Abstract
Several non-redundant features of the tumour microenvironment facilitate immunosuppression and limit anticancer immune responses. These include physical barriers to immune infiltration, the recruitment of suppressive immune cells and the upregulation of ligands on tumour cells that bind to inhibitory receptors on immune cells. Recent insights into the importance of the metabolic restrictions imposed by the tumour microenvironment on antitumour T cells have begun to inform immunotherapeutic anticancer strategies. Therapeutics that target metabolic restrictions, such as low glucose levels, a low pH, hypoxia and the generation of suppressive metabolites, have shown promise as combination therapies for different types of cancer. In this Review, we discuss the metabolic aspects of the antitumour T cell response in the context of immune checkpoint blockade, adoptive cell therapy and treatment with oncolytic viruses, and discuss emerging combination strategies.
This is a preview of subscription content, access via your institution
Access options
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$209.00 per year
only $17.42 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Muir, A. & Vander Heiden, M. G. The nutrient environment affects therapy. Science 360, 962–963 (2018).
Warburg, O., Wind, F. & Negelein, E. The metabolism of tumors in the body. J. Gen. Physiol. 8, 519–530 (1927).
DeBerardinis, R. J. & Chandel, N. S. We need to talk about the Warburg effect. Nat. Metab. 2, 127–129 (2020).
Ho, P.-C. et al. Phosphoenolpyruvate is a metabolic checkpoint of anti-tumor T cell responses. Cell 162, 1217–1228 (2015). This study shows the mechanism by which glucose deprivation in the tumour can restrict antitumour T cell activity.
Chang, C.-H. et al. Posttranscriptional control of T cell effector function by aerobic glycolysis. Cell 153, 1239–1251 (2013).
Fukumura, D., Duda, D. G., Munn, L. L. & Jain, R. K. Tumor microvasculature and microenvironment: novel insights through intravital imaging in pre-clinical models. Microcirculation 17, 206–225 (2010).
Hensley, C. T. et al. Metabolic heterogeneity in human lung tumors. Cell 164, 681–694 (2016).
DeBerardinis, R. J. & Chandel, N. S. Fundamentals of cancer metabolism. Sci. Adv. 2, e1600200 (2016).
Viallard, C. & Larrivée, B. Tumor angiogenesis and vascular normalization: alternative therapeutic targets. Angiogenesis 20, 409–426 (2017).
Sullivan, M. R. et al. Quantification of microenvironmental metabolites in murine cancers reveals determinants of tumor nutrient availability. eLife 8, e44235 (2019).
Erra Díaz, F., Dantas, E. & Geffner, J. Unravelling the interplay between extracellular acidosis and immune cells. Mediators Inflamm. 2018, 1218297 (2018).
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).
Angelin, A. et al. Foxp3 reprograms T cell metabolism to function in low-glucose, high-lactate environments. Cell Metab. 25, 1282–1293.e7 (2017). This study shows that FOXP3 can reprogramme T cells to downregulate glycolysis and upregulate OXPHOS to increase survival in low-glucose environments.
Colegio, O. R. et al. Functional polarization of tumour-associated macrophages by tumour-derived lactic acid. Nature 513, 559–563 (2014).
Sugiura, A. & Rathmell, J. C. Metabolic barriers to T cell function in tumors. J. Immunol. 200, 400–407 (2018).
Andrejeva, G. & Rathmell, J. C. Similarities and distinctions of cancer and immune metabolism in inflammation and tumors. Cell Metab. 26, 49–70 (2017).
Shyer, J. A., Flavell, R. A. & Bailis, W. Metabolic signaling in T cells. Cell Res. 30, 649–659 (2020).
Zhang, L. & Romero, P. Metabolic control of CD8+ T cell fate decisions and antitumor immunity. Trends Mol. Med. 24, 30–48 (2018).
Dai, Z., Ramesh, V. & Locasale, J. W. The evolving metabolic landscape of chromatin biology and epigenetics. Nat. Rev. Genet. 21, 737–753 (2020).
Wahl, D. R., Byersdorfer, C. A., Ferrara, J. L. M., Opipari, A. W. & Glick, G. D. Distinct metabolic programs in activated T cells: opportunities for selective immunomodulation. Immunol. Rev. 249, 104–115 (2012).
Kishton, R. J., Sukumar, M. & Restifo, N. P. Metabolic regulation of T cell longevity and function in tumor immunotherapy. Cell Metab. 26, 94–109 (2017).
Makowski, L., Chaib, M. & Rathmell, J. C. Immunometabolism: from basic mechanisms to translation. Immunol. Rev. 295, 5–14 (2020).
Menk, A. V. et al. Early TCR signaling induces rapid aerobic glycolysis enabling distinct acute T cell effector functions. Cell Rep. 22, 1509–1521 (2018).
Wang, R. et al. The transcription factor Myc controls metabolic reprogramming upon T lymphocyte activation. Immunity 35, 871–882 (2011). This article shows that MYC signalling is required for the induction of glycolysis and glutaminolysis that occurs with T cell activation.
MacIver, N. J., Michalek, R. D. & Rathmell, J. C. Metabolic regulation of T lymphocytes. Annu. Rev. Immunol. 31, 259–283 (2013).
Howie, D., Ten Bokum, A., Necula, A. S., Cobbold, S. P. & Waldmann, H. The role of lipid metabolism in T lymphocyte differentiation and survival. Front. Immunol. 8, 1949 (2017).
Michalek, R. D. et al. Cutting edge: distinct glycolytic and lipid oxidative metabolic programs are essential for effector and regulatory CD4+ T cell subsets. J. Immunol. 186, 3299–3303 (2011).
Kempkes, R. W. M., Joosten, I., Koenen, H. J. P. M. & He, X. Metabolic pathways involved in regulatory T cell functionality. Front. Immunol. 10, 2839 (2019).
Marelli-Berg, F. M. & Jangani, M. Metabolic regulation of leukocyte motility and migration. J. Leukoc. Biol. 104, 285–293 (2018).
van der Windt, G. J. W. et al. Mitochondrial respiratory capacity is a critical regulator of CD8+ T cell memory development. Immunity 36, 68–78 (2012). This article shows that IL-15 signalling promotes mitochondrial biogenesis and induction of CPT1A to increase fatty acid oxidation and spare respiratory capacity to enable long-lived memory cells.
Araki, K. et al. mTOR regulates memory CD8 T-cell differentiation. Nature 460, 108–112 (2009).
Kalbasi, A. & Ribas, A. Tumour-intrinsic resistance to immune checkpoint blockade. Nat. Rev. Immunol. 20, 25–39 (2020).
Pavlova, N. N. & Thompson, C. B. The emerging hallmarks of cancer metabolism. Cell Metab. 23, 27–47 (2016).
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).
Riley, J. L. PD-1 signaling in primary T cells. Immunol. Rev. 229, 114–125 (2009).
Patsoukis, N. et al. PD-1 alters T-cell metabolic reprogramming by inhibiting glycolysis and promoting lipolysis and fatty acid oxidation. Nat. Commun. 6, 6692 (2015).
Ogando, J. et al. PD-1 signaling affects cristae morphology and leads to mitochondrial dysfunction in human CD8+ T lymphocytes. J. Immunother. Cancer 7, 151 (2019).
Tkachev, V. et al. Programmed death-1 controls T cell survival by regulating oxidative metabolism. J. Immunol. 194, 5789–5800 (2015).
Spinelli, J. B. & Haigis, M. C. The multifaceted contributions of mitochondria to cellular metabolism. Nat. Cell Biol. 20, 745–754 (2018).
Scharping, N. E. et al. The tumor microenvironment represses T cell mitochondrial biogenesis to drive intratumoral T cell metabolic insufficiency and dysfunction. Immunity 45, 374–388 (2016). This article shows that tumour-infiltrating T cells lose their capacity to generate mitochondria through the loss of PGC1α, which controls mitochondrial biogenesis, and that restoration of PGC1α restores T cell function.
Bengsch, B. et al. Bioenergetic insufficiencies due to metabolic alterations regulated by the inhibitory receptor PD-1 are an early driver of CD8+ T cell exhaustion. Immunity 45, 358–373 (2016).
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).
Siska, P. J. et al. Mitochondrial dysregulation and glycolytic insufficiency functionally impair CD8 T cells infiltrating human renal cell carcinoma. JCI Insight 2, e93411 (2017).
Chowdhury, P. S., Chamoto, K., Kumar, A. & Honjo, T. PPAR-induced fatty acid oxidation in T cells increases the number of tumor-reactive CD8+ T cells and facilitates anti-PD-1 therapy. Cancer Immunol. Res. 6, 1375–1387 (2018).
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).
Teijeira, A. et al. Mitochondrial morphological and functional reprogramming following CD137 (4-1BB) costimulation. cancer. Immunol. Res. 6, 798–811 (2018).
Scharping, N. E. et al. Mitochondrial stress induced by continuous stimulation under hypoxia rapidly drives T cell exhaustion. Nat. Immunol. 22, 205–215 (2021). Together with Yu et al. (2020) and Vardhana et al. (2020) this article shows that T cell mitochondrial reactive oxygen species, triggered by chronic antigen stimulation, drive epigenetic alterations and T cell exhaustion.
Yu, Y.-R. et al. Disturbed mitochondrial dynamics in CD8+ TILs reinforce T cell exhaustion. Nat. Immunol. 21, 1540–1551 (2020).
Vardhana, S. A. et al. Impaired mitochondrial oxidative phosphorylation limits the self-renewal of T cells exposed to persistent antigen. Nat. Immunol. 21, 1022–1033 (2020).
McKeown, S. R. Defining normoxia, physoxia and hypoxia in tumours-implications for treatment response. Br. J. Radiol. 87, 20130676 (2014).
Bertout, J. A., Patel, S. A. & Simon, M. C. The impact of O2 availability on human cancer. Nat. Rev. Cancer 8, 967–975 (2008).
Najjar, Y. G. et al. Tumor cell oxidative metabolism as a barrier to PD-1 blockade immunotherapy in melanoma. JCI Insight 4, e124989 (2019). This article shows in patients with melanoma that oxidative tumours, which are more hypoxic than glycolytic tumours, are resistant to PD1 blockade through a more exhausted T cell phenotype.
Hatfield, S. M. et al. Immunological mechanisms of the antitumor effects of supplemental oxygenation. Sci. Transl Med. 7, 277ra30 (2015).
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).
Jayaprakash, P. et al. Targeted hypoxia reduction restores T cell infiltration and sensitizes prostate cancer to immunotherapy. J. Clin. Invest. 128, 5137–5149 (2018).
Voron, T. et al. VEGF-A modulates expression of inhibitory checkpoints on CD8+ T cells in tumors. J. Exp. Med. 212, 139–148 (2015).
Ward, P. S. & Thompson, C. B. Metabolic reprogramming: a cancer hallmark even Warburg did not anticipate. Cancer Cell 21, 297–308 (2012).
Gerriets, V. A. et al. Metabolic programming and PDHK1 control CD4+ T cell subsets and inflammation. J. Clin. Invest. 125, 194–207 (2015).
Fischer, K. et al. Inhibitory effect of tumor cell-derived lactic acid on human T cells. Blood 109, 3812–3819 (2007).
Watson, M. J. et al. Metabolic support of tumour-infiltrating regulatory T cells by lactic acid. Nature https://doi.org/10.1038/s41586-020-03045-2 (2021).
Renner, K. et al. Restricting glycolysis preserves T cell effector functions and augments checkpoint therapy. Cell Rep. 29, 135–150.e9 (2019).
Cluntun, A. A., Lukey, M. J., Cerione, R. A. & Locasale, J. W. Glutamine metabolism in cancer: understanding the heterogeneity. Trends Cancer 3, 169–180 (2017).
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).
Leone, R. D. et al. Glutamine blockade induces divergent metabolic programs to overcome tumor immune evasion. Science 366, 1013–1021 (2019).
Roy, D. G. et al. Methionine metabolism shapes T helper cell responses through regulation of epigenetic reprogramming. Cell Metab. 31, 250–266.e9 (2020).
Bian, Y. et al. Cancer SLC43A2 alters T cell methionine metabolism and histone methylation. Nature 585, 277–282 (2020).
Tripathi, P. et al. STAT5 is critical to maintain effector CD8+ T cell responses. J. Immunol. 185, 2116–2124 (2010).
Emran, A. A. et al. Targeting DNA methylation and EZH2 activity to overcome melanoma resistance to immunotherapy. Trends Immunol. 40, 328–344 (2019).
Rodriguez, P. C. et al. L-arginine consumption by macrophages modulates the expression of CD3 zeta chain in T lymphocytes. J. Immunol. 171, 1232–1239 (2003).
Grzywa, T. M. et al. Myeloid cell-derived arginase in cancer immune response. Front. Immunol. 11, 938 (2020).
Porter, D. L. et al. Chimeric antigen receptor T cells persist and induce sustained remissions in relapsed refractory chronic lymphocytic leukemia. Sci. Transl Med. 7, 303ra139 (2015).
Lugli, E. et al. Superior T memory stem cell persistence supports long-lived T cell memory. J. Clin. Invest. 123, 594–599 (2013). This article demonstrates that a pool of stem cell memory T cells are the precursors to central memory cells.
Berger, C. et al. Adoptive transfer of effector CD8+ T cells derived from central memory cells establishes persistent T cell memory in primates. J. Clin. Invest. 118, 294–305 (2008).
Sukumar, M. et al. Inhibiting glycolytic metabolism enhances CD8+ T cell memory and antitumor function. J. Clin. Invest. 123, 4479–4488 (2013).
Pilipow, K. et al. Antioxidant metabolism regulates CD8+ T memory stem cell formation and antitumor immunity. JCI Insight 3, e122299 (2018).
Sabatino, M. et al. Generation of clinical-grade CD19-specific CAR-modified CD8+ memory stem cells for the treatment of human B-cell malignancies. Blood 128, 519–528 (2016).
Alizadeh, D. et al. IL15 enhances CAR-T cell antitumor activity by reducing mTORC1 activity and preserving their stem cell memory phenotype. Cancer Immunol. Res. 7, 759–772 (2019).
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).
Geiger, R. et al. L-arginine modulates T cell metabolism and enhances survival and anti-tumor activity. Cell 167, 829–842.e13 (2016). This articles shows that elevating l-arginine levels can shift T cell metabolism from glycolysis to OXPHOS and support the antitumour function of T cells.
Rao, R. R., Li, Q., Odunsi, K. & Shrikant, P. A. The mTOR kinase determines effector versus memory CD8+ T cell fate by regulating the expression of transcription factors T-bet and eomesodermin. Immunity 32, 67–78 (2010).
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).
Ackermann, T. & Tardito, S. Cell culture medium formulation and its implications in cancer metabolism. Trends Cancer 5, 329–332 (2019).
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).
Cascone, T. et al. Increased tumor glycolysis characterizes immune resistance to adoptive T cell therapy. Cell Metab. 27, 977–987.e4 (2018).
Nabe, S. et al. Reinforce the antitumor activity of CD8+ T cells via glutamine restriction. Cancer Sci. 109, 3737–3750 (2018).
Majmundar, A. J., Wong, W. J. & Simon, M. C. Hypoxia-inducible factors and the response to hypoxic stress. Mol. Cell 40, 294–309 (2010).
Berahovich, R. et al. Hypoxia selectively impairs CAR-T cells in vitro. Cancers 11, 602 (2019).
Gropper, Y. et al. Culturing CTLs under hypoxic conditions enhances their cytolysis and improves their anti-tumor function. Cell Rep. 20, 2547–2555 (2017).
Palazon, A. et al. An HIF-1α/VEGF-A axis in cytotoxic t cells regulates tumor progression. Cancer Cell 32, 669–683.e5 (2017).
Xu, Y. et al. Glycolysis determines dichotomous regulation of T cell subsets in hypoxia. J. Clin. Invest. 126, 2678–2688 (2016).
Ohta, A. et al. A2A adenosine receptor protects tumors from antitumor T cells. Proc. Natl Acad. Sci. USA 103, 13132–13137 (2006).
Huang, S., Apasov, S., Koshiba, M. & Sitkovsky, M. Role of A2a extracellular adenosine receptor-mediated signaling in adenosine-mediated inhibition of T-cell activation and expansion. Blood 90, 1600–1610 (1997).
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).
Hatfield, S. M. & Sitkovsky, M. A2A adenosine receptor antagonists to weaken the hypoxia-HIF-1α driven immunosuppression and improve immunotherapies of cancer. Curr. Opin. Pharmacol. 29, 90–96 (2016).
Beavis, P. A. et al. Targeting the adenosine 2A receptor enhances chimeric antigen receptor T cell efficacy. J. Clin. Invest. 127, 929–941 (2017).
Kato, Y. et al. Acidic extracellular microenvironment and cancer. Cancer Cell Int. 13, 89 (2013).
Kunzelmann, K. Ion channels and cancer. J. Membr. Biol. 205, 159–173 (2005).
Kunzelmann, K. Ion channels in regulated cell death. Cell Mol. Life Sci. 73, 2387–2403 (2016).
Eil, R. et al. Ionic immune suppression within the tumour microenvironment limits T cell effector function. Nature 537, 539–543 (2016). This article shows that necrosis within the tumour can release K+ ions into the microenvironment, which interferes with T cell receptor signalling.
Omilusik, K. et al. The CaV1.4 calcium channel is a critical regulator of T cell receptor signaling and naive T cell homeostasis. Immunity 35, 349–360 (2011).
Trebak, M. & Kinet, J.-P. Calcium signalling in T cells. Nat. Rev. Immunol. 19, 154–169 (2019).
Fultang, L. et al. Metabolic engineering against the arginine microenvironment enhances CAR-T cell proliferation and therapeutic activity. Blood 136, 1155–1160 (2020).
Bommareddy, P. K., Shettigar, M. & Kaufman, H. L. Integrating oncolytic viruses in combination cancer immunotherapy. Nat. Rev. Immunol. 18, 498–513 (2018).
Davola, M. E. & Mossman, K. L. Oncolytic viruses: how “lytic” must they be for therapeutic efficacy? Oncoimmunology 8, e1581528 (2019).
Meng, G. et al. Targeting aerobic glycolysis by dichloroacetate improves Newcastle disease virus-mediated viro-immunotherapy in hepatocellular carcinoma. Br. J. Cancer 122, 111–120 (2020).
Li, C. et al. Dichloroacetate blocks aerobic glycolytic adaptation to attenuated measles virus and promotes viral replication leading to enhanced oncolysis in glioblastoma. Oncotarget 6, 1544–1555 (2015).
Kennedy, B. E. et al. Inhibition of pyruvate dehydrogenase kinase enhances the antitumor efficacy of oncolytic reovirus. Cancer Res. 79, 3824–3836 (2019).
Dyer, A. et al. Antagonism of glycolysis and reductive carboxylation of glutamine potentiates activity of oncolytic adenoviruses in cancer cells. Cancer Res. 79, 331–345 (2019).
Pant, A., Cao, S. & Yang, Z. Asparagine is a critical limiting metabolite for vaccinia virus protein synthesis during glutamine deprivation. J. Virol. https://doi.org/10.1128/JVI.01834-18 (2019).
Greseth, M. D. & Traktman, P. De novo fatty acid biosynthesis contributes significantly to establishment of a bioenergetically favorable environment for vaccinia virus infection. PLoS Pathog. 10, e1004021 (2014).
Liang, J. et al. Inhibition of the mevalonate pathway enhances cancer cell oncolysis mediated by M1 virus. Nat. Commun. 9, 1524 (2018).
Moskovskich, A. et al. The transporters SLC35A1 and SLC30A1 play opposite roles in cell survival upon VSV virus infection. Sci. Rep. 9, 10471 (2019).
Andtbacka, R. H. I. et al. Talimogene laherparepvec improves durable response rate in patients with advanced melanoma. J. Clin. Oncol. 33, 2780–2788 (2015).
Fountzilas, C., Patel, S. & Mahalingam, D. Review: oncolytic virotherapy, updates and future directions. Oncotarget 8, 102617–102639 (2017).
Rivadeneira, D. B. et al. Oncolytic viruses engineered to enforce leptin expression reprogram tumor-infiltrating T cell metabolism and promote tumor clearance. Immunity 51, 548–560.e4 (2019).
Matuszewska, K. et al. Combining vascular normalization with an oncolytic virus enhances immunotherapy in a preclinical model of advanced-stage ovarian cancer. Clin. Cancer Res. 25, 1624–1638 (2019).
Twumasi-Boateng, K., Pettigrew, J. L., Kwok, Y. Y. E., Bell, J. C. & Nelson, B. H. Oncolytic viruses as engineering platforms for combination immunotherapy. Nat. Rev. Cancer 18, 419–432 (2018).
Guo, Z. S. et al. Oncolytic immunotherapy: conceptual evolution, current strategies, and future perspectives. Front. Immunol. 8, 555 (2017).
Guedan, S. & Alemany, R. CAR-T cells and oncolytic viruses: joining forces to overcome the solid tumor challenge. Front. Immunol. 9, 2460 (2018).
Nishio, N. et al. Armed oncolytic virus enhances immune functions of chimeric antigen receptor-modified T cells in solid tumors. Cancer Res. 74, 5195–5205 (2014).
Watanabe, K. et al. Pancreatic cancer therapy with combined mesothelin-redirected chimeric antigen receptor T cells and cytokine-armed oncolytic adenoviruses. JCI Insight 3, e99573 (2018).
Warmoes, M. O. & Locasale, J. W. Heterogeneity of glycolysis in cancers and therapeutic opportunities. Biochem. Pharmacol. 92, 12–21 (2014).
Cordain, L. et al. Origins and evolution of the Western diet: health implications for the 21st century. Am. J. Clin. Nutr. 81, 341–354 (2005).
Deng, T., Lyon, C. J., Bergin, S., Caligiuri, M. A. & Hsueh, W. A. Obesity, inflammation, and cancer. Annu. Rev. Pathol. 11, 421–449 (2016).
Moore, L. L., Chadid, S., Singer, M. R., Kreger, B. E. & Denis, G. V. Metabolic health reduces risk of obesity-related cancer in Framingham study adults. Cancer Epidemiol. Biomarkers Prev. 23, 2057–2065 (2014).
McQuade, J. L. et al. Association of body-mass index and outcomes in patients with metastatic melanoma treated with targeted therapy, immunotherapy, or chemotherapy: a retrospective, multicohort analysis. Lancet Oncol. 19, 310–322 (2018). This article, along with Cortellini et al. (2019) and Wang et al. (2019), shows that patients with obesity based on body mass index respond better to immunotherapy than patients with normal body mass index.
Cortellini, A. et al. A multicenter study of body mass index in cancer patients treated with anti-PD-1/PD-L1 immune checkpoint inhibitors: when overweight becomes favorable. J. Immunother. Cancer 7, 57 (2019).
Wang, Z. et al. Paradoxical effects of obesity on T cell function during tumor progression and PD-1 checkpoint blockade. Nat. Med. 25, 141–151 (2019).
Matarese, G. & La Cava, A. The intricate interface between immune system and metabolism. Trends Immunol. 25, 193–200 (2004).
Cao, H. Adipocytokines in obesity and metabolic disease. J. Endocrinol. 220, T47–T59 (2014).
Jung, U. J. & Choi, M.-S. Obesity and its metabolic complications: the role of adipokines and the relationship between obesity, inflammation, insulin resistance, dyslipidemia and nonalcoholic fatty liver disease. Int. J. Mol. Sci. 15, 6184–6223 (2014).
Weidinger, C., Ziegler, J. F., Letizia, M., Schmidt, F. & Siegmund, B. Adipokines and their role in intestinal inflammation. Front. Immunol. 9, 1974 (2018).
Francisco, V. et al. Adipokines and inflammation: is it a question of weight? Br. J. Pharmacol. 175, 1569–1579 (2018).
Wikoff, W. R. et al. Metabolomics analysis reveals large effects of gut microflora on mammalian blood metabolites. Proc. Natl Acad. Sci. USA 106, 3698–3703 (2009).
Chang, P. V., Hao, L., Offermanns, S. & Medzhitov, R. The microbial metabolite butyrate regulates intestinal macrophage function via histone deacetylase inhibition. Proc. Natl Acad. Sci. USA 111, 2247–2252 (2014).
Furusawa, Y. et al. Commensal microbe-derived butyrate induces the differentiation of colonic regulatory T cells. Nature 504, 446–450 (2013). This article shows that butyrate produced in the gut by commensal microorganisms can induce Treg cell differentiation and maintain gut homeostasis.
Smith, P. M. et al. The microbial metabolites, short-chain fatty acids, regulate colonic Treg cell homeostasis. Science 341, 569–573 (2013).
Luu, M. et al. Regulation of the effector function of CD8+ T cells by gut microbiota-derived metabolite butyrate. Sci. Rep. 8, 14430 (2018).
Bachem, A. et al. Microbiota-derived short-chain fatty acids promote the memory potential of antigen-activated CD8+ T cells. Immunity 51, 285–297.e5 (2019). This article shows that germ-free mice have impaired memory T cell responses and that microbially derived butyrate promotes CD8+ T cell memory formation by shifting the metabolism of CD8+ T cells to OXPHOS.
Sun, M. et al. Microbiota-derived short-chain fatty acids promote Th1 cell IL-10 production to maintain intestinal homeostasis. Nat. Commun. 9, 3555 (2018).
Zmora, N., Suez, J. & Elinav, E. You are what you eat: diet, health and the gut microbiota. Nat. Rev. Gastroenterol. Hepatol. 16, 35–56 (2019).
Castaner, O. et al. The gut microbiome profile in obesity: a systematic review. Int. J. Endocrinol. 2018, 4095789 (2018).
David, L. A. et al. Diet rapidly and reproducibly alters the human gut microbiome. Nature 505, 559–563 (2014).
Hodi, F. S. et al. Improved survival with ipilimumab in patients with metastatic melanoma. N. Engl. J. Med. 363, 711–723 (2010). The clinical trial results in this article resulted in the approval of ipilimumab for treatment of patients with melanoma.
Topalian, S. L. et al. Safety, activity, and immune correlates of anti-PD-1 antibody in cancer. N. Engl. J. Med. 366, 2443–2454 (2012).
Harding, F. A., McArthur, J. G., Gross, J. A., Raulet, D. H. & Allison, J. P. CD28-mediated signalling co-stimulates murine T cells and prevents induction of anergy in T-cell clones. Nature 356, 607–609 (1992).
Brunet, J. F. et al. A new member of the immunoglobulin superfamily–CTLA-4. Nature 328, 267–270 (1987).
Linsley, P. S. et al. Human B7-1 (CD80) and B7-2 (CD86) bind with similar avidities but distinct kinetics to CD28 and CTLA-4 receptors. Immunity 1, 793–801 (1994).
Linsley, P. S. et al. CTLA-4 is a second receptor for the B cell activation antigen B7. J. Exp. Med. 174, 561–569 (1991).
Lenschow, D. J. et al. Long-term survival of xenogeneic pancreatic islet grafts induced by CTLA4lg. Science 257, 789–792 (1992).
Qureshi, O. S. et al. Trans-endocytosis of CD80 and CD86: a molecular basis for the cell-extrinsic function of CTLA-4. Science 332, 600–603 (2011).
Takahashi, T. et al. Immunologic self-tolerance maintained by CD25+CD4+ regulatory T cells constitutively expressing cytotoxic T lymphocyte-associated antigen 4. J. Exp. Med. 192, 303–310 (2000).
Tivol, E. A. et al. Loss of CTLA-4 leads to massive lymphoproliferation and fatal multiorgan tissue destruction, revealing a critical negative regulatory role of CTLA-4. Immunity 3, 541–547 (1995).
Waterhouse, P. et al. Lymphoproliferative disorders with early lethality in mice deficient in Ctla-4. Science 270, 985–988 (1995).
Wing, K. et al. CTLA-4 control over Foxp3+ regulatory T cell function. Science 322, 271–275 (2008).
Leach, D. R., Krummel, M. F. & Allison, J. P. Enhancement of antitumor immunity by CTLA-4 blockade. Science 271, 1734–1736 (1996).
Wei, S. C., Duffy, C. R. & Allison, J. P. Fundamental mechanisms of immune checkpoint blockade therapy. Cancer Discov. 8, 1069–1086 (2018).
Ishida, Y., Agata, Y., Shibahara, K. & Honjo, T. Induced expression of PD-1, a novel member of the immunoglobulin gene superfamily, upon programmed cell death. EMBO J. 11, 3887–3895 (1992).
Nishimura, H., Honjo, T. & Minato, N. Facilitation of beta selection and modification of positive selection in the thymus of PD-1-deficient mice. J. Exp. Med. 191, 891–898 (2000).
Nishimura, H. et al. Developmentally regulated expression of the PD-1 protein on the surface of double-negative (CD4-CD8-) thymocytes. Int. Immunol. 8, 773–780 (1996).
Agata, Y. et al. Expression of the PD-1 antigen on the surface of stimulated mouse T and B lymphocytes. Int. Immunol. 8, 765–772 (1996).
Freeman, G. J. et al. Engagement of the PD-1 immunoinhibitory receptor by a novel B7 family member leads to negative regulation of lymphocyte activation. J. Exp. Med. 192, 1027–1034 (2000).
Latchman, Y. et al. PD-L2 is a second ligand for PD-1 and inhibits T cell activation. Nat. Immunol. 2, 261–268 (2001).
Sharpe, A. H. & Pauken, K. E. The diverse functions of the PD1 inhibitory pathway. Nat. Rev. Immunol. 18, 153–167 (2018).
Pardoll, D. M. The blockade of immune checkpoints in cancer immunotherapy. Nat. Rev. Cancer 12, 252–264 (2012).
Ribas, A. Adaptive immune resistance: how cancer protects from immune attack. Cancer Discov. 5, 915–919 (2015).
Wherry, E. J. & Kurachi, M. Molecular and cellular insights into T cell exhaustion. Nat. Rev. Immunol. 15, 486–499 (2015).
McLane, L. M., Abdel-Hakeem, M. S. & Wherry, E. J. CD8 T cell exhaustion during chronic viral infection and cancer. Annu. Rev. Immunol. 37, 457–495 (2019).
Zajac, A. J. et al. Viral immune evasion due to persistence of activated T cells without effector function. J. Exp. Med. 188, 2205–2213 (1998).
Gallimore, A. et al. Induction and exhaustion of lymphocytic choriomeningitis virus-specific cytotoxic T lymphocytes visualized using soluble tetrameric major histocompatibility complex class I-peptide complexes. J. Exp. Med. 187, 1383–1393 (1998).
Lee, P. P. et al. Characterization of circulating T cells specific for tumor-associated antigens in melanoma patients. Nat. Med. 5, 677–685 (1999).
Barber, D. L. et al. Restoring function in exhausted CD8 T cells during chronic viral infection. Nature 439, 682–687 (2006).
Iwai, Y. et al. Involvement of PD-L1 on tumor cells in the escape from host immune system and tumor immunotherapy by PD-L1 blockade. Proc. Natl Acad. Sci. USA 99, 12293–12297 (2002).
Miller, B. C. et al. Subsets of exhausted CD8+ T cells differentially mediate tumor control and respond to checkpoint blockade. Nat. Immunol. 20, 326–336 (2019).
Sade-Feldman, M. et al. Defining T cell states associated with response to checkpoint immunotherapy in melanoma. Cell 175, 998–1013.e20 (2018). This article shows the correlation of TCF7+ T cells and response to immunotherapy.
Chen, Z. et al. TCF-1-centered transcriptional network drives an effector versus exhausted CD8 T cell-fate decision. Immunity 51, 840–855.e5 (2019). This article, along with Miller et al. (2019) and Sade-Feldman (2018), demonstrates the importance of a TCF7+ progenitor CD8+ T cell population in response to anti-PD1 therapy (TCF7 is also known as TCF1).
Sadelain, M. CD19 CAR T cells. Cell 171, 1471 (2017).
Rosenberg, S. A. et al. Durable complete responses in heavily pretreated patients with metastatic melanoma using T-cell transfer immunotherapy. Clin. Cancer Res. 17, 4550–4557 (2011).
Guedan, S., Ruella, M. & June, C. H. Emerging cellular therapies for cancer. Annu. Rev. Immunol. 37, 145–171 (2019).
Schuster, S. J. et al. Chimeric antigen receptor T cells in refractory B-cell lymphomas. N. Engl. J. Med. 377, 2545–2554 (2017).
Kalos, M. et al. T cells with chimeric antigen receptors have potent antitumor effects and can establish memory in patients with advanced leukemia. Sci. Transl Med. 3, 95ra73 (2011).
Brentjens, R. J. et al. CD19-targeted T cells rapidly induce molecular remissions in adults with chemotherapy-refractory acute lymphoblastic leukemia. Sci. Transl Med. 5, 177ra38 (2013).
Maude, S. L. et al. Chimeric antigen receptor T cells for sustained remissions in leukemia. N. Engl. J. Med. 371, 1507–1517 (2014).
Park, J. H. et al. Long-term follow-up of CD19 CAR therapy in acute lymphoblastic Leukemia. N. Engl. J. Med. 378, 449–459 (2018).
Turtle, C. J. et al. CD19 CAR-T cells of defined CD4+:CD8+composition in adult B cell ALL patients. J. Clin. Invest. 126, 2123–2138 (2016).
Lee, D. W. et al. T cells expressing CD19 chimeric antigen receptors for acute lymphoblastic leukaemia in children and young adults: a phase 1 dose-escalation trial. Lancet 385, 517–528 (2015).
Maude, S. L. et al. Tisagenlecleucel in children and young adults with B-cell lymphoblastic leukemia. N. Engl. J. Med. 378, 439–448 (2018).
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).
Hamid, O., Ismail, R. & Puzanov, I. Intratumoral immunotherapy — update 2019. Oncologist 25, e423–e438 (2020).
Filley, A. C. & Dey, M. Immune system, friend or foe of oncolytic virotherapy? Front. Oncol. 7, 106 (2017).
Lemos de Matos, A., Franco, L. S. & McFadden, G. Oncolytic viruses and the immune system: the dynamic duo. Mol. Ther. Methods Clin. Dev. 17, 349–358 (2020).
Gujar, S., Pol, J. G., Kim, Y., Lee, P. W. & Kroemer, G. Antitumor benefits of antiviral immunity: an underappreciated aspect of oncolytic virotherapies. Trends Immunol. 39, 209–221 (2018).
Rojas, J. J., Sampath, P., Hou, W. & Thorne, S. H. Defining effective combinations of immune checkpoint blockade and oncolytic virotherapy. Clin. Cancer Res. 21, 5543–5551 (2015).
Woller, N. et al. Viral infection of tumors overcomes resistance to PD-1-immunotherapy by broadening neoantigenome-directed T-cell responses. Mol. Ther. 23, 1630–1640 (2015).
Ricca, J. M. et al. Pre-existing immunity to oncolytic virus potentiates its immunotherapeutic efficacy. Mol. Ther. 26, 1008–1019 (2018).
Liu, Z., Ravindranathan, R., Kalinski, P., Guo, Z. S. & Bartlett, D. L. Rational combination of oncolytic vaccinia virus and PD-L1 blockade works synergistically to enhance therapeutic efficacy. Nat. Commun. 8, 14754 (2017).
Nakao, S. et al. Intratumoral expression of IL-7 and IL-12 using an oncolytic virus increases systemic sensitivity to immune checkpoint blockade. Sci. Transl Med. 12, eaax7992 (2020).
Zamarin, D. et al. Localized oncolytic virotherapy overcomes systemic tumor resistance to immune checkpoint blockade immunotherapy. Sci. Transl Med. 6, 226ra32 (2014).
Oh, E., Choi, I.-K., Hong, J. & Yun, C.-O. Oncolytic adenovirus coexpressing interleukin-12 and decorin overcomes Treg-mediated immunosuppression inducing potent antitumor effects in a weakly immunogenic tumor model. Oncotarget 8, 4730–4746 (2017).
Fend, L. et al. Oncolytic virotherapy with an armed vaccinia virus in an orthotopic model of renal carcinoma is associated with modification of the tumor microenvironment. Oncoimmunology 5, e1080414 (2016).
Patel, M. R. et al. Vesicular stomatitis virus expressing interferon-β is oncolytic and promotes antitumor immune responses in a syngeneic murine model of non-small cell lung cancer. Oncotarget 6, 33165–33177 (2015).
Acknowledgements
The authors thank members of the Delgoffe laboratory for helpful discussions during the course of writing this Review. K.D. was supported by T32CA082084 and 1F31CA247129. G.M.D. is supported by an NIH Director’s New Innovator Award (DP2AI136598), the Mark Foundation for Cancer Research Emerging Leader Award, the Cancer Research Institute Lloyd J. Old STAR Award and the Sy Holzer Endowed Immunotherapy Fund.
Author information
Authors and Affiliations
Contributions
The authors contributed equally to all aspects of the article.
Corresponding author
Ethics declarations
Competing interests
G.M.D. declares competing financial interests and has submitted patents covering the use of PGC1α in cell therapies, metabolic manipulation of culture conditions for cell therapies, and the use of leptin and other adipokines in oncolytic viruses that are licensed or pending and is entitled to a share in net income generated from the licencing of these patent rights for commercial development. G.M.D. consults for and/or is on the scientific advisory boards of BlueSphere Bio, Century Therapeutics, Novasenta, Pieris Pharmaceuticals and Kalivir, has grants from bluebird bio, Novasenta, Pfizer, Pieris Pharmaceuticals, TCR2 and Western Oncolytics/Kalivir, and owns stock in Novasenta and BlueSphere Bio. K.D. declares no competing interests.
Additional information
Peer review information
Nature Reviews Immunology thanks Jeffrey Rathmell and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Publisher’s note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Glossary
- Immune editing
-
The process by which tumour cells mutate and evolve to evade detection and elimination by the immune system.
- Tumoural escape
-
The ability of tumour cells to escape from elimination by immune cells, for example through immune editing.
- Glycolysis
-
The process by which cytosolic glucose is converted into pyruvate, which is then shuttled into the mitochondria, where it feeds into the tricarboxylic acid cycle. However, under conditions of low oxygen, pyruvate is usually fermented into lactic acid, which regenerates oxidized nicotinamide adenine dinucleotide (NAD+) and accelerates upstream glycolysis. Otto Warburg discovered that in tumour cells this fermentation can also occur under oxygen-sufficient conditions. This phenomenon is termed ‘aerobic glycolysis’ or the ‘Warburg effect’.
- Oxidative phosphorylation
-
(OXPHOS). The process by which metabolic intermediates from the tricarboxylic acid cycle are oxidized, generating a proton gradient that powers ATP synthase to produce ATP.
- Tricarboxylic acid (TCA) cycle
-
A series of biochemical reactions in which acetyl-CoA is metabolized to generate reducing intermediates for the electron transport chain (generating ATP through oxidative phosphorylation), as well as numerous metabolic intermediates for various synthetic pathways. Acetyl-CoA can be fed into the TCA cycle by the decarboxylation of pyruvate, β-oxidation of fatty acids or α-ketoglutarate through the production of amino acids.
- Fatty acid oxidation
-
The process by which fatty acid chains are broken down by β-oxidation and subsequently combined with CoA to form acetyl-CoA and enter the tricarboxylic acid cycle.
- PI3K–AKT–mTOR axis
-
A signalling pathway that is activated through extracellular or intracellular stimulation of phosphoinositide-3 kinase (PI3K), which leads to the activation of AKT (also known as protein kinase B) and mechanistic target of rapamycin (mTOR). The manner in which PI3K and AKT are activated can alter the downstream effects of mTOR activation and the activity of its various ternary complexes. This axis controls proliferation, translation, the cell cycle, glucose metabolism, autophagy and cell survival.
- Flux analysis
-
The analysis of the oxygen consumption rate and extracellular acidification rate of a cell population to determine its rate of glycolysis or oxidative phosphorylation.
Rights and permissions
About this article
Cite this article
DePeaux, K., Delgoffe, G.M. Metabolic barriers to cancer immunotherapy. Nat Rev Immunol 21, 785–797 (2021). https://doi.org/10.1038/s41577-021-00541-y
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41577-021-00541-y
This article is cited by
-
The quest for nanoparticle-powered vaccines in cancer immunotherapy
Journal of Nanobiotechnology (2024)
-
Shifting the paradigm: engaging multicellular networks for cancer therapy
Journal of Translational Medicine (2024)
-
Activation of mucosal insulin receptor exacerbates intestinal inflammation by promoting tissue resident memory T cells differentiation through EZH2
Journal of Translational Medicine (2024)
-
Cancer metabolism and carcinogenesis
Experimental Hematology & Oncology (2024)
-
Effects of dietary intervention on human diseases: molecular mechanisms and therapeutic potential
Signal Transduction and Targeted Therapy (2024)
