Empagliflozin (EMPA, Cat: HY-108682) and Compound C (Dorsomorphin; Cat HY-13418A) were purchased from MedChemExpress (Monmouth Junction, NJ, United States). Other chemicals, including bovine serum albumin (BSA; 2207008), PA (P5585), and insulin (I9278) were purchased from Sigma-Aldrich (Burlington, MA, United States). 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-amino]-2-deoxyglucose (2-NBDG; N13195) was obtained from Thermo Fisher Scientific (Waltham, MA, United States). Cell Death Detection enzyme-linked immunosorbent assay (ELISA^plus) kit was purchased from Roche Applied Science (Roche Applied Science, Mannheim, Germany). AMP colorimetric assay kit was purchased from Biovision (Cat; K229, Milpitas, CA, United States). Antibodies targeting phospho-AKT (#9271), T-AKT (#9272), phospho-GSK3 α/β (#9331), T-GSK3β (#9315), phospho-JNK (#9251), T-JNK (#9252), phospho-eIF2A (#9721), T-eIF2A (#9722), CHOP (#2895), phospho-NF-κB p65 (#3033), NF-κB p65 (#3034), phospho-p38 (#9211), cleaved caspase-3(#9661), phospho-AMP (#2531), and T-AMP (#2532) were obtained from Cell Signaling Technology (Beverly, MA, United States). Anti-β-actin (A300-491A) antibody was purchased from Bethyl Laboratories (Montgomery, TX, United States).

PA/bovine serum albumin (BSA) complex was prepared by soaping PA with sodium hydroxide and mixing it with BSA. Briefly, a 20 mM solution of PA in 0.01 M NaOH was incubated at 70°C for 30 min and the fatty acid soaps were then complexed with 5% fatty acid-free BSA in phosphate-buffered saline (PBS) at a 1:3 volume ratio. The complexed fatty acids consisted of 5 mM PA and 3.75% BSA. The pA/BSA conjugates were administered to cultured cells at the indicated concentration of PA.

Rat cardiomyoblast cell line H9c2 was obtained from American Type Culture Collection (ATCC; Manassas, VA, United States) and maintained in low-glucose (1 g/L) Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum and antibiotics (100 IU/mL of penicillin and 10 μg/mL of streptomycin) at 37°C in a humidified atmosphere of 5% CO₂. Rat tissue lysates including liver (RT129), pancreas (RT133), muscle (RT135), heart (RT127), and kidney (RT128) were purchased from Cell Biologics Inc. (Chicago, IL, United States).

Cells were lysed with cell lysis buffer [150 mM NaCl, 1% NP-40, 0.5% deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 50 mM Tris–Cl, pH 7.5, proteinase inhibitor and protease inhibitor cocktail (Roche Applied Science, Mannheim, Germany)] in 4°C for 30 min. Whole proteins were extracted by differential centrifugation (10,000 g, 10 min), and protein concentrations in lysates were determined using a Bio-Rad Protein Assay Kit (Bio-Rad, Hercules, CA, United States). An equal volume of 2× SDS sample buffer (125 mM Tris–Cl, pH 6.8, 4% SDS, 4% 2-mercaptoethanol, and 20% glycerol) was added to the cell lysates, and equivalent amounts of protein (10 μg) were separated by SDS-PAGE and transferred to polyvinylidene fluoride membranes (Millipore, Bedford, MA, United States). After blocking the membranes with 5% skim milk for 30 min, target proteins were immunoblotted to primary antibodies, followed by secondary antibodies (horseradish peroxidase-conjugated anti-mouse IgG or anti-rabbit IgG). Immunoreactive bands were visualized by enhanced chemiluminescence (Amersham Pharmacia Biotech, Arlington Height, IL, United States).

Total RNA from cells was extracted with RNAiso Plus reagent (Takara Bio, Shiga, Japan). cDNA was synthesized using the AMV reverse transcriptase and random 9-mers supplied with the TaKaRa RNA PCR Kit (version 3.0; TaKaRa Bio, Shiga, Japan). The primer sets for PCR amplification are listed in Supplementary Table S1. qRT-PCR was performed with SYBR Green (TaKaRa Bio, Shiga, Japan) using a TaKaRa TP-815 instrument. Relative quantities of amplified DNA were analyzed using the software bundled with the TP-815 instrument and normalized to mouse RPL32 mRNA levels.

Heart-derived H9c2 cells were treated with or without PA for 8 h and starved for 4 h. Cells were then incubated in Krebs-Ringer bicarbonate buffer (pH 7.4) containing 2% BSA at 37°C for 30 min and then treated with 500 μM 2-NBDG with or without 100 nM insulin at 37°C for 2 h. Collected cells were lysed with cell lysis buffer and centrifuged at 12,000 rpm for 30 min. The fluorescence intensity of 2-NBDG in the separated supernatant was measured (excitation: 475 nM; emission: 550 nM) using a SpectraMax iD3 Fluorescence microplate reader (Molecular Devices, Sunnyvale, CA, United States).

H9c2 cells were plated into XF24 cell culture microplates and cells were either treated or not treated with drugs, depending on the experimental condition. Following treatment, the H9c2 cells were pre-washed with Krebs-Ringer bicarbonate (KRB) buffer and equilibrated with XF assay medium supplemented with 2.5 mM glucose/50 mM carnitine/0.2 mM PA (for PA OCR) at 37°C in a CO₂-free incubator for 1 h. The OCR of PA as a carbon substrate was measured using an XF24 extracellular analyzer (Seahorse Bioscience, North Billerica, MA, United States).

AMP level was determined using an AMP colorimetric assay kit (Biovision, Milpitas, CA, United States). Briefly, cells were scraped, washed with phosphate-buffered saline (PBS), and homogenized in cold assay buffer supplied by the kit. Supernatant after differential centrifugation (10,000 g, 10 min) was collected and used to measure AMP level. Samples (2–20 ul) were added into a 96-well plate and sample reaction mix supplied by the kit was then added to each test sample. Enzymatic reaction was performed at 37°C for 60 min. Absorbance was measured at 570 nm. AMP level was determined regarding the AMP standard curve.

Cell death was determined by measuring fragmented DNAs using Cell Death Detection enzyme-linked immunosorbent assay (ELISA^plus) kit (Roche Applied Science, Mannheim, Germany). Briefly, cells were lysed with lysis buffer supplied with the kit. After centrifugation (200 g, 10 min), the supernatant was transferred to an anti-streptavidin-coated microplate. Anti-DNA monoclonal antibody conjugated with peroxidase (anti-DNA-POD) and anti-histone-biotin was added. After incubation at 25°C for 90 min, wells were rinsed with incubation buffer (supplied by the kit) three times. Color was developed by adding 2, 20-azino-di-[3-ethylbenzthiazoline sulphonate] (ABTS) substrate solution, followed by incubation with shaking at 250 rpm for 10–20 min. The amount of peroxidase retained in the nucleosome complex was determined by measuring the absorbance value at 405 nm on a microplate reader.

All experiments were repeated at least three times. All data are expressed as the mean ± standard error of the mean (SEM) and were analyzed using GraphPad Prism 9.2.0 (GraphPad Software Inc., San Diego, CA, United States). Statistical analysis was performed via one-way analysis of variance (ANOVA) with the Bonferroni post hoc test. A probability (p) value less than 0.05 (p < 0.05) was considered statistically significant.

To investigate the impact of SGLT2 inhibitors on cardiomyocytes, we first assessed both mRNA and protein expression levels of SGLT2 in different rat tissues. We found that mRNA expression of SGLT2 is present in cardiac tissues, although at levels much lower than in the kidney and pancreas, but higher than in the liver and muscle (Supplementary Figure S1A). Conversely, protein expression levels of SGLT2 were similar across several tissues (Supplementary Figure S1C). Additionally, we observed mRNA and protein expression of SGLT2 in the H9c2 cardiomyocyte cell line and the FaO hepatoma cell line (Supplementary Figures S1B, D). This suggests that the effects of SGLT2 inhibitors on cardiomyocytes extend beyond their role in glycosuria.

It is well-known that factors like hyperlipidemia, hyperglycemia, insulin resistance, high levels of AGEs, and lipotoxicity contribute to diabetic cardiomyocyte injury (Tan et al., 2020). To establish an in vitro system mimicking diabetic cardiomyocytes, H9c2 cells were treated with either high concentrations of glucose or PA, followed by assessments of insulin signaling and apoptotic signaling. In our system, long-term treatment with high glucose did not alter insulin signaling or levels of cleaved caspase-3, a representative apoptotic signal (Supplementary Figures S2A, B). However, PA treatment did result in cardiomyocyte injury, as evidenced by changes in insulin resistance, inflammatory gene expression, and cellular apoptotic signaling. H9c2 cells were treated with various concentrations of PA, followed by assessments of insulin signaling and glucose uptake. PA treatment decreased insulin-induced p-AKT and p-GSK-3α/β expression levels without changing total protein levels of Akt and GSK-3α/β (Figure 1A). Glucose uptake, measured via the 2-NBDG assay, showed that PA administration inhibited insulin-induced 2-NBDG uptake, implicating a decrease in glucose uptake via suppression of p-AKT and p-GSK-3α/β expression (Figure 1B).

Lipotoxicity also negatively impacts cardiomyocyte injury by increasing the expression of proinflammatory cytokines like TNF-α and IL1-β. These cytokines have been implicated in heart failure, affecting the phenotype and function of myocardial cells, contractile function in cardiomyocytes, and inflammatory activation in macrophages, among other roles (Hanna and Frangogiannis, 2020; Kaur et al., 2021). To examine whether PA elevates the expression of inflammatory cytokines, H9c2 cells were treated with 0.2 mM of PA for 8 h, followed by gene expression assessments for TNF-α and IL1-β (Figure 1C). PA treatment significantly increased the gene expression of both cytokines. Additionally, PA elevated levels of cleaved caspase-3 and DNA fragmentation, markers of cellular apoptosis (Figures 1D, E). These results suggest that PA treatment induces cardiomyocyte injury through impaired glucose uptake, upregulated inflammatory gene expression, and increased apoptotic signaling.

Next, we investigated intracellular stress signals such as ER-stress and inflammation signaling under treatment of PA. Salvadó et al. reported that lipotoxicity induces insulin resistance via the activation of ER-stress signals such as p-IRE and p-JNK, which lead to the inhibition of IRS-1 phosphorylation (Salvadó et al., 2015). PA treatment increased ER-stress signals like p-JNK, p-eIF2α, and CHOP (Figure 2A). We also observed that PA administration stimulated inflammatory stress signals such as p-p65 and p-p38 (Figure 2B), which subsequently promoted the expression of proinflammatory cytokines like TNF-α and IL-1β (Figure 1C).

We then sought to examine the effects of the SGLT2 inhibitor, EMPA, on PA-induced diabetic cardiomyocyte injury in H9c2 cells. EMPA treatment ameliorated the PA-induced reduction in p-AKT and p-GSK protein expression levels (Figure 3A), leading to improved 2-NBDG uptake in H9c2 cells (Figure 3B). These findings suggest that EMPA restores insulin-stimulated glucose uptake through upregulation of p-AKT expression, thereby increasing insulin sensitivity in H9c2 cells. EMPA also attenuated PA-induced expression of proinflammatory cytokines such as TNF-α and IL-β (Figure 3C). Moreover, PA-induced cleaved caspase-3 expression and DNA fragmentation were mitigated by EMPA (Figures 3D, E). We next explored whether EMPA could inhibit ER stress markers like p-JNK that affect insulin signaling. Treatment with 1 µM of EMPA reduced PA-induced ER stress signals including p-JNK, p-elF2, and CHOP expression (Figure 4A). EMPA also significantly lowered inflammation signals such as p-p65, which influence the production of proinflammatory cytokines like TNF-α and IL-β (Figure 4B), although it did not affect p-p38 levels. These results suggest that EMPA mitigates PA-induced cardiomyocyte injury, including insulin resistance, proinflammatory cytokine expression, and cellular apoptosis.

Since Figure 3 illustrated that EMPA ameliorates lipotoxic conditions such as insulin resistance and cellular apoptosis, we sought to identify the specific role of EMPA in cellular lipid metabolism. To this end, we assessed CPT1 gene expression, which is associated with lipid beta-oxidation, in the context of PA treatment with or without EMPA. EMPA restored the PA-induced decrease in CPT1 gene expression in a dose-dependent manner (Figure 5A), but not PDH gene expression or PDH activity (Supplementary Figures S3A, B). Intriguingly, EMPA alone also increased CPT1 gene expression (Supplementary Figure S3C). Subsequently, we measured the oxygen consumption rate (OCR), an indicator of lipid metabolism. PA-treated H9c2 cells exhibited a 75% reduction in OCR compared to untreated cells; however, EMPA significantly restored OCR in a dose-dependent manner (Figure 5B).

We hypothesized that EMPA might induce AMPK activation and enhance mitochondrial metabolic activity, thereby increasing FFA oxidation. Interestingly, we found that EMPA elevates AMP levels in H9c2 cells in a dose-dependent manner (Figure 5C). Given that increased AMP levels can activate AMPK, we assessed the levels of activated AMPK and its upstream proteins through immunoblotting with p-AMPK and p-ACC antibodies. As anticipated, EMPA increased p-AMPK and p-ACC levels dose-dependently (Figure 5D). To understand which subunits are involved in AMPK activation by EMPA treatment, we assessed mRNA expression levels of AMPK subunits. As a result, the γ1 subunit of AMPK significantly increased when administered with EMPA (Supplementary Figure S4). These results suggest that EMPA has beneficial effects through stimulating AMP production and subsequently activating AMPK. To validate that the beneficial effect of EMPA is dependent on AMPK activation, we co-treated the cells with compound C, a specific AMPK inhibitor. We observed that EMPA significantly improved 2-NBDG uptake and CPT1 expression in PA-treated H9c2 cells, but co-treatment with compound C notably diminished these beneficial effects (Figures 5E, F). Similarly, EMPA also reduced palmitate-induced levels of cleaved caspase-3; however, co-treatment with compound C negated this positive effect (Figure 5G). Co-treatment with compound C also considerably inhibited EMPA’s protective effects against palmitate-induced DNA fragmentation (Figure 5H). These data support our hypothesis that AMPK plays a crucial role in the protective effects of EMPA against PA-induced injury in H9c2 cells.