Methionine-SAM metabolism-dependent ubiquinone synthesis is crucial for ROS accumulation in ferroptosis induction | Nature Communications

Nature Communications volume 15, Article number: 8971 (2024) Cite this article

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Ferroptosis is a cell death modality in which iron-dependent lipid peroxides accumulate on cell membranes. Cysteine, a limiting substrate for the glutathione system that neutralizes lipid peroxidation and prevents ferroptosis, can be converted by cystine reduction or synthesized from methionine. However, accumulating evidence shows methionine-based cysteine synthesis fails to effectively rescue intracellular cysteine levels upon cystine deprivation and is unable to inhibit ferroptosis. Here, we report that methionine-based cysteine synthesis is tissue-specific. Unexpectedly, we find that rather than inhibiting ferroptosis, methionine in fact plays an essential role during cystine deprivation-induced ferroptosis. Methionine-derived S-adenosylmethionine (SAM) contributes to methylation-dependent ubiquinone synthesis, which leads to lipid peroxides accumulation and subsequent ferroptosis. Moreover, SAM supplementation synergizes with Imidazole Ketone Erastin in a tumor growth suppression mouse model. Inhibiting the enzyme that converts methionine to SAM protects heart tissue from Doxorubicin-induced and ferroptosis-driven cardiomyopathy. This study broadens our understanding about the intersection of amino acid metabolism and ferroptosis regulation, providing insight into the underlying mechanisms and suggesting the methionine-SAM axis is a promising therapeutic strategy to treat ferroptosis-related diseases.

Ferroptosis is a newly identified regulated cell death caused by iron-dependent lipid peroxidation, and is morphologically, biochemically, and genetically distinct from other forms of cell death including apoptosis, necrosis and autophagy1. Although the physiological function of ferroptosis is still unclear, its involvement in several pathological conditions has been established. Ferroptosis is a major cause for cell death associated with ischemic organ injury, such as ischemic cardiomyopathy and kidney failure2,3. Emerging evidence has suggested that ferroptosis is found in the neuronal cell death associated neurological diseases including Alzheimer’s disease4,5, Parkinson’s disease6, and Huntington’s disease7. Moreover, ferroptosis induction hold great potential for tumor suppression in clinical treatment8,9.

Lipid peroxidation is the direct cause of ferroptosis. To combat lipid peroxidation, glutathione peroxidase 4 (GPX4), a phospholipid hydroperoxidase, reduces phospholipid hydroperoxide into the corresponding phospholipid alcohol using glutathione (GSH) as an indispensable catalyzing co-factor10,11. GPX4/GSH has been known as the primary ferroptosis prevention system that can efficiently reduce lipid peroxides accumulated on cell membranes. Growing evidence revealed that many types of cancer cells are sensitive to ferroptosis induced by inactivation of GPX4 or depletion of GSH10,12. GSH is a tripeptide composed of cysteine, glutamate and glycine, and is synthesized in two steps by the ATP-dependent enzymes glutamate-cysteine ligase and glutathione synthase13. For generation of GSH, sulfur-containing cysteine is the rate-limiting precursor14. Therefore, ferroptosis is a very unique type of cell death because it is not only regulated by lipid metabolism and iron metabolism, but also by amino acid metabolism15.

Intracellular cysteine is mainly converted from cystine reduction, that is transported into the cell through the solute carrier family 7 member 11 (SLC7A11, also known as xCT)16. xCT is an amino acid antiporter that mediates the transmembrane exchange of extracellular cystine and intracellular glutamate across the cell membrane17,18. Besides being converted from the exogenously obtained cystine, cysteine may be converted from another sulfur-containing amino acid methionine. Methionine is an essential amino acid and contributes to the synthesis of the cellular cysteine via the transsulfuration pathway19,20. However, numerous studies have shown that solely depleting extracellular cystine or inhibiting cystine uptake by xCT inhibitors such as Erastin or Imidazole Ketone Erastin (IKE) is enough to result in the depletion of intracellular cysteine and GSH pools, subsequently, the inactivation of GPX4 and the accumulation of peroxidized phospholipids, thereby the induction of ferroptosis1,9,21. This implicitly suggests that methionine-based endogenous cysteine synthesis pathway fails to effectively rescue intracellular cysteine level and is unable to prevent ferroptosis. The question spontaneously arises as that whether the transsulfuration pathway is broken in the test cells used in the studies, or the producing capacity is limited.

In the present study, a tissue specificity of transsulfuration pathway is clarified. And we find that, not only having no significant inhibitory role, but reversely, methionine is essential for cystine deprivation-induced ferroptosis. Mechanistically, methionine-derived S-adenosylmethionine (SAM) is exploited for ubiquinone synthesis and is essential for ROS release, which accounts for cystine deprivation-induced lipid peroxidation. As a consequence, supplementation of SAM can synergize with IKE in tumor growth suppression; inhibition of methionine adenosyltransferase 2A (MAT2A) that converts methionine to SAM can protect the heart tissue from Doxorubicin (DOX)-induced and ferroptosis-driven cardiomyopathy.

To clarify why the methionine cannot rescue the cysteine supply to combat cystine-deprivation-induced ferroptosis, mouse embryonic fibroblast (MEF) cells and human HT1080 fibrosarcoma cells were cultured in cystine-free medium or treated with Erastin or IKE to inhibit xCT-mediated cystine transport. Lipid peroxidation and cell death were induced, however, despite the presence of methionine in medium, restriction of cystine is potent to induce ferroptosis (Fig. S1A, B) as described in numerous other studies. First, we questioned whether the methionine level in culture medium was insufficient to prevent ferroptosis. To test, MEF and HT1080 cells were incubated with methionine with increasing concentrations, that were far beyond the physiological level, and we found that no significant inhibitory effects on ferroptosis were observed (Figs. 1A, B and S2A, B). Importantly, addition of methionine to MEF and HT1080 cells culture medium failed to restore their intracellular levels of cysteine and GSH (Figs. 1C, D and S2C, D). The methionine-based cysteine synthesis embraces five steps of enzymatically catalyzed reactions, producing the intermediates SAM, S-adenosylhomocysteine (SAH), homocysteine (Hcy) and cystathionine (Cta)22,23,24 (Fig. 1E). Next, we speculated that methionine-based cysteine synthesis pathway may be defective. To test, we added exogenous methionine and its intermediate metabolites towards to the cysteine synthesis into the cell culture medium with cystine deprivation. Delivery of methionine, SAM or SAH into the cells (Fig. S2E, F) could not inhibit the upregulation of intracellular lipid peroxidation levels and cell death induced by cystine restriction (Figs. 1F, G and S2G, H). However, addition of Hcy, Cta or cysteine could (Figs. 1F, G and S2G, H). This implied that the methionine-mediated cysteine synthesis may be unimpeded only under specific condition(s).

A–D, F, G MEF cells were cultured ± cystine as indicated for 8–12 h. The levels of lipid ROS (A, F), cysteine (C) and total GSH (D) were measured respectively at the 8 h. The cell death levels (B, G) were quantified at the 10 h. E Schematic of metabolic intermediates and enzymes in methionine-based cysteine synthesis pathway. H The protein levels of the indicated metabolic enzymes in the mouse tissues were measured by western blot (WB). One representative experiment out of three is shown. I The mRNA levels of the indicated metabolic enzymes in the human tissues were analyzed basing the data from TCGA database and shown in heatmap. Blue color indicated high expression, and the red color indicated low expression. N normal tissues, T tumor tissues. J–M The cells were cultured in cystine-free medium as indicated for 8–24 h. The levels of cysteine (J), total GSH (K), and lipid ROS (L) were measured respectively (AML12 and MIHA for 24 h, MEF for 8 h, HT1080 for 10 h). The cell death levels (M) were quantified respectively (AML12 and MIHA for 30 h, MEF for 10 h, HT1080 for 14 h). CC cystine, 0.4 mM, Met methionine, 0.4 mM, SAM S-adenosylmethionine, 0.4 mM, SAH S-adenosylhomocysteine, 0.4 mM, Hcy homocysteine, 0.4 mM, Cta cystathionine, 0.4 mM, Cys cysteine, 0.4 mM, Ser serine, R methylated substrates, MAT1A methionine adenosyltransferase 1A, MAT2A methionine adenosyltransferase 2A, MAT2B methionine adenosyltransferase 2B, GAMT guanidinoacetate N-methyltransferase, GNMT glycine N-methyltransferase, NNMT nicotinamide N-methyltransferase, PEMT phosphatidylethanolamine N-methyltransferase, AHCY adenosylhomocysteinase, AHCYL1 adenosylhomocysteinase like 1, AHCYL2 adenosylhomocysteinase like 2, CBS cystathionine beta-synthase, CTH cystathionine gamma-lyase, MTR methionine synthase, MTRR methionine synthase reductase, BHMT1 betaine-homocysteine S-methyltransferase 1, BHMT2 betaine-homocysteine S-methyltransferase 2. For A–D, F, G, J–M, n = 3 biological replicates, data were represented as mean ± SD with p values determined by one-way (A–D, F, G) or two-way (J–M) ANOVA test. Source data are provided as a Source Data file.

The tissue specificity of the methionine-cysteine conversion was then investigated. Mouse tissues from 12 types of main organs were harvested and then subjected to be examined the expression of methionine-based cysteine synthesis-related metabolic enzymes. We found that metabolic enzymes of methionine-cysteine conversion were almost exclusively expressed in the liver tissue (Figs. 1E, H and S3A–J). Moreover, the Non-Human Primate Cell Atlas (NHPCA) database was mined, in which the single cell transcriptomics data showed that the expressions of methionine-cysteine conversion-related metabolic enzymes in monkey were mainly enriched in liver cells (Fig. S4A–I), which was consistent with the data of mouse liver tissue (Figs. 1H and S2, S3). At the same time, we selected 10 pairs of human organ tissues and the corresponding tumor tissues from the Cancer Genome Atlas (TCGA) database, and analyzed the mRNA levels of methionine-based cysteine synthesis-related metabolic enzymes. We found that the expression of these enzymes was much higher in normal human liver tissues and liver tumor tissues than that in other tissues (Fig. 1I). This indicated that the liver tissue had a complete methionine-initiated cysteine synthesis pathway, that is not fully weaponed in non-liver tissues. Subsequently, we found that cystine limitation mildly reduced the intracellular cysteine and GSH levels in human liver-derived MIHA cells and mouse liver-derived AML12 cells, but almost completely reduced that in MEF and HT1080 cells (Fig. 1J, K). Unlike in MEF and HT1080 cells, cystine limitation did not induce lipid peroxidation and cell death in MIHA and AML12 cells (Fig. 1L, M), probably due to the remained GSH in MIHA and AML12 cells sufficient to neutralize lipid peroxide. These suggested that methionine-initiated cysteine synthesis via the transsulfuration pathway might function in a tissue-specific manner.

Considering the tissue specificity of methionine-cysteine conversion, we expected that co-deprivation of cystine and methionine may induce ferroptosis in hepatocytes. Surprisingly, MIHA and AML12 cells did not undergo ferroptosis even though methionine and cystine were co-deprivated, which was also observed with MEF and HT1080 cells (Fig. 2A, B). Further analysis revealed that methionine was likewise required for Erastin- or IKE-induced ferroptosis (Fig. S5A–D). This suggests that the lack of methionine inhibits cystine-deprivation induced ferroptosis in general. Moreover, we examined the effect of methionine deficiency on the expression of prostaglandin-endoperoxide synthase 2 (PTGS2), which is a putative molecular marker of ferroptosis25,26. The result showed that PTGS2 gene expression was upregulated by Erastin or IKE, which was inhibited by methionine deficiency (Fig. S5E, F). In addition, we used another ferroptosis inducer RSL3, an enzymatic inhibitor of GPX4, to induce ferroptosis, and found that methionine withdrawals did not affect RSL3-induced ferroptosis (Fig. S5G–L). The combined data indicated methionine is required for ferroptosis induced by cystine deprivation or pharmacological blockage of cystine uptake.

A, B The cells were cultured as indicated for 8–24 h. A The lipid ROS levels were determined (AML12/MIHA for 24 h, MEF for 8 h, HT1080 for 10 h). B The cell death levels were quantified (AML12/MIHA for 30 h, HT1080 for 14 h). C HT1080 cells were cultured as indicated for 10 h. The metabolites levels were measured by UHPLC-HRMS. Metabolites with FC ≥ 2 and p < 0.05 were considered as hits. Blue color indicated the decreased metabolites; red color indicated the increased ones. D–J The cells were simultaneously treated as indicated for 8–30 h. D, G, I The lipid ROS levels were determined (MEF for 8 h, HT1080 for 10 h, OS-RC-2 for 14 h, HL-1 for 24 h). F The mRNA levels of PTGS2 were determined at the 8 h. E, H, J The cell death levels were determined (MEF for 12 h, HT1080 for 14 h, OS-RC-2 for 18 h, HL-1 for 30 h). K, L HT1080 was transfected with non-targeting siRNA (siControl) or targeting MAT2A siRNA for 48 h, and then cultured as indicated for another 10–14 h. K The lipid ROS levels were determined at the 10 h. L The cell death levels were quantified at the 14 h. M, N HT1080 cells were transfected without or with CHAC1 plasmid for 48 h, then cultured as indicated for another 10–14 h. M The levels of total GSH and CHAC1 proteins were determined at the 10 h. N The cell death levels were quantified at the 14 h. CC, 0.4 mM; Met, 0.4 mM; SAM, 0.4 mM; SAH, 0.4 mM; Hcy, 0.4 mM; Cta, 0.4 mM; Cys, 0.4 mM; Erastin, 2 μM; IKE, 2 μM; FIDAS-5, 5 μM. n = 3 (A–M) or n = 5 (N) biological replicates, data were represented as mean ± SD with p values determined by unpaired two-tailed t test (C), one-way (A, B, D–J, M, N) or two-way (K, L) ANOVA test. Source data are provided as a Source Data file.

Methionine might regulate cystine deprivation-induced ferroptosis through either itself or downstream metabolites. To test, we firstly performed a metabolomics analysis of HT1080 cells that were cultured in medium lacking cystine or lacking both cystine and methionine. The metabolomics results containing full profile of the metabolites was subjected to cluster assay, and the triplicates of each group were automatically clustered together (Fig. S6A), indicating better homogeneity of the data in each condition. Then, we filtrated the metabolomics data with the threshold fold change (FC) ≥ 2 and p < 0.05. The results showed that the cellular cysteine and GSH levels were expectedly depressed upon cystine limitation (Fig. S6B, C); meanwhile, cellular SAM level was significantly reduced once methionine restriction (Figs. 2C and S2C). Then, MEF cells were cultured in methionine-free media supplied with SAM, SAH, Hcy, Cta or cysteine, respectively. We found that only supply of SAM could reverse methionine restriction-inhibited ferroptosis (lipid peroxidation and cell death) that was induced by cystine deprivation (Fig. 2D, E). Likewise, only SAM rescued PTGS2 expression, that was blocked once methionine deprived (Fig. 2F). In a variety of types of cells including HT1080, human OS-RC-2, Caki-1, 786-O and ACHN renal carcinoma cells, human HT29 colorectal adenocarcinoma cells and mouse HL-1 cardiomyocytes, methionine deprivation significantly impairs ferroptosis, and methionine-derived SAM is required by ferroptosis (Figs. 2G, H and S6D–G), revealing the universality of the role of methionine metabolism in cystine restriction induced-ferroptosis. Although these results indicated that methionine may regulate ferroptosis through SAM, the possibility that methionine regulates ferroptosis by its own had not been ruled out. Methionine adenosyltransferases 2A (MAT2A), the key enzyme for methionine metabolism, catalyzes the biosynthesis of SAM from methionine and expresses in various tissues (Figs. 1H, I and S3 and 4). To further confirm that methionine functions through SAM, we sought to analyze the effect of MAT2A on ferroptosis. MAT2A inhibitor FIDAS-5 could significantly inhibit MEF, HT1080, OS-RC-2 and HL-1 cells undergoing ferroptosis that was induced by deprivation of cystine or blockage of cystine uptake in the presence of methionine (Figs. 2I, J and S7A–D). Likewise, another MAT2A inhibitor PF-936627 exhibited the similar effect as expected (Fig. S7E–H). In addition, we designed siRNA targeting MAT2A and found that knockdown of MAT2A in HT1080 cells suppressed ferroptosis induced by cystine deprivation, which was reversed by SAM compensation (Figs. 2K, L and S8A). Taken together, these results clearly demonstrated that SAM is the executor in methionine-implicated ferroptosis.

Recently, it was reported that methionine deprivation could inhibit protein synthesis of cation transport regulator homolog 1 (CHAC1), an enzyme that catalyzes the cleavage of GSH, thereby relieving cystine deprivation-caused GSH depletion and ferroptosis21. To reconcile these two studies and to clarify the contribution of each mechanism, we overexpressed CHAC1 or supplemented SAM under the conditions of both methionine and cystine deprivation. While overexpression of CHAC1 significantly diminished the restoration of GSH level caused by methionine limitation in the cells subjected to cystine-restriction, and significantly reversed methionine restriction-inhibited ferroptosis (Fig. 2M, N). So did the supply of SAM (Fig. 2N), however, which did not affect CHAC1 expression and total GSH level (Fig. 2M). Moreover, overexpression of CHAC1 along with the supply of SAM completely counteracted the inhibitory effect of methionine withdrawal on cystine restriction-induced ferroptosis (Fig. 2M, N). Taken together, the combined data suggested that the promoting effect of methionine-SAM metabolism on cystine restriction-induced ferroptosis is independent of CHAC1 expression and GSH levels.

SAM is the primary methyl donor for the methylation of nucleic acids, protein, and small molecules. Besides, SAM is also an aminopropyl groups donor for polyamine biosynthesis. Thus, it needs to further confirm SAM contributes to ferroptosis by mediating polyamine anabolism or methylation. Inactivation S-adenosyl-methionine decarboxylase, the key enzyme in polyamine biosynthesis that converts SAM to 5′-adenosyl-methylthiopropylamine, with inhibitor mitoguazone (MGBG)28 nearly did not inhibit cystine deprivation induced lipid peroxidation and cell death in MEF and HT1080 cells (Fig. 3A, B). On the contrary, the methylation inhibitor adenosine dialdehyde (ADOX)29, that generally inhibit all the SAM-dependent methyltransferases, significantly blocked the levels of lipid peroxidation and cell death induced by cystine deprivation in MEF and HT1080 cells (Fig. 3C, D). Moreover, ADOX significantly inhibited IKE-induced, or SAM supplementation-conferred lipid peroxidation and cell death (Fig. 3E–H). Although we cannot completely rule out the possibility that SAM regulating ferroptosis in a methylation-independent manner, our results collectively suggested that inhibition of methylation can significantly inhibit ferroptosis.

A, B The cells were cultured in cystine-free medium in the absence or presence of MGBG, Fer-1 or DFO as indicated for 8–14 h. A The lipid ROS levels were determined (MEF for 8 h, HT1080 for 10 h). B The cell death levels were quantified (MEF for 12 h, HT1080 for 14 h). C–H The cells were cultured in the medium ± cystine in the absence or presence of ADOX, Fer-1, DFO, IKE or FIDAS-5 for 8–14 h. C, E, G The Lipid ROS levels were determined (MEF for 8 h, HT1080 for 10 h). D, F, H The cell death levels were quantified (MEF for 12 h, HT1080 for 14 h). I–K MEF cells were cultured in the medium ± cystine or ± methionine in the absence or presence of SAM, FIDAS-5, ADOX or PBN for 8–12 h. I, J The cellular levels of ROS were respectively quantified at the 8 h. K The cell death levels were quantified at the 12 h. CC, 0.4 mM; Met, 0.4 mM; SAM, 0.4 mM; MGBG, mitoguazone, 40 μM; ADOX, adenosine dialdehyde, 20 μM; Fer-1, ferrostatin-1, 10 μM; DFO, deferoxamine, 80 μM; FIDAS-5, 5 μM; IKE, 2 μM; PBN, alpha-phenyl-tert-N-butylnitrone, 1 mM. For A–K, n = 3 biological replicates, data were represented as mean ± SD with p values determined by one-way ANOVA test. Source data are provided as a Source Data file.

Given that ferroptosis is directly caused by lipid peroxidation, we speculated that methionine-SAM axis may contribute to ferroptosis through augmenting the accumulation of reactive oxygen species (ROS) since ferroptosis was induced by suppression of cystine uptake and GSH synthesis, a context that the antioxidation system has been disabled. To this end, we detected the impact of methionine deprivation on ROS accumulation induced by cystine deprivation. The results showed that cystine deprivation-caused increase in ROS levels was significantly lowered by methionine deprivation in MEF, HT1080, OS-RC-2 and HL-1 cells (Figs. 3I and S8B–D). As expected, we found that compensation of SAM increase ROS accumulation (Figs. 3I and S8B–D). Importantly, MAT2A inhibitor or MAT2A knockdown also prevented the elevation of ROS induced by cystine deprivation (Figs. 3I and S8E–H). Moreover, we examined the effect of methylation inhibition on the rise of ROS caused by cystine deprivation. As expected, ADOX could inhibit the rise of ROS level in the presence of methionine or SAM (Fig. 3I). Then, we treated cells with ROS scavenger alpha-phenyl-tert-N-butylnitrone (PBN) and found that PBN alleviated SAM supplementation-induced ROS accumulation and ferroptosis (Fig. 3J, K). These results implied that the methylation process downstream of methionine-SAM metabolic axis is involved in ferroptosis-required ROS accumulation.

Besides ferroptosis, ROS accumulation causes various types of cell death30,31. To test the specificity of methionine-SAM mechanism in ferroptosis, we examined the effect of methionine-SAM on other forms of cell death including parthanatos, apoptosis, necroptosis and pyroptosis. Similarly, the absence of methionine moderately alleviated TNF-α/SM-164/Z-VAD-FMK (TSZ)-indued HT29 cells’ necroptosis (Fig. S9A) and LPS/Nigericin (LN)-induced human THP1 monocytic leukemia cells’ pyroptosis (Fig. S9B). In contrast, methylnitronitrosoguanidine (MNNG)-induced MEF cells’ parthanatos and TNF-α/SM-164 (TS) treatment-triggered apoptosis of HT1080 cells were enhanced by methionine restriction (Fig. S9C, D). Notably, it seems that methionine-SAM metabolism is implicated in the influence of methionine restriction on cell death to a certain extend because SAM supplementation reversed the outcomes of methionine restriction (Fig. S9A–D). The results suggested an extensive participation of methionine in regulation of cell death, yet the particular molecular mechanisms may be different and are beyond the scope of the present study, which need further investigation in the future.

ROS is produced in large quantities as a by-product from electron transfer chain, that contributes to ATP synthesis through oxidative phosphorylation (OXPHOS) in mitochondria. Therefore, we questioned whether the methionine-SAM axis regulates mitochondrial OXPHOS through methylation reaction, thereby affecting intracellular ROS levels. To test, oxygen consumption rate (OCR), an integrative and comprehensive readout of cellular metabolism and mitochondrial function coupled with respiration efficiency, was measured. The results showed that methionine restriction or the blockade of methionine-SAM conversion by MAT2A inhibitor FIDAS-5 mitigated OCR, which was replenished by supplementation of SAM (Fig. 4A, B). Further inhibition of SAM-based methylation by the methyltransferase inhibitor ADOX downregulated OCR (Fig. 4A, B). Moreover, OXPHOS inhibitor oligomycin (Oligo) significantly decreased SAM-based ROS accumulation and ferroptosis (Fig. 4C, D). These results indicated that methionine-SAM axis is required for mitochondrial OXPHOS, and is responsible for the generation of mitochondrial ROS (mtROS).

A, B MEF cells were cultured in the medium ± cystine, ± methionine, ± SAM or ± ADOX as indicated for 6 h. A The OCR was determined with Seahorse XFp. B The max OCR was calculated by Wave software 2.3.0. C, D MEF cells were cultured in the medium ± cystine or ± methionine in the absence or presence of SAM or Oligo for 8–12 h. C The cellular levels of ROS were respectively quantified at the 8 h. D The cell death levels were quantified at the 12 h. E, F HT1080 cells were transfected with the GFP-tagged Mito-pHyPer plasmid for 48 h, and then cultured as indicated for another 8 h. The fluorescence intensities of Mito-pHyPer were analyzed and quantified by fluorescent confocal microscopy. E Representative images and (F) dot plots showing intensities of Mito-pHyPer were shown. From left, n = 266, 78, 318, 476, 395, 130, 128, 150 cells. Scale bar, 5 μm. a.u. arbitrary unit. For all box plots, the bottom, middle line, and top of the box and the whiskers indicate the 25th, 50th, 75th and 10th–90th percentiles, respectively, and means are shown as green “+” symbols. G, H HT1080 cells were cultured in the medium ± cystine or ± methionine in the absence or presence of SAM or Mito-TEMPO for 10–14 h. CC, 0.4 mM; Met, 0.4 mM; SAM, 0.4 mM; FIDAS-5, 5 μM; ADOX, 20 μΜ; Oligo, oligomycin, 1 μM for OCR and 10 μM for lipid ROS and cell death; FCCP, 2 μM; Rot, rotenone, 0.5 μM; AA, antimycin A, 0.5 μM; MT, Mito-TEMPO, 5 μM. n = 3 (B–D) or n = 4 (G, H) biological replicates, data were represented as mean ± SD with p values determined by one-way ANOVA test (B–D, F–H). Source data are provided as a Source Data file.

To get insight into the causality of mtROS generation for the induction of ferroptosis, ectopic overexpression of a mitochondrial localized oxidant-sensitive green fluorescent protein (pHyPer-Mito)32,33 was conducted. Cystine deprivation resulted in a mild activation of the pHyPer-Mito probe in HT1080 cells (Fig. 4E, F); whereas, deprivation of methionine or inhibition of MAT2A significantly reduced the cellular sensitivity of the pHyPer-Mito probe (Fig. 4E, F). Supplementation of SAM restored the activation of the pHyPer-Mito probe, which was blocked by ADOX, Oligo, as well as the canonical mitochondrial-targeted antioxidants Mito-TEMPO (MT) (Fig. 4E, F), indicating that methionine-SAM axis contributed to mtROS accumulation. Moreover, SAM-based lipid peroxidation and ferroptosis were significantly quenched by mitochondrial-targeted antioxidant MT (Fig. 4G, H). In summary, these data demonstrated that mitochondrial OXPHOS-based ROS accumulation downstream methionine-SAM axis is implicated in the induction of ferroptosis.

Next, how the methionine-SAM axis regulates mitochondrial ROS generation through methylation arose as a question. Ubiquinone (UQ) is a center electron carrier in the respiratory chain that receives electrons from mitochondrial complex I or II and transfers them to complex III, where electron may be leaked to oxygen to form superoxide anion (O2·−). Importantly, UQ is the unique electron carrier that requires methylation to involve in the de novo synthesis (Fig. 5A). Therefore, we speculated that ROS is majorly generated from ubiquinone-based electron transfer during ferroptosis. To test, we firstly used antimycin A (AA) to block the electron transfer from UQ to cytochrome c. The results shown that AA completely inhibited SAM-based mtROS or total ROS accumulation, as well as cell death (Fig. 5B–D), indicating the UQ-mediated electron transmitting is indeed required for SAM-based ferroptosis. Meanwhile, we found that restriction of methionine in the absence of cystine significantly reduced cellular UQ levels in HT1080 cells (Fig. 5E). Tyrosine-derived 4-hydroxybenzoate is the precursor of the UQ quinone ring34. And the polyisoprenoid side-chain is made from mevalonate-originated farnesyl pyrophosphate and several isopentenyl pyrophosphate molecules34. In parallel with the observation that methionine deprivation decreased the content of UQ (Fig. 5E), the levels of the metabolites upstream of UQ synthesis, such as tyrosine and mevalonate markedly increased (Fig. S10A, B). However, while the level of UQ was rescued by replenishment of SAM, that of tyrosine or mevalonate was restrained (Figs. 5E and S10A, B). Moreover, inhibition of methylation reaction by ADOX treatment, which eliminated the effect of SAM on the variations of the amount of UQ, tyrosine or mevalonate (Figs. 5E and S10A, B). Taken together, these data indicated that methionine limitation affects UQ synthesis through SAM-based methylation, and UQ-based electron transfer is the major source for ROS generation during cystine limitation-induced ferroptosis.

A Schematic of key proteins in SAM-based ubiquinone synthesis pathway. B HT1080 cells were transfected with the GFP-tagged Mito-pHyPer plasmid for 48 h, and then cultured as indicated for another 8 h. The mean fluorescence intensities (MFI) of Mito-pHyPer were quantified. C–E HT1080 cells were cultured as indicate to quantify the cellular levels of ROS (C) at the 10 h, the cell death levels (D) at the 14 h and the UQ levels (E) at the 8 h. F–I HT1080 cells were respectively transfected with siRNA targeting SLC25A26 (siSLC25A26-1/2 mixture), CoQ3 (siCoQ3-1/2 mixture) or CoQ5 (siCoQ5-1/2 mixture) for 48 h, and then cultured as indicated for another 8–14 h. (F) The UQ levels were detected at the 8 h. G, H The levels of cellular ROS and lipid ROS were determined at the 10 h. I The cell death levels were quantified at the 14 h. J HT1080 cells were transfected with SLC25A26 plasmid for 48 h, and then cultured in the medium ± cystine, ± methionine as indicated for another 14 h to quantify the cell death. The proteins of SLC25A26 and tubulin were determined by WB. K–M HT1080 cells were co-transfected with SLC25A26 and Mito-pHyper for 48 h, and then cultured in the medium ± cystine, ± methionine in the absence or presence of Mito-TEMPO as indicated for another 8–10 h. K, L The UQ levels and the MFI of Mito-pHyper were respectively quantified at the 8 h. M The lipid ROS levels were determined at the 10 h. CC, 0.4 mM; Met, 0.4 mM; SAM, 0.4 mM; FIDAS-5, 5 μM; ADOX, 20 μM; AA, 10 μM; MT, 5 μM. For B-M, n = 3 biological replicates, data were represented as mean ± SD with p values determined by one-way ANOVA test. Source data are provided as a Source Data file.

The de novo synthesis of UQ involves multi-step reactions, including mitochondrial import of SAM through the importer SLC25A26, transfer of methyl group from SAM for the synthesis of UQ by methyltransferases such as 3-demethylubiquinol-10 3-O-methyltransferase (CoQ3) and 2-decaprenyl-6-methoxy-1,4-benzoquinone methyltransferase (CoQ5)34,35 (Fig. 5A). Next, we designed siRNAs targeting SLC25A26, CoQ3 or CoQ5, to further specify the importance of SAM-dependent UQ synthesis in ferroptosis induction. The result showed that knockdown of SLC25A26, CoQ3 or CoQ5 significantly decreased SAM-based UQ synthesis and ROS accumulation (Figs. 5F, G and S10C–E), as well as SAM-based lipid peroxidation and cell death induced by cystine deprivation (Fig. 5H, I). Moreover, we inhibited 5-demethoxyubiquinone hydroxylase (CoQ7), another enzyme for UQ synthesis but not a methyltransferase34 (Fig. 5A), with inhibitor COQ7-IN-136, to block the synthesis of UQ in HT1080 cells (Fig. S10F). The results showed that COQ7-IN-1 also inhibited cystine deprivation- or SAM supplementation-induced ROS accumulation, lipid peroxidation and cell death (Fig. S10G–I). Importantly, when HT1080 cells were cultured in the absence of cystine but with methionine at a very low concentration (such as 1 μM as indicated), ferroptosis is impaired (Fig. 5J). However, overexpression of SLC25A26 restored the levels of UQ, mtROS and lipid peroxidation (Fig. 5K–M), thereby rescuing ferroptosis (Fig. 5J). Additional treatment of mitochondrial-targeted antioxidants MT restrained the effect of SLC25A26 overexpression (Fig. 5K–M). Collectively, these data suggested that methionine-SAM metabolic axis, which is essential for the synthesis of the mobile electron carrier UQ, is responsible for mtROS accumulation and ferroptosis induction.

So far, ferroptosis induction-based tumor growth suppression emerges effective cancer therapeutic potential. Our findings from in vitro investigations prompted us to further analyze of the role of the methionine-SAM axis in ferroptosis-based tumor therapeutic application for cancer treatment in vivo. Firstly, IKE was employed to block xCT-mediated cystine transport to induce OS-RC-2 cells undergoing ferroptosis. Consistent with the observation on ferroptosis induced by cystine withdraw (Fig. 3G–I), IKE-induced ROS accumulation, lipid peroxidation and cell death were also compromised by methionine deprivation or the inhibition of methionine-SAM conversion by FIDAS-5 (Fig. 6A–C). In addition, supply of SAM restored ferroptosis markers, which was wiped off by methylation inhibitor ADOX (Fig. 6A–C). Then, OS-RC-2 cells were subcutaneously injected into athymic nude mice. Eighteen days post tumor colonization, IKE with or without FIDAS-5, SAM or ADOX was administrated intraperitoneally or intragastrically every other day. Another 12 days later, samples were harvested (Fig. 6D). Both the volume and the mass of the tumors in the mice injected with IKE were significantly less than that in mice with saline (Fig. 6E, F). Intriguingly, co-administration of IKE with FIDAS-5 impaired the inhibitory effect of IKE on tumor growth as expected (Fig. 6E, F); whereas, co-administration of IKE with SAM mildly but significantly strengthened the inhibition effect of IKE (Fig. 6E, F). However, further injection of ADOX reversed the inhibitory effect of IKE and SAM (Fig. 6E, F).

A–C OS-RC-2 cells were cultured in the medium ± IKE, ± methionine, ± FIDAS-5, ± SAM, or ± ADOX as indicated for 24–30 h. A The cellular levels of ROS were respectively quantified at the 24 h. B The lipid ROS levels were determined at the 24 h. C The cell death levels were quantified at the 30 h. IKE, 2 μM; Met, 0.4 mM; SAM, 0.4 mM; FIDAS-5, 5 μM; ADOX, 20 μΜ. D–F The effect of IKE combined with FIDAS-5, SAM, or ADOX on tumors. OS-RC-2 cells were injected into athymic nude mice. Eighteen days after tumor colonization, IKE (40 mg/kg), SAM (250 mg/kg) or ADOX (2 mg/kg) was injected intraperitoneally every 2 day, and FIDAS-5 (20 mg/kg) was injected intragastrically every 2 day. The xenograft tumors were sampled and photographed after 12 days. D The tumors dissected from the mice were photographed. Tumor volume (E) and tumor mass (F) were measured. G The PTGS2 mRNA level in tumor xenografts were determined by qPCR. H Representative immunohistochemical images of MDA, 4-HNE and Ki67 are shown. Scale bars, 10 μm; Representative images of TUNEL staining are shown. Scale bars, 10 μm. I, J, L Relative intensities of MDA, 4-HNE or Ki67 in tumor xenografts were calculated. K The TUNEL-positive ratios in tumor xenografts were calculated. n = 3 biological replicates (A–C); n = 5 mice per group (E, F); n = 3 random samples from each tumor xenograft tissue of the five analyzed mice per group (G); n = 5 random fields from each tumor xenograft tissue of the five analyzed mice per group (I–L); data were represented as mean ± SD with p values determined by one-way ANOVA test (A–C, E–G, I–L). Source data are provided as a Source Data file.

In the tumor xenograft tissues, IKE-induced lipid peroxidation markers, including PTGS2 upregulation, as well as 4-hydroxynonenal (4-HNE) or malondialdehyde (MDA) accumulation were significantly alleviated due to injection of FIDAS-5 (Fig. 6G–J). On the other hand, co-administration of SAM enhanced the markers induced by injection of IKE, which was reversed by further administration of ADOX (Fig. 6G–J). Parallelly, we carried out the terminal deoxynucleotidyl transferase dUTP nick-end labeling (TUNEL) assay to evaluate the ferroptosis levels as reported37,38, and the results showed that the ratio of TUNEL-positive cells was positively correlated with lipid peroxidation levels (Fig. 6H, K); whereas, Ki67, a classic marker of cell proliferation, was negatively correlated with the levels of lipid peroxidation and cell death in tumor xenografts (Fig. 6H, L). Taken together, these results indicated that SAM supplementation synergizes with IKE in tumor growth suppression, and suggested that targeting SAM-based methylation may hold promise of being applied for cancer therapies.

Other than cancer treatment, we analyze the role of the methionine-SAM axis in ferroptosis-related pathogenesis in vivo. Doxorubicin (DOX) is a clinical drug and widely used to treat breast cancer, leukemia, and many other types of cancer. However, it has been found that DOX can also induce cardiomyocyte ferroptosis, which causes cardiomyopathy in clinical applications25,26. Then, we treated mouse HL-1 cardiomyocytes with DOX and found that the ferroptosis markers were induced upon DOX treatment, such as ROS accumulation, lipid peroxidation, upregulation of PTGS2, as well as the release of myocardial enzyme lactate dehydrogenase (LDH), the well-known cardiomyopathy marker (Fig. 7A–D). However, these observations were significantly blocked by restriction of methionine (Fig. 7A–D). Collectively, the data suggested that methionine-SAM axis is required for DOX-induced ferroptosis in cardiomyocytes.

A–D HL-1 cells were cultured ± methionine and simultaneously treated with DOX as indicated for 12 h. A The cellular levels of ROS were respectively quantified. B The lipid ROS levels were determined. C The PTGS2 mRNA levels were determined by qPCR. D The LDH levels were measured by kits. DOX, Doxorubicin, 4 μM; Met, 0.4 mM; FIDAS-5, 5 μM; Fer-1, 10 μM. E The mice were injected with DOX (10 mg/kg) only or combined with FIDAS-5 (20 mg/kg) or Fer-1 (1 mg/kg) as indicated for 4 days. The mice heart tissues were subjected to histological examination by HE-staining. Representative images were shown. Scale bars, 500 μm. F The LDH levels in mice serum were measured by kits. G The PTGS2 mRNA levels in mice heart tissues were determined by qPCR. H Representative immunohistochemical images of MDA and 4-HNE are shown. Scale bars, 10 μm; Representative images of TUNEL staining are shown. Scale bars, 10 μm. I Relative intensities of MDA and 4-HNE in mice heart were calculated. J TUNEL-positive ratios in mice heart were analyzed. n = 3 biological replicates (A–D); n = 3 samples from each mouse serum of the three analyzed mice per group (F); n = 3 samples from each mouse heart tissue of the three analyzed mice per group (G); n = 9, three random fields from each mouse heart tissue of the three analyzed mice per group (I, J); data were represented as mean ± SD with p values determined by one-way ANOVA test (A–D, F, G, I, J). Source data are provided as a Source Data file.

In vivo, we established a mouse cardiotoxicity model with intraperitoneal injection of DOX to investigate the role of methionine in ferroptosis. FIDAS-5, as well as lipid peroxide scavenger Fer-1, significantly alleviated DOX-induced cardiomyopathy shown by hematoxylin-eosin (HE) staining in mice heart tissue and by LDH release in mice serum (Fig. 7E, F). In addition, we detected the other key markers of cell death and lipid peroxidation, and the results showed that injection of DOX induced a significant increase in ratios of TUNEL-positive cells and lipid peroxidation marker PTGS2, 4-HNE or MDA in mouse heart tissues, which was rescued by MAT2A inhibitor FIDAS-5 or lipid peroxide scavenger Fer-1 (Fig. 7G–J). These data indicated that methionine-SAM metabolic axis contributes to DOX-induced lipid peroxidation and cardiomyocyte ferroptosis, and suggested MAT2A inhibition could be a therapeutic strategy to alleviate the cardiomyopathy induced by DOX.

Ferroptosis is a very unique type of cell death because it is not only regulated by lipid metabolism and iron metabolism, but also by amino acid metabolism. Cysteine, a limiting substrate for GPX4/GSH system that neutralized lipid peroxides, can prevent ferroptosis. Cysteine can be synthesized from methionine via the transsulfuration pathway. As a sulfur-containing amino acid, the conversion of methionine to cysteine is supposed to be enhanced under cystine limiting conditions; however, numerous studies did not observe an effective rescue of intracellular cysteine level under cystine deprivation despite the presence of methionine. In the present study, we clarified the tissue specificity of the methionine-based endogenous cysteine synthesis. We demonstrated that methionine not only scarcely has effect on inhibiting ferroptosis, instead, is indispensable in cystine deprivation-induced ferroptosis. SAM, the methionine-derived metabolite, contributes to ROS generation through methylation-based ubiquinone synthesis. SAM supplementation synergizes with IKE in tumor growth suppression. Inhibition of SAM production by targeting MAT2A alleviates DOX-induced and ferroptosis-driven cardiomyopathy.

Although both cysteine and methionine are sulfur-containing amino acids, we found that cysteine and methionine play different roles in ferroptosis induction. Cysteine exerts an inhibitory effect on ferroptosis, while the occurrence of ferroptosis is methionine-dependent. The cells were unable to acquire cysteine from methionine and failed to restore the intracellular levels of cysteine and GSH, suggesting the metabolic pathway of methionine to cysteine may be interrupted. Similarly, a recent study has indicated that in the non-small-cell lung cancer cells, the transsulfuration pathway that synthesizes cysteine de novo from methionine-derived homocysteine and serine, cannot support intracellular cysteine levels39. Likewise, cancer cell lines including liver cancer, small cell lung cancer, pancreatic cancer cannot support the cysteine pool via transsulfuration in genetically engineered mouse cancer models of these cancers in vivo40. On the contrary, in the thioredoxin reductase-1 (TrxR1)- and glutathione reductase (GR)- null mouse hepatocytes, that could not reduce GSSG to GSH due to lacking of disulfide reducing power, the [35S]-labeled methionine can be converted to into cysteine through the transsulfuration pathway and thereby sustain the de novo synthesis of GSH41, indicating that cysteine can synthesized from methionine in mouse hepatocytes. Coincidentally, it has been reported that upon extracellular cystine limitation in a wide variety of cancer cell lines, the expression of transsulfuration enzymes increased, but the rate of transsulfuration is limited by the cellular capacity to SAM-SAH conversion which mainly take place in the liver24. In our study, we unveil a liver specificity of the methionine-based cysteine synthesis. In addition, out of expectation, we find that, not only having no significant inhibitory role, but reversely, methionine plays an essential role in cystine deprivation-induced ferroptosis. Methionine-derived SAM contributes to the methylation-dependent ubiquinone synthesis, which is the causality of mtROS generation for the induction of ferroptosis.

Ubiquinone (UQ), also named as coenzyme Q10 (CoQ10), is a mobile component of the mitochondrial respiratory chain34, where CoQ10 undergoes a redox cycle (fully oxidized ubiquinone versus fully reduced ubiquinol) that is central to its role in the electron transport20. However, the plasma membrane-situated CoQ10, at its reduced form (ubiquinol, UQH2), serves as an antioxidant and helps prevent ferroptosis through radical trapping and suppression of lipid peroxidation42. Generally, treatment with exogenous CoQ10 is often ineffective, likely due to its extreme hydrophobicity and high molecular weight42. We supplement a cell permeable and functional version of exogenous ubiquinone CoQ4 (a synthetic substitute for CoQ10)42 in the present study, and found that exogenous CoQ4 could restore the oxygen consumption rate (OCR) under the condition of methionine deprivation (Fig. S11A), indicating that CoQ4 could be delivered into the mitochondria and integrated into electron transport chain. Nevertheless, supplementation of CoQ4 failed to restore methionine deprivation-reduced accumulation of mtROS or total ROS, lipid peroxidation and ferroptosis (Fig. S11B–E). Potentially, although exogenous CoQ4 could enter the mitochondrial inner membrane, most of CoQ4 would still localize within the plasma membrane. Hence, mitochondrial CoQ4-produced ROS are supposed to be quenched by the CoQ4 at the plasma membrane. This may explain why the exogenous supplementation of CoQ4 did not drive cells undergoing ferroptosis under methionine deprivation in the present study.

The GSH-dependent lipid hydroperoxidase GPX4 prevents ferroptosis by converting lipid hydroperoxides into non-toxic lipid alcohols. Ferroptosis can be induced by either depletion of antioxidant GSH through cystine limitation, or by direct GPX4 inhibition induce ferroptosis widely. However, we revealed a specificity of methionine starvation in protection against ferroptosis induced by cystine limitation but not GPX4 inhibition, which is consistent with previous report21. Our data substantially demonstrate methionine-SAM-mtROS axis is the major cause for lipid peroxidation and ferroptosis if cystine is not available. Coincidentally, another study reported that mitochondria play a crucial and proactive role in cystine deprivation-induced ferroptosis but not in GPX4 inhibition/knockout-induced ferroptosis43. Similar to our conclusion, the mitochondrial electron transfer promotes cystine deprivation-induced ferroptosis by serving as the major source of ROS generation and subsequent cellular lipid peroxide production2. GPX4-inhibition results in no detoxification of lipid peroxides even though without ROS release from mitochondrial OXPHOS, implying the lipid peroxidation might also stem from other oxidants. Our present study together with those of others suggested that the condition of cystine limitation is not equal to the direct GPX4 inhibition. Obviously, in response to cystine deprivation, GPX4 still can counteract with the lipid peroxides which are imposed by the rest few other cellular ROS upon methionine deprivation, yet it cannot if completely inhibited by RSL3.

FSP1/CoQ axis was identified as a nonmitochondrial CoQ antioxidant system and acts parallel to GPX4 to inhibit ferroptosis44,45. However, as documented in the previous studies, FSP1/CoQ inhibition was functional under a condition to synergize with GPX4/GSH system44,46. In the present study, inhibition of FSP1/CoQ axis alone indeed was unable to induce ferroptosis (Fig. S12A–C). In contrast, the combination of iFSP1 (FSP1 inhibitor) caused more severe ferroptosis than cystine deprivation alone in human kidney-2 (HK-2) cells (Fig. S12D). Nevertheless, utilization of iFSP1 under the condition absent of both cystine and methionine could not elicit ferroptosis (Fig. S12D). Although methionine deficiency-resulted impairment UQ synthesis may limit amount of antioxidant CoQ and dampen the function of FSP1 in the plasma membrane, the outage of the electron transfer through the mobile electron carrier UQ is supposed to cease mtROS accumulation. Thus, a defect in UQ production caused by methionine starvation may alleviate the cell ferroptosis that is induced by either invalidation of GSH/GPX4 system or imperfect FSP1/CoQ axis.

A recent study also observed that methionine is required for ferroptosis execution in response to the blockade of cystine uptake21. They found that deprivation of cystine alone depletes intracellular GSH, while deprivation of cystine and methionine unexpectedly increased intracellular GSH level21. In our study, we also found deprivation of both methionine and cystine resulted in a mild increase in GSH levels compared with cystine deprivation alone (Figs. 2C and S6A, C). However, the level of SAM, the metabolite only converted from methionine, was largely decreased (Figs. 2C and S6A, C), which impelled us to determine the effect of SAM in cystine deprivation-induced ferroptosis. In the present study, we unveiled that methionine-derived SAM is exploited for the synthesis of UQ to maintain the integrity of mitochondrial respiratory chain, thereby methionine deprivation suppresses the accumulation of intracellular ROS that are released from electron transfer. We have clarified that the promoting effect of methionine-SAM metabolism on cystine restriction-induced ferroptosis is independent of GSH levels. This deciphers the mechanism how methionine deprivation inhibits ferroptosis from a perspective of limiting the generation of ROS; whereas, the investigation of Xue et al. was from the view of blocking the exhaustion of antioxidant21. Methionine deprivation simultaneously reduces the oxidant and increases the antioxidant, thereby efficiently suppressing ferroptosis.

Roles of amino acid metabolism in ferroptosis regulation have been drawing increasing attention. A recent study observed that deprivation of glutamine, lysine, valine, arginine, as well as methionine suppressed ferroptosis in both U-2OS and HT1080 cells in response to Erastin2 or cystine deprivation47. In our study, we supplemented SAM to MEF cells under the conditions of cystine limitation along with deprivation of glutamine, lysine, valine, arginine or methionine. As expected, as a control, methionine restriction-inhibited ferroptosis was almost completely resumed upon SAM supplementation. Yet, the outcome from arginine restriction was moderately restored; whereas, SAM scarcely affected other amino acid deprivation-caused consequences (Fig. S13). Recently, arginine is identified as a ferroptosis-promoting factor due to that the conversion of arginine to polyamines contributes to H2O2 production48. In the present study, methionine-derived SAM also displayed the potent ferroptosis-promoting property though mediating ROS accumulation. This may be the reason why SAM supplementation rescued arginine deficiency-induced ferroptosis to some extent (Fig. S13). Besides, glutaminolysis could replenish TCA cycle and then maintain mitochondrial membrane potential hyperpolarization, thereby contributing an increase of electron transfer chain activity and subsequent ROS generation43,49. Obviously, SAM could not complement electron transfer chain activity if TCA cycle is suspended (Fig. S13). To date, the mechanisms of lysine and valine involved in ferroptosis have remained unknown, however, which may not be related with ROS generation/accumulation but with other alternative mechanisms such as impairing translation or mTOR signaling. Taken together, SAM supplementation cannot efficiently rescue the ferroptosis induced by deprivation of glutamine, lysine and valine, indicating that amino acids may work through different mechanisms to modulate ferroptosis.

In summary, the present study unveiled an unexpected role of methionine in ferroptosis induction, and provided a new insight into methionine-SAM metabolic axis-mediated ROS production. The study broadened our understanding about the role of amino acid metabolism in regulation of ferroptosis, and suggested that targeting the methionine-SAM metabolic axis, for example, by inhibiting MAT2A, could be a potential therapeutic strategy for diseases associated with ferroptosis.

The use of animal models in this research complies with all relevant ethical regulations of the Ethics Committee of Science and Technology of Northeast Normal University (202302037).

Human HT1080 fibrosarcoma cells, human OS-RC-2, Caki-1, 786-O and ACHN renal carcinoma cells, human HT29 colorectal adenocarcinoma cells, human THP1 monocytic leukemia cells, human hepatocytes MIHA, human kidney-2 (HK-2) cells, mouse embryonic fibroblast (MEF) cells, mouse cardiomyocytes HL-1 and hepatocytes AML12 were purchased from the American Tissue Culture Collection (ATCC) and authenticated by the distributors. All cell lines were routinely tested for the presence of mycoplasma by a PCR-based method.

MEF, HT1080 and HL-1 cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin and streptomycin (P/S) at 37 °C with 5% CO2. OS-RC-2, 786-O and MIHA cells were cultured in RPMI-1640 medium supplemented with 10% FBS and 1% P/S at 37 °C with 5% CO2. ACHN and HK-2 MEM cells were cultured in MEM medium supplemented with 10% FBS and 1% P/S at 37 °C with 5% CO2. Caki-1 cells were cultured in McCoy’s 5A medium supplemented with 10% FBS and 1% P/S at 37 °C with 5% CO2. AML12 cells were cultured in DMEM/F12 supplemented with 10% FBS and 1% P/S at 37 °C with 5% CO2. THP1 cells were cultured in RPMI-1640 supplemented with 10% FBS, 0.05 mM β-mercaptoethanol and 1% P/S at 37 °C with 5% CO2.

Primary antibodies used were anti-MAT2A (1:1000 for WB, A19272, Abclonal), anti-MAT1A (1:1000 for WB, A2630, Abclonal), anti-MAT2B (1:1000 for WB, A11608, Abclonal), anti-GAMT (1:1000 for WB, A6597, Abclonal), anti-GNMT (1:1000 for WB, A6608, Abclonal), anti-NNMT (1:1000 for WB, A14030, Abclonal), anti-AHCY (1:1000 for WB, A2756, Abclonal), anti-CBS (1:1000 for WB, A11612, Abclonal), anti-CTH (1:1000 for WB, A6121, Abclonal), anti-CHAC1 (1:1000 for WB, 15207, Proteintech), anti-SLC25A26 (1:1000 for WB, A15557, Abclonal), anti-MDA (1:100 for IHC, ab27642, Abcam), anti-4HNE (1:100 for IHC, bs-6313R, Bioss Antibodies), anti-Ki67 (1:100 for IHC, 34330, CST), anti-Tubulin (1:1000 for WB, HC101-01, TransGen), anti-Actin (1:1000 for WB, HC201-01, TransGen), anti-CoQ3 (1:1000 for WB, 28051-1-AP, Proteintech), and anti-CoQ5 (1:1000 for WB, K009369P, Solarbio). The secondary antibodies included goat anti-mouse IgG secondary antibody HRP conjugated (1:5000 for WB, L3032, SAB) and goat anti-rabbit IgG secondary antibody HRP conjugated (1:5000 for WB, L3012, SAB). The compounds used are Ferrostain-1 (Fer-1) (S7243, Selleck), Deferoxamine (DFO) (D9533, Sigma), Propidium iodide (PI) (P4170, Sigma), BODIPY581/591 C11 (D3861, ThermoFisher), Lipofectamine 2000 (11668500, Invitrogen), DCFH2-DA (D6883, Sigma), Cystine (30200, Sigma), Homocysteine (Hcy) (H4628, Sigma), Cystathionine (Cta) (C7505, Sigma), S-adenosylmethionine (SAM) (A7007, Sigma), S-adenosylhomocysteine (SAH) (A9384, Sigma), Methionine (Met) (M9625, Sigma), FIDAS-5 (HY-136144, MCE), PF-9366 (S0435, Selleck), Adenosine Dialdehyde (ADOX) (S8608, Selleck), Mitoguazone (MGBG) (HY-15763, MCE), Erastin (HY-15763, MCE), Imidazole Ketone Erastin (IKE) (S8877, Selleck), Mito-TEMPO (HY-112879, MCE), Oligomycin (HY-N6782, MCE), Antimycin A (MS0070, MaoKangBio), Coenzyme Q4 (C921665, Maklin), RSL3 (HY-100218A, MCE), Doxorubicin (DOX) (HY-15142, MCE), Apoptosis Inducer Kit (TNF-α/SM-164, C0006, Beyotime), Necroptosis Inducer Kit (TNF-α/SM-164/Z-VAD-FMK, C1058, Beyotime), Annexin V-FITC Apoptosis Detection Kit (C1062, Beyotime), LPS (Sigma, L2630), Nigericin (HY-100381, MCE), Methylnitronitrosoguanidine (MNNG) (HY-128612, MCE), alpha-phenyl-tert-N-butylnitrone (PBN, B100838, Aladdin), iFSP1 (HY-136057, MCE), MitoTracker (M7512, Thermo), Total Glutathione Assay Kit (S0052, Beyotime), Cysteine Content Assay Kit (D799571-0050, Sangon Biotech), LDH Cytotoxicity Assay Kit (C0016, Beyotime), CoQ10 Test Kit (PM23627, PERFEMIKER), Mevalonate ELISA Kit (YX-132201M, Sino Best Biological Technology), Tyrosine Assay Kit (ab185435, Abcam), In Situ Cell Death Detection (TUNEL) Kit (11684817910, Roche), S-adenosylmethionine (SAM) ELISA Kit (BY-JZF0361, BYabscience, Nanjing, China), S-adenosylhomocysteine (SAH) ELISA Kit (BY-JZF0362, BYabscience), Mito Stress Test Kits (101706-100, Agilent). Dulbecco’s Modified Eagle’s Medium high glucose without L-methionine, L-cystine and L-glutamine (D0422-100 ml, Sigma), L-glutamine (49419, Sigma), L-lysine (L5501, Sigma), L-valine (V0500, Sigma), L-arginine (A5006, Sigma), Penicillin-Streptomycin (P06-07100, Pan Biotech). Other reagents were obtained from Sigma.

Cells were seeded at a density of 5 × 104 per well in a 24-well dish for 12 h in fresh medium. After indicated treatment, 4 μM BODIPY 581/591 C11 was added into cell culture medium and incubated for 25 min. Cells were then trypsinized, washed and resuspended in 0.3 ml of PBS with 5% fetal bovine serum for flow cytometry analysis. Cells positive for BODIPY 581/591 C11 were defined as lipid peroxidation or lipid ROS accumulation. Flow cytometry analysis was carried out with BD FACS Canto II flow cytometer (BD Biosciences), and acquired data were analyzed using FlowJo (version 10).

Cells were seeded at a density of 1 × 105 per well in a 24-well dish for 12 h in fresh medium. After indicated treatment, 1 μg/ml propidium iodide (PI) was added into cell culture medium and incubated for 30 min. Then cells were trypsinized, washed and resuspended in 0.3 ml of PBS with 5% fetal bovine serum for flow cytometry analysis of the cell death level. Flow cytometry analysis was carried out with BD FACS Canto II flow cytometer (BD Biosciences), and acquired data were analyzed using FlowJo, version 10, software (FlowJo).

Cells were seeded at a density of 5 × 104 per well in a 24-well dish and grown for 12 h in fresh medium. After indicated treatment, 10 μM DCFH2-DA was added into cell culture medium and incubated for 30 min. Cells were then trypsinized, washed and resuspended in 0.3 ml of PBS with 5% fetal bovine serum for flow cytometry analysis of the cellular ROS levels. Flow cytometry analysis was carried out with BD FACS Canto II flow cytometer (BD Biosciences), and acquired data were analyzed using FlowJo, version 10, software (FlowJo).

To measure mitochondrial-generated ROS, pHyPer-Mito, a mitochondrial localized oxidant-activated fluorescent protein, was transfected into cells for 48 h, then cells were treated with subsequent treatments such as cystine deprivation, and then stained with MitoTracker. Fluorescence intensity, corresponding to mitochondrial-generated ROS, was quantified by flow cytometry, or captured by fluorescent confocal microscopy (Zeiss LSM980) and analyzed with software NIS-Elements AR version 5.01 (Nikon).

As reported, the cDNA of human CHAC1 was amplified by PCR using the following primers: human CHAC1 forward: ATGAAGCAGGAGTCTGCAGCCCCG, human CHAC1 reverse: CACCAGCGCCAGAGCCTGCTCGGT21. The cDNA of human SLC25A26 was amplified by PCR using the following primers: human SLC25A26 forward: ATGGACCGGCCGGGGTTCGTGG, human SLC25A26 reverse: TCAAGGACTCTTTCTGCCAACTTC. PCR-amplified cDNAs of human CHAC1 or SLC25A26 were cloned into pcDNA 3.1 vector. The plasmid DNAs were validated by Sanger sequencing and purified with an EndoFree Plasmid Midi Kit (CWBio) for transfection experiments.

Glutathione level was measured using the Total Glutathione Assay Kit (S0052, Beyotime) according to the instructions from the manufacturer. In brief, cells were seeded at a density of 2 × 105 per well in a 6-well dish and grown for 12 h in fresh medium. Cells were harvested, and cell numbers were determined, lysed by using liquid nitrogen and 37 °C for multiple rapid freeze-thaw cycles, and centrifugated at 10,000 × g, 4 °C for 10 min. The samples from supernatant of cell and standards from the kit were incubated with appropriate volumes of the reaction mix (Glutathione Reductase, DTNB, and Total Glutathione detection buffer) in 96 well plate. For colorimetric assays, the absorbance at 412 nm was measured with a microplate reader. The total oxidized and reduced glutathione levels of the samples were calculated based on the absorbance of the standards.

The assessment of cysteine content was performed with Cysteine Content Assay Kit (D799571-0050, Sangon Biotech) according to the manufacturer’s instructions. In brief, cells were collected and subjected to cysteine extraction. The extraction sample, as well as the standard solution with different concentrations were respectively mixed with reaction buffer containing the critical reagent, phosphotungstic acid. Cysteine, exited in the extraction sample or standard solution, could reduce the phosphotungstic acid to produce tungsten blue, which has an absorption peak at 600 nm. The content of cysteine can be calculated with the absorbance at 600 nm and normalized with the same cell number of each sample.

SAM levels were assayed using SAM ELISA kit (BYabscience, Nanjing, China) according to the manufacturers’ instructions. Briefly, the supernatants of the cells and the standards from the kit were collected incubated with appropriate volumes of the reaction mix (Chromogen Solution A, Chromogen Solution B, HRP-Conjugate Regent, Stop Solution) in an enzyme-linked 96 well plate. For colorimetric assays, the absorbance at 450 nm was measured with a microplate reader. SAM concentrations of the samples were calculated based on the absorbance of the standards.

SAH levels were assayed using SAH ELISA kit (BYabscience, Nanjing, China) according to the manufacturers’ instructions as described50: the supernatants of the cells and the standards from the kit were collected incubated with appropriate volumes of the reaction mix (Chromogen Solution A, Chromogen Solution B, HRP-Conjugate Regent, Stop Solution) in an enzyme-linked 96 well plate. For colorimetric assays, the absorbance at 450 nm was measured with a microplate reader. SAH concentrations of the samples were calculated based on the absorbance of the standards.

Mevalonate levels were assayed using Mevalonate ELISA kit (Sino Best Biological Technology, Shanghai, China) according to the manufacturers’ instructions as described51: the supernatants of the cells and the standards from the kit were collected incubated with appropriate volumes of the reaction mix (Chromogen Solution A, Chromogen Solution B, HRP-Conjugate Regent, Stop Solution) in an enzyme-linked 96 well plate. For colorimetric assays, the absorbance at 450 nm was measured with a microplate reader. Mevalonate concentrations of the samples were calculated based on the absorbance of the standards.

As reported, tyrosine levels were measured using the Tyrosine Assay Kit (Abcam)52 according to the instructions from the manufacturer. In brief, samples from the cells and standards from the kit were incubated with appropriate volumes of the reaction mix (Tyr Assay Buffer, Tyrosine Enzyme Mix) in 96 well plate. For colorimetric assays, the absorbance at 492 nm was measured with a microplate reader. Pyruvate concentrations of the samples were calculated based on the absorbance of the standards.

Cells were seeded at a density of 2 × 106 per well in a 10 cm glass bottom cell culture dish and treated as indicated, and then analyzed at LipidALL Technologies (Changzhou, China). Briefly, polar metabolites were extracted using 1000 µl of ice-cold methanol: H2O (4:1, v/v), and incubated at 2000 × g for 30 min at 4 °C. At the end of the incubation, samples were centrifuged for 10 min at 13,000 × g at 4 °C. Clean supernatant was transferred to a new tube. Extracts were dried in a centrifugal concentrator. The dried extract was reconstituted in 2% acetonitrile in water and subjected to ultra-high performance liquid chromatography-high resolution mass spectrometry (UHPLC-HRMS) analysis. The UHPLC conditions were as previously described53,54: for reverse phase chromatography, polar metabolites were separated on a Waters ACQUITY UPLC HSS T3 columns (1.7 μm, 2.1 × 100 mm, Dublin, Ireland), and mobile phase A was water containing 0.1% formic acid (v/v), and mobile phase B was acetonitrile. The column oven temperature was maintained at 40 °C, and autosampler was set at 10 °C. The injection volume was 10 μL. Flow rate was 0.35 mL/min. The following linear gradient was used: 0–1.0 min with 2% B, 1.0–6.0 min with 2%–42% B, 6.0–8.0 min with 42%–65% B, 8.0–10.0 min with 65%–76% B, 10.0–11.0 min with 76%–100% B, 11.0–14.0 min with 100%–100% B. The eluents were analyzed on an Agilent 1290 II UPLC coupled to Sciex 5600 + quadrupole-TOF MS. HRMS parameters for detection were: ESI source voltage positive ion mode 5.5 kV, negative ion mode −4.5 kV; vaporizer temperature, 450 °C; drying gas (N2) pressure, 50 psi; nebulizer gas (N2) pressure, 50 psi; curtain gas (N2) pressure, 35 psi; The scan range was m/z 60-800. Information-dependent acquisition mode was used for MS/MS analyses of the metabolites. Collision energy was set at (±) 35 ± 15 eV. Data acquisition and processing were performed using Analyst® TF 1.7.1 Software (AB Sciex, Concord, ON, Canada). All detected ions were extracted using MarkerView 1.3 (AB Sciex, Concord, ON, Canada) into Excel in the format of two-dimensional matrix, including mass to charge ratio (m/z), retention time, and peak areas, and isotopic peaks were filtered. PeakView 2.2 (AB Sciex, Concord, ON, Canada) was applied to extract MS/MS data and perform comparisons with the Metabolites database (AB Sciex, Concord, ON, Canada), HMDB, METLIN, and standard references to annotate ion identities. R (version 4.3.1) was used to analyze the UHPLC-HRMS data and then showed that in heatmap (Fig. S6A) and volcano maps (Figs. 2C and S2B).

Total RNA was extracted from cells using TRNzol (TIANGEN, China). Total RNA was then reverse transcribed using the PrimeScript TM RT reagent Kit with gDNA Eraser (TaKaRa) and real-time PCR was performed on the QuantStudio 3 real-time PCR instrument with the TB Green Premix Ex Taq TM (Tli RNase H Plus) (TaKaRa). The mRNA expression of genes was normalized to the expression of β-actin gene. Data were analyzed using the comparative cycling threshold method. The sequences of the primers used for real-time PCR were: PTGS2 (Mouse), forward TGAGCAACTATTCCAAACCAGC, reverse GCACGTAGTCTTCGATCACTATC; β-actin (Mouse), forward GGCTGTATTCCCCTCCATCG, reverse CCAGTTGGTAACAATGCCATGT; MAT1A (Mouse), forward GTGCTGGATGCTCACCTCAAG, reverse CCACCCGCTGGTAATCAACC; MAT2A (Mouse), forward GCTTCCACGAGGCGTTCAT, reverse AGCATCACTGATTTGGTCACAA; MAT2B (Mouse), forward AGGGAACCTTTCACTGGTCTG, reverse ATTTGGAGCAATCGAGCTGAG; AHCY (Mouse), forward CCCTACAAAGTCGCGGACATC, reverse GAGGCTGAGTACATCTCCCG; CBS (Mouse), forward GGGACAAGGATCGAGTCTGGA, reverse AGCACTGTGTGATAATGTGGG; CTH (Mouse), forward TTCCTGCCTAGTTTCCAGCAT, reverse GGAAGTCCTGCTTAAATGTGGTG; NNMT (Mouse), forward TGTGCAGAAAACGAGATCCTC, reverse AGTTCTCCTTTTACAGCACCCA; GAMT (Mouse), forward CACGCACCTGCAAATCCTG, reverse TACCGAAGCCCACTTCCAAGA; GNMT (Mouse), forward AAGAGGGCTTCAGCGTGATG, reverse CTGGCAAGTGAGCAAAACTGT; PEMT (Mouse), forward TGGCTGCTGGGTTACATGG, reverse GCTTCCGAGTTCTCTGCTCC; CoQ3 (Human), forward AAGCTGCGCGTCCCTTAATTT, reverse TCGTCCTGTAGGATTTGAACCAT; CoQ5 (Human), forward GAGCTGGACTTGCATGGGTAT, reverse GGGGATTGTTCACTTGGCTAAAT; SLC25A26 (Human), forward AGTTTCCCTTATGGGAGTCCTT, reverse CACCTGCAAAAGCTCCACAGA; β-actin (Human), forward GTTGTCGACGACGAGCG, reverse GCACAGAGCCTCGCCTT.

Transfections were carried out using Lipofectamine™ 2000 Transfection Reagent (Invitogen) according to a protocol provided by the manufacturer. For reporter assays, cells seeded in 6-well dish were transfected with 40 ng siRNA, siControl-UUCUCCGAACGUGUCACGUUTT, siMAT2A-GGAUCGAGGUGCUGUGCUUTT, siCoQ3-1-CCTGAAACACTAGAGAGCATT, siCoQ3-2- CCGGTGGTTCTTTATTCATTA, siCoQ5-1-GTCTTGGTATCCATCGTGTTT, siCoQ5-2-CGAAAGTCTAACATCAGGCAT, siSLC25A26-1-GCUAUUGGAUCCUUUCCUATT, siSLC25A26-2-GCACAGGUAUCUGCUUCUATT. After 48 h, cells were lysed, and the interference efficiency was measured using western blotting.

Cells were lysed in cell lysis buffer (P0013, Beyotime) containing 20 mM Tris (pH7.5), 150 mM NaCl, 1% Triton X-100, sodium pyrophosphate, β-glycerophosphate, EDTA, Na3VO4, leupeptin for 30 min, and subjected to SDS-PAGE for western blotting. Membranes were blocked with 5% (wt/vol) nonfat dried milk in TBS (pH7.2) containing 0.1% Tween 20 (TBST) and incubated with the appropriate antibodies in 5% (wt/vol) nonfat dried milk in TBST overnight at 4 °C. All primary antibodies (1:1000 in TBST, overnight at 4 °C) incubations were followed by incubation with secondary HRP-conjugated antibody (1:5000 in TBST, 1 h at room temperature), and then visualized using chemiluminescent HRP substrate on chemiluminescence image system (Tanon Science and Technology Company).

For the oxygen consumption rate (OCR) assay, the Seahorse XFp Analyzer (Agilent) and Mito Stress Test Kits (Agilent) were used to assess the OCR of cells followed the instructions provided with the kit. In brief, cells were collected XFp plates (5 × 103/well) via centrifugation in serum-free XF Base Medium (Agilent). OCR was measured under basal state and in response to 1 μM Oligo (Agilent), 2 μM FCCP (Agilent), and 0.5 μM Rot (Agilent) plus 0.5 μM AA (Agilent) employing the Seahorse XFp Analyzer (Agilent) using Wave software, version 2.3.0 (Agilent). Oligo, oligomycin, an inhibitor of mitochondrial complex V; FCCP, fluorocarbonyl cyanide phenylhydrazone, oxidative phosphorylation uncoupling agent; Rot, rotenone, an inhibitor of mitochondrial complex I; AA, antimycin A, an inhibitor of mitochondrial complex III.

Ubiquinone level was measured using the CoQ10 Test Kit (PM23627, PERFEMIKER) according to the instructions from the manufacturer. In brief, cells were seeded at a density of 1 × 106 per well in a 10 cm glass bottom cell culture dish for 12 h in fresh medium. After different treatments, cells were fragmented by sonication, and centrifugated at 10,000 × g, 4 °C for 10 min. The cell supernatants were mixed with chloroform (1:1, v/v) for 30 min, and then subjected to centrifugation at 10,000 × g, 4 °C for 10 min. The samples from the organic phase and standards from the kit were incubated with appropriate volumes of the reaction mix (EC buffer, EC solution, and CoQ10 assay buffer) in 96 well plate. For colorimetric assays, the absorbance at 620 nm was measured with a microplate reader. Ubiquinone levels in the samples were calculated based on the absorbance of the standards.

Lactate dehydrogenase (LDH) release levels were measured using the LDH Release Assay Kit (C0016, Beyotime) according to the instructions from the manufacturer. In brief, the samples from the mouse serum and standards from the kit were incubated with appropriate volumes of the reaction mix (Lactic acid, INT, and Diaphorase Enzyme Mix) in 96 well plate. For colorimetric assays, the absorbance at 490 nm was measured with a microplate reader. LDH release of the samples was calculated based on the absorbance of the standards.

Four-week-old female homozygous (Foxn1nu) mut/mut BALB/c nude mice were purchased from Beijing HFK Bioscience (Beijing, China) and housed under specific-pathogen-free conditions with a 12 h light-12 h dark cycle. The ambient temperature was 22–25 °C, with 45% humidity and the mice had ad libitum access to water and standard chow (1010001, XieTong biology, Nanjing, China). OS-RC-2 cells (1 × 107) were resuspended with 50 μL VitroGel Hydrogel Matrix (The Well Bioscience) and 100 μL PBS. Subsequently, the mixtures were injected into BALB/c-nude mice (6–8 weeks). Eighteen days after tumor colonization, IKE (40 mg/kg), SAM (250 mg/kg) or ADOX (2 mg/kg) was injected intraperitoneally every 2 day, and FIDAS-5 (20 mg/kg) was injected intragastrically every 2 day. The xenograft tumors were sampled and photographed after 12 days. The nude mice were obtained from the Beijing HFK Bioscience Co., Ltd. (Beijing, China), and housed in a 12 h light and black cycle at a 20 degrees centigrade and 40% humidity-controlled room. All animal work procedures were approved by the Ethics Committee of Science and Technology of Northeast Normal University (202302037).

Four-week-old male C57BL/6 mice were obtained from Liaoning Experimental Animal Resource Center (Liaoning, China) and housed under specific-pathogen-free conditions with a 12 h light-12 h dark cycle. The ambient temperature was 22–25 °C, with 45% humidity and the mice had ad libitum access to water and standard chow (1010001, XieTong biology, Nanjing, China). The mice (6–8 weeks) were treated with FIDAS-5 (20 mg/kg) and Fer-1 (1 mg/kg) by gavage and intraperitoneal injection daily, respectively. Then, the next day, the mouse treated with DOX (10 mg/kg) and injected intraperitoneally, after 4 days, mouse blood was collected from the eyeballs and mouse heart was removed for subsequent testing. The disposal methods for animals during the experiment refer to the guiding opinions on treating experimental animals well issued by ministry of science and technology of China. All animal work procedures were approved by the Ethics Committee of Science and Technology of Northeast Normal University (202302037).

The TUNEL assay was used to evaluate the ferroptosis levels in tumor xenografts or mice heart tissues using the In Situ Cell Death Detection (TUNEL) Kit (11684817910, Roche) according to the manufacturer’s instructions. The positive control was used here for quality control, and 4′,6-diamidino-2-phenylindole (DAPI) was used to stain the nuclei. TUNEL labeling sections were scanned by Pannoramic DESK, P-MIDI, P250 (3D HISTECH) and imaged by Pannoramic viewer (P.V 1.15.3). TUNEL-positive cells were analyzed and quantified by NIS-Elements AR (Nikon, Version 5.01.00).

Mouse hearts or tumor xenografts were fixed with 4% paraformaldehyde for 24 h, dehydration, embedded in paraffin, and serially sectioned at 4 µm thickness. The sections were respectively stained with Hematoxylin and Eosin (HE), MDA, 4-HNE, or Ki67. For HE, the sections were scanned by Pannoramic DESK, P-MIDI, P250 (3D HISTECH) and imaged by Pannoramic viewer (P.V 1.15.3). For MDA, 4-HNE and Ki67, the sections were blindly and microscopically imaged by Leica DM3000LED. All the images were analyzed and quantified by NIS-Elements AR (Nikon, Version 5.01.00).

All experiments were performed a minimum of three biological replications. Statistical analysis was carried out with Prism 6.0. All data are shown as mean ± SD. The mean values for biochemical data from each group were compared by unpaired two-tailed t test. Comparisons between multiple time points were analyzed by repeated-measurements analysis of variance (ANOVA). In all tests, p values of less than 0.05 were considered statistically significant.

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

All data to support the conclusions in this paper can be found in the Article, Supplementary Information, or Source Data file, which are provided alongside this paper. The mass spectrometry based-metabolomics data of cellular metabolites levels have been deposited to Metabolomics Workbench55 repository with the study ID ST003504. Source data are provided with this paper.

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This work was supported by National Natural Science Foundation of China (32070896, 32101048 and 32101028), the Young Scientific and Technological Talents Support Project of Jilin Province (QT202105), the China Postdoctoral Science Foundation (2020T130090 and 2019M661190). Metabolomics workbench repository is supported by NIH grant U2C-DK119886 and OT2-OD030544 grants.

These authors contributed equally: Chaoyi Xia, Pinghui Peng, Wenxia Zhang.

Key Laboratory of Molecular Epigenetics of the Ministry of Education (MOE), Northeast Normal University, 5268 Renmin Street, Changchun, 130024, Jilin, China

Chaoyi Xia, Pinghui Peng, Wenxia Zhang, Xiyue Xing, Xin Jin, Jianlan Du, Wanting Peng, Fengqi Hao, Zhexuan Zhao, Kejian Dong, Miaomiao Tian, Yunpeng Feng, Xueqing Ba, Min Wei & Yang Wang

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C.X., P.P., and W.Z. conceived the idea, designed the experiments, and composed the paper. C.X., P.P., and W.Z. conducted experiments; W.P., F.H., and W.Z. assisted in animal experiments; X.X., X.J., and J.D. assisted in flow cytometry analysis; J.D. assisted in qPCR; K.D. and Y.W. assisted in tissue analysis; Z.Z. assisted in bioinformatics analysis; X.B., M.W., and Y.W. contributed to the interpretation of the results. Y.F. and M.T. supervised the project.

Correspondence to Xueqing Ba, Min Wei or Yang Wang.

The authors declare no competing interests.

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

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Xia, C., Peng, P., Zhang, W. et al. Methionine-SAM metabolism-dependent ubiquinone synthesis is crucial for ROS accumulation in ferroptosis induction. Nat Commun 15, 8971 (2024). https://doi.org/10.1038/s41467-024-53380-5

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Received: 17 November 2023

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DOI: https://doi.org/10.1038/s41467-024-53380-5

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