Approval for the present study was obtained from the Shaanxi Provincial People's Hospital Ethics Committee (Xi'an, China) and the present study adhered to the National Institutes of Health's Guide for the Care and Use of Laboratory Animals (28). In the present study, a total of 32 male C57BL/6 mice (age, 8 weeks; weight, 18–22 g), obtained from GemPharmatech Co. Ltd., were acclimated for 1 week at a temperature of 25°C, 55% humidity with a 12 h light/dark cycle and ad libitum access to food and water.

The mice were randomly divided into four groups (n=8/group). The two groups received a single intraperitoneal injection of LPS (10 mg/kg; MilliporeSigma) dissolved in PBS, to establish a model of septic cardiomyopathy (29). The remaining groups were pre-treated with 2% AKG (MilliporeSigma) in the drinking water for 9 weeks prior to LPS administration (23). Animals were anesthetized by isoflurane inhalation (4% induction and 2% maintenance), blood samples were collected by cardiac puncture, and then cardiac tissue was collected following cervical dislocation.

Cardiac function was evaluated by transthoracic echocardiography using a Vevo 2100 ultrasound system (VisualSonics, Inc.) equipped with an MS550D transducer, as described previously (25). For imaging, 2D images of the left ventricle (LV) were captured at the papillary muscle level. M-mode tracings, encompassing both the anterior and posterior LV walls, were subsequently recorded. Key parameters, including LV end-diastolic dimension (LVEDD), LV end-systolic dimension (LVESD), ejection fraction and fractional shortening, were measured in a blinded manner. These measurements were derived from an average of five cardiac cycles to ensure accuracy and reliability.

Western blotting was performed as described previously (24). Following protein transfer, polyvinylidene fluoride membranes were blocked using 5% non-fat milk for 1 h at room temperature. This was followed by an overnight incubation at 4°C with primary antibodies against atrial natriuretic protein (ANP; cat. no. ab225844; 1:500; Abcam), brain natriuretic peptide (BNP; cat. no. ab239510; 1:1,000; Abcam), β-major histocompatibility complex (β-MHC; cat. no. ab172967; 1:1,000; Abcam), BCL2 interacting protein 3 (Bnip3; cat. no. ab10433; 1:1,000; Abcam), LC3 (cat. no. 4599S; 1:1,000; Cell Signaling Technology, Inc.), Fis1 (cat. no. ab71498; 1:1,000; Abcam), dynamin-related protein 1 (DRP1; cat. no. 8570S; 1:1,000; Cell Signaling Technology, Inc.), NADH dehydrogenase (ubiquinone) 1α subcomplex subunit 12 (Ndufa12; cat. no. ab192617; 1:4,000; Abcam), NADH:ubiquinone oxidoreductase subunit AB1 (Ndufab1; cat. no. ab181021; 1:1,000; Abcam), mitochondrial NADH-ubiquinone oxidoreductase chain 1 (MT-ND1; cat. no. ab181848; 1:5,000; Abcam), succinate dehydrogenase (SDHA; cat. no. 11998; 1:4,000; Cell Signaling Technology, Inc.), complex IV (COX IV; cat. no. 11242-1-AP; 1:3,000; Proteintech Group, Inc.), oxoglutarate dehydrogenase (OGDH; cat. no. ab137773; 1:5,000; Abcam), pyruvate dehydrogenase (PDH; cat. no. 3205; 1:3,000; Cell Signaling Technology, Inc.), hypoxia inducible factor-1α (HIF-1α; cat. no. 36169; 1:1,000; Cell Signaling Technology, Inc.), NADPH oxidase 2 (NOX2; cat. no. bs-3889R; 1:1,000; BIOSS), NOX4 (cat. no. bs-1091R; 1:1,000; BIOSS), Bax (cat. no. 2772S; 1:3,000; Cell Signaling Technology, Inc.) and Bcl-2 (cat. no. 3498S; 1:1,000; Cell Signaling Technology, Inc.), GAPDH (cat. no. 10494-1-AP; 1:8,000; Proteintech Group, Inc.), α-tubulin (cat. no. 11224-1-AP; 1:4,000; Proteintech Group, Inc.) and HSP90 (cat. no. 11224-1-AP; 1:2,000; Proteintech Group, Inc.). Following incubation with the primary antibody, the membranes were treated with horseradish peroxidase-conjugated secondary antibodies (anti-Mouse secondary antibodies; cat. no. 62-6520; 1:10,000; Thermo Fisher Scientific, Inc. and anti-Rabbit secondary antibodies; cat. no. VJ313046; 1:10,000; Thermo Fisher Scientific, Inc.) for 1 h at room temperature. Signals were visualized using Clarity^™ Western ECL Substrate (Bio-Rad Laboratories, Inc.). Densitometry analysis was performed using ImageJ software (version 4.5.2; National Institutes of Health).

Total RNA was isolated from LV myocardial tissues using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.). From the extracted RNA, 1 µg was reverse-transcribed into cDNA using a RT kit (cat. no. RR047A; Takara Bio, Inc.) at 37°C for 15 min and 85°C for 5 sec. qPCR was performed using TB Green^™ Premix Ex Taq^™ II (Tli RNaseH Plus; cat. no. RR820A; Takara Bio, Inc.) on a Bio-Rad CFX96 Real-time PCR Detection System (Bio-Rad Laboratories, Inc.). The amplification protocol involved an initial denaturation at 95°C for 30 sec, followed by 40 cycles of denaturation at 95°C for 5 sec and annealing/extension at 60°C for 30 sec. Gene expression levels were quantified relative to the housekeeping gene GAPDH for normalization using the 2^−ΔΔCq method (30). The sequences of the primers used for qPCR were: ANP forward, 5′-AAGAACCTGCTAGACCACCTGGAG-3′ and reverse, 5′-TGCTTCCTCAGTCTGCTCACTCAG-3′; BNP forward, 5′-GGAAGTCCTAGCCAGTCTCCAGAG-3′ and reverse, 5′-GCCTTGGTCCTTCAAGAGCTGTC-3′; β-MHC forward, 5′-CAGAACACCAGCCTCATCAACCAG-3′ and reverse, 5′-TTCTCCTCTGCGTTCCTACACTCC-3′; and GAPDH forward, 5′-AGGTCGGTGTGAACGGATTTG-3′ and reverse, 5′-TGTAGACCATGTAGTTGAGGTCA-3′.

TUNEL staining of the LV tissue sections was performed according to the manufacturer's protocol. Briefly, heart tissues were fixed in 4% paraformaldehyde at room temperature overnight. Subsequently, tissues were embedded in paraffin and cut into slices with a thickness of 4 µm. After dewaxing and hydration, slices were stained with TUNEL reagent (cat. no. C1088; Beyotime Institute of Biotechnology) at room temperature for 1 h. Then, antifade mounting medium with DAPI (cat. no. P0131; Beyotime Institute of Biotechnology) solution was incubated at room temperature for 10 min for nuclear counterstaining. TUNEL-stained sections were imaged using an Olympus DP-72 fluorescence microscope (Olympus Corporation), focusing on the identification of apoptotic cells within the myocardium. Images were analyzed using ImageJ. The number of cardiomyocyte nuclei exhibiting green fluorescence (indicating apoptosis) was counted. To ensure objectivity, all analyses were carried out under double-blinded conditions. For each histological section, five random visual fields were selected for measurement. The average value from these fields was then used for statistical analysis.

TEM was carried out as described previously (24). Fresh apical myocardial tissue from the LV was first fixed using 2.5% glutaraldehyde at 4°C for 2 h. After fixation, a graded series of ethanol/acetone solutions was employed for dehydration, with the final solution being absolute acetone. The dehydrated samples were then infiltrated with Epon 812 resin. Ultrathin sections of the embedded tissues were prepared using an ultramicrotome (LKB-V/NOVA; Leica Microsystems GmbH) and stained with acidified uranyl acetate for enhanced contrast. Observation of the prepared samples was carried out using a Hitachi Model H-7650 TEM (Hitachi, Ltd.). ImageJ was used for quantitative analysis of the TEM images. Specifically, the count of mitochondria was determined from five images at a magnification of ×10,000 per LV sample. Additionally, the proportion of mitochondria exhibiting structural impairments, such as incomplete outer membranes or dissolved cristae, was assessed from another set of five images at a magnification of ×30,000 per LV sample. The count of aberrant mitochondria is presented as a percentage of the total mitochondrial count.

ROS levels in myocardial tissues were quantified using two distinct methods as described previously (24). Initially, O²⁻ content was assessed using 5-µm frozen myocardial sections. These sections were incubated with dihydroethidium (DHE; 5 µM) for 1 h at 37°C. Fluorescence images were captured at a magnification of ×400 (five fields per heart) using an Olympus DP-72 fluorescence microscope (Olympus Corporation), with excitation and emission wavelengths set at 488 and 610 nm, respectively. In the second method, cardiomyocytes were isolated via enzyme digestion using the Langendorff perfusion system. Briefly, the aorta was retrogradely perfused with collagenase II (cat. no. LS004177; Worthington Biochemical Corporation) at room temperature for 15 min to digest the heart. Subsequently, the heart was minced into small pieces and centrifuged at 1,400 × g for 3 min at room temperature. Finally, the isolated cardiomyocytes were transferred onto the cell culture dish. The isolated cells were then incubated with chloromethyl derivative CM-H₂DCFDA (DCF; 5 µM) for 30 min at 37°C. Fluorescence imaging was performed using a Leica TCS SP8 STED 3X confocal microscope (Leica Microsystems GmbH), with a ×40 1.3 NA oil immersion objective lens, with excitation at 488 nm and emission at 525 nm, using standardized scanning parameters. The intensity of DCF fluorescence was quantified using ImageJ, with an average of 80 cells analyzed per heart.

Briefly, the heart was fixed in 4% paraformaldehyde at room temperature overnight. Subsequently, it was embedded in paraffin and sliced into 4 µm slices. After deparaffinization and hydration, heart sections were stained with hematoxylin for 5 min and eosin for 1 min at room temperature using the H&E Staining kit (cat. no. 0105; Beyotime Institute of Biotechnology). The fluorescence images were captured using an Olympus DP-72 fluorescence microscope (Olympus Corporation) from 3–5 random fields.

Fresh LV tissue and plasma was harvested to determine the content of ATP, lactate and MDA using commercial kits. ATP kits (cat. no. A095-2-1; Nanjing Jiancheng Bioengineering Institute), lactate kits (cat. no. E-BC-Ko44-M; Wuhan Elabscience Biotechnology Co., Ltd.) and MDA kits (cat. no. R21869; Shanghai Yuanye Biotechnology Co., Ltd.) were carried out according to the manufacturer's protocol as described previously (25).

Plasma levels of AKG were quantified using a commercial kit (cat. no. G0861W; Grace Biotech Co., Ltd.) according to the manufacturer's protocol.

A total of 24 Neonatal Sprague-Dawley rats (age, 1–2 days; weight, 6–7 g) were purchased from the Laboratory Animal Center of Xi'an Jiaotong University (Xi'an, China), and humanely euthanized via cervical dislocation. Subsequently, their hearts were excised and subjected to enzymatic digestion using 1 ml 0.2% collagenase II for 5–6 min at 37°C for six cycles. The digested tissue was centrifuged at 2,200 × g for 5 min at room temperature and the resultant cardiomyocytes were resuspended in F12 medium (Thermo Fisher Scientific, Inc.) supplemented with 15% FBS (Thermo Fisher Scientific, Inc.). This suspension was incubated for 60 min at 37°C to facilitate differential adhesion, allowing for the separation of cardiomyocytes from non-myocyte cells. To further purify the culture, the cardiomyocytes were then treated with 5-bromo-2-deoxyuridine, an agent used to inhibit the proliferation of non-cardiomyocyte cells (31). After a 48-h incubation period at 37°C, the NRVMs given the following treatments (Control, 2 mM AKG, 0.5 µg/ml LPS or a combination of 2 mM AKG and 0.5 µg/ml LPS) were used for TUNEL staining, western blotting and Seahorse assays. The control group was treated with F12 medium containing 10% FBS. These treatments were administered in F12 medium containing 10% FBS for a duration of 24 h at 37°C as previously described (7,23).

The respiratory capacity of mitochondria in NRVMs was evaluated using the Agilent Seahorse XF24 Extracellular Flux Analyzer (Agilent Technologies, Inc.). NRVMs were initially seeded onto Seahorse XF24 cell culture microplates and subjected to the aforementioned treatments. The Seahorse XF sensor cartridge was prepared by hydrating its probe plate with calibration solution in a CO₂-free incubator maintained at 37°C overnight. Subsequently, during the assay, specific mitochondrial inhibitors were sequentially added to the wells: Oligomycin A (1.5 µM; cat. no. 103672-100; Agilent) to inhibit ATP synthase, trifluoromethoxy carbonylcyanide phenylhydrazone (1 µM; cat. no. 103672-100; Agilent) to uncouple the proton gradient and antimycin A (0.5 µM; cat. no. 103672-100; Agilent) to inhibit the mitochondrial respiratory chain. The oxygen consumption rate, a key indicator of mitochondrial respiration, was measured following these additions, according to the manufacturer's protocol.

Data are presented as the mean ± standard error of the mean (n=6-8 repeats/group). To determine the normality and homogeneity of variance of the data, Shapiro-Wilk's and Levene's tests were respectively employed. For comparisons among multiple groups, one-way ANOVA was carried out, followed by Tukey's post hoc test. All statistical analyses were carried out using GraphPad Prism (version 9.0; Dotmatics). P<0.05 was considered to indicate a statistically significant difference.

In the present study, the cardiotoxic effects of LPS in wild-type mice were investigated, focusing on LPS-induced cardiac dysfunction and LV remodeling. Echocardiographic assessments were carried out following exposure to vehicle control, AKG, LPS or a combination of AKG and LPS. Notably, the LV ejection fraction (LVEF) and LV fractional shortening (LVFS) in the LPS group were reduced by 33 and 32%, respectively, compared with those in the control group. Concurrently, there was a 40 and 60% increase in LVEDD and LVESD (Fig. 1A and B). These findings indicated cardiac dysfunction characteristic of septic cardiomyopathy induced by LPS.

The cardioprotective effect of AKG supplementation was evident. AKG treatment led to a 22 and 21% improvement in LVEF and LVFS, along with a 20 and 32% reduction in LVEDD and LVESD, respectively, compared with the LPS group (Fig. 1A and B). There were no significant differences in the cardiac function indices between the AKG group and the control group. This suggested that AKG supplementation did not affect cardiac function when administered alone (Fig. 1B).

Cardiomyocyte remodeling was evaluated by assessing the protein and transcriptional levels of ANP, BNP and β-MHC in the LV myocardium. LPS-treated mice exhibited a notable increase in the mRNA and protein expression levels of these markers, indicative of cardiomyocyte remodeling. However, AKG supplementation may partially mitigate the LPS-induced remodeling (Fig. 1C).

As shown in Fig. S1, the circulating AKG concentration was decreased in the LPS group compared with that in the control group, but the difference was not significant. Although the concentration of circulating AKG increased following AKG supplementation in the LPS group, the differences were not statistically significant.

Given that mitochondria constitute ~35% of the volume of cardiomyocytes and serve a key role in cellular function (11), a detailed analysis of mitochondrial ultrastructure was performed in heart tissues from mice in the different groups using TEM. LPS treatment induced notable mitochondrial morphological damage, evidenced by increased proportions of mitochondria with incomplete outer membranes or dissolved cristae, compared with those in hearts from the control group (Fig. 2A and B). Additionally, quantitative TEM analysis revealed a reduction in the mitochondrial number. This was further corroborated by assessing the protein levels of Ndufa12 and Ndufab1, which serve as mitochondrial markers (Fig. 2A-C).

Mitochondrial quality control, encompassing processes such as biosynthesis, fusion, fission and mitophagy, is essential for maintaining mitochondrial integrity (11). In hearts from the group subjected to LPS, there was a marked decrease in the expression levels of proteins indicative of mitochondrial fission (DRP1 and Fis1) and mitophagy (LC3 II/I and Bnip3) compared with the control group (Fig. 2C). These findings suggested impaired mitochondrial quality control mechanisms following LPS exposure.

By contrast, AKG treatment mitigated these adverse effects. It restored mitochondrial morphology by reducing the fraction of mitochondria with structural impairments, and increased the overall mitochondrial count, as evidenced by the increased expression levels of mitochondrial content markers (Ndufa12; Fig. 2A-C). Furthermore, AKG positively influenced mitochondrial quality control, demonstrated by the upregulated expression levels of proteins associated with mitochondrial fission and mitophagy (Fig. 2C). This suggested that AKG supplementation counteracted the mitochondrial dysfunction observed in LPS-induced cardiomyopathy.

Generation of ATP through OXPHOS is the primary role of mitochondria and this was further investigated in the context of LPS-induced mitochondrial damage (13). Specifically, the changes in mitochondrial energy metabolism in mouse hearts treated with LPS were assessed.

Initially, immunoblotting was used to analyze heart samples for the presence of specific marker proteins. These included proteins associated with mitochondrial respiratory electron transport chain complexes I, II and IV (MT-ND1, SDHA and COX IV), and key enzymes involved in acetyl-CoA production and the TCA cycle (PDH and OGDH). A notable decrease in the expression levels of these proteins was observed, indicating impaired mitochondrial aerobic oxidation. Conversely, the protein levels of HIF-1α, indicative of anaerobic glycolysis, were revealed to be elevated following LPS treatment (Fig. 3A and B).

Furthermore, to directly assess mitochondrial functionality, the levels of ATP and lactate were measured using an ELISA. Analysis revealed a significant reduction in ATP levels in the hearts of mice treated with LPS, accompanied by an increase in lactate content in LV tissues and plasma (Fig. 3C). AKG supplementation led to a partial increase in the abundance of proteins associated with aerobic oxidation and ATP production, while concurrently decreasing the levels of HIF-1α and lactate (Fig. 3A-C). This suggested that AKG reversed the alterations in mitochondrial energy metabolism induced by LPS treatment.

Given the key role of mitochondria in the regulation of ROS generation and apoptosis (15), the impact of AKG on these parameters in LPS-induced cardiomyopathy was assessed.

To assess oxidative stress, DCF and DHE staining were carried out to determine ROS levels in myocardial tissue (Fig. 4A and B). Additionally, the expression levels of key enzymes involved in ROS production, namely NOX2 and NOX4, were examined (Fig. 4D). MDA, a biomarker of lipid peroxidation, was also measured to provide further insights into oxidative stress (Fig. 4C). Compared with the control group, a significant increase in ROS generation was observed in the LPS-treated group. This elevation in ROS levels was mitigated in the LPS + AKG group, suggesting that AKG reduced oxidative stress in LPS-induced septic cardiomyopathy.

To evaluate apoptosis, TUNEL staining was performed and the protein expression levels of Bax and Bcl-2 in myocardial tissue were measured. Enhanced apoptosis was evident in the LPS group compared with both the control and AKG groups (Fig. 4A, B and E). However, AKG administration in the LPS-treated group led to a reduction in apoptosis. These findings indicated that AKG supplementation attenuated both oxidative stress and apoptosis induced by LPS in the context of septic cardiomyopathy.

To examine the morphology of the myocardium, H&E staining was performed. In the LPS group, the myocardium showed a disorderly arrangement of myocytes, karyolysis and an increased presence of inflammatory cells. However, AKG administration improved the abnormal histological structure (Fig. 4A).

The effects of AKG on NRVMs subjected to LPS treatment were explored (Fig. 5). The impact on apoptosis was initially assessed using TUNEL staining and measurement of Bcl-2 protein levels. Compared with the control group, a seven-fold increase in the optical density of TUNEL-positive cells was observed in the LPS group, coupled with a significant decrease in Bcl-2 protein levels, indicating increased apoptosis due to LPS exposure. However, simultaneous administration of AKG significantly mitigated LPS-induced apoptosis in NRVMs (Fig. 5A, D and E).

Additionally, the expression levels of marker proteins integral to mitochondrial aerobic oxidation, including MT-ND1, SDHA, COX IV, OGDH and PDH, were explored. LPS treatment resulted in a significant reduction in the expression levels of these proteins. By contrast, AKG administration alleviated this reduction, suggesting its efficacy in improving mitochondrial aerobic oxidation when this is compromised by LPS (Fig. 5B and C).

Immunoblotting analysis revealed alterations in mitochondrial turnover induced by LPS, evidenced by decreased expression levels of proteins associated with mitophagy (Bnip3) and fission (DRP1), along with a reduction in the mitochondrial content marker Ndufa12. Notably, AKG supplementation significantly counteracted these effects, suggesting its potential to restore mitochondrial turnover disrupted by LPS treatment (Fig. 5D and E). These findings collectively underscore the therapeutic potential of AKG in mitigating mitochondrial dysfunction and apoptosis in LPS-induced cardiomyopathy.

Whether AKG directly enhanced mitochondrial function was subsequently assessed. A Seahorse XF mitochondrial stress test analyzer was used to evaluate the mitochondrial respiratory capacity in NRVMs (Fig. 6A).

Key parameters of mitochondrial respiratory function, including basal and maximal respiratory capacity (Fig. 6C and D), ATP production (Fig. 6E) and spare respiratory capacity (Fig. 6G), were assessed. These parameters were found to be significantly compromised in NRVMs treated with LPS, compared with those in the control group. Notably, when AKG was administered in conjunction with LPS, an improvement in mitochondrial respiration was observed.

This enhancement suggested that AKG supplementation effectively counteracted the mitochondrial dysfunction induced by LPS in NRVMs. In the present study, no significant differences in non-mitochondrial respiration in NRVMs were observed.