Review Article | Published:

Metabolic interventions in the immune response to cancer

Nature Reviews Immunology (2019) | Download Citation


At the centre of the therapeutic dilemma posed by cancer is the question of how to develop more effective treatments that discriminate between normal and cancerous tissues. Decades of research have shown us that universally applicable principles are rare, but two well-accepted concepts have emerged: first, that malignant transformation goes hand in hand with distinct changes in cellular metabolism; second, that the immune system is critical for tumour control and clearance. Unifying our understanding of tumour metabolism with immune cell function may prove to be a powerful approach in the development of more effective cancer therapies. Here, we explore how nutrient availability in the tumour microenvironment shapes immune responses and identify areas of intervention to modulate the metabolic constraints placed on immune cells in this setting.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Additional information

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.


  1. 1.

    Vyas, S., Zaganjor, E. & Haigis, M. C. Mitochondria and cancer. Cell 166, 555–566 (2016).

  2. 2.

    Aras, S. & Zaidi, M. R. TAMeless traitors: macrophages in cancer progression and metastasis. Br. J. Cancer 117, 1583–1591 (2017).

  3. 3.

    Wellenstein, M. D. & de Visser, K. E. Cancer-cell-intrinsic mechanisms shaping the tumor immune landscape. Immunity 48, 399–416 (2018).

  4. 4.

    Chang, C.-H. et al. Metabolic competition in the tumor microenvironment is a driver of cancer progression. Cell 162, 1229–1241 (2015).

  5. 5.

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

  6. 6.

    Pagès, F. et al. International validation of the consensus Immunoscore for the classification of colon cancer: a prognostic and accuracy study. Lancet 391, 2128–2139 (2018).

  7. 7.

    Pavlova, N. N. & Thompson, C. B. The emerging hallmarks of cancer metabolism. Cell Metab. 23, 27–47 (2016).

  8. 8.

    O’Neill, L. A. J. & Pearce, E. J. Immunometabolism governs dendritic cell and macrophage function. J. Exp. Med. 213, 15–23 (2016).

  9. 9.

    Buck, M. D., Sowell, R. T., Kaech, S. M. & Pearce, E. L. Metabolic instruction of immunity. Cell 169, 570–586 (2017).

  10. 10.

    Assmann, N. et al. Srebp-controlled glucose metabolism is essential for NK cell functional responses. Nat. Immunol. 18, 1197–1206 (2017).

  11. 11.

    Badur, M. G. & Metallo, C. M. Reverse engineering the cancer metabolic network using flux analysis to understand drivers of human disease. Metab. Eng. 45, 95–108 (2018).This review highlights different approaches to understand metabolic fluxes in the context of cancer while also covering the fundamentals of cancer metabolism.

  12. 12.

    Andrejeva, G. & Rathmell, J. C. Similarities and distinctions of cancer and immune metabolism in inflammation and tumors. Cell Metab. 26, 49–70 (2017).

  13. 13.

    Mehta, M. M., Weinberg, S. E. & Chandel, N. S. Mitochondrial control of immunity: beyond ATP. Nat. Rev. Immunol. 17, 608–620 (2017).

  14. 14.

    Grzes, K. M. et al. Control of amino acid transport coordinates metabolic reprogramming in T cell malignancy. Leukemia 31, 2771–2779 (2017).

  15. 15.

    Crompton, J. G. et al. Akt inhibition enhances expansion of potent tumor-specific lymphocytes with memory cell characteristics. Cancer Res. 75, 296–305 (2015).

  16. 16.

    Palm, W., Araki, J., King, B., DeMatteo, R. G. & Thompson, C. B. Critical role for PI3-kinase in regulating the use of proteins as an amino acid source. Proc. Natl Acad. Sci. USA 114, E8628–E8636 (2017).

  17. 17.

    Mayers, J. R. et al. Tissue of origin dictates branched-chain amino acid metabolism in mutant Kras-driven cancers. Science 353, 1161–1165 (2016).

  18. 18.

    Reznik, E. et al. A landscape of metabolic variation across tumor types. Cell Syst. 6, 301–313 (2018).This study integrates diverse metabolic data sets of clinical relevance and does a thorough analysis of the common and distinct metabolic features of human cancer.

  19. 19.

    Hensley, C. T. et al. Metabolic heterogeneity in human lung tumors. Cell 164, 681–694 (2016).This paper explores the metabolic profiles of human lung tumours, highlighting the differences encountered between tumours across patients but also within the same individual.

  20. 20.

    Porporato, P. E., Filigheddu, N., Pedro, J. M. B.-S., Kroemer, G. & Galluzzi, L. Mitochondrial metabolism and cancer. Cell Res. 28, 265–280 (2018).

  21. 21.

    LeBleu, V. S. et al. PGC-1α mediates mitochondrial biogenesis and oxidative phosphorylation in cancer cells to promote metastasis. Nat. Cell Biol. 16, 992–1003 (2014).

  22. 22.

    Yang, C. et al. Glutamine oxidation maintains the TCA cycle and cell survival during impaired mitochondrial pyruvate transport. Mol. Cell 56, 414–424 (2014).

  23. 23.

    Pavlova, N. N. et al. As extracellular glutamine levels decline, asparagine becomes an essential amino acid. Cell Metab. 27, 428–438 (2018).

  24. 24.

    Kuo, C.-Y. & Ann, D. K. When fats commit crimes: fatty acid metabolism, cancer stemness and therapeutic resistance. Cancer Commun. 38, 47 (2018).

  25. 25.

    Qu, Q., Zeng, F., Liu, X., Wang, Q. J. & Deng, F. Fatty acid oxidation and carnitine palmitoyltransferase I: emerging therapeutic targets in cancer. Cell Death Dis. 7, e2226 (2016).

  26. 26.

    Spinelli, J. B. et al. Metabolic recycling of ammonia via glutamate dehydrogenase supports breast cancer biomass. Science 358, 941–946 (2017).This study describes the way in which breast cancer cells utilize ammonia to replenish their amino acid pools via glutamate dehydrogenase.

  27. 27.

    Corbet, C. et al. Interruption of lactate uptake by inhibiting mitochondrial pyruvate transport unravels direct antitumor and radiosensitizing effects. Nat. Commun. 9, 1208 (2018).

  28. 28.

    Faubert, B. et al. Lactate metabolism in human lung tumors. Cell 171, 358–371 (2017). This paper describes the way in which human lung cancer tumours can utilize lactate in vivo as an alternative source of carbon.

  29. 29.

    Gatto, F., Nookaew, I. & Nielsen, J. Chromosome 3p loss of heterozygosity is associated with a unique metabolic network in clear cell renal carcinoma. Proc. Natl Acad. Sci. USA 111, E866–E875 (2014).

  30. 30.

    Hu, J. et al. Heterogeneity of tumor-induced gene expression changes in the human metabolic network. Nat. Biotechnol. 31, 522–529 (2013).

  31. 31.

    Nilsson, R. et al. Metabolic enzyme expression highlights a key role for MTHFD2 and the mitochondrial folate pathway in cancer. Nat. Commun. 5, 3128 (2014).

  32. 32.

    Buescher, J. M. & Driggers, E. M. Integration of omics: more than the sum of its parts. Cancer Metab. 4, 4 (2016).

  33. 33.

    de Bruin, E. C. et al. Spatial and temporal diversity in genomic instability processes defines lung cancer evolution. Science 346, 251–256 (2014).

  34. 34.

    Maj, T. et al. Oxidative stress controls regulatory T cell apoptosis and suppressor activity and PD-L1-blockade resistance in tumor. Nat. Immunol. 18, 1332–1341 (2017).

  35. 35.

    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).

  36. 36.

    Gentles, A. J. et al. The prognostic landscape of genes and infiltrating immune cells across human cancers. Nat. Med. 21, 938–945 (2015).

  37. 37.

    Thorsson, V. et al. The immune landscape of cancer. Immunity 48, 812–830 (2018). This paper describes the way in which human lung cancer tumours can utilize lactate in vivo as an alternative source of carbon.

  38. 38.

    Ho, P.-C. et al. Phosphoenolpyruvate is a metabolic checkpoint of anti-tumor T cell responses. Cell 162, 1217–1228 (2015).

  39. 39.

    Keating, S. E. et al. Metabolic reprogramming supports IFN-γ production by CD56bright NK cells. J. Immunol. 196, 2552–2560 (2016).

  40. 40.

    Cong, J. et al. Dysfunction of natural killer cells by FBP1-induced inhibition of glycolysis during lung cancer progression. Cell Metab. 28, 243–255 (2018).

  41. 41.

    Cascone, T. et al. Increased tumor glycolysis characterizes immune resistance to adoptive T cell therapy. Cell Metab. 27, 977–987 (2018).

  42. 42.

    Chang, C.-H. et al. Posttranscriptional control of T cell effector function by aerobic glycolysis. Cell 153, 1239–1251 (2013).

  43. 43.

    Zhao, E. et al. Cancer mediates effector T cell dysfunction by targeting microRNAs and EZH2 via glycolysis restriction. Nat. Immunol. 17, 95–103 (2015).

  44. 44.

    Song, M. et al. IRE1α-XBP1 controls T cell function in ovarian cancer by regulating mitochondrial activity. Nature 562, 423–428 (2018).

  45. 45.

    Xia, H. et al. Suppression of FIP200 and autophagy by tumor-derived lactate promotes naïve T cell apoptosis and affects tumor immunity. Sci. Immunol. 2, eaan4631 (2017).

  46. 46.

    Angelin, A. et al. Foxp3 reprograms T cell metabolism to function in low-glucose, high-lactate environments. Cell Metab. 25, 1282–1293 (2017).

  47. 47.

    Haas, R. et al. Lactate regulates metabolic and pro-inflammatory circuits in control of T cell migration and effector functions. PLOS Biol. 13, e1002202 (2015).

  48. 48.

    Li, W. et al. Aerobic glycolysis controls myeloid-derived suppressor cells and tumor immunity via a specific CEBPB isoform in triple-negative breast cancer. Cell Metab. 28, 87–103 (2018).

  49. 49.

    Klysz, D. et al. Glutamine-dependent α-ketoglutarate production regulates the balance between T helper 1 cell and regulatory T cell generation. Sci. Signal. 8, ra97 (2015).

  50. 50.

    Araujo, L., Khim, P., Mkhikian, H., Mortales, C.-L. & Demetriou, M. Glycolysis and glutaminolysis cooperatively control T cell function by limiting metabolite supply to N-glycosylation. eLife 6, 1239 (2017).

  51. 51.

    Ma, E. H. et al. Serine is an essential metabolite for effector T cell expansion. Cell Metab. 25, 482 (2017).

  52. 52.

    Ron-Harel, N. et al. Mitochondrial biogenesis and proteome remodeling promote one-carbon metabolism for T cell activation. Cell Metab. 24, 104–117 (2016).

  53. 53.

    Swamy, M. et al. Glucose and glutamine fuel protein O-GlcNAcylation to control T cell self-renewal and malignancy. Nat. Immunol. 17, 712–720 (2016).

  54. 54.

    Loftus, R. M. et al. Amino acid-dependent cMyc expression is essential for NK cell metabolic and functional responses in mice. Nat. Commun. 9, 2341 (2018).

  55. 55.

    Ren, W. et al. Amino-acid transporters in T cell activation and differentiation. Cell Death Dis. 8, e2655 (2017).

  56. 56.

    Malinarich, F. et al. High mitochondrial respiration and glycolytic capacity represent a metabolic phenotype of human tolerogenic dendritic cells. J. Immunol. 194, 5174–5186 (2015).

  57. 57.

    Goffaux, G., Hammami, I. & Jolicoeur, M. A. Dynamic metabolic flux analysis of myeloid-derived suppressor cells confirms immunosuppression-related metabolic plasticity. Sci. Rep. 7, 9850 (2017).

  58. 58.

    Cantelmo, A. R. et al. Inhibition of the glycolytic activator PFKFB3 in endothelium induces tumor vessel normalization, impairs metastasis, and improves chemotherapy. Cancer Cell 30, 968–985 (2016).This paper explores the metabolism of endothelial cells in the context of cancer.

  59. 59.

    Liu, X. et al. Regulatory T cells trigger effector T cell DNA damage and senescence caused by metabolic competition. Nat. Commun. 9, 249 (2018).

  60. 60.

    Li, L. et al. TLR8-mediated metabolic control of human Treg function: a mechanistic target for cancer immunotherapy. Cell Metab. 29, 103–123 (2018).

  61. 61.

    Ladanyi, A. et al. Adipocyte-induced CD36 expression drives ovarian cancer progression and metastasis. Oncogene 37, 2285–2301 (2018).

  62. 62.

    Rozovski, U. et al. STAT3-activated CD36 facilitates fatty acid uptake in chronic lymphocytic leukemia cells. Oncotarget 9, 21268–21280 (2018).

  63. 63.

    Huang, S. C.-C. et al. Cell-intrinsic lysosomal lipolysis is essential for alternative activation of macrophages. Nat. Immunol. 15, 846–855 (2014).

  64. 64.

    Mantovani, A., Sozzani, S., Locati, M., Allavena, P. & Sica, A. Macrophage polarization: tumor-associated macrophages as a paradigm for polarized M2 mononuclear phagocytes. Trends Immunol. 23, 549–555 (2002).

  65. 65.

    Kim, N. H. et al. Snail reprograms glucose metabolism by repressing phosphofructokinase PFKP allowing cancer cell survival under metabolic stress. Nat. Commun. 8, 14374 (2017).

  66. 66.

    Romero, R. et al. Keap1 loss promotes Kras-driven lung cancer and results in dependence on glutaminolysis. Nat. Med. 23, 1362–1368 (2017).

  67. 67.

    Frossi, B., De Carli, M., Piemonte, M. & Pucillo, C. Oxidative microenvironment exerts an opposite regulatory effect on cytokine production by Th1 and Th2 cells. Mol. Immunol. 45, 58–64 (2008).

  68. 68.

    Mills, E. L. & O’Neill, L. A. Reprogramming mitochondrial metabolism in macrophages as an anti-inflammatory signal. Eur. J. Immunol. 46, 13–21 (2016).

  69. 69.

    Tannahill, G. M. et al. Succinate is an inflammatory signal that induces IL-1β through HIF-1α. Nature 496, 238–242 (2016).

  70. 70.

    Zhang, Y. et al. ROS play a critical role in the differentiation of alternatively activated macrophages and the occurrence of tumor-associated macrophages. Cell Res. 23, 898–914 (2013).

  71. 71.

    Cubillos-Ruiz, J. R. et al. ER stress sensor XBP1 controls anti-tumor immunity by disrupting dendritic cell homeostasis. Cell 161, 1527–1538 (2015).

  72. 72.

    Tardito, S. et al. Glutamine synthetase activity fuels nucleotide biosynthesis and supports growth of glutamine-restricted glioblastoma. Nat. Cell Biol. 17, 1556–1568 (2015).

  73. 73.

    Cormerais, Y. et al. The glutamine transporter ASCT2 (SLC1A5) promotes tumor growth independently of the amino acid transporter LAT1 (SLC7A5). J. Biol. Chem. 293, 2877–2887 (2018).

  74. 74.

    Schulte, M. L. et al. Pharmacological blockade of ASCT2-dependent glutamine transport leads to antitumor efficacy in preclinical models. Nat. Med. 24, 194–202 (2018).

  75. 75.

    Johnson, M. O. et al. Distinct regulation of Th17 and Th1 cell differentiation by glutaminase-dependent metabolism. Cell 175, 1780–1795 (2018).

  76. 76.

    Nakaya, M. et al. Inflammatory T cell responses rely on amino acid transporter ASCT2 facilitation of glutamine uptake and mTORC1 kinase activation. Immunity 40, 692–705 (2014).

  77. 77.

    Kim, B., Li, J., Jang, C. & Arany, Z. Glutamine fuels proliferation but not migration of endothelial cells. EMBO J. 36, 2321–2333 (2017).

  78. 78.

    Eelen, G. et al. Role of glutamine synthetase in angiogenesis beyond glutamine synthesis. Nature 561, 63–69 (2018).

  79. 79.

    Yang, L. et al. Targeting stromal glutamine synthetase in tumors disrupts tumor microenvironment-regulated cancer cell growth. Cell Metab. 24, 685–700 (2016).

  80. 80.

    Palmieri, E. M. et al. Pharmacologic or genetic targeting of glutamine synthetase skews macrophages toward an M1-like phenotype and inhibits tumor metastasis. Cell Rep. 20, 1654–1666 (2017).

  81. 81.

    Liu, P.-S. et al. α-Ketoglutarate orchestrates macrophage activation through metabolic and epigenetic reprogramming. Nat. Immunol. 18, 985–994 (2017).This study establishes the impact of changing the relative amounts of α-ketoglutarate in macrophages with regard to the phenotype of these cells.

  82. 82.

    Mayers, J. R. et al. Elevation of circulating branched-chain amino acids is an early event in human pancreatic adenocarcinoma development. Nat. Med. 20, 1193–1198 (2014).

  83. 83.

    Katagiri, R. et al. Increased levels of branched-chain amino acid associated with increased risk of pancreatic cancer in a prospective case-control study of a large cohort. Gastroenterology 155, 1474–1482 (2018).

  84. 84.

    Albaugh, V. L., Pinzon-Guzman, C. & Barbul, A. Arginine-dual roles as an onconutrient and immunonutrient. J. Surg. Oncol. 115, 273–280 (2017).

  85. 85.

    Fletcher, M. et al. L-arginine depletion blunts antitumor T cell responses by inducing myeloid-derived suppressor cells. Cancer Res. 75, 275–283 (2015).

  86. 86.

    Geiger, R. et al. L-arginine modulates T cell metabolism and enhances survival and anti-tumor activity. Cell 167, 829–842 (2016).

  87. 87.

    Steggerda, S. M. et al. Inhibition of arginase by CB-1158 blocks myeloid cell-mediated immune suppression in the tumor microenvironment. J. Immunother. Cancer 5, 101 (2017).

  88. 88.

    Munn, D. H. & Mellor, A. L. Indoleamine 2,3 dioxygenase and metabolic control of immune responses. Trends Immunol. 34, 137–143 (2013).

  89. 89.

    Mondanelli, G. et al. A relay pathway between arginine and tryptophan metabolism confers immunosuppressive properties on dendritic cells. Immunity 46, 233–244 (2017).

  90. 90.

    Chuang, S.-C. et al. Circulating biomarkers of tryptophan and the kynurenine pathway and lung cancer risk. Cancer Epidemiol. Biomarkers Prev. 23, 461–468 (2014).

  91. 91.

    Mullard, A. IDO takes a blow. Nat. Rev. Drug Discov. 17, 307–307 (2018).

  92. 92.

    Sarrouilhe, D. & Mesnil, M. Serotonin and human cancer: a critical view. Biochimie. (2018).

  93. 93.

    Wu, H., Denna, T. H., Storkersen, J. N. & Gerriets, V. A. Beyond a neurotransmitter: the role of serotonin in inflammation and immunity. Pharmacol. Res. 140, 100–114 (2018).

  94. 94.

    Wang, Q. et al. 5-HTR3 and 5-HTR4 located on the mitochondrial membrane and functionally regulated mitochondrial functions. Sci. Rep. 6, 37336 (2016).

  95. 95.

    Tan, D.-X., Manchester, L. C., Qin, L. & Reiter, R. J. Melatonin: a mitochondrial targeting molecule involving mitochondrial protection and dynamics. Int. J. Mol. Sci. 17, 2124 (2016).

  96. 96.

    Triplett, T. A. et al. Reversal of indoleamine 2,3-dioxygenase-mediated cancer immune suppression by systemic kynurenine depletion with a therapeutic enzyme. Nat. Biotechnol. 13, 5 (2018).

  97. 97.

    Kumar, V. et al. Cancer-associated fibroblasts neutralize the anti-tumor effect of CSF1 receptor blockade by inducing PMN-MDSC infiltration of tumors. Cancer Cell 32, 654–668 (2017).

  98. 98.

    Cannarile, M. A. et al. Colony-stimulating factor 1 receptor (CSF1R) inhibitors in cancer therapy. J. Immunother. Cancer 5, 53 (2017).

  99. 99.

    Gao, A., Sun, Y. & Peng, G. ILT4 functions as a potential checkpoint molecule for tumor immunotherapy. Biochim. Biophys. Acta Rev. Cancer 1869, 278–285 (2018).

  100. 100.

    Lyons, Y. A. et al. Macrophage depletion through colony stimulating factor 1 receptor pathway blockade overcomes adaptive resistance to anti-VEGF therapy. Oncotarget 8, 96496–96505 (2017).

  101. 101.

    Ridker, P. M. et al. Effect of interleukin-1β inhibition with canakinumab on incident lung cancer in patients with atherosclerosis: exploratory results from a randomised, double-blind, placebo-controlled trial. Lancet 390, 1833–1842 (2017).

  102. 102.

    Ribas, A. & Wolchok, J. D. Cancer immunotherapy using checkpoint blockade. Science 359, 1350–1355 (2018).

  103. 103.

    Lim, W. A. & June, C. H. The principles of engineering immune cells to treat cancer. Cell 168, 724–740 (2017).

  104. 104.

    Hartmann, J., Schüßler-Lenz, M., Bondanza, A. & Buchholz, C. J. Clinical development of CAR T cells-challenges and opportunities in translating innovative treatment concepts. EMBO Mol. Med. 9, 1183–1197 (2017).

  105. 105.

    Vyas, M., Müller, R. & Pogge von Strandmann, E. Antigen loss variants: catching hold of escaping foes. Front. Immunol. 8, 991 (2017).

  106. 106.

    Leslie, M. New cancer-fighting cells enter trials. Science 361, 1056–1057 (2018).

  107. 107.

    Fraietta, J. A. et al. Determinants of response and resistance to CD19 chimeric antigen receptor (CAR) T cell therapy of chronic lymphocytic leukemia. Nat. Med. 24, 563–571 (2018).This paper correlates the clinical outcome of CAR T cell therapy with the prevalence of a memory precursor phenotype at the time of leukapheresis.

  108. 108.

    Buck, M. D. et al. Mitochondrial dynamics controls T cell fate through metabolic programming. Cell 166, 63–76 (2016).

  109. 109.

    Kawalekar, O. U. et al. Distinct signaling of coreceptors regulates specific metabolism pathways and impacts memory development in CAR T cells. Immunity 44, 380–390 (2016).

  110. 110.

    Salter, A. I. et al. Phosphoproteomic analysis of chimeric antigen receptor signaling reveals kinetic and quantitative differences that affect cell function. Sci. Signal. 11, eaat6753 (2018).This paper characterizes the phosphoproteome of CAR T cells with different co-stimulatory signalling domains.

  111. 111.

    Feucht, J. et al. Calibration of CAR activation potential directs alternative T cell fates and therapeutic potency. Nat. Med. 545, 423 (2018).

  112. 112.

    Hickman, T. et al. Adaptability of antibody-coupled T cell receptor (ACTR) engineered autologous T cells in combination with daratumumab over CAR-based approaches. Blood 130, 3189 (2017).

  113. 113.

    Raj, D. et al. Switchable CAR-T cells mediate remission in metastatic pancreatic ductal adenocarcinoma. Gut. (2018).This study demonstrates the potential of using an antibody-based switch to control and target CAR T cells.

  114. 114.

    Yoon, D. H., Osborn, M. J., Tolar, J. & Kim, C. J. Incorporation of immune checkpoint blockade into chimeric antigen receptor T cells (CAR-Ts): combination or built-in CAR-T. Int. J. Mol. Sci. 19, 340 (2018).

  115. 115.

    Sengupta, S., Katz, S. C., Sengupta, S. & Sampath, P. Glycogen synthase kinase 3 inhibition lowers PD-1 expression, promotes long-term survival and memory generation in antigen-specific CAR-T cells. Cancer Lett. 433, 131–139 (2018).

  116. 116.

    Tang, L. et al. Enhancing T cell therapy through TCR-signaling-responsive nanoparticle drug delivery. Nat. Biotechnol. 36, 707–716 (2018).This paper highlights a novel approach for drug delivery.

  117. 117.

    Krenciute, G. et al. Transgenic expression of IL15 improves antiglioma activity of IL13Rα2-CAR T cells but results in antigen loss variants. Cancer Immunol. Res. 5, 571–581 (2017).

  118. 118.

    Prestipino, A. et al. Oncogenic JAK2V617F causes PD-L1 expression, mediating immune escape in myeloproliferative neoplasms. Sci. Transl Med. 10, eaam7729 (2018).

  119. 119.

    Zhang, Y. et al. Enhancing CD8+T cell fatty acid catabolism within a metabolically challenging tumor microenvironment increases the efficacy of melanoma immunotherapy. Cancer Cell 32, 377–391 (2017).

  120. 120.

    Patsoukis, N. et al. PD-1 alters T cell metabolic reprogramming by inhibiting glycolysis and promoting lipolysis and fatty acid oxidation. Nat. Commun. 6, 269 (2015).

  121. 121.

    Chamoto, K. et al. Mitochondrial activation chemicals synergize with surface receptor PD-1 blockade for T cell-dependent antitumor activity. Proc. Natl Acad. Sci. USA 114, E761–E770 (2017).

  122. 122.

    Parry, R. V. et al. CTLA-4 and PD-1 receptors inhibit T cell activation by distinct mechanisms. Mol. Cell. Biol. 25, 9543–9553 (2005).

  123. 123.

    Baumeister, S. H., Freeman, G. J., Dranoff, G. & Sharpe, A. H. Coinhibitory pathways in immunotherapy for cancer. Annu. Rev. Immunol. 34, 539–573 (2016).

  124. 124.

    Hui, E. et al. T cell costimulatory receptor CD28 is a primary target for PD-1-mediated inhibition. Science 355, 1428–1433 (2017).

  125. 125.

    Frauwirth, K. A. et al. The CD28 signaling pathway regulates glucose metabolism. Immunity 16, 769–777 (2002).

  126. 126.

    Menk, A. V. et al. 4-1BB costimulation induces T cell mitochondrial function and biogenesis enabling cancer immunotherapeutic responses. J. Exp. Med. 215, 1091–1100 (2018).

  127. 127.

    Geltink, R. I. K. et al. Mitochondrial priming by CD28. Cell 171, 385–390 (2017).

  128. 128.

    Afzal, M. Z., Mercado, R. R. & Shirai, K. Efficacy of metformin in combination with immune checkpoint inhibitors (anti-PD-1/anti-CTLA-4) in metastatic malignant melanoma. J. Immunother. Cancer 6, 64 (2018).

  129. 129.

    Eikawa, S. et al. Immune-mediated antitumor effect by type 2 diabetes drug, metformin. Proc. Natl Acad. Sci. USA 112, 1809–1814 (2015).

  130. 130.

    Liu, X., Romero, I. L., Litchfield, L. M., Lengyel, E. & Locasale, J. W. Metformin targets central carbon metabolism and reveals mitochondrial requirements in human cancers. Cell Metab. 24, 728–739 (2016).

  131. 131.

    Foretz, M., Guigas, B., Bertrand, L., Pollak, M. & Viollet, B. Metformin: from mechanisms of action to therapies. Cell Metab. 20, 953–966 (2014).

  132. 132.

    Scharping, N. E., Menk, A. V., Whetstone, R. D., Zeng, X. & Delgoffe, G. M. Efficacy of PD-1 blockade is potentiated by metformin-induced reduction of tumor hypoxia. Cancer Immunol. Res. 5, 9–16 (2017).

  133. 133.

    Cha, J.-H. et al. Metformin promotes antitumor immunity via endoplasmic-reticulum-associated degradation of PD-L1. Mol. Cell 71, 606–620 (2018).

  134. 134.

    Matson, V. et al. The commensal microbiome is associated with anti-PD-1 efficacy in metastatic melanoma patients. Science 359, 104–108 (2018).

  135. 135.

    Gopalakrishnan, V. et al. Gut microbiome modulates response to anti-PD-1 immunotherapy in melanoma patients. Science 359, 97–103 (2018).

  136. 136.

    Routy, B. et al. Gut microbiome influences efficacy of PD-1-based immunotherapy against epithelial tumors. Science 359, 91–97 (2018).The results presented in studies 134, 135 and 136 highlight the impact that the commensal microbiota has on the clinical outcomes of ICB therapy.

  137. 137.

    Nicholson, J. K. et al. Host-gut microbiota metabolic interactions. Science 336, 1262–1267 (2012).

  138. 138.

    Rodriguez, J., Hiel, S. & Delzenne, N. M. Metformin: old friend, new ways of action-implication of the gut microbiome? Curr. Opin. Clin. Nutr. Metab. Care 21, 294–301 (2018).

  139. 139.

    Zheng, X. et al. Increased vessel perfusion predicts the efficacy of immune checkpoint blockade. J. Clin. Invest. 128, 2104–2115 (2018).

  140. 140.

    Tian, L. et al. Mutual regulation of tumour vessel normalization and immunostimulatory reprogramming. Nature 544, 250–254 (2017).

  141. 141.

    O’Sullivan, D. & Pearce, E. L. Targeting T cell metabolism for therapy. Trends Immunol. 36, 71–80 (2015).

  142. 142.

    Miyajima, M. et al. Metabolic shift induced by systemic activation of T cells in PD-1-deficient mice perturbs brain monoamines and emotional behavior. Nat. Immunol. 18, 1342–1352 (2017).This paper demonstrates how a strong T cell response can impact systemic metabolite levels.

  143. 143.

    He, X., Lin, H., Yuan, L. & Li, B. Combination therapy with L-arginine and α-PD-L1 antibody boosts immune response against osteosarcoma in immunocompetent mice. Cancer Biol. Ther. 18, 94–100 (2017).

  144. 144.

    Berger, S. L. & Sassone-Corsi, P. Metabolic signaling to chromatin. Cold Spring Harb. Perspect. Biol. 8, a019463 (2016).

  145. 145.

    Hopkins, B. D. et al. Suppression of insulin feedback enhances the efficacy of PI3K inhibitors. Nature 560, 499–503 (2018).

  146. 146.

    Weber, D. D., Aminazdeh-Gohari, S. & Kofler, B. Ketogenic diet in cancer therapy. Aging 10, 164–165 (2018).

  147. 147.

    Malvi, P. et al. Weight control interventions improve therapeutic efficacy of dacarbazine in melanoma by reversing obesity-induced drug resistance. Cancer Metab. 4, 21 (2016).

  148. 148.

    Pascual, G. et al. Targeting metastasis-initiating cells through the fatty acid receptor CD36. Nature 541, 41–45 (2017).

  149. 149.

    O’Flanagan, C. H., Smith, L. A., McDonell, S. B. & Hursting, S. D. When less may be more: calorie restriction and response to cancer therapy. BMC Med. 15, 106 (2017).

  150. 150.

    Raffaghello, L. et al. Starvation-dependent differential stress resistance protects normal but not cancer cells against high-dose chemotherapy. Proc. Natl Acad. Sci. USA 105, 8215–8220 (2008).

  151. 151.

    Kalaany, N. Y. & Sabatini, D. M. Tumours with PI3K activation are resistant to dietary restriction. Nature 458, 725–731 (2009).

  152. 152.

    Farazi, M. et al. Caloric restriction maintains OX40 agonist-mediated tumor immunity and CD4 T cell priming during aging. Cancer Immunol. Immunother. 63, 615–626 (2014).

  153. 153.

    Rubio-Patiño, C. et al. Low-protein diet induces IRE1α-dependent anticancer immunosurveillance. Cell Metab. 27, 828–842 (2018).

  154. 154.

    Baek, A. E. et al. The cholesterol metabolite 27 hydroxycholesterol facilitates breast cancer metastasis through its actions on immune cells. Nat. Commun. 8, 864 (2017).

  155. 155.

    Wang, Z. et al. Paradoxical effects of obesity on T cell function during tumor progression and PD-1 checkpoint blockade. Nat. Med. 372, 2521 (2018).

  156. 156.

    Fabbiano, S. et al. Caloric restriction leads to browning of white adipose tissue through type 2 immune signaling. Cell Metab. 24, 434–446 (2016).

  157. 157.

    Messaoudi, I. et al. Delay of T cell senescence by caloric restriction in aged long-lived nonhuman primates. Proc. Natl Acad. Sci. USA 103, 19448–19453 (2006).

  158. 158.

    Sukumar, M., Kishton, R. J. & Restifo, N. P. Metabolic reprograming of anti-tumor immunity. Curr. Opin. Immunol. 46, 14–22 (2017).

  159. 159.

    Lawless, S. J. et al. Glucose represses dendritic cell-induced T cell responses. Nat. Commun. 8, 15620 (2017).

  160. 160.

    Thwe, P. M. & Amiel, E. The role of nitric oxide in metabolic regulation of Dendritic cell immune function. Cancer Lett. 412, 236–242 (2018).

  161. 161.

    Hashimoto, M. et al. CD8 T cell exhaustion in chronic infection and cancer: opportunities for interventions. Annu. Rev. Med. 69, 301–318 (2018).

  162. 162.

    Ghoneim, H. E. et al. De novo epigenetic programs inhibit PD-1 blockade-mediated T cell rejuvenation. Cell 170, 142–157 (2017).

  163. 163.

    Balmer, M. L. et al. Memory CD8(+) T cells require increased concentrations of acetate induced by stress for optimal function. Immunity 44, 1312–1324 (2016).

  164. 164.

    Ghassemi, S. et al. Reducing ex vivo culture improves the antileukemic activity of chimeric antigen receptor (CAR) T cells. Cancer Immunol. Res. 6, 1100–1109 (2018).

  165. 165.

    Zheng, W. et al. PI3K orchestration of the in vivo persistence of chimeric antigen receptor-modified T cells. Leukemia 32, 1157–1167 (2018).

  166. 166.

    Klebanoff, C. A. et al. Inhibition of AKT signaling uncouples T cell differentiation from expansion for receptor-engineered adoptive immunotherapy. JCI Insight 2, 95103 (2017).

  167. 167.

    Sukumar, M. et al. Inhibiting glycolytic metabolism enhances CD8+T cell memory and antitumor function. J. Clin. Invest. 123, 4479–4488 (2013).

  168. 168.

    Cantor, J. R. et al. Physiologic medium rewires cellular metabolism and reveals uric acid as an endogenous inhibitor of UMP synthase. Cell 169, 258–272 (2017).

  169. 169.

    Ecker, C. et al. Differential reliance on lipid metabolism as a salvage pathway underlies functional differences of T cell subsets in poor nutrient environments. Cell Rep. 23, 741–755 (2018).

  170. 170.

    Xu, Y. et al. Closely related T-memory stem cells correlate with in vivo expansion of CAR. CD19-T cells and are preserved by IL-7 and IL-15. Blood 123, 3750–3759 (2014).

Download references


The authors thank M. D. Buck and members of the Pearce laboratories for discussion and critical reading of the manuscript. This work was supported by grants from the National Institutes of Health (NIH) (AI110481 to E.J.P.; AI091965 and CA158823 to E.L.P.) and the Max Planck Society.

Reviewer information

Nature Reviews Immunology thanks M. Haigis and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information

Author notes

  1. These authors contributed equally: David O’Sullivan, David E. Sanin.


  1. Max Planck Institute of Immunobiology and Epigenetics, Freiburg, Germany

    • David O’Sullivan
    • , David E. Sanin
    • , Edward J. Pearce
    •  & Erika L. Pearce
  2. University of Freiburg, Freiburg, Germany

    • David O’Sullivan
    • , David E. Sanin
    •  & Edward J. Pearce


  1. Search for David O’Sullivan in:

  2. Search for David E. Sanin in:

  3. Search for Edward J. Pearce in:

  4. Search for Erika L. Pearce in:


All the authors contributed to the discussion of content and to the writing, review and editing of the manuscript. D.O. and D.E.S. were involved in researching data for the article.

Competing interests

E.L.P. is a scientific advisory board member of Immunomet and a founder of Rheos Medicines. E.J.P. is a founder of Rheos Medicines.

Corresponding authors

Correspondence to Edward J. Pearce or Erika L. Pearce.


Warburg metabolism

Diversion of glucose metabolism towards lactate production in the presence of oxygen.

Oxidative phosphorylation

(OXPHOS). An electron transport chain-mediated process in which the energy resulting from the oxidation of carbon compounds is used to produce ATP.

Electron transport chain

Protein complexes spanning the inner mitochondrial membrane, which couple redox reactions to the establishment of a proton gradient used to generate ATP.

Pentose phosphate pathway

A metabolic pathway that generates NADPH and intermediates for nucleotide synthesis.


A laboratory procedure to extract the cellular component from blood.


A subset of the proteins in a cell that either contain a phosphate group or can be phosphorylated.


A type of biguanide that can induce AMPK activity and can disrupt the electron transport chain by reducing the activity of complex I.


A class of compounds that are used for the treatment of type 2 diabetes, as they regulate glucose metabolism.

About this article

Publication history



Further reading