological replicates). Scale bar: 2 mm. Statistical analysis was performed by unpaired Student’s t-test, ****p < 0.0001. (D) Representative images of immunofluorescence staining and quantification of the TUNEL positive cardiomyocytes in the hearts of p16^INK4a^ cKO mice and control mice treated with acute myocardial I/R injury (n = 6 per group, biological replicates). Scale bar: 20 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test, ****p < 0.0001. (E) LDH assays analysis serum levels of LDH in the hearts of p16^INK4a^ cKO mice and control mice treated with acute myocardial I/R injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test, ****p < 0.0001. (F) The schedule of myocardial I/R injury remodeling model establishment in p16^INK4a flox/flox^ (control) and α-MHC-Cre; p16^INK4a flox/flox^ (cKO) mice. (G) Representative echocardiographic images and left ventricular ejection fraction (EF), ventricular fractional shortening (FS) of p16^INK4a^ cKO mice and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Comparison of Ejection Fraction (%): *p = 0.0348; ****p < 0.0001. Comparison of Fractional Shortening (%): *p = 0.0262; ****p < 0.0001. (H) Representative images of Masson’s trichrome staining and quantification of fibrotic area (%) in the hearts of p16^INK4a^ cKO mice and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Scale bar, 50 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test, ****p < 0.0001. (I) qRT-PCR analysis of Col1a1, Col3a1 and Ctgf expression in the hearts of p16^INK4a^ cKO mice and control mice a following I/R remodeling injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Comparison of Col1a1expression: **p = 0.0048; ****p < 0.0001. Comparison of Col3a1expression: **p = 0.0012; ****p < 0.0001. Comparison of Ctgf expression: *p = 0.0204; ***p = 0.0002; ****p < 0.0001. (J) qRT-PCR analysis of Anp, Bnp and β-MHC expression in the hearts of p16^INK4a^ cKO mice and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Comparison of Anp expression: Sham: Control vs. I/R(3 W): Control, ***p = 0.0009; I/R(3 W): Control vs. I/R(3 W): p16^INK4a^ cKO, ***p = 0.0009. Comparison of Bnp expression: ****p < 0.0001. Comparison of β-MHC expression: **p = 0.0021; ***p = 0.0008. Data represent mean ± SD. Source data are available online for this figure.
p16INK4a transcriptionally induces Slco1a4 to modulate cardiomyocyte apoptosis
In an effort to elucidate the potential mechanisms underlying the regulation of cardiac I/R injury by p16^INK4a^, we conducted a comprehensive transcriptome analysis, utilizing RNA sequencing (RNA-seq) in p16^INK4a^ cKO mice (fold change > 2; p < 0.05) and a mouse model of I/R remodeling injury at 3 weeks (GEO, GSE241079) (fold change > 1.5; p < 0.05) (Fig. 4A,B). Cross-comparison of differentially expressed genes from both RNA-seq datasets highlighted three shared genes: Arnt1, Myh7, and Slco1a4, with only Slco1a4 showing decreased expression in p16^INK4a^ cKO mice and increased expression in a mouse model of I/R remodeling injury at 3 weeks (Fig. 4C). Simultaneously, Slco1a4 expression was decreased in cardiomyocytes transfected with sh-p16^INK4a^ lentivirus, while it was increased in those transfected with p16^INK4a^ overexpression lentivirus (Appendix Fig. S3A,B). Additionally, Slco1a4 expression was elevated in the cellular OGD/R model (Appendix Fig. S3C). We further assessed the expression of Slco1a4 in p16^INK4a^ cKO mice subjected to cardiac I/R injury. Notably, Slco1a4 was consistently upregulated in control mice undergoing remodeling 3 weeks post-I/R injury, as well as in the infarct and border zones of hearts from control mice after acute I/R. However, it was downregulated in p16^INK4a^ cKO mice that were subjected to cardiac I/R injury (Fig. 4D,E). Given that Slco1a4 is a member of the solute carrier organic anion transporter family, which is involved in the transport of bile acids, conjugated steroids, and prostaglandins (Granados and Nigam, 2024; Shan et al, 2023; Zhang et al, 2013), we measured the levels of three representative bioactive molecules: bile acid; dehydroepiandrosterone sulfate (DHEAS), a major conjugated steroid; and prostaglandin E2 (PGE2), a key prostaglandin extensively studied in cardiac physiology. Among these, only bile acid levels in cardiac tissue showed a positive correlation with Slco1a4 expression (Fig. 4F,G; Appendix Fig. S3D–G). Subsequently, we utilized Slco1a4 overexpression lentivirus and Slco1a4 siRNA to modulate Slco1a4 expression at the NRCM level (Appendix Fig. S3H,I). In the OGD/R model, we observed that Slco1a4 overexpression reversed the anti-apoptotic effects of p16^INK4a^ inhibition, whereas Slco1a4 inhibition counteracted the pro-apoptotic effects of p16^INK4a^ overexpression (Fig. 4H,I; Appendix Fig. S3J,K). Collectively, we identified Slco1a4 as a potential downstream target of p16^INK4a^ during cardiac I/R injury. Reducing Slco1a4 may inhibit cardiomyocyte apoptosis through inhibiting bile acid transport.Figure 4p16^INK4a^ transcriptionally upregulates Slco1a4 to modulate bile acid transport and cardiomyocyte apoptosis.(A) Heat map presenting RNA-sequencing data from p16^INK4a^ cKO mice and control mice. (B) Heat map presenting RNA-sequencing data from Sham mice and mice with 3-weeks of I/R injury. (C) Venn diagram showing RNA-seq data from hearts of p16^INK4a^ cKO mice and control mice (p < 0.05; fold-change>2) were intersected with RNA-seq data from hearts of Sham mice and mice with 3-weeks of I/R injury (p < 0.05; fold-change>1.5). (D) qRT-PCR analysis of Slco1a4 mRNA expression in the hearts of p16^INK4a^ cKO mice and control mice following 3-weeks of I/R injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. *p = 0.0275; Sham: Control vs. I/R (3 W): Control, **p = 0.0017; Sham: p16^INK4a^ cKO vs. I/R (3 W): p16^INK4a^ cKO, **p = 0.0077; I/R (3 W): Control vs. I/R (3 W): p16^INK4a^ cKO, **p = 0.0062. (E) qRT-PCR analysis of Slco1a4 mRNA expression in the hearts of p16^INK4a^ cKO mice and control mice following acute I/R injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Left: Infarct zone, **p = 0.0032; ***p = 0.0002; ****p < 0.0001. Right: Border zone. *p = 0.0348; **p = 0.0082; ****p < 0.0001. (F) ELISA analysis of blie acid concentration in the hearts of p16^INK4a^ cKO mice and control mice following 3-weeks of I/R injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (G) ELISA analysis of blie acid concentration in the hearts of p16^INK4a^ cKO mice and control mice following acute I/R injury (n = 6 per group, biological replicates). Left: Infarct zone; Right: Border zone. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (H) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with sh-p16^INK4a^ and Slco1a4 OE in OGD/R induced apoptosis model (*n *= 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (I) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with p16^INK4a^ OE and si-Slco1a4 in OGD/R induced apoptosis model (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
Bile acids exacerbate OGD/R-induced cardiomyocyte apoptosis and cardiac I/R injury
To further explore the potential role of bile acids in OGD/R-induced cardiomyocyte injury, we added bile acids to NRCMs subjected to OGD/R. While bile acids alone did not induce apoptosis under baseline conditions, they markedly aggravated OGD/R-induced cardiomyocyte apoptosis, as demonstrated by an increased number of TUNEL-positive cells (Fig. 5A), along with elevated Bax/Bcl-2 and Cleaved-caspase3/Caspase3 protein expression ratios (Fig. 5B). These results indicate that bile acids exacerbate OGD/R-induced cardiomyocyte apoptosis.Figure 5. Bile acid exacerbates OGD/R-induced apoptosis in vitro.(A) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with bile acid or vehicle under OGD/R or Control condition (n = 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. ****p < 0.0001. (B) Immunoblotting analysis of NRCMs apoptosis by detection of Bax, Bcl2, caspase-3, and Cleaved-caspase3 in Control or OGD/R-induced apoptosis model treated with bile acid or vehicle (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by the Tukey post hoc test. **p = 0.0014; ***p = 0.0001; ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
To investigate the in vivo effects of bile acids on cardiac I/R injury, mice were administered with bile acid injections every other day for 2 weeks prior to acute I/R surgery (Fig. 6A). Histological analysis revealed no morphological damage to the cardiac tissue induced by bile acids treatment alone (Fig. EV5A). Notably, bile acids quantification confirmed a significant increase in cardiac bile acid levels following administration (Fig. 6B). Compared to vehicle-treated controls, mice treated with bile acids displayed significantly larger myocardial infarct areas (Fig. 6C), accompanied with elevated numbers of TUNEL-positive cells and higher Bax/Bcl-2 and Cleaved-caspase3/Caspase3 protein expression ratios (Figs. 6D and EV5B,C). Moreover, cardiomyocyte proliferation was suppressed in mice treated with bile acids, as evidenced by a decreased number of Ki67-positive nuclei in I/R-injured cardiac tissue (Fig. EV5D). To further evaluate the chronic effects of bile acids post-I/R, mice were administered with bile acids every other day for 1 week prior to I/R surgery, and bile acid treatment continued throughout the 3-week remodeling phase following I/R injury (Fig. 6E). Similar to the acute model, bile acids did not induce structural abnormalities in the heart (Fig. EV5E), but a significant elevation in cardiac bile acid levels was again confirmed (Fig. 6F). Importantly, bile acid-treated mice exhibited worsened cardiac outcomes, including impaired cardiac function, increased myocardial fibrosis, and exacerbated pathological cardiac hypertrophy three weeks after I/R injury (Fig. 6G–J). Collectively, these findings demonstrate that bile acids aggravate both acute and chronic myocardial injury following I/R.Figure 6. Bile acid exacerbates myocardial I/R injury.(A) The schedule of acute myocardial I/R injury model establishment in mice gavaged with bile acid or vehicle control. (B) ELISA analysis of bile acid concentration in the hearts of mice administered bile acid by gavage and control mice following acute I/R injury (n = 8–10 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Left: Infarct zone, **p = 0.0015; ***p = 0.0008; ****p < 0.0001. Right: Border zone. ****p < 0.0001. (C) Representative images of TTC staining and quantification of the area at risk/left ventricular weight (AAR/LV) ratio and the infarct size/area at risk (INF/AAR) ratio in the hearts of mice administered bile acid by gavage and control mice following acute I/R injury (n = 7–8 per group, biological replicates). Scale bar, 2 mm. Statistical analysis was performed by unpaired Student’s t-test. ****p < 0.0001. (D) Representative images of immunofluorescence staining and quantification of the TUNEL positive cardiomyocytes in the hearts of mice administered bile acid by gavage and control mice following acute I/R injury (n = 6 per group, biological replicates). Scale bar: 20 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (E) The schedule of myocardial I/R remodeling injury model establishment in mice gavaged with bile acid or vehicle control. (F) ELISA analysis of bile acid concentration in the hearts of mice administered bile acid by gavage and control mice following I/R remodeling injury (n = 10–13 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (G) Representative echocardiographic images and left ventricular ejection fraction (EF), ventricular fractional shortening (FS) of mice administered bile acid by gavage and control mice following I/R remodeling injury (n = 10–13 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (H) Representative images of Masson’s trichrome staining and quantification of fibrotic area (%) in the hearts of mice administered bile acid by gavage and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Scale bar, 50 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (I) qRT-PCR analysis of Col1a1, Col3a1 and Ctgf expression in the hearts of mice administered bile acid by gavage and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (J) qRT-PCR analysis of Anp, Bnp and β-MHC expression in the hearts of mice administered bile acid by gavage and control mice following I/R remodeling injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ***p = 0.0002; ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
p16INK4a regulates the binding of Npas2 to Slco1a4 promoter region
Through Slco1a4 sequence analysis, we sought to identify promoter regulatory sequences. To achieve this, motif analysis was performed on genomic regions where p16^INK4a^ binds within Slco1a4 promoters using the JASPAR 2020 database. However, no p16^INK4a^ binding sites were identified within the proximal region (upstream 2000nt) of the Slco1a4 DNA promoter. This suggests that p16^INK4a^ may exert its regulatory influence on Slco1a4 indirectly. Further analysis of the Slco1a4 promoter sequence identified binding sites for Arnt1 and Npas2, which exhibit strong binding affinities and p16^INK4a^-responsive regulatory elements, according to transcription factor binding predictions using the JASPAR database and RNA-seq data from p16^INK4a^ cKO mice (fold change>2; p < 0.05) (Fig. 7A,B). Subsequent chromatin immunoprecipitation (ChIP) validation revealed that Arnt1 does not bind to the Slco1a4 promoter, whereas Npas2 binds at both sites (Fig. 7C). The luciferase reporter experiment further confirmed Npas2’s ability to directly bind to and enrich the promoter region of Slco1a4 in NRCMs (Appendix Fig. S4A). Additionally, ChIP results demonstrated that overexpression of p16^INK4a^ reduced the binding of Npas2 to the promoter regions of Slco1a4 (Fig. 7D). Afterwards, qPCR further validated that Npas2 is a negative regulator responsive to p16^INK4a^ in NRCMs, along with the observed decrease in Npas2 expression within the OGD/R cellular model (Appendix Fig. S4B–D). Additionally, qPCR revealed decreased Npas2 expression in myocardial tissue of control mice subjected to cardiac I/R injury. However, p16^INK4a^ cKO mice were able to counteract the reduction in Npas2 expression induced by cardiac I/R injury (Fig. 7E,F). Subsequently, in the OGD/R model, we observed that knockdown of Npas2 attenuated the resistance conferred by sh-p16^INK4a^ against OGD/R-induced cell apoptosis (Fig. 7G; Appendix Fig. S4E,G). Conversely, overexpression of Npas2 alleviated the exacerbating effect of p16^INK4a^ overexpression on OGD/R-induced cell apoptosis (Appendix Figs. S4F,H,I). Finally, qPCR analysis revealed that overexpression of Npas2 in vitro resulted in decreased Slco1a4 expression, while knocking down Npas2 had the opposite effect (Appendix Fig. S5A,B). In the OGD/R model, we also observed that Slco1a4 overexpression reversed the anti-apoptotic effects of Npas2 overexpression, whereas Slco1a4 inhibition counteracted the pro-apoptotic effects of Npas2 inhibition (Fig. 7H; Appendix Fig. S5C–E). These results strongly suggest that p16^INK4a^ modulates the transcription of Slco1a4 by influencing the binding of Npas2 to its promoter region, thereby affecting the myocardial I/R injury process.Figure 7p16^INK4a^ regulates the binding of Npas2 to Slco1a4 promoter region.(A) Venn diagram showing the intersection between Jaspar-identified transcription factor binding to Slco1a4 promoter (site score>10) and RNA-seq data from hearts of p16^INK4a^ cKO mice and control mice (p < 0.05; fold-change > 2). (B) Motif analysis of Arnt1 and Npas2 binding sites in the Slco1a4 promoter. (C) ChIP analysis of Arnt1 and Npas2 enrichment at the Slco1a4 promoter region (n = 6 per group, biological replicates). Statistical analysis was performed by unpaired Student’s *t-*test. **p = 0.001; ***p = 0.0001. (D) ChIP analysis of the enrichment of Npas2 binding to the Slco1a4 promoter region in NRCMs infected with the p16^INK4a^ OE lentivirus (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. *p = 0.0227; ****p < 0.0001. (E) qRT-PCR analysis of Npas2 mRNA expression in the hearts of p16^INK4a^ cKO mice and control mice following 3-weeks of I/R injury (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. *p = 0.0365; **p = 0.0072; ****p < 0.0001. (F) qRT-PCR analysis of Npas2 mRNA expression in the hearts of p16^INK4a^ cKO mice and control mice following acute I/R injury (*n *= 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. Left: Infarct zone (Sham: Control vs. Acute I/R (Infarct): Control, *p = 0.042; Acute I/R (Infarct): Control vs. Acute I/R (Infarct): p16^INK4a^ cKO, *p = 0.0165; ****p < 0.0001. Right: Border zone (Sham: Control vs. Acute I/R (Border): Control, *p = 0.0403; Acute I/R (Border): Control vs. Acute I/R (Border): p16^INK4a^ cKO, *p = 0.0116; ****p < 0.0001). (G) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with sh-p16^INK4a^ and si-Npas2 in OGD/R induced apoptosis model (n = 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. *p = 0.0191; ****p < 0.0001. (H) Representative images of immunofluorescence staining and quantification of the TUNEL positive NRCMs treated with Npas2 OE and Slco1a4 OE in OGD/R induced apoptosis model (*n *= 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
p16INK4a-CDK4 complex regulates Npas2 mRNA stability and translation via CUGBP1 to promote bile acid uptake and OGD/R-induced apoptosis in cardiomyocytes
To further elucidate the mechanism by which p16^INK4a^ regulates the activity of Npas2, we conducted in vitro experiments to modulate the expression of p16^INK4a^ while concurrently treating with Actinomycin D to halt transcription, enabling the assessment of Npas2 mRNA stability under varying levels of p16^INK4a^. Our data revealed that suppressing p16^INK4a^ expression led to increased stability of Npas2 mRNA, whereas overexpression of p16^INK4a^ reduced its stability (Fig. 8A). Analysis of Npas2 mRNA revealed the presence of a GRE motif within its 3’UTR, which can affect mRNA half-life through interactions with RNA-binding proteins (RBPs). CUGBP1, a recognized RBP capable of modulating mRNA degradation in the presence of GRE motifs, was identified (Vlasova et al, 2008). Consequently, we developed a specific biotin-labeled probe targeting Npas2-GRE (Npas2-GRE-Bio) and a negative control mutant probe (mut-Npas2-GRE-Bio) for RNA pulldown assays in cardiomyocytes (Fig. 8B). Our results demonstrate that Npas2-GRE-Bio effectively binds and precipitates CUGBP1, whereas mut-Npas2-GRE-Bio does not. Moreover, the competitive binding of Npas2-GRE probe to CUGBP1 in the presence of modified Npas2-GRE-Bio probe was observed (Fig. 8C). Additional experiments revealed that overexpression of p16^INK4a^ enhances the association between Npas2-GRE-Bio and CUGBP1, whereas sh-p16^INK4a^ attenuates this interaction (Fig. 8D). We further investigated the mechanism by which p16^INK4a^ regulates the interplay between CUGBP1 and Npas2. Recognizing that both p16^INK4a^ and Npas2 are pivotal in cell cycle regulation, we first modulated p16^INK4a^ expression in NRCMs under OGD/R conditions. qPCR analysis demonstrated that p16^INK4a^ overexpression markedly suppressed the expression of cell cycle-related genes, including CDK4, CDK6, Cyclin D1, Cyclin D2, Cyclin D3, Cyclin E2, Cyclin A2, and PCNA, while p16^INK4a^ inhibition yielded the opposite effects (Appendix Fig. S6). Given that p16^INK4a^ is known to bind and inhibit CDK4/6 activity (Li et al, 1994; Sandhu et al, 2000), we validated this interaction through co-immunoprecipitation (Co-IP), while also revealing that p16^INK4a^ does not directly interact with CUGBP1 (Fig. 8E), Therefore, we hypothesize that p16^INK4a^ regulates the binding of CUGBP1 to the GRE motif on Npas2 mRNA through its interaction with CDK4 or CDK6, thereby modulating the stability of Npas2 mRNA.
To test this hypothesis, we performed Actinomycin D chase assays to evaluate Npas2 mRNA stability. The results showed that p16^INK4a^ overexpression significantly decreased the half-life of Npas2 mRNA, whereas co-overexpression with CDK4, but not CDK6, restored its stability (Fig. 8F). Furthermore, polysome profiling revealed that p16^INK4a^ overexpression reduced the proportion of Npas2 mRNA in polysome fractions, with a corresponding increase in monosome-associated fractions, and this effect was reversed when CDK4 was co-expressed, suggesting that p16^INK4a^ decreases Npas2 translation efficiency through CDK4 inhibition (Fig. 8G). Western blot analysis confirmed that p16^INK4a^ overexpression reduced Npas2 protein levels, whereas CDK4 co-expression rescued its expression (Fig. 8H).
Functionally, based on CCK8 assays, suitable concentrations of cholic acid, chenodeoxycholic acid, and taurocholic acid were chosen for subsequent experiments (Appendix Fig. S7A). Bile acid uptake assays demonstrated that reduced Npas2 expression promoted bile acid accumulation in NRCMs under both basal and OGD/R conditions (Fig. 8I). Consistently, apoptosis analyses confirmed that decreased Npas2 expression enhanced cardiomyocyte apoptosis, as evidenced by increased TUNEL-positive cells and elevated Bax/Bcl-2 and Cleaved-caspase3/Caspase3 ratios, whereas restoration of Npas2 expression by CDK4 co-expression markedly attenuated these effects (Fig. 8J; Appendix Fig. S7B). Together, these findings indicate that the p16^INK4a^-CDK4 complex regulates Npas2 mRNA stability and translation via CUGBP1, thereby promoting bile acid uptake and OGD/R-induced apoptosis in cardiomyocytes.Figure 8p16^INK4a^-CDK4 complex regulates Npas2 mRNA stability and translation via CUGBP1 to promote bile acid uptake and OGD/R-induced apoptosis in cardiomyocytes.(A) qRT-PCR analysis of Npas2 mRNA stability in NRCMs infected with sh-p16^INK4a^ or p16^INK4a^ OE lentivirus under treatment of Actinomycin D (*n *= 6 per group/per time point, biological replicates). Statistical analysis was performed by unpaired Student’s t-test for (A) (NRCMs infected with shScr and sh-p16^INK4a^: 0 h, 4 h (****p < 0.0001), 6 h (**p = 0.003), 8 h (****p < 0.0001), and 10 h (***p = 0.0003); NRCMs infected with Fugw and p16^INK4a^ OE: 0 h, 2 h (**p = 0.007), 6 h (**p = 0.006), 8 h (***p = 0.0005) and10h (**p = 0.009)), by Mann-Whitney U test for (A) (NRCMs infected with shScr and sh-p16^INK4a^: 2 h (**p = 0.0022); NRCMs infected with Fugw and p16^INK4a^ OE: 4 h (*p = 0.0152)). (B) Representative segments of GRE motifs within the Npas2 3’UTR and sequences of Npas2-GRE-Bio and mutated Npas2-GRE-Bio were used for the binding reactions. (C) mRNA pull-down assay using Npas2-GRE-Bio or mut-Npas2-GRE-Bio with cold Npas2-GRE probe competing for binding to CUGBP1 (n = 3 per group, independent experiments). (D) mRNA pull down assay was performed by mixing Npas2-GRE-Bio or mut-Npas2-GRE-Bio with p16^INK4a^ OE or sh-p16^INK4a^ lentivirus-treated NRCMs (n = 3 per group, independent experiments). (E) Co-IP analysis of the binding status of p16^INK4a^ with CDK4, CDK6 and CUGBP1 (n = 3 per group, independent experiments). (F) qRT-PCR analysis of Npas2 mRNA stability in NRCMs infected with p16^INK4a^ OE, or combined p16^INK4a^ and CDK4 OE, or combined p16^INK4a^ and CDK6 OE lentivirus under treatment with Actinomycin D (n = 6 per group/per time point, biological replicates). Statistical analysis was performed by one-way ANOVA followed by Bonferroni post hoc test. ****p < 0.0001. (G) RT-qPCR analysis of Npas2 mRNA expression in polyribosome and monoribosome fractions of NRCMs infected with p16^INK4a^ OE, or combined p16^INK4a^ and CDK4 OE lentivirus. (H) Immunoblotting analysis of Npas2 protein expression in NRCMs treated with p16^INK4a^ OE, or combined p16^INK4a^ and CDK4 OE lentivirus (n = 4 per group, biological replicates). Statistical analysis was performed by one-way ANOVA followed by Dunnett T3 post hoc test. *p = 0.028; **p = 0.006. (I) ELISA analysis of bile acid concentration in NRCMs treated with p16^INK4a^ OE, or combined p16^INK4a^ and CDK4 OE lentivirus (n = 6 per group, biological replicates). Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test. ****p < 0.0001. (J) Representative images of immunofluorescence staining and quantification of the TUNEL positive cardiomyocytes in NRCMs treated with p16^INK4a^ OE, or combined p16^INK4a^ and CDK4 OE lentivirus (*n *= 6 per group, biological replicates). Scale bar: 50 μm. Statistical analysis was performed by two-way ANOVA test followed by Tukey post hoc test, ****p < 0.0001. Data represent mean ± SD. Source data are available online for this figure.
Discussion
In silico analyses of clinical databases cross-referenced with gene expression studies implicated p16^INK4a^ as one of a small number of genes associated with ischemic injury. p16^INK4a^ levels were elevated in mouse and canine hearts following cardiac I/R, as well as in NRCMs and hESC-CMs subjected to OGD/R. Similarly, increased p16^INK4a^ expression was also observed in NRCFs upon TGF-β stimulation. Importantly, the elevation of p16^INK4a^ was also confirmed in human ischemic cardiomyopathy, as well as in human cardiomyocytes and fibroblasts after myocardial infarction. Functionally, p16^INK4a^ overexpression exacerbated apoptosis in NRCMs and hESC-CMs, whereas its knockdown reduced OGD/R-induced NRCMs and hESC-CMs apoptosis in vitro. In contrast, p16^INK4a^ had no significant effect on fibrotic activation in TGF-β-stimulated NRCFs, indicating that the functional role of p16^INK4a^ is confined to cardiomyocytes rather than cardiac fibroblasts. Consistently, in vivo knockdown of p16^INK4a^ reduced infarct size and enhanced cardiac function after I/R injury, whereas its overexpression exacerbated cardiac damage.
While our findings confirm the protective role of p16^INK4a^ knockdown in myocardial I/R injury, the clinical implications extend beyond the scope of this study. The observation that p16^INK4a^ exerts protective effects upon knockdown but promotes injury when overexpressed supports the rationale for developing precision-targeted therapeutic strategies. This could potentially be translated into clinical strategies where p16^INK4a^ modulation serves as a novel approach to mitigate the impacts of I/R injury. We further demonstrate that p16^INK4a^, in conjunction with CDK4, regulates CUGBP1 to reduce the stability and translation of Npas2 mRNA, thereby enhancing the transcription of Slco1a4 and bile acid transport. This study underscores the protective role of p16^INK4a^ knockdown against myocardial I/R injury and expands our understanding of its regulatory mechanisms in cardiovascular diseases.
I/R injury involves oxidative stress, cellular apoptosis, and mitochondrial dysfunction (Ferdinandy et al, 2014; Logue et al, 2005; Wang et al, 2018). The Cdkn2a gene, encoding tumor suppressors p16^INK4a^, p19^ARF^ and p14^ARF^, exhibits anti-angiogenic properties (Kawagishi et al, 2010; Takeuchi et al, 2004) and downregulates vascular endothelial growth factor (VEGF) in various cancers (Gibson et al, 2003; Harada et al, 1999). Additionally, Cdkn2a regulates genes involved in oxidative stress and apoptosis (Che et al, 2022; Javaheri et al, 2016). As an inhibitor of CDK4/6, p16^INK4a^ plays a role in cellular aging and may mitigate aging effects (Liu et al, 2023). Upregulation of p16^INK4a^ is linked to aging in human cardiac progenitor cells (hCPCs), where its downregulation enhances proliferation, survival, and antioxidative gene expression (Khatiwala et al, 2018). Furthermore, p16^INK4a^ is implicated in myocardial regeneration through CDK4/6 and autophagy pathways, with its downregulation promoting cell proliferation (Sun et al, 2023). Here we find that p16^INK4a^, a central mediator of cellular senescence that increases with age (Baker et al, 2016; Liu et al, 2009), plays an important role in modulating the initial and chronic response to I/R, suggesting it could provide just such a link between aging and cardiac dysfunction after I/R. We noted consistent p16^INK4a^ upregulation in acute myocardial I/R injury, I/R remodeling, OGD/R-induced apoptosis, and TGF-β-induced fibrotic activation in cardiac fibroblasts. Inhibition of p16^INK4a^ in cardiomyocytes decreased apoptosis, while its overexpression increased it, with no detectable effect in cardiac fibroblasts. Cardiomyocyte-specific p16^INK4a^ knockout mice showed resistance to acute I/R injury, reduced myocardial apoptosis, and mitigated I/R remodeling effects. These findings suggest that p16^INK4a^ plays a significant role in myocardial I/R injury. Notably, while TTC staining and LDH levels primarily serve as indicators of myocardial necrosis, they also offer insights into the broader spectrum of cellular damage, including apoptosis. This highlights the critical need for in-depth exploration into the regulatory mechanisms of p16^INK4a^ in mediating cellular necrosis during myocardial I/R injury. Furthermore, oxidative stress and impaired angiogenesis also contribute to I/R injury, warranting further research into p16^INK4a^’s multifaceted role (Shi et al, 2017). Moreover, besides cardiomyocytes and fibroblasts, there is a suggestion to investigate the role of p16^INK4a^ in other cell types, such as endothelial cells and immune cells, during cardiac I/R injury, although it is beyond the scope of this study.
Through transcriptomic analysis, we identified Slco1a4 as a potential p16^INK4a^ target. Slco1a4 upregulation occurs in I/R-injured mouse hearts and correlates with p16^INK4a^ levels. The protective effect of p16^INK4a^ knockdown against OGD/R-induced apoptosis was abrogated by Slco1a4 overexpression, while knocking down Slco1a4 mitigated the negative effects of p16^INK4a^ overexpression. Slco1a4’s role in cardiovascular diseases is yet to be fully explored (Qi and Ma, 2017). Slco1a4 is a member of the solute carrier organic anion transporter family involved in the transport of bile acids, conjugated steroids, and prostaglandins (Granados and Nigam, 2024; Shan et al, 2023; Zhang et al, 2013). Emerging evidence suggests that the bile acid receptor farnesoid-X-receptor is a novel apoptosis mediator and contributes to myocardial ischemia/reperfusion injury, potentially indicating a link between bile acids and myocardial damage (Pu et al, 2013). Conversely, other studies have demonstrated that mesenchymal stromal cells overexpressing farnesoid X receptor exerted cardioprotective effects against acute ischemic heart injury by binding endogenous bile acids (Xia et al, 2022). In addition, lower circulating levels of dehydroepiandrosterone sulfate (DHEAS), a major conjugated steroid, have been associated with increased risk of heart failure and all-cause mortality in older adults, suggesting a potential role in cardiovascular health (Jia et al, 2020). Among prostaglandins, prostaglandin E2 (PGE2) has been shown to exert cardioprotective effects in the setting of myocardial ischemia/reperfusion (I/R) injury, primarily through activation of the EP4 receptor pathway (Xiao et al, 2004). However, other studies have indicated that following myocardial infarction, elevated PGE2 levels may not only regulate local inflammatory responses but also contribute to remote myocardial remodeling during the chronic phase of injury, potentially impairing long-term cardiac function (Sun and Wang, 2012).
Our study reveals that p16^INK4a^ does not directly bind to the promoter of Slco1a4; instead, it modulates the indirect binding of Npas2 to the promoter region of Slco1a4, thereby influencing the transcription of Slco1a4 and bile acid transport, while having no direct impact on the regulation of DHEAS and PGE2. Furthermore, we found that bile acids exacerbate OGD/R-induced cardiomyocyte apoptosis in vitro and worsen myocardial injury in vivo following I/R. This discovery provides evidence that the regulatory role of p16^INK4^ on Slco1a4 is mediated through Npas2. Thus, we provide evidence that p16^INK4a^ indirectly enhances Slco1a4 transcription through a reduction of the combined Npas2 to the promoter region of Slco14.
Some limitations should be highlighted. Due to limited availability, we did not perform studies in aged mice, but given the well-documented increase in p16^INK4^ with aging, it is plausible to infer that it contributes to the increased dysfunction and adverse outcomes seen with aging in I/R injury. Our study findings suggest that its upregulation in myocardial cells may lead to increased apoptosis. Therefore, we posit that the upregulation of p16^INK4^ may be a contributing factor to the heightened susceptibility of aging myocardial cells to apoptosis following I/R-injured myocardial injury. However, further research is required to elucidate the exact relationship between p16^INK4^ and apoptosis in aging myocardium, as well as other potential mechanistic implications. This will aid in gaining a more comprehensive understanding of the impact of aging on cellular survival following myocardial injury. Besides, to identify the downstream targets of p16^INK4a^, we performed comparisons between control and p16^INK4a^ cardiac-specific knockout (cKO) hearts under basal conditions, as well as between these groups following ischemia/reperfusion (I/R) injury relative to sham controls. It would be interesting to compare wild-type and p16^INK4a^ cKO hearts after I/R injury in future studies. Moreover, in this study, we only used p16^INK4a flox/flox^ mice carrying loxP sites but lacking Cre expression as control animals, it will be better to add α-MHC-Cre mice as the other control in the future study. Our study also does not fully explore the broader implications of p16^INK4a^’s interactions within the cell, nor does it address the potential compensatory mechanisms that may be at play in the absence of p16^INK4a^. Future research should aim to fill these gaps, offering a more comprehensive understanding of p16^INK4a^’s role in cardiac pathology. Also, further exploration should clarify how p16^INK4a^ modulates Npas2. Distinct from existing research, our work reveals that p16^INK4a^’s regulation of myocardial injury involves CDK4-mediated control of Npas2 stability. This novel mechanism could reveal new targets for mitigating I/R injury and improving cardiac repair. p16^INK4a^’s interaction with CDK4/6 impacts cell proliferation and tumor suppression (Romagosa et al, 2011). For example, p16^INK4a^ facilitates cervical cancer cell proliferation via the CDK6-HuR-IL1A pathway, affecting IL1A mRNA stability (Li et al, 2020). RNA-binding proteins like HuR and CUGBP1 modulate mRNA stability (Dasgupta and Ladd, 2012; DeMaria and Brewer, 1996; Ladd and Cooper, 2004; Nadar et al, 2011; Philips et al, 1998; Timchenko et al, 1996; Yeh et al, 2008). The elevated expression of CUGBP1 has been associated with cardiac pathologies induced by type 1 myotonic dystrophy (DM1) (Misra et al, 2020; Sovari et al, 2007; Wang et al, 2007). However, CUGBP1 has been shown to protect cardiomyocytes from ischemia-induced injury by promoting angiogenesis and inhibiting apoptosis through the regulation of the VEGF-A gene (Gu et al, 2017). Our findings suggest that p16^INK4a^ interacts with CDK4 to regulate CUGBP1, which in turn reduces Npas2 mRNA stability and translation. This informs potential post-transcriptional regulation by p16^INK4a^ during ischemia/reperfusion injury. Therefore, further investigations should ascertain if other elements also regulate Npas2 stability via CUGBP1. Additionally, we further found that NPAS2 binds to the Slco1a4 promoter and represses its transcription. Although NPAS2 is traditionally recognized as a transcriptional activator within the bHLH-PAS family through its interaction with BMAL1 (also known as ARNTL), it has also been reported to function as a transcriptional repressor. For instance, BMAL1/NPAS2 heterodimers suppress c-Myc transcription by binding to E-box elements in the c-Myc P1 promoter (Fu et al, 2002; Peng et al, 2021). Moreover, several bHLH-PAS members, including IPAS and DEC1/DEC2, have been shown to exert transcriptional repression through noncanonical mechanisms, such as competitive binding to partner proteins or E-box elements (Honma et al, 2002; Makino et al, 2001). Based on this, we propose that NPAS2 may repress Slco1a4 via similar noncanonical pathways, such as competing for E-box occupancy, forming low-activity complexes, or recruiting corepressors under pathological conditions. These findings suggest a context-dependent and versatile role of NPAS2 in transcriptional regulation. While our study results show promising potential, the translation of this research pathway into clinical applications for humans warrants further exploration. For instance, we may consider utilizing genome editing techniques to modulate the p16^INK4a^/Npas2/Slco1a4 axis to validate its potential therapeutic efficacy in human cardiovascular diseases.
In summary, our study demonstrates that in myocardial ischemia-reperfusion injury, p16^INK4a^, in conjunction with CDK4, regulates CUGBP1 to reduce the mRNA stability, translation, and expression of Npas2, which in turn induces Slco1a4 transcription and enhances bile acid transport. These findings not only elucidate key molecular mechanisms of myocardial ischemia-reperfusion injury but also identify promising targets for the development of novel therapeutic strategies to improve cardiac function and clinical outcomes.
Methods
Reagents and tools tableReagent/resourceReference or sourceIdentifier or catalog number Experimental models Mouse: C57BL/6JCavens Laboratory AnimalN/AMouse: α-MHC CreFrom Professor Anthony Rosenzweig, Harvard Medical SchoolN/AMouse: p16^INK4a loxp/loxp^From Professor Anthony Rosenzweig, Harvard Medical SchoolN/AMouse: α-MHC Cre p16^INK4a loxp/loxp^This paperN/ANRCMsThis paperN/AhESC cellFrom Professor Guoping Li, School of Medicine, Tongji University.N/AhESC-CMsThis paperN/ANRCFsThis paperN/AH9C2 cellAmerican Type Culture CollectionCRL-1446HEK293TAmerican Type Culture CollectionCRL-3216 Recombinant DNA ScrambleAddgene14883pLKO.1Addgene8453pLKO.1-sh-p16^INK4a^This paperN/APlasmid: FugwAddgene8453Fugw-p16^INK4a^ OEThis paperN/AFugw-Npas2 OEThis paperN/AFugw-slco1a4 OEThis paperN/AFugw-CDK4 OEThis paperN/AFugw-CDK6 OEThis paperN/ApMD2.GAddgene12259psPAX2Addgene12260 Antibodies p16INK4a Monoclonal Antibody (4A10E1)Thermo FisherMA5-38494p16 INK4A Rabbit pAbABclonalA23882CDKN2A/p19ARF Rabbit mAbABclonalA26879Anti-CDKN2A/p14ARF antibodyAbcamab185620Bax Rabbit pAbABclonalA12009Bcl-2 Rabbit pAbABclonalA0208[KO Validated] Caspase-3 Rabbit pAbABclonalA2156PCNA Rabbit mAbABclonalA12427Cyclin D1 Rabbit pAbABclonalA1301CUGBP1 Polyclonal antibodyProteintech13002-1-APCDK4 Rabbit pAbABclonalA0366CDK6 Rabbit mAbABclonalA0106NPAS2Abcamab55833Arnt 1(A-3)Santa Cruzsc-17811β-actin Rabbit mAbABclonalAC038α-Actin antibodySigmaA7811Ki67Abcamab15580Cy3-AffiniPure rabbit anti-mouse immunoglobulin G (IgG) (H + L)Jackson ImmunoResearch715-165-151Cy3-conjugated monoclonal anti-SMASigmaC6198Anti-Rabbit IgG (Fc specific) antibody produced in goatSigmaSAB3700848 Oligonucleotides and sequence-based reagents Primers sequences for qPCRThis paperAppendix Table S1Target sequences of siRNAsThis paperAppendix Table S2ShRNA sequence of p16^INK4a^This paperAppendix Table S3Primers sequences for overexpression plasmid constructionThis paperAppendix Table S4Primer sequences for ChIPThis paperAppendix Table S5 Chemicals, enzymes and other reagents DMEMCorning10-013-CVFetal Bovine SerumBiological Industries04-001-1ACSPenicillin and StreptomycinKeyGENKGY0023Horse SerumGibco16050-122Lipofectamine® 2000Invitrogen11668-019Anaero packsMGCC-1Recombinant Human TGF-beta1 (CHO derived)Peprotech100-21 CRNAiso PlusTaKaRa9109TB Green Premix Ex Taq II kitTakaraRR820ARIPA lysis bufferKenGENKGP-701-100Polyvinylidene Fluoride membrane (PVDF)PALLBSP0161FormaldehydeSCIENTZSCIENTZ08-IIIProtein G Dynabeads™Thermo Fisher Scientific1004DActinomycin DSigma-AldrichSBR00013Biotin RNA Labeling MixSigma-Aldrich11685597910Dynabeads^TM^ MyOne^TM^ Streptavidin T1Invitrogen65604DCell lysis bufferSigma-AldrichC2978Protease Inhibitor Cocktail (100×)Thermo Fisher Scientific1862209HochestKeyGEN33342RevertAid First Strand cDNA Synthesis KitThermo scientificK1622Pierce™ BCA Protein Assay KitTaKaRaT9300AHigh-sig ECLTanon180-5001Cholic Acid (CA)Sigma-AldrichC1129Chenodeoxycholic Acid (CDCA)Sigma-AldrichC9377Taurocholic Acid (TCA)Sigma-AldrichT4009CMC-NaTargetMolT19232Tris-HCl (pH 7.5)Sigma-AldrichT2319KClSigma-AldrichP3911MgCl₂Sigma-AldrichM8266Nonidet P-40 SubstituteThermo Fisher85124DithiothreitolSigma-AldrichD9779CycloheximideSigma-AldrichC4859RNase InhibitorThermo FisherAM2694cOmplete™ Mini EDTA-free Protease Inhibitor CocktailRoche11836170001Total Bile Acid (TBA) Content Assay KitEnzyme-linked BioM122CL96DHEAS ELISA kitEnzyme-linked Bioml104863PGE2 ELISA kitEnzyme-linked Bioml037542Dual-LumiTM Luciferase Reporter Gene Assay KitBeyotimeRG088MSimple ChIP Enzymatic Chromatin IP KitCell Signaling Technology9003TUNEL FITC Apoptosis Detection KitVazymeA111-03Dead End Fluorometric TUNEL SystemPromegaG3250Cell Counting Kit-8BeyotimeC0038Masson’s Trichrome stainingServicebioG1006EdU staining kitKeyGENKGA9602-500hematoxylin and eosin (HE) solutionsKeyGENKGE1204-50 Software GraphPad Prism 8.0GraphPad https://www.graphpad.com/ SPSS Statistics 20IBM https://www.graphpad.com/ ImageJFIJI https://imagej.net/software/fiji/
Other Vevo 2100 Imaging System deviceVisualsonics, Inc.LightCycler 480RocheOlympus FLUOVIEW FV3000 confocal laser scanning microscopeOlympusSpectraMax iD3Molecular Devices, LLC
Animal treatment
All animal experiments complied with the Guidelines on the Use and Care of Laboratory Animals for biomedical research, as published by the National Institutes of Health (No. 85-23, revised 1996), and the protocol was approved by the Ethics Committees of Shanghai University (NO.2022212). Mice were housed in a specific pathogen-free (SPF) facility under controlled conditions (22 ± 2 °C, 50–60% relative humidity, 12-h light/dark cycle) with free access to standard chow and water. Canine ischemia-reperfusion injury model samples were obtained from a study conducted by our group (Wang et al, 2021), with all procedures adhering to the Institutional Animal Care Guidelines and receiving ethical approval. The human data utilized in this study were derived from publicly accessible datasets and represent a secondary analysis of these resources. The original studies complied with the ethical principles outlined in the Declaration of Helsinki, obtained the necessary institutional ethical approvals, and documented informed consent from all participants as specified in their respective publications.
The role of p16^INK4a^ in cardiac ischemia/reperfusion (I/R) injury was investigated using both loss-of-function and gain-of-function models. For loss-of-function studies, cardiac-specific p16^INK4a^ conditional knockout (p16^INK4a^ cKO) mice were generated by crossing floxed p16^INK4a^ (p16^INK4a flox/flox^) mice with transgenic mice expressing Cre recombinase under the control of the alpha-myosin heavy chain (α-MHC) promoter. Upon reaching adulthood (8–9 weeks), these mice underwent cardiac I/R injury induced by left anterior descending artery (LAD) ligation for 30 min, followed by either 24 h of reperfusion (acute I/R injury) or 3 weeks of reperfusion (I/R remodeling). For gain-of-function studies, cardiac-specific overexpression of p16^INK4a^ was achieved via adeno-associated virus serotype 9 (AAV9). Male C57BL/6 J mice (8–9 weeks old) received a single intravenous tail vein injection of either AAV9 carrying p16^INK4a^ under the cardiac troponin T (cTNT) promoter (10¹² vg/mL) or a control virus (10¹² vg/mL), one week before LAD ligation. As previously described (Bei et al, 2019; Gao et al, 2021), surgeries were performed under 2.0% isoflurane anesthesia with mechanical ventilation to ensure passive respiration. Following thoracotomy, the LAD was ligated with 7-0 silk for 30 min, after which reperfusion was initiated before the thoracic cavity was closed. Sham-operated mice underwent the same procedure without actual LAD ligation.
To assess the effect of bile acids on myocardial injury, mice were orally gavaged with a mixture of cholic acid (CA, 50 mg/kg), chenodeoxycholic acid (CDCA, 30 mg/kg), and taurocholic acid (TCA, 10 mg/kg), dissolved in 0.5% sodium carboxymethyl cellulose (CMC-Na). For the acute I/R injury model, bile acids administration began 2 weeks prior to LAD ligation and continued every other day until the day before surgery. For the I/R remodeling model, mice received bile acids gavage starting 1 week before surgery and continuing throughout the 3-week reperfusion period.
For the evaluation of acute I/R injury, mice were anesthetized with 2.0% isoflurane, and 1 mL of 1% Evans Blue dye was injected into the left ventricle (LV). The hearts were excised, sectioned into 1-mm-thick slices, stained with 1% 2,3,5-triphenyltetrazolium chloride (TTC) solution in a 37 °C water bath, and fixed in 4% paraformaldehyde before imaging. For the assessment of I/R remodeling, cardiac ultrasound was performed. Finally, cardiac tissue was collected following euthanasia via intraperitoneal injection of sodium pentobarbital (60 mg/kg).
H9 human embryonic stem cell (hESC) differentiation and culture
H9 human embryonic stem cell lines (WiCell) were cultured on Matrigel-coated cell culture plates and maintained in E8 medium at 37 °C with 5% CO_2_. When the cells reached 80% confluency, they were induced to differentiate using a differentiation medium composed of RPMI 1640 (Life Technologies, 72400047), 0.5 mg/ml recombinant human albumin and 0.2 mg/ml ascorbic acid phosphate. After differentiation, the cells were screened with glucose-deprived RPMI 1640 medium supplemented with DL-lactic acid (Sigma, L3263), followed by culturing with glucose-containing RPMI 1640 medium supplemented with B27 serum substitute (Thermo Fisher, 17504-044) for 30–60 days to obtain mature cardiomyocytes derived from hESC (hESC-CMs).
Primary cardiomyocyte and fibroblast isolation and culture
Primary neonatal rat cardiomyocytes (NRCMs) and cardiac fibroblasts (NRCFs) were isolated from 1- to 3-day-old Sprague-Dawley rats as described previously (Bei et al, 2022; Bei et al, 2024). Briefly, ventricles were enzymatically digested using Collagenase II (Gibco, 17101015) and pancreatin (Sigma, 8049-47-6). Following differential adhesion to enrich fibroblasts, cardiomyocytes were further purified via Percoll (GE Healthcare, 17-0891-01) density gradient centrifugation. NRCMs were cultured in high-glucose DMEM (Corning, 10-013-CVR) supplemented with 10% horse serum (Gibco, 16050-122) and 5% fetal bovine serum (Biological Industries, 04-001-1ACS), while NRCFs were maintained in DMEM containing 10% horse serum.
Oxygen–glucose deprivation and reperfusion (OGD/R) model
To induce oxygen and glucose deprivation, DMEM medium without glucose was added to neonatal rat cardiomyocytes (NRCMs) before placing them into an anaerobic chamber containing Anaero packs (MGC, C-1) at 37 °C containing 1% CO_2_. After 8 h, NRCMs were removed from the anaerobic chamber and subjected to reoxygenation, followed by the replacement of the medium with DMEM containing glucose for 12 h. hESC-CMs were subjected to 16 h of hypoxia, followed by 12 h of reoxygenation.
Quantitative real-time polymerase chain reactions (qRT-PCRs)
Based on the manufacturer’s protocol, total RNAs were extracted from cells and tissues with RNAiso Plus (TaKaRa, 9109), and a reverse transcription on 400 ng of RNA using RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific, K1622) was performed. The cDNA products were quantified using Real-Time PCR Detection System (Roche LightCycler480) with TB Green Premix Ex Taq II kit (Takara, RR820A). By utilizing the 2^−ΔΔCT^ method, the relative mRNA expression was calculated and normalized to 18S ribosomal RNA. The primer sequences are listed in Appendix Table 1.
Immunoblotting
The total protein was extracted using RIPA lysis buffer (KenGEN, KGP-701-100), followed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Separated proteins were transferred to PVDF (PALL, BSP0161) and incubated with specific primary antibodies at 4 °C overnight. After that, proteins were incubated with goat anti-rabbit or mouse secondary antibodies for 2 h at room temperature. Immunopositive bands were visualized by High-sig ECL (Tanon, 180-5001) using ChemiDoc Imaging System (Bio-Rad, 17001402). The density of bands was normalized to β-actin and quantified using Image J.
CCK-8 assay
The cytotoxicity of bile acids toward NRCMs was evaluated using the Cell Counting Kit-8 (CCK-8; Beyotime, C0038) according to the manufacturer’s protocol. NRCMs were treated with different concentrations of cholic acid (CA), chenodeoxycholic acid (CDCA), and taurocholic acid (TCA) for 12 h, and cell viability was then determined by measuring the absorbance at 450 nm with a microplate reader. Relative cell viability was expressed as a percentage of the vehicle control group.
Cardiomyocyte transfection, lentiviral infection and bile acid treatment
When cultured cells reached 70% confluency, 100 nM of target siRNA was transfected using Lipofectamine® 2000 (Invitrogen, 11668-019). Medium was changed 8 h after transfection and the assays performed at 48 h post-transfection. For lentiviral infection, lentiviral particles were firstly produced by transfecting HEK293T cells with the plasmid and lentiviral packaging vectors pSPAX2 and pMD2.G. Subsequently, NRCMs were infected with lentivirus (3 multiplicity of infection), after 8 h incubation, the medium was replaced and cultures were maintained for another 48 h. The effects of gene interference were verified using qRT-PCR and immunoblotting. The sequences for siRNAs were listed in Appendix Table 2. The sequences for shRNAs were listed in Appendix Table 3. The primers used to construct overexpression plasmid were listed in Appendix Table 4.
For bile acid treatment, NRCMs were pre-incubated with a mixture of cholic acid (CA), chenodeoxycholic acid (CDCA), and taurocholic acid (TCA), each at a final concentration of 10 μM, for 12 h prior to being subjected to OGD/R.
Adenoviral construction and preparation
In animal study, for overexpression of p16^INK4a^ in vivo, overexpression p16^INK4a^ adeno-associated virus 9 (AAV9) carrying a cTNT-promoter (10^12^ vg/mL) or control adeno-associated virus (10^12^ vg/mL) were established and virus packaging were completed by Shanghai Hanbio Biotechnology Co.,Ltd according to standard procedures. AAV9 was injected intravenously at a dose of 3 × 10^12^ viral particles.
ELISA
Heart tissue samples were collected, and each sample was accurately weighed to the same mass of 20 mg. The concentration of bile acids in the heart tissue was determined and statistically analyzed according to the instructions of the Total Bile Acid (TBA) Content Assay Kit (Enzyme Cycling Colorimetric Method) (Enzyme-linked Bio, M122CL96). Levels of DHEAS and PGE2 were measured using the DHEAS ELISA kit (Enzyme-linked Bio, ml104863) and the PGE2 ELISA kit (Enzyme-linked Bio, ml037542), respectively.
For detecting bile acid uptake in NRCMs, cells were incubated with a mixture of cholic acid (CA), chenodeoxycholic acid (CDCA), and taurocholic acid (TCA) at a final concentration of 10 μM each. After incubation, cells were subjected to subsequent treatments, and bile acid uptake was quantified using the TBA assay kit as described above.
Luciferase reporter assay
For the analysis of the regulation between Npas2 and Slco1a4, the promoter sequence of Slco1a4 was cloned into the PGL3-basic vector. NRCMs were transfected with this construct using Lipofectamine 2000 according to the manufacturer’s instructions. After 48 h of transfection, the cells were lysed, and luciferase activity was measured using a dual-luciferase reporter assay system. The firefly luciferase activity was normalized to Renilla luciferase activity to account for transfection efficiency.
Chromatin immunoprecipitation (ChIP) assay
The ChIP assay in H9C2 cells was performed using the SimpleChIP Enzymatic Chromatin IP Kit (CST, 9003) according to manufacturer’s instructions. Briefly, 4 × 10^6^ cells were cross-linked with 1% formaldehyde and brief sonication (SCIENTZ, SCIENTZ08-III) were added after enzyme digestion. Then, 10% supernatant as the input control and the remaining was divided equally into specific antibodies (Anti-NPAS2, Abcam, ab55833; Arnt 1(A-3), Santa Cruz, sc-17811) and negative control rabbit IgG (Sigma, SAB3700848). Following this, the immunocomplexes were precipitated with protein agarose beads (Thermo Fisher Scientific, 10004D). After immunoprecipitation, DNA was eluted and reversed crosslink and then subjected to PCR analysis. All primer sequences were listed in Appendix Table 5.
RNA stability assay
Actinomycin D (ActD, Sigma-Aldrich, SBR00013) treatment assay was performed for detecting Npas2 mRNA stability. NRCMs were subjected to indicated treatments. After treatment, cells were treated with 5 μg/mL Actinomycin D and collected at different time points upon Actinomycin D treatment. Quantitative RT-qPCR was used to detect the relative expression level of Npas2 mRNA at various time points.
RNA pulldown
The synthetic single-strand RNA sequences of Npas2 GRE and its mutated counterpart were procured from Tsingke. Biotinylated RNA for the pulldown assay was labeled using in vitro transcription with Biotin RNA Labeling Mix (Sigma-Aldrich, 11685597910). The Biotin-RNA pulldown assay was conducted as follows: Cells were lysed in buffer and incubated on ice for 10 min. After centrifugation of the cell lysates (12,000 × g, 4 °C, 10 min), 100 μL of the supernatant was collected as the input, while the remaining portion was incubated along with either 10 µg of Biotin-labeled Npas2 GRE probe or mutated Npas2 GRE probe, or 500 nM of Npas2 GRE. The mixture was incubated for 2 h at room temperature. Subsequently, 100 µL of DynabeadsTM MyOneTM Streptavidin T1 (Invitrogen, #65604D) were added and mixed on a rotor for 1 h at room temperature. After washing the beads five times with wash buffer, SDS-PAGE was performed to analyze the binding of CUGBP1 to the RNA probes.
Co-immunoprecipitation
Cells were harvested and lysed in cell lysis buffer (Sigma-Aldrich, C2978) with protease inhibitor cocktail (100×) (Thermo Fisher Scientific, 1862209) for 2 h. After centrifugation (30 min, 12,000 rpm, 4 °C), 1% supernatant was collected as input and the remaining lysate was incubated with indicated antibodies or normal IgG at 4 °C overnight with rotation. The following day, Protein G Dynabeads™ (Thermo Fisher Scientific, 10004D) were added and incubated for 2 h at 4 °C. After collecting by centrifugation, protein complexes were washed three times with lysis buffer on the magnetic rack (ThermoFisher DynaMag-2, 75004061). Protein complexes were dissociated with 2 × SDS loading buffer at 95 °C for 10 min followed by immunoblotting.
Cardiomyocytes apoptosis and proliferation staining
Cardiomyocytes or frozen heart tissue sections were fixed in 4% PFA at room temperature for 20 min, permeabilized with 0.5% Triton X-100 for 20 min, blocked with 5% BSA at room temperature for 1 h to prevent non-specific binding, and incubated overnight with primary α-Actin antibody (1:200, Sigma, A7811) at 4 °C. Following incubation with Cy3 AffiniPure Donkey Anti-Mouse IgG (H + L) (1:200, Jackson, 715-165-151) for 2 h. Cardiomyocytes apoptosis were evaluated using TUNEL FITC apoptosis detection kit (Vazyme, A111-03), while apoptosis in heart tissue was assessed using Dead End Fluorometric TUNEL System (Promega, G3250) according to the manufacturer’s protocol. The cell nuclei were stained with Hoechst (1:2000, KeyGEN, 33342) for 20 min, and the extent of TUNEL positivity was assessed as the proportion of TUNEL-positive cells to cardiomyocyte nuclei. Heart tissue sections were stained with a rabbit polyclonal anti-Ki67 antibody (Abcam, ab15580). Proliferation was quantified as the percentage of Ki67-positive nuclei within α-actinin-positive cardiomyocytes. Confocal images were acquired using a Carl Zeiss confocal microscope (Thuringia, Germany).
Echocardiography
Echocardiography was performed with the Vevo2100 system (VisualSonics Inc, Toronto, Ontario, Canada) while controlling the heart rate of mice at 450–500 beats per minute. Anesthesia induction was achieved by administering a 3% isoflurane concentration at a controlled air flow rate of 1.0 L/min. The probe was used to locate the standard parasternal long axis section of the left ventricle to ensure an unobstructed left ventricular outflow tract and visualize the left ventricular papillary muscle. The measuring axis was placed in the center of the long axis of the left ventricle, and left ventricular long axis M mode and B mode images were collected to measure left ventricular ejection fraction (EF) and left ventricular shortening fraction (FS). All the echocardiography parameters measured in this study were listed in Appendix Tables 6, 7 and 8.
TGF-β-induced activation and α-SMA/EdU staining of NRCFs
Neonatal rat cardiac fibroblasts (NRCFs) were first subjected to lentiviral infection for 48 h. Following infection, cells were treated with 20 ng/mL recombinant human TGF-β1 (Peprotech, 100-21C) for an additional 48 h to induce myofibroblast activation. After treatment, cells were washed with PBS and fixed with 4% paraformaldehyde for 20 min at room temperature. Fixed cells were washed three times with PBS, permeabilized using 0.5% Triton X-100 for 20 min, and then blocked with 5% bovine serum albumin (BSA) for 1 h. Cells were then incubated overnight at 4 °C with Cy3-conjugated anti-α-SMA antibody (Sigma, C6198; 1:200 dilution in 5% BSA). After washing, EdU staining was performed using the EdU assay kit (KeyGEN, KGA9602-500) according to the manufacturer’s instructions. EdU A solution was added to the culture medium 24 h prior to fixation. Nuclei were counterstained with Hoechst 33342 for 20 min at room temperature in the dark. Fluorescence images were acquired using a fluorescence microscope, and both the number of EdU-positive cells and α-SMA fluorescence intensity were quantified using ImageJ software.
Polysome fractionation and Npas2 mRNA detection in NRCMs
NRCMs were infected with lentivirus for 48 h. After infection, cells were lysed in polysome extraction buffer containing 20 mM Tris-HCl (pH 7.4), 100 mM KCl, 5 mM MgCl₂, 1% Nonidet P-40 Substitute (Thermo Fisher, 85124), 0.5% sodium deoxycholate, 1 mM dithiothreitol (DTT, Sigma-Aldrich, D9779), 100 μg/mL cycloheximide (Sigma-Aldrich, C4859), RNase inhibitor (Thermo Fisher, AM2694), and cOmplete™ Mini EDTA-free protease inhibitor cocktail (Roche, 11836170001). Lysates were centrifuged at 12,000 × g for 10 min at 4 °C, and supernatants were loaded onto 10–50% sucrose gradients prepared in polysome buffer. Gradients were ultracentrifuged at 35,000 rpm for 2 h at 4 °C in an SW41Ti rotor (Beckman Coulter). Fractionation was performed using an ISCO gradient fractionation system while continuously monitoring absorbance at 254 nm. RNA was extracted from each fraction using RNAiso Plus reagent (TaKaRa, 9109), followed by reverse transcription. Npas2 mRNA in each fraction was measured by RT-qPCR and expressed as a percentage of total Npas2 mRNA across all fractions. The percentages were plotted for each fraction, with monosome- and polysome-associated regions indicated.
Masson’s trichrome staining
The heart tissue was immediately fixed with 4% PFA after collection, followed by embedding in paraffin, sectioning, deparaffinization, and hydration. Masson’s Trichrome staining (Servicebio, G1006) was performed for the visualization of fibrosis according to the manufacturer’s instruction. Images were obtained by Leica microscope (DM3000 LED, Germany), and the percentage of fibrosis area in the total myocardial area was calculated using Image J software.
Hematoxylin and eosin (HE) staining
Heart tissues were immediately fixed in 4% paraformaldehyde (PFA) after collection, followed by paraffin embedding, sectioning, deparaffinization, and hydration. Hematoxylin and eosin (HE) staining was performed using the HE staining solutions kit (KeyGEN, KGE1204-50) according to the manufacturer’s instructions. Images were captured using a Leica microscope (DM3000 LED, Germany).
Statistical analysis
Data collection and analysis were performed by investigators blinded to the experimental groups. All data were presented as mean ± SD. Analyses were performed using SPSS 20.0 software (SPSS Inc., Chicago, IL) or GraphPad Prism 8.0. The distribution of the data was tested for normality before statistical analysis. Statistical comparisons between two groups were performed using an unpaired Student’s t-test to normally distributed variables and a non-parametric Mann-Whitney U test to non-normally distributed variables. Statistical comparisons between three groups were performed using one-way ANOVA test followed by Bonferroni or Dunnett T3 post hoc tests to normally distributed variables and Kruskal-Wallis test with the original FDR method of Benjamini and Hochberg to non-normally distributed variables. Statistical comparisons between four or more groups were performed using two-way ANOVA test with Tukey post hoc to normally distributed variables and Kruskal-Wallis test with the original FDR method of Benjamini and Hochberg to non-normally distributed variables. P values < 0.05 were considered significantly different.
Supplementary information
Appendix Peer Review File Source data Fig. 1 Source data Fig. 2 Source data Fig. 3 Source data Fig. 4 Source data Fig. 5 Source data Fig. 6 Source data Fig. 7 Source data Fig. 8 Figure EV1 Source Data Figure EV2 Source Data Figure EV3 Source Data Figure EV4 Source Data Figure EV5 Source Data Appendix Figure S1 Source Data Appendix Figure S2 Source Data Appendix Figure S3 Source Data Appendix Figure S4 Source Data Appendix Figure S5 Source Data Appendix Figure S6 Source Data Appendix Figure S7 Source Data Expanded View Figures
The reference list from the paper itself. Each links out to its DOI / PubMed record.
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