Mitophagy mitigates mitochondrial fatty acid β-oxidation deficient cardiomyopathy

Sun, Nuo; Barta, Hayley; Chaudhuri, Samhita; Chen, Kangxuan; Jin, Jiacheng; Luo, Hongke; Yang, Mingchong; Krigman, Judith; Zhang, Ruohan; Sanghvi, Shridhar; Sekine, Shiori; Sanders, Hannah; Kolonay, Dominic; Patel, Mudra; Baskin, Kedryn; Singh, Harpreet; Zhang, Pengyi; Xin, Gang; Finkel, Toren

doi:10.1038/s41467-025-60670-z

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Published: 01 July 2025

Mitophagy mitigates mitochondrial fatty acid β-oxidation deficient cardiomyopathy

Nature Communications volume 16, Article number: 5465 (2025) Cite this article

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Abstract

The healthy heart relies on mitochondrial fatty acid β-oxidation (FAO) to sustain its high energy demands. FAO deficiencies can cause muscle weakness, cardiomyopathy, and, in severe cases, neonatal/infantile mortality. Although FAO deficits are thought to induce mitochondrial stress and activate mitophagy, a quality control mechanism that eliminates damaged mitochondria, the mechanistic link in the heart remains unclear. Here we show that mitophagy is unexpectedly suppressed in FAO-deficient hearts despite pronounced mitochondrial stress, using a cardiomyocyte-specific carnitine palmitoyltransferase 2 (CPT2) knockout model. Multi-omics profiling reveals impaired PINK1/Parkin signaling and dysregulation of PARL, a mitochondrial protease essential for PINK1 processing. Strikingly, deletion of USP30, a mitochondrial deubiquitinase that antagonizes PINK1/Parkin function, restores mitophagy, improves cardiac function, and significantly extends survival in FAO-deficient animals. These findings redefine the mitophagy response in FAO-deficient hearts and establish USP30 as a promising therapeutic target for metabolic cardiomyopathies and broader heart failure characterized by impaired FAO.

Introduction

The heart relies almost exclusively on mitochondria to generate the large amounts of ATP required to fuel cardiac contractile function^1,2. In patients with heart failure (HF), deficient cardiac energetics, altered mitochondrial substrate utilization, and impaired mitochondrial function are thought to underlie contractile dysfunction and the progression of the disease². Cardiomyocytes can exhibit metabolic flexibility and dynamic substrate utilization under differing physiological conditions^3,4,5. The healthy adult heart relies predominantly on mitochondrial fatty acid β-oxidation (FAO) to fuel ATP production^1,3,6,7. In contrast, a failing or hypertrophied heart usually exhibits impaired FAO and enhanced reliance on glucose utilization^1,4,7. Mutations or deficiencies in the FAO enzymes can disrupt the breakdown of fatty acids for energy production, leading to compromised mitochondrial function and impaired metabolism⁸. Consequently, individuals with FAO deficiency may present with a spectrum of symptoms, including muscle weakness, hypoglycemia, liver problems, and various forms of cardiomyopathy^9,10. The severity of cardiac symptoms in individuals with FAO deficiency underscores the profound impact of impaired FAO on the heart. These clinical manifestations not only result in frequent hospitalizations but can also lead to high mobility and mortality rates. In severe cases, particularly in the neonatal and infantile forms, FAO deficiency may manifest as cardiomyopathy, ultimately culminating in lethality^9,10. Prognosis and treatment for patients with the lethal neonatal/infantile form of FAO deficiency are generally poor, with death occurring within days to months after birth⁹.

The capability to adapt cellular metabolism to the challenging conditions present in FAO-deficient hearts necessitates the preservation of mitochondrial function and integrity through specific mitochondrial quality control mechanisms. Mitophagy, a process essential for mitochondrial quality control, serves as a cardio-protective mechanism by eliminating dysfunctional mitochondria, thereby preventing their accumulation and minimizing potential cellular damage¹¹. Despite the recognized role of mitophagy in myocardial recovery from mitochondrial damage in various cardiac pathological conditions^2,12,13, such as cardiac hypertrophy^14,15, myocardial infarction¹⁶, ischemic preconditioning^17,18, and ischemia/reperfusion^19,20, the underlying regulatory mechanisms of mitophagy in response to cardiac injury and bioenergetic deficits remain elusive. Notably, mitophagy is sensitive to bioenergetic deficits and may be required for the heart to adapt to altered bioenergetic conditions, particularly in the context of FAO deficiency. However, our understanding of the mechanistic correlation between mitophagy and myocardial bioenergetics, as well as the regulatory mechanisms governing mitophagy in response to bioenergetic deficits, remains limited.

Research in cellular models has uncovered multiple mechanistically distinct pathways that converge upon and activate mitophagy, with one of the most well-characterized pathways involving the mitochondrial-targeted PTEN-induced kinase 1 (PINK1) and the E3 ubiquitin ligase Parkin^11,21,22. PINK1 undergoes constitutive cleavage and degradation by mitochondrial proteases, including the mitochondrial rhomboid protease PARL (presenilin-associated rhomboid-like protein)^11,21. However, upon mitochondrial damage, PINK1 evades proteolytic degradation and accumulates on the outer mitochondrial membrane (OMM) of impaired mitochondria^11,21. In this context, PINK1 phosphorylates ubiquitin attached to OMM proteins. These phospho-ubiquitin chains bind to Parkin, recruiting it from the cytosol to the mitochondria and activating its latent E3 ubiquitin ligase activity. Parkin then ubiquitinates OMM proteins, facilitating recruitment of receptors such as optineurin (OPTN) and nuclear dot protein 52 (NDP52), signaling autophagosome assembly proximal to individual damaged mitochondria^11,21. Interestingly, recent evidence suggests that mitochondrial deubiquitinating enzymes, including USP30, can remove ubiquitin attached to OMM proteins, thereby antagonizing PINK1/Parkin-mediated mitophagy^23,24,25. The pivotal role of PINK1 and Parkin in orchestrating mitophagy is emphasized by the observation that loss-of-function mutations in either PINK1 or Parkin result in early-onset Parkinson’s disease in humans^11,21,25. The specific roles of PINK1/Parkin-dependent mitophagy mechanisms in both healthy and diseased hearts are currently in the early stages of investigation. Diminished levels of PINK1 protein are observed in human heart failure²⁶. Mice lacking PINK1 exhibit disrupted mitochondrial function, leading to cardiac hypertrophy and contractile dysfunction at an early age. However, additional research is required to elucidate the mechanistic role for PINK1/Parkin in regulating cardiac mitophagy, especially in response to myocardial bioenergetic deficits.

To gain insights into the mitophagic regulation underlying impaired FAO in the heart, we generated FAO-deficient mice with cardiomyocyte-specific deletion of the carnitine palmitoyltransferase 2 (CPT2), an inner mitochondrial membrane (IMM) protein essential for metabolizing long-chain fatty acids (LCFA) in the mitochondria^27,28,29,30. We revealed a suppressed PINK1/Parkin signaling pathway and a diminished mitophagic response in FAO-deficient hearts. Intriguingly, we identified that the loss of cardiac FAO impairs the PINK1/Parkin pathway via modulating the mitochondrial rhomboid protease PARL, establishing a previously unrecognized connection between FAO and PINK1-dependent mitophagy. In addition, using cardiomyocyte-specific USP30 knockout mice, we demonstrated that deleting USP30 activates cardiac mitophagy, alleviating the FAO defect-associated decline in cardiac function. This underscores the potential therapeutic value of enhancing mitophagy in the treatment of currently intractable FAO-deficient cardiomyopathies.

Result

Cardiac-specific deletion of CPT2 induces heart failure

The ability to metabolize long-chain fatty acids (LCFA) in the mitochondria requires the carnitine palmitoyl transferase (CPT) system, consisting of CPT1 and CPT2 positioned in the outer and inner mitochondrial membrane, respectively^27,28 (Fig. 1a). Although various CPT1 isoforms (CPT1A, CPT1B, CPT1C) have been identified, there are no tissue-specific isoforms of CPT2^29,30. In a clinical context, deficiency in CPT2 manifests as a rare inherited metabolic disorder, disrupting the transport of long-chain fatty acids into mitochondria for subsequent breakdown and energy production^31,32. This deficiency leads to compromised beta-oxidation of fatty acids, particularly in tissues heavily reliant on fatty acid metabolism for energy, such as skeletal muscle and the heart^31,32. To investigate the specific impact on cardiac function, we generated mice with cardiomyocyte-specific deletion of CPT2 by crossing CPT2^fl/fl mice^8,33 with transgenic mice expressing Cre recombinase under the control of the cardiac-specific alpha myosin-heavy chain (Myh6) promoter³⁴ (CPT2^H-KO). We demonstrated that the CPT2 protein is reduced substantially in CPT2^H-KO hearts relative to wild-type (Myh6-Cre/CPT2^WT/WT) hearts, confirming the fidelity of cardiac-specific deletion of CPT2 (Fig. 1b, c). In contrast, CPT2 levels were maintained in brain, liver, and skeletal muscle tissues (Supplementary Figs. 1a, b).

**Fig. 1: Cardiac-specific CPT2 deletion disrupts mitochondrial FAO.**

To directly probe lipid metabolic remodeling in hearts deficient in FAO, we conducted liquid chromatography-tandem mass spectrometry (LC-MS/MS)-based unbiased lipidomic analysis on both wild-type and CPT2^H-KO hearts. In the hearts of CPT2^H-KO mice, our analysis identified 2521 metabolites exhibiting significant alterations compared to their levels in wild-type hearts (Fig. 1d) (p < 0.05 and fold change > 2.0; n = 5). Principal-Component Analysis (PCA)^35,36 underscored a notable shift in the lipid metabolic profile between wild-type and CPT2^H-KO hearts (Fig. 1e). To delve into the functional implications of CPT2 deletion on lipid metabolic pathways in the heart, we employed the mummichog algorithm alongside an adapted gene set enrichment analysis (GSEA) method^35,36. The lipid pathway analysis uncovered key metabolic pathways, with acylcarnitines and lysophosphatidylcholine (LPC) metabolism emerging as top hits (Fig. 1f). Consistent with the known role of CPT2 in converting long-chain acylcarnitines to their respective long-chain acyl-CoAs, we observed significantly elevated levels of long-chain acylcarnitines, including palmitoylcarnitine, stearoylcarnitine, and arachidylcarnitine, in CPT2 deficient hearts (Fig. 1g). Interestingly, levels of short-chain and medium-chain acylcarnitines, which operate independently from the CPT system, exhibited minimal to no alteration (Fig. 1g). These findings conclusively demonstrate that the absence of CPT2 in the heart leads to a substantial accumulation of long-chain acylcarnitines, indicative of impaired FAO in CPT2^H-KO hearts.

CPT2 deficiency profoundly impacts the heart, leading to diverse cardiac symptoms, including cardiomyopathy^32,37. We noted that CPT2^H-KO mutant mice exhibited increased mortality after postnatal day 28 (P28) (Fig. 2a). Necropsies revealed an increased heart-to-body weight ratio in CPT2^H-KO mice (Fig. 2b). Furthermore, histological examination (Fig. 2c) and Wheat Germ Agglutinin staining (Supplementary Figs. 1c, d) demonstrated marked cardiac hypertrophy in CPT2^H-KO hearts compared to controls. We next examined the effect of CPT2 depletion on left ventricular function. Longitudinal analyses of mice by echocardiographic imaging conducted in wild-type and CPT2^H-KO mice (Fig. 2d–f) revealed a decline in left ventricle (LV) systolic function, as indicated by reduced fractional shortening (FS) (Fig. 2d) and ejection fraction (EF) (Fig. 2e) in CPT2^H-KO mice. Concurrently, there was an increase in LV diameter measured using the left ventricular internal dimension (Fig. 2f). Thus, the absence of FAO precipitates the cardiomyopathy observed in CPT2-deficient mice.

**Fig. 2: Cardiac phenotypes of CPT2^H-KO mice.**

The loss of FAO modulates the mitochondrial transcriptome and proteome

To gain further mechanistic insights regarding the functional consequences resulting from CPT2 deletion, we initiated a transcriptomic analysis on heart samples obtained from both CPT2^H-KO and control mice. Employing Gene Set Enrichment Analysis (GSEA), we explored statistically significant and concordant molecular signatures in CPT2^H-KO and control RNA sequencing samples, revealing divergent gene enrichment patterns related to mitochondrial energy production (Fig. 3a–c). Particularly noteworthy was the significant downregulation of genes associated with the Electron Transport Chain (ETC) and the tricarboxylic acid (TCA) cycle following CPT2 deletion (Fig. 3b, c). Since transcriptomic changes often do not correlate with changes at the protein level³⁸, we next interrogated the proteomic landscape of CPT2-deficient hearts. We conducted quantitative proteomic analysis on heart samples from both wild-type and CPT2^H-KO mice (4 weeks of age, n = 4). From a total of 6663 quantified proteins, we generated fold-change values by normalizing the protein abundance of CPT2^H-KO samples against control samples. We identified 185 upregulated and 239 downregulated proteins, each exhibiting a fold change > 1.5 (p < 0.05) (Fig. 3d). As anticipated, we observed significantly reduced CPT2 protein levels in myocardial samples from CPT2^H-KO mice (Fig. 3d), consistent with the western blot analysis (Fig. 1b). Among the downregulated proteins, we identified a significant enrichment of proteins related to cardiac muscle contraction, aligning with the GSEA analysis of gene expression levels (Supplementary Figs. 2a, b). Surprisingly, we noted increased levels of mitochondrial protein expression in CPT2^H-KO samples (Fig. 3d, e), contradicting findings from the transcriptomic analysis (Fig. 3a–c). To further elucidate this discrepancy, we mapped the upregulated proteins in a cellular component analysis, revealing a widespread distribution across various mitochondrial components, including the mitochondrial membrane, matrix, and the translocase of the inner membrane (TIM) complex (Fig. 3e). In alignment with the proteomic profile, we confirmed a substantial increase in TOM40, TIM22, and TIM23 protein levels in CPT2^H-KO hearts compared to their wild-type counterparts (Fig. 3f, g). These inconsistent observations between mitochondrial protein levels and transcription suggest a potential decrease in mitochondria degradation, possibly indicating a mitophagy defect in CPT2^H-KO hearts.

**Fig. 3: Transcriptomics and proteomics analysis of CPT2^H-KO mice.**

Impaired FAO disrupts PINK1/Parkin-mediated mitophagy via PARL activation

Mitophagy plays an essential role in maintaining mitochondrial homeostasis and is sensitive to bioenergetic deficits^22,39,40. Leveraging the mt-Keima analysis⁴¹, we observed an approximate 80% reduction in mitophagy following the deletion of cardiac CPT2 (Fig. 4a, b). Furthermore, we noted an augmented accumulation of mitophagy receptors, p62 and OPTN^21,42,43 in CPT2^H-KO hearts (Fig. 4c, d). These receptors are typically recruited to damaged mitochondria and are subsequently degraded during mitophagy²¹. Conversely, in conditions of impaired mitophagy, these adapter proteins can accumulate^21,44. Notably, OPTN, a crucial adapter in initiating the formation of autophagosomes on the surface of damaged mitochondria during PINK1 and Parkin-mediated mitophagy²¹, exhibited a remarkable 9.8-fold increase in CPT2^H-KO hearts (Fig. 4c, d).

**Fig. 4: Loss of cardiac FAO inhibits mitophagy.**

The mitochondrial-targeted kinase PINK1 and the E3 ubiquitin ligase Parkin, recognized for their involvement in Parkinson’s disease, play pivotal roles in governing mitophagy^11,21. Emerging evidence further suggests that PINK1-mediated mitophagy serves as a major mechanism for mitochondrial quality control in the heart during cardiac stress and injury^26,45. Interestingly, GSEA analysis reveals a pronounced downregulation of genes associated with PINK1 inactivation and inefficient electron transport following CPT2 deletion (Supplementary Figs. 3a, b), implying that the suppression of mitophagy by CPT2 deletion may occur through the PINK1-mediated signaling pathway. Consistent with this hypothesis, there was a substantial decrease in PINK1 abundance in CPT2^H-KO hearts relative to wild-type hearts (Fig. 4e, f). Furthermore, mitochondria-associated Parkin was reduced in CPT2^H-KO hearts, despite total Parkin protein levels remaining unchanged relative to wild-type controls (Supplementary Figs. 3c, d).

We next sought to explore the regulatory mechanisms impacting PINK1 following CPT2 deletion. The mitochondrial rhomboid protease PARL plays a key role in mediating the cleavage of PINK1, thereby influencing its cellular localization and levels^39,46. Remarkably, the protein levels of PARL exhibited a significant increase in CPT2^H-KO hearts compared to their wild-type counterparts (Fig. 4g, h), suggesting that CPT2 deletion may impede PINK1 through the modulation of PARL levels or activity. PARL can also mediate the cleavage of phosphoglycerate mutase 5 (PGAM5) in response to the loss of mitochondrial membrane potential⁴⁷. We confirmed that treatment with FCCP (carbonyl cyanide p-trifluoromethoxyphenylhydrazone), a potent mitochondrial uncoupler, induced the cleavage of full-length PGAM5 (PGAM5-FL) into its cleaved form (PGAM5-CL) in HeLa cells (Fig. 4i). Consistent with increased PARL activity, we observed a notable increase in PGAM5 cleavage in CPT2-deficient hearts, a known substrate of PARL^48,49,50,51 (Fig. 4i, j). The precise mechanism underlying PARL upregulation remains unclear. Altered expression of Pyruvate Dehydrogenase Kinases (PDKs) post CPT2 deletion (Fig. 4e–g) may play a role in regulating PARL activity^39,46,52,53. Metabolic re-wiring, specifically reduced FAO, could induce an increase in glycolysis by inhibiting the pyruvate dehydrogenase complex through PDK activation^54,55. Previous studies indicate that PDK2 can phosphorylate PARL and inhibit β cleavage, a vertebrate-specific autocatalytic proteolytic processing that negatively regulates PARL activity³⁹. In support of this hypothesis, Phos-tag gel analysis revealed increased PARL phosphorylation in CPT2^H-KO hearts compared to wild-type controls (Supplementary Figs. 3e, f). While the specific PDK isoform(s) regulating PARL activity remain to be determined, these results reinforce the notion that the loss of cardiac FAO may compromise the PINK1/Parkin pathway by modulating PARL activity. These compelling findings establish a connection between FAO deficiencies and impaired mitophagy, offering a potential therapeutic avenue for treating FAO-deficient cardiomyopathy by targeting mitophagy.

Cardiac-specific deletion of USP30 restores mitophagy and rescues FAO-deficient hearts

Impaired mitophagy in the heart can have deleterious consequences on mitochondrial function, potentially contributing to the development of heart failure. Conversely, restoring mitophagy holds promise as a strategy to protect mitochondrial function and alleviate cardiac dysfunction resulting from deficient FAO. Recent evidence suggests that inhibiting PARL could result in the accumulation of PINK1^48,56, potentially triggering mitophagy⁵⁶. Nevertheless, it is crucial to note that PARL has multiple substrates and PARL deletion has been associated with defects in Complex III activity and the onset of progressive mitochondrial dysfunction⁴⁸. Consequently, inhibiting PARL may intensify cardiac damage in FAO-deficient mice. An alternative strategy for stimulating mitophagy involves the manipulation of the mitochondrial deubiquitinating enzyme USP30^24,57. Recent evidence from experiments utilizing both cell culture and flies suggests that USP30 acts to antagonize the activity of PINK1/Parkin-mediated mitophagy^23,24. Building upon these insights, we generated USP30^H-KO mice (Myh6-Cre/USP30^fl/fl) lacking cardiac USP30 (Fig. 5a, b), while, as expected, USP30 expression was retained in non-cardiac tissues (Supplementary Figs. 4a, b). We next asked whether inhibiting USP30 could restore mitophagy and ameliorate cardiac function in FAO-deficient hearts. Interestingly, we observed a significant enhancement in cardiac mitophagy upon the loss of USP30 in the context of CPT2 deletion (Fig. 5c, d), despite indistinguishable baseline cardiac mitophagy between USP30^H-KO and wild-type animals (Supplementary Fig. 4c). Furthermore, CPT2/USP30^H-KO hearts exhibited a notable reduction in TIM22, TIM23, and OPTN protein levels compared to CPT2^H-KO hearts, suggesting a decrease in overall mitochondrial mass likely resulting from enhanced mitophagic clearance (Supplementary Fig. 4d, e). However, we found that the absence of USP30 had minimal impact on the levels of PINK1 in the context of CPT2 deletion (Supplementary Fig. 4f, g). Consistent with this, levels of PDKs and PARL remain elevated following USP30 deletion in the context of FAO deficiency (Supplementary Figs. 4f, g), supporting the notion that PINK1 expression is suppressed via PARL-dependent mechanisms. These results suggest that USP30 inhibition may enhance mitophagy during cardiac stress by lowering the OMM ubiquitination threshold required for mitophagy activation^58,59,60,61. To further explore this mechanism, we assessed mitochondrial protein ubiquitination and observed a marked increase following USP30 deletion in the context of CPT2 deficiency (Supplementary Figs. 4h, i). Notably, this increase occurred despite unchanged Parkin recruitment to mitochondria (Supplementary Figs. 4h, i), suggesting that USP30 deletion enhances mitochondrial ubiquitination through a mechanism independent of Parkin translocation. These findings are consistent with previous studies^58,59,60,61.

**Fig. 5: Deletion of USP30 promotes cardiac mitophagy and rescues CPT2-deficient mice.**

To characterize mitochondrial ultrastructure and morphological features, we employed transmission electron microscopy (TEM), revealing conspicuous alterations in morphology featuring damaged cristae morphology and reduced cristae density in CPT2^H-KO hearts compared to their wild-type counterparts (Fig. 5e–g). These observations suggest compromised fitness of cardiac mitochondria in the absence of FAO. Intriguingly, the absence of USP30 leads to a marked improvement in mitochondrial ultrastructural abnormalities, characterized by a significant reduction in swelling and enhanced integrity in the context of CPT2 deletion (Fig. 5e–g, and Supplementary Fig. 5a). This is consistent with the notion that USP30 deletion may preserve mitochondrial quality and stabilize cristae structure. In alignment with the cardioprotective effects of mitophagy, the decline in cardiac function was significantly ameliorated in the combined CPT2/USP30^H-KO animals compared to CPT2^H-KO mice (Fig. 5h–k). The previously observed reduction in systolic function in CPT2-cKO mice was notably less pronounced in age-matched mice lacking both USP30 and CPT2 (Fig. 5h–k). The left ventricular mass was significantly reduced in the CPT2/USP30^H-KO mice compared to age-matched CPT2^H-KO mice (Fig. 5k). Importantly, the absence of USP30 extended the lifespan of animals lacking CPT2. In our colony, the median survival of CPT2^H-KO mice was less than 7 weeks, and all mutant mice succumbed by 11 weeks (Fig. 5l). Deletion of USP30 provided a significant survival advantage, extending the median survival of the CPT2/USP30^H-KO mice to 15 weeks with the maximum survival exceeding a year (Fig. 5l). Collectively, these results indicate that USP30 inhibition enhances mitophagy and alleviates cardiac dysfunction induced by CPT2 deletion.

Deletion of USP30 improves mitochondrial metabolism in FAO-deficient hearts

We next conducted a metabolomics screen for an unbiased analysis of the most differentially affected pathways between CPT2^H-KO and CPT2/USP30^H-KO mice. Principal component analysis (PCA) highlighted a profound shift in the metabolite composition of the heart following CPT2 deletion compared to CPT2/USP30^H-KO and wild-type animals (Fig. 6a). In addition, the PCA revealed that the deletion of USP30 in the context of CPT2 deficiency promoted a metabolic switch toward the wild-type, compensating for defective FAO. To explore which metabolites were predominantly altered by USP30 deletion in the settings of FAO deficiency, we employed the mummichog algorithm of MetaboAnalyst to identify global pathway activity changes in CPT2^H-KO and CPT2/USP30^H-KO mice^35,62. Interestingly, we observed a significant impact on amino acid metabolism in the CPT2/USP30^H-KO animals compared to CPT2^H-KO mice (Fig. 6b, and Supplementary Data 1). Consistent with the metabolomics profile, RNA-Seq analysis also reveals a significant alteration in gene signatures associated with amino acid metabolism following USP30 deletion (Fig. 6c). Specifically, the metabolism of Proline, Glutamine Phenylalanine, Tyrosine, and Branch-Chain Amino Acids (BCAA) exhibited the most substantial changes (Fig. 6d, e). As the heart undergoes a decline in FAO and experiences increased mitochondrial damage, glucose and glycolysis may emerge as the primary means of ATP production (Fig. 6d). However, this process yields less energy per molecule and produces two protons per molecule of glucose consumed⁶³. The resulting energy deficit and acidification may lead to a lower pH, eventually inhibiting glycolysis. Amino acids can function as substrates to replenish the tricarboxylic acid cycle. With augmented mitophagy and improved mitochondrial quality, amino acids may become more crucial as a fuel source. Notably, glutamine can be converted to α-ketoglutarate⁶⁴, maintaining the levels of TCA cycle intermediates (Fig. 6d, e). Recent studies also reveal the critical role of BCAA in regulating cardiac metabolism and stress response⁶⁵. Although these fuels may not sustain the cardiac metabolic demand for an extended period, any improvement in cardiac function could potentially lead to lower mortality, as observed in CPT2/USP30^H-KO mice (Fig. 5l). In line with this, in FAO-deficient hearts, the deletion of USP30 significantly impacted the intracellular levels of TCA cycle intermediates (Fig. 6d, e).

Prompted by these findings, we examined whether USP30 deletion in the setting of CPT2 deficiency enhances mitochondrial metabolism and oxidative capacity. Seahorse assays using palmitoyl-carnitine as a substrate confirmed that FAO-driven respiration was markedly reduced in mitochondria from both CPT2^H-KO and CPT2/USP30^H-KO hearts (Supplementary Fig. 5b). However, respiration supported by pyruvate and glutamate was significantly increased in CPT2/USP30^H-KO mitochondria compared to CPT2^H-KO (Supplementary Figs. 5c, d), indicating enhanced utilization of non-FAO substrates and improved mitochondrial function. Furthermore, mitochondrial reactive oxygen species (ROS) levels were significantly reduced in CPT2/USP30^H-KO hearts (Supplementary Figs. 5e, f), suggesting that USP30 deletion mitigates oxidative stress likely via improved mitochondrial quality control. Together, these results support a model in which USP30 ablation enhances mitophagy in FAO-deficient hearts and promotes a metabolic switch toward the preferential use of amino acids for energy production, compensating for defective FAO.

Discussion

Preserving mitochondrial function and integrity through specific mitochondrial quality control mechanisms is critical to ensure normal cardiac function. Mitophagy, a specialized autophagic pathway that mediates the lysosomal clearance of damaged mitochondria, plays a pivotal role in cardiac mitochondrial quality control^11,66,67. However, our current understanding of mitophagy in pathological circumstances of FAO deficiency remains limited. Mitophagy acts as a cardio-protective mechanism, orchestrating the removal of dysfunctional mitochondria to prevent their accumulation and minimize potential cellular damage. When confronted with compromised fatty acid metabolism, mitophagy is expected to become a cellular response dedicated to removing damaged mitochondria, thereby alleviating metabolic stress and dysfunction. Contrary to our initial assumptions, our findings reveal a previously unrecognized role for disrupted FAO in restraining cardiac mitophagy, mediated through the regulation of the PINK1/Parkin signaling. We have elucidated that the impairment of FAO disrupts PINK1/Parkin-mediated mitophagy via the activation of PARL. The intricate interplay among FAO deficiency, PARL activation, and mitophagy unveils a complex regulatory network. We anticipate that the upregulation of PARL may be mediated by the pyruvate dehydrogenase kinases (PDKs), particularly PDK2, which has been previously identified for phosphorylating PARL and regulating β cleavage³⁹. These findings integrate cardiac metabolism with mitophagy, emphasizing the significance of upholding optimal mitochondrial quality control, especially in the face of impaired myocardial FAO.

Compromised mitophagy in the heart may lead to detrimental effects on mitochondrial function, potentially exacerbating cardiac symptoms associated with FAO deficiency. Conversely, enhanced mitophagy could serve as a compensatory mechanism to mitigate mitochondrial damage within the context of these metabolic disorders. We have demonstrated that the deletion of USP30 activates stress-induced mitophagy and improves cardiac resilience to metabolic stress. USP30 inhibition may promote OMM ubiquitination, potentially regulating the ubiquitin threshold required for mitophagy activation^58,59,60,61. Improved mitophagy following USP30 deletion could potentially reverse certain aspects of cardiac dysfunction in FAO-deficient mice. It is noteworthy that USP30 can presumably have functions beyond mitophagy^25,58,59,68. Recent studies, for instance, indicate a crucial role for USP30 in tumor metabolism⁶⁹. In this context, we also explored how USP30 deletion might influence the metabolomic profile of the heart (Supplementary Fig. 5g). Despite observed differences, which likely stem from a few outlier features, PCA revealed minimal alterations in the metabolic or lipidomic profile between wild-type and USP30^H-KO hearts (Supplementary Fig. 5g), suggesting that USP30 deletion alone does not markedly alter the overall metabolism of the heart.

As assessed by mt-Keima analysis, the basal mitophagy levels were comparable between USP30^H-KO and wild-type animals. A similar absence of a basal phenotype occurs in Parkin-deficient mice, which is not surprising, given the apparent critical role of the balance between E3 ubiquitin ligases, such as Parkin, and the activity of mitochondrial deubiquitinases, exemplified by USP30^25,70. In energetically demanding tissues such as the heart, this mitophagic removal would presumably require exquisite regulatory controls. An optimal balance is crucial to prevent the heart from becoming maladapted to its environment, with both excessive and insufficient mitophagic removal posing potential risks. Consequently, the absence of a significant basal disparity in mitophagy levels in the absence of cardiac stress in USP30-deficient mice is not unexpected. Of particular interest is the observation that, in the context of FAO deficiency, the genetic deletion of USP30 restores the mitophagic response, thereby contributing to the pathophysiological response to cardiac stress. This bears significance, as the ultimate response to USP30 genetic deletion offers valuable insights for potential pharmacological interventions. The variations in phenotype resulting from the same gene deletion may reflect stress condition-dependent differences in the induction of mitophagy through the inhibition of USP30.

The specific manifestations and severity of cardiomyopathy among individuals with CPT2 deficiency exhibit a broad spectrum³². Factors such as the extent of enzyme deficiency, genetic variations, and environmental determinants collectively influence the development and progression of cardiac complications^9,31,32. Managing cardiomyopathy in CPT2 deficiency necessitates a comprehensive, multidisciplinary approach, incorporating interventions such as dietary modifications, avoidance of fasting, and supplementation with alternative energy sources^9,31,32. Ongoing research endeavors seek to elucidate the underlying mechanisms that link CPT2 deficiency to cardiomyopathy, with the ultimate goal of developing targeted therapeutic interventions capable of alleviating cardiac complications. Activation of cardio-protective mitophagy emerges as a promising and innovative strategy for addressing diverse cardiac symptoms associated with FAO deficiency. Notably, the use of a specific inhibitor targeting USP30 may represent an appealing therapeutic approach to enhance mitochondrial function in FAO-deficient hearts, particularly considering the clinical impracticality of direct genetic manipulation of this pathway. Future investigations could explore the efficacy of combining specific dietary interventions, such as a ketogenic diet or a coconut-oil diet rich in Medium-Chain Fatty Acids (MCFAs), with USP30 deletion. This combination approach holds the potential for increased effectiveness in rescuing CPT2^H-KO phenotypes compared to USP30 deletion alone. Intriguingly, previous studies suggest that a ketogenic diet may ameliorate pathological cardiac remodeling^71,72,73. Collectively, our findings highlight opportunities for addressing rare FAO-deficient cardiomyopathies. They also suggest a potential strategy for treating heart failure precipitated by a myriad of diverse etiologies, since these more prevalent conditions are often associated with altered cardiac metabolism characterized by impaired FAO^1,7,74. As such, mitophagy-directed therapies, by restoring mitochondrial function, have the potential to improve cardiac outcomes across a broad population of patients affected by heart failure.

Methods

Ethics declaration

All animal studies followed the Guide for the Care and Use of Laboratory Animals (NIH Publication, 8th Edition, 2011) and all experiments involving animals were approved by the Institutional Animal Care and Use Committee at The Ohio State University.

Mice

The mice were housed in ventilated cages under controlled environmental conditions. The housing facility maintained a 12 h light/dark cycle (lights on at 6 AM), with an ambient temperature of 20–24 °C and relative humidity of 40–60%. Animals had ad libitum access to standard chow food and water unless otherwise specified. Both male and female mice were used in this study. We have previously described the mt-Keima mouse⁷⁵. The mt-Keima mouse line was backcrossed for over 10 generations and maintained on a C57BL/6 J background. CPT2^H-KO mice were generated by crossing the CPT2^fl/fl mice^8,33 with transgenic mice expressing Cre recombinase under the control of the cardiac-specific alpha myosin-heavy chain (Myh6) promoter³⁴. USP30^fl/fl mice were obtained from the European Mouse Mutant Archive (EMMA, EM:09734, C57BL/6N-A^tm1Brd Usp30^{tm2a(EUCOMM)Hmgu}/WtsiH). Echocardiographic measurements were taken using a Vevo3100 Visual Sonics (Visual Sonics) system. The mice were lightly anesthetized with isoflurane, and the ejection fraction (EF), fractional shortening (FS), and ventricular chamber dimensions were determined using 2-D-guided M-mode images. EF, FS, ventricular chamber dimensions, and left ventricular mass were calculated automatically using the VevoLAB program.

Cell Culture

HeLa cells were grown in Dulbecco’s minimum essential medium (DMEM) with 10% fetal bovine serum (FBS) supplemented and 1% penicillin-streptomycin.

Western blotting

Tissues were lysed in RIPA buffer (50 mM Tris-HCl, at pH 8.0; 150 mM NaCl; 1% (vol/vol) Nonidet P-40; 0.5% sodium deoxycholate, 0.1% SDS and protease inhibitor cocktail (Roche) on ice. Primary antibodies were used at the following concentrations: USP30 (Santa Cruz, sc-109455, 1:200); TOM40 (Proteintech, 18409-1-AP, 1:1000); TIM23 (Proteintech, 11123-1-AP, 1:1000); TIM22 (Proteintech, 14927-1-AP, 1:1000); PINK1 (Cayman Chemical, 10006283, 1:500); PDK1 (Proteintech, 10026-1-AP, 1:1000); PDK2 (Proteintech, 15647-1-AP, 1:1000); PDK3 (Proteintech, 12215-1-AP, 1:1000); PDK4 (Proteintech, 12949-1-AP, 1:1000); CPT2 (Abcam, ab181114, 1:1000); OPTN (Proteintech, 10837-1-AP, 1:1000); p62/SQSTM1 (Abnova, H00008878-M01, 1:1000); PGAM5 (Abcam, ab126534, 1:1000), PARL (Abcam, ab118554, 1:1000); GAPDH (Cell Signaling Technology, 51332, 1:1000); Ubiquitin (MilliporeSigma, 04-263, 1:1000); Parkin (Proteintech, 14060-1-AP, 1:1000); GAPDH (Cell Signaling Technology, 51332, 1:1000); ACTIN (Cell Signaling Technology, 3700S, 1:1000). The membranes were incubated with anti-rabbit (LI-COR, 926-32211, 1:15000) or anti-mouse (LI-COR, 926-68072, 1:15000) IgG secondary antibodies for 1 h at room temperature. Images were captured using the Odyssey system (LI-Cor).

Transmission electron microscope

Hearts from wild-type or CPT2^H-KO and CPT2/USP30^H-KO mice were dissected and fixed in 2.5% glutaraldehyde (product #18426, Ted Pella Inc., Redding, CA) in 0.1 M phosphate buffer. Samples were postfixed with 1% osmium tetroxide (product #18456, Ted Pella Inc.) and then en bloc stained with 1% aqueous uranyl acetate (product #19481, Ted Pella Inc.), dehydrated in a graded series of ethanol and 100% acetone, and embedded in Eponate 12 epoxy resin (product #18012, Ted Pella Inc.). Ultrathin sections were cut with a Leica EM UC7 ultramicrotome (Leica Microsystems Inc., Deerfield, IL) and collected on copper grids. Images were acquired with an FEI Technai G2 Spirit BioTwin transmission electron microscope (Thermo Fisher Scientific, Waltham, MA) operating at 80 kV, and a Macrofire digital camera (Optronics, Inc., Chelmsford, MA) and AMT image capture software (Advanced Microscopy Techniques, Woburn, MA).

Mitochondrial cristae volume density was determined using NIH ImageJ (v1.53). The outer mitochondrial membrane and individual cristae membranes were manually traced to calculate their respective areas. The total cristae area was divided by the mitochondrial outer membrane area and multiplied by 100 to yield cristae volume density (%). Mitochondrial cristae morphology was assessed using a standardized scoring system ranging from 0 (severely disrupted) to 4 (normal), as previously described⁷⁶. In brief, each mitochondrion was evaluated based on cristae abundance and structure: 0-no clearly defined cristae; 1- > 50% of the area lacking cristae; 2- > 25% lacking cristae; 3-cristae in > 75% of the area but irregular in form; 4-abundant, well-organized cristae⁷⁶.

Immunofluorescence

For immunofluorescence, fresh frozen tissue sections embedded in OCT were cut onto slides at a thickness of 12 μm. Tissue samples were fixed in 4% PFA in PBS for 10 min, permeabilized with 0.5% Triton X-100 for an additional 15 min at room temperature and then blocked for 1 h with PBS + 3% BSA. After three washes with PBS, samples were incubated overnight at 4 °C with Wheat Germ Agglutinin (WGA) Alexa Fluor 488 (Invitrogen, W11261). The slides were sealed by coverslips and mounted with mounting medium (Invitrogen, P36934) before imaging.

Transcriptomics

Related to Fig. 3, RNA was isolated from cardiac samples of wild-type or CPT2^H-KO mice (n = 5) with the PureLink RNA Mini Kit (Thermo Fisher). Genomic DNA was removed from RNA preparations by DNAse digestion using RNase-Free DNase Set (QIAGEN) according to the manufacturer’s instructions. Sequencing libraries were constructed using Illumina’s TruSeq Stranded Total RNA kit with Ribo-Zero following the manufacturer’s instructions. The fragment size of RNA-Seq libraries was verified using the Agilent 2100 Bioanalyzer (Agilent), and the concentrations were determined using the Qubit instrument (LifeTech). The libraries were loaded onto the Illumina HiSeq 3000 for 2 × 75 bp paired-end read sequencing and generated about 60 M reads per sample. The fastq files were generated using the bcl2fastq software for further analysis.

Related to Fig. 6, total RNA was isolated from mouse cardiac samples of wild-type, CPT2^H-KO or CPT2/USP30^H-KO mice (n = 3) and validated on the Agilent Technologies 2100 bioanalyzer for quality control. Oligo(dT)-attached magnetic beads were used to purify mRNA, which was fragmented into small pieces using a fragmentation reagent. For cDNA synthesis, first-strand cDNA was generated using random hexamer-primed reverse transcription, then was followed by a second-strand cDNA synthesis. The synthesized cDNA was subjected to end-repair and then was 3′ adenylated. Adapters were ligated to the ends of these 3′ adenylated cDNA fragments. The cDNA fragments were amplified by PCR, and the products were purified with XP Beads and then dissolved in EB solution. The product was validated on the Agilent Technologies 2100 bioanalyzer. The double-stranded PCR products were heat-denatured and circularized by the splint oligo sequence. The single-strand circle DNA (ssCir DNA) was formatted as the final library. The final library was amplified with phi29 to make DNA nanoball (DNB), which had more than 300 copies of one molecular. The DNBs were loaded into the patterned nanoarray, and single-end 50 (pair-end 100/150) bases reads were generated in the way of combinatorial Probe-Anchor Synthesis (cPAS).

Different gene expression (DEG) analysis was performed on the clean reads with DESeq2 v1.42.0. Heatmaps were generated with the R package pheatmap. Up- and down-regulated genes were determined by fold change and significance, and subsequent KEGG- and GO-enrichment was performed with clusterProfiler v4.10.0. Unweighted Gene Set Enrichment Analysis (GSEA) was performed on a pre-ranked gene list using the GSEA desktop software v4.3.3.

Proteomics

Protein levels were compared in wild-type and CPT2^H-KO hearts (n = 4) using quantitative tandem mass tags (TMT10plex #90110, Thermo Fisher). In brief, hearts were homogenized using Precellys homogenizing beads in the following buffer: 280 mM sucrose, 10 mM HEPES, 1 mM EGTA, and protease inhibitor. Buffer was then supplemented with 1% maltoside, and samples were spun down at 10,000 x g for 10 min. Protein concentration was determined using the BCA Protein Assay Kit, and 100 µg of each sample was precipitated overnight with acetone (4 °C). Protein was resuspended in 0.5 M TEAB, reduced with 12 mM TCEP, and alkylated with 19 mM iodoacetamide. Samples were digested overnight with 10 µg trypsin (37 °C). All samples were added with their assigned TMT channels. The TMT-added samples were incubated at room temperature for 1 h for labeling. Tag labeling was quenched by adding 5% hydroxylamine, and protein samples were acidified with 1% trifluoroacetic acid. All labeled samples were combined and cleaned up with detergent-removal spin columns (Pierce #87777). The resulting mixture was further desalted using HLB cartridges (Oasis #WAT094225). The eluted sample was then fractionated into 12 fractions using high-pH reversed-phase liquid chromatography. Each fraction was dried by SpeedVac and resuspended in 0.1% formic acid for mass spec analysis on an Orbitrap Fusion (Thermo Fisher Scientific). Identified peptides were then quantified based on the intensities of TMT reporter ions using Proteome Discoverer (Thermo Fisher Scientific).

Confocal microscopy

Fluorescent samples were examined with a Zeiss LSM 780 confocal microscope (Carl Zeiss MicroImaging). As previously described⁷⁵, the fluorescence of mt-Keima was imaged in two channels via two sequential excitations (458 nm, green; 561 nm, red) and using a 570–695 nm emission range. Representative confocal images were processed using Imaris software by contrast linear stretch only. Calculation of mitophagy based on mt-Keima signal was performed using Zeiss ZEN software as previously described⁷⁵. The images were analyzed on a pixel-by-pixel basis. Each pixel’s intensity level from both channels was plotted on a scatter diagram, with green intensity on the x-axis and red intensity on the y-axis. Pixels with high red intensity are identified by the software using crosshairs and quadrants, while those with low intensity in both channels are considered background and excluded from the analysis. The fold change in mitophagy was calculated by comparing the number of pixels with high red intensity to the total number of pixels. The average of four images from each tissue sample was taken, and the values were normalized to the average value seen in the controls, assigned the value of one.

Untargeted metabolomics and lipidomics analysis

Heart tissues were pre-weighed and an extraction volume of MeOH was added to each sample at a ratio of 5 µL/mg of tissue. The samples were homogenized in a Precellys 24 soft tissue bead tube for 10 cycles consisting of 30 s on and 30 s off, followed by a centrifugation step for 5 min at 4 °C.

For metabolomics analysis (n = 3 with 3 technical replicates each), 250 µL of supernatant was dried down and reconstituted in 100 µL of 5% MeOH with 0.1% formic acid, and then placed in LC vials for LCMS analysis. The samples were injected at 5 µL onto a Thermo Scientific Vanquish Horizon using an Agilent Poroshell 120 SB-C18 column (2.1 × 100 mm, 2.7 µm particle size). The solvent system consisted of 100% water with 0.1% formic acid for solvent A and 100% MeOH with 0.1% formic acid for solvent B. A flow rate of 200 µL/min was run throughout with an initial gradient of 5% solvent B with a linear increase to 95% B by 15 min, holding 95% until minute 16, back down to 15% by minute 17, and ending the run at 5% at minute 30. Ionization of samples following LC separation was done with a Heated Electrospray ionization (HESI) source. Runs were performed in positive and negative modes with ESI voltages of 4.5 kV and 4 kV, respectively. The sheath gas was set to 15, the auxiliary gas to 5, and the capillary temperature to 320 °C. MS data collection was performed using Data-Dependent Analysis with a mass range of 80–1200 m/z, an MS resolution of 70,000, an MSMS resolution of 35,000, and 30-second dynamic exclusion with top 10 ion selection in each MS cycle.

For lipidomic analysis (n = 5 with 2 technical replicates each), 300 µL of supernatant was put aside and 300 µL of CHCl₃ was added to the homogenization tube and homogenized for 10 cycles again. This tube was then centrifuged, and 100 µL of the resulting CHCl₃ extract was combined with 100 µL of the first MeOH extract. This combined solution was then dried down and reconstituted in 50:50 MeOH: CHCl₃ before being placed in LC vials for LCMS analysis. The samples were injected at 5 µL onto a Thermo Scientific Vanquish Horizon using an Agilent Poroshell 120 SB-C18 column (2.1 × 100 mm, 2.7 µm particle size). The solvent system consisted of 60:40 water: acetonitrile with 13 mM ammonium formate for solvent A and 90:10 IPA: ACN with 13 mM ammonium formate for solvent B. A flow rate of 200 µL/min was run throughout with an initial gradient of 32% solvent B until 1.5 min then a linear increase to 45% B ending at minute 4, another increase to 52% ending at minute 5, up to 55% by minute 8, up to 60% by minute 11, up to 70% by minute 14, up to 75% by minute 18, and up to 97% by minute 21, holding 97% until minute 25, back down to 32% by minute 27, and ending the run at 32% at minute 30. Ionization of samples following LC separation was performed using a HESI source. Runs were performed in both positive and negative modes with ESI voltages set at 4 kV and 3 kV, respectively. The sheath gas was set to 15, the auxiliary gas to 5, and the capillary temperature to 320 °C. MS data collection was performed using Data-Dependent Analysis with a mass range of 106–1600 m/z, an MS resolution of 70,000, an MSMS resolution of 35,000, and 30 s dynamic exclusion with top 10 ion selection in each MS cycle.

For data analysis, the files were uploaded into Thermo Compound Discoverer 3.3. Metabolomics data was processed using online databases including mzCloud, mzVault, and Chemspider, and applying mzLogic. Lipidomics data processing also included LipidBlast and LipiMaps database comparisons for all tentative identifications. All datasets included Metabolika pathway analysis and applied SERRF QC corrections and background compound removal using pooled QC samples. In addition, lipidomic and metabolomic data, including the m/z features, p values, and statistical scores, were integrated into the MS Peaks to Pathways module of MetaboAnalyst 5.0, and the mummichog algorithm was then applied to generate mummichog pathway analysis plots with a global lipid network^35,62. Accession numbers of detected metabolites (HMDB, PubChem, and KEGG Identifiers) were generated, manually inspected, and utilized to map the canonical pathways. The mummichog pathway analysis plots display all the matched pathways as circles, with the color and size of each circle corresponding to its p-value and enrichment factor, respectively, where the enrichment factor is the ratio between the number of significant (p < 0.05) pathway hits and the expected number of pathway hits.

Isolated mitochondrial reactive oxygen species (ROS) measurement

Amplex Red was used to detect ROS generation by cardiac mitochondria using a fluorescence spectrophotometer (Hitachi F-7100). Briefly, mouse hearts were rapidly excised and homogenized in ice-cold ROS Buffer A (210 mM mannitol, 70 mM sucrose, 50 mM Tris-Base, 1 mM EDTA-2Na, pH7.4). The homogenate was centrifuged at 2500 × g for 10 min at 4 °C. The supernatant was transferred to a clean 1.7 mL microcentrifuge tube and centrifuged at 12,000 x g for 10 min 4 °C. The mitochondrial pellet was resuspended in ice-cold ROS Buffer B (210 mM mannitol, 70 mM sucrose, 50 mM Tris-Base, and 0.1 mM EDTA, pH 7.4) and centrifuged at 12,000 x g for 10 min at 4 °C. The supernatant was discarded, and the mitochondrial pellet was resuspended in ROS Buffer C (250 mM sucrose, 20 mM Tris-Base, 1 mM EGTA, 1 mM EDTA, pH 7.4, 0.15% (w/v) BSA) followed by a final centrifugation step at 12,000 × g for 10 min at 4 °C. The final mitochondrial pellet was resuspended in ROS buffer C for use in the ROS detection assay. For ROS detection 5 μg horseradish peroxidase (Sigma-Aldrich) was added to 2 mL of ROS Buffer C. Baseline fluorescence was recorded at 37 °C using a fluorescence spectrophotometer (excitation: 560 nm; emission: 590 nm). After 60 s, 10 μM Amplex Red (Life Technologies) was added, followed by 25 μL of isolated mitochondria after an additional 60 s. Mitochondrial activity was stimulated by the addition of 3 mM succinate (Sigma-Aldrich) 90 s later. Antimycin A (2.5 μM), an inhibitor of complex III, was added at 1100 s as a control to enhance ROS production. Fluorescence was continuously recorded for 2010 seconds at 0.5-second resolution. ROS production was plotted as fluorescence intensity over time and normalized to mitochondrial protein concentration. Total ROS production was quantified by calculating the area under the fluorescence-time curve (AUC) using GraphPad Prism 10.

Measurement of mitochondria respiration

Mitochondria were isolated from cardiac tissue by homogenization in MSHE buffer (70 mM sucrose, 210 mM mannitol, 1 mM HEPES, 1 mM EGTA, 0.5% fatty acid-free BSA; pH 7.2) at 4 °C using a KIMBLE ® Potter-Elvehjem tissue grinder and glass pestle with 5–10 strokes at 2500 rpm, followed by centrifugation at 800 × g for 10 min. The supernatant was filtered through a 40 µm Nylon cell strainer and centrifuged at 8000 × g for 10 min at 4 °C. The resulting pellet was resuspended in MSHE buffer and centrifuged at 8000 × g for 10 min. The pellet was then washed in MSHE buffer without BSA, centrifuged at 8000 × g for 5 min at 4 °C, and the final mitochondrial pellet was used for respiration assays. Mitochondrial respiration was assessed by measuring Oxygen Consumption Rates (OCRs) using the XF96 Analyzer (Agilent)^5,77. Briefly, isolated cardiac mitochondria were resuspended in 1× mitochondrial assay buffer (MAS; 70 mM sucrose, 220 mM mannitol, 10 mM KH₂PO₄, 5 mM MgCl₂, 2 mM HEPES, 1 mM EGTA, and 0.2% fatty acid-free BSA; pH 7.2). A total of 6 μg of mitochondria was seeded per well in the presence of palmitoyl carnitine/malate (0.04 mM/0.8 mM), pyruvate/malate (10 mM/2 mM), or glutamate/malate (10 mM/2 mM). Basal OCR was recorded at three consecutive time points at 37 °C, followed by measurement of state 3 respiration, initiated by the addition of ADP (1 mM). OCR values were normalized to mitochondrial protein concentration for each sample.

Statistics

All data were expressed as mean ± s.d. In comparisons between two groups with equal variance, statistical analysis was performed with an unpaired 2-tailed t test. When comparing multiple groups, One-way and repeated measures analysis of variance (ANOVA) was used to evaluate the statistical significance. Heart weight and echocardiography were measured in a blinded manner.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The RNA-seq data sets generated for this study are in the Gene Expression Omnibus (GEO) under accession number GSE266487. The mass spectrometry data have been deposited to the ProteomeXchange via the PRIDE partner repository with the dataset identifier PXD052216. Source data are provided in this paper.

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Acknowledgements

We thank Dr. Richard Youle for generously providing us with the PARL KO cells. We thank the Wellcome Trust Sanger Institute Mouse Genetics Project (Sanger MGP) and its funders for providing the USP30^f/f mouse line. We thank Dr. Michael Wolfgang for providing the CPT2^f/f mouse line. We thank Matthew Bernier (Mass Spectrometry and Proteomics Facility, The Ohio State University) for his help in LC-MS and the metabolomic analysis. The authors acknowledge resources for confocal microscopy from the Campus Microscopy and Imaging Facility (CMIF), and the OSU Comprehensive Cancer Center (OSUCCC) Microscopy Shared Resource (MSR) at the Ohio State University with NIH S10 OD025008 and NIH NIC P30CA016058. This work was supported by National Institutes of Health (NIH) grants HL135051 and HL160581 (N.S.). H.S. and D.K. were supported by T32HL134616, K.B. was supported by R01HL166520.

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Department of Physiology and Cell Biology, The Ohio State University Wexner Medical Center, Columbus, Ohio, USA
Nuo Sun, Hayley Barta, Samhita Chaudhuri, Kangxuan Chen, Hongke Luo, Mingchong Yang, Judith Krigman, Ruohan Zhang, Shridhar Sanghvi, Hannah Sanders, Dominic Kolonay, Mudra Patel, Kedryn Baskin, Harpreet Singh & Pengyi Zhang
Dorothy M. Davis Heart and Lung Research Institute, The Ohio State University Wexner Medical Center, Columbus, Ohio, USA
Nuo Sun, Hayley Barta, Samhita Chaudhuri, Kangxuan Chen, Shridhar Sanghvi, Hannah Sanders, Dominic Kolonay, Mudra Patel, Kedryn Baskin & Harpreet Singh
Department of Microbial Infection and Immunity, The Ohio State University Wexner Medical Center, Columbus, OH, USA
Jiacheng Jin & Gang Xin
Aging Institute, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA
Shiori Sekine & Toren Finkel

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Nuo Sun
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Hayley Barta
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Samhita Chaudhuri
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Kangxuan Chen
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Jiacheng Jin
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Gang Xin
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Contributions

N.S. and T.F. conceived the study. N.S., H.B., S.C., K.C., H.L., M.Y., J.K., R.Z., S.Sa., S.Se., H.S., D.K., M.P., K.B., H.S. and P.Z. performed experiments and/or analyzed or interpreted data. J.J. and G.X. analyzed transcriptomic data. N.S. wrote the paper. All authors read and approved the final version of the manuscript prior to submission for publication.

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Correspondence to Nuo Sun.

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Competing interests

T.F. is a co-founder and stockholder in Generian Pharmaceuticals and Coloma Therapeutics. The other authors declare no competing interests.

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Nature Communications thanks Rong Tian and the other anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Sun, N., Barta, H., Chaudhuri, S. et al. Mitophagy mitigates mitochondrial fatty acid β-oxidation deficient cardiomyopathy. Nat Commun 16, 5465 (2025). https://doi.org/10.1038/s41467-025-60670-z

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Received: 16 April 2024
Accepted: 29 May 2025
Published: 01 July 2025
DOI: https://doi.org/10.1038/s41467-025-60670-z