Oncometabolite d-2-hydroxyglutarate impairs α-ketoglutarate dehydrogenase and contractile function in rodent heart

Significance

We show that the oncometabolite d-2-hydroxyglutarate (D2-HG) affects cardiac function in the isolated working heart by inhibiting α-KGDH, a key regulatory enzyme of cellular energy metabolism. Analyzing metabolic flux rates by using in vitro and ex vivo approaches in combination with integrative mathematical modeling enabled us to identify the mechanisms by which D2-HG perturbs metabolic flux and induces epigenetic modifications in the heart. The results provide knowledge about malignancy-related changes in enzymatic activity and posttranslational modifications in the context of cardiac remodeling.

Abstract

Hematologic malignancies are frequently associated with cardiac pathologies. Mutations of isocitrate dehydrogenase 1 and 2 (IDH1/2) occur in a subset of acute myeloid leukemia patients, causing metabolic and epigenetic derangements. We have now discovered that altered metabolism in leukemic cells has a profound effect on cardiac metabolism. Combining mathematical modeling and in vivo as well as ex vivo studies, we found that increased amounts of the oncometabolite d-2-hydroxyglutarate (D2-HG), produced by IDH2 mutant leukemic cells, cause contractile dysfunction in the heart. This contractile dysfunction is associated with impaired oxidative decarboxylation of α-ketoglutarate, a redirection of Krebs cycle intermediates, and increased ATP citrate lyase (ACL) activity. Increased availability of D2-HG also leads to altered histone methylation and acetylation in the heart. We propose that D2-HG promotes cardiac dysfunction by impairing α-ketoglutarate dehydrogenase and induces histone modifications in an ACL-dependent manner. Collectively, our results highlight the impact of cancer cell metabolism on function and metabolism of the heart.

Metabolic dysregulation in cancer cells changes the way nutrients are consumed and macromolecules are produced to meet the increased demands for cell growth. Somatic mutations in isocitrate dehydrogenase 1 and 2 (IDH1/2) are common and are described in several cancer types (i.e., gliomas and acute myeloid leukemia). IDH mutations lead to increased production and accumulation of the oncometabolite d-2-hydroxyglutarate (D2-HG) through a neomorphic enzymatic function (1). WT IDH1/2 catalyzes the oxidative decarboxylation of isocitrate to α-ketoglutarate (α-KG), while reducing NADP⁺ to NADPH either in the cytosol and peroxisome (IDH1), or in mitochondria (IDH2). In this reaction, D2-HG is produced in small amounts but converted back to its structural homolog α-KG by D2-HG dehydrogenase. Common features of tumors with IDH1/2 mutations are abnormal histone and DNA methylation, connecting metabolic changes with epigenetic control of gene expression (2). In hematologic malignancies, IDH1/2 are often co-mutated with epigenetic regulatory genes encoding enzymes that are important in DNA hydroxymethylation (i.e., tet methylcytosine dioxygenase 2, TET2) and methylation (i.e., DNA methyltransferase 3 A, DNMT3A) (3). Accumulation of D2-HG contributes to leukemogenesis, likely due to inhibition of α-KG–dependent dioxygenases, including histone lysine demethylases (KDMs) and TET2 (4). This hypothesis has been supported by recent reports linking the hypermethylation phenotype in cancer cells to IDH, fumarate hydratase, and succinate dehydrogenase mutations (5, 6).

The starting point for the present work were reports that myeloid malignancies are associated with cardiac pathologies, which are commonly considered a side effect of chemotherapy (7). Other recent reports suggest that systemically produced D2-HG by neomorphic IDH2 is associated with cardiomyopathy, suggesting that D2-HG influences cardiac cellular responses (8, 9). However, the extent to which D2-HG can directly affect cardiac function and metabolism, and which processes are involved, has remained unknown. It is known, however, that the heart adapts to stress by remodeling, both metabolically and structurally. Similar to cancer cells, metabolic remodeling in the heart is characterized by a shift from fatty acid utilization toward glucose utilization and, ultimately, mitochondrial dysfunction. This remodeling involves changes in pathways that regulate energy and redox homeostasis, growth, and autophagy, resulting in altered enzyme activities (10, 11). Consequently, we proposed that D2-HG mediates metabolic stress in the heart, and we tested this hypothesis using a targeted multiomics approach, together with the predictive and integrative value of mathematical modeling.

Results

D2-HG Promotes Cardiac Remodeling.

To investigate the impact of Idh2 mutations on cardiac remodeling, we generated mice bearing hematopoietic cells with an Idh2^R140Q mutation, which mimics one of the most common IDH1/2 mutations in acute myeloid leukemia (AML) patients. Wild-type (WT) C57BL/6 mice were lethally irradiated and reconstituted with hematopoietic stem/progenitor cells (HSPCs) transduced with a retrovirus expressing either WT Idh2 or Idh2^R140Q generating WT control (Idh2^WT) or single-mutant (Idh2^R140Q) HSPCs (Fig. S1A). We found no difference in the survival rate between mice with Idh2^WT and mice with Idh2^R140Q (Fig. S1B). We used liquid chromatography–mass spectrometry (LC-MS) to measure serum total 2-hydroxyglutarate (2-HG) in mice with neomorphic IDH2 (Idh2^R140Q) and WT control HSPCs 6 mo after bone marrow transplantation (BMT). The serum total 2-HG markedly increased fivefold in all mice transplanted with BM cells overexpressing mutant Idh2 compared with controls (Fig. S1C). A previous report indicated that overproduction of D2-HG, caused by a broadly expressed mutant IDH2, is associated with cardiac hypertrophy, dilatation, and failure (8). We found no cardiac hypertrophy in transplanted mice with Idh2^WT HSPCs 6 mo after BMT. In contrast, we observed substantial changes at the molecular level in the hearts of mice with Idh2^R140Q HSPCs 6 mo after BMT. In this group, levels of myosin-heavy-chain α (α-MHC) expression were decreased and myosin-heavy-chain β (β-MHC) expression levels were increased ([β-MHC]:[α-MHC] ratio increased) (Fig. S1 D–F). We and others have previously observed these same isoform changes in both hypertrophied and atrophied heart, suggesting cardiac remodeling (10–13). A shift in gene expression of MHC isoforms occurs only in response to stress and is dependent on the severity and duration of the stress. In other words, our findings support the hypothesis that D2-HG mediates metabolic stress, causing cardiac remodeling in myeloid malignancies.

Fig. S1.

Generation and characterization of acute myeloid leukemia model harboring Idh2^R140Q mutation. (A) Schematic representation of model generation. Hematopoietic progenitors isolated from the BM of the mice irradiated were transduced with empty vector or MSCV-IDH2R140Q and injected into recipient C57BL/6 mice (6–8 wk old). (B) Kaplan–Meier survival analysis in the recipient mice after transplantation of marrow progenitors transduced with retrovirus encoding empty vector (Idh2^WT, n = 10) or MSCV-IDH2R140Q (Idh2^R140Q, n = 15). (C) Total serum 2-HG levels from mice overexpression Idh2^R140Q and WT controls (Idh2^WT). The total 2-HG level was measured 6 mo after BMT by LC-MS. (D–F) The mRNA expression of myosin heavy chain (MHC) α (D) and β (E) and the ratio of β-MHC to α-MHC (F) were assessed in hearts from Idh2^R140Q mutant and Idh2^WT mice 6 mo after BMT. In C, n = 3–4 mice per group; in D–F, n = 5 mice per group. All data shown are mean ± SEM. Statistical analysis by ANOVA and Student’s t test. *P < 0.05; **P < 0.01; NS, not significant.

D2-HG Impairs Cardiac Energy Substrate Metabolism.

To understand whether overproduction of D2-HG alone was responsible for the effects observed in the Idh2 mutant mouse model, we measured rates of substrate metabolism in the isolated working rat heart and conducted computational flux rate analysis using the CardioNet model of mammalian cardiac metabolism (14). Rat hearts were perfused ex vivo in the presence or absence of D2-HG in concentrations similar to those found in the plasma of Idh2^R140Q mutant mice (0.5 mM) and AML patients (8, 15–17) and those reported by Latini et al. (18) to promote inhibition of ATP synthase in cardiac muscle in vitro (range 0.05–5 mM; F₀/F₁ ATP synthase K_i = 0.47 ± 0.18 mM) (Fig. 1A). Pretreatment levels of serum 2-HG in AML patients range from 19 to 96,000 ng/mL (15–17). Glucose and lactate were the only other substrates provided. The stressed heart (e.g., adrenergic stimulation) shifts to increased oxidation of glucose, whereas in the fasted state the heart oxidizes predominantly fatty acids for energy provision (19). Importantly, we limited the substrate supply to carbohydrates to assess any potential de novo fatty acid synthesis.

Fig. 1.

D2-HG impairs cardiac energy metabolism by inhibiting α-KGDH. (A) Protocol for the isolated working rat heart with (0.5 mM or 1.0 mM) or without D2-HG. (B) LC-MS analysis of D2-HG concentration in hearts perfused with (0.5 mM or 1.0 mM) or without D2-HG (n = 3). (C and D) Hydraulic power (C) and glucose oxidation rate (D) at near-physiologic (100 cmH₂O, 45–55 min) and increased workload (140 cmH₂O, acute stimulation 55–58 min, prolonged stimulation 65–75 min). (E) Chemical structures of α-KG and D2-HG. (F) DARTs blotting showing D2-HG as a substrate of α-KGDH and ATP5B. Susceptibility of both α-KGDH and ATP5B to pronase digestion is increased in the presence of D2-HG. (G–I) Effect of D2-HG on α-KGDH activity (G), H₂O₂ production rate (H), and catalase activity (I) in mitochondria isolated from hearts perfused with or without D2-HG. (J) MMP assessed by 3,3′-dipropylthiadicarbocyanine iodide (DiSC₃5) staining and corrected by carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP) of mitochondria isolated from hearts perfused with or without D2-HG. n = 3 rats per group. All data shown are mean ± SEM. Statistical analyses were performed with Kruskal–Wallis test, ANOVA, and Student’s t test. *P < 0.05; **P < 0.01; NS, not significant.

D2-HG was taken up by the perfused heart at a constant rate and accumulated in the tissue (Fig. 1B and Fig. S2A). The D2-HG tissue content was 110 ± 48 µg/g dry weight (0.5 mM D2-HG group) and 442 ± 65 µg/g dry weight (1.0 mM D2-HG group). With high D2-HG levels, we observed a significant decrease in cardiac power before and after an imposed increase in cardiac work (Fig. 1C). However, myocardial oxygen consumption was the same in all groups, likely due to inefficient oxidative phosphorylation of ADP, as indicated by the decline in hydraulic power and cardiac efficiency (Fig. S2 B and C). Next, we determined the effect of D2-HG on glucose oxidation and measured ¹⁴CO₂ production from d-[U-¹⁴C]glucose (19). In perfusions with 1.0 mM D2-HG, glucose oxidation rates increased significantly at high workloads (Fig. 1D). At the same time ATP levels were reduced and AMP levels were increased ([ATP]:[AMP] ratio decreased) (Fig. S2D). The results are consistent with prior reports that an increased supply of D2-HG leads to impaired ATP provision through inhibition of ATP synthase by D2-HG both in vivo and in vitro (18, 20). To assess whether D2-HG differentially affects energy substrate metabolism in the heart, we perfused rat hearts (n = 4 animals per group) with or without D2-HG (1 mM) in presence of glucose (5 mM) and oleate (0.4 mM) (Fig. S3A). Likewise, D2-HG accumulated in the tissue of perfused rat hearts (Fig. S3B). At normal workload we observed after 5 min of perfusion with D2-HG a steady decrease in cardiac power (Fig. S3C). Hearts perfused with D2-HG showed a decreased adrenergic response upon stimulation with epinephrine and significant decline in cardiac power. The oxidation of both glucose and oleate was significantly reduced in presence of D2-HG (Fig. S3D). We now found that with deficient oxidative phosphorylation at high D2-HG concentration cardiac metabolism shifted toward glycolysis, as demonstrated by higher glucose uptake and increased lactate release (Fig. S3E).

Fig. S2.

D2-HG impairs cardiac performance in the isolated working rat heart. (A) D2-HG uptake rate during acute (55–58 min) and prolonged (68–75 min) stimulation in isolated working rat hearts. (B and C) Myocardial oxygen consumption (B) and cardiac efficiency (C) are shown at near-physiologic (100 cmH₂O, 45–55 min) and increased workload (140 cmH₂O, 55–75 min) in perfused hearts. (D) ATP and AMP concentration and calculated [ATP]:[AMP] ratio in perfused rat hearts freeze-clamped at the end of the protocol. In A–D, n = 3 rats per group. Data are mean ± SEM. *P < 0.05; **P < 0.01; NS, not significant (Kruskal–Wallis test for perfusion data analysis and Student’s t test for pairwise comparisons).

Fig. S3.

D2-HG affects energy substrate metabolism in the isolated working rat heart. (A) Protocol for the isolated working rat heart with physiologic concentrations of glucose (5 mM) and oleate (0.4 mM) in the presence or absence of D2-HG (1.0 mM). (B) LC-MS analysis of tissue D2-HG content in perfused rat hearts freeze-clamped at the end of the protocol. (C) Hydraulic power at near-physiologic (100 cmH₂O, 45–55 min) and increased workload (140 cmH₂O, acute stimulation 55–58 min, prolonged stimulation 65–75 min) in rat hearts perfused with or without D2-HG. (D) Measurement of glucose oxidation and oleate oxidation rates in isolated working rat hearts with or without D2-HG. (E) Analysis of glucose uptake and lactate release. (F) Effect of D2-HG on α-KGDH activity in mitochondria isolated from hearts perfused with or without D2-HG at physiologic concentrations of glucose and oleate. In A–F, n = 4 rats per group. Data are mean ± SEM *P < 0.05; **P < 0.01; ***P < 0.001; NS, not significant (Kruskal–Wallis test for perfusion data analysis, ANOVA and Student’s t test for pairwise comparisons).

D2-HG Inhibits α-KG Dehydrogenase Activity.

Next, we hypothesized that D2-HG, as a structural homolog to α-KG (Fig. 1E), can bind to α-KG dehydrogenase (α-KGDH). Using drug-affinity responsive target stability (DARTS) (21), an unbiased molecular approach, we found decreased protease susceptibility of α-KGDH and ATP5B, the beta subunit of the catalytic core of the ATP synthase, in the presence of D2-HG in a concentration-dependent manner (Fig. 1F). This decreased susceptibility suggests that D2-HG binds to and stabilizes both α-KGDH and ATP5B, affecting their activity, as evidenced by the reduced ATP levels and consistent with previous reports (18, 20). We measured the level of α-KGDH activity and production rate of hydrogen peroxide (H₂O₂) fluorometrically in the ex vivo perfused rat hearts. There was a distinct reduction in α-KGDH activity at both 0.5 mM and 1.0 mM D2-HG (Fig. 1G and Fig. S3F), indicating an impaired NADH generation in the Krebs cycle and increased reductive carboxylation. H₂O₂ was produced at the same rate in both control and at 0.5 mM D2-HG perfused hearts (Fig. 1H). However, H₂O₂ formation rose markedly at high extracellular D2-HG concentrations (1.0 mM), implying that D2-HG impairs mitochondrial membrane potential (MMP) and induces generation of reactive oxygen species (ROS). To assess the activation of ROS defense mechanisms, we determined the catalase activity colorimetrically in isolated mitochondria from ex vivo perfused rat hearts. Catalase activity was increased at both 0.5 mM and 1.0 mM D2-HG (Fig. 1I), indicating that D2-HG promotes an acute rise in ROS generation and cellular adaptation. The production of ROS correlates with the MMP. A slight decrease in the membrane potential can cause a significant decrease in ROS production (22). We found the MMP markedly decreased in D2-HG perfused rat hearts (Fig. 1J). The diminished MMP impairs the ability of perfused hearts to provide ATP by oxidative phosphorylation. Together, these data indicate that the observed changes in α-KGDH activity were induced by D2-HG rather than a rise in H₂O₂ and show that D2-HG impairs Krebs cycle function and mitochondrial electron transport.

Computational Estimation of Metabolic Changes in the Presence of D2-HG.

To understand the effect of D2-HG and the impact of α-KGDH inhibition on metabolic reactions in the heart, we next performed computational simulations by flux balance analysis (23). We applied this algorithm to identify the metabolic reactions involved in the metabolic adaptation to high levels of D2-HG. To estimate flux rates, we used uptake and release rates of D2-HG, glucose, oxygen, and lactate, which were determined during the isolated working rat heart perfusions. We combined these measurements with previously reported rates for glycogen, protein, and lipid turnover to constrain the flux bounds of associated reactions during calculations (Table S1). Furthermore, we incorporated the experimentally measured decrease in α-KGDH activity to reflect the inhibition caused by D2-HG in the computations. We used the metabolic model of mammalian cardiac metabolism, CardioNet (14), and flux balance analysis to identify flux distributions that would optimally fit the experimental datasets. The objective of the optimization problem was to maximize ATP hydrolysis to reflect cardiac work. To validate the model predictions, we compared estimated glucose oxidation rates with tracer measurements and found a high correlation (R² = 0.9–0.98) between model simulations and flux rate measurements (Fig. S4A). Predicted flux rates were used to calculate the contribution of glycolysis, β-oxidation of fatty acids, and oxidative phosphorylation to ATP provision. Increased D2-HG supply caused a decrease in ATP synthesis, with most ATP being provided oxidatively (Fig. S4B). Glucose oxidation was the main source of ATP with or without D2-HG. However, ATP provision from glycolysis and β-oxidation of endogenous fatty acids was increased in simulations with D2-HG supply. D2-HG caused a shift in cardiac metabolism, increasing the reliance on endogenous substrates under experimental conditions (Fig. 2).

Table S1.

Constraints for CardioNet flux rate analysis

Constraint	Control		D2-HG		Source
	LB	UB	LB	UB
Glucose uptake	0.294	1.316	0.372	1.610	This study
Oxygen uptake	1.294	80.000	60.203	78.359	This study
D2-HG uptake	2.294	0.000	3.888	18.730	This study
Lactate release	3.294	0.046	0.099	0.109	This study
Glutamine release	0.000	Inf	0.000	Inf
NH3 release	0.000	Inf	0.000	Inf
Urate release	0.000	Inf	0.000	Inf
Glycogen
Synthesis	0.000	0.015	0.000	0.015	(19)
Degradation	0.000	0.136	0.000	0.136	(19)
Triglyceride turnover	−0.026	0.026	−0.026	0.026	(41)
Protein turnover (myosin)	−0.015	0.01	−0.015	0.01	(42)

Upper and lower bounds for network reactions are given in the table. These bounds define the maximum and minimum allowable fluxes of respective reaction. The stoichiometric matrix and reaction bounds define the solution space of flux distributions of a system, or, in other words, the rates at which every metabolite is consumed or produced by each reaction. Values are given in micromoles per minute per gram dry weight. Inf, Infinite; LB, lower flux bound; and UB, upper flux bound.

Fig. S4.

Validation and computational estimation of metabolic flux rates. (A) Estimated glucose oxidation rates were compared with measured glucose oxidation rates from isolated working rat hearts with or without D2-HG. For each experimental group, flux rate estimations and measurements were fitted to a regression line. The shaded area denotes the 95% confidence interval. (B) Estimations of ATP synthesis with or without D2-HG supply.

Fig. 2.

Flux rate analysis reveals dysregulation of cardiac energy substrate metabolism with increased D2-HG supply. Schematic of in silico flux rate analysis for glucose and D2-HG metabolism in ex vivo working heart perfusions. Colors indicate flux changes in the presence of D2-HG compared with control conditions. Ac-CoA, acetyl-CoA; Asp, aspartate; Fum, fumarate; Glut, glutamate; Homocys, homocysteine; Mal, malate; OAA, oxaloacetate; SAM, S-adenosylmethionine; Succ, succinate; Succ-CoA, succinyl-CoA; Tryp, tryptophane; and 5,10-Met-THF, 5,10-methenyl-tetrahydrofolate.

We identified metabolic differences by performing pairwise analysis of estimated flux rates with or without D2-HG supply (P < 0.05). Metabolic reactions and their metabolic subsystems, classified in the Kyoto Encyclopedia of Genes and Genomes database (24), are presented (Fig. S5 A and B). Increased supply of D2-HG is associated with distinct metabolic alterations, including increased β-oxidation of short chain fatty acids and degradation of amino acids. These reactions provided intermediates directly feeding into the Krebs cycle through succinyl-CoA, fumarate, and oxaloacetate. For example, mitochondrial provision of oxaloacetate through malate dehydrogenase (MDH) in the Krebs cycle decreased by 0.65-fold (Fig. 2). Further, D2-HG caused increased flux in the D2-HG dehydrogenase reaction and conversion of D2-HG to α-KG. Impairment of the Krebs cycle at the α-KGDH reaction shifted the model toward increased production of citrate through the NADPH-dependent reverse function of IDH2 (Fig. 2). This shift, in turn, increased fluxes for the transport of citrate into the cytosol and conversion of citrate to acetyl-CoA and oxaloacetate via the ATP citrate lyase (ACL). Our simulations indicate that D2-HG–dependent inhibition of α-KGDH shifts cardiac metabolism toward increased reductive carboxylation, which increases the reliance on glucose oxidation and endogenous substrates to maintain flux through the Krebs cycle.

Fig. S5.

Identification of metabolic flux rate changes promoted by D2-HG. (A and B) Experimental measurements including uptake and release rates were input into a flux balance analysis using the metabolic model CardioNet (Table S1). Metabolic differences between the flux distributions in cardiac metabolism with or without D2-HG supply were analyzed to identify significant changes. Colors indicate calculated P values for each metabolic reaction in simulations without D2-HG supply compared with simulations with D2-HG supply. Metabolic reactions are shown according to their subcellular localization in the cytosol (A) or mitochondria (B) and clustered according to their association metabolic pathways. The minus logarithms of the P values are presented.

D2-HG Promotes Metabolic and Epigenetic Alterations in the Heart.

Based on the model predictions, we determined the impact of D2-HG on the cytosolic and mitochondrial redox states. We measured the [pyruvate]:[lactate] and the [acetoacetate]:[β-hydroxybutyrate] ratios (25). The conversion of pyruvate to lactate and the formation of β-hydroxybutyrate to acetoacetate increased in the presence of D2-HG (Fig. S6 A and B). The differential expression pattern corresponds to a decreased cytosolic and mitochondrial [NAD⁺]:[NADH] ratio. We further quantified citrate, α-KG, and succinate levels and determined the glucose-derived triglyceride turnover by measuring ¹⁴C labeling in extracted lipids from perfused hearts. Increased amounts of D2-HG significantly increased the incorporation of glucose-derived glycerol into lipids (Fig. S6C). Decreased α-KGDH activity suggested that Krebs cycle intermediates might not be fully oxidized in the heart. Indeed, the levels of both citrate and succinate were elevated, whereas α-KG levels were decreased, resulting in a decreased [α-KG]:[succinate] ratio (Fig. 3 A and B). NAD⁺-dependent decarboxylation of pyruvate to acetyl-CoA by pyruvate dehydrogenase is followed by the condensation of acetyl-CoA with oxaloacetate to form citrate, which is, in turn, transported into the cytosol (26). In the cytosol, citrate is converted to acetyl-CoA and oxaloacetate by ACL. In silico modeling suggested an increased flux rate in presence of D2-HG. Therefore, we determined ACL and MDH activity in perfused hearts freeze-clamped at the end of the protocol. As expected, high levels of D2-HG (1.0 mM) increased ACL activity by 37% and MDH activity by 58% compared with controls (Fig. 3 C and D). These results suggest an increased conversion of citrate to acetyl-CoA and oxaloacetate in the presence of D2-HG (Fig. 3E). Previous studies have shown that changes in metabolite concentrations of Krebs cycle intermediates (e.g., citrate and succinate) and ACL activity affect epigenetic mechanisms regulating the posttranslational modification of the chromatin-modifying machinery (27, 28).

Fig. S6.

Effect of D2-HG on [NAD⁺]/[NADH] redox state, lipid remodeling, and histone modifcations. (A and B) Cytosolic and mitochondrial [NAD⁺]/[NADH] redox state was assessed from the tissue concentrations of lactate and pyruvate (A) as well as β-hydroxybutyrate and acetoacetate (B) in perfused rat hearts freeze-clamped at the end of the protocol. (C) d-[U-¹⁴C]glucose-derived ¹⁴C-enrichment of triglycerides. (D) Analysis of HAT activities in nuclear fractions from ex vivo working rat hearts in the absence and presence of D2-HG. (E) Analysis of global histone 3 acetylation (Ac-H3), and lysine 9 histone 3 acetylation (Ac-H3K9), and trimethylation (M3-H3K9) in histone extractions. The ratio of Ac-H3K9 normalized to Me3-K9 shows that acetylation is reduced during 30 min of perfusions with D2-HG. In A–E, n = 3 rats per group. Data are the mean ± SEM. *P < 0.05; NS, not significant (Student’s t test for pairwise comparisons).

Fig. 3.

Perfusion with D2-HG redirects Krebs cycle intermediates. (A and B) Analysis of citrate, α-KG, and succinate concentrations (A) and [α-KG]:[succinate] ratio (B) in perfused hearts freeze-clamped at the end of the protocol. (C and D) Effect of D2-HG on ACL (C) and MDH (D) activity in hearts perfused with or without D2-HG. (E) Schematic overview of ACL and α-KGDH reaction in the cytosol (cyto) and mitochondria (mito). In A–D, n = 3 rats per group; data are mean ± SEM. ANOVA and Student’s t test. *P < 0.05; **P < 0.01; NS, not significant.

To test whether the observed changes in ACL activity from D2-HG–perfused hearts affected the acetylation and methylation of histones, we determined the activity of global histone acetyltransferases (HATs), and protein levels of acetylated and methylated histone 3. Enzymatic assays performed on nuclear extracts from the perfused hearts revealed that the activity of HATs increased when exposed to D2-HG in a concentration-dependent manner (Fig. S6D), suggesting that the level of D2-HG affects the histone acetylation state of cardiomyocytes. We also assessed the ability of D2-HG to induce histone acetylation in histone extractions from perfused hearts. Pan-acetylation of histone 3 increased in D2-HG–perfused hearts compared with controls, whereas histone 3 K9 acetylation (Ac-H3K9) and trimethylation (Me3-H3K9) decreased (Fig. S6E). Thus, the balance between histone acetylation and deacetylation shifted toward increased histone deacetylation in isolated working rat heart perfusions with D2-HG. The observed effects can most likely be attributed to the relatively short perfusion time with D2-HG (30 min).

Prolonged D2-HG Supply Promotes Unique Metabolic and Epigenetic Changes.

To test whether the D2-HG–dependent metabolic and epigenetic changes in the heart also exist in vivo, we injected WT mice daily for 32 d with PBS (control, 0.2 mL) or D2-HG (250 mg D2-HG/ kg body weight). We observed significant skeletal muscle atrophy and reduction in total body weight (Fig. 4A and Fig. S7A). The serum D2-HG level was 657 ± 164 ng/mL in controls and 3,962 ± 356 ng/mL in with D2-HG–treated mice, thus within the range of reported serum D2-HG levels from AML patients (Fig. S7B). Next, we perfused mouse hearts from those groups ex vivo to assess the metabolic consequences of prolonged exposure to D2-HG on the heart. Consistent with the perfused rat hearts, we observed increased concentrations of D2-HG in the hearts of mice injected with D2-HG (Fig. S7B). Glucose was oxidized at a higher rate in hearts from D2-HG–treated mice, suggesting that continuous D2-HG supply causes a shift of cardiac metabolism toward higher glucose utilization rates (Fig. 4B). We also found a marked reduction in α-KGDH activity in the chronic exposure model, whereas there was no difference in H₂O₂ formation and catalase activity between control and D2-HG–treated animals (Fig. S7 C–E). The results suggest that glucose-derived pyruvate is decarboxylated to acetyl-CoA, promoting increased citrate synthesis. Under these conditions, ROS generation rises acutely, and with chronic exposure to D2-HG in vivo antioxidant defense mechanisms seem to be activated. We also asked whether prolonged exposure to D2-HG affected the activity of ACL and MDH, and histone methylation and acetylation. Consistent with measurements from rat heart perfusions, treating animals for 4 wk with D2-HG leads to a decreased [α-KG]:[succinate] ratio and a rise in ACL and MDH activity (Fig. 4 C and D and Fig. S7F). Furthermore, we observed that with D2-HG both the activity of HATs (Fig. S7G) and pan-acetylation of histone 3 (Ac-H3) increased, whereas specifically H3K9 acetylation (Ac-H3K9) and trimethylation decreased (M3-H3K9) (Fig. 4E). Increased demethylation of H3K9 resulted in a decreased [Ac-H3K9]:[Me-H3K9] ratio in D2-HG–treated hearts compared with controls. The results suggest an increased turnover rate for H3K9 deacetylation and demethylation during short periods of increased D2-HG supply, as observed in the isolated working rat heart perfusions. However, prolonged supply of D2-HG redirects pyruvate toward citrate and acetyl-CoA and consequently increases histone acetylation. Our data provide evidence that chronic exposure of the heart to D2-HG leads to reductive carboxylation, which contributes to both increased intracellular citrate concentrations and modifications of the cardiac epigenome (Fig. 4F).

Fig. 4.

Mice injected with D2-HG exhibit metabolic and epigenetic alterations in the heart. (A) Body weight and heart weight to tibia length ratio in mice after 32 d of PBS (control, Cnt) or D2-HG injection. (B) Rates of glucose oxidation (d-[U-¹⁴C]glucose) in hearts from mice treated with or without D2-HG. (C and D) Measurement of ACL (C) and MDH (D) activity in heart tissue from mice treated with PBS (control, Cnt) or D2-HG. (E) Analysis of global histone 3 acetylation (Ac-H3), lysine 9 histone 3 acetylation (Ac-H3K9), lysine 9 histone 3 trimethylation (M3-H3K9), and [Ac-H3K9]:[M3-H3K9] ratio in histone extractions from heart tissue in mice treated with PBS (control, Cnt) or D2-HG. (F) Schematic summarizing the proposed concept of metabolic and epigenetic modifications in the heart. In A–E, n = 10 mice per group. In F–H, n = 8 mice per group. Data expressed as mean ± SEM. Student’s t test. *P < 0.05, **P < 0.01; NS, not significant.

Fig. S7.

Effect of D2-HG on skeletal muscle weight and cardiac metabolism. (A) Weight of left and right gastrocnemius muscles in mice after 32 d of D2-HG injection. (B) LC-MS analysis of D2-HG concentration in serum and heart tissue from mice treated with or without D2-HG. (C–E) Effect of D2-HG on α-KGDH activity (C), H₂O₂ production rate (D), and catalase activity (E) in isolated mitochondria from mice injected with D2-HG (n = 10). (F) Analysis of α-KG and succinate concentrations and [α-KG]:[succinate] ratio in perfused hearts freeze-clamped at the end of the protocol. (G) Analysis of HAT activities in nuclear fractions from heart tissue from mice injected with D2-HG. In A–G, n = 10 mice per group. Data expressed as mean ± SEM. Student’s t test. *P < 0.05, **P < 0.01; NS, not significant.

Discussion

Metabolic alterations in AML caused by IDH2 mutations initiate the production of a small molecule, D2-HG, which is a structural homolog to the Krebs cycle intermediate α-KG. We show that IDH2 mutation in HSPC contributes to accelerated serum total 2-HG production, and that this increase promotes cardiac remodeling as indicated by the shift of α-MHC to β-MHC expression. Thus, increased levels of D2-HG impose metabolic stress not only in cancer cells but also in other tissues, driving adaptive changes in cellular processes. We demonstrate that cardiac dysfunction can be initiated directly by D2-HG through inhibition of α-KGDH, which impairs oxidative phosphorylation and promotes compensatory epigenetic modifications in the heart.

In the isolated working rat heart, D2-HG initiates cardiac dysfunction and metabolic alterations. We focused on glucose and lactate as energy-providing substrates because we were interested in assessing the possible effects of increased D2-HG supply on lipid remodeling, which cannot be assessed in the presence of fatty acids. Our results indicate that in response to increased levels of D2-HG cardiac metabolism shifts toward increased glycolysis and intermediary metabolites such as pyruvate and citrate are redirected into pathways other than the Krebs cycle. These processes are driven by both decreased α-KGDH activity and decreased production of reducing equivalents in the form of NADH. D2-HG is known to be a potent inhibitor of F₀/F₁ ATP synthase (18) and mediator of histone methylation modifications by inhibiting α-KG–dependent dioxygenases (e.g., KDMC4) in IDH2 mutant cells (4, 5).

We have now demonstrated that D2-HG also perturbs mitochondrial metabolism in the heart by inhibiting α-KGDH and decreasing the MMP. Analysis of substrate binding by DARTS demonstrated that D2-HG binds to α-KGDH, and that this binding leads to decreased α-KGDH activity and contributes to a decreased mitochondrial [NAD⁺]:[NADH] ratio. The regenerative function of many antioxidative and ROS-scavenging enzymes requires NADH or NADPH, and thus changes in the redox state can affect the antioxidant capacity of cells. In the isolated working rat heart, we show that increased D2-HG supply is associated with increased mitochondrial ROS production. A corresponding increase in catalase activity confirmed the cellular adaptation to increased mitochondrial ROS. Previous studies in isolated mitochondria demonstrated that the mitochondrial [NAD⁺]:[NADH] ratio and ROS production are positively correlated with the MMP (29). However, ROS changes can also be transient. A drop in MMP can be associated with ROS production and indicates a dysfunction of respiratory chain components (30). We observed in the isolated working rat heart that D2-HG supply decreased the MMP, which reduces the electron transfer across the mitochondrial membrane and decreases the ability to provide ATP by oxidative phosphorylation. This decrease in MMP was most likely caused by several mechanisms including deficiency of oxidizable substrates, due to impaired α-KGDH activity, and the uncoupling of the inner membrane, as indicated by the rise in ROS production. Prolonged exposure to D2-HG in mice did not display the increase in ROS observed in perfusions with D2-HG for 30 min, and thus catalase activation and MMP decrease were sufficient to normalize ROS production over time. Computational flux rate analysis indicates that specific metabolic processes are part of a response system to meet cellular energy demands to maintain cardiac output. Our findings suggest that, in conditions of increased D2-HG, citrate and acetyl-CoA formation is enhanced by two converging processes: (i) the decarboxylation of glucose to pyruvate driving acetyl-CoA formation through ACL and (ii) reductive carboxylation of α-KG by the reverse function of mitochondrial IDH (6). Reductive carboxylation requires an intracellular reducing environment (low [NAD(P)⁺]:[NAD(P)H] ratio) to generate NAD(P)H for the conversion of D2-HG to α-KG. Our results suggest that prolonged exposure to D2-HG shifts cardiac redox homeostasis toward NAD(P)H oxidation.

Dynamic regulation of histone modification is critical to the maintenance of cellular processes in response to stress (31). D2-HG affects histone methylation by inhibiting α-KG–dependent dioxygenases, resulting in epigenetic changes and promoting tumorigenesis in cancer cells (4). Prolonged systemic exposure to D2-HG in mice altered histone methylation and acetylation in the heart, specifically increasing pan-acetylation of histones and H3K9 demethylation. Recent studies have shown a link between histone trimethylation status and the development of cardiac hypertrophy, corroborating our conclusion that D2-HG acts as a metabolic signal promoting metabolic and structural adaptation in the heart. ACL-dependent production of acetyl-CoA has been shown to contribute to increased histone acetylation (28). Further, the activity of ACL and HATs synergize to maintain global histone acetylation and regulate chromatin structure by changes in the generation of acetyl-CoA. We now show that activities of ACL and HATs are increased in the presence of D2-HG. Thus, it seems that D2-HG–induced alterations in cardiac substrate utilization and energy metabolism are linked to changes in the acetylation of histones by ACL activity.

In conclusion, we demonstrate that cardiac dysfunction is initiated directly by D2-HG through inhibition of α-KGDH, promoting both impairment of oxidative phosphorylation and compensatory epigenetic modifications. We offer insight into the synergy of epigenetic and metabolic modifications caused by D2-HG in hematologic malignancies, which underscores the interaction of tumor metabolism and heart metabolism. The interaction between metabolism and epigenetics is of particular interest in light of recent investigations showing that the use of hypomethylating agents and histone deacetylase inhibitors may be beneficial in the treatment of cancer and heart failure (32, 33). However, our study is limited to fully elucidate all of the molecular mechanisms linking metabolic dysregulation with structural remodeling in the heart in response to elevated D2-HG levels. Further studies on the mechanisms leading to metabolic and epigenetic dysregulation may offer new opportunities to treat cancer and prevent (rather than cause) heart failure at the same time.

Methods

Animals.

All animal experiments were approved by the Institutional Animal Care and Use Committee and conducted according to the guidelines issued by The University of Texas Health Science Center at Houston and by Baylor College of Medicine, respectively. Animals were fed a standard laboratory chow (LabDiet 5001; PMI Nutrition International). All rats were male Sprague-Dawley rats obtained from Harlan Laboratories, and all mice were with a C57BL/6 background, obtained from Charles River Laboratories.

Metabolic Assays.

α-KGDH activity and H₂O₂ production rate were measured fluorometrically as described by Starkov et al. (34). ACL activity was determined as described by Srere (35). Metabolite concentrations were assessed colorimetrically using established enzymatic assays as described previously (19). Further detailed information on reagents and antibodies, isolated working rat heart and Langendorff mouse perfusions, in silico analysis of metabolic flux rates, Western blotting, and other methods is given in Supporting Information.

Generation of Idh2 Mutant Mouse Model

The mouse Idh2 cDNA clone was purchased from OpenBiosystem and cloned into pEntr-dTOPO (Invitrogen) by PCR using the following primer: Idh2-forward: caccatggccggctacctgc; Idh2-reverse: ctactgcttgcccagagctct, following the manufacturer’s instructions. Introduction of the point mutation was achieved by QuikChange II XL Site-Directed Mutagenesis Kit (Agilent Technologies). Mutant and WT Idh2 in pEntr-dTOPO were cloned into MSCV-RFB-IRES-GFP vector by Gateway recombination. Mice were killed and Sca1+ cells were enriched as described by Challen et al. (36). Next, 293T cells were transfected with MSCV-Idh2R140Q–IRES-GFP or MSCV-Idh2–IRES-GFP together with pCL-Eco vector. Sca1+ (0.5 × 10⁵) BM cells were plated in 1 mL Stempro-34 medium with mouse TPO (100 ng/mL), mouse SCF (10 ng/mL), and polybrene (4 µg/mL). Then, Sca1+ cells were spin-transduced at 1,100 rpm with 0.5 mL virus-containing supernatant for 2 h at room temperature and kept at 37 °C for 3 h. Infected Sca1+ cells were transplanted by retroorbital injection into C57BL/6 mice (6–8 wk old) after irradiation (9.5 Gy; Fig. S1A).

Isolated Working Rat Heart Perfusions

Hearts were perfused by the method described earlier (37). Briefly, rats (10–12 wk old, 340–380 g) were anesthetized with chloral hydrate (600 mg/kg, intraperitoneal) and heparinized (200 U) through direct injection into the inferior vena cava. Next, the chest was opened and the heart rapidly excised and arrested in ice-cold Krebs–Henseleit (KH) buffer (120 mM NaCl, 5mM KCl, 1.2 mM MgSO₄, 1.2 mM KH₂PO₄, 25 NaHCO₃, 2.5 mM Ca²⁺, 5 mM glucose, 0.4 mM oleate, and 0.5 mM lactate) at pH 7.4. Hearts were mounted on a cannula assembly and perfused retrogradely until they resumed beating and switched to the working mode thereafter, all at 37 °C with KH buffer equilibrated with 95% O₂ and 5% CO₂. The buffer of the experimental groups contained either 0.5 mM or 1.0 mM D2-HG (sodium salt, ≥95% purity; Cayman Chemical). The filling pressure was 15 cmH₂O with an afterload of 100 cmH₂O from start to min 55 of the perfusion. At this time epinephrine (1 μM) was added to the buffer, and the afterload was raised to 140 cmH₂O. Cardiac contractile performance was calculated as the product of cardiac output (sum of coronary flow and aortic flow, cubic meters per minute) and the afterload (pascals). Aortic pressure and heart rate were measured continuously with a 3 French manometer-tipped catheter (Millar Instruments) connected to a PowerLab 8/30 recording system (ADInstruments). At the end of the experiments the hearts were freeze-clamped between aluminum tongs cooled in liquid N₂ and stored at −80 °C until further use.

In Vivo Mouse Experiments

To assess the consequences of chronic D2-HG supply on cardiac function and metabolism, male mice (8–10 wk old; Charles River Laboratories) were divided into two groups (n = 10 animals per group), control or D2-HG treatment group, and injected over 32 d with either PBS (control group, 0.2 mL, intraperitoneal) or D2-HG (treatment group, 250 mg/kg body weight, intraperitoneal). At the end of each protocol, hearts were removed for ex vivo heart perfusions by the Langendorff method, and skeletal muscle tissue (gastrocnemius) was removed, weighed, and frozen in liquid N₂. The tibia length was documented as well.

Ex Vivo Mouse Heart Perfusions

We used the Langendorff method for isolated mouse heart perfusions, as described earlier (38). Mice were anesthetized with chloral hydrate (400 mg⋅kg⁻¹, intraperitoneal) and heparinized (200 U) through direct injection into the inferior vena cava. The chest was then opened and the heart rapidly excised and arrested in KH buffer (120 mM NaCl, 5mM KCl, 1.2 mM MgSO₄, 1.2 mM KH₂PO₄, 25 NaHCO₃, and 2.5 mM Ca²⁺) containing 5 mM glucose and 0.5 mM lactate at pH 7.4. The aorta was cannulated with a 20-gauge needle and the heart was transferred to the perfusion apparatus, which was a modification of the perfusion apparatus for isolated working rat heart. The heart was perfused with KH buffer (37 °C, 95% O₂ and 5% CO₂) at a constant perfusion pressure of 80 mmHg. After an equilibration period of 15 min hearts were perfused for another 20 min. At the end of the experiments the hearts were freeze-clamped with aluminum tongs aluminum tongs cooled in liquid N₂ and stored at −80 °C until further use.

Determination of Glucose and Oleate Oxidation Rates with d-[U-¹⁴C]Glucose and [9,10-³H]Oleate

In working rat heart experiments, hearts were perfused with KH buffer containing d-[U-¹⁴C]glucose (20 μCi/L, 9 dpm/nmol) with or without 0.4 mM sodium oleate prebound to 1% (wt/vol) BSA (fatty acid-free; Millipore), and [9,10-³H]oleate (30 μCi/L). The coronary effluent was collected every minute. In ex vivo Langendorff mouse hearts experiments, hearts were perfused for 20 min with KH buffer containing d-[U-¹⁴C]glucose (20 μCi/L, 9 dpm/nmol). Rates of glucose and oleate oxidation were determined by quantitative collection of ¹⁴CO₂ and ³H₂O released in the coronary effluent as described previously (19). Myocardial oxygen consumption (MVO₂) was measured by using YSI 5300A biological monitor (YSI Life Sciences). Electrodes were calibrated with air-saturated water (19.6% O₂ saturation after correction for water vapor, 47 mmHg at 37 °C). MVO₂ was calculated from the product of the arterial–venous difference and the coronary flow, using 1.06 mM for the concentration of dissolved O₂ at 100% saturation.

Gene Expression Analysis

Total RNA was prepared from freeze-clamped heart tissue using TRI Reagent according to the manufacturer’s protocol (Molecular Research Center, Inc.) and treated for potential DNA contamination with DNA-free DNA removal kit (ThermoFisher Scientific). Total RNA was then quantified using the NanoDrop ND-1000 (Thermo Fisher Scientific) and a 1.5-µg aliquot was reverse-transcribed into cDNA using RevertAid Reverse Transcriptase (Thermo Fisher Scientific). Relative quantification of target mRNA levels was performed on an ABI PRISM 7000 Sequence Detection System (Thermo Fisher Scientific) using specially designed primers and Taqman probes (Integrated DNA Technologies). RT-PCR amplification conditions were as follows: 10 min at 50 °C, 10 min at 95 °C, and PCR amplification (each of 40 cycles) at 95 °C for 15 s (melt) and 52 °C for 1 min (anneal/extend). All real-time PCR reactions included a “no reverse transcriptase control” to confirm the absence of DNA contamination in RNA preparations. Target transcript levels were normalized against cyclophilin A mRNA levels. The sequence and position of primers and probes are as follows:

Mus musculus myosin, heavy polypeptide 6, cardiac muscle, alpha (Myh6), National Center for Biotechnology Information (NCBI) reference sequence: NM_001164171.1

• Forward primer: 5′-GCAAAGGAGGCAAGAAGAAAGG-3′ (exon 16)

• Reverse primer: 5′-TGAGGGTGGGTGGTCTTCAG-3′ (exon 17)

• Taqman probe: 5′-FAM-ACAGTGTCTGCTCTCCACCGGGAA-TAMRA-3′ (exons 16–17)

Mus musculus myosin, heavy polypeptide 7, cardiac muscle,  beta (Myh7), NCBI reference sequence: NM_080728.2

• Forward primer: 5′-AGGGCGACCTCAACGAGAT-3′ (exon 35)

• Reverse primer: 5′-CAGCAGACTCTGGAGGCTCTT-3′ (exon 35)

• Taqman probe: 5′-FAM-AGCTCAGCCATGCCAACCGTA-TAMRA-3′ (exon 35)

Mus musculus peptidylprolyl isomerase A/cyclophilin A (Ppia/ CypA), NCBI reference sequence: NM_008907.1

• Forward primer: 5′-CCGATGACGAGCCCTTG-3′ (exon 1)

• Reverse primer: 5′-TCTGCTGTCTTTGGAACTTTGTC-3′ (exon 2)

• Taqman probe: 5′-FAM-CGCGTCTCCTTCGAGCTGTTTGCA-TAMRA-3′ (exons 1–2)

Western Blotting

Tissue homogenates were prepared in presence of phosphatase (Sigma-Aldrich) and protease (Roche Applied Science) inhibitors. Total histone extracts from tissue samples were prepared using the EpiQuik Total Histone Extraction Kit (OP-0006; Epigentek). Proteins were separated on 4–20% (wt/vol) SDS/PAGE gels (20 µg protein per well), transferred to PVDF membranes, and probed with antibodies [diluted 1:1,000 in 5% (wt/vol) BSA, 1× TBS, and 0.1% Tween 20, at 4 °C overnight] against total histone H3, histone H3 acetylated or trimethylated at lysine 9 (Abcam), and GAPDH (Fitzgerald Industries International). Standard anti-rabbit or anti-mouse secondary antibodies conjugated to HRP [rabbit: diluted 1:1,000; mouse: diluted 1:5,000 in 5% (wt/vol) skim milk in 1× TBS and 0.1% Tween 20; Santa Cruz Biotechnology] were used following incubation with primary antibodies. Levels of proteins were detected by immunoblotting using horseradish peroxidase-conjugated secondary antibodies and chemiluminescence (Santa Cruz Biotechnology). Signals were quantified by densitometry using NIH ImageJ software.

Target Identification Using DARTS

Hearts from WT C57BL/6 mice were lysed using M-PER (Thermo Fisher Scientific) with the addition of protease inhibitors and phosphatase inhibitors. TNC buffer (50 mM Tris⋅HCl, pH 8.0, 50 mM NaCl, and 10 mM CaCl₂) was added to the lysate and protein concentration was then determined using BCA protein assay. Tissue lysates were incubated with either vehicle (purified water) or varying concentrations of D2-HG (100 and 500 µM; Cayman Chemical Company) for 30 min at 37 °C followed by an additional 20 min in at room temperature. Digestion was performed using pronase (10 mg/mL stock, 1:3,000 dilution), and stopped by adding 4× Laemli loading buffer and immediately heating at 95 °C for 5 min. Proteins were separated on 4–20% (wt/vol) SDS/PAGE gels (20 µg protein per well), transferred to PVDF membranes, and probed with antibodies [dilution 1:1,000 in 5% (wt/vol) BSA, 1× TBS, and 0.1% Tween 20, at 4 °C overnight] against α-KGDH (Proteintech), ATP synthase subunit ATP5B (Sigma-Aldrich), α-actinin (Cell Signaling Technology), and GAPDH (Santa Cruz Biotechnology).

Absorption Spectroscopic Analysis of Intracellular Metabolites

Equal amounts of frozen heart tissue were deproteinized in ice-cold 6% (vol/vol) perchloric acid and homogenized with a 1-mL Dounce homogenizer. The suspension was immediately centrifuged (3,000 × g, 10 min at 4 °C). The supernatant fraction was transferred into a new microtube and neutralized with buffered KOH. Energy-rich phosphates (ATP, ADP, AMP, and glucose 6-phosphate) and pyruvate were measured in extracts, adjusted to pH 5 with KOH. Metabolite concentrations were assessed colorimetrically using established enzymatic assays as described previously (19). Citrate, α-KG, and succinate concentrations were measured using commercially available kits (Citrate Assay Kit ab83396, alpha KG assay Kit ab83431, and Succinate Assay Kit ab204718; Abcam). Tissue extractions were performed according to the manufacturer’s protocol. Next, samples were loaded to 96-well plates and either the absorbance was measured at 450 nm (succinate assay) or the fluorescence was followed at 587-nm emission after excitation at 535 nm (citrate assay). Extractions of total lipid fraction were prepared as reported previously by Harmancey et al. (39). Lipid extracts were emulsified by sonication for 30 s in buffer containing 28.75 mM 1,4-piperazinediethanesulfonic acid, 57.76 mM MgCl₂⋅6H₂O, 8.76 μM BSA (fatty acid-free), and 0.1% SDS. Next, the triglyceride content was quantified using the L-Type TG H assay (Wako Chemicals), following the manufacturer’s instructions.

Isolation of Mitochondria

Mitochondria were isolated from freeze-clamped hearts using a Dounce homogenizer in mitochondria isolation buffer (22 mM mannitol, 75 mM sucrose, 5 mM Hepes⋅KOH, pH 7.4, 1 mM EGTA, and 1 mg/L BSA) supplemented with protease and phosphatase inhibitors at 4 °C. Tissue homogenates were centrifuged at 3,000 × g for 4 min at 4 °C. The supernatants were subsequently centrifuged at 14,000 × g for 10 min at 4 °C, and pellets were resuspended in mitochondria isolation buffer. This step was repeated at least twice. After the last centrifugation, pellets were washed in mitochondria isolation buffer without EGTA and BSA, and stored at −80 °C until further processing.

Measurement of MMP

The MMP (ΔΨ) was determined as described before by Farrelly et al. (40). We measured the distribution of a ΔΨ sensitive probe 3,3*-dipropylthiadicarbocyanine iodide (DiSC₃5) in permeabilized mitochondria isolated from perfused and freeze-clamped hearts. The addition of the positively fluorescent probe DiSC₃5 to the liposomes results in a transient fluorescence that is self-quenched as the dye concentrates within the mitochondrial membrane. The addition of FCCP dissipates ΔΨ with the magnitude of the increase in fluorescence due to the released DiSC₃5 being proportional to ΔΨ. Mitochondria samples (50 mg/mL) were diluted into 100 µL of mitochondria isolation buffer, 5 µM valinomycin, and 1 µM DiSC₃5. The addition of 5 µM FCCP was used to dissipate the membrane potential to measure the increased fluorescence of DiSC₃5, and ΔΨ was monitored at the excitation wavelength of 650 nm and an emission wavelength of 590 nm. The fluorescence measurements were performed at 30 °C using a BioTek Synergy HT Multimode microplate reader (BioTek Instruments, Inc.). Data were collected and analyzed using BioTek Gen5 software (BioTek Instruments, Inc.).

Determination of α-KGDH Activity

α-KGDH activity was measured fluorometrically as described by Starkov et al. (34). In brief, mitochondria were isolated from freeze-clamped hearts in ice-cold isolation buffer (22 mM mannitol, 75 mM sucrose, 5 mM Hepes⋅KOH, pH 7.4, 1 mM EGTA, and 1 mg/L BSA) by differential centrifugation. Mitochondrial protein content was measured using the Bradford protein method. Samples of 0.1–0.25 mg/mL permeabilized mitochondria were added to the assay buffer (50 mM KCl, 10 mM Hepes, pH 7.4, 20 g/mL alamethicin, 0.3 mM thiamine pyrophosphate, 10 mM CaCl₂, 0.2 mM MgCl₂, 5 mM α-ketoglutarate, 1 μM rotenone, and 0.2 mM NAD⁺), and the reaction was started by adding 0.14 mM CoASH. The reduction of NAD⁺ to NADH/H⁺ was followed at emission after excitation at 346 nm for 10–60 min at 37 °C using a Quanta-Master model QM3-SS (Photon Technology International) cuvette-based fluorescence spectrometer. Samples were held at a constant temperature using a Peltier TEC temperature controller. Data were collected and analyzed using Felix 32 software (Photon Technology International).

Determination of H₂O₂ Production Rate

The measurement of H₂O₂ production rate was performed as described by Starkov et al. (34). Samples of permeabilized mitochondria (0.1–0.25 mg/mL) were incubated in assay buffer containing 125 mM KCl, 20 mM Hepes, pH 7.0, 2 mM KH₂PO₄, 4 mM ATP, 5 mM MgCl₂, 1 μM Amplex Red, 5 U/mL HRP, and 20 U/mL Cu,Zn-SOD. The change in concentration of H₂O₂ was assessed at 37 °C as an increase in Amplex Red fluorescence at 550 nm emission following excitation at 585 nm.

Measurement of ACL Activity

ACL activity was determined as described previously by Srere (35). Samples of 0.25–0.5 mg/mL of extracted proteins were added to the ACL assay buffer (21 mM potassium citrate, 5 mM ATP, 0.3 mM CoA, 0.1 mM NADH, 10 mM MgCl₂, 100 mM Tris⋅HCl, pH 7.5, 10 mM DTT, and 500 U of MDH). The reaction was initiated immediately after mixing all of the reagents and the sample. The oxidation of NADH to NAD⁺ was followed at 37 °C by measuring the absorbance at 340 nm.

Measurement of MDH Activity

Samples of extracted proteins (0.5 mg/mL) were added to the MDH assay buffer (0.1 mM phosphate buffer at pH 7.5, 2 mg/mL oxaloacetic acid, and 10 mg/mL NADH). The reaction was initiated immediately after mixing all of the reagents and the sample. The oxidation of NADH/H⁺ to NAD⁺ was followed at 37 °C by measuring the absorbance at 340 nm.

Measurement of Catalase Activity

Samples of extracted proteins (0.5 mg/mL) were added to the catalase assay buffer (50 mM sodium-phosphate buffer at pH 7.0 and 10 nM H₂O₂). The reaction was initiated immediately after mixing all of the reagents and the sample. The decomposition of H₂O₂ was followed at 37 °C by measuring the decrease in absorbance at 240 nm.

Measurement of HAT Activity

HAT activity was measured using a commercially available kit (ab65352; Abcam). Extractions of nuclear fractions and measurements were performed according to the manufacturer’s protocol. Briefly, nuclear fractions were isolated from freeze-clamped tissue in ice-cold extraction buffer (22 mM mannitol, 75 mM sucrose, 5 mM Hepes⋅KOH, pH 7.4, 1 mM EGTA, and 1 mg/L BSA) by differential centrifugation. Next, samples were loaded to 96-well plates and incubated at 37 °C for 3 h. The absorbance was measured at 440 nm every 20 min.

D2-HG Content in Tissue and Coronary Effluent

Analysis of serum total 2-HG and tissue D2-HG was conducted at Baylor College of Medicine Metabolomics Core Facility and CLIA Molecular Diagnostics Laboratory. Heart tissue was lysed in 500 μL extraction solution [50% (vol/vol) methanol and 50% (vol/vol) chloroform] per 50 mg tissue in a Dounce homogenizer. The suspension was immediately centrifuged (18,000 × g, 15 min at 4 °C) and the supernatant fraction was used for LC-MS analysis. For quantification, ratios of analytes and stable isotope-labeled internal standard, ¹³C₄-succinate (Sigma-Aldrich), were used. Heart tissue was lysed in 500 μL extraction solution [50% (vol/vol) methanol and 50% (vol/vol) chloroform] per 50 mg tissue in a Dounce homogenizer. The suspension was immediately centrifuged (18,000 × g, 15 min at 4 °C) and the supernatant fraction was used for LC-MS analysis. Chromatographic separation was conducted with a Shimadzu Nexera LC-30 UPLC system equipped with Restek Raptor Biphenyl 2.7 µm, 50 mm × 2.1 mm column using the following conditions. The mobile phase A was water/0.1% formic acid/4 mM ammonium formate; the mobile phase B was methanol/0.1% formic acid/4 mM ammonium formate. Samples were running with gradient (%B): 0 min at 2%, 1 min at 2%, 1.5 min at 100% followed by 1 min washing with 100% and equilibrating 1 min at 2%. The flow rate was 0.4 mL/min and the injection volume was 10 μL. Detection was performed in a Shimadzu 8040 mass spectrometer using electron spray ionization source at positive ion mode and the following parameters: nebulizing gas flow 2.8 L/min, drying gas flow 15 L/min, DL temperature 220 °C, BH temperature 400 °C. D2-HG was identified by multiple reaction monitoring mode, monitoring the transition of m/z 147.0 for precursor ion to m/z 129.1 and 101.2 for product ions. For quantification, ratios of analytes and stable isotope-labeled internal standard, ¹⁴C₄-succinate (Sigma-Aldrich), were used. Identification of the D2-HG peaks was confirmed using purified D2-HG. Signal intensities were quantified by integration of peak areas. The analytical data were processed by Shimadzu LC solution software (Shimadzu Scientific Instruments).

In Silico Analysis of Metabolic Flux During D2-HG Supply

Simulations were performed using the mathematical model of mammalian cardiac metabolism CardioNet (14). Simulations were run with boundary conditions reflecting the metabolite composition of the perfusion buffer and experimentally measured uptake and release rates for glucose, lactate, oxygen, and D2-HG (Table S1). At the same time, various metabolites, including amino acids and lipids, were set to previously reported values (14) to determine whether they contribute to ATP production and anaplerosis of Krebs-cycle intermediates. The following flux balance analysis was applied to identify steady-state flux distributions that are in agreement with applied substrate uptake and release rates:

$$\max v_{ATPase}$$

subject to

$$S\cdot v=0,$$

$$v_i^{(-)}\le v_i\le v_i^{(+)},$$

$$L_j^{(-)}\le v_j\le L_j^{(+)}\hspace{0.17em}\left(j=j_1,\hspace{0.17em}{j}_2,\dots \right),$$

where $v_j$ denotes the measured uptake or secretion rate through reaction j.

Statistical Analysis

Comparison between groups was performed by nonparametric (Kruskal–Wallis test) and parametric (Student’s t test or ANOVA) test methods and Kaplan–Meier curves for survival analysis. A P value less than 0.05 was considered statistically significant.