SIRT2 functions as an N-acetylaspartate hydrolase that counteracts cardiac metabolic remodeling in kidney disease

SIRT2 functions as an N-acetylaspartate hydrolase that counteracts cardiac metabolic remodeling in kidney disease | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Biological Sciences - Article SIRT2 functions as an N-acetylaspartate hydrolase that counteracts cardiac metabolic remodeling in kidney disease Jian-Yuan Zhao, Ze-Yu Zhou, Xi-Ji Qin, Jiang-Yu Yan, Ke Cai, Xin Sun, and 6 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8510072/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract N-acetylated amino acids systemically accumulate in the circulatory system in patients with chronic kidney disease (CKD), raising questions about their roles in peripheral tissues1,2. Here, targeted metabolomics in patients with CKD identified that circulating N-acetylaspartate (NAA) as the metabolite most closely linked to early cardiac dysfunction. In mice, both CKD and exogenous NAA administration induced pronounced cardiac NAA accumulation, resulting in systolic dysfunction and pathological hypertrophy. Activity-based protein profiling identified cytosolic malate dehydrogenase 1 (MDH1) and sirtuin 2 (SIRT2) as direct targets of NAA. NAA acts as a substrate analogue that occupies the MDH1 substrate-binding pocket, competitively inhibits its enzymatic activity, and disrupts the malate–aspartate shuttle, thereby lowering cytosolic and mitochondrial NAD⁺/NADH ratios, suppressing tricarboxylic acid cycle flux, and compromising cardiomyocyte energy metabolism. Notably, we identified SIRT2 as a previously unrecognized NAA hydrolase that specifically binds and hydrolyzes NAA in an NAD⁺-dependent manner, thereby mitigating NAA-induced metabolic stress. Cardiomyocyte-specific restoration of SIRT2 activity in mice reduced cardiac NAA levels, improved systolic dysfunction, and attenuated hypertrophy. These findings expand sirtuins from protein deacylases to direct regulators of small-molecule metabolites and establish NAA as a kidney–heart metabolic mediator, revealing an amino acid acetylation-dependent layer of energy homeostasis. Biological sciences/Biochemistry/Enzyme mechanisms Health sciences/Diseases/Cardiovascular diseases Figures Figure 1 Figure 2 Figure 3 Figure 3 Figure 4 Figure 4 Figure 5 Introduction N -acetylated amino acids have generally been regarded as terminal metabolic derivatives, and their circulating levels rise in chronic kidney disease (CKD) due to impaired renal clearance 1,3-5 . CKD-associated cardiomyopathy involves complex cardiac remodelling partly driven by the accumulation of circulating uremic toxins 2 , and offers a relevant endogenous disease context to investigate the physiological roles of N -acetylated amino acids. Protein acetylation is one of the major post-translational modifications involved in many key cellular processes 6,7 ; however, whether N -acetylated amino acids act as functional modulators of cellular metabolism remains unknown. Among these, N -acetylaspartate (NAA) is notable for being the second most abundant metabolite in the central nervous system 8,9 , yet its functional roles outside the brain remain poorly understood. In this context, whether elevated circulating NAA acts as a signalling metabolite and contributes to cardiac metabolic remodelling remains unknown. In mammals, the seven sirtuins (SIRT1–SIRT7) constitute an evolutionarily conserved family of NAD + -dependent enzymes with diverse biological functions 10-19 . These enzymatic activities share a common mechanism of amide bond hydrolysis, suggesting the potential existence of additional, yet-undiscovered enzymatic functions. This study investigates the relationship between circulating NAA levels and cardiac metabolic homeostasis. Mechanistically, we show that NAA suppresses the activity of malate dehydrogenase 1 (MDH1), thereby impairing mitochondrial NADH transport and cardiac metabolism. Moreover, we discover that SIRT2 is a previously unrecognized NAA hydrolase that mitigates NAA-induced metabolic disturbances. Altogether, these findings reveal the detrimental impact of NAA on cardiac metabolic homeostasis and broaden the functional landscape of sirtuins. Results NAA levels are elevated and associated with cardiac impairment in CKD To investigate the potential association between circulating N -acetylated amino acids and cardiac impairment, we analyzed a well-characterized cohort of 62 patients with CKD. Participants were divided by plasma B-type natriuretic peptide (BNP) levels, using a threshold ≥ 200 ng/L as a marker of early cardiac dysfunction 20 . Individuals above this threshold exhibited reduced left ventricular ejection fraction (LVEF), enlarged left ventricular and left atrial dimensions, and increased interventricular septal and posterior wall thickness, consistent with early cardiac remodelling and subclinical dysfunction ( Extended Data Table 1 ). Targeted metabolomics analysis of circulating metabolites, including 12 circulating N -acetylated amino acids, revealed that NAA and N -acetylglutamate (NAG) levels were significantly elevated in individuals with high BNP levels, whereas their unmodified precursors, aspartate and glutamate, remained unchanged ( Fig. 1a, Extended Data Fig. 1a, b ). Among all N -acetylated amino acids measured, NAA and NAG showed the strongest correlations with BNP levels, exceeding those of known risk factors for CKD-associated cardiomyopathy, such as p-cresyl sulfate and indoxyl sulfate 21 ( Fig. 1b, Extended Data Fig. 1c, d ). Together, these results indicate that elevated circulating NAA and NAG levels are closely linked to early cardiac dysfunction in CKD. To investigate whether dysregulation of N -acetylated amino acids contributes to CKD-associated cardiomyopathy, we employed the 5/6 subtotal nephrectomy model. Eight weeks post-surgery, mice exhibited elevated blood urea nitrogen (BUN) levels, confirming renal impairment, accompanied by systemic accumulation of N -acetylated amino acids and cardiac dysfunction, including systolic impairment and myocardial hypertrophy. ( Extended Data Fig. 2a-e ). Only NAA and NAG levels increased significantly in the heart, with NAA displaying the most pronounced elevation ( Fig. 1c, Extended Data Fig. 2f ). This pattern mirrors our observations in the human cohort ( Fig. 1a, b) , supporting a potential link between these metabolites and early cardiac metabolic remodelling. NAG is a well-characterized allosteric activator of carbamoyl phosphate synthetase 1 (CPS1) in the urea cycle 22 . However, the biological functions of NAA remain largely unexplored. Considering the marked elevation of NAA levels in mice and humans, we focused the subsequent mechanistic studies on NAA. Tissue profiling of NAA‐metabolizing enzymes revealed that NAA was predominantly synthesized in non-cardiac tissues, where both its synthetase and hydrolase were highly expressed; however, the heart highly expressed the dicarboxylate transporter SLC13A3 but showed minimal hydrolase expression ( Fig. 1d ). Together, these findings indicate that renal injury drives systemic accumulation of N -acetylated amino acids, and results in disproportionate cardiac exposure to circulating NAA, owing to robust uptake capacity but limited hydrolytic activity in the heart. NAA impairs cardiac function and induces cardiac hypertrophy To investigate the effects of NAA on cardiac function, NAA was intraperitoneally injected in sham mice and mice with CKD. After eight weeks of treatment, echocardiographic analysis revealed impaired cardiac function in both groups ( Extended Data Fig. 3a ). NAA induced systolic dysfunction and myocardial hypertrophy in sham mice, with these deleterious effects being further exacerbated under CKD conditions ( Fig. 1e ). Analysis of hematoxylin and eosin (H&E)-stained tissues revealed that NAA treatment induced cardiac hypertrophy under physiological and pathological conditions ( Extended Data Fig. 3b ). This finding was confirmed by wheat germ agglutinin (WGA) staining results, which showed enlargement of cardiomyocytes following NAA administration ( Fig. 1f ). Moreover, Masson’s trichrome staining showed no significant fibrosis in the Sham + NAA group, suggesting that NAA alone did not induce cardiac fibrosis under normal conditions ( Fig. 1g ). In contrast, NAA treatment markedly aggravated interstitial collagen deposition in the CKD + NAA group, indicating that NAA exacerbates cardiac fibrosis in the CKD setting ( Fig. 1g ). Injection of NAA in sham and mice with CKD did not cause significant changes in renal function, as evidenced by comparable BUN levels and unaltered renal histology ( Extended Data Fig. 3c-e ); these findings indicated that the cardiac effects of NAA were not secondary to altered renal function. Overall, NAA induces maladaptive cardiac remodelling and functional deterioration, which is exacerbated in the context of renal dysfunction. Furthermore, NAA treatment led to a significant increase in the primary neonatal rat cardiomyocytes (NRCMs) cell area ( Fig. 1h ), and levels of atrial natriuretic peptide (ANP), BNP, and β-myosin heavy chain (β-MHC), which are molecular markers indicative of pathological hypertrophy ( Fig. 1i ). In contrast, treatment with aspartate did not upregulate the expression of these markers, indicating that the pro-hypertrophic effect of NAA is not mediated through its hydrolysis to aspartate ( Extended Data Fig. 3f ). Together, these data support the notion that NAA impairs cardiac function and promotes hypertrophy, contributing to cardiac remodelling at the organ and cellular levels. NAA binds to and inhibits MDH1 To elucidate the molecular mechanism by which NAA contributes to cardiac dysfunction, we employed an activity-based protein profiling (ABPP) approach using NAA probes as bait. Photoaffinity-tagged NAA probes were synthesized to enable enrichment and visualisation of interacting proteins ( Extended Data Fig. 4a ). The pro-hypertrophic effects of probe-tagged NAA in cardiomyocytes were comparable to those of non-tagged NAA ( Extended Data Fig. 4b ), validating the functional integrity probe-tagged NAA. The NAA probes were incubated with freshly prepared heart lysates for target labelling, and an equivalent concentration of non-tagged NAA was used as a control ( Extended Data Fig. 4c ). In the NAA probe group, 56 candidate proteins were specifically enriched ( Extended Data Fig. 4d ). The top 10 proteins were ranked according to their peptide-spectrum matches (PSMs) and coverage ( Fig. 2a ). Among the 10 candidate proteins, MDH1 was prioritized as a potential target because it is a key enzyme in the malate–aspartate shuttle 23-25 ; moreover, the aspartate-derived metabolite NAA is structurally similar to the MDH1 substrate oxaloacetate (OAA) ( Extended Data Fig. 4e ). In this shuttle, aspartate undergoes transamination to form OAA, which is subsequently reduced to malate (Mal) by MDH1 ( Extended Data Fig. 4f ). Drug affinity responsive target stability (DARTS) assay results revealed that incubation of purified recombinant MDH1 with NAA increased its resistance to Pronase digestion, suggesting that NAA binds to and stabilizes MDH1 ( Fig. 2b ). In contrast, no such protective effect was observed for purified recombinant MDH2, the mitochondrial isoform that converts Mal to OAA ( Extended Data Fig. 4g ). Since the cardiac dysfunction and hypertrophy phenotypes were more pronounced than fibrosis, and that MDH1 primarily functions in cardiomyocytes, we analysed the effects of NAA on cardiomyocytes. Tail-vein injection of adeno-associated virus 9 (AAV9) carrying the cardiac troponin T (cTnT) promoter–driven sh Mdh1 resulted in cardiomyocyte-specific knockdown of Mdh1 , which was consistent with the predominant expression of MDH1 in cardiomyocytes ( Extended Data Fig. 4h ). In Mdh1 -knockdown mice, NAA treatment did not exacerbate cardiac dysfunction compared with that in untreated knockdown mice, indicating that MDH1 is the primary target that mediates the effects of NAA on cardiac function ( Fig. 2c, Extended Data Fig. 4i) . Next, we elucidated the effects of NAA on MDH1 function. Surface plasmon resonance (SPR) assays were conducted to evaluate the binding affinities of OAA and NAA to MDH1 ( Fig. 2d, e ). The equilibrium dissociation constant ( K D ) of NAA (7.19 μM) was higher than that of OAA (2.03 μM), indicating that although NAA binds MDH1, its affinity to MDH1 is weaker than that of OAA; however, when MDH1 was pre-incubated with 10 μM NAA, the K D value for OAA binding increased to 8.62 μM, suggesting that NAA inhibits the interaction between OAA and MDH1 ( Fig. 2f, g ). To explore the molecular basis of NAA binding, we performed molecular docking and molecular dynamics simulations. NAA occupies the substrate-binding pocket of MDH1 26 , engaging key residues also involved in OAA recognition ( Fig. 2h ). The protein Root Mean Square Deviation (RMSD) remained stable for both MDH1-OAA (1.0–2.0 Å) and MDH1-NAA (0.75–2.0 Å) complexes ( Extended Data Fig. 5a, b ). OAA interacted with pocket residues mainly through hydrogen bonds (binding energy, -16.94 kcal/mol), whereas NAA engaged more residues via additional hydrogen bonds and water bridges, resulting in stronger binding than that of OAA (-36.00 kcal/mol) ( Extended Data Fig. 5c, d ). These simulations suggest that NAA engages MDH1 in a binding mode analogous to that of OAA but with more extensive interactions. NAA binding did not occur in the MDH1 quintuple mutant (R92A/R98A/N131A/R162A/S242A, MDH1-5A) ( Extended Data Fig. 5e ), confirming the critical role of these residues and validating the binding mode of NAA to MDH1; these residues correspond to high-frequency interaction sites and constitute the OAA-binding pocket 26 . Notably, OAA interacted frequently with NADH, whereas NAA did not ( Extended Data Fig. 5c, d ); the probable reason is that the amide carbonyl in NAA is resonance-stabilized and thus less reducible than the keto group in OAA ( Extended Data Fig. 4e ). This finding suggests that NAA may not readily undergo the reduction reaction catalysed by MDH1. Overall, NAA can engage MDH1, potentially interfering with OAA binding and enzymatic activity. To assess enzymatic inhibition, we conducted in vitro assays to monitor NADH consumption at 340 nm during MDH1-catalysed OAA reduction ( Fig. 2i, Extended Data Fig. 4 f ). NAA is not reduced by MDH1; however, NAA inhibits the catalysis of OAA by MDH1 ( Fig. 2i, j ). Together, these findings demonstrate that NAA occupies the substrate-binding pocket of MDH1 and inhibits its activity, providing a mechanistic basis for the effects of NAA on cardiac metabolism and function. NAA-induced MDH1 inhibition disrupts cardiomyocyte metabolic homeostasis To assess whether NAA inhibits MDH1 activity in primary cardiomyocytes, we measured intracellular levels of OAA and Mal following NAA treatment. The MDH1 substrate OAA accumulated, whereas Mal levels remained largely unchanged ( Fig. 3a, b ); the probable reason is that intracellular OAA exists at low concentrations and readily reflects perturbations, whereas Mal is generated and consumed through multiple metabolic pathways. Using the Mal/OAA ratio as a proxy for MDH1 enzymatic activity, we observed that NAA markedly inhibited MDH1 activity ( Fig. 3c ). Since the malate–aspartate shuttle is essential for transferring cytosolic reducing equivalents into mitochondria to sustain oxidative phosphorylation in cardiomyocytes 23-25 , we examined whether NAA disrupts metabolic homeostasis in primary cardiomyocytes. To assess the effects of NAA on redox homeostasis, we monitored NAD⁺/NADH dynamics in primary cardiomyocytes using non-targeted SoNar (cyto-SoNar) or mitochondria-targeted SoNar (mito-SoNar). Real-time measurements revealed that NAA treatment led to a decrease in the cytosolic NAD⁺/NADH ratio, while slowing the rate of NAD⁺/NADH increase in mitochondria ( Fig. 3d, e ). Consistently, at the whole-cell level, NAA treatment markedly decreased the overall NAD⁺/NADH ratio in cardiomyocytes ( Fig. 3f ). This metabolic impairment was accompanied by a significant decrease in intracellular ATP levels, reflecting compromised mitochondrial energy production ( Fig. 3g ). These findings indicate that NAA lowers cytosolic NAD⁺/NADH ratio and dampens the mitochondrial redox response, which is consistent with inhibition of the MDH1-dependent malate–aspartate shuttle. To elucidate how NAA perturbs central carbon metabolism, we performed 13 C 6 -glucose and 13 C 5 -glutamine tracing in primary cardiomyocytes to assess the effects of NAA on glycolysis and tricarboxylic acid (TCA) cycle activity ( Extended Data Fig. 5f, g ). After NAA treatment, levels of labelled upstream glycolytic intermediates glucose-6-phosphate (G6P), fructose-1,6-bisphosphate (FBP), and glyceraldehyde-3-phosphate (G3P) remained unaltered, whereas labelled 3-phosphoglycerate (3PG) and pyruvate M + 3 levels decreased ( Fig. 3h ). This finding aligns with the observed cytosolic NAD⁺/NADH decline caused by MDH1 inhibition, because glyceraldehyde-3-phosphate dehydrogenase (GAPDH) requires NAD⁺ for its activity. Moreover, acetyl-CoA M + 2 labeling remained unchanged, but incorporation of both 13 C 6 -glucose and 13 C 5 -glutamine into TCA cycle intermediates was markedly reduced, indicating suppression of mitochondrial oxidative metabolism ( Fig. 3i, j ). Accumulation of unlabelled OAA, predominantly derived from aspartate, suggested dysfunction of the cytosolic MDH1-mediated reaction ( Fig. 3k ). Collectively, these findings demonstrate that NAA inhibits MDH1 activity and disrupts redox-linked metabolite shuttling and mitochondrial metabolism, ultimately leading to energy insufficiency and cardiomyocyte hypertrophy. SIRT2 is an NAA hydrolase Beyond MDH1, our ABPP screen also identified SIRT2 and SIRT3 as NAA-binding candidates ( Fig. 2a ). Since sirtuins are NAD⁺-dependent enzymes with amide bond-hydrolysing activity 14-18 , this raised the possibility that certain sirtuins might recognise and hydrolyse NAA as a small-molecule amide substrate, in addition to their canonical protein deacetylase activity. To investigate whether members of the sirtuin family possess NAA hydrolase activity, we purified recombinant homo SIRT1–SIRT7 heterologously expressed in bacteria and conducted an enzyme-coupled assay to continuously monitor the hydrolysis process ( Extended Data Fig. 6a ). This assay is analogous to that used for measuring the deacetylase activity of sirtuins and uses NADPH consumption as a surrogate readout for the first-step reaction 27 . NADPH levels were quantified based on the intrinsic fluorescence (excitation at approximately 340 nm; emission at 460 nm). Among the seven sirtuins, SIRT2 and SIRT3 exhibited robust NAA hydrolase activity in an NAD + -dependent manner ( Fig. 4a, b ), whereas the others showed minimal or undetectable activity ( Extended Data Fig. 6b ). To validate the NAA hydrolase activity of SIRT2 and SIRT3, we incubated these enzymes with NAA and quantified NAA and its hydrolysis product, aspartate, using LC-MS/MS ( Fig. 4c ). NAA hydrolysis was determined to be a previously unrecognized enzymatic activity of SIRT2 and SIRT3, with estimated catalytic efficiencies ( K cat /K m ) of 1.39 × 10 4 and 2.97 × 10 4 s -1 M -1 , respectively ( Fig. 4d ). Subsequently, we examined whether SIRT2 and SIRT3 physically associate with NAA. DARTS assay results revealed that NAA increased the Pronase resistance of SIRT2 and SIRT3 ( Fig. 4e, f ), indicating the occurrence of a specific interaction, whereas no such effect was observed using aspartate ( Extended Data Fig. 6c, d ). To quantify the binding affinities, we performed SPR analysis. NAA exhibited concentration-dependent binding to SIRT2 and SIRT3 ( Fig. 4g, h ), with K D values of 6.32 and 2.85 μM, respectively ( Fig. 4i ). Since NAA treatment did not alter SIRT2 nuclear–cytosolic distribution 28 ( Extended Data Fig. 6e ) and MDH1 is a cytosolic enzyme, we focused on the hydrolytic activity of SIRT2 toward NAA, in contrast to that of SIRT3, which is predominantly localized in mitochondria 29 . To gain structural insights into the interaction between SIRT2 and NAA, we performed molecular docking and molecular dynamics simulations. NAA stably interacted with the L4 loop of SIRT2, positioning it within the active-site pocket of the enzyme 30,31 ( Extended Data Fig. 7a ). The overall protein structure remained stable during molecular dynamics simulations (RMSD: 1.6–3.0 Å; Extended Data Fig. 7b ), supporting a well-defined binding mode. NAA further interacted with key residues in the L1 and L5 loops involved in substrate recognition and entry 31 ( Extended Data Fig. 7c ), indicating a small-molecule binding mode stabilised by coordinated non-covalent interactions. In contrast, interactions between SIRT2 and its canonical protein substrate tubulin alpha-1A (TUBA1A) involve 93 residues across two spatially separated sites, enriched in charged and polar residues, forming extensive hydrogen-bond networks characteristic of protein–protein interactions ( Extended Data Fig. 7d, e ). Together, these observations suggest that SIRT2 can recognise and hydrolyse NAA, showing that NAA binds in a small-molecule-specific mode within the active pocket. In Sirtuin2 -global knockout mice, NAA levels were significantly elevated in the circulation and heart, suggesting that SIRT2 functions as a physiological NAA hydrolase in vivo ( Fig. 4j, k ). In contrast, renal NAA levels remained largely unchanged, likely due to the high expression levels of known NAA hydrolases in the kidney ( Fig. 1d and 4l ). Based on these results, we propose the mechanism of NAA hydrolysis by SIRT2. In this process, the acetyl group of NAA is transferred to NAD + , yielding nicotinamide, O -acetyl-ADP-ribose, and free aspartate ( Fig. 4m ). These findings support the conclusion that SIRT2 acts as a physiological NAA hydrolase. SIRT2-mediated hydrolysis of NAA relieves MDH1 inhibition and improves cardiac function We next investigated whether SIRT2-mediated hydrolysis of NAA could mitigate the detrimental effects of NAA on the heart. Overexpression of SIRT2 in primary cardiomyocytes significantly reduced NAA levels, confirming the physiological NAA-hydrolysing activity of SIRT2 ( Fig. 5a ). SIRT2 overexpression restored cytosolic and mitochondrial NAD⁺/NADH ratios, indicating alleviation of MDH1 inhibition ( Fig. 5b, c ). In addition, the hypertrophic phenotype of the cardiomyocytes was markedly alleviated ( Fig. 5d ). Overall, SIRT2-mediated NAA hydrolysis exerted a protective effect on cardiomyocyte metabolism and function. Given the reduction in cardiac SIRT2 expression levels in mine with CKD ( Extended Data Fig. 8a ), we next examined whether restoration of SIRT2 could counteract NAA-induced cardiac toxicity. To separately evaluate the ability of SIRT2 to rescue NAA-induced toxicity and rectify CKD-associated endogenous NAA elevation, we performed two parallel sets of experiments using exogenous administration and chronic disease models. Tail-injected AAV9 enabled cardiomyocyte-specific cTnT promoter–driven SIRT2 overexpression ( Extended Data Fig. 8b, c ). In control mice, SIRT2 overexpression effectively reversed the intraperitoneal administration-induced NAA accumulation in cardiac tissues and sera ( Fig. 5e, Extended Data Fig. 8d ), thereby restoring cardiac MDH1 activity and ATP levels ( Fig. 5f, g ). Similarly, in mice with CKD, SIRT2 overexpression normalized cardiac NAA levels ( Fig. 5h ), partially alleviated the decrease in MDH1 activity and ATP levels ( Fig. 5i, j ), and slightly reduced the elevated serum NAA levels ( Extended Data Fig. 8e ), likely due to the persistent systemic metabolic impairment associated with renal dysfunction. Furthermore, SIRT2 overexpression improved cardiac systolic performance ( Fig. 5k, l, Extended Data Fig. 8f, g ) and reduced cardiomyocyte hypertrophy ( Fig. 5m, n ) in NAA-treated mice and mice with CKD. Moreover, western blotting confirmed downregulation of hypertrophic markers in SIRT2-overexpressing hearts ( Fig. 5o, p ). Masson’s trichrome staining analysis revealed that NAA treatment alone did not result in cardiac fibrosis, which is consistent with previous observations (Fig. 1g, Extended Data Fig. 8h) ; moreover, SIRT2 overexpression markedly attenuated cardiac fibrosis in mice with CKD ( Extended Data Fig. 8i). Together, these results demonstrate that cardiomyocyte-specific SIRT2 overexpression counteracts exogenous NAA overload and CKD-associated endogenous NAA accumulation, thereby protecting against cardiac dysfunction and remodelling. Discussion We discovered that NAA is a pathogenic metabolic signal that links renal dysfunction to cardiac remodelling. NAA aberrantly accumulates under CKD conditions and drives cardiac dysfunction and cardiomyocyte hypertrophy. Mechanistically, NAA competitively inhibits MDH1, leading to disruption of mitochondrial metabolism and impairment of energy homeostasis in cardiomyocytes. Therefore, NAA is a maladaptive metabolic mediator in the kidney–heart axis. Moreover, SIRT2 is a previously unrecognized NAA hydrolase, which detoxifies excessive NAA and counteracts NAA-induced metabolic stress, cardiac dysfunction, and structural remodelling ( Extended Data Fig. 8j) . These findings underscore NAA as a potential mediator of kidney–heart metabolic crosstalk in a broad range of renal pathologies 32 . Further studies are warranted to determine whether circulating NAA levels predicts cardiac outcomes across kidney diseases and to evaluate its value as a diagnostic biomarker in cardiorenal syndromes. Moreover, SIRT2 exerts a cardioprotective effect by mitigating NAA-induced hypertrophic remodelling, suggesting that enhancing its activity may help preserve cardiac function under metabolic stress 33 . Sirtuins are NAD⁺-dependent metabolic sensors 10-12,34 , and we discovered that SIRT2 acts as an NAA hydrolase, expanding the known functions of sirtuins to include the regulation of small-molecule metabolites. This finding raises the possibility that SIRT2-mediated hydrolysis of NAA constitutes an adaptive response to energy fluctuations and modulates energy homeostasis in the heart and other high-energy-demanding tissues. Although identified in the heart, the SIRT2–NAA–MDH1 axis is likely to represent a general mechanism, as SIRT2 and MDH1 are widely expressed core metabolic enzymes. Dysregulation of the SIRT2–NAA–MDH1 axis may contribute to mitochondrial dysfunction-related diseases. Future studies need to delineate its regulatory circuitry and broader physiological and therapeutic relevance in metabolic disorders and aging. Besides metabolic regulation, our findings may have broader translational implications. Deficiency of ACY2, the canonical NAA hydrolase, causes Canavan disease, a lethal leukodystrophy characterized by NAA accumulation 9,35,36 . The intrinsic NAA hydrolase activity of SIRT2 raises the possibility of compensatory or therapeutic strategies to reduce NAA toxicity, though this remains to be tested. Besides the conventionally known roles of N -acetylated amino acids as terminal metabolites, the physiological and pathological functions of these metabolites remain largely unexplored 5,37 . While the idea that NAA-derived acetate feeds acetyl-CoA pools is conceptually intriguing 38 , ACY2 or SIRT2 hydrolysis produces acetate or O -acetyl-ADP-ribose rather than acetyl-CoA, underscoring the need to reconsider the metabolic efficiency of this route. Interestingly, a recent study proposed that NAA may act as a structural analog of NAG to activate carbamoyl-phosphate synthetase 2, aspartate transcarbamylase, and dihydroorotase (CAD) 39 , conceptually supporting the idea that structural similarity guides NAA target identification. NAA has also been implicated in protein acetylation, though the observed effects are variable 40,41 . Thus, although NAA contains an acetyl moiety, focusing solely on the acetyl group risks overlooking the biological specificity conferred by its amino acid backbone. The biological specificity of individual N -acetylated amino acids is likely conferred by the chemical properties of their underlying amino acid scaffold. Acetylation alters reactivity, p K a , and polarity, with potential consequences for transport, compartmentalization, and enzymatic recognition. Importantly, the incorporation of an acetyl group via acetyl-CoA confers nutrient-related signalling properties to N -acetylated amino acids 42,43 , positioning them as integrators of amino acid and energy metabolism. In line with this view, both NAG and NAA function as metabolic signalling modulators, activating CPS1 or inhibiting MDH1, respectively, coupling their abundance to broader metabolic network activity. Together, these insights suggest that N -acetylated amino acids constitute an underappreciated class of metabolic signals with roles extending far beyond their traditional designation as terminal metabolites. In a broader context, small-molecule acyl-amino acids are emerging as metabolic regulators. N -acetyltaurine modulates body weight control and energy balance via GDF15-GFRAL pathway 44 , while N -lactoyl-phenylalanine (Lac-Phe) acts as a blood-borne metabolic that suppress feeding and mitigates obesity 45,46 . Additionally, ketone body-amino acid conjugates were recently shown to suppress feeding and contribute to the maintenance of energy homeostasis 47 . Together with our findings, these studies highlight acyl-amino acids as an emerging class of metabolic signals that merits further investigation in energy homeostasis and disease pathogenesis. This study has several limitations. First, although both NAG and NAA were associated with CKD-associated cardiomyopathy, we primarily focused on NAA for mechanistic investigations. NAG may also exert cardiac effects apart from its known role in CPS1 activation. Second, SIRT3 showed NAA-hydrolysis activity in vitro, raising the possibility of mitochondrial-specific regulation. Third, we did not assess whether the SIRT2–NAA–MDH1 axis operates in other tissues or contributes to systemic metabolic regulation, and this warrants further investigation. In summary, our study reveals that NAA is a previously unrecognized metabolic risk factor linking renal dysfunction to cardiac remodelling. NAA is a bioactive acetylated metabolite that modulates MDH1 activity and undergoes SIRT2-mediated hydrolysis, which suggests that NAA is embedded in pathways tightly coupled to cellular energy homeostasis. Moreover, the discovery of SIRT2 as an NAA hydrolase expands the enzymatic landscape of sirtuins and uncovers a direct role for them in the regulating of small-molecule metabolites, thereby integrating the control of protein function and signalling metabolites to enable rapid, systemic metabolic adaptation and the maintenance of homeostasis. Overall, N -acetylated amino acids emerge as underappreciated metabolic signals with systemic relevance, warranting further study to inform strategies for cardiorenal and metabolic disease management. Methods Materials Chemicals and reagents were obtained from the following sources: N -acetylaspartate (Cat. No.: 00920, Sigma-Aldrich), NADH (Cat. No.: HY-F0001R, MedChemExpress), NAD + (Cat. No.: HY-B0445, MedChemExpress), NADPH (Cat. No.: HY-113324, MedChemExpress), oxaloacetate (Cat. No.: O4126, Sigma-Aldrich), glutamate dehydrogenase (Cat. No.: G2626, Sigma-Aldrich), Pronase (Cat. No.: 10165921001, Sigma-Aldrich), type II collagenase (Cat. No.: C6885, Sigma-Aldrich), photoaffinity-tagged NAA probe (Cat. No.: 1132432, WuXi AppTec), tris(2-carboxyethyl)phosphine hydrochloride (TCEP; Cat. No.: C4706, Sigma-Aldrich), tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA; Cat. No.: 678937, Sigma-Aldrich), CuSO4 (Cat. No.: C1297, Sigma-Aldrich), Biotin-azide (Cat. No.: HY-129832, MedChemExpress), Ni-NTA beads 6FF (Cat. No.: N30210, LABLEAD), [U- 13 C 6 ]-D-glucose (Cat. No.: HY-B0389A, MedChemExpress), [U- 13 C 5 ]-L-glutamine (Cat. No.: HY-N0390S1, MedChemExpress), cyto-SoNar (Cat. No.: ADSON1001-S, Provoson), mito-SoNar (Cat. No.: ADSON1003-S, Provoson), anti-MDH1 (Cat. No.: 15904-1-AP, Proteintech), anti-β-MHC (Cat. No.: 22280-1-AP, Proteintech), anti-ANP (Cat. No.: A14755, ABclonal), anti-BNP (Cat. No.: A23996, ABclonal), anti-SIRT2 (Cat. No.: A12575, ABclonal), anti-ACY1 (Cat. No.: A6351, ABclonal), anti-ACY2 (Cat. No.: A7271, ABclonal), anti-NAT8L (Cat. No.: 23841-1-AP, Proteintech), anti-SLC13A3 (Cat. No.: 26184-1-AP, Proteintech), anti-GAPDH (Cat. No.: A19056, ABclonal), anti-α-Tubulin (Cat. No.: A6830, ABclonal), goat anti-mouse IgG antibody (Cat. No.: A21001S, Abmart), and goat anti-rabbit IgG antibody (Cat. No.: M21002L, Abmart). Study participants and blood collection Adult patients with CKD were recruited at Shanghai Jiao Tong University School of Medicine, Xinhua Hospital, Department of Nephrology and Cardiology. The study design was approved and supervised by the Ethics Committee of Xinhua Hospital (Approval No.: XHEC-C-2024-185-2) in accordance with the Declaration of Helsinki. All participants were fully informed and signed the informed consent prior to enrolment. CKD was defined as an estimated glomerular filtration rate (eGFR) < 60 mL/min/1.73 m 2 and peritoneal dialysis persisting for at least three months, and further supported by elevated BUN and serum creatinine (Scr) levels 48 . Participants were excluded if they had any of the following conditions 49 : hemodialysis, history of heart failure, hypertrophic or restrictive cardiomyopathy, infiltrative cardiomyopathy (including cardiac amyloidosis), congenital heart disease, constrictive pericarditis, moderate or greater valvular disease, isolated pulmonary arterial hypertension, heart transplantation, diabetes mellitus, lupus nephritis, anti-neutrophil cytoplasmic antibody-associated vasculitis, or any known malignancy. Venous blood (3 mL) was collected from each patient into vacuum blood collection tubes containing a clot activator and inert separation gel. After standing at room temperature for 15 min to allow clot formation, samples were centrifuged at 1,500 × g for 10 min. The resulting serum was carefully transferred into a new 1.5 mL microcentrifuge tube and stored in -80 °C refrigerators. Quantification of serum metabolites Participants’ serum metabolites were analysed at Applied Protein Technology (APTBIO, Shanghai, China) using ultra-performance liquid chromatography coupled to tandem mass spectrometry (UPLC-MS/MS). Serum samples were thawed on ice, and 400 μL of cold methanol/acetonitrile (1:1, v/v) extraction solvent containing stable-isotope internal standards was added to precipitate proteins and extract metabolites. The mixture was vortexed, transferred to a new centrifuge tube, and centrifuged at 14,000 × g for 20 min at 4 °C. The supernatant was collected and dried under vacuum. For UPLC-MS/MS analysis, the dried extracts were reconstituted in 100 μL of acetonitrile/water (1:1; v/v), mixed, and centrifuged at 14,000 × g for 15 min at 4 °C. The resulting supernatant was injected for analysis. Analyses were performed using a UHPLC system (1290 Infinity LC, Agilent Technologies) coupled to a QTRAP mass spectrometer (6500+; AB Sciex). Metabolites were separated on both HILIC and reversed-phase C18 columns (Waters UPLC BEH Amide and Waters UPLC BEH C18, respectively; 2.1 × 100 mm, 1.7 μm). Data acquisition was conducted in both positive and negative ion modes using multiple reaction monitoring (MRM), and quantification was performed using MultiQuant or Analyst software. Quality control (QC) samples were interspersed throughout the run to assess system stability, and metabolites with a coefficient of variation (CV) < 30% in QC samples were retained for further analysis. For metabolite determination, a comprehensive set of rigorous quality control/assurance procedures was employed to ensure consistently high-quality analytical results by monitoring every step from sample receipt at the laboratory to final deliverables. Experimental animals and animal care All animal experiments were approved and conducted in accordance with the guidelines of the Animal Care Committee at Xinhua Hospital, Shanghai Jiao Tong University School of Medicine (Approval No.: XHEC-NSFC-2023-100). The CKD mouse models were established using the 5/6 nephrectomy surgery 50,51 . Male C57BL/6 mice, aged six weeks, were purchased from GemPharmatech Co., Ltd. (Nanjing, China). All mice were housed in a specific pathogen-free (SPF) facility under controlled temperature (20–22 °C), humidity (40–60%), and a 12 h light/dark cycle, with free access to food and water. Prior to surgery, mice were acclimatized to the new environment for two weeks. At eight weeks of age, CKD was induced using a two-step 5/6 nephrectomy. Briefly, mice were anesthetized with 2% isoflurane and placed on a warming pad to maintain body temperature. In the first surgery, approximately two-thirds of the left kidney was removed by resecting the upper and lower poles while carefully controlling hemostasis. After a seven-day recovery, the second surgery was performed to completely remove the right kidney. Sham-operated mice underwent identical surgical procedures without renal tissue removal. Mice were monitored daily for wound healing and general health. For pharmacological intervention, NAA (00920, Sigma-Aldrich) was dissolved in phosphate buffer (pH 7.2–7.4) and administered intraperitoneally at 200 mg/kg daily for eight weeks. Control mice received equal volumes of phosphate buffer. Four-week-old C57BL/6 Sirtuin2 -/- mice were purchased from GemPharmatech Co., Ltd. (Nanjing, China). Genotypes were determined via PCR using genomic DNA obtained from tails; two primer pairs were used ( Sirtuin2 -/- forward 1: 5′-CCTTAAACAGAACACCCCAGGG-3’, reverse 1: 5′-GAGATAGGTAGTTTTGGCTGACCTTG-3′; Sirtuin2 -/- forward 2: 5′-TAAGTTCCCCGTCCTTCAGCCA-3′, reverse 2: 5′-CCCAGAAGCTAGGTTCTTAATCATGT-3′). All mice were housed in a specific SPF facility under controlled temperature (20–22 °C), humidity (40–60%), and a 12 h light/dark cycle, with free access to food and water. AAV9-mediated cardiomyocyte-specific Mdh1 -knockdown To achieve Mdh1 cardiomyocyte-specific knockdown, mice received a single tail-vein injection of AAV9-cTnT-sh Mdh1 (GIEV5032496, GeneChem) at 1 × 10 12 vg per mouse. AAV9 carrying shRNA to interfere with the mouse Mdh1 gene were constructed and inserted into GV683, which contained the expression cassette cTnTp-EGFP-MIR155(MCS)-WPRE-SV40 PolyA. Knockdown efficiency was assessed by fluorescence, and the validated target sequence was: GACCCAGTATCCAGATGTCAA. AAV9-mediated cardiomyocyte-specific SIRT2 overexpression For cardiomyocyte-specific overexpression of SIRT2, mice received a single tail-vein injection of AAV9-cTnT-SIRT2 (GOSV5019825, GeneChem) at 1 × 10 12 vg per mouse. AAV9 carrying the mouse Sirtuin2 -coading gene was constructed and inserted into GV831, which contained the expression cassette cTnTp-MCS-3Flag-FT2A-EGFP-WPRE-BGH polyA; the cassette was generated using the forward primer (5′-AAGGCTAGAGTACTGCTAGCCGCCACCATGGCCGAGCCGGACCCCTCTGACCCTCTGGAG-3′) and reverse primer (5′-TAGTCCATGGTGGCACCGGTCTGCTGTTCCTCTTTCTCTTTGG-3′). Successful transduction and SIRT2 overexpression were validated by fluorescence in myocardial tissue. Echocardiography Transthoracic echocardiography was performed to assess cardiac function using a high-frequency ultrasound system (Vevo 3100, VisualSonics) equipped with an MX550 transducer. Two-dimensional (B-mode) and M-mode images were obtained from the parasternal long-axis and short-axis views at the level of the papillary muscles. Left ventricular diameters (end-diastolic, LVEDD; end-systolic, LVESD), volumes (end-diastolic, LVEDV; end-systolic, LVESV), and posterior wall thickness at end-diastole (LVPWd) were measured from M-mode tracings. Fractional shortening (LVFS) and ejection fraction (LVEF) were determined, with three consecutive cardiac cycles analysed per mouse. Detection of blood urea nitrogen (BUN) levels in mice Mouse BUN levels were measured using a Mouse Urea Nitrogen ELISA Kit (FT-P9S949X, Shanghai Fantai Biotechnology) according to the manufacturer’s protocol. In brief, 10 μL of serum was mixed with 40 μL diluent in precoated wells, followed by 100 μL HRP-conjugated reagent. Plates were incubated at 37 °C for 1 h, washed thoroughly, and incubated with chromogenic substrate for color development. After 15 min of dark incubation at 37 °C, the reaction was terminated, and absorbance at 450 nm was recorded using a Multimode Microplate Reader (Spark, Tecan). Data were background-corrected, and BUN concentrations were calculated from standard curves. Histology and staining Mice tissues were harvested, fixed overnight in 4% paraformaldehyde, dehydrated, and embedded in paraffin. Five-micrometer sections were cut and stained with hematoxylin and eosin (H&E) for morphological evaluation, wheat germ agglutinin (WGA) to visualize cardiomyocyte borders, and Masson’s trichrome to detect collagen deposition. For immunofluorescence, sections were incubated with primary antibodies at 4 °C overnight, followed by appropriate fluorophore-conjugated secondary antibodies. Nuclei were counterstained with DAPI, and images were acquired using a fluorescence microscope (Leica Microsystems). Quantification was performed using ImageJ software (NIH). Isolation and culture of primary neonatal rat cardiomyocytes (NRCMs) NRCMs were isolated from 1-day-old Sprague-Dawley rats as previously described 52,53 . Hearts were excised under sterile conditions, atria removed, and ventricles minced into small fragments in ice-cold CBFHH buffer (NaCl 137 mM, KCl 5.4 mM, KH 2 PO 4 0.44 mM, HEPES 10 mM, Na 2 HPO 4 0.17 mM, D-glucose 3 mM, MgSO 4 ·7H 2 O 0.4 mM, pH 7.4). Tissue was digested at 37 °C with type II collagenase (0.45 mg·ml⁻¹, 2 × 10 min; C6885, Sigma-Aldrich) and subsequent trypsin digestion (0.125%, five min per cycle; G4022, Servicebio) until completely dispersed. Supernatants from all digestions were pooled, neutralized using low-glucose DMEM (L170KJ, BasalMedia) supplemented with 20% FBS (A5256701, Gibco), filtered through a 70 µm cell strainer (CLS431751, Sigma-Aldrich), and centrifuged at 800 rpm for five min. Cell pellets were resuspended in low-glucose DMEM with 20% FBS and pre-plated for one hour to minimize fibroblast contamination via differential adhesion. Non-adherent NRCMs were collected and seeded onto laminin-coated (10 µg/mL; CC095, Sigma-Aldrich) plates. Cells were maintained in low-glucose DMEM supplemented with 20% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin (Invitrogen) at 37 °C in a humidified atmosphere of 5% CO 2 /95% O 2 . Culture medium was replaced 24 h post-plating before experimental use. NRCMs were treated with NAA (final concentration: 0.5 mM) prepared in phosphate buffer (pH 7.2–7.4). Control cells received an equal volume of phosphate buffer. AAV-mediated SIRT2 overexpression in NRCMs Recombinant AAV9 harboring the rat Sirtuin2 -coding gene was constructed in the GV345 vector, which contains the CMV-MCS-3FLAG-SV40-Cherry expression cassette. The construct was cloned using the following primers: forward 5′-AGGTCGACTCTAGAGGATCCCGCCACCATGGACTTCCTACGGAATTTATTC-3′ and reverse 5′-TCCTTGTAGTCCATACCGGTGTGTTCCTCTTTCTCTTTGGTCC-3′, and packaged by GeneChem (GOSA5022468) at a titer of 2.5 × 10 10 PFU/mL. NRCMs were infected at a multiplicity of infection (MOI) of 50 in serum-free medium for 2 h. Complete medium was added after 2 h, and the medium was replaced 12 h later. Cells were cultured for 36–48 h before downstream assays and analyses. Metabolite quantification Metabolites quantification was performed by Metabo-Profile Biotechnology (Shanghai, China) using UPLC-MS/MS 54 . Samples were thawed on ice to minimize degradation before extraction. For mice serum, 50 μL of sample was mixed with 250 μL of precipitation reagent containing internal standards, followed by vortexing at 1,200 rpm for 20 min at 10 °C. After centrifugation (18,000 × g , 20 min, 4 °C), 100 μL of the supernatant was transferred into a 96-well plate for UPLC-MS/MS analysis. For mice heart and kidney tissues, ~10 mg of each sample was homogenized in 50 μL of deionized water with 10 zirconium oxide beads for 3 min, followed by the addition of 250 μL precipitation reagent (with internal standards). Samples were further homogenized (speed 8, 3 min) and centrifuged at 18,000 × g for 20 min at 4 °C. Then, 100 μL of supernatant was transferred into a 96-well plate for UPLC-MS/MS analysis. For cells, 120 μL of methanol was added to each tube, and metabolites were extracted by ultrasonic disruption. After centrifugation at 18,000 × g for 20 min at 4 °C, 30 μL of the supernatant was transferred into a 96-well plate for automated derivatization on a Biomek 4000 workstation (Beckman Coulter, Brea, USA). Briefly, 20 μL of freshly prepared derivatization reagent was added to each well, and the plate was shaken at 1,450 rpm for 60 min at 30 °C. Following derivatization, 330 μL of 50% methanol was added, and the plate was mixed at 650 rpm for 5 min at 30 °C, then centrifuged at 4,000 × g for 30 min at 4 °C. The supernatants were sealed in 96-well plates for UPLC-MS/MS analysis. Chromatographic separation was achieved on a BEH Amide column (Waters) coupled to a triple quadrupole tandem mass spectrometer (ACQUITY UPLC–Xevo TQ-S, Waters Corp., Milford, MA, USA) operating in both positive and negative multiple reaction monitoring modes. Authentic and isotopically labeled standards were used for calibration, and pooled QC samples were injected periodically to monitor analytical stability. Data were processed with vendor software, and metabolite concentrations were determined from standard curves. Rigorous quality control/assurance procedures were implemented throughout to ensure high-quality analytical results. Identification of NAA-binding proteins NAA-binding proteins were identified as previously described 55,56 . Heart tissues were harvested from adult mice and immediately washed with ice-cold phosphate buffer. Apex tissues (50 mg) were minced and lysed in 500 μL ice-cold 0.5% NP-40 buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% NP-40, 1 μg/mL aprotinin, 1 μg/mL leupeptin, 1 μg/mL pepstatin and 1 mM PMSF) using a tissue grinder. Lysates were clarified by centrifugation at 15,000 × g for 20 min at 4 °C. The supernatants were incubated with 100 μM synthesized NAA probes (1132432, AppTec) or NAA at 4 °C for 2 h, followed by exposure to 365 nm UV light (UCL-3200L, LUYOR) for 10 min to crosslink NAA-interacting proteins. Subsequently, the reaction mixtures were adjusted to final concentrations of 1 mM TCEP (C4706, Sigma-Aldrich), 0.1 mM TBTA (678937, Sigma-Aldrich), 1 mM CuSO4 (C1297, Sigma-Aldrich), and 1 mM Biotin-azide (HY-129832, MedChemExpress), and incubated at 4 °C for 1 h for click chemistry. Protein aggregates were removed by centrifugation at 20,000 × g for 15 min, and the supernatants were incubated with streptavidin magnetic beads (HY-K0208, MedChemExpress) for 2 h with gentle rotation at 4 °C. Beads were washed three times with 700 µL 0.5% NP-40 buffer and then resuspended in 100 µL SDS-PAGE loading buffer. Samples were resolved by SDS-PAGE, stained with Coomassie Blue Fast Staining Solution (P0017, Beyotime), and protein bands were excised for mass spectrometry analysis. Protein expression and purification The coding sequences of SIRT1–7, MDH1, MDH2, and nicotinamidase were subcloned into a modified pGEX-6p-1 vector carrying an N-terminal 6 × His tag. Plasmids were transformed into E. coli BL21 (DE3), and protein expression was induced with 0.5 mM IPTG when the OD 600 reached 0.6–0.8, followed by incubation at 4 °C overnight. Bacterial pellets were lysed by sonication in lysis buffer (50 mM Tris-HCl, pH 7.5, 300 mM NaCl, 10 mM imidazole, 1 mM DTT, protease inhibitors), and lysates were clarified by centrifugation at 15,000 × g for 30 min at 4 °C. Recombinant proteins were purified using Ni-NTA Beads 6FF (N30210, LABLEAD), eluted with 250 mM imidazole, and purity was verified by SDS-PAGE. Purified proteins were concentrated, flash-frozen in liquid nitrogen, and stored at -80 °C until use. The following primers were used: hSIRT1: forward 5′-CCCCTGGGATCCCCGGAATTCATGGCGGACGAGGCGGCC-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGTGATTTGTTTGATGGATAGTTCATGT-3′. hSIRT2: forward 5′-CCCCTGGGATCCCCGGAATTCATGGCAGAGCCAGACCCC-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGCTGGGGTTTCTCCCTCTCTGTT-3′. hSIRT3: forward 5′-CCCCTGGGATCCCCGGAATTCATGGCGTTCTGGGGTTGG-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGTTTGTCTGGTCCATCAAGCTTCC-3′. hSIRT4: forward 5′-CCCCTGGGATCCCCGGAATTCATGAAGATGAGCTTTGCGTTGA-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGGCATGGGTCTATCAAAGGCAGC-3′. hSIRT5: forward 5′-CCCCTGGGATCCCCGGAATTCATGCGACCTCTCCAGATTGTCC-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGAGAAACAGTTTCATTTTCATGACAGG-3′. hSIRT6: forward 5′-CCCCTGGGATCCCCGGAATTCATGTCGGTGAATTACGCGGC-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGGCTGGGGACCGCCTTGGC-3′. hSIRT7: forward 5′-CCCCTGGGATCCCCGGAATTCATGGCAGCCGGGGGTCTG-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGCGTCACTTTCTTCCTTTTTGTGC-3′. mMDH1: forward 5′-CCCCTGGGATCCCCGGAATTCATGTCTGAACCAATCAGAGTCCTTG-3′, reverse 5′-GTCACGATGCGGCCGCTCGAGCGCAGAGGAGAGAAACTCAAAAG-3′. mMDH2: forward 5′-TAGTCCAGTGTGGTGGAATTCATGCTGTCCGCTCTCGCC-3′, reverse 5′-AACGGGCCCTCTAGACTCGAGCTTCATGTTCTTGACAAAGTCCTCG-3′. Site-directed mutagenesis of mMDH1 was performed to generate alanine substitution mutants (R92A/R98A/N131A/R162A/S242A) using the following primer pairs: R92A: forward 5′-GTCCTAGTGGGCTCCATGCCAGCTAGGGAAGGCATGGAGAGGAAG-3′, reverse 5′-CTTCCTCTCCATGCCTTCCCTAGCTGGCATGGAGCCCACTAGGAC-3′. R98A: forward 5′-GAAGGGAAGGCATGGAGGCAAAGGACCTACTGAAAG-3′, reverse 5′-CTTTCAGTAGGTCCTTTGCCTCCATGCCTTCCCTTC-3′. N131A: forward 5′-GTCATTGTTGTGGGAGCCCCAGCCAATACGAAC-3′, reverse 5′-GTTCGTATTGGCTGGGGCTCCCACAACAATGAC-3′. R162A: forward 5′-CTCGCTTGGACCACAACGCAGCAAAATCTCAAATTG-3′, reverse 5′-CAATTTGAGATTTTGCTGCGTTGTGGTCCAAGCGAG-3′. S242A: forward 5′-CTCGGAAGCTGTCCGCTGCAATGTCTGCTG-3′, reverse 5′-CAGCAGACATTGCAGCGGACAGCTTCCGAG-3′. Nicotinamidase: forward 5′- CCCCTGGGATCCCCGGAATTCATGCCCCCTCGCGCCCTG-3′, reverse 5′- GTCACGATGCGGCCGCTCGAGCCCCTGTGTCTCTTCCCAGTC-3′. MDH1 enzymatic activity assay MDH1 enzymatic activity was determined by monitoring NADH consumption at 340 nm. For recombinant assays, purified MDH1 (0.1 mg/mL) was incubated in 150 µL of reaction buffer: 50 mM Tris-HCl pH 7.4, 6 mM MgCl 2 , 1 mM DTT, 0.1 mM NADH (HY-F0001R, MedChemExpress), and initiated with freshly prepared 100 µM oxaloacetate (O4126, Sigma-Aldrich). The reactions were continuously analysed in a Multimode Microplate Reader (Spark, Tecan) at 25 °C, and the NADH consumption was measured every minute for 15 min. For tissue assays, MDH1 was immunoprecipitated with an anti-MDH1 antibody from freshly isolated mouse heart apex and assayed for enzymatic activity under the same reaction conditions. Enzymatic activity was calculated from the rate of NADH consumption, using the molar extinction coefficient of NADH (ε = 6.22 mM⁻¹·cm⁻¹) to convert absorbance changes at 340 nm into reaction rates. SIRT2 enzymatic activity assay SIRT2-mediated NAA hydrolysis was evaluated using complementary enzyme-coupled and LC-MS/MS-based assays. Enzyme-coupled assays were conducted as previously described 27,57 . The following reagents were used in the assay mixtures: 1 mM NAA (00920, Sigma-Aldrich), 1 mM NAD + (HY-B0445, MedChemExpress), 1 mM DTT, 6 mM MgCl 2 , 0.1 mg/mL nicotinamidase, 2.5 mM α-ketoglutarate, 0.2 mM NADPH (HY-113324, MedChemExpress) and 0.175 units of glutamate dehydrogenase (G2626, Sigma-Aldrich), 20 mM phosphate buffer pH 7.4. All assay components except SIRT2 were preincubated at 25 °C for 5 min, and 2.5 mg/mL purified SIRT2 was added to start the reactions. The final volume was 100 µL. NADPH was quantified by its intrinsic fluorescence with excitation at approximately 340 nm and emission at 460 nm in a solid black, flat-bottomed, 96-well plate. The reactions were continuously analysed in a Multimode Microplate Reader (Spark, Tecan) at 25 °C, and the NADPH consumption was measured every 5 min for 2 h. The raw data from three replicates were fitted to Akima splines using GraphPad Prism (v. 10.0, Dotmatics) and kinetic parameters ( K m and V max ) by varying NAA concentrations from 0.1 to 5 mM. For LC-MS/MS analysis, reactions containing 1 mM NAA, 1 mM NAD + , 1 mM DTT, 6 mM MgCl 2 and SIRT2 (2.5 mg/mL) in 50 mM HEPES (pH 7.4) were incubated at 37 °C for 2 h. Reactions were quenched with ice-cold methanol (3 × reaction volume) and centrifuged at 15,000 × g for 10 min at 4 °C. Supernatants were analysed on a Waters H-Class UPLC equipped with a Welch Ultimate AQ-C18 column (2.1 × 250 mm, 5 μm) using a 0.2 mL/min flow rate and a water–acetonitrile gradient (0.1% formic acid; 0-1 min, 0% B; 1-8 min, 0→90% B; 8-10 min, 90% B; 10-10.1 min, 90→0% B; 10.1-15 min, 0% B). Detection was performed on a Sciex API 6500 QTRAP in positive electrospray ionization mode using multiple reaction monitoring (MRM), with NAA and L-aspartate unambiguously distinguished based on their specific parent-to-fragment ion transitions and chromatographic retention times. MRM transitions were optimized with authentic standards. Peak areas were extracted in Skyline for relative quantification and plotted using GraphPad Prism (v. 10.0, Dotmatics). Drug Affinity Responsive Target Stability (DARTS) Purified protein was diluted to 1 mg/mL with TNM buffer (50 mM Tris-HCl, 50 mM NaCl, 6 mM MgCl 2 , pH 7.4) and incubated with 1 mM NAA or 1 mM L-aspartate for 1 h at 25 °C. Pronase (10165921001, Sigma-Aldrich) was added at varying mass ratios relative to the protein, and reactions were incubated for 30 min at 25 °C. Reactions were terminated by addition of SDS-PAGE loading buffer. Samples were resolved by SDS-PAGE, stained with Coomassie Blue Fast Staining Solution (P0017, Beyotime), and imaged using a Typhoon FLA 9500 scanner (GE Healthcare). Surface-Plasmon Resonance (SPR) Binding kinetics and affinity of NAA to target proteins were measured on a Biacore T200 at 25 °C using CM5 sensor chips, with data analyzed via Biacore T200 Evaluation software (v. 2.0GE Healthcare). Purified proteins were covalently immobilized on the CM5 sensor chip via amine groups in 10 mM sodium acetate buffer (pH 5.0). Carboxyl groups on the sensor surface were activated by injection of 0.2 M N-ethyl-N′-(3-dimethylaminopropyl) carbodiimide (EDC) and 0.05 M N-hydroxysuccinimide (NHS) at 10 µL/min. Proteins in 10 mM sodium acetate buffer (pH 5.0) were injected at 10 µL/min to couple to the sensor surface, and remaining active sites were blocked with 1M ethanolamine (EA). A reference flow cell was activated and blocked in the absence of protein. For direct binding experiments, immobilization levels were adjusted to 10,000 response units (RU), and NAA solutions containing 5% DMSO were serially injected from low to high concentrations at 30 µL/min for 150 s. Association and dissociation rates were determined by fitting the data to a 1:1 Langmuir binding model using Biacore T200 Evaluation software. Molecular docking MDH1 or SIRT2 structures were obtained from Protein Data Bank (PDB IDs: 7RM9 and 4RMG, respectively) and prepared in MOE at pH 7.4 (AMBER10: EHT). Binding pockets were identified via structural analysis or MOE Site Finder. Molecular docking was performed with the triangle matcher algorithm, initial poses were scored using London ΔG, and the top 10-20 poses per pocket were refined via the induced fit algorithm with GBVI/WSA ΔG scoring, and the pose with the best docking score was used for visualization in the manuscript. SIRT2-TUBA1A complexes were predicted using AlphaFold-Multimer, and resulting models were inspected to identify plausible interaction interfaces. Molecular dynamics simulations Molecular dynamics simulations of protein–ligand complexes were conducted with Desmond 2020. Docking-derived best-scoring poses were placed in a TIP3P water box and neutralized with Na + /Cl - ions. Systems were parameterized with the OPLS3e force field. Energy minimization was carried out with SHAKE constraints on bond lengths and angles of heavy atoms and water geometry. Periodic boundary conditions and the particle mesh Ewald method were applied. Production MD simulations were run for 100 ns in the NPT ensemble at 300 K and 1 atm with a 2 fs integration step, and snapshots were recorded every 10 ps. Post-MD analyses included assessment of structural stability (backbone RMSD) and characterization of protein-ligand interactions, including interacting residues, interaction types, interaction frequencies, and estimated binding free energies. Metabolic flux analysis NRCMs were seeded in 6-well plates and cultured to ~80% confluency. Cells were washed twice with sterile PBS and incubated with isotope-labeled tracing medium. For [U- 13 C 6 ]-D-glucose tracing, cells were cultured in glucose-free DMEM (D5030, Sigma-Aldrich; with 0.584 g/L glutamine) supplemented with 20% dialyzed FBS (04-011-1A, Biological Industries) and 10 mM [U- 13 C 6 ]-D-glucose (HY-B0389A, MedChemExpress) for 6 h to assess glycolytic and TCA cycle fluxes 58,59 . For [U- 13 C 5 ]-L-glutamine tracing, cells were cultured in glutamine-free DMEM (D5030, Sigma-Aldrich; with 1.0 g/L D-Glucose) supplemented with 20% dialyzed FBS (04-011-1A, Biological Industries) and 2 mM [U- 13 C 5 ]-L-glutamine (HY-N0390S1, MedChemExpress) for 24 h to assess TCA cycle fluxes 60 . Metabolites were subjected to UPLC-MS/MS-based metabolic flux analysis (Metabo-Profile Biotechnology, China) using an ultrahigh-pressure liquid chromatography-triple quadrupole mass spectrometer (ACQUITY UPLC-Xevo TQ-S, Waters Corp., Milford, MA, USA) 61 . For data processing, the raw data files generated by UPLC-MS/MS were processed using MassLynx software (v 4.1, Waters Corp., Milford, MA, USA）for peak extraction, integration, identification, and quantification of individual metabolites. NAD + /NADH ratio determination Cellular NAD + /NADH ratios were determined using two complementary approaches. For live-cell measurements, cells were seeded evenly in black, opaque 96-well plates (~2 × 10 4 cells per well) and transduced with adenovirus expressing cyto-Sonar (ADSON1001-S) or mito-Sonar (ADSON1003-S, Provoson) following the manufacturer’s instructions 62,63 . Fluorescence was measured using a Multimode Microplate Reader (Spark, Tecan) at 420/528 nm (NADH) and 485/528 nm (NAD + ). The NAD + /NADH ratio was determined ratiometrically as R 485/420 , by dividing the fluorescence intensity at 485 nm by that at 420 nm. For biochemical measurements, cells were lysed and processed according to the NAD + /NADH Assay Kit protocol (S0176S, Beyotime). Lysates were subjected to enzymatic cycling reactions, and absorbance at 450 nm was measured using a microplate reader. NAD + /NADH ratios were calculated and normalized to total protein. ATP quantification Cellular ATP levels were measured using the ATP Assay Kit (S002, Beyotime) according to the manufacturer’s instructions. Briefly, cells were washed with PBS and lysed using the provided lysis buffer. After centrifugation at 12,000 × g for 5 min at 4 °C, the supernatant was collected. An aliquot of the lysate was mixed with the reaction solution and incubated for 5 min at room temperature in the dark. The luminescence was measured using a Multimode Microplate Reader (Spark, Tecan). ATP concentrations were calculated based on a standard curve generated with ATP standard solutions provided in the kit, and results were normalized to protein concentration determined by BCA assay (P0009, Beyotime). Western blotting For cell samples, proteins were extracted directly using 1× SDS loading buffer. Tissue samples were homogenized in 0.5% NP-40 lysis buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% NP-40, 1 μg/mL aprotinin, 1 μg/mL leupeptin, 1 μg/mL pepstatin and 1 mM PMSF). The lysates were centrifuged at 4 °C for 20 min at 15,000 × g , and the supernatants were used as whole-cell extracts. Protein samples were subjected to western blotting according to standard procedures. Detection was performed by measuring chemiluminescence using an ECL Plus Western Blotting Detection System on a Typhoon FLA 9500 (both GE Healthcare). The following antibodies were used for western blot analysis: anti-β-MHC (1:1000, 22280-1-AP, Proteintech), anti-ANP (1:1000, A14755, ABclonal), anti-BNP (1:1000, A23996, ABclonal), anti-SIRT2 (1:1000, A12575, ABclonal), anti-ACY1 (1:1000, A6351, ABclonal), anti-ACY2 (1:1000, A7271, ABclonal), anti-NAT8L (1:1000, 23841-1-AP, Proteintech), anti-SLC13A3 (1:1000, 26184-1-AP, Proteintech), anti-GAPDH (1:3000, A19056, ABclonal), anti-α-Tubulin (1:3000, A6830, ABclonal), goat anti-mouse IgG antibody (1:5000, A21001S, Abmart), goat anti-rabbit IgG antibody (1:5000, M21002L, Abmart). Phalloidin staining NRCMs were cultured on glass coverslips in 12-well plates and fixed with 4% paraformaldehyde for 10 min at room temperature. After washing three times with PBS, the cells were permeabilized with 0.1% Triton X-100 in PBS for 5 min. Following PBS washes, the samples were incubated with Phalloidin-iFluor™ 488 (BL1190A, Biosharp) diluted 1:1000 in PBS for 30 min at room temperature in the dark. Nuclei were counterstained with DAPI (D9542, Sigma-Aldrich). Coverslips were mounted with antifade mounting medium (G1226-7, ServiceBio) and imaged using a Leica fluorescence microscope. The size of individual cells was quantified by ImageJ software (NIH). Statistical analysis All statistical analyses were performed using GraphPad Prism (v. 10.0, Dotmatics). For all experiments, data are presented as the mean ± standard error of the mean (SEM) unless otherwise specified. All experiments (except those described otherwise in the legend) were performed independently at least three times with a similar outcome. The statistical tests, n values, and the P values are all indicated in the figures and/or legends. Depending on the type of experiment, the P values were calculated using different analytical methods. Two-group comparisons were performed using two-tailed Student’s t -tests. For single-variable comparisons among three or more groups, one-way analysis of variance (ANOVA) followed by Tukey’s multiple comparison test was applied. For experiments involving two independent variables, two-way ANOVA with Tukey’s or Bonferroni’s post hoc test was used to obtain adjusted P values. P < 0.05 was considered significant (* P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001). Declarations Acknowledgements This work was supported by the Grants from National Natural Science Foundation of China (82330048, 82501473, 32370824, 82300428, 82101750), Program of Shanghai Academic Research Leader (21XD1421700), Innovation Program of the Shanghai Municipal Education Commission (2023ZKZD24), China National Postdoctoral Program for Innovative Talents fellowship (BX20250188), Innovative research team of high-level local universities in Shanghai (SHSMU-ZDCX20211100), the Science and Technology Commission of Shanghai Municipality (25ZR1402362, 25ZR1402037), Shanghai Sailing Program (23YF1425500), and The Construction Project of the “Discipline Peak-Climbing Plan” of Xinhua Hospital Affiliated to Shanghai Jiao Tong University School of Medicine (XKPF2024A5001). Author contributions J.-Y.Z. and Z.-Y.Z. conceived the project. Z.-Y.Z. and J.-Y.Z. wrote the manuscript. J.-Y.Z. and H.T. supervised the study. Z.-Y.Z., K.C., Y.Y., and F.-J.L. performed human samples studies. 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Genetically encoded biosensors for evaluating NAD(+)/NADH ratio in cytosolic and mitochondrial compartments. Cell Rep Methods 1 (2021). https://doi.org:10.1016/j.crmeth.2021.100116 Additional Declarations There is NO Competing Interest. Supplementary Files ExtendedTable2026.pdf Supplementary Table ExtendedFigs2026.pdf Supplementary Figure Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. 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4","display":"","copyAsset":false,"role":"figure","size":673611,"visible":true,"origin":"","legend":"","description":"","filename":"4.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8510072/v1/592228a7a6d8fe6cf44e6e3c.jpg"},{"id":100134126,"identity":"9d526121-709b-41ed-a3e1-ec9b89ac497c","added_by":"auto","created_at":"2026-01-13 10:31:51","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":434785,"visible":true,"origin":"","legend":"\u003cp\u003eSee image above for figure legend.\u003c/p\u003e","description":"","filename":"Fig5.png","url":"https://assets-eu.researchsquare.com/files/rs-8510072/v1/ad1ddb9d48abcd78e27cd2a5.png"},{"id":101202391,"identity":"343d19f9-0b42-48f6-a894-fcc907609c80","added_by":"auto","created_at":"2026-01-27 09:29:51","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":4222269,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8510072/v1/900bd2f0-5cab-45ea-be2f-0c57be450150.pdf"},{"id":100134118,"identity":"6df3fda3-6195-4a0b-80dc-b6922961e509","added_by":"auto","created_at":"2026-01-13 10:31:51","extension":"pdf","order_by":1,"title":"","display":"","copyAsset":false,"role":"supplement","size":93325,"visible":true,"origin":"","legend":"Supplementary Table","description":"","filename":"ExtendedTable2026.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8510072/v1/472b58401fbed893accc6387.pdf"},{"id":100134129,"identity":"04262208-56c1-48ac-a95e-fba04e8c6b6c","added_by":"auto","created_at":"2026-01-13 10:31:52","extension":"pdf","order_by":2,"title":"","display":"","copyAsset":false,"role":"supplement","size":21416686,"visible":true,"origin":"","legend":"Supplementary Figure","description":"","filename":"ExtendedFigs2026.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8510072/v1/e7c05f97077c330a639307d3.pdf"}],"financialInterests":"There is \u003cb\u003eNO\u003c/b\u003e Competing Interest.","formattedTitle":"SIRT2 functions as an N-acetylaspartate hydrolase that counteracts cardiac metabolic remodeling in kidney disease","fulltext":[{"header":"Introduction","content":"\u003cp\u003e\u003cem\u003eN\u003c/em\u003e-acetylated amino acids have generally been regarded as terminal metabolic derivatives, and their circulating levels rise in chronic kidney disease (CKD) due to impaired renal clearance\u003csup\u003e1,3-5\u003c/sup\u003e. CKD-associated cardiomyopathy involves complex cardiac remodelling partly driven by the accumulation of circulating\u0026nbsp;uremic toxins\u003csup\u003e2\u003c/sup\u003e, and offers a relevant endogenous disease context to investigate the physiological roles of \u003cem\u003eN\u003c/em\u003e-acetylated amino acids.\u0026nbsp;Protein acetylation is one of the major post-translational modifications involved in many key cellular processes\u003csup\u003e6,7\u003c/sup\u003e; however, whether \u003cem\u003eN\u003c/em\u003e-acetylated amino acids act as functional modulators of cellular metabolism remains unknown. Among these, \u003cem\u003eN\u003c/em\u003e-acetylaspartate (NAA) is notable for being the second most abundant metabolite in the central nervous system\u003csup\u003e8,9\u003c/sup\u003e, yet its functional roles outside the brain remain poorly understood. In this context, whether elevated circulating NAA acts as a signalling metabolite and contributes to cardiac metabolic remodelling remains unknown. In mammals, the seven sirtuins (SIRT1\u0026ndash;SIRT7) constitute an evolutionarily conserved family of NAD\u003csup\u003e+\u003c/sup\u003e-dependent enzymes with diverse biological functions\u003csup\u003e10-19\u003c/sup\u003e. These enzymatic activities share a common mechanism of amide bond hydrolysis, suggesting the potential existence of additional, yet-undiscovered enzymatic functions.\u003c/p\u003e\n\u003cp\u003eThis study investigates the relationship between circulating NAA levels and cardiac metabolic homeostasis. Mechanistically, we show that NAA suppresses the activity of malate dehydrogenase 1 (MDH1), thereby impairing mitochondrial NADH transport and cardiac metabolism. Moreover, we discover that SIRT2 is a previously unrecognized NAA hydrolase that mitigates NAA-induced metabolic disturbances. Altogether, these findings reveal the detrimental impact of NAA on cardiac metabolic homeostasis and broaden the functional landscape of sirtuins.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003eNAA levels are elevated and associated with cardiac impairment in CKD\u003c/p\u003e\n\u003cp\u003eTo investigate the potential association between circulating \u003cem\u003eN\u003c/em\u003e-acetylated amino acids and cardiac impairment, we analyzed a well-characterized cohort of 62 patients with CKD. Participants were divided by plasma B-type natriuretic peptide (BNP) levels, using a threshold \u003cstrong\u003e\u0026ge;\u0026nbsp;\u003c/strong\u003e200 ng/L as a marker of early cardiac dysfunction\u003csup\u003e20\u003c/sup\u003e. Individuals above this threshold exhibited reduced left ventricular ejection fraction (LVEF), enlarged left ventricular and left atrial dimensions, and increased interventricular septal and posterior wall thickness, consistent with early cardiac remodelling and subclinical dysfunction\u0026nbsp;(\u003cstrong\u003eExtended Data\u003c/strong\u003e \u003cstrong\u003eTable\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;1\u003c/strong\u003e). Targeted metabolomics analysis of circulating metabolites, including 12 circulating \u003cem\u003eN\u003c/em\u003e-acetylated amino acids, revealed that NAA and \u003cem\u003eN\u003c/em\u003e-acetylglutamate (NAG) levels were significantly elevated in individuals with high BNP levels, whereas their unmodified precursors, aspartate and glutamate, remained unchanged (\u003cstrong\u003eFig. 1a,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 1a, b\u003c/strong\u003e). Among all \u003cem\u003eN\u003c/em\u003e-acetylated amino acids measured, NAA and NAG showed the strongest correlations with BNP levels, exceeding those of known risk factors for CKD-associated cardiomyopathy, such as p-cresyl sulfate and indoxyl sulfate\u003csup\u003e21\u003c/sup\u003e (\u003cstrong\u003eFig. 1b,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 1c, d\u003c/strong\u003e). Together, these results indicate that elevated circulating NAA and NAG levels are closely linked to early cardiac dysfunction in CKD.\u003c/p\u003e\n\u003cp\u003eTo investigate whether dysregulation of \u003cem\u003eN\u003c/em\u003e-acetylated amino acids contributes to CKD-associated cardiomyopathy, we employed the 5/6 subtotal nephrectomy model. Eight weeks post-surgery, mice exhibited elevated blood urea nitrogen (BUN) levels, confirming renal impairment, accompanied by systemic accumulation of \u003cem\u003eN\u003c/em\u003e-acetylated amino acids and cardiac dysfunction, including systolic impairment and myocardial hypertrophy. (\u003cstrong\u003eExtended Data Fig. 2a-e\u003c/strong\u003e).\u0026nbsp;Only NAA and NAG levels increased significantly in the heart, with NAA displaying the most pronounced elevation (\u003cstrong\u003eFig. 1c,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 2f\u003c/strong\u003e). This pattern mirrors our observations in the human cohort (\u003cstrong\u003eFig. 1a, b)\u003c/strong\u003e, supporting a potential link between these metabolites and early cardiac metabolic remodelling. NAG is a well-characterized allosteric activator of carbamoyl phosphate synthetase 1 (CPS1) in the urea cycle\u003csup\u003e22\u003c/sup\u003e. However, the biological functions of NAA remain largely unexplored. Considering the marked elevation of NAA levels in mice and humans, we focused the subsequent mechanistic studies on NAA.\u003c/p\u003e\n\u003cp\u003eTissue profiling of NAA‐metabolizing enzymes revealed that NAA was predominantly synthesized in non-cardiac tissues, where both its synthetase and hydrolase were highly expressed; however, the heart highly expressed the dicarboxylate transporter SLC13A3 but showed minimal hydrolase expression (\u003cstrong\u003eFig. 1d\u003c/strong\u003e). Together, these findings indicate that renal injury drives systemic accumulation of \u003cem\u003eN\u003c/em\u003e-acetylated amino acids, and results in disproportionate cardiac exposure to circulating NAA, owing to robust uptake capacity but limited hydrolytic activity in the heart.\u003c/p\u003e\n\u003cp\u003eNAA\u0026nbsp;impairs cardiac function and induces cardiac hypertrophy\u003c/p\u003e\n\u003cp\u003eTo investigate the effects of NAA on cardiac function, NAA was intraperitoneally injected in sham mice and mice with CKD. After eight weeks of treatment, echocardiographic analysis revealed impaired cardiac function in both groups (\u003cstrong\u003eExtended Data Fig. 3a\u003c/strong\u003e). NAA induced systolic dysfunction and myocardial hypertrophy in sham mice, with these deleterious effects being further exacerbated under CKD conditions (\u003cstrong\u003eFig. 1e\u003c/strong\u003e). Analysis of hematoxylin and eosin (H\u0026amp;E)-stained tissues revealed that NAA treatment induced cardiac hypertrophy under physiological and pathological conditions (\u003cstrong\u003eExtended Data Fig. 3b\u003c/strong\u003e). This finding was confirmed by wheat germ agglutinin (WGA) staining results, which showed enlargement of cardiomyocytes following NAA administration (\u003cstrong\u003eFig. 1f\u003c/strong\u003e). Moreover, Masson\u0026rsquo;s trichrome staining showed no significant fibrosis in the Sham + NAA group, suggesting that NAA alone did not induce cardiac fibrosis under normal conditions (\u003cstrong\u003eFig. 1g\u003c/strong\u003e). In contrast, NAA treatment markedly aggravated interstitial collagen deposition in the CKD + NAA group, indicating that NAA exacerbates cardiac fibrosis in the CKD setting (\u003cstrong\u003eFig. 1g\u003c/strong\u003e). Injection of NAA in sham and mice with CKD did not cause significant changes in renal function, as evidenced by comparable BUN levels and unaltered renal histology (\u003cstrong\u003eExtended Data Fig. 3c-e\u003c/strong\u003e); these findings indicated that the cardiac effects of NAA were not secondary to altered renal function.\u0026nbsp;Overall,\u0026nbsp;NAA induces maladaptive cardiac remodelling and functional deterioration, which is exacerbated in the context of renal dysfunction.\u003c/p\u003e\n\u003cp\u003eFurthermore, NAA treatment led to a significant increase in the primary neonatal rat cardiomyocytes (NRCMs) cell area (\u003cstrong\u003eFig. 1h\u003c/strong\u003e), and levels of atrial natriuretic peptide (ANP), BNP, and \u0026beta;-myosin heavy chain (\u0026beta;-MHC), which are molecular markers indicative of pathological hypertrophy (\u003cstrong\u003eFig. 1i\u003c/strong\u003e). In contrast, treatment with aspartate did not upregulate the expression of these markers, indicating that the pro-hypertrophic effect of NAA is not mediated through its hydrolysis to aspartate (\u003cstrong\u003eExtended Data Fig. 3f\u003c/strong\u003e). Together, these data support the notion that NAA impairs cardiac function and promotes hypertrophy, contributing to cardiac remodelling at the organ and cellular levels.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eNAA binds to and inhibits MDH1\u003c/p\u003e\n\u003cp\u003eTo elucidate the molecular mechanism by which NAA contributes to cardiac dysfunction, we employed an activity-based protein profiling (ABPP) approach using NAA probes as bait. Photoaffinity-tagged NAA probes were synthesized to enable enrichment and visualisation of interacting proteins (\u003cstrong\u003eExtended Data Fig. 4a\u003c/strong\u003e). The pro-hypertrophic effects of probe-tagged NAA in cardiomyocytes were comparable to those of non-tagged NAA (\u003cstrong\u003eExtended Data Fig. 4b\u003c/strong\u003e), validating the functional integrity probe-tagged NAA. The NAA probes were incubated with freshly prepared heart lysates for target labelling, and an equivalent concentration of non-tagged NAA was used as a control (\u003cstrong\u003eExtended Data Fig. 4c\u003c/strong\u003e). In the NAA probe group, 56 candidate proteins were specifically enriched (\u003cstrong\u003eExtended Data Fig. 4d\u003c/strong\u003e). The top 10 proteins were ranked according to their peptide-spectrum matches (PSMs) and coverage (\u003cstrong\u003eFig. 2a\u003c/strong\u003e).\u003c/p\u003e\n\u003cp\u003eAmong the 10 candidate proteins, MDH1 was prioritized as a potential target because it is a key enzyme in the malate\u0026ndash;aspartate shuttle\u003csup\u003e23-25\u003c/sup\u003e; moreover, the aspartate-derived metabolite NAA is structurally similar to the MDH1 substrate oxaloacetate (OAA) (\u003cstrong\u003eExtended Data Fig. 4e\u003c/strong\u003e). In this shuttle, aspartate undergoes transamination to form OAA, which is subsequently reduced to malate (Mal) by MDH1 (\u003cstrong\u003eExtended Data Fig.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e4f\u003c/strong\u003e). Drug affinity responsive target stability (DARTS) assay results revealed that incubation of purified recombinant MDH1 with NAA increased its resistance to Pronase digestion, suggesting that NAA binds to and stabilizes MDH1 (\u003cstrong\u003eFig. 2b\u003c/strong\u003e). In contrast, no such protective effect was observed for purified recombinant MDH2, the mitochondrial isoform that converts Mal to OAA (\u003cstrong\u003eExtended Data Fig.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e4g\u003c/strong\u003e).\u003c/p\u003e\n\u003cp\u003eSince the cardiac dysfunction and hypertrophy phenotypes were more pronounced than fibrosis, and that MDH1 primarily functions in cardiomyocytes, we analysed the effects of NAA on cardiomyocytes. Tail-vein injection of adeno-associated virus 9 (AAV9) carrying the cardiac troponin T (cTnT) promoter\u0026ndash;driven sh\u003cem\u003eMdh1\u003c/em\u003e resulted in cardiomyocyte-specific knockdown of \u003cem\u003eMdh1\u003c/em\u003e, which was consistent with the predominant expression of MDH1 in cardiomyocytes (\u003cstrong\u003eExtended Data Fig. 4h\u003c/strong\u003e). In \u003cem\u003eMdh1\u003c/em\u003e-knockdown mice, NAA treatment did not exacerbate cardiac dysfunction compared with that in untreated knockdown mice, indicating that MDH1 is the primary target that mediates the effects of NAA on cardiac function (\u003cstrong\u003eFig. 2c,\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;Extended Data Fig. 4i)\u003c/strong\u003e.\u003c/p\u003e\n\u003cp\u003eNext, we elucidated the effects of NAA on MDH1 function. Surface plasmon resonance (SPR) assays were conducted to evaluate the binding affinities of OAA and NAA to MDH1 (\u003cstrong\u003eFig. 2d, e\u003c/strong\u003e). The equilibrium dissociation constant (\u003cem\u003eK\u003csub\u003eD\u003c/sub\u003e\u003c/em\u003e) of NAA (7.19 \u0026mu;M) was higher than that of OAA (2.03 \u0026mu;M),\u0026nbsp;indicating that although NAA binds MDH1, its affinity to MDH1 is weaker than that of OAA; however, when MDH1 was pre-incubated with 10 \u0026mu;M NAA, the \u003cem\u003eK\u003csub\u003eD\u003c/sub\u003e\u003c/em\u003e value for OAA binding increased to 8.62 \u0026mu;M, suggesting that NAA inhibits the interaction between OAA and MDH1 (\u003cstrong\u003eFig. 2f, g\u003c/strong\u003e).\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eTo explore the molecular basis of NAA binding, we performed molecular docking and molecular dynamics simulations. NAA occupies the substrate-binding pocket of MDH1\u003csup\u003e26\u003c/sup\u003e, engaging key residues also involved in OAA recognition (\u003cstrong\u003eFig. 2h\u003c/strong\u003e). The protein Root Mean Square Deviation (RMSD) remained stable for both MDH1-OAA (1.0\u0026ndash;2.0 \u0026Aring;) and MDH1-NAA (0.75\u0026ndash;2.0 \u0026Aring;) complexes (\u003cstrong\u003eExtended Data Fig. 5a, b\u003c/strong\u003e). OAA interacted with pocket residues mainly through hydrogen bonds (binding energy, -16.94 kcal/mol), whereas NAA engaged more residues via additional hydrogen bonds and water bridges, resulting in stronger binding than that of OAA (-36.00 kcal/mol) (\u003cstrong\u003eExtended Data Fig. 5c, d\u003c/strong\u003e). These simulations suggest that NAA engages MDH1 in a binding mode analogous to that of OAA but with more extensive interactions. NAA binding did not occur in the MDH1 quintuple mutant (R92A/R98A/N131A/R162A/S242A, MDH1-5A)\u0026nbsp;(\u003cstrong\u003eExtended Data Fig. 5e\u003c/strong\u003e), confirming the critical role of these residues and validating the binding mode of NAA to MDH1; these residues correspond to high-frequency interaction sites and constitute the OAA-binding pocket\u003csup\u003e26\u003c/sup\u003e.\u0026nbsp;Notably, OAA interacted frequently with NADH, whereas NAA did not (\u003cstrong\u003eExtended Data Fig. 5c, d\u003c/strong\u003e); the probable reason is that the amide carbonyl in NAA is resonance-stabilized and thus less reducible than the keto group in OAA\u0026nbsp;(\u003cstrong\u003eExtended Data Fig. 4e\u003c/strong\u003e).\u0026nbsp;This finding suggests that NAA may not readily undergo the reduction reaction catalysed by MDH1. Overall, NAA can engage MDH1, potentially interfering with OAA binding and enzymatic activity.\u003c/p\u003e\n\u003cp\u003eTo assess enzymatic inhibition, we conducted in vitro assays to monitor NADH consumption at 340 nm during MDH1-catalysed OAA reduction (\u003cstrong\u003eFig. 2i,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 4\u003c/strong\u003e\u003cstrong\u003ef\u003c/strong\u003e). NAA is not reduced by MDH1; however, NAA inhibits the catalysis of OAA by MDH1 (\u003cstrong\u003eFig. 2i, j\u003c/strong\u003e). Together, these findings demonstrate that NAA occupies the substrate-binding pocket of MDH1 and inhibits its activity, providing a mechanistic basis for the effects of NAA on cardiac metabolism and function.\u003c/p\u003e\n\u003cp\u003eNAA-induced MDH1 inhibition disrupts cardiomyocyte metabolic homeostasis\u003c/p\u003e\n\u003cp\u003eTo assess whether NAA inhibits MDH1 activity in primary cardiomyocytes, we measured intracellular levels of OAA and Mal following NAA treatment. The MDH1 substrate OAA accumulated, whereas Mal levels remained largely unchanged (\u003cstrong\u003eFig. 3a, b\u003c/strong\u003e); the probable reason is that intracellular OAA exists at low concentrations and readily reflects perturbations, whereas Mal is generated and consumed through multiple metabolic pathways. Using the Mal/OAA ratio as a proxy for MDH1 enzymatic activity, we observed that NAA markedly inhibited MDH1 activity (\u003cstrong\u003eFig. 3c\u003c/strong\u003e).\u003c/p\u003e\n\u003cp\u003eSince the malate\u0026ndash;aspartate shuttle is essential for transferring cytosolic reducing equivalents into mitochondria to sustain oxidative phosphorylation in cardiomyocytes\u003csup\u003e23-25\u003c/sup\u003e, we examined whether NAA disrupts metabolic homeostasis in primary cardiomyocytes. To assess the effects of NAA on redox homeostasis, we monitored NAD⁺/NADH dynamics in primary cardiomyocytes using non-targeted SoNar (cyto-SoNar) or mitochondria-targeted SoNar (mito-SoNar). Real-time measurements revealed that NAA treatment led to a decrease in the cytosolic NAD⁺/NADH ratio, while slowing the rate of NAD⁺/NADH increase in mitochondria (\u003cstrong\u003eFig. 3d, e\u003c/strong\u003e). Consistently, at the whole-cell level, NAA treatment markedly decreased the overall NAD⁺/NADH ratio in cardiomyocytes (\u003cstrong\u003eFig. 3f\u003c/strong\u003e). This metabolic impairment was accompanied by a significant decrease in intracellular ATP levels, reflecting compromised mitochondrial energy production (\u003cstrong\u003eFig. 3g\u003c/strong\u003e). These findings indicate that NAA lowers cytosolic NAD⁺/NADH ratio and dampens the mitochondrial redox response, which is consistent with inhibition of the MDH1-dependent malate\u0026ndash;aspartate shuttle.\u003c/p\u003e\n\u003cp\u003eTo elucidate how NAA perturbs central carbon metabolism, we performed \u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e6\u003c/sub\u003e-glucose and \u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e5\u003c/sub\u003e-glutamine tracing in primary cardiomyocytes to assess the effects of NAA on glycolysis and tricarboxylic acid (TCA) cycle activity (\u003cstrong\u003eExtended Data Fig. 5f, g\u003c/strong\u003e). After NAA treatment, levels of labelled upstream glycolytic intermediates glucose-6-phosphate (G6P), fructose-1,6-bisphosphate (FBP), and glyceraldehyde-3-phosphate (G3P) remained unaltered, whereas labelled 3-phosphoglycerate (3PG) and pyruvate M + 3 levels decreased (\u003cstrong\u003eFig. 3h\u003c/strong\u003e). This finding aligns with the observed cytosolic NAD⁺/NADH decline caused by MDH1 inhibition, because glyceraldehyde-3-phosphate dehydrogenase (GAPDH) requires NAD⁺ for its activity. Moreover, acetyl-CoA M + 2 labeling remained unchanged, but incorporation of both \u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e6\u003c/sub\u003e-glucose and \u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e5\u003c/sub\u003e-glutamine into TCA cycle intermediates was markedly reduced, indicating suppression of mitochondrial oxidative metabolism (\u003cstrong\u003eFig. 3i, j\u003c/strong\u003e). Accumulation of unlabelled OAA, predominantly derived from aspartate, suggested dysfunction of the cytosolic MDH1-mediated reaction (\u003cstrong\u003eFig. 3k\u003c/strong\u003e). Collectively, these findings demonstrate that NAA inhibits MDH1 activity and disrupts redox-linked metabolite shuttling and mitochondrial metabolism, ultimately leading to energy insufficiency and cardiomyocyte hypertrophy.\u003c/p\u003e\n\u003cp\u003e\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eSIRT2 is an NAA hydrolase\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eBeyond MDH1, our ABPP screen also identified SIRT2 and SIRT3 as NAA-binding candidates (\u003cstrong\u003eFig. 2a\u003c/strong\u003e). Since sirtuins are NAD⁺-dependent enzymes with amide bond-hydrolysing activity\u003csup\u003e14-18\u003c/sup\u003e, this raised the possibility that certain sirtuins might recognise and hydrolyse NAA as a small-molecule amide substrate, in addition to their canonical protein deacetylase activity.\u003c/p\u003e\n\u003cp\u003eTo investigate whether members of the sirtuin family possess NAA hydrolase activity, we purified recombinant homo SIRT1\u0026ndash;SIRT7 heterologously expressed in bacteria and conducted an enzyme-coupled assay to continuously monitor the hydrolysis process (\u003cstrong\u003eExtended Data Fig. 6a\u003c/strong\u003e). This assay is analogous to that used for measuring the deacetylase activity of sirtuins and uses NADPH consumption as a surrogate readout for the first-step reaction\u003csup\u003e27\u003c/sup\u003e. NADPH levels were quantified based on the intrinsic fluorescence (excitation at approximately 340 nm; emission at 460 nm). Among the seven sirtuins, SIRT2 and SIRT3 exhibited robust NAA hydrolase activity in an NAD\u003csup\u003e+\u003c/sup\u003e-dependent manner\u0026nbsp;(\u003cstrong\u003eFig. 4a, b\u003c/strong\u003e), whereas the others showed minimal or undetectable activity (\u003cstrong\u003eExtended Data Fig.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e6b\u003c/strong\u003e). To validate the NAA hydrolase activity of SIRT2 and SIRT3, we incubated these enzymes with NAA and quantified NAA and its hydrolysis product, aspartate, using LC-MS/MS (\u003cstrong\u003eFig. 4c\u003c/strong\u003e). NAA hydrolysis was determined to be a previously unrecognized enzymatic activity of SIRT2 and SIRT3, with estimated catalytic efficiencies (\u003cem\u003eK\u003csub\u003ecat\u003c/sub\u003e/K\u003csub\u003em\u003c/sub\u003e\u003c/em\u003e) of 1.39 \u0026times; 10\u003csup\u003e4\u003c/sup\u003e and 2.97 \u0026times; 10\u003csup\u003e4\u0026nbsp;\u003c/sup\u003es\u003csup\u003e-1\u003c/sup\u003eM\u003csup\u003e-1\u003c/sup\u003e, respectively (\u003cstrong\u003eFig. 4d\u003c/strong\u003e).\u003c/p\u003e\n\u003cp\u003eSubsequently, we examined whether SIRT2 and SIRT3 physically associate with NAA. DARTS assay results revealed that NAA increased the Pronase resistance of SIRT2 and SIRT3 (\u003cstrong\u003eFig. 4e, f\u003c/strong\u003e), indicating the occurrence of a specific interaction, whereas no such effect was observed using aspartate (\u003cstrong\u003eExtended Data Fig.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e6c, d\u003c/strong\u003e). To quantify the binding affinities, we performed SPR analysis. NAA exhibited concentration-dependent binding to SIRT2 and SIRT3 (\u003cstrong\u003eFig. 4g,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eh\u003c/strong\u003e), with \u003cem\u003eK\u003csub\u003eD\u003c/sub\u003e\u003c/em\u003e values of 6.32 and 2.85 \u0026mu;M, respectively (\u003cstrong\u003eFig. 4i\u003c/strong\u003e).\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eSince NAA treatment did not alter SIRT2 nuclear\u0026ndash;cytosolic distribution\u003csup\u003e28\u003c/sup\u003e (\u003cstrong\u003eExtended Data Fig. 6e\u003c/strong\u003e) and MDH1 is a cytosolic enzyme, we focused on the hydrolytic activity of SIRT2 toward NAA, in contrast to that of SIRT3, which is predominantly localized in mitochondria\u003csup\u003e29\u003c/sup\u003e. To gain structural insights into the interaction between SIRT2 and NAA, we performed molecular docking and molecular dynamics simulations. NAA stably interacted with the L4 loop of SIRT2, positioning it within the active-site pocket of the enzyme\u003csup\u003e30,31\u003c/sup\u003e (\u003cstrong\u003eExtended Data Fig. 7a\u003c/strong\u003e). The overall protein structure remained stable during molecular dynamics simulations (RMSD: 1.6\u0026ndash;3.0 \u0026Aring;;\u003cstrong\u003e\u0026nbsp;Extended Data Fig. 7b\u003c/strong\u003e), supporting a well-defined binding mode.\u0026nbsp;NAA further interacted with key residues in the L1 and L5 loops involved in substrate recognition and entry\u003csup\u003e31\u003c/sup\u003e (\u003cstrong\u003eExtended Data Fig. 7c\u003c/strong\u003e), indicating a small-molecule binding mode stabilised by coordinated non-covalent interactions. In contrast, interactions between SIRT2 and its canonical protein substrate tubulin alpha-1A (TUBA1A) involve 93 residues across two spatially separated sites, enriched in charged and polar residues, forming extensive hydrogen-bond networks characteristic of protein\u0026ndash;protein interactions (\u003cstrong\u003eExtended Data Fig. 7d, e\u003c/strong\u003e). Together, these observations suggest that SIRT2 can recognise and hydrolyse NAA, showing that NAA binds in a small-molecule-specific mode within the active pocket.\u003c/p\u003e\n\u003cp\u003eIn \u003cem\u003eSirtuin2\u003c/em\u003e-global knockout mice, NAA levels were significantly elevated in the circulation and heart, suggesting that SIRT2 functions as a physiological NAA hydrolase in vivo (\u003cstrong\u003eFig. 4j, k\u003c/strong\u003e). In contrast, renal NAA levels remained largely unchanged, likely due to the high expression levels of known NAA hydrolases in the kidney (\u003cstrong\u003eFig.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003e1d and 4l\u003c/strong\u003e). Based on these results, we propose the mechanism of NAA hydrolysis by SIRT2. In this process, the acetyl group of NAA is transferred to NAD\u003csup\u003e+\u003c/sup\u003e, yielding nicotinamide, \u003cem\u003eO\u003c/em\u003e-acetyl-ADP-ribose, and free aspartate (\u003cstrong\u003eFig. 4m\u003c/strong\u003e). These findings support the conclusion that SIRT2 acts as a physiological NAA hydrolase.\u003c/p\u003e\n\u003cp\u003eSIRT2-mediated hydrolysis of NAA relieves MDH1 inhibition and improves cardiac function\u003c/p\u003e\n\u003cp\u003eWe next investigated whether SIRT2-mediated hydrolysis of NAA could mitigate the detrimental effects of NAA on the heart. Overexpression of SIRT2 in primary cardiomyocytes significantly reduced NAA levels, confirming the physiological NAA-hydrolysing activity of SIRT2 (\u003cstrong\u003eFig. 5a\u003c/strong\u003e). SIRT2 overexpression restored cytosolic and mitochondrial NAD⁺/NADH ratios, indicating alleviation of MDH1 inhibition (\u003cstrong\u003eFig. 5b, c\u003c/strong\u003e). In addition, the hypertrophic phenotype of the cardiomyocytes was markedly alleviated (\u003cstrong\u003eFig. 5d\u003c/strong\u003e). Overall, SIRT2-mediated NAA hydrolysis exerted a protective effect on cardiomyocyte metabolism and function.\u003c/p\u003e\n\u003cp\u003eGiven the reduction in cardiac SIRT2 expression levels in mine with CKD (\u003cstrong\u003eExtended Data Fig. 8a\u003c/strong\u003e), we next examined whether restoration of SIRT2 could counteract NAA-induced cardiac toxicity. To separately evaluate the ability of SIRT2 to rescue NAA-induced toxicity and rectify CKD-associated endogenous NAA elevation, we performed two parallel sets of experiments using exogenous administration and chronic disease models. Tail-injected AAV9 enabled cardiomyocyte-specific cTnT promoter\u0026ndash;driven SIRT2 overexpression (\u003cstrong\u003eExtended Data Fig. 8b, c\u003c/strong\u003e). In control mice, SIRT2 overexpression effectively reversed the intraperitoneal administration-induced NAA accumulation in cardiac tissues and sera (\u003cstrong\u003eFig. 5e,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 8d\u003c/strong\u003e), thereby restoring cardiac MDH1 activity and ATP levels (\u003cstrong\u003eFig. 5f, g\u003c/strong\u003e). Similarly, in mice with CKD, SIRT2 overexpression normalized cardiac NAA levels (\u003cstrong\u003eFig. 5h\u003c/strong\u003e), partially alleviated the decrease in MDH1 activity and ATP levels (\u003cstrong\u003eFig. 5i, j\u003c/strong\u003e), and slightly reduced the elevated serum NAA levels (\u003cstrong\u003eExtended Data Fig. 8e\u003c/strong\u003e), likely due to the persistent systemic metabolic impairment associated with renal dysfunction.\u0026nbsp;Furthermore, SIRT2 overexpression improved cardiac systolic performance (\u003cstrong\u003eFig. 5k, l,\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;Extended Data Fig. 8f, g\u003c/strong\u003e) and reduced cardiomyocyte hypertrophy (\u003cstrong\u003eFig. 5m, n\u003c/strong\u003e) in NAA-treated mice and mice with CKD. Moreover, western blotting confirmed downregulation of hypertrophic markers in SIRT2-overexpressing hearts (\u003cstrong\u003eFig. 5o, p\u003c/strong\u003e). Masson\u0026rsquo;s trichrome staining analysis revealed that NAA treatment alone did not result in cardiac fibrosis, which is consistent with previous observations \u003cstrong\u003e(Fig. 1g,\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 8h)\u003c/strong\u003e; moreover, SIRT2 overexpression markedly attenuated cardiac fibrosis in mice with CKD \u003cstrong\u003e(\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 8i).\u003c/strong\u003e Together, these results demonstrate that cardiomyocyte-specific SIRT2 overexpression counteracts exogenous NAA overload and CKD-associated endogenous NAA accumulation, thereby protecting against cardiac dysfunction and remodelling.\u003c/p\u003e"},{"header":"Discussion","content":"\u003cp\u003eWe discovered that NAA is a pathogenic metabolic signal that links renal dysfunction to cardiac remodelling. NAA aberrantly accumulates under CKD conditions and drives cardiac dysfunction and cardiomyocyte hypertrophy. Mechanistically, NAA competitively inhibits MDH1, leading to disruption of mitochondrial metabolism and impairment of energy homeostasis in cardiomyocytes. Therefore, NAA is a maladaptive metabolic mediator in the kidney\u0026ndash;heart axis. Moreover, SIRT2 is a previously unrecognized NAA hydrolase, which detoxifies excessive NAA and counteracts NAA-induced metabolic stress, cardiac dysfunction, and structural remodelling \u003cstrong\u003e(\u003c/strong\u003e\u003cstrong\u003eExtended Data Fig. 8j)\u003c/strong\u003e.\u003c/p\u003e\n\u003cp\u003eThese findings underscore NAA as a potential mediator of kidney\u0026ndash;heart metabolic crosstalk in a broad range of renal pathologies\u003csup\u003e32\u003c/sup\u003e. Further studies are warranted to determine whether circulating NAA levels predicts cardiac outcomes across kidney diseases and to evaluate its value as a diagnostic biomarker in cardiorenal syndromes. Moreover, SIRT2 exerts a cardioprotective effect by mitigating NAA-induced hypertrophic remodelling, suggesting that enhancing its activity may help preserve cardiac function under metabolic stress\u003csup\u003e33\u003c/sup\u003e.\u003c/p\u003e\n\u003cp\u003eSirtuins are NAD⁺-dependent metabolic sensors\u003csup\u003e10-12,34\u003c/sup\u003e, and we discovered that SIRT2 acts as an NAA hydrolase, expanding the known functions of sirtuins to include the regulation of small-molecule metabolites. This finding raises the possibility that SIRT2-mediated hydrolysis of NAA constitutes an adaptive response to energy fluctuations and modulates energy homeostasis in the heart and other high-energy-demanding tissues. Although identified in the heart, the SIRT2\u0026ndash;NAA\u0026ndash;MDH1 axis is likely to represent a general mechanism, as SIRT2 and MDH1 are widely expressed core metabolic enzymes. Dysregulation of the SIRT2\u0026ndash;NAA\u0026ndash;MDH1 axis may contribute to mitochondrial dysfunction-related diseases. Future studies need to delineate its regulatory circuitry and broader physiological and therapeutic relevance in metabolic disorders and aging.\u003c/p\u003e\n\u003cp\u003eBesides metabolic regulation, our findings may have broader translational implications. Deficiency of ACY2, the canonical NAA hydrolase, causes Canavan disease, a lethal leukodystrophy characterized by NAA accumulation\u003csup\u003e9,35,36\u003c/sup\u003e. The intrinsic NAA hydrolase activity of SIRT2 raises the possibility of compensatory or therapeutic strategies to reduce NAA toxicity, though this remains to be tested.\u003c/p\u003e\n\u003cp\u003eBesides the conventionally known roles of \u003cem\u003eN\u003c/em\u003e-acetylated amino acids as terminal metabolites, the physiological and pathological functions of these metabolites remain largely unexplored\u003csup\u003e5,37\u003c/sup\u003e. While the idea that NAA-derived acetate feeds acetyl-CoA pools is conceptually intriguing\u003csup\u003e38\u003c/sup\u003e, ACY2 or SIRT2 hydrolysis produces acetate or \u003cem\u003eO\u003c/em\u003e-acetyl-ADP-ribose rather than acetyl-CoA, underscoring the need to reconsider the metabolic efficiency of this route. Interestingly, a recent study proposed that NAA may act as a structural analog of NAG to activate carbamoyl-phosphate synthetase 2, aspartate transcarbamylase, and dihydroorotase (CAD)\u003csup\u003e39\u003c/sup\u003e, conceptually supporting the idea that structural similarity guides NAA target identification. NAA has also been implicated in protein acetylation, though the observed effects are variable\u003csup\u003e40,41\u003c/sup\u003e. Thus, although NAA contains an acetyl moiety, focusing solely on the acetyl group risks overlooking the biological specificity conferred by its amino acid backbone.\u003c/p\u003e\n\u003cp\u003eThe biological specificity of individual \u003cem\u003eN\u003c/em\u003e-acetylated amino acids is likely conferred by the chemical properties of their underlying amino acid scaffold. Acetylation alters reactivity, p\u003cem\u003eK\u003csub\u003ea\u003c/sub\u003e\u003c/em\u003e, and polarity, with potential consequences for transport, compartmentalization, and enzymatic recognition. Importantly, the incorporation of an acetyl group via acetyl-CoA confers nutrient-related signalling properties to \u003cem\u003eN\u003c/em\u003e-acetylated amino acids\u003csup\u003e42,43\u003c/sup\u003e, positioning them as integrators of amino acid and energy metabolism. In line with this view, both NAG and NAA function as metabolic signalling modulators, activating CPS1 or inhibiting MDH1, respectively, coupling their abundance to broader metabolic network activity.\u0026nbsp;Together, these insights suggest that \u003cem\u003eN\u003c/em\u003e-acetylated amino acids constitute an underappreciated class of metabolic signals with roles extending far beyond their traditional designation as terminal metabolites.\u003c/p\u003e\n\u003cp\u003eIn a broader context, small-molecule acyl-amino acids are emerging as metabolic regulators. \u003cem\u003eN\u003c/em\u003e-acetyltaurine modulates body weight control and energy balance via GDF15-GFRAL pathway\u003csup\u003e44\u003c/sup\u003e, while \u003cem\u003eN\u003c/em\u003e-lactoyl-phenylalanine (Lac-Phe) acts as a blood-borne metabolic that suppress feeding and mitigates obesity\u003csup\u003e45,46\u003c/sup\u003e. Additionally, ketone body-amino acid conjugates were recently shown to suppress feeding and contribute to the maintenance of energy homeostasis\u003csup\u003e47\u003c/sup\u003e. Together with our findings, these studies highlight acyl-amino acids as an emerging class of metabolic signals that merits further investigation in energy homeostasis and disease pathogenesis.\u003c/p\u003e\n\u003cp\u003eThis study has several limitations. First, although both NAG and NAA were associated with CKD-associated cardiomyopathy, we primarily focused on NAA for mechanistic investigations. NAG may also exert cardiac effects apart from its known role in CPS1 activation. Second, SIRT3 showed NAA-hydrolysis activity in vitro, raising the possibility of mitochondrial-specific regulation. Third, we did not assess whether the SIRT2\u0026ndash;NAA\u0026ndash;MDH1 axis operates in other tissues or contributes to systemic metabolic regulation, and this warrants further investigation.\u003c/p\u003e\n\u003cp\u003eIn summary, our study reveals that NAA is a previously unrecognized metabolic risk factor linking renal dysfunction to cardiac remodelling. NAA is a bioactive acetylated metabolite that modulates MDH1 activity and undergoes SIRT2-mediated hydrolysis, which suggests that NAA is embedded in pathways tightly coupled to cellular energy homeostasis. Moreover, the discovery of SIRT2 as an NAA hydrolase expands the enzymatic landscape of sirtuins and uncovers a direct role for them in the regulating of small-molecule metabolites, thereby integrating the control of protein function and signalling metabolites to enable rapid, systemic metabolic adaptation and the maintenance of homeostasis. Overall, \u003cem\u003eN\u003c/em\u003e-acetylated amino acids emerge as underappreciated metabolic signals with systemic relevance, warranting further study to inform strategies for cardiorenal and metabolic disease management.\u003c/p\u003e"},{"header":"Methods","content":"\u003cp\u003eMaterials\u003c/p\u003e\n\u003cp\u003eChemicals and reagents were obtained from the following sources: \u003cem\u003eN\u003c/em\u003e-acetylaspartate (Cat. No.: 00920, Sigma-Aldrich), NADH (Cat. No.: HY-F0001R, MedChemExpress), NAD\u003csup\u003e+\u003c/sup\u003e (Cat. No.: HY-B0445, MedChemExpress), NADPH (Cat. No.: HY-113324, MedChemExpress), oxaloacetate (Cat. No.: O4126, Sigma-Aldrich), glutamate dehydrogenase (Cat. No.: G2626, Sigma-Aldrich), Pronase (Cat. No.: 10165921001, Sigma-Aldrich), type II collagenase (Cat. No.: C6885, Sigma-Aldrich), photoaffinity-tagged NAA probe (Cat. No.: 1132432, WuXi AppTec), tris(2-carboxyethyl)phosphine hydrochloride (TCEP; Cat. No.: C4706, Sigma-Aldrich), tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA; Cat. No.: 678937, Sigma-Aldrich), CuSO4 (Cat. No.: C1297, Sigma-Aldrich), Biotin-azide (Cat. No.: HY-129832, MedChemExpress), Ni-NTA beads 6FF (Cat. No.: N30210, LABLEAD), [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e6\u003c/sub\u003e]-D-glucose (Cat. No.: HY-B0389A, MedChemExpress), [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e5\u003c/sub\u003e]-L-glutamine (Cat. No.: HY-N0390S1, MedChemExpress), cyto-SoNar (Cat. No.: ADSON1001-S, Provoson), mito-SoNar (Cat. No.: ADSON1003-S, Provoson), anti-MDH1 (Cat. No.: 15904-1-AP, Proteintech), anti-\u0026beta;-MHC (Cat. No.: 22280-1-AP, Proteintech), anti-ANP (Cat. No.: A14755, ABclonal), anti-BNP (Cat. No.: A23996, ABclonal), anti-SIRT2 (Cat. No.: A12575, ABclonal), anti-ACY1 (Cat. No.: A6351, ABclonal), anti-ACY2 (Cat. No.: A7271, ABclonal), anti-NAT8L (Cat. No.: 23841-1-AP, Proteintech), anti-SLC13A3 (Cat. No.: 26184-1-AP, Proteintech), anti-GAPDH (Cat. No.: A19056, ABclonal), anti-\u0026alpha;-Tubulin (Cat. No.: A6830, ABclonal), goat anti-mouse IgG antibody (Cat. No.: A21001S, Abmart), and goat anti-rabbit IgG antibody (Cat. No.: M21002L, Abmart).\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eStudy participants and blood collection\u003c/p\u003e\n\u003cp\u003eAdult patients with CKD were recruited at Shanghai Jiao Tong University School of Medicine, Xinhua Hospital, Department of Nephrology and Cardiology. The study design was approved and supervised by the Ethics Committee of Xinhua Hospital (Approval No.: XHEC-C-2024-185-2) in accordance with the Declaration of Helsinki. All participants were fully informed and signed the informed consent prior to enrolment. CKD was defined as an estimated glomerular filtration rate (eGFR) \u0026lt; 60 mL/min/1.73 m\u003csup\u003e2\u003c/sup\u003e and peritoneal dialysis persisting for at least three months, and further supported by elevated BUN and serum creatinine (Scr) levels\u003csup\u003e48\u003c/sup\u003e. Participants were excluded if they had any of the following conditions\u003csup\u003e49\u003c/sup\u003e: hemodialysis, history of heart failure, hypertrophic or restrictive cardiomyopathy, infiltrative cardiomyopathy (including cardiac amyloidosis), congenital heart disease, constrictive pericarditis, moderate or greater valvular disease, isolated pulmonary arterial hypertension, heart transplantation, diabetes mellitus, lupus nephritis, anti-neutrophil cytoplasmic antibody-associated vasculitis, or any known malignancy.\u003c/p\u003e\n\u003cp\u003eVenous blood (3 mL) was collected from each patient into vacuum blood collection tubes containing a clot activator and inert separation gel. After standing at room temperature for 15 min to allow clot formation, samples were centrifuged at 1,500 \u0026times;\u003cem\u003e\u0026nbsp;g\u003c/em\u003e for 10 min. The resulting serum was carefully transferred into a new 1.5 mL microcentrifuge tube and stored in -80 \u0026deg;C refrigerators.\u003c/p\u003e\n\u003cp\u003eQuantification of serum\u0026nbsp;metabolites\u003c/p\u003e\n\u003cp\u003eParticipants\u0026rsquo; serum metabolites were analysed at Applied Protein Technology (APTBIO, Shanghai, China) using ultra-performance liquid chromatography coupled to tandem mass spectrometry (UPLC-MS/MS). Serum samples were thawed on ice, and 400 \u0026mu;L of cold methanol/acetonitrile (1:1, v/v) extraction solvent containing stable-isotope internal standards was added to precipitate proteins and extract metabolites. The mixture was vortexed, transferred to a new centrifuge tube, and centrifuged at 14,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 20 min at 4 \u0026deg;C. The supernatant was collected and dried under vacuum. For UPLC-MS/MS analysis, the dried extracts were reconstituted in 100 \u0026mu;L of acetonitrile/water (1:1; v/v), mixed, and centrifuged at 14,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 15 min at 4 \u0026deg;C. The resulting supernatant was injected for analysis.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eAnalyses were performed using a UHPLC system (1290 Infinity LC, Agilent Technologies) coupled to a QTRAP mass spectrometer (6500+; AB Sciex). Metabolites were separated on both HILIC and reversed-phase C18 columns (Waters UPLC BEH Amide and Waters UPLC BEH C18, respectively; 2.1 \u0026times; 100 mm, 1.7 \u0026mu;m). Data acquisition was conducted in both positive and negative ion modes using multiple reaction monitoring (MRM), and quantification was performed using MultiQuant or Analyst software. Quality control (QC) samples were interspersed throughout the run to assess system stability, and metabolites with a coefficient of variation (CV) \u0026lt; 30% in QC samples were retained for further analysis. For metabolite determination, a comprehensive set of rigorous quality control/assurance procedures was employed to ensure consistently high-quality analytical results by monitoring every step from sample receipt at the laboratory to final deliverables.\u003c/p\u003e\n\u003cp\u003eExperimental animals and animal care\u003c/p\u003e\n\u003cp\u003eAll animal experiments were approved and conducted in accordance with the guidelines of the Animal Care Committee at Xinhua Hospital, Shanghai Jiao Tong University School of Medicine (Approval No.: XHEC-NSFC-2023-100).\u003c/p\u003e\n\u003cp\u003eThe CKD mouse models were established using the 5/6 nephrectomy surgery\u003csup\u003e50,51\u003c/sup\u003e. Male C57BL/6 mice, aged six weeks, were purchased from GemPharmatech Co., Ltd. (Nanjing, China). All mice were housed in a specific pathogen-free (SPF) facility under controlled temperature (20\u0026ndash;22 \u0026deg;C), humidity (40\u0026ndash;60%), and a 12 h light/dark cycle, with free access to food and water. Prior to surgery, mice were acclimatized to the new environment for two weeks. At eight weeks of age, CKD was induced using a two-step 5/6 nephrectomy. Briefly, mice were anesthetized with 2% isoflurane and placed on a warming pad to maintain body temperature. In the first surgery, approximately two-thirds of the left kidney was removed by resecting the upper and lower poles while carefully controlling hemostasis. After a seven-day recovery, the second surgery was performed to completely remove the right kidney. Sham-operated mice underwent identical surgical procedures without renal tissue removal. Mice were monitored daily for wound healing and general health.\u003c/p\u003e\n\u003cp\u003eFor pharmacological intervention, NAA (00920, Sigma-Aldrich) was dissolved in phosphate buffer (pH 7.2\u0026ndash;7.4) and administered intraperitoneally at 200 mg/kg daily for eight weeks. Control mice received equal volumes of phosphate buffer.\u003c/p\u003e\n\u003cp\u003eFour-week-old C57BL/6 \u003cem\u003eSirtuin2\u003csup\u003e-/-\u003c/sup\u003e\u0026nbsp;\u003c/em\u003emice were purchased from GemPharmatech Co., Ltd. (Nanjing, China). Genotypes were determined via PCR using genomic DNA obtained from tails; two primer pairs were used (\u003cem\u003eSirtuin2\u003csup\u003e-/-\u003c/sup\u003e\u0026nbsp;\u003c/em\u003eforward 1: 5\u0026prime;-CCTTAAACAGAACACCCCAGGG-3\u0026rsquo;, reverse 1: 5\u0026prime;-GAGATAGGTAGTTTTGGCTGACCTTG-3\u0026prime;; \u003cem\u003eSirtuin2\u003csup\u003e-/-\u003c/sup\u003e\u0026nbsp;\u003c/em\u003eforward 2: 5\u0026prime;-TAAGTTCCCCGTCCTTCAGCCA-3\u0026prime;, reverse 2: 5\u0026prime;-CCCAGAAGCTAGGTTCTTAATCATGT-3\u0026prime;). All mice were housed in a specific SPF facility under controlled temperature (20\u0026ndash;22 \u0026deg;C), humidity (40\u0026ndash;60%), and a 12 h light/dark cycle, with free access to food and water.\u003c/p\u003e\n\u003cp\u003eAAV9-mediated cardiomyocyte-specific \u003cem\u003eMdh1\u003c/em\u003e-knockdown\u003c/p\u003e\n\u003cp\u003eTo achieve \u003cem\u003eMdh1\u003c/em\u003e cardiomyocyte-specific knockdown, mice received a single tail-vein injection of AAV9-cTnT-sh\u003cem\u003eMdh1\u003c/em\u003e (GIEV5032496, GeneChem) at 1 \u0026times; 10\u003csup\u003e12\u003c/sup\u003e vg per mouse. AAV9 carrying shRNA to interfere with the mouse \u003cem\u003eMdh1\u003c/em\u003e gene were constructed and inserted into GV683, which contained the expression cassette cTnTp-EGFP-MIR155(MCS)-WPRE-SV40 PolyA. Knockdown efficiency was assessed by fluorescence, and the validated target sequence was: GACCCAGTATCCAGATGTCAA.\u003c/p\u003e\n\u003cp\u003eAAV9-mediated cardiomyocyte-specific SIRT2 overexpression\u003c/p\u003e\n\u003cp\u003eFor cardiomyocyte-specific overexpression of SIRT2, mice received a single tail-vein injection of AAV9-cTnT-SIRT2 (GOSV5019825, GeneChem) at 1 \u0026times; 10\u003csup\u003e12\u003c/sup\u003e vg per mouse. AAV9 carrying the mouse \u003cem\u003eSirtuin2\u003c/em\u003e-coading gene was constructed and inserted into GV831, which contained the expression cassette cTnTp-MCS-3Flag-FT2A-EGFP-WPRE-BGH polyA; the cassette was generated using the forward primer (5\u0026prime;-AAGGCTAGAGTACTGCTAGCCGCCACCATGGCCGAGCCGGACCCCTCTGACCCTCTGGAG-3\u0026prime;) and reverse primer (5\u0026prime;-TAGTCCATGGTGGCACCGGTCTGCTGTTCCTCTTTCTCTTTGG-3\u0026prime;). Successful transduction and SIRT2 overexpression were validated by fluorescence in myocardial tissue.\u003c/p\u003e\n\u003cp\u003eEchocardiography\u003c/p\u003e\n\u003cp\u003eTransthoracic echocardiography was performed to assess cardiac function using a high-frequency ultrasound system (Vevo 3100, VisualSonics) equipped with an MX550 transducer. Two-dimensional (B-mode) and M-mode images were obtained from the parasternal long-axis and short-axis views at the level of the papillary muscles. Left ventricular diameters (end-diastolic, LVEDD; end-systolic, LVESD), volumes (end-diastolic, LVEDV; end-systolic, LVESV), and posterior wall thickness at end-diastole (LVPWd) were measured from M-mode tracings. Fractional shortening (LVFS) and ejection fraction (LVEF) were determined, with three consecutive cardiac cycles analysed per mouse.\u003c/p\u003e\n\u003cp\u003eDetection of blood urea nitrogen (BUN) levels in mice\u003c/p\u003e\n\u003cp\u003eMouse BUN levels were measured using a Mouse Urea Nitrogen ELISA Kit (FT-P9S949X, Shanghai Fantai Biotechnology) according to the manufacturer\u0026rsquo;s protocol. In brief, 10 \u0026mu;L of serum was mixed with 40 \u0026mu;L diluent in precoated wells, followed by 100 \u0026mu;L HRP-conjugated reagent. Plates were incubated at 37 \u0026deg;C for 1 h, washed thoroughly, and incubated with chromogenic substrate for color development. After 15 min of dark incubation at 37 \u0026deg;C, the reaction was terminated, and absorbance at 450 nm was recorded using a\u0026nbsp;Multimode Microplate Reader (Spark, Tecan). Data were background-corrected, and BUN concentrations were calculated from standard curves.\u003c/p\u003e\n\u003cp\u003eHistology and staining\u003c/p\u003e\n\u003cp\u003eMice tissues were harvested, fixed overnight in 4% paraformaldehyde, dehydrated, and embedded in paraffin. Five-micrometer sections were cut and stained with hematoxylin and eosin (H\u0026amp;E) for morphological evaluation, wheat germ agglutinin (WGA) to visualize cardiomyocyte borders, and Masson\u0026rsquo;s trichrome to detect collagen deposition. For immunofluorescence, sections were incubated with primary antibodies at 4 \u0026deg;C overnight, followed by appropriate fluorophore-conjugated secondary antibodies. Nuclei were counterstained with DAPI, and images were acquired using a fluorescence microscope (Leica Microsystems). Quantification was performed using ImageJ software (NIH).\u003c/p\u003e\n\u003cp\u003eIsolation and culture of primary neonatal rat cardiomyocytes (NRCMs)\u003c/p\u003e\n\u003cp\u003eNRCMs were isolated from 1-day-old Sprague-Dawley rats as previously described\u003csup\u003e52,53\u003c/sup\u003e. Hearts were excised under sterile conditions, atria removed, and ventricles minced into small fragments in ice-cold CBFHH buffer (NaCl 137 mM, KCl 5.4 mM, KH\u003csub\u003e2\u003c/sub\u003ePO\u003csub\u003e4\u003c/sub\u003e 0.44 mM, HEPES 10 mM, Na\u003csub\u003e2\u003c/sub\u003eHPO\u003csub\u003e4\u003c/sub\u003e 0.17 mM, D-glucose 3 mM, MgSO\u003csub\u003e4\u003c/sub\u003e\u0026middot;7H\u003csub\u003e2\u003c/sub\u003eO 0.4 mM, pH 7.4). Tissue was digested at 37 \u0026deg;C with type II collagenase (0.45 mg\u0026middot;ml⁻\u0026sup1;, 2 \u0026times; 10 min; C6885, Sigma-Aldrich) and subsequent trypsin digestion (0.125%, five min per cycle; G4022, Servicebio) until completely dispersed. Supernatants from all digestions were pooled, neutralized using low-glucose DMEM (L170KJ, BasalMedia) supplemented with 20% FBS (A5256701, Gibco), filtered through a 70 \u0026micro;m cell strainer (CLS431751, Sigma-Aldrich), and centrifuged at 800 rpm for five min. Cell pellets were resuspended in low-glucose DMEM with 20% FBS and pre-plated for one hour to minimize fibroblast contamination via differential adhesion. Non-adherent NRCMs were collected and seeded onto laminin-coated (10 \u0026micro;g/mL; CC095, Sigma-Aldrich) plates. Cells were maintained in low-glucose DMEM supplemented with 20% FBS, 100 U/mL penicillin, and 100 \u0026micro;g/mL streptomycin (Invitrogen) at 37 \u0026deg;C in a humidified atmosphere of 5% CO\u003csub\u003e2\u003c/sub\u003e/95% O\u003csub\u003e2\u003c/sub\u003e. Culture medium was replaced 24 h post-plating before experimental use.\u003c/p\u003e\n\u003cp\u003eNRCMs were treated with NAA (final concentration: 0.5 mM) prepared in phosphate buffer (pH 7.2\u0026ndash;7.4). Control cells received an equal volume of phosphate buffer.\u003c/p\u003e\n\u003cp\u003eAAV-mediated SIRT2 overexpression in NRCMs\u003c/p\u003e\n\u003cp\u003eRecombinant AAV9 harboring the rat \u003cem\u003eSirtuin2\u003c/em\u003e-coding gene was constructed in the GV345 vector, which contains the CMV-MCS-3FLAG-SV40-Cherry expression cassette. The construct was cloned using the following primers: forward 5\u0026prime;-AGGTCGACTCTAGAGGATCCCGCCACCATGGACTTCCTACGGAATTTATTC-3\u0026prime; and reverse 5\u0026prime;-TCCTTGTAGTCCATACCGGTGTGTTCCTCTTTCTCTTTGGTCC-3\u0026prime;, and packaged by GeneChem (GOSA5022468) at a titer of 2.5 \u0026times; 10\u003csup\u003e10\u003c/sup\u003e PFU/mL. NRCMs were infected at a multiplicity of infection (MOI) of 50 in serum-free medium for 2 h. Complete medium was added after 2 h, and the medium was replaced 12 h later. Cells were cultured for 36\u0026ndash;48 h before downstream assays and analyses.\u003c/p\u003e\n\u003cp\u003eMetabolite quantification\u003c/p\u003e\n\u003cp\u003eMetabolites quantification was performed by Metabo-Profile Biotechnology (Shanghai, China) using UPLC-MS/MS\u003csup\u003e54\u003c/sup\u003e. Samples were thawed on ice to minimize degradation before extraction. For mice serum, 50 \u0026mu;L of sample was mixed with 250 \u0026mu;L of precipitation reagent containing internal standards, followed by vortexing at 1,200 rpm for 20 min at 10 \u0026deg;C. After centrifugation (18,000 \u0026times; \u003cem\u003eg\u003c/em\u003e, 20 min, 4 \u0026deg;C), 100 \u0026mu;L of the supernatant was transferred into a 96-well plate for UPLC-MS/MS analysis. For mice heart and kidney tissues, ~10 mg of each sample was homogenized in 50 \u0026mu;L of deionized water with 10 zirconium oxide beads for 3 min, followed by the addition of 250 \u0026mu;L precipitation reagent (with internal standards). Samples were further homogenized (speed 8, 3 min) and centrifuged at 18,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 20 min at 4 \u0026deg;C. Then, 100 \u0026mu;L of supernatant was transferred into a 96-well plate for UPLC-MS/MS analysis. For cells, 120 \u0026mu;L of methanol was added to each tube, and metabolites were extracted by ultrasonic disruption. After centrifugation at 18,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 20 min at 4 \u0026deg;C, 30 \u0026mu;L of the supernatant was transferred into a 96-well plate for automated derivatization on a Biomek 4000 workstation (Beckman Coulter, Brea, USA). Briefly, 20 \u0026mu;L of freshly prepared derivatization reagent was added to each well, and the plate was shaken at 1,450 rpm for 60 min at 30 \u0026deg;C. Following derivatization, 330 \u0026mu;L of 50% methanol was added, and the plate was mixed at 650 rpm for 5 min at 30 \u0026deg;C, then centrifuged at 4,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 30 min at 4 \u0026deg;C. The supernatants were sealed in 96-well plates for\u0026nbsp;UPLC-MS/MS analysis. Chromatographic separation was achieved on a BEH Amide column (Waters) coupled to a triple quadrupole tandem mass spectrometer (ACQUITY UPLC\u0026ndash;Xevo TQ-S, Waters Corp., Milford, MA, USA) operating in both positive and negative multiple reaction monitoring modes. Authentic and isotopically labeled standards were used for calibration, and pooled QC samples were injected periodically to monitor analytical stability. Data were processed with vendor software, and metabolite concentrations were determined from standard curves. Rigorous quality control/assurance procedures were implemented throughout to ensure high-quality analytical results.\u003c/p\u003e\n\u003cp\u003eIdentification of NAA-binding proteins\u003c/p\u003e\n\u003cp\u003eNAA-binding proteins were identified as previously described\u003csup\u003e55,56\u003c/sup\u003e. Heart tissues were harvested from adult mice and immediately washed with ice-cold phosphate buffer. Apex tissues (50 mg) were minced and lysed in 500 \u0026mu;L ice-cold 0.5% NP-40 buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% NP-40, 1 \u0026mu;g/mL aprotinin, 1 \u0026mu;g/mL leupeptin, 1 \u0026mu;g/mL pepstatin and 1 mM PMSF) using a tissue grinder. Lysates were clarified by centrifugation at 15,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 20 min at 4 \u0026deg;C. The supernatants were incubated with 100 \u0026mu;M synthesized NAA probes (1132432, AppTec) or NAA at 4 \u0026deg;C for 2 h, followed by exposure to 365 nm UV light (UCL-3200L, LUYOR) for 10 min to crosslink NAA-interacting proteins. Subsequently, the reaction mixtures were adjusted to final concentrations of 1 mM TCEP (C4706, Sigma-Aldrich), 0.1 mM TBTA (678937, Sigma-Aldrich), 1 mM CuSO4 (C1297, Sigma-Aldrich), and 1 mM Biotin-azide (HY-129832, MedChemExpress), and incubated at 4 \u0026deg;C for 1 h for click chemistry. Protein aggregates were removed by centrifugation at 20,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 15 min, and the supernatants were incubated with streptavidin magnetic beads (HY-K0208, MedChemExpress) for 2 h with gentle rotation at 4 \u0026deg;C. Beads were washed three times with 700 \u0026micro;L 0.5% NP-40 buffer and then resuspended in 100 \u0026micro;L SDS-PAGE loading buffer. Samples were resolved by SDS-PAGE, stained with Coomassie Blue Fast Staining Solution (P0017, Beyotime), and protein bands were excised for mass spectrometry analysis.\u003c/p\u003e\n\u003cp\u003eProtein expression and purification\u003c/p\u003e\n\u003cp\u003eThe coding sequences of SIRT1\u0026ndash;7, MDH1, MDH2, and nicotinamidase were subcloned into a modified pGEX-6p-1 vector carrying an N-terminal 6 \u0026times; His tag. Plasmids were transformed into \u003cem\u003eE. coli\u003c/em\u003e BL21 (DE3), and protein expression was induced with 0.5 mM IPTG when the OD\u003csub\u003e600\u003c/sub\u003e reached 0.6\u0026ndash;0.8, followed by incubation at 4 \u0026deg;C overnight. Bacterial pellets were lysed by sonication in lysis buffer (50 mM Tris-HCl, pH 7.5, 300 mM NaCl, 10 mM imidazole, 1 mM DTT, protease inhibitors), and lysates were clarified by centrifugation at 15,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 30 min at 4 \u0026deg;C. Recombinant proteins were purified using Ni-NTA Beads 6FF (N30210, LABLEAD), eluted with 250 mM imidazole, and purity was verified by SDS-PAGE. Purified proteins were concentrated, flash-frozen in liquid nitrogen, and stored at -80 \u0026deg;C until use.\u003c/p\u003e\n\u003cp\u003eThe following primers were used: hSIRT1: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGGCGGACGAGGCGGCC-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGTGATTTGTTTGATGGATAGTTCATGT-3\u0026prime;. hSIRT2: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGGCAGAGCCAGACCCC-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGCTGGGGTTTCTCCCTCTCTGTT-3\u0026prime;. hSIRT3: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGGCGTTCTGGGGTTGG-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGTTTGTCTGGTCCATCAAGCTTCC-3\u0026prime;. hSIRT4: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGAAGATGAGCTTTGCGTTGA-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGGCATGGGTCTATCAAAGGCAGC-3\u0026prime;. hSIRT5: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGCGACCTCTCCAGATTGTCC-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGAGAAACAGTTTCATTTTCATGACAGG-3\u0026prime;. hSIRT6: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGTCGGTGAATTACGCGGC-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGGCTGGGGACCGCCTTGGC-3\u0026prime;. hSIRT7: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGGCAGCCGGGGGTCTG-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGCGTCACTTTCTTCCTTTTTGTGC-3\u0026prime;. mMDH1: forward 5\u0026prime;-CCCCTGGGATCCCCGGAATTCATGTCTGAACCAATCAGAGTCCTTG-3\u0026prime;, reverse 5\u0026prime;-GTCACGATGCGGCCGCTCGAGCGCAGAGGAGAGAAACTCAAAAG-3\u0026prime;. mMDH2: forward 5\u0026prime;-TAGTCCAGTGTGGTGGAATTCATGCTGTCCGCTCTCGCC-3\u0026prime;, reverse 5\u0026prime;-AACGGGCCCTCTAGACTCGAGCTTCATGTTCTTGACAAAGTCCTCG-3\u0026prime;. Site-directed mutagenesis of mMDH1 was performed to generate alanine substitution mutants (R92A/R98A/N131A/R162A/S242A) using the following primer pairs: R92A: forward 5\u0026prime;-GTCCTAGTGGGCTCCATGCCAGCTAGGGAAGGCATGGAGAGGAAG-3\u0026prime;, reverse 5\u0026prime;-CTTCCTCTCCATGCCTTCCCTAGCTGGCATGGAGCCCACTAGGAC-3\u0026prime;. R98A: forward 5\u0026prime;-GAAGGGAAGGCATGGAGGCAAAGGACCTACTGAAAG-3\u0026prime;, reverse 5\u0026prime;-CTTTCAGTAGGTCCTTTGCCTCCATGCCTTCCCTTC-3\u0026prime;. N131A: forward 5\u0026prime;-GTCATTGTTGTGGGAGCCCCAGCCAATACGAAC-3\u0026prime;, reverse 5\u0026prime;-GTTCGTATTGGCTGGGGCTCCCACAACAATGAC-3\u0026prime;. R162A: forward 5\u0026prime;-CTCGCTTGGACCACAACGCAGCAAAATCTCAAATTG-3\u0026prime;, reverse 5\u0026prime;-CAATTTGAGATTTTGCTGCGTTGTGGTCCAAGCGAG-3\u0026prime;. S242A: forward 5\u0026prime;-CTCGGAAGCTGTCCGCTGCAATGTCTGCTG-3\u0026prime;, reverse 5\u0026prime;-CAGCAGACATTGCAGCGGACAGCTTCCGAG-3\u0026prime;. Nicotinamidase: forward 5\u0026prime;- CCCCTGGGATCCCCGGAATTCATGCCCCCTCGCGCCCTG-3\u0026prime;, reverse 5\u0026prime;- GTCACGATGCGGCCGCTCGAGCCCCTGTGTCTCTTCCCAGTC-3\u0026prime;.\u003c/p\u003e\n\u003cp\u003eMDH1 enzymatic activity assay\u003c/p\u003e\n\u003cp\u003eMDH1 enzymatic activity was determined by monitoring NADH consumption at 340 nm. For recombinant assays, purified MDH1 (0.1 mg/mL) was incubated in 150 \u0026micro;L of reaction buffer: 50 mM Tris-HCl pH 7.4, 6 mM MgCl\u003csub\u003e2\u003c/sub\u003e, 1 mM DTT, 0.1 mM NADH (HY-F0001R, MedChemExpress), and initiated with freshly prepared 100 \u0026micro;M oxaloacetate (O4126, Sigma-Aldrich). The reactions were continuously analysed in a Multimode Microplate Reader (Spark, Tecan) at 25 \u0026deg;C, and the NADH consumption was measured every minute for 15 min. For tissue assays, MDH1 was immunoprecipitated with an anti-MDH1 antibody from freshly isolated mouse heart apex and assayed for enzymatic activity under the same reaction conditions. Enzymatic activity was calculated from the rate of NADH consumption, using the molar extinction coefficient of NADH (\u0026epsilon; = 6.22 mM⁻\u0026sup1;\u0026middot;cm⁻\u0026sup1;) to convert absorbance changes at 340 nm into reaction rates.\u003c/p\u003e\n\u003cp\u003eSIRT2 enzymatic activity assay\u003c/p\u003e\n\u003cp\u003eSIRT2-mediated NAA hydrolysis was evaluated using complementary enzyme-coupled and LC-MS/MS-based assays.\u003c/p\u003e\n\u003cp\u003eEnzyme-coupled assays were conducted as previously described\u003csup\u003e27,57\u003c/sup\u003e. The following reagents were used in the assay mixtures: 1 mM NAA (00920, Sigma-Aldrich), 1 mM NAD\u003csup\u003e+\u003c/sup\u003e (HY-B0445, MedChemExpress), 1 mM DTT, 6 mM MgCl\u003csub\u003e2\u003c/sub\u003e, 0.1 mg/mL nicotinamidase, 2.5 mM \u0026alpha;-ketoglutarate, 0.2\u0026nbsp;mM NADPH (HY-113324, MedChemExpress) and 0.175 units of glutamate dehydrogenase (G2626, Sigma-Aldrich), 20\u0026nbsp;mM phosphate buffer pH 7.4. All assay components except SIRT2 were preincubated at 25 \u0026deg;C for 5 min, and 2.5 mg/mL purified SIRT2 was added to start the reactions. The final volume was 100 \u0026micro;L. NADPH was quantified by its intrinsic fluorescence with excitation at approximately 340 nm and emission at 460 nm in a solid black, flat-bottomed, 96-well plate. The reactions were continuously analysed in a Multimode Microplate Reader (Spark, Tecan) at 25 \u0026deg;C, and the NADPH consumption was measured every 5 min for 2 h. The raw data from three replicates were fitted to Akima splines using GraphPad Prism (v. 10.0, Dotmatics) and kinetic parameters (\u003cem\u003eK\u003csub\u003em\u003c/sub\u003e\u003c/em\u003e and V\u003csub\u003emax\u003c/sub\u003e) by varying NAA concentrations from 0.1 to 5 mM.\u003c/p\u003e\n\u003cp\u003eFor LC-MS/MS analysis, reactions containing 1 mM NAA, 1 mM NAD\u003csup\u003e+\u003c/sup\u003e, 1 mM DTT, 6 mM MgCl\u003csub\u003e2\u0026nbsp;\u003c/sub\u003eand SIRT2 (2.5 mg/mL) in 50 mM HEPES (pH 7.4) were incubated at 37 \u0026deg;C for 2 h. Reactions were quenched with ice-cold methanol (3 \u0026times; reaction volume) and centrifuged at 15,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 10 min at 4 \u0026deg;C. Supernatants were analysed on a Waters H-Class UPLC equipped with a Welch Ultimate AQ-C18 column (2.1 \u0026times; 250 mm, 5 \u0026mu;m) using a 0.2 mL/min flow rate and a water\u0026ndash;acetonitrile gradient (0.1% formic acid; 0-1 min, 0% B; 1-8 min, 0\u0026rarr;90% B; 8-10 min, 90% B; 10-10.1 min, 90\u0026rarr;0% B; 10.1-15 min, 0% B). Detection was performed on a Sciex API 6500 QTRAP in positive electrospray ionization mode using multiple reaction monitoring (MRM), with NAA and L-aspartate unambiguously distinguished based on their specific parent-to-fragment ion transitions and chromatographic retention times. MRM transitions were optimized with authentic standards. Peak areas were extracted in Skyline for relative quantification and plotted using GraphPad Prism (v. 10.0, Dotmatics).\u003c/p\u003e\n\u003cp\u003eDrug Affinity Responsive Target Stability (DARTS)\u003c/p\u003e\n\u003cp\u003ePurified protein was diluted to 1 mg/mL with TNM buffer (50 mM Tris-HCl, 50 mM NaCl, 6 mM MgCl\u003csub\u003e2\u003c/sub\u003e, pH 7.4) and incubated with 1 mM NAA or 1 mM L-aspartate for 1 h at 25 \u0026deg;C. Pronase (10165921001, Sigma-Aldrich) was added at varying mass ratios relative to the protein, and reactions were incubated for 30 min at 25 \u0026deg;C. Reactions were terminated by addition of SDS-PAGE loading buffer. Samples were resolved by SDS-PAGE, stained with Coomassie Blue Fast Staining Solution (P0017, Beyotime), and imaged using a Typhoon FLA 9500 scanner (GE Healthcare).\u003c/p\u003e\n\u003cp\u003eSurface-Plasmon Resonance (SPR)\u003c/p\u003e\n\u003cp\u003eBinding kinetics and affinity of NAA to target proteins were measured on a Biacore T200 at 25 \u0026deg;C using CM5 sensor chips, with data analyzed via Biacore T200 Evaluation software (v. 2.0GE Healthcare). Purified proteins were covalently immobilized on the CM5 sensor chip via amine groups in 10 mM sodium acetate buffer (pH 5.0). Carboxyl groups on the sensor surface were activated by injection of 0.2 M N-ethyl-N\u0026prime;-(3-dimethylaminopropyl) carbodiimide (EDC) and 0.05 M N-hydroxysuccinimide (NHS) at 10 \u0026micro;L/min. Proteins in 10 mM sodium acetate buffer (pH 5.0) were injected at 10 \u0026micro;L/min to couple to the sensor surface, and remaining active sites were blocked with 1M ethanolamine (EA). A reference flow cell was activated and blocked in the absence of protein. For direct binding experiments, immobilization levels were adjusted to 10,000 response units (RU), and NAA solutions containing 5% DMSO were serially injected from low to high concentrations at 30 \u0026micro;L/min for 150 s. Association and dissociation rates were determined by fitting the data to a 1:1 Langmuir binding model using Biacore T200 Evaluation software.\u003c/p\u003e\n\u003cp\u003eMolecular docking\u003c/p\u003e\n\u003cp\u003eMDH1 or SIRT2 structures were obtained from Protein Data Bank (PDB IDs: 7RM9 and 4RMG, respectively) and prepared in MOE at pH 7.4 (AMBER10: EHT). Binding pockets were identified via structural analysis or MOE Site Finder. Molecular docking was performed with the triangle matcher algorithm, initial poses were scored using London \u0026Delta;G, and the top 10-20 poses per pocket were refined via the induced fit algorithm with GBVI/WSA \u0026Delta;G scoring, and the pose with the best docking score was used for visualization in the manuscript. SIRT2-TUBA1A complexes were predicted using AlphaFold-Multimer, and resulting models were inspected to identify plausible interaction interfaces.\u003c/p\u003e\n\u003cp\u003eMolecular dynamics simulations\u003c/p\u003e\n\u003cp\u003eMolecular dynamics simulations of protein\u0026ndash;ligand complexes were conducted with Desmond 2020. Docking-derived best-scoring poses were placed in a TIP3P water box and neutralized with Na\u003csup\u003e+\u003c/sup\u003e/Cl\u003csup\u003e-\u003c/sup\u003e ions. Systems were parameterized with the OPLS3e force field. Energy minimization was carried out with SHAKE constraints on bond lengths and angles of heavy atoms and water geometry. Periodic boundary conditions and the particle mesh Ewald method were applied. Production MD simulations were run for 100 ns in the NPT ensemble at 300 K and 1 atm with a 2 fs integration step, and snapshots were recorded every 10 ps. Post-MD analyses included assessment of structural stability (backbone RMSD) and characterization of protein-ligand interactions, including interacting residues, interaction types, interaction frequencies, and estimated binding free energies.\u003c/p\u003e\n\u003cp\u003eMetabolic flux analysis\u003c/p\u003e\n\u003cp\u003eNRCMs were seeded in 6-well plates and cultured to ~80% confluency. Cells were washed twice with sterile PBS and incubated with isotope-labeled tracing medium. For [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e6\u003c/sub\u003e]-D-glucose tracing, cells were cultured in glucose-free DMEM (D5030, Sigma-Aldrich; with 0.584 g/L glutamine) supplemented with 20% dialyzed FBS (04-011-1A, Biological Industries) and 10 mM [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e6\u003c/sub\u003e]-D-glucose (HY-B0389A, MedChemExpress) for 6 h to assess glycolytic and TCA cycle fluxes\u003csup\u003e58,59\u003c/sup\u003e. For [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e5\u003c/sub\u003e]-L-glutamine tracing, cells were cultured in glutamine-free DMEM (D5030, Sigma-Aldrich; with 1.0 g/L D-Glucose) supplemented with 20% dialyzed FBS (04-011-1A, Biological Industries) and 2 mM [U-\u003csup\u003e13\u003c/sup\u003eC\u003csub\u003e5\u003c/sub\u003e]-L-glutamine (HY-N0390S1, MedChemExpress) for 24 h to assess TCA cycle fluxes\u003csup\u003e60\u003c/sup\u003e. Metabolites were subjected to UPLC-MS/MS-based metabolic flux analysis (Metabo-Profile Biotechnology, China) using an ultrahigh-pressure liquid chromatography-triple quadrupole mass spectrometer (ACQUITY UPLC-Xevo TQ-S, Waters Corp., Milford, MA, USA)\u003csup\u003e61\u003c/sup\u003e. For data processing, the raw data files generated by UPLC-MS/MS were processed using MassLynx software (v 4.1, Waters Corp., Milford, MA, USA）for peak extraction, integration, identification, and quantification of individual metabolites.\u003c/p\u003e\n\u003cp\u003eNAD\u003csup\u003e+\u003c/sup\u003e/NADH ratio determination\u003c/p\u003e\n\u003cp\u003eCellular NAD\u003csup\u003e+\u003c/sup\u003e/NADH ratios were determined using two complementary approaches. For live-cell measurements, cells were seeded evenly in black, opaque 96-well plates (~2 \u0026times; 10\u003csup\u003e4\u003c/sup\u003e cells per well) and transduced with adenovirus expressing cyto-Sonar (ADSON1001-S) or mito-Sonar (ADSON1003-S, Provoson) following the manufacturer\u0026rsquo;s instructions\u003csup\u003e62,63\u003c/sup\u003e. Fluorescence was measured using a Multimode Microplate Reader (Spark, Tecan) at 420/528 nm (NADH) and 485/528 nm (NAD\u003csup\u003e+\u003c/sup\u003e). The NAD\u003csup\u003e+\u003c/sup\u003e/NADH ratio was determined ratiometrically as R\u003csub\u003e485/420\u003c/sub\u003e, by dividing the fluorescence intensity at 485 nm by that at 420 nm. For biochemical measurements, cells were lysed and processed according to the NAD\u003csup\u003e+\u003c/sup\u003e/NADH Assay Kit protocol (S0176S, Beyotime). Lysates were subjected to enzymatic cycling reactions, and absorbance at 450 nm was measured using a microplate reader. NAD\u003csup\u003e+\u003c/sup\u003e/NADH ratios were calculated and normalized to total protein.\u003c/p\u003e\n\u003cp\u003eATP quantification\u003c/p\u003e\n\u003cp\u003eCellular ATP levels were measured using the ATP Assay Kit (S002, Beyotime) according to the manufacturer\u0026rsquo;s instructions. Briefly, cells were washed with PBS and lysed using the provided lysis buffer. After centrifugation at 12,000 \u0026times; \u003cem\u003eg\u003c/em\u003e for 5 min at 4 \u0026deg;C, the supernatant was collected. An aliquot of the lysate was mixed with the reaction solution and incubated for 5 min at room temperature in the dark. The luminescence was measured using a Multimode Microplate Reader (Spark, Tecan). ATP concentrations were calculated based on a standard curve generated with ATP standard solutions provided in the kit, and results were normalized to protein concentration determined by BCA assay (P0009, Beyotime).\u003c/p\u003e\n\u003cp\u003eWestern blotting\u003c/p\u003e\n\u003cp\u003eFor cell samples, proteins were extracted directly using 1\u0026times; SDS loading buffer. Tissue samples were homogenized in 0.5% NP-40 lysis buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% NP-40, 1 \u0026mu;g/mL aprotinin, 1 \u0026mu;g/mL leupeptin, 1 \u0026mu;g/mL pepstatin and 1 mM PMSF). The lysates were centrifuged at 4 \u0026deg;C for 20\u0026thinsp;min at 15,000 \u0026times; \u003cem\u003eg\u003c/em\u003e, and the supernatants were used as whole-cell extracts. Protein samples were subjected to western blotting according to standard procedures. Detection was performed by measuring chemiluminescence using an ECL Plus Western Blotting Detection System on a Typhoon FLA 9500 (both GE Healthcare). The following antibodies were used for western blot analysis: anti-\u0026beta;-MHC (1:1000, 22280-1-AP, Proteintech), anti-ANP (1:1000, A14755, ABclonal), anti-BNP (1:1000, A23996, ABclonal), anti-SIRT2 (1:1000, A12575, ABclonal), anti-ACY1 (1:1000, A6351, ABclonal), anti-ACY2 (1:1000, A7271, ABclonal), anti-NAT8L (1:1000, 23841-1-AP, Proteintech), anti-SLC13A3 (1:1000, 26184-1-AP, Proteintech), anti-GAPDH (1:3000, A19056, ABclonal), anti-\u0026alpha;-Tubulin (1:3000, A6830, ABclonal), goat anti-mouse IgG antibody (1:5000, A21001S, Abmart), goat anti-rabbit IgG antibody (1:5000, M21002L, Abmart).\u003c/p\u003e\n\u003cp\u003ePhalloidin staining\u003c/p\u003e\n\u003cp\u003eNRCMs were cultured on glass coverslips in 12-well plates and fixed with 4% paraformaldehyde for 10 min at room temperature. After washing three times with PBS, the cells were permeabilized with 0.1% Triton X-100 in PBS for 5 min. Following PBS washes, the samples were incubated with Phalloidin-iFluor\u0026trade; 488 (BL1190A, Biosharp) diluted 1:1000 in PBS for 30 min at room temperature in the dark. Nuclei were counterstained with DAPI (D9542, Sigma-Aldrich). Coverslips were mounted with antifade mounting medium (G1226-7, ServiceBio) and imaged using a Leica fluorescence microscope. The size of individual cells was quantified by ImageJ software (NIH).\u003c/p\u003e\n\u003cp\u003eStatistical analysis\u003c/p\u003e\n\u003cp\u003eAll statistical analyses were performed using GraphPad Prism (v. 10.0, Dotmatics). For all experiments, data are presented as the mean \u0026plusmn; standard error of the mean (SEM) unless otherwise specified. All experiments (except those described otherwise in the legend) were performed independently at least three times with a similar outcome. The statistical tests, n values, and the \u003cem\u003eP\u003c/em\u003e values are all indicated in the figures and/or legends. Depending on the type of experiment, the \u003cem\u003eP\u003c/em\u003e values were calculated using different analytical methods. Two-group comparisons were performed using two-tailed Student\u0026rsquo;s \u003cem\u003et\u003c/em\u003e-tests. For single-variable comparisons among three or more groups, one-way analysis of variance (ANOVA) followed by Tukey\u0026rsquo;s multiple comparison test was applied. For experiments involving two independent variables, two-way ANOVA with Tukey\u0026rsquo;s or Bonferroni\u0026rsquo;s post hoc test\u0026nbsp;was used to obtain adjusted \u003cem\u003eP\u003c/em\u003e values. \u003cem\u003eP\u003c/em\u003e \u0026lt; 0.05 was considered significant (*\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.05, **\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.01, ***\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.001, ****\u003cem\u003eP\u003c/em\u003e \u0026lt; 0.0001).\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003eAcknowledgements\u003c/p\u003e\n\u003cp\u003eThis work was supported by the Grants from National Natural Science Foundation of China (82330048, 82501473, 32370824, 82300428, 82101750), Program of Shanghai Academic Research Leader (21XD1421700), Innovation Program of the Shanghai Municipal Education Commission (2023ZKZD24), China National Postdoctoral Program for Innovative Talents fellowship (BX20250188), Innovative research team of high-level local universities in Shanghai (SHSMU-ZDCX20211100), the Science and Technology Commission of Shanghai Municipality (25ZR1402362, 25ZR1402037), Shanghai Sailing Program (23YF1425500), and The Construction Project of the\u0026nbsp;\u0026ldquo;Discipline Peak-Climbing Plan\u0026rdquo;\u0026nbsp;of Xinhua Hospital Affiliated to Shanghai Jiao Tong University School of Medicine (XKPF2024A5001).\u003c/p\u003e\n\u003cp\u003eAuthor contributions\u003c/p\u003e\n\u003cp\u003eJ.-Y.Z. and Z.-Y.Z. conceived the project. Z.-Y.Z. and J.-Y.Z. wrote the manuscript. J.-Y.Z. and H.T. supervised the study. Z.-Y.Z., K.C., Y.Y., and F.-J.L. performed human samples studies. Z.-Y.Z., X.-J.Q., J.-Y.Y., and H.T. performed animal experiments and prepared figures. Z.-Y.Z., J.-Y.Y., X.S., Y.S., and R.Z. performed biochemistry experiments. Z.-Y.Z., J.-Y.Z., F.L., H.H., and K.S. carried out analyses and interpreted the results. All authors reviewed the results and approved the final version of the manuscript.\u003c/p\u003e\n\u003cp\u003eCompeting interests\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u003c/p\u003e\n\u003cp\u003eData availability\u003c/p\u003e\n\u003cp\u003eThe datasets generated and analysed during the current study are available from the corresponding author upon reasonable request. All data supporting the findings of this study will be deposited in a public repository prior to publication, and accession codes will be provided at that time.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n\u003cli\u003eTanaka, H., Sirich, T. L., Plummer, N. S., Weaver, D. S. \u0026amp; Meyer, T. W. An Enlarged Profile of Uremic Solutes. \u003cem\u003ePLoS One\u003c/em\u003e \u003cstrong\u003e10\u003c/strong\u003e, e0135657 (2015). https://doi.org:10.1371/journal.pone.0135657\u003c/li\u003e\n\u003cli\u003ePatel, N., Yaqoob, M. M. \u0026amp; Aksentijevic, D. Cardiac metabolic remodelling in chronic kidney disease. \u003cem\u003eNat Rev Nephrol\u003c/em\u003e \u003cstrong\u003e18\u003c/strong\u003e, 524-537 (2022). https://doi.org:10.1038/s41581-022-00576-x\u003c/li\u003e\n\u003cli\u003eLindner, H. A., Tafler-Naumann, M. \u0026amp; Rohm, K. H. 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Here, targeted metabolomics in patients with CKD identified that circulating N-acetylaspartate (NAA) as the metabolite most closely linked to early cardiac dysfunction. In mice, both CKD and exogenous NAA administration induced pronounced cardiac NAA accumulation, resulting in systolic dysfunction and pathological hypertrophy. Activity-based protein profiling identified cytosolic malate dehydrogenase 1 (MDH1) and sirtuin 2 (SIRT2) as direct targets of NAA. NAA acts as a substrate analogue that occupies the MDH1 substrate-binding pocket, competitively inhibits its enzymatic activity, and disrupts the malate–aspartate shuttle, thereby lowering cytosolic and mitochondrial NAD⁺/NADH ratios, suppressing tricarboxylic acid cycle flux, and compromising cardiomyocyte energy metabolism. Notably, we identified SIRT2 as a previously unrecognized NAA hydrolase that specifically binds and hydrolyzes NAA in an NAD⁺-dependent manner, thereby mitigating NAA-induced metabolic stress. Cardiomyocyte-specific restoration of SIRT2 activity in mice reduced cardiac NAA levels, improved systolic dysfunction, and attenuated hypertrophy. These findings expand sirtuins from protein deacylases to direct regulators of small-molecule metabolites and establish NAA as a kidney–heart metabolic mediator, revealing an amino acid acetylation-dependent layer of energy homeostasis.","manuscriptTitle":"SIRT2 functions as an N-acetylaspartate hydrolase that counteracts cardiac metabolic remodeling in kidney disease","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-01-13 10:31:47","doi":"10.21203/rs.3.rs-8510072/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"165ffcd1-bc2d-4744-ac4d-185628084eef","owner":[],"postedDate":"January 13th, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[{"id":60887520,"name":"Biological sciences/Biochemistry/Enzyme mechanisms"},{"id":60887521,"name":"Health sciences/Diseases/Cardiovascular diseases"}],"tags":[],"updatedAt":"2026-01-15T21:12:00+00:00","versionOfRecord":[],"versionCreatedAt":"2026-01-13 10:31:47","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8510072","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8510072","identity":"rs-8510072","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}