2.1 Animals

WT C57BL/6 male mice (8–10 weeks old) were sourced from the Southern Medical University Animal Centre, while PINK1-knockout mice (stock no. 017946) came from the Jackson Laboratory. A cardiac-specific PINK1-overexpressing mouse model was established by Shanghai Biomodel Organism Science & Technology Development.¹⁶ All animals were housed under pathogen-free conditions. At the end of the study, all the animals were anaesthetized via isoflurane inhalation (4%) and then euthanized by cervical dislocation. All experimental protocols were approved by the Southern Medical University Institutional Animal Care and Use Committee (No. K2019022) and adhered to the NIH Guide for the Care and Use of Laboratory Animals.

2.2 Cell culture and treatments

NRCMs were isolated from 1- to 3-day-old SD rats following previously established methods. The cells were cultured in Dulbecco's modified Eagle's medium (DMEM) with 10% foetal bovine serum (FBS) in a humidified incubator at 37°C with 5% CO₂. NRCMs were further transfected with plasmid or adenovirus (OBiO) according to the experimental design. PINK1 and Prdx2 siRNAs were purchased from Jiyuan Biotech (Guangzhou). Transfections of the siRNAs and negative control siRNAs were carried out via Lipofectamine 3000 in accordance with the manufacturer's guidelines. Following 48–72 h of transfection, NRCMs were exposed to palmitic acid (500 µmol/L) for 24 h. The generation and sequence information of the PINK1 (full length)-GFP, PINK1 (1-139aa)-GFP and PINK1 (140-580aa)-GFP plasmids are presented in Supporting Information 3, and the H9C2 cell lineage was used in the structural domain experiments.

2.3 HFpEF model

The HFpEF model was constructed as described in a previous study.²³ The mice were exposed to a combination of a HFD (60% calories from lard) and L-NAME (.5 g/L in drinking water) for 15 weeks, while the control group was fed with standard (CHOW) diet for 15 weeks. The L-NAME used in our experiments was purchased from Yuan Ye (S20013; Shanghai) and has a solubility of 100 mg/mL in water. The L-NAME powder can be stored at −20°C for 2 years. Once dissolved, the solution can be stored at −20°C for 1 month and remains stable for 7 days at room temperature. In our experiment, we prepared a .5 g/L L-NAME solution in real time by dissolving the powder in SPF animal drinking water. The solution was changed every 5 days throughout the study. Finally, the effect of L-NAME on blood pressure in mice was assessed.²⁴

2.4 Chemicals and reagents

DMEM and FBS were sourced from Gibco, while collagenase, trypsin, palmitic acid and fatty acid-free BCA were obtained from Sigma. High-fat feed (60%) was purchased from the Guangdong Animal Experiment Centre. The antibodies used in this study included anti-PINK1 (Santa Cruz, sc-517353, rabbit) and the following from Proteintech: anti-Prdx2 (10545-2-AP, rabbit), anti-P-JNK (80024-1-RR, rabbit), anti-P-p38 (28796-1-AP, rabbit), anti-JNK (24164-1-AP, rabbit), anti-p38 (14064-1-AP, rabbit), anti-Bcl2 (26593-1-AP, rabbit), anti-Bax (50599-2-Ig, rabbit), anti-C-Caspase3 (68773-1-Ig, mouse), anti-GFP (50430-2-AP, rabbit), anti-ANP (27426-1-AP, rabbit) and anti-β-actin (66009-1-Ig, mouse).

2.5 Detection of LDs via fluorescence microscopy

To visualize LDs, NRCMs were isolated and inoculated on confocal cell dishes for 3–5 days and were incubated with 2 g/mL BODIPY 493/503. Fluorescent labelling was stopped by washing with PBS, and then immediately fixed with 4% paraformaldehyde. The image scale was set to 20 µm, and the green fluorescence intensity represents the amount of LDs.

2.6 ROS determination

Dihydroethidium (DHE; Sigma) staining of tissue sections and 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA; Beyotime) staining of cells were used to assess ROS production. According to previously established protocols,¹⁵ harvested hearts were embedded in OCT, frozen at −80°C and sectioned using a cryostat. The sections were thawed and incubated with DHE staining for 30 min, followed by three PBS washes. Intracellular ROS levels of cells were quantified using a DCFH-DA assay. Cells were incubated with 10 µM DCFH-DA for 30 min, after which they were stained with DAPI and followed by three PBS washes. All the above incubation processes were carried out at 37°C. Confocal images were acquired using a Leica SP8 microscope, and analysis was performed using ImageJ software. Red fluorescence (DHE; scale bar 100 µm) and green fluorescence (DCFH-DA; scale bar 50 µm) indicate the levels of ROS.

2.7 TdT-mediated dUTP nick-end labelling (TUNEL) assay

A TUNEL assay (Beyotime) was conducted to assess cardiomyocyte apoptosis both in vivo and in vitro. Harvested hearts were embedded, sectioned, dewaxed and stained as instruction. At room temperature, both heart and cells were fixed with 4% formaldehyde for 15 min and permeabilized using .2% Triton X-100 for 20 min. Afterward, the samples were incubated with TUNEL solution at 37°C for 30 min. DAPI staining was then applied, followed by three additional PBS washes. Images were obtained using a Leica SP8 confocal microscope, and analysis was performed with ImageJ software. The image scale was set to 100 µm, and red fluorescence indicates the level of apoptosis.

2.8 Mitochondrial membrane potential (MMP) analysis

JC-1 mitochondrial membrane potential assay kit (Beyotime) was employed to access MMP. Cells were washed by PBS three times and stained with JC-1 solution for 30 min. Then cells were stained with DAPI and followed by three PBS washes. Images were captured with confocal microscope. The green fluorescence represents reduced MMP and red fluorescence corresponds to active MMP. Image-J was used for analysis and the scale bar represents 100 µm.

2.9 Statistical analysis

Data are presented as mean ± SEM. For comparisons between two groups, Student's t-test was applied, while one-way analysis of variance followed by Tukey's test was used for multiple group comparisons. Statistical significance was defined as p < .05, with all tests being two-sided. Analyses were performed using SPSS version 25.

3.1 PINK1 deficiency exacerbates cardiac dysfunction and reduces Prdx2 expression in HFpEF mice

To assess whether PINK deficiency exacerbates HFpEF, we induced HFpEF via a 60% high-fat diet and L-NAME for 15 weeks. Compared with control mice, HFpEF mice presented an increased body mass gain rate (Figure S1A), heart weight-to-body weight (HW/BW) ratio and lung weight-to-heart weight ratio (LW/HW) (Figure S1B). HFpEF mice had higher systolic blood pressure (SBP), serum glucose, serum cholesterol and triglyceride (TG) levels than control mice (Figure S1B,C). These factors were re-evaluated in the PINK1-KO-HFpEF mice; the final body weights of the PINK1-KO-HFPEF mice were reduced, which resulted in an increase in HW/BW. Serum blood glucose and TG levels were greater in PINK1-KO-HFPEF group than WT-HFpEF group (Figure S1B,C). Cardiac function was assessed via echocardiography after the 15 weeks induction of HFpEF. Compared with those in the control group, HFpEF mice exhibited diastolic dysfunction, represented by the increasing ratio between mitral E wave to A wave (E/A) and the mitral E wave to E´ wave (E/E´) (Figure 1C–F). In addition, the left ventricular posterior wall (LVPW) thickness and the opposite wall delay indicated severe left ventricular remodelling (Figure 1B,G). What is more, PINK1 deficiency further exacerbated diastolic dysfunction and left ventricular remodelling in HFpEF mice. Left ventricular ejection fraction was not significantly different between WT control and HFpEF mice but was distinctly reduced in PINK1-deficient HFpEF mice (Figure 1B,D). HFpEF mice exhibited elevated serum BNP than control mice, and PINK1 deficiency further exacerbated BNP levels in HFpEF mice relative to WT mice (Figure S1C), as shown by ELISA. Moreover, serum IFN-β, IL-6, TGF-β and TNFα levels were elevated in HFpEF group than WT control group, and PINK1 deficiency further increased these factors in HFpEF mice (Figure S1C). Masson's staining and wheat germ agglutinin stain (WGA) staining revealed that PINK1 deficiency exacerbated cardiac fibrosis and LV remodelling (Figure 1I,J). In addition, DHE staining revealed that ROS levels were greater in PINK1-deficient HFpEF group (Figure 1K). These results further confirmed that PINK1 deficiency exacerbates HFpEF-induced cardiac dysfunction in mice.

Next, we examined protein expression in PINK1-deficient HFpEF mice and found significantly decreased protein expression of Prdx2. Surprisingly, PINK1 deficiency affected Prdx2 expression (Figure 2A), as shown by immunoblotting. We subsequently examined PINK1 and Prdx2 expression and confirmed the results via immunohistochemistry (Figure 2C,D). To explore the impact of the reduction in Prdx2, we examined several downstream proteins associated with changes in Prdx2 by western blotting. Our findings revealed an increase in JNK and p38 phosphorylation during HFpEF and even more pronounced alterations in PINK1-deficient HFpEF mice (Figure 2A). B-cell lymphoma 2 (bcl2) was decreased, and increases in Bcl-2-associated X protein (Bax) and cleaved caspase-3 cysteine protease (c-caspase 3) were also observed (Figure 2A). We further performed TUNEL staining to compare apoptosis between four groups (Figure 2E), which confirmed an increase in apoptosis in PINK1-deficient mice in the HFpEF group.

3.2 PINK1 overexpression increases Prdx2 and attenuates cardiac diastolic dysfunction in HFpEF mice

We further investigated the effect of PINK1 overexpression following a high-fat diet and L-NAME induction. Therefore, we constructed transgenic mice that overexpressed PINK1 (PINK1-Tg) and compared cardiac function and Prdx2 protein expression among the groups. PINK1-Tg mice presented superior diastolic function and improved cardiac remodelling compared to WT mice after HFpEF induction by a high-fat diet and L-NAME. E/A, E/E´, opposite wall delay and LVPW in HFpEF mice were partially rescued by PINK overexpression (Figure 3B–H). Masson and WGA staining revealed that cardiac fibrosis and left ventricular remodelling were attenuated in the PINK1-Tg mice (Figure 3I,J). In addition, less oxidative stress was observed in the HFpEF hearts of the PINK1-Tg mice than in those of the WT mice (Figure 3K). The PINK1-Tg-HFpEF mice presented improved HW/BW, LW/HW and SBP (Figure S2A,B), and the serum levels of BNP, IFN-β, IL-6, TGF-β and TNF-α were notably lower in the PINK1-Tg mice than in the WT mice (Figure S2C). Immunoblotting revealed an increase in Prdx2 in the PINK1-Tg-HFpEF mice (Figure 4A), which was accompanied by the attenuation of cardiac apoptosis. Immunohistochemistry and TUNEL staining further supported these findings (Figure 4C–E). These results suggest that PINK1 may alleviate cardiomyocyte apoptosis via Prdx2, thereby improving cardiac diastolic function.

3.3 PINK1 interacts with Prdx2, and both are impaired in palmitic acid-induced lipotoxicity

To identify the PINK1 interactome, we immunoprecipitated mouse heart tissue with an anti-PINK1 antibody and performed mass spectrometry to analyse the obtained proteins. A collection of the proteins targeted by PINK1 is provided in Table S1. Intriguingly, the proteins targeted by PINK1 included Prdx2 peptides. To verify the results obtained by mass spectrometry, we performed endogenous co-IP and found that PINK1 was associated with Prdx2 (Figure 5A). We next examined the colocalization of PINK1 with Prdx2 via immunofluorescence analysis (Figure 5E) and found that PINK1 colocalized with Prdx2 in NRCMs. These results suggest that PINK1 binds to Prdx2. According to the UniProt database,²⁵ the supersecond structure of PINK1 includes two alpha helices: the N-terminal region (amino acids 1–133) and the C-terminal region (amino acids 138–500) (Figure S6A). Moreover, the two structures are predicted to be protein modification sites that interact with the other proteins. To identify the structural component of PINK1 that interacts with Prdx2, H9C2 cells were transfected with PINK1 (full-length)-GFP, PINK1 (1-139 aa)-GFP and PINK1 (140-580 aa)-GFP plasmids (Figure 5B). The cells were collected after 48 h for coimmunoprecipitation (co-IP) experiments. After affinity purification, mass spectrometry was used to identify differences in the obtained proteins among the three groups (Table S2), and PEAKS software was used to search the protein database. Additionally, Figure S7 displays the abundance of the obtained proteins in the biological process, cellular component and molecular function categories. According to the mass spectrometry data, distinct signals of the Prdx2 protein were observed in the full-length PINK1 and 1–139 aa PINK1 groups but were absent in the 140–580 aa PINK1 group (Figure S6B and Table S2). The areas enriched with the Prdx2 peptide were essentially identical between the PINK1 (full-length) and PINK1 (1-139 aa) groups (2.28 × 10⁶ and 2.44 × 10⁶) but significantly lower in the PINK1 (140–580 aa) group (1.62 × 10⁶) (Figure S6B and Table S2). These data suggest that the interaction between PINK1 and Prdx2 primarily occurs within the N-terminal region. To verify the results obtained by mass spectrometry, we examined the endogenous interactions of the three groups. Proteins were coimmunoprecipitated with anti-GFP (Figure 5C), and the pulldown samples were immunoblotted with GFP and Prdx2 antibodies. Immunoblotting revealed that Prdx2 was captured by the PINK1 (full length) and PINK1 (1–139 aa) groups (Figure 5D), which suggested that the N-terminal region (amino acids 1–133) of PINK1 interacts directly with Prdx2.

To determine the appropriate concentration for myocardial lipotoxicity, we treated NRCMs with increasing concentrations (0, 125, 250 and 500 µmol/L) of palmitic acid for 24 h. Lipid droplet staining revealed a large increase in the number of intracellular droplets in response to 500 µmol/L palmitic acid (Figure 5F). We then examined PINK1 and Prdx2 protein expression and found that both PINK1 and Prdx2 showed similar trends: an initial increase in response to 125 µmol/L, followed by a decrease as palmitic acid content continued to increase (Figure 5H). In addition, ANP expression increased, and cell viability gradually decreased with increasing palmitic acid concentration (Figure 5H,I). Thus, 500 µmol/L palmitic acid was selected to induce myocardial lipotoxicity to examine the function of PINK1.

3.4 PINK1 ablation increases palmitic acid-induced ROS production and apoptosis by reducing Prdx2

To determine whether PINK1 ameliorates apoptosis in the context of myocardial lipotoxicity, we used siPINK1- and adPINK1-infected NRCMs treated with 500 µmol/L palmitic acid. We examined the expression of apoptosis-associated proteins and found that palmitic acid reduced the expression of PINK1 and Prdx2 and stimulated ROS production (Figure 6A,E). TUNEL staining revealed that apoptosis in NRCMs increased significantly after palmitic acid treatment (Figure S3B). After NRCMs were transfected with siPINK1, western blot and immunofluorescence analyses confirmed that siPINK1 inhibited PINK1 expression (Figure 6A,B). The expression of Prdx2 was also reduced as PINK1 expression decreased (Figure 6A). The increases in the P-JNK/JNK and P-p38/p38 ratios in response to myocardial lipotoxicity were significantly exacerbated by the reduction in Prdx2 (Figure 6A). Subsequent immunoblotting revealed the upregulation of Bax and caspase-3 and the downregulation of Bcl-2, which indicated activated apoptotic flux in palmitic acid-treated NRCMs (Figure 6A) that was greatly exacerbated by siPINK1 transfection. DCHF-DA staining revealed that siPINK1 exacerbated palmitic acid-induced increases in ROS levels (Figure 6E). TUNEL and JC-1 staining further indicated that siPINK1 increased lipotoxicity-induced myocardial apoptosis and decreased the MMP (Figure S3A,B).

3.5 PINK1 overexpression attenuates palmitic acid-induced ROS production and apoptosis by increasing Prdx2

In contrast, the palmitic acid-induced increase in ROS production and apoptosis in NRCMs was significantly reversed by adPINK1 transfection. Immunofluorescence and immunoblotting revealed that the palmitic acid-induced disordered expression of Prdx2 was reversed by PINK1 overexpression (Figure 7A–D); P-JNK/JNK and P-p38/p38 were also suppressed (Figure 7A). Bcl2 was increased and Bax and C-Caspase3 were decreased in palmitic acid-treated NRCMs transfected with adPINK1 (Figure 7A), which indicated that apoptotic flux was obstructed. Moreover, DCFH-DA indicated high levels of ROS generation in palmitic acid-treated NRCMs (Figure 7E), which was improved by adPINK1 treatment. In addition, transfection with adPINK1 restored palmitic acid-induced apoptotic flux and mitochondrial dysfunction, as shown by TUNEL and JC-1 staining (Figure S4A,B), which was consistent with the immunoblot results. Furthermore, an analysis of Prdx2 gene expression and a cycloheximide chase (CHX) assay under PINK1 overexpression intervention revealed that while adPINK1 does not significantly affect Prdx2 mRNA levels, it notably enhances the stability of the Prdx2 protein (Figure S10A,B). The results from the CHX assay provide further evidence of PINK1's involvement in the regulation of Prdx2 at the protein level. These results revealed that PINK1 protected the myocardium from lipid toxicity via Prdx2, which reduced ROS production and apoptosis.

3.6 Prdx2 ablation attenuates PINK1 overexpression-mediated alleviation of palmitic acid-induced lipotoxicity

To confirm the role of Prdx2 in the mechanism by which PINK1 counteracts lipotoxicity, NRCMs were transfected with adPINK1 and siPrdx2, and ROS production and apoptosis flux were examined after palmitic acid treatment. As expected, western blot analysis revealed that Prdx2 blockade reactivated the P-JNK and P-p38 pathways (Figure 8A). Pretreatment with siPrdx2 efficiently reversed the adPINK1-mediated reduction in ROS levels (Figure 8G), as shown by DCFH staining. Moreover, TUNEL staining revealed a decrease in the antiapoptotic effect of adPINK1 after siPrdx2 pretreatment (Figure 8I), which suggested that Prdx2 is essential for PINK1-mediated reductions in ROS production and apoptosis. JC-1 staining also revealed a decrease in the MMP (Figure S5A), which suggested that adPINK1 induced impaired cellular energy metabolism after siPINK1 pretreatment. Taken together, these findings suggest that Prdx2 is important for the PINK1-mediated alleviation of myocardial lipotoxicity.

To further confirm the role of mitochondrial ROS in Prdx2 ablation-induced apoptosis, we performed additional experiments treating cells with palmitic acid and siPrdx2 to induce oxidative stress, while using SS-31 to reduce mitochondrial ROS. MitoSOX staining and confocal fluorescence microscopy revealed that SS-31 significantly attenuated the increase in mitochondrial ROS induced by siPrdx2 (Figure S12A). Moreover, SS-31 reduced malondialdehyde levels elevated by siPrdx2 (Figure S12B). Additionally, we examined the levels of key apoptotic proteins, including Prdx2, Bcl-2, Bax and cleaved Caspase-3. Our results demonstrated that SS-31 effectively decreased mitochondrial ROS levels and blocked the apoptotic effects induced by siPrdx2 (Figure S13), supporting the conclusion that mitochondrial ROS plays a significant role in Prdx2 reduction-induced apoptosis.