Review Article | Published:

Mitochondrial function — gatekeeper of intestinal epithelial cell homeostasis

Nature Reviews Gastroenterology & Hepatologyvolume 15pages497516 (2018) | Download Citation


The intestinal epithelium is a multicellular interface in close proximity to a dense microbial milieu that is completely renewed every 3–5 days. Pluripotent stem cells reside at the crypt, giving rise to transient amplifying cells that go through continuous steps of proliferation, differentiation and finally anoikis (a form of programmed cell death) while migrating upwards to the villus tip. During these cellular transitions, intestinal epithelial cells (IECs) possess distinct metabolic identities reflected by changes in mitochondrial activity. Mitochondrial function emerges as a key player in cell fate decisions and in coordinating cellular metabolism, immunity, stress responses and apoptosis. Mediators of mitochondrial signalling include molecules such as ATP and reactive oxygen species and interrelate with pathways such as the mitochondrial unfolded protein response (MT-UPR) and AMP kinase signalling, in turn affecting cell cycle progression and stemness. Alterations in mitochondrial function and MT-UPR activation are integral aspects of pathologies, including IBD and cancer. Mitochondrial signalling and concomitant changes in metabolism contribute to intestinal homeostasis and regulate IEC dedifferentiation–differentiation programmes in the context of diseases, suggesting that mitochondrial function as a cellular checkpoint critically contributes to disease outcome. This Review highlights mitochondrial function and MT-UPR signalling in epithelial cell stemness, differentiation and lineage commitment and illustrates mitochondrial function in intestinal diseases.

Key points

  • The intestinal epithelium is a constantly renewing monolayer of cells undergoing continuous steps of proliferation, differentiation and finally cell death, representing an excellent model system to study stem cell regulation.

  • Intestinal epithelial cells (IECs) are key players in intestinal diseases such as IBD and colorectal cancer (CRC), constituting a dynamic interface between microbiota and host.

  • Mitochondrial function and metabolism determine and regulate IEC properties, such as differentiation status and proliferation.

  • Mitochondrial unfolded protein response (MT-UPR) coordinates mitochondrial function, metabolism and cellular phenotype and is activated in various diseases, including IBD and CRC.

  • MT-UPR might act as a sensor of the luminal and host environment, orchestrating epithelial tissue responses.

  • Determining the proliferative and regenerative capacity of IECs, the MT-UPR constitutes an attractive target for future therapeutic approaches for intestinal diseases.

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  1. 1.

    Gribble, F. M. & Reimann, F. Signalling in the gut endocrine axis. Physiol. Behav. 176, 183–188 (2017).

  2. 2.

    Peterson, L. W. & Artis, D. Intestinal epithelial cells: regulators of barrier function and immune homeostasis. Nat. Rev. Immunol. 14, 141–153 (2014).

  3. 3.

    Rath, E. & Haller, D. Inflammation and cellular stress: a mechanistic link between immune-mediated and metabolically driven pathologies. Eur. J. Nutr. 50, 219–233 (2011).

  4. 4.

    Barker, N. Adult intestinal stem cells: critical drivers of epithelial homeostasis and regeneration. Nat. Rev. Mol. Cell Biol. 15, 19–33 (2014).

  5. 5.

    Potten, C. S. A comprehensive study of the radiobiological response of the murine (BDF1) small intestine. Int. J. Radiat. Biol. 58, 925–973 (1990).

  6. 6.

    Leushacke, M. & Barker, N. Ex vivo culture of the intestinal epithelium: strategies and applications. Gut 63, 1345–1354 (2014).

  7. 7.

    Bellafante, E. et al. PGC-1beta promotes enterocyte lifespan and tumorigenesis in the intestine. Proc. Natl Acad. Sci. USA 111, E4523–E4531 (2014).

  8. 8.

    D’Errico, I. et al. Peroxisome proliferator-activated receptor-gamma coactivator 1-alpha (PGC1alpha) is a metabolic regulator of intestinal epithelial cell fate. Proc. Natl Acad. Sci. USA 108, 6603–6608 (2011). This paper identified PGC1α as driver of mitochondrial biogenesis and respiration in IECs and the balance of ROS accumulation and antioxidant enzyme activities as metabolic regulator of IEC fate.

  9. 9.

    Rodriguez-Colman, M. J. et al. Interplay between metabolic identities in the intestinal crypt supports stem cell function. Nature 543, 424–427 (2017). This research shows that Lgr5 + CBCs and Paneth cells display different metabolic programmes, with Paneth cells supporting intestinal stem cell function by providing lactate to sustain the enhanced mitochondrial oxidative phosphorylation in the Lgr5 + CBCs.

  10. 10.

    Sethi, J. K. & Vidal-Puig, A. Wnt signalling and the control of cellular metabolism. Biochem. J. 427, 1–17 (2010).

  11. 11.

    Berger, E. et al. Mitochondrial function controls intestinal epithelial stemness and proliferation. Nat. Commun. 7, 13171 (2016). This paper demonstrates that mitochondrial function has a critical role in maintaining intestinal stemness and homeostasis and links mitochondrial unfolded protein response to mitochondrial dysfunction.

  12. 12.

    Heijmans, J. et al. ER stress causes rapid loss of intestinal epithelial stemness through activation of the unfolded protein response. Cell Rep. 3, 1128–1139 (2013).

  13. 13.

    Ohashi, W. et al. Zinc transporter SLC39A7/ZIP7 promotes intestinal epithelial self-renewal by resolving ER stress. PLoS Genet. 12, e1006349 (2016).

  14. 14.

    Rath, E. & Haller, D. Mitochondria at the interface between danger signaling and metabolism: role of unfolded protein responses in chronic inflammation. Inflamm. Bowel Dis. 18, 1364–1377 (2012).

  15. 15.

    Ryan, M. T. & Hoogenraad, N. J. Mitochondrial-nuclear communications. Annu. Rev. Biochem. 76, 701–722 (2007).

  16. 16.

    Cabibi, D. et al. CD1A-positive cells and HSP60 (HSPD1) levels in keratoacanthoma and squamous cell carcinoma. Cell Stress Chaperones 21, 131–137 (2016).

  17. 17.

    Tong, W. W., Tong, G. H., Kong, H. & Liu, Y. The tumor promoting roles of HSP60 and HIF2alpha in gastric cancer cells. Tumour Biol. 37, 9849–9854 (2016).

  18. 18.

    Rappa, F. et al. Quantitative patterns of Hsps in tubular adenoma compared with normal and tumor tissues reveal the value of Hsp10 and Hsp60 in early diagnosis of large bowel cancer. Cell Stress Chaperones 21, 927–933 (2016).

  19. 19.

    Hwang, Y. J. et al. Expression of heat shock protein 60 kDa is upregulated in cervical cancer. Yonsei Med. J. 50, 399–406 (2009).

  20. 20.

    Castilla, C. et al. Immunohistochemical expression of Hsp60 correlates with tumor progression and hormone resistance in prostate cancer. Urology 76, 1017.e1–1017.e6 (2010).

  21. 21.

    Hamrita, B. et al. Identification of tumor antigens that elicit a humoral immune response in breast cancer patients’ sera by serological proteome analysis (SERPA). Clin. Chim. Acta 393, 95–102 (2008).

  22. 22.

    Piselli, P. et al. Different expression of CD44, ICAM-1, and HSP60 on primary tumor and metastases of a human pancreatic carcinoma growing in scid mice. Anticancer Res. 20, 825–831 (2000).

  23. 23.

    De Cecco, L. et al. Gene expression profiling of advanced ovarian cancer: characterization of a molecular signature involving fibroblast growth factor 2. Oncogene 23, 8171–8183 (2004).

  24. 24.

    Rath, E. et al. Induction of dsRNA-activated protein kinase links mitochondrial unfolded protein response to the pathogenesis of intestinal inflammation. Gut 61, 1269–1278 (2012). This study identifies dsRNA-activated protein kinase as mediator of the MT-UPR and demonstrates MT-UPR activation in intestinal epithelial cells from patients with IBD and from mouse models of intestinal inflammation.

  25. 25.

    Scorrano, L. Keeping mitochondria in shape: a matter of life and death. Eur. J. Clin. Invest. 43, 886–893 (2013).

  26. 26.

    Brookes, P. S. et al. Control of mitochondrial respiration by NO*, effects of low oxygen and respiratory state. J. Biol. Chem. 278, 31603–31609 (2003).

  27. 27.

    Zong, H. et al. AMP kinase is required for mitochondrial biogenesis in skeletal muscle in response to chronic energy deprivation. Proc. Natl Acad. Sci. USA 99, 15983–15987 (2002).

  28. 28.

    Kelly, D. P. & Scarpulla, R. C. Transcriptional regulatory circuits controlling mitochondrial biogenesis and function. Genes Dev. 18, 357–368 (2004).

  29. 29.

    Schell, J. C. et al. Control of intestinal stem cell function and proliferation by mitochondrial pyruvate metabolism. Nat. Cell Biol. 19, 1027–1036 (2017). This study shows that mitochondrial pyruvate metabolism is important for ISC maintenance and that limiting mitochondrial pyruvate metabolism is necessary and sufficient to maintain ISC proliferation.

  30. 30.

    Bukau, B., Weissman, J. & Horwich, A. Molecular chaperones and protein quality control. Cell 125, 443–451 (2006).

  31. 31.

    Ron, D. & Walter, P. Signal integration in the endoplasmic reticulum unfolded protein response. Nat. Rev. Mol. Cell Biol. 8, 519–529 (2007).

  32. 32.

    Aldridge, J. E., Horibe, T. & Hoogenraad, N. J. Discovery of genes activated by the mitochondrial unfolded protein response (mtUPR) and cognate promoter elements. PLoS ONE 2, e874 (2007).

  33. 33.

    Zhao, Q. et al. A mitochondrial specific stress response in mammalian cells. EMBO J. 21, 4411–4419 (2002). This paper was the first description and characterization of the mitochondrial unfolded protein stress response in mammalian cells.

  34. 34.

    Martinus, R. D. et al. Selective induction of mitochondrial chaperones in response to loss of the mitochondrial genome. Eur. J. Biochem. 240, 98–103 (1996).

  35. 35.

    Nargund, A. M., Fiorese, C. J., Pellegrino, M. W., Deng, P. & Haynes, C. M. Mitochondrial and nuclear accumulation of the transcription factor ATFS-1 promotes OXPHOS recovery during the UPR(mt). Mol. Cell 58, 123–133 (2015). This study identifies the transcription factor ATFS-1 as important factor for the recovery of the OXPHOS machinery under MT-UPR conditions, next to regulating genes involved in proteostasis in C. elegans.

  36. 36.

    He, C., Hart, P. C., Germain, D. & Bonini, M. G. SOD2 and the mitochondrial UPR: partners regulating cellular phenotypic transitions. Trends Biochem. Sci. 41, 568–577 (2016).

  37. 37.

    Horibe, T. & Hoogenraad, N. J. The chop gene contains an element for the positive regulation of the mitochondrial unfolded protein response. PLoS ONE 2, e835 (2007).

  38. 38.

    Pellegrino, M. W., Nargund, A. M. & Haynes, C. M. Signaling the mitochondrial unfolded protein response. Biochim. Biophys. Acta 1833, 410–416 (2013).

  39. 39.

    Yano, M. ABCB10 depletion reduces unfolded protein response in mitochondria. Biochem. Biophys. Res. Commun. 486, 465–469 (2017).

  40. 40.

    Jovaisaite, V., Mouchiroud, L. & Auwerx, J. The mitochondrial unfolded protein response, a conserved stress response pathway with implications in health and disease. J. Exp. Biol. 217, 137–143 (2014).

  41. 41.

    Ishikawa, F. et al. Gene expression profiling identifies a role for CHOP during inhibition of the mitochondrial respiratory chain. J. Biochem. 146, 123–132 (2009).

  42. 42.

    Quiros, P. M. et al. Multi-omics analysis identifies ATF4 as a key regulator of the mitochondrial stress response in mammals. J. Cell Biol. 216, 2027–2045 (2017). This study identifies activating transcription factor 4 as the main regulator of the mammalian response towards mitochondrial stress.

  43. 43.

    Fiorese, C. J. et al. The transcription factor ATF5 mediates a mammalian mitochondrial UPR. Curr. Biol. 26, 2037–2043 (2016).

  44. 44.

    Baker, B. M., Nargund, A. M., Sun, T. & Haynes, C. M. Protective coupling of mitochondrial function and protein synthesis via the eIF2alpha kinase GCN-2. PLoS Genet. 8, e1002760 (2012).

  45. 45.

    Hood, D. A., Irrcher, I., Ljubicic, V. & Joseph, A. M. Coordination of metabolic plasticity in skeletal muscle. J. Exp. Biol. 209, 2265–2275 (2006).

  46. 46.

    Lin, Y. F. & Haynes, C. M. Metabolism and the UPR(mt). Mol. Cell 61, 677–682 (2016).

  47. 47.

    Kim, H. E. et al. Lipid biosynthesis coordinates a mitochondrial-to-cytosolic stress response. Cell 166, 1539–1552.e16 (2016). This paper demonstrates a conserved mechanism linking mitochondrial protein homeostasis and the cytosolic folding environment through changes in lipid homeostasis.

  48. 48.

    Zhang, Y. et al. SIRT3 and SIRT5 regulate the enzyme activity and cardiolipin binding of very long-chain acyl-CoA dehydrogenase. PLoS ONE 10, e0122297 (2015).

  49. 49.

    Berendzen, K. M. et al. Neuroendocrine coordination of mitochondrial stress signaling and proteostasis. Cell 166, 1553–1563.e10 (2016). This research shows that neuronal mitochondrial stress induces a cell-non-autonomous MT-UPR, eliciting a global induction of MT-UPR-specific changes, thereby affecting whole-animal physiology.

  50. 50.

    Durieux, J., Wolff, S. & Dillin, A. The cell-non-autonomous nature of electron transport chain-mediated longevity. Cell 144, 79–91 (2011).

  51. 51.

    Murphy, M. P. How mitochondria produce reactive oxygen species. Biochem. J. 417, 1–13 (2009).

  52. 52.

    Muller, F. L., Liu, Y. & Van Remmen, H. Complex III releases superoxide to both sides of the inner mitochondrial membrane. J. Biol. Chem. 279, 49064–49073 (2004).

  53. 53.

    Lustgarten, M. S. et al. Complex I generated, mitochondrial matrix-directed superoxide is released from the mitochondria through voltage dependent anion channels. Biochem. Biophys. Res. Commun. 422, 515–521 (2012).

  54. 54.

    Tatsuta, T., Scharwey, M. & Langer, T. Mitochondrial lipid trafficking. Trends Cell Biol. 24, 44–52 (2014).

  55. 55.

    Cadenas, E. & Davies, K. J. Mitochondrial free radical generation, oxidative stress, and aging. Free Radic. Biol. Med. 29, 222–230 (2000).

  56. 56.

    Riemer, J., Schwarzlander, M., Conrad, M. & Herrmann, J. M. Thiol switches in mitochondria: operation and physiological relevance. Biol. Chem. 396, 465–482 (2015).

  57. 57.

    Smeitink, J. A., Zeviani, M., Turnbull, D. M. & Jacobs, H. T. Mitochondrial medicine: a metabolic perspective on the pathology of oxidative phosphorylation disorders. Cell Metab. 3, 9–13 (2006).

  58. 58.

    Chandel, N. S. et al. Mitochondrial reactive oxygen species trigger hypoxia-induced transcription. Proc. Natl Acad. Sci. USA 95, 11715–11720 (1998).

  59. 59.

    Bell, E. L. et al. The Qo site of the mitochondrial complex III is required for the transduction of hypoxic signaling via reactive oxygen species production. J. Cell Biol. 177, 1029–1036 (2007).

  60. 60.

    Raimundo, N. et al. Mitochondrial stress engages E2F1 apoptotic signaling to cause deafness. Cell 148, 716–726 (2012). This study demonstrates that mitochondrial stress and dysfunction engage specific cellular signalling cascades in the context of a human pathology caused by a mitochondrial DNA mutation.

  61. 61.

    Formentini, L., Sanchez-Arago, M., Sanchez-Cenizo, L. & Cuezva, J. M. The mitochondrial ATPase inhibitory factor 1 triggers a ROS-mediated retrograde prosurvival and proliferative response. Mol. Cell 45, 731–742 (2012).

  62. 62.

    Raimundo, N. Mitochondrial pathology: stress signals from the energy factory. Trends Mol. Med. 20, 282–292 (2014).

  63. 63.

    Hamanaka, R. B. et al. Mitochondrial reactive oxygen species promote epidermal differentiation and hair follicle development. Sci. Signal. 6, ra8 (2013).

  64. 64.

    West, A. P. et al. TLR signalling augments macrophage bactericidal activity through mitochondrial ROS. Nature 472, 476–480 (2011).

  65. 65.

    Schroeder, E. A., Raimundo, N. & Shadel, G. S. Epigenetic silencing mediates mitochondria stress-induced longevity. Cell Metab. 17, 954–964 (2013).

  66. 66.

    Zarse, K. et al. Impaired insulin/IGF1 signaling extends life span by promoting mitochondrial L-proline catabolism to induce a transient ROS signal. Cell Metab. 15, 451–465 (2012).

  67. 67.

    Senapedis, W. T., Kennedy, C. J., Boyle, P. M. & Silver, P. A. Whole genome siRNA cell-based screen links mitochondria to Akt signaling network through uncoupling of electron transport chain. Mol. Biol. Cell 22, 1791–1805 (2011).

  68. 68.

    Lim, J. H., Lee, H. J., Ho Jung, M. & Song, J. Coupling mitochondrial dysfunction to endoplasmic reticulum stress response: a molecular mechanism leading to hepatic insulin resistance. Cell Signal. 21, 169–177 (2009).

  69. 69.

    Fukushima, K. & Fiocchi, C. Paradoxical decrease of mitochondrial DNA deletions in epithelial cells of active ulcerative colitis patients. Am. J. Physiol. Gastrointest. Liver Physiol. 286, G804–G813 (2004).

  70. 70.

    Haga, N. et al. Mitochondria regulate the unfolded protein response leading to cancer cell survival under glucose deprivation conditions. Cancer Sci. (2010).

  71. 71.

    Barker, N., van Oudenaarden, A. & Clevers, H. Identifying the stem cell of the intestinal crypt: strategies and pitfalls. Cell Stem Cell 11, 452–460 (2012).

  72. 72.

    Tan, S. & Barker, N. Epithelial stem cells and intestinal cancer. Semin. Cancer Biol. 32, 40–53 (2015).

  73. 73.

    Barriga, F. M. et al. Mex3a Marks a Slowly Dividing Subpopulation of Lgr5+ Intestinal Stem Cells. Cell Stem Cell 20, 801–816.e7 (2017).

  74. 74.

    Roth, S. et al. Paneth cells in intestinal homeostasis and tissue injury. PLoS ONE 7, e38965 (2012).

  75. 75.

    Nakanishi, Y. et al. Dclk1 distinguishes between tumor and normal stem cells in the intestine. Nat. Genet. 45, 98–103 (2013).

  76. 76.

    Tsubouchi, S. & Leblond, C. P. Migration and turnover of entero-endocrine and caveolated cells in the epithelium of the descending colon, as shown by radioautography after continuous infusion of 3H-thymidine into mice. Am. J. Anat. 156, 431–451 (1979).

  77. 77.

    Folmes, C. D., Dzeja, P. P., Nelson, T. J. & Terzic, A. Metabolic plasticity in stem cell homeostasis and differentiation. Cell Stem Cell 11, 596–606 (2012).

  78. 78.

    Zhang, J., Nuebel, E., Daley, G. Q., Koehler, C. M. & Teitell, M. A. Metabolic regulation in pluripotent stem cells during reprogramming and self-renewal. Cell Stem Cell 11, 589–595 (2012).

  79. 79.

    Xu, X. et al. Mitochondrial regulation in pluripotent stem cells. Cell Metab. 18, 325–332 (2013).

  80. 80.

    Cho, Y. M. et al. Dynamic changes in mitochondrial biogenesis and antioxidant enzymes during the spontaneous differentiation of human embryonic stem cells. Biochem. Biophys. Res. Commun. 348, 1472–1478 (2006). This study shows that differentiation of human embryonic stem cells is accompanied by dynamic changes in mitochondrial mass, ATP and ROS production, as well as antioxidant enzyme expressions.

  81. 81.

    Hom, J. R. et al. The permeability transition pore controls cardiac mitochondrial maturation and myocyte differentiation. Dev. Cell 21, 469–478 (2011).

  82. 82.

    Tormos, K. V. et al. Mitochondrial complex III ROS regulate adipocyte differentiation. Cell Metab. 14, 537–544 (2011).

  83. 83.

    Kasahara, A. et al. Mitochondrial fusion directs cardiomyocyte differentiation via calcineurin and Notch signaling. Science 342, 734–737 (2013).

  84. 84.

    Yuan, D. et al. Kupffer cell-derived tnf triggers cholangiocellular tumorigenesis through JNK due to chronic mitochondrial dysfunction and ROS. Cancer Cell 31, 771–789.e6 (2017).

  85. 85.

    Waldschmitt, N. et al. C/EBP homologous protein inhibits tissue repair in response to gut injury and is inversely regulated with chronic inflammation. Mucosal Immunol. 7, 1452–1466 (2014).

  86. 86.

    Chandel, N. S., Jasper, H., Ho, T. T. & Passegue, E. Metabolic regulation of stem cell function in tissue homeostasis and organismal ageing. Nat. Cell Biol. 18, 823–832 (2016).

  87. 87.

    Mohrin, M. et al. Stem cell aging. A mitochondrial UPR-mediated metabolic checkpoint regulates hematopoietic stem cell aging. Science 347, 1374–1377 (2015). This research identifies a regulatory branch of the mitochondrial unfolded protein response, which is coupled to cellular energy metabolism and proliferation, and determines hematopoietic stem cell quiescence and regenerative capacity.

  88. 88.

    Kobayashi, M. et al. The antioxidant defense system Keap1-Nrf2 comprises a multiple sensing mechanism for responding to a wide range of chemical compounds. Mol. Cell. Biol. 29, 493–502 (2009).

  89. 89.

    Li, Q. & Engelhardt, J. F. Interleukin-1beta induction of NFkappaB is partially regulated by H2O2-mediated activation of NFkappaB-inducing kinase. J. Biol. Chem. 281, 1495–1505 M511153200 (2006).

  90. 90.

    Rera, M. et al. Modulation of longevity and tissue homeostasis by the Drosophila PGC-1 homolog. Cell Metab. 14, 623–634 (2011).

  91. 91.

    Laplante, M. & Sabatini, D. M. mTOR signaling in growth control and disease. Cell 149, 274–293 (2012).

  92. 92.

    Jasper, H. & Jones, D. L. Metabolic regulation of stem cell behavior and implications for aging. Cell Metab. 12, 561–565 (2010).

  93. 93.

    Sampson, L. L., Davis, A. K., Grogg, M. W. & Zheng, Y. mTOR disruption causes intestinal epithelial cell defects and intestinal atrophy postinjury in mice. FASEB J. 30, 1263–1275 (2016).

  94. 94.

    Chen, T. et al. Rapamycin and other longevity-promoting compounds enhance the generation of mouse induced pluripotent stem cells. Aging Cell 10, 908–911 (2011).

  95. 95.

    Chen, C. et al. TSC-mTOR maintains quiescence and function of hematopoietic stem cells by repressing mitochondrial biogenesis and reactive oxygen species. J. Exp. Med. 205, 2397–2408 (2008).

  96. 96.

    Yilmaz, O. H. et al. mTORC1 in the Paneth cell niche couples intestinal stem-cell function to calorie intake. Nature 486, 490–495 (2012). This paper demonstrates that caloric restriction augments intestinal stem cell function via mTOR-signalling in Paneth cells to couple this process to organismal physiology.

  97. 97.

    Kaiko, G. E. et al. The colonic crypt protects stem cells from microbiota-derived metabolites. Cell 165, 1708–1720 (2016).

  98. 98.

    Donohoe, D. R. et al. The microbiome and butyrate regulate energy metabolism and autophagy in the mammalian colon. Cell Metab. 13, 517–526 (2011). This study shows that the microbiome via butyrate production ensures energy homeostasis in colonocytes, and that colonocytes from germ-free mice exhibit decreased oxidative phosphorylation, decreased expression of enzymes in the intermediary metabolism, and decreased ATP levels.

  99. 99.

    Stringari, C. et al. Metabolic trajectory of cellular differentiation in small intestine by Phasor Fluorescence Lifetime Microscopy of NADH. Sci. Rep. 2, 568 (2012).

  100. 100.

    Jeynes, B. J. & Altmann, G. G. A region of mitochondrial division in the epithelium of the small intestine of the rat. Anat. Rec. 182, 289–296 (1975).

  101. 101.

    Lin, J. E. et al. The hormone receptor GUCY2C suppresses intestinal tumor formation by inhibiting AKT signaling. Gastroenterology 138, 241–254 (2010).

  102. 102.

    Cristofaro, M. et al. Adenomatous polyposis coli (APC)-induced apoptosis of HT29 colorectal cancer cells depends on mitochondrial oxidative metabolism. Biochim. Biophys. Acta 1852, 1719–1728 (2015).

  103. 103.

    Wu, Z. et al. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC-1. Cell 98, 115–124 (1999).

  104. 104.

    Lin, J. et al. Transcriptional co-activator PGC-1 alpha drives the formation of slow-twitch muscle fibres. Nature 418, 797–801 (2002).

  105. 105.

    Kumar, N. et al. A YY1-dependent increase in aerobic metabolism is indispensable for intestinal organogenesis. Development 143, 3711–3722 (2016).

  106. 106.

    Blattler, S. M. et al. Defective mitochondrial morphology and bioenergetic function in mice lacking the transcription factor Yin Yang 1 in skeletal muscle. Mol. Cell. Biol. 32, 3333–3346 (2012).

  107. 107.

    Cunningham, J. T. et al. mTOR controls mitochondrial oxidative function through a YY1-PGC-1alpha transcriptional complex. Nature 450, 736–740 (2007).

  108. 108.

    Kluck, R. M., Bossy-Wetzel, E., Green, D. R. & Newmeyer, D. D. The release of cytochrome c from mitochondria: a primary site for Bcl-2 regulation of apoptosis. Science 275, 1132–1136 (1997).

  109. 109.

    Turan, A. & Mahmood, A. The profile of antioxidant systems and lipid peroxidation across the crypt-villus axis in rat intestine. Dig. Dis. Sci. 52, 1840–1844 (2007).

  110. 110.

    D’Errico, I. et al. Bax is necessary for PGC1alpha pro-apoptotic effect in colorectal cancer cells. Cell Cycle 10, 2937–2945 (2011).

  111. 111.

    Boohaker, R. J., Zhang, G., Carlson, A. L., Nemec, K. N. & Khaled, A. R. BAX supports the mitochondrial network, promoting bioenergetics in nonapoptotic cells. Am. J. Physiol. Cell Physiol. 300, C1466–1478 (2011).

  112. 112.

    McCarthy, N. Metabolism: a TIGAR tale. Nat. Rev. Cancer 13, 522 (2013).

  113. 113.

    Cheung, E. C. et al. TIGAR is required for efficient intestinal regeneration and tumorigenesis. Dev. Cell 25, 463–477 (2013).

  114. 114.

    Cheung, E. C. et al. Opposing effects of TIGAR- and RAC1-derived ROS on Wnt-driven proliferation in the mouse intestine. Genes Dev. 30, 52–63 (2016). This paper shows that two key targets in the Wnt pathway function to integrate pro-proliferative and anti-proliferative effects of ROS, modulating cell proliferation.

  115. 115.

    O’Hara, A. M. & Shanahan, F. The gut flora as a forgotten organ. EMBO Rep. 7, 688–693 (2006).

  116. 116.

    Macpherson, A. J. & McCoy, K. D. Standardised animal models of host microbial mutualism. Mucosal Immunol. 8, 476–486 (2015).

  117. 117.

    Chu, H. & Mazmanian, S. K. Innate immune recognition of the microbiota promotes host-microbial symbiosis. Nat. Immunol. 14, 668–675 (2013).

  118. 118.

    Haller, D., Russo, M. P., Sartor, R. B. & Jobin, C. IKK beta and phosphatidylinositol 3-kinase/Akt participate in non-pathogenic Gram-negative enteric bacteria-induced RelA phosphorylation and NF-kappa B activation in both primary and intestinal epithelial cell lines. J. Biol. Chem. 277, 38168–38178 (2002).

  119. 119.

    Haller, D. et al. Transforming growth factor-beta 1 inhibits non-pathogenic Gram negative bacteria-induced NF-kappa B recruitment to the interleukin-6 gene promoter in intestinal epithelial cells through modulation of histone acetylation. J. Biol. Chem. 278, 23851–23860 (2003).

  120. 120.

    Nenci, A. et al. Epithelial NEMO links innate immunity to chronic intestinal inflammation. Nature 446, 557–561 (2007).

  121. 121.

    Rakoff-Nahoum, S., Paglino, J., Eslami-Varzaneh, F., Edberg, S. & Medzhitov, R. Recognition of commensal microflora by toll-like receptors is required for intestinal homeostasis. Cell 118, 229–241 (2004).

  122. 122.

    Rakoff-Nahoum, S. & Medzhitov, R. Regulation of spontaneous intestinal tumorigenesis through the adaptor protein MyD88. Science 317, 124–127 (2007). This study identifies bacterial sensing and innate immune signalling as important contributors to spontaneous and carcinogen-induced intestinal tumour development.

  123. 123.

    Koh, A., De Vadder, F., Kovatcheva-Datchary, P. & Backhed, F. From dietary fiber to host physiology: short-chain fatty acids as key bacterial metabolites. Cell 165, 1332–1345 (2016).

  124. 124.

    Wahlstrom, A., Sayin, S. I., Marschall, H. U. & Backhed, F. Intestinal crosstalk between bile acids and microbiota and its impact on host metabolism. Cell Metab. 24, 41–50 (2016).

  125. 125.

    Lamas, B. et al. CARD9 impacts colitis by altering gut microbiota metabolism of tryptophan into aryl hydrocarbon receptor ligands. Nat. Med. 22, 598–605 (2016).

  126. 126.

    Li, Y. et al. Exogenous stimuli maintain intraepithelial lymphocytes via aryl hydrocarbon receptor activation. Cell 147, 629–640 (2011).

  127. 127.

    Bjeldanes, L. F., Kim, J. Y., Grose, K. R., Bartholomew, J. C. & Bradfield, C. A. Aromatic hydrocarbon responsiveness-receptor agonists generated from indole-3-carbinol in vitro and in vivo: comparisons with 2,3,7,8-tetrachlorodibenzo-p-dioxin. Proc. Natl Acad. Sci. USA 88, 9543–9547 (1991).

  128. 128.

    Hwang, H. J. et al. Mitochondrial-targeted aryl hydrocarbon receptor and the impact of 2,3,7,8-tetrachlorodibenzo-p-dioxin on cellular respiration and the mitochondrial proteome. Toxicol. Appl. Pharmacol. 304, 121–132 (2016).

  129. 129.

    Buffie, C. G. et al. Precision microbiome reconstitution restores bile acid mediated resistance to Clostridium difficile. Nature 517, 205–208 (2015).

  130. 130.

    O’Keefe, S. J. Diet, microorganisms and their metabolites, and colon cancer. Nat. Rev. Gastroenterol. Hepatol. 13, 691–706 (2016).

  131. 131.

    Wichmann, A. et al. Microbial modulation of energy availability in the colon regulates intestinal transit. Cell Host Microbe 14, 582–590 (2013).

  132. 132.

    Greiner, T. U. & Backhed, F. Microbial regulation of GLP-1 and L-cell biology. Mol. Metab. 5, 753–758 (2016).

  133. 133.

    Donohoe, D. R., Wali, A., Brylawski, B. P. & Bultman, S. J. Microbial regulation of glucose metabolism and cell-cycle progression in mammalian colonocytes. PLoS ONE 7, e46589 (2012).

  134. 134.

    Kien, C. L. et al. Cecal infusion of butyrate increases intestinal cell proliferation in piglets. J. Nutr. 137, 916–922 (2007).

  135. 135.

    Guzman, J. R., Conlin, V. S. & Jobin, C. Diet, microbiome, and the intestinal epithelium: an essential triumvirate? Biomed. Res. Int. 2013, 425146 (2013).

  136. 136.

    Zietek, T. & Rath, E. Inflammation meets metabolic disease: gut feeling mediated by GLP-1. Front. Immunol. 7, 154 (2016).

  137. 137.

    Ni, J., Wu, G. D., Albenberg, L. & Tomov, V. T. Gut microbiota and IBD: causation or correlation? Nat. Rev. Gastroenterol. Hepatol. 14, 573–584 (2017).

  138. 138.

    Huda-Faujan, N. et al. The impact of the level of the intestinal short chain fatty acids in inflammatory bowel disease patients versus healthy subjects. Open Biochem. J. 4, 53–58 (2010).

  139. 139.

    Sun, M., Wu, W., Liu, Z. & Cong, Y. Microbiota metabolite short chain fatty acids, GPCR, and inflammatory bowel diseases. J. Gastroenterol. 52, 1–8 (2017).

  140. 140.

    Fuentes, S. et al. Reset of a critically disturbed microbial ecosystem: faecal transplant in recurrent Clostridium difficile infection. ISME J. 8, 1621–1633 (2014).

  141. 141.

    Gribble, F. M. & Reimann, F. Enteroendocrine cells: chemosensors in the intestinal epithelium. Annu. Rev. Physiol. 78, 277–299 (2016).

  142. 142.

    Cipriani, S. et al. The bile acid receptor GPBAR-1 (TGR5) modulates integrity of intestinal barrier and immune response to experimental colitis. PLoS ONE 6, e25637 (2011).

  143. 143.

    Neal, M. D. et al. Toll-like receptor 4 is expressed on intestinal stem cells and regulates their proliferation and apoptosis via the p53 up-regulated modulator of apoptosis. J. Biol. Chem. 287, 37296–37308 (2012).

  144. 144.

    Santaolalla, R. et al. TLR4 activates the beta-catenin pathway to cause intestinal neoplasia. PLoS ONE 8, e63298 (2013).

  145. 145.

    Nigro, G., Rossi, R., Commere, P. H., Jay, P. & Sansonetti, P. J. The cytosolic bacterial peptidoglycan sensor Nod2 affords stem cell protection and links microbes to gut epithelial regeneration. Cell Host Microbe 15, 792–798 (2014).

  146. 146.

    Kato, M. et al. The ROS-generating oxidase Nox1 is required for epithelial restitution following colitis. Exp. Anim. 65, 197–205 (2016).

  147. 147.

    Datta, A. et al. Mouse lung development and NOX1 induction during hyperoxia are developmentally regulated and mitochondrial ROS dependent. Am. J. Physiol. Lung Cell. Mol. Physiol. 309, L369–377 (2015).

  148. 148.

    Chiarugi, P. et al. Reactive oxygen species as essential mediators of cell adhesion: the oxidative inhibition of a FAK tyrosine phosphatase is required for cell adhesion. J. Cell Biol. 161, 933–944 (2003).

  149. 149.

    Ogier-Denis, E., Mkaddem, S. B. & Vandewalle, A. NOX enzymes and Toll-like receptor signaling. Semin. Immunopathol. 30, 291–300 (2008).

  150. 150.

    Jones, R. M., Mercante, J. W. & Neish, A. S. Reactive oxygen production induced by the gut microbiota: pharmacotherapeutic implications. Curr. Med. Chem. 19, 1519–1529 (2012).

  151. 151.

    Lee, S. R. et al. Reversible inactivation of the tumor suppressor PTEN by H2O2. J. Biol. Chem. 277, 20336–20342 (2002).

  152. 152.

    Rhodes, J. M. & Campbell, B. J. Inflammation and colorectal cancer: IBD-associated and sporadic cancer compared. Trends Mol. Med. 8, 10–16 (2002).

  153. 153.

    Miyoshi, H. et al. Prostaglandin E2 promotes intestinal repair through an adaptive cellular response of the epithelium. EMBO J. 36, 5–24 (2017). This study characterizes formation of metabolically distinct wound-associated epithelial cells via the PGE2-Ptger4 pathway by adaptive cellular reprogramming of the intestinal epithelium following intestinal injury.

  154. 154.

    Marino Gammazza, A. et al. Doxorubicin anti-tumor mechanisms include Hsp60 post-translational modifications leading to the Hsp60/p53 complex dissociation and instauration of replicative senescence. Cancer Lett. 385, 75–86 (2017).

  155. 155.

    Tsai, Y. P. et al. Interaction between HSP60 and beta-catenin promotes metastasis. Carcinogenesis 30, 1049–1057 (2009).

  156. 156.

    Molodecky, N. A. et al. Increasing incidence and prevalence of the inflammatory bowel diseases with time, based on systematic review. Gastroenterology 142, 46–54.e2 (2012).

  157. 157.

    Ng, S. C. et al. Worldwide incidence and prevalence of inflammatory bowel disease in the 21st century: a systematic review of population-based studies. Lancet 390, 2769–2778 (2018).

  158. 158.

    de Souza, H. S. P., Fiocchi, C. & Iliopoulos, D. The IBD interactome: an integrated view of aetiology, pathogenesis and therapy. Nat. Rev. Gastroenterol. Hepatol. 14, 739–749 (2017).

  159. 159.

    Gersemann, M., Stange, E. F. & Wehkamp, J. From intestinal stem cells to inflammatory bowel diseases. World J. Gastroenterol. 17, 3198–3203 (2011).

  160. 160.

    Roediger, W. E. The colonic epithelium in ulcerative colitis: an energy-deficiency disease? Lancet 2, 712–715 (1980).

  161. 161.

    Beltran, B. et al. Mitochondrial dysfunction, persistent oxidative damage, and catalase inhibition in immune cells of naive and treated Crohn’s disease. Inflamm. Bowel Dis. 16, 76–86 (2010).

  162. 162.

    Barrett, J. C. et al. Genome-wide association defines more than 30 distinct susceptibility loci for Crohn’s disease. Nat. Genet. 40, 955–962 (2008).

  163. 163.

    Yu, X. et al. Association of UCP2 -866 G/A polymorphism with chronic inflammatory diseases. Genes Immun. 10, 601–605 (2009).

  164. 164.

    Waller, S. et al. Evidence for association of OCTN genes and IBD5 with ulcerative colitis. Gut 55, 809–814 (2006).

  165. 165.

    Rinaldo, P., Matern, D. & Bennett, M. J. Fatty acid oxidation disorders. Annu. Rev. Physiol. 64, 477–502 (2002).

  166. 166.

    Shekhawat, P. S. et al. Spontaneous development of intestinal and colonic atrophy and inflammation in the carnitine-deficient jvs (OCTN2(−/−)) mice. Mol. Genet. Metab. 92, 315–324 (2007).

  167. 167.

    Roediger, W. E. & Nance, S. Metabolic induction of experimental ulcerative colitis by inhibition of fatty acid oxidation. Br. J. Exp. Pathol. 67, 773–782 (1986).

  168. 168.

    Santhanam, S., Venkatraman, A. & Ramakrishna, B. S. Impairment of mitochondrial acetoacetyl CoA thiolase activity in the colonic mucosa of patients with ulcerative colitis. Gut 56, 1543–1549 (2007).

  169. 169.

    Treem, W. R., Ahsan, N., Shoup, M. & Hyams, J. S. Fecal short-chain fatty acids in children with inflammatory bowel disease. J. Pediatr. Gastroenterol. Nutr. 18, 159–164 (1994).

  170. 170.

    Ritzhaupt, A., Wood, I. S., Ellis, A., Hosie, K. B. & Shirazi-Beechey, S. P. Identification and characterization of a monocarboxylate transporter (MCT1) in pig and human colon: its potential to transport L-lactate as well as butyrate. J. Physiol. 513, 719–732 (1998).

  171. 171.

    Thibault, R. et al. Down-regulation of the monocarboxylate transporter 1 is involved in butyrate deficiency during intestinal inflammation. Gastroenterology 133, 1916–1927 (2007).

  172. 172.

    Baur, P. et al. Metabolic phenotyping of the Crohn’s disease-like IBD etiopathology in the TNF(DeltaARE/WT) mouse model. J. Proteome Res. 10, 5523–5535 (2011).

  173. 173.

    Glover, L. E. & Colgan, S. P. Hypoxia and metabolic factors that influence inflammatory bowel disease pathogenesis. Gastroenterology 140, 1748–1755 (2011).

  174. 174.

    Colgan, S. P., Curtis, V. F. & Campbell, E. L. The inflammatory tissue microenvironment in IBD. Inflamm. Bowel Dis. 19, 2238–2244 (2013).

  175. 175.

    Colgan, S. P. & Taylor, C. T. Hypoxia: an alarm signal during intestinal inflammation. Nat. Rev. Gastroenterol. Hepatol. 7, 281–287 (2010).

  176. 176.

    Campbell, E. L. et al. Transmigrating neutrophils shape the mucosal microenvironment through localized oxygen depletion to influence resolution of inflammation. Immunity 40, 66–77 (2014).

  177. 177.

    Ledoux, S. et al. Hypoxia enhances Ecto-5′-Nucleotidase activity and cell surface expression in endothelial cells: role of membrane lipids. Circ. Res. 92, 848–855 (2003).

  178. 178.

    Taylor, C. T. & Cummins, E. P. The role of NF-kappaB in hypoxia-induced gene expression. Ann. NY Acad. Sci 1177, 178–184 (2009).

  179. 179.

    Giatromanolaki, A. et al. Hypoxia inducible factor 1alpha and 2alpha overexpression in inflammatory bowel disease. J. Clin. Pathol. 56, 209–213 (2003).

  180. 180.

    Cummins, E. P. et al. The hydroxylase inhibitor dimethyloxalylglycine is protective in a murine model of colitis. Gastroenterology 134, 156–165 (2008).

  181. 181.

    Ogura, Y. et al. A frameshift mutation in NOD2 associated with susceptibility to Crohn’s disease. Nature 411, 603–606 (2001).

  182. 182.

    Franchimont, D. et al. Deficient host-bacteria interactions in inflammatory bowel disease? The toll-like receptor (TLR)-4 Asp299gly polymorphism is associated with Crohn’s disease and ulcerative colitis. Gut 53, 987–992 (2004).

  183. 183.

    Hampe, J. et al. A genome-wide association scan of nonsynonymous SNPs identifies a susceptibility variant for Crohn disease in ATG16L1. Nat. Genet. 39, 207–211 (2007).

  184. 184.

    Parkes, M. et al. Sequence variants in the autophagy gene IRGM and multiple other replicating loci contribute to Crohn’s disease susceptibility. Nat. Genet. 39, 830–832 (2007).

  185. 185.

    Travassos, L. H. et al. Nod1 and Nod2 direct autophagy by recruiting ATG16L1 to the plasma membrane at the site of bacterial entry. Nat. Immunol. 11, 55–62 (2010).

  186. 186.

    Adolph, T. E. et al. Paneth cells as a site of origin for intestinal inflammation. Nature 503, 272–276 (2013).

  187. 187.

    Cadwell, K. et al. A key role for autophagy and the autophagy gene Atg16l1 in mouse and human intestinal Paneth cells. Nature 456, 259–263 (2008). This study links the ATG16L1 Crohn’s disease risk allele to autophagy, Paneth cell granule abnormalities and degenerating mitochondria.

  188. 188.

    Wehkamp, J. et al. Reduced Paneth cell alpha-defensins in ileal Crohn’s disease. Proc. Natl Acad. Sci. USA 102, 18129–18134 (2005).

  189. 189.

    Singh, S. B. et al. Human IRGM regulates autophagy and cell-autonomous immunity functions through mitochondria. Nat. Cell Biol. 12, 1154–1165 (2010).

  190. 190.

    Chauhan, S., Mandell, M. A. & Deretic, V. IRGM governs the core autophagy machinery to conduct antimicrobial defense. Mol. Cell 58, 507–521 (2015).

  191. 191.

    Pecqueur, C. et al. Uncoupling protein-2 controls proliferation by promoting fatty acid oxidation and limiting glycolysis-derived pyruvate utilization. FASEB J. 22, 9–18 (2008).

  192. 192.

    Emre, Y. et al. Mitochondria contribute to LPS-induced MAPK activation via uncoupling protein UCP2 in macrophages. Biochem. J. 402, 271–278 (2007).

  193. 193.

    Seth, R. B., Sun, L., Ea, C. K. & Chen, Z. J. Identification and characterization of MAVS, a mitochondrial antiviral signaling protein that activates NF-kappaB and IRF 3. Cell 122, 669–682 (2005).

  194. 194.

    Zhou, R., Yazdi, A. S., Menu, P. & Tschopp, J. A role for mitochondria in NLRP3 inflammasome activation. Nature 469, 221–225 (2011).

  195. 195.

    Schroder, K. & Tschopp, J. The inflammasomes. Cell 140, 821–832 (2010).

  196. 196.

    Villani, A. C. et al. Common variants in the NLRP3 region contribute to Crohn’s disease susceptibility. Nat. Genet. 41, 71–76 (2009).

  197. 197.

    Zhernakova, A. et al. Genetic analysis of innate immunity in Crohn’s disease and ulcerative colitis identifies two susceptibility loci harboring CARD9 and IL18RAP. Am. J. Hum. Genet. 82, 1202–1210 (2008).

  198. 198.

    Lei-Leston, A. C., Murphy, A. G. & Maloy, K. J. Epithelial Cell Inflammasomes in Intestinal Immunity and Inflammation. Front. Immunol. 8, 1168 (2017).

  199. 199.

    Ip, W. K. E., Hoshi, N., Shouval, D. S., Snapper, S. & Medzhitov, R. Anti-inflammatory effect of IL-10 mediated by metabolic reprogramming of macrophages. Science 356, 513–519 (2017). This research shows that the anti-inflammatory cytokine IL-10 controls immune responses by opposing the metabolic switch induced by inflammatory stimuli in macrophages, and promotes mitophagy to eliminate dysfunctional mitochondria.

  200. 200.

    Kuhn, R., Lohler, J., Rennick, D., Rajewsky, K. & Muller, W. Interleukin-10-deficient mice develop chronic enterocolitis. Cell 75, 263–274 (1993).

  201. 201.

    Aithal, G. P. et al. Role of polymorphisms in the interleukin-10 gene in determining disease susceptibility and phenotype in inflamatory bowel disease. Dig. Dis. Sci 46, 1520–1525 (2001).

  202. 202.

    Glocker, E. O., Kotlarz, D., Klein, C., Shah, N. & Grimbacher, B. IL-10 and IL-10 receptor defects in humans. Ann. NY Acad. Sci. 1246, 102–107 (2011).

  203. 203.

    Mosser, D. M. & Zhang, X. Interleukin-10: new perspectives on an old cytokine. Immunol. Rev. 226, 205–218 (2008).

  204. 204.

    Choi, C. R., Bakir, I. A., Hart, A. L. & Graham, T. A. Clonal evolution of colorectal cancer in IBD. Nat. Rev. Gastroenterol. Hepatol. 14, 218–229 (2017).

  205. 205.

    Schwitalla, S. et al. Intestinal tumorigenesis initiated by dedifferentiation and acquisition of stem-cell-like properties. Cell 152, 25–38 (2013). This study characterizes the contribution of NF-κB signalling in intestinal epithelial cells to Wnt activation, dedifferentiation of nonstem cells and generation of intestinal tumor-initiating cells.

  206. 206.

    Andersson-Rolf, A., Zilbauer, M., Koo, B. K. & Clevers, H. Stem cells in repair of gastrointestinal epithelia. Physiology 32, 278–289 (2017).

  207. 207.

    Seno, H. et al. Efficient colonic mucosal wound repair requires Trem2 signaling. Proc. Natl Acad. Sci. USA 106, 256–261 (2009).

  208. 208.

    Dignass, A. U. Mechanisms and modulation of intestinal epithelial repair. Inflamm. Bowel Dis. 7, 68–77 (2001).

  209. 209.

    Miyoshi, H., Ajima, R., Luo, C. T., Yamaguchi, T. P. & Stappenbeck, T. S. Wnt5a potentiates TGF-beta signaling to promote colonic crypt regeneration after tissue injury. Science 338, 108–113 (2012).

  210. 210.

    Kim, T. H. et al. Broadly permissive intestinal chromatin underlies lateral inhibition and cell plasticity. Nature 506, 511–515 (2014).

  211. 211.

    Mills, J. C. & Sansom, O. J. Reserve stem cells: Differentiated cells reprogram to fuel repair, metaplasia, and neoplasia in the adult gastrointestinal tract. Sci. Signal. 8, re8 (2015).

  212. 212.

    Shao, J., Sheng, G. G., Mifflin, R. C., Powell, D. W. & Sheng, H. Roles of myofibroblasts in prostaglandin E2-stimulated intestinal epithelial proliferation and angiogenesis. Cancer Res. 66, 846–855 (2006).

  213. 213.

    Li, P. et al. Aspirin use after diagnosis but not prediagnosis improves established colorectal cancer survival: a meta-analysis. Gut 64, 1419–1425 (2015).

  214. 214.

    Oshima, M. et al. Suppression of intestinal polyposis in Apc delta716 knockout mice by inhibition of cyclooxygenase 2 (COX-2). Cell 87, 803–809 (1996).

  215. 215.

    Biancone, L., Tosti, C., De Nigris, F., Fantini, M. & Pallone, F. Selective cyclooxygenase-2 inhibitors and relapse of inflammatory bowel disease. Gastroenterology 125, 637–638 (2003).

  216. 216.

    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).

  217. 217.

    Medema, J. P. & Vermeulen, L. Microenvironmental regulation of stem cells in intestinal homeostasis and cancer. Nature 474, 318–326 (2011).

  218. 218.

    Vermeulen, L., Sprick, M. R., Kemper, K., Stassi, G. & Medema, J. P. Cancer stem cells — old concepts, new insights. Cell Death Differ. 15, 947–958 (2008).

  219. 219.

    Vermeulen, L. et al. Wnt activity defines colon cancer stem cells and is regulated by the microenvironment. Nat. Cell Biol. 12, 468–476 (2010).

  220. 220.

    Barker, N. et al. Crypt stem cells as the cells-of-origin of intestinal cancer. Nature 457, 608–611 (2009).

  221. 221.

    van Es, J. H. et al. Notch/gamma-secretase inhibition turns proliferative cells in intestinal crypts and adenomas into goblet cells. Nature 435, 959–963 (2005).

  222. 222.

    Todaro, M. et al. Colon cancer stem cells dictate tumor growth and resist cell death by production of interleukin-4. Cell Stem Cell 1, 389–402 (2007).

  223. 223.

    de Sousa, E. M., Vermeulen, L., Richel, D. & Medema, J. P. Targeting Wnt signaling in colon cancer stem cells. Clin. Cancer Res. 17, 647–653 (2011).

  224. 224.

    Todaro, M., Francipane, M. G., Medema, J. P. & Stassi, G. Colon cancer stem cells: promise of targeted therapy. Gastroenterology 138, 2151–2162 (2010).

  225. 225.

    Song, I. S. et al. FOXM1-induced PRX3 regulates stemness and survival of colon cancer cells via maintenance of mitochondrial function. Gastroenterology 149, 1006–1016.e9 (2015).

  226. 226.

    Siegelin, M. D. et al. Exploiting the mitochondrial unfolded protein response for cancer therapy in mice and human cells. J. Clin. Invest. 121, 1349–1360 (2011).

  227. 227.

    Pace, A. et al. Hsp60, a novel target for antitumor therapy: structure-function features and prospective drugs design. Curr. Pharm. Des. 19, 2757–2764 (2013).

  228. 228.

    Hartl, M. The quest for targets executing MYC-dependent cell transformation. Front. Oncol. 6, 132 (2016).

  229. 229.

    Tsai, Y. P., Teng, S. C. & Wu, K. J. Direct regulation of HSP60 expression by c-MYC induces transformation. FEBS Lett. 582, 4083–4088 (2008).

  230. 230.

    Yan, F. Q., Wang, J. Q., Tsai, Y. P. & Wu, K. J. HSP60 overexpression increases the protein levels of the p110alpha subunit of phosphoinositide 3-kinase and c-Myc. Clin. Exp. Pharmacol. Physiol. 42, 1092–1097 (2015).

  231. 231.

    Tang, H. et al. Down-regulation of HSP60 suppresses the proliferation of glioblastoma cells via the ROS/AMPK/mTOR pathway. Sci. Rep. 6, 28388 (2016).

  232. 232.

    Lachat, J. J. & Goncalves, R. P. Influence of autonomic denervation upon the kinetics of the ileal epithelium of the rat. Cell Tissue Res. 192, 285–297 (1978).

  233. 233.

    Kaur, P. & Potten, C. S. Circadian variation in migration velocity in small intestinal epithelium. Cell Tissue Kinet. 19, 591–599 (1986).

  234. 234.

    Williams, J. M. et al. Epithelial cell shedding and barrier function: a matter of life and death at the small intestinal villus tip. Vet. Pathol. 52, 445–455 (2015).

  235. 235.

    Burrin, D. G. et al. Glucagon-like peptide 2 dose-dependently activates intestinal cell survival and proliferation in neonatal piglets. Endocrinology 146, 22–32 (2005).

  236. 236.

    Beyaz, S. et al. High-fat diet enhances stemness and tumorigenicity of intestinal progenitors. Nature 531, 53–58 (2016).

  237. 237.

    Shirkey, T. W. et al. Effects of commensal bacteria on intestinal morphology and expression of proinflammatory cytokines in the gnotobiotic pig. Exp. Biol. Med. 231, 1333–1345 (2006).

  238. 238.

    Fallingborg, J. Intraluminal pH of the human gastrointestinal tract. Dan. Med. Bull. 46, 183–196 (1999).

  239. 239.

    Taylor, C. T. & Colgan, S. P. Hypoxia and gastrointestinal disease. J. Mol. Med. 85, 1295–1300 (2007).

  240. 240.

    Bohlen, H. G. Intestinal tissue PO2 and microvascular responses during glucose exposure. Am. J. Physiol. 238, H164–H171 (1980).

  241. 241.

    Arco, A. D. & Satrustegui, J. New mitochondrial carriers: an overview. Cell. Mol. Life Sci. 62, 2204–2227 (2005).

  242. 242.

    Lehmann, G., Udasin, R. G. & Ciechanover, A. On the linkage between the ubiquitin-proteasome system and the mitochondria. Biochem. Biophys. Res. Commun. 473, 80–86 (2016).

  243. 243.

    Frey, T. G. & Mannella, C. A. The internal structure of mitochondria. Trends Biochem. Sci. 25, 319–324 (2000).

  244. 244.

    Koehler, C. M., Beverly, K. N. & Leverich, E. P. Redox pathways of the mitochondrion. Antioxid. Redox Signal. 8, 813–822 (2006).

  245. 245.

    Amiri, M. & Hollenbeck, P. J. Mitochondrial biogenesis in the axons of vertebrate peripheral neurons. Dev. Neurobiol. 68, 1348–1361 (2008).

  246. 246.

    Chen, H. et al. Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J. Cell Biol. 160, 189–200 (2003).

  247. 247.

    Hayashi, T., Rizzuto, R., Hajnoczky, G. & Su, T. P. MAM: more than just a housekeeper. Trends Cell Biol. 19, 81–88 (2009).

  248. 248.

    James, A. M., Collins, Y., Logan, A. & Murphy, M. P. Mitochondrial oxidative stress and the metabolic syndrome. Trends Endocrinol. Metab. 23, 429–434 (2012).

  249. 249.

    Kaelin, W. G. Jr & McKnight, S. L. Influence of metabolism on epigenetics and disease. Cell 153, 56–69 (2013).

  250. 250.

    Raimundo, N., Baysal, B. E. & Shadel, G. S. Revisiting the TCA cycle: signaling to tumor formation. Trends Mol. Med. 17, 641–649 (2011).

  251. 251.

    Selak, M. A. et al. Succinate links TCA cycle dysfunction to oncogenesis by inhibiting HIF-alpha prolyl hydroxylase. Cancer Cell 7, 77–85 (2005).

  252. 252.

    Raimundo, N., Vanharanta, S., Aaltonen, L. A., Hovatta, I. & Suomalainen, A. Downregulation of SRF-FOS-JUNB pathway in fumarate hydratase deficiency and in uterine leiomyomas. Oncogene 28, 1261–1273 (2009).

  253. 253.

    Adam, J. et al. Renal cyst formation in Fh1-deficient mice is independent of the Hif/Phd pathway: roles for fumarate in KEAP1 succination and Nrf2 signaling. Cancer Cell 20, 524–537 (2011).

  254. 254.

    Zhang, Z. et al. Identification of lysine succinylation as a new post-translational modification. Nat. Chem. Biol. 7, 58–63 (2011).

  255. 255.

    Killela, P. J. et al. Mutations in IDH1, IDH2, and in the TERT promoter define clinically distinct subgroups of adult malignant gliomas. Oncotarget 5, 1515–1525 (2014).

  256. 256.

    Dang, L. et al. Cancer-associated IDH1 mutations produce 2-hydroxyglutarate. Nature 465, 966 (2010).

  257. 257.

    Charitou, P. et al. FOXOs support the metabolic requirements of normal and tumor cells by promoting IDH1 expression. EMBO Rep. 16, 456–466 (2015).

  258. 258.

    Figueroa, M. E. et al. Leukemic IDH1 and IDH2 mutations result in a hypermethylation phenotype, disrupt TET2 function, and impair hematopoietic differentiation. Cancer Cell 18, 553–567 (2010).

  259. 259.

    Hall, J. A., Dominy, J. E., Lee, Y. & Puigserver, P. The sirtuin family’s role in aging and age-associated pathologies. J. Clin. Invest. 123, 973–979 (2013).

  260. 260.

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

  261. 261.

    Zhang, J. et al. UCP2 regulates energy metabolism and differentiation potential of human pluripotent stem cells. EMBO J. 30, 4860–4873 (2011).

  262. 262.

    Wang, J. et al. Dependence of mouse embryonic stem cells on threonine catabolism. Science 325, 435–439 (2009).

  263. 263.

    Ito, K. et al. A PML-PPAR-delta pathway for fatty acid oxidation regulates hematopoietic stem cell maintenance. Nat. Med. 18, 1350–1358 (2012).

  264. 264.

    Shyh-Chang, N. & Ng, H. H. The metabolic programming of stem cells. Genes Dev. 31, 336–346 (2017).

  265. 265.

    Arnould, T., Michel, S. & Renard, P. Mitochondria retrograde signaling and the UPR mt: where are we in mammals? Int. J. Mol. Sci. 16, 18224–18251 (2015).

  266. 266.

    Quiros, P. M., Mottis, A. & Auwerx, J. Mitonuclear communication in homeostasis and stress. Nat. Rev. Mol. Cell Biol. 17, 213–226 (2016).

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  1. Technische Universität München, Chair of Nutrition and Immunology, Freising, Germany

    • Eva Rath
    •  & Dirk Haller
  2. Department of Interdisciplinary Medicine, “Aldo Moro” University of Bari, Bari, Italy

    • Antonio Moschetta
  3. National Cancer Research Center, IRCCS Istituto Oncologico “Giovanni Paolo II”, Bari, Italy

    • Antonio Moschetta
  4. Technische Universität München, ZIEL — Institute for Food & Health, Munich, Germany

    • Dirk Haller


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Correspondence to Dirk Haller.

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