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Nuclear metabolism and the regulation of the epigenome

Abstract

Cellular metabolism has emerged as a major biological node governing cellular behaviour. Metabolic pathways fuel cellular energy needs, providing basic chemical molecules to sustain cellular homeostasis, proliferation and function. Changes in nutrient consumption or availability therefore can result in complete reprogramming of cellular metabolism towards stabilizing core metabolite pools, such as ATP, S-adenosyl methionine, acetyl-CoA, NAD/NADP and α-ketoglutarate. Because these metabolites underlie a variety of essential metabolic reactions, metabolism has evolved to operate in separate subcellular compartments through diversification of metabolic enzyme complexes, oscillating metabolic activity and physical separation of metabolite pools. Given that these same core metabolites are also consumed by chromatin modifiers in the establishment of epigenetic signatures, metabolite consumption on and release from chromatin directly influence cellular metabolism and gene expression. In this Review, we highlight recent studies describing the mechanisms determining nuclear metabolism and governing the redistribution of metabolites between the nuclear and non-nuclear compartments.

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Fig. 1: Metabolic fluxes are redirected to replenish core metabolites.
Fig. 2: Metabolic fluxes are compartmentalized in time, space and subcellular organelles.
Fig. 3: The nucleus is an underappreciated but separated metabolic compartment.
Fig. 4: Regulatory mechanisms balance metabolites between the nuclear and non-nuclear compartments.
Fig. 5: The formation of complexes on chromatin, including metabolic enzymes, drives hyperlocal metabolic conversions.

References

  1. DeBerardinis, R. J. & Chandel, N. S. Fundamentals of cancer metabolism. Sci. Adv. 2, e1600200 (2016).

    PubMed  PubMed Central  Google Scholar 

  2. Zecchin, A., Stapor, P. C., Goveia, J. & Carmeliet, P. Metabolic pathway compartmentalization: an underappreciated opportunity? Curr. Opin. Biotechnol. 34, 73–81 (2015).

    CAS  PubMed  Google Scholar 

  3. Wente, S. R. & Rout, M. P. The nuclear pore complex and nuclear transport. Cold Spring Harb. Perspect. Biol. 2, a000562 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. Walsh, C. T., Tu, B. P. & Tang, Y. Eight kinetically stable but thermodynamically activated molecules that power cell metabolism. Chem. Rev. 118, 1460–1494 (2018).

    CAS  PubMed  Google Scholar 

  5. Vander Heiden, M. G. & DeBerardinis, R. J. Understanding the intersections between metabolism and cancer biology. Cell 168, 657–669 (2017).

    PubMed Central  Google Scholar 

  6. Vander Heiden, M. G., Cantley, L. C. & Thompson, C. B. Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324, 1029–1033 (2009).

    Google Scholar 

  7. Folmes, C. D. L. et al. Somatic oxidative bioenergetics transitions into pluripotency-dependent glycolysis to facilitate nuclear reprogramming. Cell Metab. 14, 264–271 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. Warburg, O., Wind, F. & Negelein, E. The metabolism of tumors in the body. J. Gen. Physiol. 8, 519–530 (1927).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. DeBerardinis, R. J. & Chandel, N. S. We need to talk about the Warburg effect. Nat. Metab. 2, 127–129 (2020).

    PubMed  Google Scholar 

  10. Lunt, S. Y. & Vander Heiden, M. G. Aerobic glycolysis: meeting the metabolic requirements of cell proliferation. Annu. Rev. Cell Dev. Biol. 27, 441–464 (2011).

    CAS  PubMed  Google Scholar 

  11. Locasale, J. W. Serine, glycine and one-carbon units: cancer metabolism in full circle. Nat. Rev. Cancer 13, 572–583 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Comerford, S. A. et al. Acetate dependence of tumors. Cell 159, 1591–1602 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. DeBerardinis, R. J. et al. Beyond aerobic glycolysis: transformed cells can engage in glutamine metabolism that exceeds the requirement for protein and nucleotide synthesis. Proc. Natl Acad. Sci. USA 104, 19345–19350 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Hosios, A. M. & Vander Heiden, M. G. The redox requirements of proliferating mammalian cells. J. Biol. Chem. 293, 7490–7498 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Agostini, M. et al. Metabolic reprogramming during neuronal differentiation. Cell Death Differ. 23, 1502–1514 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  16. Wanet, A. et al. Mitochondrial remodeling in hepatic differentiation and dedifferentiation. Int. J. Biochem. Cell Biol. 54, 174–185 (2014).

    CAS  PubMed  Google Scholar 

  17. Chung, S. et al. Mitochondrial oxidative metabolism is required for the cardiac differentiation of stem cells. Nat. Clin. Pract. Cardiovasc. Med. 4, S60–S67 (2007). (Suppl. 1).

    CAS  PubMed  PubMed Central  Google Scholar 

  18. Kim, H. et al. Core pluripotency factors directly regulate metabolism in embryonic stem cell to maintain pluripotency. Stem Cells 33, 2699–2711 (2015).

    CAS  PubMed  Google Scholar 

  19. Vardhana, S. A. et al. Glutamine independence is a selectable feature of pluripotent stem cells. Nat. Metab. 1, 676–687 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Kasahara, A., Cipolat, S., Chen, Y., Dorn, G. W. II & Scorrano, L. Mitochondrial fusion directs cardiomyocyte differentiation via calcineurin and Notch signaling. Science 342, 734–737 (2013).

    CAS  PubMed  Google Scholar 

  21. Salazar-Roa, M. & Malumbres, M. Fueling the cell division cycle. Trends Cell Biol. 27, 69–81 (2017).

    CAS  PubMed  Google Scholar 

  22. Tu, B. P., Kudlicki, A., Rowicka, M. & McKnight, S. L. Logic of the yeast metabolic cycle: temporal compartmentalization of cellular processes. Science 310, 1152–1158 (2005).

    CAS  PubMed  Google Scholar 

  23. Wang, C. et al. Cyclin D1 repression of nuclear respiratory factor 1 integrates nuclear DNA synthesis and mitochondrial function. Proc. Natl Acad. Sci. USA 103, 11567–11572 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Lee, Y. et al. Cyclin D1–Cdk4 controls glucose metabolism independently of cell cycle progression. Nature 510, 547–551 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Hsieh, M. C. F., Das, D., Sambandam, N., Zhang, M. Q. & Nahlé, Z. Regulation of the PDK4 isozyme by the Rb-E2F1 complex. J. Biol. Chem. 283, 27410–27417 (2008).

    CAS  PubMed  Google Scholar 

  26. Wang, H. et al. The metabolic function of cyclin D3–CDK6 kinase in cancer cell survival. Nature 546, 426–430 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Harbauer, A. B. et al. Cell cycle–dependent regulation of mitochondrial preprotein translocase. Science 346, 1109–1113 (2014).

    CAS  PubMed  Google Scholar 

  28. Liang, J. et al. PKM2 dephosphorylation by Cdc25A promotes the Warburg effect and tumorigenesis. Nat. Commun. 7, 12431 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Slavov, N., Macinskas, J., Caudy, A. & Botstein, D. Metabolic cycling without cell division cycling in respiring yeast. Proc. Natl Acad. Sci. 108, 19090–19095 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Pendleton, K. E. et al. The U6 snRNA m6A methyltransferase METTL16 regulates SAM synthetase intron retention. Cell 169, 824–835.e14 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Sander, T. et al. Allosteric feedback inhibition enables robust amino acid biosynthesis in E. coli by enforcing enzyme overabundance. Cell Syst. 8, 66–75.e8 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Lane, A. N. & Fan, T. W.-M. Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Res. 43, 2466–2485 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Vinnakota, K. C., Bazil, J. N., Van den Bergh, F., Wiseman, R. W. & Beard, D. A. Feedback regulation and time hierarchy of oxidative phosphorylation in cardiac mitochondria. Biophys. J. 110, 972–980 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Panda, S. Circadian physiology of metabolism. Science 354, 1008–1015 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. Etchegaray, J.-P. & Mostoslavsky, R. Interplay between metabolism and epigenetics: a nuclear adaptation to environmental changes. Mol. Cell 62, 695–711 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. Eckel-Mahan, K. & Sassone-Corsi, P. Metabolism and the circadian clock converge. Physiol. Rev. 93, 107–135 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  37. Hatanaka, F. et al. Genome-wide profiling of the core clock protein BMAL1 targets reveals a strict relationship with metabolism. Mol. Cell. Biol. 30, 5636–5648 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. Sinturel, F., Petrenko, V. & Dibner, C. Circadian clocks make metabolism run. J. Mol. Biol. 432, 3680–3699 (2020).

    CAS  PubMed  Google Scholar 

  39. Zhang, E. E. et al. Cryptochrome mediates circadian regulation of cAMP signaling and hepatic gluconeogenesis. Nat. Med. 16, 1152–1156 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Doi, R., Oishi, K. & Ishida, N. CLOCK regulates circadian rhythms of hepatic glycogen synthesis through transcriptional activation of Gys2. J. Biol. Chem. 285, 22114–22121 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Aviram, R. et al. Lipidomics analyses reveal temporal and spatial lipid organization and uncover daily oscillations in intracellular organelles. Mol. Cell 62, 636–648 (2016).

    CAS  PubMed  Google Scholar 

  42. Patel, V. R., Eckel-Mahan, K., Sassone-Corsi, P. & Baldi, P. CircadiOmics: integrating circadian genomics, transcriptomics, proteomics and metabolomics. Nat. Methods 9, 772–773 (2012).

    CAS  PubMed  Google Scholar 

  43. Peek, C. B. et al. Circadian clock interaction withith HIF1α mediates oxygenic metabolism and anaerobic glycolysis in skeletal muscle. Cell Metab. 25, 86–92 (2017).

    CAS  PubMed  Google Scholar 

  44. Schmitt, K. et al. Circadian control of DRP1 activity regulates mitochondrial dynamics and bioenergetics. Cell Metab. 27, 657–666.e5 (2018).

    CAS  PubMed  Google Scholar 

  45. Lamia, K. A. et al. AMPK regulates the circadian clock by cryptochrome phosphorylation and degradation. Science 326, 437–440 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  46. Walton, Z. E. et al. Acid suspends the circadian clock in hypoxia through inhibition of mTOR. Cell 174, 72–87.e32 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Kastritis, P. L. & Gavin, A.-C. Enzymatic complexes across scales. Essays Biochem. 62, 501–514 (2018).

    PubMed  PubMed Central  Google Scholar 

  48. De Bock, K. et al. Role of PFKFB3-driven glycolysis in vessel sprouting. Cell 154, 651–663 (2013).

    PubMed  Google Scholar 

  49. Sweetlove, L. J. & Fernie, A. R. The role of dynamic enzyme assemblies and substrate channelling in metabolic regulation. Nat. Commun. 9, 2136 (2018).

    PubMed  PubMed Central  Google Scholar 

  50. Pareek, V., Tian, H., Winograd, N. & Benkovic, S. J. Metabolomics and mass spectrometry imaging reveal channeled de novo purine synthesis in cells. Science 368, 283–290 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  51. Panicot, M. et al. A polyamine metabolon involving aminopropyl transferase complexes in Arabidopsis. Plant Cell 14, 2539–2551 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Sanyal, N., Arentson, B. W., Luo, M., Tanner, J. J. & Becker, D. F. First evidence for substrate channeling between proline catabolic enzymes: a validation of domain fusion analysis for predicting protein-protein interactions. J. Biol. Chem. 290, 2225–2234 (2015).

    CAS  PubMed  Google Scholar 

  53. Ishikawa, M., Tsuchiya, D., Oyama, T., Tsunaka, Y. & Morikawa, K. Structural basis for channelling mechanism of a fatty acid beta-oxidation multienzyme complex. EMBO J. 23, 2745–2754 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Herzig, S. et al. Identification and functional expression of the mitochondrial pyruvate carrier. Science 337, 93–96 (2012).

    CAS  PubMed  Google Scholar 

  55. Kory, N. et al. SFXN1 is a mitochondrial serine transporter required for one-carbon metabolism. Science 362, eaat9528 (2018).

    PubMed  PubMed Central  Google Scholar 

  56. Yoo, H. C. et al. A variant of SLC1A5 is a mitochondrial glutamine transporter for metabolic reprogramming in cancer cells. Cell Metab. 31, 267–283.e12 (2020).

    CAS  PubMed  Google Scholar 

  57. Agrimi, G. et al. Identification of the human mitochondrial S-adenosylmethionine transporter: bacterial expression, reconstitution, functional characterization and tissue distribution. Biochem. J. 379, 183–190 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  58. Davila, A. et al. Nicotinamide adenine dinucleotide is transported into mammalian mitochondria. eLife 7, e33246 (2018).

    PubMed  PubMed Central  Google Scholar 

  59. Diehl, F. F., Lewis, C. A., Fiske, B. P. & Vander Heiden, M. G. Cellular redox state constrains serine synthesis and nucleotide production to impact cell proliferation. Nat. Metab. 1, 861–867 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. Feldman, J. L. et al. Kinetic and structural basis for acyl-group selectivity and NAD+ dependence in sirtuin-catalyzed deacylation. Biochemistry 54, 3037–3050 (2015).

    CAS  PubMed  Google Scholar 

  61. Ryu, K. W. et al. Metabolic regulation of transcription through compartmentalized NAD+ biosynthesis. Science 360, eaan5780 (2018). This study shows a switch from nuclear to cytoplasmic NAD+ production that sustains adipogenic differentiation through the PARP-dependent regulation of C/EBPβ.

    PubMed  PubMed Central  Google Scholar 

  62. Cambronne, X. A. et al. Biosensor reveals multiple sources for mitochondrial NAD+. Science 352, 1474–1477 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. Tibbetts, A. S. & Appling, D. R. Compartmentalization of mammalian folate-mediated one-carbon metabolism. Annu. Rev. Nutr. 30, 57–81 (2010).

    CAS  PubMed  Google Scholar 

  64. Meiser, J. & Vazquez, A. Give it or take it: the flux of one-carbon in cancer cells. FEBS J. 283, 3695–3704 (2016).

    CAS  PubMed  Google Scholar 

  65. Davis, S. R. et al. Tracer-derived total and folate-dependent homocysteine remethylation and synthesis rates in humans indicate that serine is the main one-carbon donor. Am. J. Physiol. Endocrinol. Metab. 286, E272–E279 (2004).

    CAS  PubMed  Google Scholar 

  66. Ducker, G. S. et al. Reversal of cytosolic one-carbon flux compensates for loss of the mitochondrial folate pathway. Cell Metab. 23, 1140–1153 (2016). This study shows that the cytosolic arm can compensate to produce one-carbon units when the default mitochondrial pathway is inhibited. It also identifies a strong dependency on exogenous serine for fuelling the cytosolic pathway.

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Vandekeere, S. et al. Serine synthesis via PHGDH is essential for heme production in endothelial cells. Cell Metab. 28, 573–587.e13 (2018).

    CAS  PubMed  Google Scholar 

  68. Possemato, R. et al. Functional genomics reveal that the serine synthesis pathway is essential in breast cancer. Nature 476, 346–350 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Kawai, S., Suzuki, S., Mori, S. & Murata, K. Molecular cloning and identification of UTR1 of a yeast Saccharomyces cerevisiae as a gene encoding an NAD kinase. FEMS Microbiol. Lett. 200, 181–184 (2001).

    CAS  PubMed  Google Scholar 

  70. Ding, C. C. et al. MESH1 is a cytosolic NADPH phosphatase that regulates ferroptosis. Nat. Metab. 2, 270–277 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  71. Hoxhaj, G. et al. Direct stimulation of NADP+ synthesis through Akt-mediated phosphorylation of NAD kinase. Science 363, 1088–1092 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  72. Lewis, C. A. et al. Tracing compartmentalized NADPH metabolism in the cytosol and mitochondria of mammalian cells. Mol. Cell 55, 253–263 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. Chen, L. et al. NADPH production by the oxidative pentose-phosphate pathway supports folate metabolism. Nat. Metab. 1, 404–415 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  74. Fan, J. et al. Quantitative flux analysis reveals folate-dependent NADPH production. Nature 510, 298–302 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Rosenzweig, A., Blenis, J. & Gomes, A. P. Beyond the Warburg effect: how do cancer cells regulate one-carbon metabolism? Front. Cell Dev. Biol. 6, 90 (2018).

    PubMed  PubMed Central  Google Scholar 

  76. Trefely, S., Lovell, C. D., Snyder, N. W. & Wellen, K. E. Compartmentalised acyl-CoA metabolism and roles in chromatin regulation. Mol. Metab. 38, 100941 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  77. Diehl, K. L. & Muir, T. W. Chromatin as a key consumer in the metabolite economy. Nat. Chem. Biol. 16, 620–629 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. Yanes, O. et al. Metabolic oxidation regulates embryonic stem cell differentiation. Nat. Chem. Biol. 6, 411–417 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. Boon, R. et al. Amino acid levels determine metabolism and CYP450 function of hepatocytes and hepatoma cell lines. Nat. Commun. 11, 1393 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Stillman, B. Histone modifications: insights into their influence on gene expression. Cell 175, 6–9 (2018).

    CAS  PubMed  Google Scholar 

  81. Jenuwein, T. & Allis, C. D. Translating the histone code. Science 293, 1074–1080 (2001).

    CAS  PubMed  Google Scholar 

  82. Brehove, M. et al. Histone core phosphorylation regulates DNA accessibility. J. Biol. Chem. 290, 22612–22621 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. Dawson, M. A. et al. JAK2 phosphorylates histone H3Y41 and excludes HP1α from chromatin. Nature 461, 819–822 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. Mateescu, B., England, P., Halgand, F., Yaniv, M. & Muchardt, C. Tethering of HP1 proteins to chromatin is relieved by phosphoacetylation of histone H3. EMBO Rep. 5, 490–496 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Sabari, B. R., Zhang, D., Allis, C. D. & Zhao, Y. Metabolic regulation of gene expression through histone acylations. Nat. Rev. Mol. Cell Biol. 18, 90–101 (2017).

    CAS  PubMed  Google Scholar 

  86. Fujiki, R. et al. GlcNAcylation of histone H2B facilitates its monoubiquitination. Nature 480, 557–560 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  87. Morris, J. P. IV et al. α-Ketoglutarate links p53 to cell fate during tumour suppression. Nature 573, 595–599 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. Carey, B. W., Finley, L. W. S., Cross, J. R., Allis, C. D. & Thompson, C. B. Intracellular α-ketoglutarate maintains the pluripotency of embryonic stem cells. Nature 518, 413–416 (2015). This study shows that a high αKG/succinate ratio maintains a naive phenotype in pluripotent stem cells through the maintenance of epigenetic hypomethylation and additionally shows the utility of αKG and succinate supplementation in boosting pluripotency and differentiation, respectively.

    CAS  PubMed  Google Scholar 

  89. TeSlaa, T. et al. α-Ketoglutarate accelerates the initial differentiation of primed human pluripotent stem cells. Cell Metab. 24, 485–493 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. Tran, T. Q. et al. α-Ketoglutarate attenuates Wnt signaling and drives differentiation in colorectal cancer. Nat. Can. 1, 345–358 (2020).

    Google Scholar 

  91. Baksh, S. C. et al. Extracellular serine controls epidermal stem cell fate and tumour initiation. Nat. Cell Biol. 22, 779–790 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Su, X., Wellen, K. E. & Rabinowitz, J. D. Metabolic control of methylation and acetylation. Curr. Opin. Chem. Biol. 30, 52–60 (2016).

    CAS  PubMed  Google Scholar 

  93. Banerjee, S., Bennion, G. R., Goldberg, M. W. & Allen, T. D. ATP dependent histone phosphorylation and nucleosome assembly in a human cell free extract. Nucleic Acids Res. 19, 5999–6006 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  94. Zhang, D. et al. Metabolic regulation of gene expression by histone lactylation. Nature 574, 575–580 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  95. Sakabe, K., Wang, Z. & Hart, G. W. Beta-N-acetylglucosamine (O-GlcNAc) is part of the histone code. Proc. Natl Acad. Sci. USA 107, 19915–19920 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. Simithy, J. et al. Characterization of histone acylations links chromatin modifications with metabolism. Nat. Commun. 8, 1141 (2017).

    PubMed  PubMed Central  Google Scholar 

  97. Miousse, I. R. et al. Short-term dietary methionine supplementation affects one-carbon metabolism and DNA methylation in the mouse gut and leads to altered microbiome profiles, barrier function, gene expression and histomorphology. Genes Nutr. 12, 22 (2017).

    PubMed  PubMed Central  Google Scholar 

  98. Mattocks, D. A. L. et al. Short term methionine restriction increases hepatic global DNA methylation in adult but not young male C57BL/6J mice. Exp. Gerontol. 88, 1–8 (2017).

    CAS  PubMed  Google Scholar 

  99. Mentch, S. J. et al. Histone methylation dynamics and gene regulation occur through the sensing of one-carbon metabolism. Cell Metab. 22, 861–873 (2015). This study links diet, methionine metabolism, SAMe levels and histone methylation to an ultimate cancer phenotype.

    CAS  PubMed  PubMed Central  Google Scholar 

  100. Roy, D. G. et al. Methionine metabolism shapes T helper cell responses through regulation of epigenetic reprogramming. Cell Metab. 31, 250–266.e9 (2020).

    CAS  PubMed  Google Scholar 

  101. Yu, W. et al. One-carbon metabolism supports S-adenosylmethionine and histone methylation to drive inflammatory macrophages. Mol. Cell 75, 1147–1160.e5 (2019).

    CAS  PubMed  Google Scholar 

  102. Cluntun, A. A. et al. The rate of glycolysis quantitatively mediates specific histone acetylation sites. Cancer Metab. 3, 10 (2015).

    PubMed  PubMed Central  Google Scholar 

  103. Gao, X. et al. Acetate functions as an epigenetic metabolite to promote lipid synthesis under hypoxia. Nat. Commun. 7, 11960 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  104. Cai, L., Sutter, B. M., Li, B. & Tu, B. P. Acetyl-CoA induces cell growth and proliferation by promoting the acetylation of histones at growth genes. Mol. Cell 42, 426–437 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Barrero, M. J., Boué, S. & Izpisúa Belmonte, J. C. Epigenetic mechanisms that regulate cell identity. Cell Stem Cell 7, 565–570 (2010).

    CAS  PubMed  Google Scholar 

  106. Christie, M. et al. Structural biology and regulation of protein import into the nucleus. J. Mol. Biol. 428, 2060–2090 (2016).

    CAS  PubMed  Google Scholar 

  107. Unnikrishnan, A., Gafken, P. R. & Tsukiyama, T. Dynamic changes in histone acetylation regulate origins of DNA replication. Nat. Struct. Mol. Biol. 17, 430–437 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Skene, P. J. et al. Neuronal MeCP2 is expressed at near histone-octamer levels and globally alters the chromatin state. Mol. Cell 37, 457–468 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  109. Ye, C. & Tu, B. P. Sink into the epigenome: histones as repositories that influence cellular metabolism. Trends Endocrinol. Metab. 29, 626–637 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  110. Martinez-Pastor, B., Cosentino, C. & Mostoslavsky, R. A tale of metabolites: the cross-talk between chromatin and energy metabolism. Cancer Discov. 3, 497–501 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Ye, C., Sutter, B. M., Wang, Y., Kuang, Z. & Tu, B. P. A metabolic function for phospholipid and histone methylation. Mol. Cell 66, 180–193.e8 (2017). This study shows the ability of yeast to either store or mobilize methyl groups on or from histones, thus maintaining the homeostatic cellular levels of SAMe needed to balance phospholipid production.

    CAS  PubMed  PubMed Central  Google Scholar 

  112. Kizer, K. O. et al. A novel domain in Set2 mediates RNA polymerase II interaction and couples histone H3 K36 methylation with transcript elongation. Mol. Cell. Biol. 25, 3305–3316 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. Bannister, A. J. et al. Spatial distribution of di- and tri-methyl lysine 36 of histone H3 at active genes. J. Biol. Chem. 280, 17732–17736 (2005).

    CAS  PubMed  Google Scholar 

  114. Kolasinska-Zwierz, P. et al. Differential chromatin marking of introns and expressed exons by H3K36me3. Nat. Genet. 41, 376–381 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  115. Lacoste, N., Utley, R. T., Hunter, J. M., Poirier, G. G. & Côte, J. Disruptor of telomeric silencing-1 is a chromatin-specific histone H3 methyltransferase. J. Biol. Chem. 277, 30421–30424 (2002).

    CAS  PubMed  Google Scholar 

  116. van Leeuwen, F., Gafken, P. R. & Gottschling, D. E. Dot1p modulates silencing in yeast by methylation of the nucleosome core. Cell 109, 745–756 (2002).

    PubMed  Google Scholar 

  117. Steger, D. J. et al. DOT1L/KMT4 recruitment and H3K79 methylation are ubiquitously coupled with gene transcription in mammalian cells. Mol. Cell. Biol. 28, 2825–2839 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Krogan, N. J. et al. The Paf1 complex is required for histone H3 methylation by COMPASS and Dot1p: linking transcriptional elongation to histone methylation. Mol. Cell 11, 721–729 (2003).

    CAS  PubMed  Google Scholar 

  119. Haws, S. A. et al. Methyl-metabolite depletion elicits adaptive responses to support heterochromatin stability and epigenetic persistence. Mol. Cell 78, 210–223.e8 (2020). This study identifies H3K9me3 and H3K9me2 as metabolic buffer marks that can be mobilized by mammalian cells to replenish cellular levels of SAMe while maintaining H3K9me1 to sustain epigenetic stability.

    CAS  PubMed  PubMed Central  Google Scholar 

  120. Fadloun, A., Eid, A. & Torres-Padilla, M.-E. Mechanisms and dynamics of heterochromatin formation during mammalian development: closed paths and open questions. Curr. Top. Dev. Biol. 104, 1–45 (2013).

    CAS  PubMed  Google Scholar 

  121. Ehrlich, M. DNA hypomethylation in cancer cells. Epigenomics 1, 239–259 (2009).

    CAS  PubMed  Google Scholar 

  122. Peters, A. H. et al. Loss of the Suv39h histone methyltransferases impairs mammalian heterochromatin and genome stability. Cell 107, 323–337 (2001).

    CAS  PubMed  Google Scholar 

  123. Sulkowski, P. L. et al. Oncometabolites suppress DNA repair by disrupting local chromatin signalling. Nature 582, 586–591 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. Lu, C. et al. Histone H3K36 mutations promote sarcomagenesis through altered histone methylation landscape. Science 352, 844–849 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  125. McLean, C. M., Karemaker, I. D. & van Leeuwen, F. The emerging roles of DOT1L in leukemia and normal development. Leukemia 28, 2131–2138 (2014).

    CAS  PubMed  Google Scholar 

  126. Ting, D. T. et al. Aberrant overexpression of satellite repeats in pancreatic and other epithelial cancers. Science 331, 593–596 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. Reid, M. A., Dai, Z. & Locasale, J. W. The impact of cellular metabolism on chromatin dynamics and epigenetics. Nat. Cell Biol. 19, 1298–1306 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  128. Wellen, K. E. et al. ATP-citrate lyase links cellular metabolism to histone acetylation. Science 324, 1076–1080 (2009). This study identifies ACLY as a modulator of histone acetylation through the nuclear production of acetyl-CoA from citrate.

    CAS  PubMed  PubMed Central  Google Scholar 

  129. Li, X. et al. Nucleus-translocated ACSS2 promotes gene transcription for lysosomal biogenesis and autophagy. Mol. Cell 66, 684–697.e9 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  130. Sutendra, G. et al. A nuclear pyruvate dehydrogenase complex is important for the generation of acetyl-CoA and histone acetylation. Cell 158, 84–97 (2014). This study identifies the pyruvate dehydrogenase complex as a modulator of histone acetylation through the nuclear production of acetyl-CoA from pyruvate.

    CAS  PubMed  Google Scholar 

  131. Bulusu, V. et al. Acetate recapturing by nuclear acetyl-CoA synthetase 2 prevents loss of histone acetylation during oxygen and serum limitation. Cell Rep. 18, 647–658 (2017). This study shows that nuclear ACSS2 recaptures acetate released from histones and maintains nuclear acetylation capacity under stress conditions.

    CAS  PubMed  PubMed Central  Google Scholar 

  132. Sivanand, S. et al. Nuclear acetyl-CoA production by ACLY promotes homologous recombination. Mol. Cell 67, 252–265.e6 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  133. Sebastián, C. et al. The histone deacetylase SIRT6 is a tumor suppressor that controls cancer metabolism. Cell 151, 1185–1199 (2012).

    PubMed  PubMed Central  Google Scholar 

  134. Choi, J.-E. & Mostoslavsky, R. Sirtuins, metabolism, and DNA repair. Curr. Opin. Genet. Dev. 26, 24–32 (2014).

    CAS  PubMed  Google Scholar 

  135. Chang, A. R., Ferrer, C. M. & Mostoslavsky, R. SIRT6, a mammalian deacylase with multitasking abilities. Physiol. Rev. 100, 145–169 (2020).

    CAS  PubMed  Google Scholar 

  136. Chalkiadaki, A. & Guarente, L. The multifaceted functions of sirtuins in cancer. Nat. Rev. Cancer 15, 608–624 (2015).

    CAS  PubMed  Google Scholar 

  137. Scher, M. B., Vaquero, A. & Reinberg, D. Sirt3 is a nuclear NAD+-dependent histone deacetylase that translocates to the mitochondria upon cellular stress. Genes Dev. 21, 920–928 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  138. Iwahara, T., Bonasio, R., Narendra, V. & Reinberg, D. SIRT3 functions in the nucleus in the control of stress-related gene expression. Mol. Cell. Biol. 32, 5022–5034 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  139. Finkel, T., Deng, C.-X. & Mostoslavsky, R. Recent progress in the biology and physiology of sirtuins. Nature 460, 587–591 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  140. Onn, L. et al. SIRT6 is a DNA double-strand break sensor. eLife 9, e51636 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  141. Zhong, L. et al. The histone deacetylase Sirt6 regulates glucose homeostasis via Hif1α. Cell 140, 280–293 (2010). This study provides the first demonstration of a metabolic adaptation regulated at the epigenetic level by a chromatin factor (SIRT6).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. Rodgers, J. T. et al. Nutrient control of glucose homeostasis through a complex of PGC-1α and SIRT1. Nature 434, 113–118 (2005).

    CAS  PubMed  Google Scholar 

  143. Gomes, A. P. et al. Declining NAD+ induces a pseudohypoxic state disrupting nuclear-mitochondrial communication during aging. Cell 155, 1624–1638 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  144. Liu, T. F., Vachharajani, V. T., Yoza, B. K. & McCall, C. E. NAD+-dependent sirtuin 1 and 6 proteins coordinate a switch from glucose to fatty acid oxidation during the acute inflammatory response. J. Biol. Chem. 287, 25758–25769 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  145. Svoboda, P. et al. Nuclear transport of nicotinamide phosphoribosyltransferase is cell cycle-dependent in mammalian cells, and its inhibition slows cell growth. J. Biol. Chem. 294, 8676–8689 (2019).

    PubMed  PubMed Central  Google Scholar 

  146. Sociali, G. et al. SIRT6 deacetylase activity regulates NAMPT activity and NAD(P)(H) pools in cancer cells. FASEB J. 33, 3704–3717 (2019).

    CAS  PubMed  Google Scholar 

  147. Mao, Z. et al. SIRT6 promotes DNA repair under stress by activating PARP1. Science 332, 1443–1446 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  148. Anderson, R. M. et al. Manipulation of a nuclear NAD+ salvage pathway delays aging without altering steady-state NAD+ levels. J. Biol. Chem. 277, 18881–18890 (2002).

    CAS  PubMed  Google Scholar 

  149. Araki, T., Sasaki, Y. & Milbrandt, J. Increased nuclear NAD biosynthesis and SIRT1 activation prevent axonal degeneration. Science 305, 1010–1013 (2004).

    CAS  PubMed  Google Scholar 

  150. Bass, J. & Lazar, M. A. Circadian time signatures of fitness and disease. Science 354, 994–999 (2016).

    CAS  PubMed  Google Scholar 

  151. Nakahata, Y., Sahar, S., Astarita, G., Kaluzova, M. & Sassone-Corsi, P. Circadian control of the NAD+ salvage pathway by CLOCK-SIRT1. Science 324, 654–657 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Ramsey, K. M. et al. Circadian clock feedback cycle through NAMPT-mediated NAD+ biosynthesis. Science 324, 651–654 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  153. Feng, D. & Lazar, M. A. Clocks, metabolism, and the epigenome. Mol. Cell 47, 158–167 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  154. Tahiliani, M. et al. Conversion of 5-methylcytosine to 5-hydroxymethylcytosine in mammalian DNA by MLL partner TET1. Science 324, 930–935 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Shi, Y. et al. Histone demethylation mediated by the nuclear amine oxidase homolog LSD1. Cell 119, 941–953 (2004).

    CAS  PubMed  Google Scholar 

  156. Tsukada, Y. et al. Histone demethylation by a family of JmjC domain-containing proteins. Nature 439, 811–816 (2006).

    CAS  PubMed  Google Scholar 

  157. Shim, E.-H. et al. L-2-Hydroxyglutarate: an epigenetic modifier and putative oncometabolite in renal cancer. Cancer Discov. 4, 1290–1298 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  158. Sciacovelli, M. & Frezza, C. Oncometabolites: unconventional triggers of oncogenic signalling cascades. Free Radic. Biol. Med. 100, 175–181 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  159. Nagaraj, R. et al. Nuclear localization of mitochondrial TCA cycle enzymes as a critical step in mammalian zygotic genome activation. Cell 168, 210–223.e11 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  160. Jiang, Y. et al. Local generation of fumarate promotes DNA repair through inhibition of histone H3 demethylation. Nat. Cell Biol. 17, 1158–1168 (2015). This study shows that the DNA-damage response activates nuclear fumarase in a specific local manner, thus inhibiting KDM2B histone demethylase activity and increasing H3K36me2 at the site of damage.

    CAS  PubMed  PubMed Central  Google Scholar 

  161. Wang, Y. et al. KAT2A coupled with the α-KGDH complex acts as a histone H3 succinyltransferase. Nature 552, 273–277 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  162. Li, S. et al. Serine and SAM responsive complex SESAME regulates histone modification crosstalk by sensing cellular metabolism. Mol. Cell 60, 408–421 (2015). This study identifies a nuclear multienzyme complex consisting of subcomponents for the simultaneous generation of ATP, SAMe and acetyl-CoA in the proximity of histone kinases, methyltransferases and acetyltransferases, thus providing the local substrate for chromatin modifiers.

    CAS  PubMed  Google Scholar 

  163. Mews, P. et al. Acetyl-CoA synthetase regulates histone acetylation and hippocampal memory. Nature 546, 381–386 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  164. Zhang, T. et al. Enzymes in the NAD+ salvage pathway regulate SIRT1 activity at target gene promoters. J. Biol. Chem. 284, 20408–20417 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  165. Zhang, T. et al. Regulation of poly(ADP-ribose) polymerase-1-dependent gene expression through promoter-directed recruitment of a nuclear NAD+ synthase. J. Biol. Chem. 287, 12405–12416 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  167. Gibson, B. A. et al. Organization of chromatin by intrinsic and regulated phase separation. Cell 179, 470–484.e21 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  168. Prouteau, M. & Loewith, R. Regulation of cellular metabolism through phase separation of enzymes. Biomolecules 8, 160 (2018).

    PubMed Central  Google Scholar 

  169. Hämäläinen, R. H. et al. Defects in mtDNA replication challenge nuclear genome stability through nucleotide depletion and provide a unifying mechanism for mouse progerias. Nat. Metab. 1, 958–965 (2019). This study shows that a defect in mitochondrial nucleotide pools is translated to the nuclear compartment, thereby leading to genomic instability.

    PubMed  Google Scholar 

  170. Mews, P. et al. Alcohol metabolism contributes to brain histone acetylation. Nature 574, 717–721 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  171. Horvath, S. DNA methylation age of human tissues and cell types. Genome Biol. 14, R115 (2013).

    PubMed  PubMed Central  Google Scholar 

  172. Reid, M. A. et al. Serine synthesis through PHGDH coordinates nucleotide levels by maintaining central carbon metabolism. Nat. Commun. 9, 5442 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  173. Martínez-Reyes, I. & Chandel, N. S. Mitochondrial one-carbon metabolism maintains redox balance during hypoxia. Cancer Discov. 4, 1371–1373 (2014).

    PubMed  PubMed Central  Google Scholar 

  174. Kottakis, F. et al. LKB1 loss links serine metabolism to DNA methylation and tumorigenesis. Nature 539, 390–395 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  175. Farber, S. & Diamond, L. K. Temporary remissions in acute leukemia in children produced by folic acid antagonist, 4-aminopteroyl-glutamic acid. N. Engl. J. Med. 238, 787–793 (1948).

    CAS  PubMed  Google Scholar 

  176. Gao, X. et al. Dietary methionine influences therapy in mouse cancer models and alters human metabolism. Nature 572, 397–401 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  177. Parkhitko, A. A., Jouandin, P., Mohr, S. E. & Perrimon, N. Methionine metabolism and methyltransferases in the regulation of aging and lifespan extension across species. Aging Cell 18, e13034 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. McDonald, O. G. et al. Epigenomic reprogramming during pancreatic cancer progression links anabolic glucose metabolism to distant metastasis. Nat. Genet. 49, 367–376 (2017). This study identifies a coevolved dependency of pancreatic ductal adenocarcinoma–derived metastasis on the oxidative branch of the PPP and the demethylation of large stretches of heterochromatin.

    CAS  PubMed  PubMed Central  Google Scholar 

  179. Gu, X. et al. SAMTOR is an S-adenosylmethionine sensor for the mTORC1 pathway. Science 358, 813–818 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  180. Ye, C. et al. Demethylation of the protein phosphatase PP2A promotes demethylation of histones to enable their function as a methyl group sink. Mol. Cell 73, 1115–1126.e6 (2019). This study identifies PP2A as a SAMe sensor capable of stimulating histone demethylation after SAMe starvation.

    CAS  PubMed  PubMed Central  Google Scholar 

  181. Dann, S. G. et al. Reciprocal regulation of amino acid import and epigenetic state through Lat1 and EZH2. EMBO J. 34, 1773–1785 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Pan, P. W. et al. Structure and biochemical functions of SIRT6. J. Biol. Chem. 286, 14575–14587 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Feldman, J. L., Baeza, J. & Denu, J. M. Activation of the protein deacetylase SIRT6 by long-chain fatty acids and widespread deacylation by mammalian sirtuins. J. Biol. Chem. 288, 31350–31356 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  184. Kim, H.-S. et al. Hepatic-specific disruption of SIRT6 in mice results in fatty liver formation due to enhanced glycolysis and triglyceride synthesis. Cell Metab. 12, 224–236 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  185. Mostoslavsky, R. et al. Genomic instability and aging-like phenotype in the absence of mammalian SIRT6. Cell 124, 315–329 (2006).

    CAS  PubMed  Google Scholar 

  186. Kugel, S. et al. SIRT6 suppresses pancreatic cancer through control of Lin28b. Cell 165, 1401–1415 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  187. Tasselli, L. et al. SIRT6 deacetylates H3K18ac at pericentric chromatin to prevent mitotic errors and cellular senescence. Nat. Struct. Mol. Biol. 23, 434–440 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  188. Ferriero, R. et al. Pyruvate dehydrogenase complex and lactate dehydrogenase are targets for therapy of acute liver failure. J. Hepatol. 69, 325–335 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  189. Attar, N. et al. The histone H3–H4 tetramer is a copper reductase enzyme. Science 369, 59–64 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  190. Machnicka, M. A. et al. MODOMICS: a database of RNA modification pathways: the 2013 update. Nucleic Acids Res. 41, D262–D267 (2013).

    CAS  PubMed  Google Scholar 

  191. Helm, M. & Motorin, Y. Detecting RNA modifications in the epitranscriptome: predict and validate. Nat. Rev. Genet. 18, 275–291 (2017).

    CAS  PubMed  Google Scholar 

  192. Frye, M., Harada, B. T., Behm, M. & He, C. RNA modifications modulate gene expression during development. Science 361, 1346–1349 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  193. Maddocks, O. D. K., Labuschagne, C. F., Adams, P. D. & Vousden, K. H. Serine metabolism supports the methionine cycle and DNA/RNA methylation through de novo ATP synthesis in cancer cells. Mol. Cell 61, 210–221 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  194. Laxman, S. et al. Sulfur amino acids regulate translational capacity and metabolic homeostasis through modulation of tRNA thiolation. Cell 154, 416–429 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  195. van der Wijst, M. G. P. & Rots, M. G. Mitochondrial epigenetics: an overlooked layer of regulation? Trends Genet. 31, 353–356 (2015).

    PubMed  Google Scholar 

  196. Bellizzi, D. et al. The control region of mitochondrial DNA shows an unusual CpG and non-CpG methylation pattern. DNA Res. 20, 537–547 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  197. Infantino, V. et al. Impairment of methyl cycle affects mitochondrial methyl availability and glutathione level in Down’s syndrome. Mol. Genet. Metab. 102, 378–382 (2011).

    CAS  PubMed  Google Scholar 

  198. Shock, L. S., Thakkar, P. V., Peterson, E. J., Moran, R. G. & Taylor, S. M. DNA methyltransferase 1, cytosine methylation, and cytosine hydroxymethylation in mammalian mitochondria. Proc. Natl Acad. Sci. USA 108, 3630–3635 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  199. Chatterjee, A. et al. MOF acetyl transferase regulates transcription and respiration in mitochondria. Cell 167, 722–738.e23 (2016).

    CAS  PubMed  Google Scholar 

  200. Montanari, A., Leo, M., De Luca, V., Filetici, P. & Francisci, S. Gcn5 histone acetyltransferase is present in the mitoplasts. Biol. Open 8, bio041244 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  201. King, G. A. et al. Acetylation and phosphorylation of human TFAM regulate TFAM-DNA interactions via contrasting mechanisms. Nucleic Acids Res. 46, 3633–3642 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  202. Dai, Z., Mentch, S. J., Gao, X., Nichenametla, S. N. & Locasale, J. W. Methionine metabolism influences genomic architecture and gene expression through H3K4me3 peak width. Nat. Commun. 9, 1955 (2018).

    PubMed  PubMed Central  Google Scholar 

  203. Shimazu, T. et al. Suppression of oxidative stress by β-hydroxybutyrate, an endogenous histone deacetylase inhibitor. Science 339, 211–214 (2013).

    CAS  PubMed  Google Scholar 

  204. Chriett, S. et al. Prominent action of butyrate over β-hydroxybutyrate as histone deacetylase inhibitor, transcriptional modulator and anti-inflammatory molecule. Sci. Rep. 9, 742 (2019).

    PubMed  PubMed Central  Google Scholar 

  205. Donohoe, D. R. et al. A gnotobiotic mouse model demonstrates that dietary fiber protects against colorectal tumorigenesis in a microbiota- and butyrate-dependent manner. Cancer Discov. 4, 1387–1397 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  206. Iida, N. et al. Commensal bacteria control cancer response to therapy by modulating the tumor microenvironment. Science 342, 967–970 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

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Acknowledgements

We thank members of the laboratory of R.M. for helpful discussions. Research in the laboratory of R.M. is supported by US NIH grants (R33ES025638, R01GM128448, R21ES027931 and R01CA235412) and CDMRP (W81XWH-17-1-0517). R.M. is supported as the Laurel Schwartz Endowed Professor of Oncology at the MGH Cancer Center/Harvard Medical School. R.B. is supported by postdoctoral fellowship LT-000311/2019-l from the Human Frontier Science Program. G.G.S. is supported by an MGH Cancer Center Excellence Award.

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R.M., G.G.S. and R.B. wrote, revised and approved the final version; R.B. prepared the figures.

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Correspondence to Raul Mostoslavsky.

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Boon, R., Silveira, G.G. & Mostoslavsky, R. Nuclear metabolism and the regulation of the epigenome. Nat Metab 2, 1190–1203 (2020). https://doi.org/10.1038/s42255-020-00285-4

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