The animal study was reviewed and approved by the Animal Care and Use Committee of Zhongshan Hospital, Fudan University, and was in strict accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (revised in 1996).

Male C57BL/6J mice (8–10 weeks old, 20–25 g/mouse) were purchased from Shanghai SLAC Laboratory Animal Company. Pressure overload was induced by transverse aortic constriction (TAC), as we described previously (Wu et al., 2012). Briefly, mice were anesthetized with 2% isoflurane, intubated through the trachea, and connected to a ventilator. Thoracotomy was performed in the left parasternal second intercostal space to separate the aortic arch. Subsequently, the 6-0 silk thread was passed below the aortic arch, followed by ligation of the separated aortic arch with 27 G needle, which was carefully pulled out to result in annular constriction of ascending aorta. Then, the muscle and skin of mice were sutured with 4-0 silk thread and disinfected.

HP model was established as we described previously with appropriate modification to reduce mortality rate during operation (Huang et al., 2017; Wu et al., 2020b). Briefly, after the 1st TAC, a long thread was buried under the chest skin. Three days after TAC, a second thoracotomy was performed, the banding knot was relieved (De-TAC) by microforceps, and the same thread was buried under the chest skin again. Four days after De-TAC, a third thoracotomy was performed and the aortic arch was rebanded (Re-TAC) using the same thread during TAC and De-TAC (Supplementary Figure S1). In order to avoid the interference caused by repeated thoracotomy, mice in the sham group and TAC group also underwent thoracotomy three times at corresponding time points, while the aortic arch in the TAC group was only banded during the third thoracotomy.

The echocardiography was evaluated by a Vevo 2100 high-frequency ultrasound system with a 30 MHz scanner (VisualSonics, Toronto, ON, Canada). Under the anesthesia of 1.5% isoflurane, the left ventricular structure and function were evaluated in M-mode of the parasternal long-axis view (Wu et al., 2010; You et al., 2012; Huang et al., 2017). The indices included the following: heart rate (HR), left ventricular posterior wall end-diastolic thickness (LVPWTd), left ventricular posterior wall end-systolic thickness (LVPWTs), left ventricular ejection fraction (LVEF), and left ventricular fractional shortening (LVFS). Then, the probe was placed on the right side of the mouse and tilted horizontally to display the aortic arch section, and the peak systolic velocity of aortic arch flow (PSVa) was obtained (Huang et al., 2017)

Left ventricular hemodynamics was assessed using the Power Laboratory system (AD Instruments, Castle Hill, NSW, Australia) connected to a micromanometric catheter (Millar 1.4-Fr, SPR 835, Millar Instruments, Houston, TX). After anesthesia with 1.5% isoflurane, the neck skin was cut off, and the right common carotid artery was isolated. The catheter was advanced into the left ventricle through the right common carotid artery. Left ventricular end-systolic pressure (LVESP) and left ventricular end-diastolic pressure (LVEDP) were acquired for left ventricular blood pressure evaluation; then, the maximal rate of pressure rising (Max dP/dt) and maximal rate of pressure fall (Min dP/dt) were collected for left ventricular systolic and diastolic function evaluation (Huang et al., 2017).

After the hemodynamic study, the mice were killed by cervical dislocation. Then, the heart was removed, dried on filter paper, and weighed. The heart weight/body weight (HW/BW) was calculated. Subsequently, the heart was fixed under 4% paraformaldehyde and embedded in paraffin. And the papillary muscle at the most expansive part of the heart was sectioned along the short axis of the horizontal direction with the standard thickness of 4 μm. Hematoxylin-eosin (HE) staining was used to observe the morphology of cardiomyocytes. For visual evaluation of cardiac hypertrophy, the Texas Red™ X-labeled wheat germ lectin (WGA, Invitrogen Corp) was further used to observe cell membranes, and the nuclei were labeled with 4′,6-diamidino-2-phenylindole (DAPI). Masson’s trichrome staining was used to observe the degree of cardiac interstitial and perivascular fibrosis. For the cross-sectional area (CSA) and fibrosis measurements, the images were quantified by an image analysis system (ImageJ 1.52v National Institutes of Health, Bethesda, United States).

The myocardial specific caspase-1 was overexpressed in vivo by using an adeno-associated virus type 9 (AAV9) vector (pAOV-cTNT-EGFP-2A-Casp1-3Flag, Obio Biotechnology, Shanghai, China). A total of 1 × 10¹¹ viral particles (vp) of AAV9-caspase-1 or vector virus AAV9-EGFP (pAOV-cTNT-EGFP-2A-MCS-3Flag, Obio Biotechnology, Shanghai, China) were injected into the tail vein of mice three weeks before surgery.

Total RNA was extracted from heart tissue samples using TRIZOL reagent, and cDNA was synthesized by Reverse Transcription Kit (Takara, Kusatsu, Japan). RT-qPCR was carried out with SYBR Premix Ex Taq II (Takara, Kusatsu, Japan). Cardiac hypertrophy and fibrosis-related genes were amplified using the following primers: ANP, forward: GCT​TCC​AGG​CCA​TAT​TGG​AG and reverse: GGG​GGC​ATG​ACC​TCA​TCT​T; BNP, forward: GAG​GTC​ACT​CCT​ATC​CTC​TGG and reverse: GCC​ATT​TCC​TCC​GAC​TTT​TCT​C; Col1a1, forward: GGA​CGC​CAT​CAA​GGT​CTA​CTG​C and reverse: GAA​CGG​GAA​TCC​ATC​GGT​CAT; Col3a1, forward: CTC​AAG​AGT​GGA​GAA​TAC​TGG​GTT and reverse: GGT​ATG​TAA​TGT​TCT​GGG​AGG​C; GAPDH, forward: GCC​ATC​ACT​GCC​ACT​CAG​AA and reverse: GGC​ATG​TCA​GAT​CCA​CAA​CG. Expression levels of hypertrophic and fibrotic-related genes were shown as 2-ΔΔCt of the target gene relative to GAPDH.

Total protein was obtained from the left ventricular heart tissue. Electrophoresis for protein (20–40 ug) was performed by 10% or 12.5% SDS-polyacrylamide gel and then protein was transferred to 0.22 μm PVDF membrane (Millipore, Billerica, United States). After blocking with 5% BSA for 2 h at room temperature, PVDF membrane with target protein was incubated with primary antibodies overnight at 4°C and secondary antibodies for 2 h at room temperature. The primary antibodies used for WB analysis were anti-p ERK1/2 (1:1,000; CST, United States) and anti-ERK1/2 antibody (1:1,000; CST, United States), anti-p p38 antibody (1:1,000; CST, United States), anti-p38 antibody (1:1,000; CST, United States), anti-p JNK antibody (1:1,000; CST, United States), anti-JNK antibody (1:1,000; CST, United States), anti-NLRP3 antibody (1:1,000; CST, United States), anti-caspase-1 antibody (1:500; Santa Cruz, United States), anti-cleaved caspase-1 antibody (1:1,000; CST, United States), anti-caspase-11 antibody (1:1,000; CST, United States), anti-GSDMD antibody (1:1,000; Abcam, United States), anti-IL-1β antibody (1:1,000; CST, United States), anti-IL-18 antibody (1:1,000; CST, United States), anti-cleaved IL-18 antibody (1:1,000; MBL, United States), and anti-GAPDH antibody (1:5,000; Abcam, United States). HRP-conjugated anti-rabbit or mouse secondary antibody was used as the secondary antibody (1:5,000; Weiao Biotechnology, Shanghai, China). WB detection system (Bio-Rad, United States) was used to detect the imprint after treatment of PVDF membrane by Millipore ECL kit (Invitrogen, Carlsbad, United States), and optical density was quantified using ImageJ analysis software (1.52v, National Institutes of Health, Bethesda, United States).

All continuous data in our study were presented as the mean ± standard error (SE). GraphPad Prism 8 (GraphPad Software, Version 8.01, San Diego, CA, United States) was used for statistical analysis. The normality tests were performed by the Shapiro–Wilk test. Based on the characteristics of data distribution, the difference between groups was analyzed by one-way ANOVA (only operation treatment) or two-way ANOVA (operation and virus treatments), followed by the Student–Newman–Keuls (SNK) test (data with normal distribution) or Kruskal–Wallis test followed by Dunn’s test (data not normally distributed) for multiple comparisons. A p value < 0.05 was considered statistically significant.

Both TAC 4W and Re-TAC 4W mice showed similar HRs and similarly elevated PSVa compared with those of sham mice (Figures 1H,I, Supplementary Figure S2). Compared with sham mice, TAC 4W mice showed prominent cardiac hypertrophy, as measured by increased HW/BW (Figure 1J), inner dimension (Figures 1K,L), LV wall thickness (Figures 1M,N), and fibrosis (Figures 1D,E,P–Q), enhanced levels of reprogramming of fetal genes ANP and BNP (Figures 1R,S), and augmentation of cardiomyocyte size (Figures 1A–C,O). Re-TAC 4W mice demonstrated that HP decreased the hypertrophic response compared with TAC 4W mice.

Meanwhile, TAC 4W mice showed marked cardiac dysfunction, as demonstrated by depressed LVEF and LVFS (Figures 1F,T,U), which was due to notable dilation of LVESD (Figure 1L). On the contrary, HP improved cardiac function and reduced LVESD to the level of sham mice. We also employed invasive hemodynamics to further assess the effects of HP on cardiac function. We found that LVESP increased in both the TAC group and HP group, while the HP group showed a significant reduction in LVEDP compared with that of the TAC group (Figures 1G,V,W). LV systolic and diastolic functions were also improved by HP, as indicated by higher maximal dP/dt and minimal dP/dt, respectively (Figures 1G,X,Y).

Mitogen-activated protein kinase (MAPK) is a type of protein kinase that is involved in directing cellular responses to hypertrophic stimuli (Zhang et al., 2017; You et al., 2018). The MAP kinases can be grouped into three main families, namely, ERK1/2 (extracellular-signal-regulated kinases), p38/SAPKs (stress-activated protein kinases), and JNK1/2 (Jun amino-terminal kinases). MAPK plays an important role in the development of cardiac hypertrophy (Li et al., 2016; Tamura et al., 2019). We, therefore, assessed the protein expression levels of ERK1/2, p38, and JNK. Compared with the sham group, the phosphorylation levels of ERK, p38, and JNK in both TAC and HP groups were significantly increased. Nevertheless, the hypertrophy signaling in Re-TAC 4W mice group was significantly downregulated compared with the TAC group (Figures 2A,B).

As for fibrosis signaling, compared with the sham group, fibrosis-related factors COL1A1 and COL3A1 were greatly elevated in TAC 4W mice, whereas they were much less elevated in Re-TAC 4W (Figure 2C).

To unveil the changes of caspase-1 in HP, we found remarkable activation of caspase-1 in TAC 4W mice (Figures 3A,B). In contrast, Re-TAC 4W mice showed significantly decreased caspase-1 activation compared with TAC 4W mice (Figures 3A,B), suggesting close relations of caspase-1 with HP. Caspase-1 induces inflammation through cytokines IL-1β and IL-18 and causes pyroptosis through executor GSDMD (Shi et al., 2015). To further answer whether its downstream targets were also associated with HP, we found that IL-1β and IL-18 were both less activated in Re-TAC 4W mice, compared with TAC 4W mice (Figures 3A,B). Intriguingly, although compared with sham mice, the activation of GSDMD was increased in both TAC and Re-TAC mice, no significant difference was observed between the two groups (Figures 3A,B). Considering that caspase-1 and caspase-11 mediate a canonical pathway and a noncanonical pathway of pyroptosis, respectively, we also investigated the changes of caspase-11. We found caspase-11 was activated in TAC 4W mice but remained markedly activated in Re-TAC 4W mice, suggesting that the augmented pyroptosis originated in a noncanonical pathway (Figures 3A,B). To unveil the mechanism of the caspase-1 variations, we also investigated the change of NLRP3, the upstream effectors of caspase-1. In accordance with the caspase-1 level, the expression of NLRP3 was upregulated in TAC 4W mice but was downregulated in Re-TAC 4W mice (Supplementary Figure S3). Taken together, our findings indicated that HP mitigated the function of caspase-1 through attenuation of inflammation rather than pyroptosis.

To further clarify the relationship between caspase-1 and HP in the development of hypertrophy and fibrosis, we constructed an AAV9 virus vector to overexpress caspase-1 in mice hearts through tail vein injection. Overexpression of caspase-1 had no significant effect on PSVa and HRs (Figures 4H,I, Supplementary Figure S4). However, caspase-1 overexpression dramatically abrogated the beneficial effects of HP, which was evidenced by increased HW/BW (Figure 4J), inner dimension (Figures 4K,L), LV wall thickness (Figures 4M,N), and fibrosis (Figures 4D,E,P,Q), enhanced levels of ANP and BNP (Figures 4R,S), and augmentation of cardiomyocyte size (Figures 4A–C,O). Consistently, caspase-1 overexpression deteriorated cardiac function in HP mice, which was indicated by lower LVEF (Figures 4F,T), LVFS (Figures 4F,U), maximal dP/dt (Figures 4G,X), minimal dP/dt (Figures 4G,Y), and higher LVEDP (Figures 4G,V), while LVESP (Figures 4G,W) did not change significantly.

To answer whether caspase-1 alters the signaling pathways underlying the hypertrophic response in the context of HP, we examined the activation levels of MAPK under a Re-TAC background. Compared with EGFP treated Re-TAC mice, caspase-1 overexpression noticeably enhanced phosphorylated ERK, p38, and JNK (Figures 5A,B). In addition, caspase-1 overexpression dramatically aggravated the gene levels of COL1A1 and COL3A1 (Figure 5C).

Since HP mitigates activation of caspase-1 mediated inflammatory cytokines IL-1β and IL-18, we examined whether IL-18 and IL-1β were involved in the detrimental effects of caspase-1 overexpression in HP mice. As expected, we found that caspase-1 overexpression aggravated the activation of IL-1β and IL-18 in Re-TAC 4W mice (Figures 6A,B), recapitulating that HP attenuates cardiac hypertrophy and regresses heart failure through downregulation of the inflammatory cascade of caspase-1.

This study indicates that myocardium caspase-1 overexpression diminishes the benefits of HP in pressure overload mouse hearts. However, the heart is composed of cardiomyocytes, fibroblasts, endothelial cells, and immune cells, not only cardiomyocytes (Litviňuková et al., 2020). We did not differentiate the effects of HP and caspase-1 on cardiomyocytes from those on other types of cells, especially immune cells, nor did we investigate the communication between cardiomyocytes and other types of cells. With the application of state-of-the-art technology in cardiovascular research, such as transcriptome analysis and single-cell sequencing technology (Murashige et al., 2020; Wang et al., 2020), more mechanisms of HP will be explored in-depth.