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Introduction

Myocardial ischemia (MI) is typically caused by inadequate blood flow in the coronary arteries. A worldwide epidemiological analysis indicated that the number of cases of MI steadily increased from 1990-2019, reaching a total of 197 million cases by 2019 (1). Myocardial infarction, caused by prolonged and severe MI, is a prevalent cardiovascular disease and a major cause of disability and mortality worldwide (2,3). The accumulation of reactive oxygen species (ROS) during MI creates conditions for the generation of oxidative stress (4). Currently, the primary treatment method for myocardial infarction is coronary artery reperfusion (5). This procedure is crucial for treating ischemic myocardial tissue damage by removing harmful metabolites and restoring normal metabolism (4). However, the sudden reintroduction of high oxygen tension due to blood flow reperfusion results in increased levels of oxygen free radicals, leading to a surge in ROS levels. This causes more severe tissue damage than that caused by ischemia (6-8), a condition known as MI/reperfusion injury (MIRI). MIRI is a complex pathological process related to the production of ROS and mitochondrial dysfunction, involving reduced ATP production and destruction of the mitochondrial ridge (9). Therefore, it is necessary to clarify the molecular mechanisms underlying MIRI to develop effective prevention strategies.

During reperfusion, uncontrolled ROS-mediated oxidative stress has been considered a key triggering factor for MIRI (10). ROS mainly originate from the byproduct of the mitochondrial respiratory chain (11). Under physiological conditions, ROS participate in signal transduction and regulate various physiological activities of the heart (12). However, uncontrolled increases in ROS levels cause damage to biological macromolecules, such as lipid peroxidation and mitochondrial dysfunction, which may lead to ferroptosis (13). Nuclear factor erythroid 2-related factor 2 (Nrf2) is a transcription factor that activates endogenous antioxidant enzymes in response to oxidative stress and ferroptosis (14). It is widely expressed in tissues of oxygen-consuming organs, such as the muscles, heart and blood vessels (15). When oxidative stress occurs, Nrf2 is activated and is transferred from the cytoplasm to the nucleus to maintain cellular redox balance and avoid the occurrence of lipid peroxidation (16).

The regulator of G-protein signaling (RGS) family serves a crucial role in the regulatory processes of the cardiovascular system (17). A previous study reported a significant increase in the mRNA and protein expression levels of RGS3 and RGS4 in failing heart tissues (18). It has been shown that the expression of RGS2 inhibits G-protein signaling in the myocardium of adult mice, which is necessary to prevent the early development of compensatory hypertrophy in response to pressure overload (19). The deletion of RGS6 exacerbates ischemic injury in the hearts of mice (20). RGS12 is currently the largest known mammalian RGS protein (21). It has been reported that the expression of RGS12 may contribute to pathological cardiac hypertrophy (22). RGS12 contains a central RGS domain, a postsynaptic density protein-95/discs-large/zona occludens homology domain, a phosphotyrosine binding domain, tandem Ras binding domains and a Gαi/o-Loco interaction motif (21), which suggests the versatility of RGS12 and its association with multiple signaling pathways. RGS12 has been reported to regulate oxidative stress through Nrf2 in various tissues, including neuronal cells (23) and osteoclasts (24); however, its role in the heart remains unexplored. The deficiency of RGS12 activates Nrf2 and impairs the production of ROS in osteoclast (24), but overexpression of RGS12 inhibited the activation of Nrf2 (24). Targeting RGS12 may thus alleviate oxidative stress and inflammation by regulating the Nrf2 pathway in the hippocampus of depressed rats (23). However, in MIRI, it is currently unknown whether RGS12 affects the Nrf2 pathway.

Penehyclidine hydrochloride (PHC) is an anticholinergic drug with antimuscarinic and antinicotinic activity (25). Previous studies have reported that preconditioning with PHC is effective in preventing ischemia/reperfusion injury in multiple organs, which is achieved by inhibiting apoptosis and relieving oxidative stress (26,27). Preconditioning with PHC alleviates the mitochondrial dynamic imbalance and protects myocardial cells against MIRI (28). This suggests a potential protective role of PHC against MIRI; however, the underlying molecular mechanisms have not yet been fully elucidated.

The present study aimed to establish a mouse model of MIRI and a cell model subjected to hypoxia/reoxygenation (H/R) to investigate the regulatory role and molecular mechanisms of RGS12 in MIRI and the myocardium after PHC preconditioning.

Materials and methods

Adenovirus (Ad) construction

The short hairpin RNA (shRNA) sequence targeting mouse RGS12 (shRGS12, 5′-GGACCTCAGCACTCGAGAAAG-3′), along with a non-specific sequence (shNC, 5′-TTCTCCGAACGTGTCACGT-3′), were synthesized and inserted into pShuttle-CMV plasmids (cat. no. BR009; Hunan Fenghui Biotechnology Co., Ltd.) by General Biology, Inc. A total of 25 μg shuttle plasmids were transfected into 293A cells at room temperature using Lipofectamine 3000 (cat. no. L3000015; Thermo Fisher Scientific, Inc.) and P3000 (cat. no. L3000015; Thermo Fisher Scientific, Inc.). The coding region of RGS12 was cloned into pDC315 plasmids by YouBio (cat. no. VT1500) to construct the RGS12 overexpression vector. The mass of nucleic acid used was 9 μg. Empty vector without any exogenous fragment was used as a negative control. A total of 27 μg plasmids (shuttle plasmid: skeleton plasmid=1:2) were transfected into the 293A cells at room temperature using Lipofectamine 3000 (41.8 μl) (cat. no. L3000015; Thermo Fisher Scientific, Inc.) and P3000 (54 μl) (cat. no. L3000015; Thermo Fisher Scientific, Inc.). After 14 days, the cell supernatant containing Ad particles was collected.

Animal experiments

The present study utilized 210 10-week-old male C57BL/6 mice (weighing 24.0±2.0 g) that underwent a 1-week acclimatization period to the feeding regimen. During the experiment, the mice were allowed to eat and drink freely. Mice were subjected to a 12 h light/dark cycle under controlled environmental conditions, including a temperature maintained at 22±1°C and humidity levels between 45-55%. The animal experiments were approved by the Ethics Committee of Hebei North University (approval no. HBNU202306022105; Zhangjiakou, China).

The mice were allocated into two groups (Sham and MIRI) through a random process, and were anesthetized using an intraperitoneal injection of 75 mg/kg ketamine and 10 mg/kg xylazine. Mice in the MIRI group were subjected to left main coronary artery ligation for a duration of 30 min to induce ischemia, followed by the restoration of blood flow for 24 h to establish the MIRI model (29). The mice in the Sham group underwent a surgical procedure without ligation as a control measure. After surgery, all the animals were administered a subcutaneous injection of 5 mg/kg carprofen for pain relief (30) and were placed in a comfortable environment that included maintaining a constant temperature and humidity, as well as providing a cage with adequate space and non-toxic, non-irritating bedding material. Animal health and behavior were monitored every 4-6 h. According to the National Institutes of Health Guidelines for Endpoints in Animal Study Proposals (31) the following humane endpoints were used: Anorexia (lack or loss of appetite), failure to drink, labored breathing, gasping, lethargy or persistent recumbency and excessive hyperthermia or hypothermia.

Next, the mice were randomly divided into two groups [MIRI + Ad-RGS12 and MIRI + Ad-empty vector (EV)]. After the mice were anesthetized, the plasmid vectors carrying the coding sequence of RGS12 or EVs were packaged into Ads and injected into the left ventricular free wall of the mice in both groups with a total volume of 30 μl and a concentration of 1×1011 plaque forming units/ml, 72 h prior to the induction of MIRI (32).

For further experiments, the mice were randomly divided into four groups, denoted as the MIRI, MIRI + PHC, MIRI + PHC + Ad-EV and MIRI + PHC + Ad-RGS12 groups. The mice in the experimental MIRI + PHC, MIRI + PHC + Ad-EV and MIRI + PHC + Ad-RGS12 groups received a tail vein injection of PHC (cat. no. HY-137976; Merck KGaA) at a dose of 1 mg/kg body weight 1 h prior to the induction of MIRI (33). The mice in the MIRI + PHC + Ad-EV and MIRI + PHC + Ad-RGS12 groups received Ad injections 72 h prior to undergoing MIRI, followed by the administration of PHC 1 h prior to the induction of MIRI.

The left ventricular function of all the mice was evaluated under anesthesia utilizing an echocardiographic imaging system. Subsequently, the mice were sacrificed via exsanguination. When the mice experienced cardiac arrest and their pupils were dilated, they were considered deceased. Blood (~1 ml) from postcava was collected. Serum and myocardial tissues were subsequently obtained for further analysis.

A total of 210 animals were used in the present study, of which 195 were euthanized (n=24/group), while 15 were found deceased during the operation. Of the animals euthanized, 2 animals experienced persistent recumbency and were euthanized at the humane endpoints and 1 animal was used to explore the appropriate antibody concentrations and incubation times in a preliminary experiment.

Cell culture

The mouse myocardial cell line HL-1 (cat. no. iCell-m077; iCell) was cultured in Minimum Essential Medium (cat. no. 41500; Beijing Solarbio Science & Technology Co., Ltd.) containing 10% FBS and 1% penicillin streptomycin and incubated at 37°C and 5% CO2.

Cell induction

HL-1 cells were exposed to various concentrations of PHC (1, 2.5 and 5 μg/ml) at 37°C for a duration of 2 h (34), after which they were subjected to H/R injury according to previously established protocols (35). Briefly, the cells were cultured in a medium devoid of FBS for a duration of 12 h. Following this, the cells were subjected to hypoxic conditions consisting of 1% O2, 94% N2 and 5% CO2 for a period of 8 h. Subsequently, the cells were returned to normal culture conditions for 12 h to induce H/R injury. Control cells were cultured under normal conditions. Cell viability was assessed using the Cell Counting Kit-8 assay in accordance with the manufacturer's protocol (cat. no. KGA317; Nanjing KeyGen Biotech Co., Ltd.). The cells were incubated with CCK-8 reagent at 37°C for 2 h.

Infection

Ad was added to the culture medium. Following a 48-h infection period, the HL-1 cells were exposed to PHC (5 μg/ml) at 37°C and H/R was induced in cells after 2 h. The concentration of 5 μg/ml was selected on the basis that 5 μg/ml PHC treatment had the strongest inhibition of RGS12 expression and did not decrease cell viability.

Staining with 2,3,5-triphenyl tetrazolium chloride (TTC)

Myocardial tissues cryopreserved at -2°C were sectioned into 1 mm slices, followed by staining with a 0.4% solution of TTC at 37°C for 15 min in the dark. The viable myocardial tissues exhibited a red stain, while the infarcted region displayed a white appearance.

Measurement of biochemical markers

The activity of LDH, CK and AST in serum were measured. The content of MDA and the activity of SOD in ischemic penumbra of myocardial tissues were detected. The content of MDA and the activity of SOD and CAT in the cells were detected. Samples were preprocessed according to the instructions provided by the kits' manufacturer. The myocardial tissues were homogenized in normal saline, then centrifuged at 4°C at 12,000 × g for 10 min and the supernatant was collected for analysis. The cells were lysed using ultrasound on ice (power, 200 W; 5 sec ultrasonic treatment; 15 sec interval, repeated 5 times) and then centrifuged at 4°C at 15,000 × g for 10 min to separate the supernatant for detection. The serum was analyzed directly without any pretreatment. The activities of lactate dehydrogenase (LDH; cat. no. A020), creatine kinase (CK; cat. no. A032), aspartate aminotransferase (AST; cat. no. C010), catalase (CAT; cat. no. A007), glutathione peroxidase (GPX; cat. no. A005) and superoxide dismutase (SOD; cat. no. A003-1), as well as the concentration of malondialdehyde (MDA; cat. no. A001), were quantified utilizing test kits procured from Nanjing Jiancheng Bioengineering Institute in accordance with the manufacturer's protocols. The activity of all enzymes was measured through enzymatic reaction at 37°C. MDA content was determined based on the Thibabituric Acid assay at 95°C (cat. no. A001; Nanjing Jiancheng Bioengineering Institute). Serum troponin T (TnT) levels were assessed using an ELISA kit (cat. no. SEB820Mu; Wuhan USCN Business Co., Ltd.).

Histopathological analysis

Myocardial tissues obtained from the left ventricle were preserved in a 4% paraformaldehyde solution at room temperature for 24 h, subsequently embedded in paraffin and sectioned into 5 μm slices. The sections underwent a dewaxing process and were stained with hematoxylin (5 min) and eosin (3 min) at room temperature. Subsequently, the samples were imaged using a light microscope (BX53; Olympus Corporation).

Immunofluorescence

The HL-1 cell and ischemic penumbra of myocardial tissue were pretreated separately. The cells (5×104) were seeded onto coverslips in a 24-well plate and subsequently fixed using 4% paraformaldehyde at room temperature for 20 min. Cells were permeabilized using 0.1% Triton X-100 (cat. no. ST795; Beyotime Institute of Biotechnology) for 30 min at room temperature. Tissues were embedded in paraffin and sectioned into 5-μm slices. The slices were rehydrated in descending alcohol series (95, 85, and 75%), and boiled in an antigen retrieval solution (1.8% 0.1 M citrate buffer and 8.2% 0.1 M sodium citrate buffer) at 95°C for 10 min. The slices were washed in a PBS buffer. Subsequently, the prepared tissue or cell specimens were treated with 1% BSA (cat. no. A602440-0050; Sangon Biotech Co., Ltd.) and incubated for 15 min at room temperature. Samples were incubated at 4°C overnight with the following primary antibodies: RGS12 (cat. no. DF4415; 1:100; Affinity Biosciences) and Nrf2 (cat. no. AF0639; 1:100; Affinity Biosciences). Following multiple washes in PBS buffer, samples were incubated for 1 h at 37°C with Alexa Fluor 555-conjugated goat anti-rabbit IgG secondary antibodies (cat. no. 4413; 1:200; Cell Signaling Technology, Inc.) in the dark. DAPI (cat. no. D106471-5mg; Shanghai Aladdin Biochemical Technology Co., Ltd.) culture for 5 min at room temperature was utilized as a nuclear counterstain. Subsequently, the stained sections were imaged using a fluorescence microscope.

ROS and lipid ROS detection

The ischemic penumbra of myocardial tissue was embedded in Optimal Cutting Temperature Compound freezing medium (cat. no. 4583; Sakura Finetek USA, Inc.), frozen at −20°C and cut into 10 μm sections for staining. Cells (1×105) were seeded into a 12-well plate and cultured to a confluence of 70-80%, and subsequently prepared for staining procedures. DHE (4 μM; cat. no. D807594; Shanghai Macklin Biochemical Co., Ltd.) was added to the tissues and cells and incubated at 37°C for 30 min. The stained sections and cells were then examined using a fluorescence microscope to determine ROS levels.

To detect lipid ROS levels, cells (5×105) were seeded into a 6-well plate and incubated with C11-BODIPY 581/591 (cat. no. MX5211; Shanghai Maokang Biotechnology Co., Ltd.) at 37°C for 30 min, and the alteration in fluorescence emission peak wavelength from 590-510 nm was assessed using a flow cytometer (NovoCyte; Agilent Technologies, Inc.).

Fe2+ detection

The presence of Fe2+ in tissues and cells was identified using a Ferrous Iron Colorimetric Assay kit [cat. no. E-BC-K773-M (for tissues); cat. no. E-BC-K881-M (for cells); Wuhan Elabscience Biotechnology Co., Ltd.] according to the manufacturer's instructions.

Reverse transcription-quantitative PCR (RT-qPCR)

The HL-1 cells and ischemic penumbra of myocardial tissue were homogenized in TRIpure lysis buffer (cat. no. RP1001; BioTeke Corporation) and the total RNA concentration was assessed using an ultraviolet spectrophotometer (NanoDrop 2000; Thermo Fisher Scientific, Inc.). Subsequently, cDNA was synthesized using the BeyoRT II M-MLV reverse transcriptase (cat. no. D7160L; Beyotime Institute of Biotechnology) as per the manufacturer's guidelines. The resulting cDNA was utilized as a template for qPCR. The qPCR procedure was conducted with SYBR Green (cat. no. SY1020; Beijing Solarbio Science & Technology Co., Ltd.) and 2×Taq PCR MasterMix (cat. no. PC1150; Beijing Solarbio Science & Technology Co., Ltd.) using a fluorescent qPCR instrument (Exicycler 96; Bioneer Corporation). The following thermocycling conditions were used: 95°C for 5 min, followed by 95°C for 10 sec, 60°C for 10 sec and 72°C for 15 s. The 2−ΔΔCq method was used to analyze the target gene expression (36). The primers used were obtained from General Biology, Inc. and the sequences were as follows: RGS12 forward (F), 5′-AAGCGGACTTTGTTTCGG-3′ and reverse (R), 5′-GGAGCACCTTTCTGTTTGT-3′; and β-actin F, 5′-CATCCGTAAAGACCTCTATGCC-3′ and R, 5′-ATGGAGCCACCGATCCACA-3′.

Western blotting

The HL-1 cells and ischemic penumbra of myocardial tissue were used for western blotting. Total protein was extracted using lysis buffer (cat. no. P0013; Beyotime Institute of Biotechnology) containing phenylmethanesulfonyl fluoride (cat. no. ST506; Beyotime Institute of Biotechnology) on ice for 5 min. The resulting lysates were centrifuged at 10,000 x g for 5 min at 4°C to obtain supernatants containing proteins. The cytoplasmic and nuclear proteins were then separated using the Nuclear Protein and Cytoplasmic Protein Extraction kit (cat. no. P0027; Beyotime Institute of Biotechnology). The protein concentration was determined using the BCA method (cat. no. P0011; Beyotime Institute of Biotechnology). The 30 μg protein samples were then separated using SDS-PAGE with varying concentrations of gel (8, 10 and 14%) and a 5% stacking gel, followed by transfer onto PVDF membranes. The membranes were blocked using a blocking buffer (cat. no. P023; Beyotime Institute of Biotechnology) at room temperature for 1 h and subsequently incubated overnight at 4°C with primary antibodies targeting RGS12 (cat. no. DF4415; 1:500; Affinity Biosciences), GPX4 (cat. no. A1933; 1:1,000; ABclonal Biotech Co., Ltd.), solute carrier family 7 member 11 (SLC7A11; cat. no. DF12509; 1:500; Affinity Biosciences) and Nrf2 (cat. no. AF0639; 1:1,000; Affinity Biosciences). After washing using a washing reagent (cat. no. P023; Beyotime Institute of Biotechnology), the blots were incubated for 45 min at 37°C with horseradish peroxidase-conjugated secondary antibodies (goat anti-rabbit IgG, cat. no. A0208; goat anti-mouse IgG, cat. no. A0216; 1:5,000; Beyotime Institute of Biotechnology). Histone H3 (cat. no. AF0009; 1:1,000; Beyotime Institute of Biotechnology) and β-actin (cat. no. sc-47778; 1:1,000; Santa Cruz Biotechnology, Inc.) were used as internal references for nuclear protein and total protein, respectively. The blots were visualized using ECL reagent (cat. no. P0018; Beyotime Institute of Biotechnology).

Transmission electron microscopy (TEM) analysis

The cells were fixed using an Electron Microscope Fixative (cat. no. G1102; Wuhan Servicebio Technology Co., Ltd.) and incubated at 4°C for 2 h before being embedded in 1% agarose. Cells were treated with 1% osmium tetroxide for at room temperature 2 h. Following dehydration in ascending alcohol series (30, 50, 70, 80, 95 and 100%) the cells were encased in 812 epoxy resin monomers (cat. no. 02659-ABl; Structure Probe, Inc.) at 37°C overnight. Embedded cells were sliced into 60-80 nm sections, stained with 2% uranyl acetate (8 min) and 2.6% lead citrate (8 min) at room temperature and subsequently examined using TEM (cat. no. H-7650; Hitachi High-Technologies Corporation). The mitochondria were manually counted.

Statistical analysis

Data were statistically analyzed using GraphPad software (version 9.0; Dotmatics). The data are presented as the mean ± SD. Statistical analysis was conducted using the unpaired Student's t-test or one-way ANOVA with Tukey's post-hoc test to identify significant differences between the experimental groups. P<0.05 was considered to indicate a statistically significant difference.

Results

RGS12 is highly expressed in a MIRI model of mice and in cells subjected to H/R

The MIRI model was developed using a 30 min ischemia and 24 h reperfusion (Fig. 1A). The present study assessed heart function and myocardial damage-related markers in mice to confirm the successful establishment of the MIRI model. The echocardiography results indicated a significant reduction in the left ventricular ejection fraction (LVEF) and left ventricular fractional shortening (LVFS) in the mice with MIRI compared with the control group (Fig. 1B). The mice with MIRI exhibited severe injury in the myocardial tissues, as indicated by an increase in the size of the infarct (Fig. 1C) and significantly increased activity levels of LDH and CK in comparison with the control mice (Fig. 1D). The intensity of red fluorescence in the ischemic penumbra of the mice with MIRI was greater compared with that of the control mice, indicating an increase in ROS levels caused by MIRI (Fig. 1E). The concentration of Fe2+ in the myocardial tissues of mice with MIRI was significantly higher compared with that in the control group (Fig. 1F), suggesting the occurrence of ferroptosis. These results indicated that MIRI was successfully induced in the mice subjected to ischemia/reperfusion. Subsequently, it was demonstrated that RGS12 was highly expressed in the mice with MIRI compared with the control group, as evidenced by the results of RT-qPCR, western blotting analysis and immunofluorescence (Fig. 1G and H). Moreover, a cell model of H/R-induced injury was developed using 12 h serum starvation, 8 h hypoxia and 12 h reoxygenation (Fig. 1I). The expression level of RGS12 was assessed. Similar patterns of RGS12 expression levels were also observed in HL-1 cells with H/R injury compared with control cells (Fig. 1J and K).

Figure 1 RGS12 is highly expressed in MIRI mice and H/R cells. (A) The MIRI model was developed using a 30 min ischemia and 24 h reperfusion. (B) Echocardiogram, LVEF and LVFS of mice. (C) Quantification of the infarct area in the myocardial tissue and images of infarct tissue. (D) Serum LDH and CK activity. (E) ROS levels (magnification, ×400) and (F) Fe2+ content in the myocardial tissue. RGS12 expression levels in the myocardial tissue was analyzed using (G) reverse-transcription quantitative PCR and western blotting and (H) immunofluorescence. Magnification, ×400. n=6. (I) A cell model of H/R-induced injury was developed using a 12 h serum starvation, 8 h hypoxia and 12 h reoxygenation. RGS12 expression levels in H/R cells was analyzed using (J) reverse transcription quantitative PCR and western blotting and (K) immunofluorescence. Magnification, ×400. n=3. Data were presented as the mean ± SD. MIRI, myocardial ischemia/reperfusion injury; H/R, hypoxia/reoxygenation; RGS12, regulator of G-protein signaling 12; LVEF, left ventricular ejection fraction; LVFS, left ventricular fractional shortening; LDH, lactate dehydrogenase; CK, creatine kinase; ROS, reactive oxygen species.

Downregulation of RGS12 relieves oxidative stress in H/R cells

To investigate the involvement of RGS12 in MIRI, the expression of RGS12 was knocked down in HL-1 cells (Fig. S1A and B). Increased protein expression levels of RGS12 were observed in the cells subjected to H/R injury, except for the cells in which RGS12 expression was suppressed (Fig. 2A). The viability of the cells subjected to H/R was significantly reduced compared with the control cells; however, the silencing of RGS12 resulted in a significant increase in cell viability (Fig. 2B). Elevated levels of ROS and MDA were detected in the cells subjected to H/R compared with those in the controls (Fig. 2C and D). Conversely, the silencing of RGS12 led to a reduction in ROS levels and the inhibition of MDA production. The activities of SOD and CAT significantly decreased following H/R-induced injury; however, these significantly increased after the silencing of RGS12 (Fig. 2E and F).

Figure 2 RGS12 silencing relieves oxidative stress in H/R cells. (A) RGS12 protein expression levels in H/R cells. (B) Cell survival rate. (C) ROS levels (magnification, ×100). (D) MDA content. (E) SOD and (F) CAT activity in H/R cells. n=3. MDA, malondialdehyde; SOD, superoxide dismutase; CAT, catalase; H/R, hypoxia/reoxygenation; RGS12, regulator of G-protein signaling 12; sh, short hairpin RNA; NC, negative control; prot, protein; ROS, reactive oxygen species. The data are presented as the mean ± SD.

Downregulation of RGS12 inhibits ferroptosis in H/R cells

Excess ROS and MDA levels often predict the occurrence of ferroptosis (13). The levels of Fe2+ in the cells subjected to H/R significantly increased, while these levels were significantly reduced by the silencing of RGS12 (Fig. 3A). Mitochondrial morphology was assessed using TEM and the control cells displayed intact mitochondria (Fig. 3B). Conversely, H/R-induced damage was evident in the mitochondrial membrane and cristae, with protective effects observed after the silencing of RGS12. The proportion of damaged mitochondria was calculated by evaluating the ratio of the number of damaged mitochondria to the total number of mitochondria in the visual field (Fig. 3B), which was consistent with the results shown in the representative images (Fig. 3C). Lipid ROS detection using C11-BODIPY 581/591 staining, combined with flow cytometric analysis, indicated that RGS12 silencing reduced lipid ROS production (Fig. 3D). Additionally, the proportion of C11-BODIPY 581/591-positive cells in the samples were measured (Fig. 3E and F). Cells undergoing ferroptosis exhibited significantly elevated lipid ROS levels, which were significantly suppressed by the silencing of RGS12. In addition, the protein expression levels of GPX4 and SLC7A11 were significantly reduced in the cells subjected to H/R compared with those in the control cells. However, the silencing of RGS12 expression significantly increased the protein expression levels of GPX4 and SLC7A11.

Figure 3 RGS12 silencing inhibits ferroptosis in H/R cells. (A) Fe2+ content. (B) Damaged mitochondria were indicated by the ratio of the number of damaged mitochondria to the total number of mitochondria. (C) Mitochondrial morphology (magnification, ×20,000). The red arrows represent damaged mitochondria and the green arrows represent normal mitochondria. Lipid reactive oxygen species levels were analyzed using (D) flow cytometry and the results were presented as (E) scatter diagrams and (F) histograms. Percentage represents the proportion of the number of C11-BODIPY staining positive cells compared with the total number of cells. (G) Protein expression levels of GPX4 and SLC7A11 and (H) GPX activity in H/R cells. Data are presented as the mean ± SD (n=3). GPX, glutathione peroxidase; SLC7A11, solute carrier family 7-member 11; H/R, hypoxia/reoxygenation; RGS12, regulator of G-protein signaling 12; sh, short hairpin RNA; NC, negative control; prot, protein.

Downregulation of RGS12 activates the Nrf2 pathway

Nrf2 serves a crucial role in the regulation of ferroptosis and oxidative stress (37). Therefore, the present study investigated the effects of RGS12 on Nrf2 expression. Upon the induction of H/R, the protein expression levels of both nuclear and total Nrf2 decreased compared with those in the control cells (Fig. 4A). Conversely, the silencing of RGS12 led to a significant increase in Nrf2 protein expression levels. Immunofluorescence analysis demonstrated a similar trend, indicating that the low expression level of RGS12 induced the expression of Nrf2 in cells subjected to H/R (Fig. 4B).

Figure 4 RGS12 silencing activates the Nrf2 signaling pathway. (A) Protein expression levels of intranuclear and total Nrf2 and (B) the cellular location of Nrf2 in H/R cells (magnification, ×400). n=3. Nrf2, nuclear factor erythroid 2-related factor 2; H/R, hypoxia/reoxygenation; RGS12, regulator of G-protein signaling 12; sh, short hairpin RNA; NC, negative control.

Overexpression of RGS12 exacerbates MIRI in mice

In the aforementioned experiments, it was demonstrated that the silencing of RGS12 mitigated H/R-injury in HL-1 cells. Subsequently, overexpression of RGS12 was induced in mice to further elucidate its potential role in MIRI. The LVEF and LVFS significantly decreased in the mice with RGS12 overexpression, compared with the control group (Fig. 5A). In addition, compared with the MIRI + Ad-EV group, the increase in TnT serum levels of the MIRI + Ad-RGS12 group also indicated the damage inflicted on cardiac function (Fig. S2). In the mice with RGS12 overexpression, a significant increase in the infarct area of the myocardial tissue (Fig. 5B), accompanied by pronounced leukocyte infiltration in the myocardial tissue (Fig. 5C) and the significantly increased activity of myocardial enzymes, such as LDH, CK and AST (Fig. 5D-F), were observed compared with the control group. The expression of RGS12 was examined in the myocardial tissue and it was demonstrated that the protein expression level of RGS12 was successfully increased in the RGS12-overexpressing mice compared with that in the control group (Fig. 5G). Furthermore, oxidative stress levels were significantly increased in the mice in which RGS12 was overexpressed (Fig. 5H), resulting in significantly increased MDA levels and decreased SOD activity compared with the control group (Fig. 5I and J). The overexpression of RGS12 in mice exacerbated ferroptosis, as evidenced by the significantly elevated levels of Fe2+ and the reduced protein expression level of GPX4 (Fig. 5K and L). Additionally, the upregulation of RGS12 inhibited the activation of the Nrf2 pathway (Fig. 5L).

Figure 5 RGS12 upregulation exacerbates MIRI in mice. (A) Echocardiogram, LVEF and LVFS of mice with MIRI. (B) Representative images of 2,3,5-triphenyl tetrazolium chloride staining and quantification of the infarct area in myocardial tissues. (C) Representative images of hematoxylin and eosin staining of myocardial tissues (magnification, ×200). The activity of (D) LDH, (E) CK and (F) AST in serum. (G) RGS12 protein expression levels in myocardial tissues. (H) ROS level (magnification, ×400), (I) MDA content, (J) SOD activity and (K) Fe2+ content. (L) Protein expression levels of GPX4, SLC7A11, intranuclear Nrf2 and total Nrf2 in myocardial tissues. Data are presented as the mean ± SD (n=6). Ad, adenovirus; Ad-EV, Ad-empty vectors; AST, aspartate aminotransferase; GPX4, glutathione peroxidase 4; SLC7A11, solute carrier family 7 member 11; MIRI, myocardial ischemia/reperfusion injury; LVEF, left ventricular ejection fraction; LVFS, left ventricular fractional shortening; LDH, lactate dehydrogenase; CK, creatine kinase; MDA, malondialdehyde; SOD, superoxide dismutase; RGS12, regulator of G-protein signaling 12; ROS, reactive oxygen species; nrf2Nrf2, nuclear factor erythroid 2-related factor 2.

RGS12 attenuates the protective effect of PHC on oxidative stress and ferroptosis in cells subjected to H/R

As aforementioned, in a preliminary experiment, the present study identified the effect of RGS12 on MIRI. Subsequently, the current study aimed to explore whether RGS12 mediates the protective effects of PHC on the heart. PHC was shown to exert a positive effect on the survival of HL-1 cells under normal conditions, with cell viability significantly increasing as the concentration of PHC treatment increased to 2.5 and 5 μg/ml, compared with the viability of control cells and those treated with 1 μg/ml PHC (Fig. 6A). Upon the induction of H/R, the beneficial effects of PHC on cell survival became more pronounced, with cell viability significantly increasing as the PHC concentration increased, peaking at 5 μg/ml (Fig. 6B). Conversely, the expression level of RGS12 decreased as the PHC concentration increased (Fig. 6C). A concentration of 5 μg/ml PHC was selected to be used in subsequent experiments, resulting in a significant downregulation of RGS12 expression in HL-1 cells. To explore the role of RGS12 in the effect of PHC against H/R damage, RGS12 was overexpressed in HL-1 cells (Fig. S3A). The elevated levels of RGS12 led to a significantly decreased cell survival rate and negated the beneficial effects of PHC treatment in H/R-induced HL-1 cells (Fig. 6D). Furthermore, PHC treatment reduced ROS production (Fig. 6E), MDA level (Fig. 6F) and SOD activity (Fig. 6G) in H/R-induced cells. PHC treatment significantly reduced the production of lipid ROS (Fig. 6H). Additionally, PHC treatment reduced the number of cell points within the gate (Fig. 6I) and a reduction in the number of cells with high fluorescence intensity (Fig. 6J). PHC treatment increased the protein expression levels of GPX4 and SLC7A11 in cells subjected to H/R (Fig. 6K) and promoted the activation of the Nrf2 signaling pathway (Fig. 6L). The overexpression of RGS12 reversed the protective effects of PHC. In addition, in cells overexpressing RGS12, increased concentrations of PHC treatment caused increased cell viability (Fig. S3B), decreased MDA levels (Fig. S3C) and increased SOD activity (Fig. S3D). This suggested that PHC prevented MIRI through inhibiting RGS12 in a concentration-dependent manner.

Figure 6 RGS12 attenuates the protective effect of PHC on oxidative stress and ferroptosis in H/R cells. (A) Survival rate of HL-1 cells treated with PHC. (B) Survival rate of HL-1 cells treated with H/R and PHC. (C) RGS12 expression levels in H/R cells induced with PHC. (D) Survival rate of HL-1 cells. (E) ROS levels (magnification, ×100), (F) MDA content, and (G) SOD activity in H/R cells. Lipid reactive oxygen species levels were analyzed using flow cytometry (H), and the result was shown by scatter diagram (I) and histogram. (J,K) Protein expression levels of GPX4 and SLC7A11 and (L) the cellular location of Nrf2 in H/R cells (magnification, ×400). Data are presented as the mean ± SD (n=3). Ad, adenovirus; Ad-EV, Ad-empty vectors; PHC, penehyclidine hydrochloride; RGS12, regulator of G-protein signaling 12; ROS, reactive oxygen species; H/R, hypoxia/reoxygenation; MDA, malondialdehyde; SOD, superoxide dismutase; GPX4, glutathione peroxidase 4; SLC7A11, solution carrier family 7 member 11; Nrf2, nuclear factor erythroid 2-related factor 2; prot, protein.

RGS12 attenuates the protective effect of PHC on MIRI

Administration of PHC resulted in the remission of MIRI in mice in vivo. PHC treatment led to significant improvements in LVEF and LVFS in mice with MIRI (Fig. 7A). Compared with MIRI + Ad-RGS12 group, the TnT level of MIRI + PHC + Ad-RGS12 group was reduced, indicating that PHC administration improved cardiac function (Fig. S2). Treatment with PHC relieved the alleviated MIRI and ameliorated the damaged myocardial tissue morphology (Fig. 7B and 7C), as myocardial fiber tissue structure was clear and inflammatory cell infiltration was decreased. Furthermore, PHC treatment significantly decreased the activity of myocardial enzymes compared with those in the control group, including LDH (Fig. 7D), CK (Fig. 7E) and AST (Fig. 7F). Notably, the RGS12 protein expression levels were lower in the myocardial tissue of all PHC-treated mice compared with the mice with MIRI, apart from those in the mice in which RGS12 was overexpressed (Fig. 7G). PHC administration reduced ROS production (Fig. 7H) and inhibited oxidative stress-induced injury, including increased MDA level (Fig. 7I) and decreased SOD activity (Fig. 7J). Ferroptosis in mice with MIRI was also alleviated by PHC administration, which was evidenced by reduced Fe2+ (Fig. 7K) and enhanced expression of GPX4 (Fig. 7L). Moreover, treatment with PHC induced the activation of the Nrf2 pathway in the model of MIRI (Fig. 7L). This was also observed in the MIRI + PHC + Ad-EV group. However, the overexpression of RGS12 abrogated the effects of PHC treatment. These findings suggested that RGS12 attenuated the protective effects of PHC in the myocardial tissue of mice with MIRI.

Figure 7 RGS12 attenuates the protective effect of PHC on a MIRI mouse model. (A) Echocardiogram, LVEF and LVFS of mice with MIRI. (B) Representative images of 2,3,5-triphenyl tetrazolium chloride staining and quantification of the infarct area in myocardial tissues. (C) Representative images of hematoxylin and eosin staining in myocardial tissues (magnification, ×200). Black boxes represent areas with significant pathological changes. The activity of (D) LDH, (E) CK and (F) AST in serum. (G) RGS12 protein expression levels in myocardial tissues. (H) ROS levels (magnification, ×400), (I) MDA content, (J) SOD activity and (K) Fe2+ content in myocardial tissues. (L) Protein expression levels of GPX4 and intranuclear Nrf2. The data are presented as the mean ± SD (n=6). RGS12, regulator of G-protein signaling 12; PHC, penehyclidine hydrochloride; MIRI, myocardial ischemia/reperfusion injury; LVEF, left ventricular ejection fraction; LVFS, left ventricular fractional shortening; LDH, lactate dehydrogenase; CK, creatine kinase; AST, aspartate aminotransferase; ROS, reactive oxygen species; MDA, malondialdehyde; SOD, superoxide dismutase; GPX4, glutathione peroxidase 4; prot, protein; Nrf2, nuclear factor erythroid 2-related factor 2.

Discussion

The present study observed a high expression level of RGS12 in a mouse model of MIRI. The mechanism of action of RGS12 in PHC pretreatment was investigated through cell experiments and verified in a mouse model. These results suggested that the silencing of RGS12 alleviated H/R-induced damage to myocardial cells, including reducing oxidative stress and lipid peroxidation, thereby preventing ferroptosis, which may involve the activation of the Nrf2 pathway. The overexpression of RGS12 led to the opposite results in vivo and in vitro. Preconditioning with PHC relied on the RGS12-mediated regulatory mechanism to protect the heart from MIRI in a concentration-dependent manner.

Oxidative stress is a key physiological process involved in MIRI (38). The present study demonstrated that RGS12 promoted oxidative stress in the MIRI process by inhibiting the Nrf2 pathway, a typical antioxidant pathway. The regulation of the Nrf2 pathway by RGS12 may involve a direct effect. For instance, it has been demonstrated that RGS12 promotes the degradation of the Nrf2 protein in RAW 264.7 cells (24). In addition, RGS12 may indirectly regulate the Nrf2 pathway as, for example, it has been reported that targeting RGS12 leads to a decrease in the expression of Kelch-like ECH-associated protein 1, an upstream regulatory molecule of Nrf2, thereby activating the Nrf2 pathway (23,39). Therefore, based on the aforementioned research findings, the effects of RGS12 on the Nrf2 pathway in MIRI may be either direct or indirect, both of which support the conclusions reported herein. In addition to the Nrf2 pathway, RGS12 may be involved in the regulation of oxidative stress by affecting other pathways. RGS12 activates the classical NF-κB inflammatory pathway and contributes to the onset of inflammatory arthritis (40). Inflammation and oxidative stress are interrelated processes that mutually promote one another, potentially due to crosstalk between the NF-κB and Nrf2 pathways (41). Thus, RGS12 may indirectly promote oxidative stress by activating the NF-κB pathway.

Ferroptosis is an iron-dependent mechanism of cell death closely related to oxidative stress. It is characterized by the accumulation of Fe2+ and lipid peroxidation (42). The regulatory effects of RGS12 on ferroptosis remain unclear; however, targeting of the Nrf2 pathway by RGS12 may be a key mechanism involved in the promotion of ferroptosis. Nrf2 is a crucial transcriptional regulator of anti-ferroptosis genes and its target genes participate in the ferroptosis cascade reaction (14). The activation of the Nrf2 pathway mediates the inhibition of ferroptosis and exerts protective effects on the heart, which supports the conclusion obtained in the present study (43). Furthermore, the regulation of ferroptosis by RGS12 may involve other pathways, such as the MAPK pathway. It has been reported that RGS12 activates the MAPK pathway in pathological cardiac hypertrophy (22). The activation of the MAPK pathway may mediate ferroptosis in rat myocardial cells (44). Therefore, RGS12 may enhance ferroptosis by activating the MAPK pathway, indicating the complex mechanism by which RGS12 regulates ferroptosis.

The present study demonstrated that RGS12 promoted oxidative stress and aggravated MIRI by inhibiting the Nrf2 pathway in myocardial cells. The Nrf2 signaling pathway is a classical pathway associated with oxidative stress and ferroptosis. Previous studies have demonstrated the therapeutic potential of the Nrf2 pathway in MIRI (15,43,45). The present study showed the regulatory effects of RGS12 on this pathway, which further highlighted the influence of RGS12 on the MIRI process. However, RGS12 may affect oxidative stress and ferroptosis through various factors. It has been reported that RGS12 is an activator of the NF-κB and MAPK pathways (22,40), which are well-established inflammatory signaling pathways. The activation of these pathways stimulates the production of inflammatory factors and amplifies the inflammatory response, while simultaneously decreasing the expression levels of certain anti-ferroptosis-related proteins, such as heme oxygenase 1, ultimately leading to ferroptosis (46). Targeting the NF-κB/MAPK pathway effectively inhibits ferroptosis in osteoarthritis (47). In addition, the activation of the NF-κB and MAPK pathways is a marker of oxidative stress in peripheral blood mononuclear cell and endothelial cells (48,49) and inhibiting the activation of the NF-κB and MAPK pathways alleviates oxidative stress in bronchial epithelial cells and nerve cells (50,51). Therefore, the promotion of oxidative stress and ferroptosis by RGS12 may involve inflammatory processes, such as the activation of the NF-κB and MAPK pathways.

The present study primarily investigated the regulatory mechanisms involving Nrf2-mediated ferroptosis in myocardial cells, which are the predominant cell type in the heart and are essential for its contractile function (52). Myocardial cell death ultimately results in the structural and functional impairment of the heart tissue. Preventing MIRI-induced myocardial cell damage effectively improves cardiac function (53). Oxidative stress leads to the ferroptosis and apoptosis of myocardial cells, which is a typical manifestation of MIRI at the cellular level (54). Therefore, the present study was conducted using the mouse myocardial cell line, HL-1. However, the regulatory pattern of RGS12 may differ between different types of cells. Macrophages are key immune cells in MIRI, and M1 and M2 macrophage polarization mediates the enhancement and decrease of MIRI-induced inflammation, respectively (55). RGS12 has been reported to induce M1 macrophage polarization (56). IL-6, IL-1β and TNF-α secreted by M1 macrophages enhance the occurrence of ferroptosis (57), and the expression of RGS12 is increased and the NF-κB pathway is activated in macrophages under inflammatory conditions (58,59). This suggests that RGS12-induced M1 macrophage polarization may mediate ferroptosis and the activation of the NF-κB pathway.

The therapeutic effects of PHC on ischemia/reperfusion injury have been previously well-established in animal models and this effect on MIRI is dose-dependent (33,60). According to previous reports (33,60), the present study adopted an optimal dose of 1 mg/kg PHC, which was shown to be effective in alleviating MIRI in the mouse model. Moreover, it was demonstrated that PHC prevented MIRI through the inhibition of RGS12 in a dose-dependent manner using rescue experiments. The present study demonstrated the molecular mechanism by which PHC attenuates MIRI; therefore, it could be suggested that targeting RGS12 may be a promising approach for preventing MIRI by alleviating oxidative stress and ferroptosis. The present study also provided insights into the pharmacological effects of the treatment of MIRI using PHC and may lead to the development of novel strategies for the gene therapy of MIRI in the future.

In conclusion, the present study demonstrated that the silencing of RGS12 mediated the protective effects of PHC by activating the Nrf2 pathway in MIRI, which contributed to the reduction of oxidative stress and the inhibition of ferroptosis (Fig. 8). This mechanism could potentially provide new insights into the prevention and treatment of MIRI.

Figure 8 Predicted molecular mechanism of RGS12 in MIRI. MIRI, myocardial ischemia/reperfusion injury; RGS12, regulator of G-protein signaling 12; Nrf2, nuclear factor erythroid 2-related factor 2; SLC7A11, solute carrier family 7 member 11; ROS, reactive oxygen species; MDA, malondialdehyde; GPX4, glutathione peroxidase 4. The data are presented as the mean ± SD.

Supplementary Data

Availability of data and materials

The data generated in the present study may be requested from the corresponding author.

Authors' contributions

CZ was responsible for the conceptualization, methodology conducting the experiment, data analysis, and drafting of the manuscript. YW was responsible for conducting the experiments, data analysis and writing, reviewing and editing of the manuscript. YZ was responsible for conducting the experiment, data analysis, data visualization and obtaining resources. JF was responsible for the use of software. YW and JF confirm the authenticity of all the raw data. All authors read and approved the final version of the manuscript.

Ethics approval and consent to participate

The present study was carried out in accordance with the requirements in the Guide for the Care and Use of Laboratory Animals of the Ministry of Science and Technology of China. All animal experiments were approved by the Ethics Committee of Hebei North University (approval no. HBNU202306022105; Zhangjiakou, China).

Patient consent for publication

Not applicable.

Competing interests

The authors declare that they have no competing interests.

Abbreviations:

Acknowledgements

Not applicable.

Funding

Funding for the present study was provided by the Hebei Province Natural Science Foundation Project (grant no. H2022405031).

References