Review Article | Published:

Immunometabolism at the interface between macrophages and pathogens

Nature Reviews Immunology (2019) | Download Citation


It is generally regarded that the progression of an infection within host macrophages is the consequence of a failed immune response. However, recent appreciation of macrophage heterogeneity, with respect to both development and metabolism, indicates that the reality is more complex. Different lineages of tissue-resident macrophages respond divergently to microbial, environmental and immunological stimuli. The emerging picture that the developmental origin of macrophages determines their responses to immune stimulation and to infection stresses the importance of in vivo infection models. Recent investigations into the metabolism of infecting microorganisms and host macrophages indicate that their metabolic interface can be a major determinant of pathogen growth or containment. This Review focuses on the integration of data from existing studies, the identification of challenges in generating and interpreting data from ongoing studies and a discussion of the technologies and tools that are required to best address future questions in the field.

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.

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

  2. 2.

    Van den Bossche, J., O’Neill, L. A. & Menon, D. Macrophage immunometabolism: where are we (going)? Trends. Immunol. 38, 395–406 (2017).

  3. 3.

    VanderVen, B. C., Yates, R. M. & Russell, D. G. Intraphagosomal measurement of the magnitude and duration of the oxidative burst. Traffic 10, 372–378 (2009).

  4. 4.

    van Furth, R. & Cohn, Z. A. The origin and kinetics of mononuclear phagocytes. J. Exp. Med. 128, 415–435 (1968).

  5. 5.

    van Furth, R. et al. The mononuclear phagocyte system: a new classification of macrophages, monocytes, and their precursor cells. Bull. World Health Organ. 46, 845–852 (1972).

  6. 6.

    Munder, M., Eichmann, K. & Modolell, M. Alternative metabolic states in murine macrophages reflected by the nitric oxide synthase/arginase balance: competitive regulation by CD4+ T cells correlates with Th1/Th2 phenotype. J. Immunol. 160, 5347–5354 (1998).

  7. 7.

    Gordon, S. & Martinez-Pomares, L. Physiological roles of macrophages. Pflugers Arch. 469, 365–374 (2017).

  8. 8.

    Sica, A. & Mantovani, A. Macrophage plasticity and polarization: in vivo veritas. J. Clin. Invest. 122, 787–795 (2012).

  9. 9.

    Xue, J. et al. Transcriptome-based network analysis reveals a spectrum model of human macrophage activation. Immunity 40, 274–288 (2014).

  10. 10.

    Ginhoux, F. & Guilliams, M. Tissue-resident macrophage ontogeny and homeostasis. Immunity 44, 439–449 (2016).

  11. 11.

    Ginhoux, F. & Jung, S. Monocytes and macrophages: developmental pathways and tissue homeostasis. Nat. Rev. Immunol. 14, 392–404 (2014).

  12. 12.

    Gordon, S. & Pluddemann, A. Tissue macrophages: heterogeneity and functions. BMC Biol. 15, 53 (2017).

  13. 13.

    Diskin, C. & Palsson-McDermott, E. M. Metabolic modulation in macrophage effector function. Front. Immunol. 9, 270 (2018).

  14. 14.

    Escoll, P. & Buchrieser, C. Metabolic reprogramming of host cells upon bacterial infection: why shift to a Warburg-like metabolism? FEBS J. 285, 2146–2160 (2018).

  15. 15.

    O’Neill, L. A., Kishton, R. J. & Rathmell, J. A guide to immunometabolism for immunologists. Nat. Rev. Immunol. 16, 553–565 (2016).

  16. 16.

    Jha, A. K. et al. Network integration of parallel metabolic and transcriptional data reveals metabolic modules that regulate macrophage polarization. Immunity 42, 419–430 (2015).

  17. 17.

    Michelucci, A. et al. Immune-responsive gene 1 protein links metabolism to immunity by catalyzing itaconic acid production. Proc. Natl Acad. Sci. USA 110, 7820–7825 (2013). This study reports the identification of Irg1 as encoding an enzyme that generates itaconate and the finding that itaconate is a potent antimicrobial molecule that can block the growth of intracellular M. tuberculosis and Salmonella spp.

  18. 18.

    MacMicking, J. D. et al. Identification of nitric oxide synthase as a protective locus against tuberculosis. Proc. Natl Acad. Sci. USA 94, 5243–5248 (1997).

  19. 19.

    Weiss, G. & Schaible, U. E. Macrophage defense mechanisms against intracellular bacteria. Immunol. Rev. 264, 182–203 (2015).

  20. 20.

    Cyster, J. G., Dang, E. V., Reboldi, A. & Yi, T. 25-hydroxycholesterols in innate and adaptive immunity. Nat. Rev. Immunol. 14, 731–743 (2014).

  21. 21.

    Tan, Z. et al. Pyruvate dehydrogenase kinase 1 participates in macrophage polarization via regulating glucose metabolism. J. Immunol. 194, 6082–6089 (2015).

  22. 22.

    Mosser, D. M. & Edwards, J. P. Exploring the full spectrum of macrophage activation. Nat. Rev. Immunol. 8, 958–969 (2008).

  23. 23.

    Gorke, B. & Stulke, J. Carbon catabolite repression in bacteria: many ways to make the most out of nutrients. Nat. Rev. Microbiol. 6, 613–624 (2008).

  24. 24.

    Olive, A. J. & Sassetti, C. M. Metabolic crosstalk between host and pathogen: sensing, adapting and competing. Nat. Rev. Microbiol. 14, 221–234 (2016).

  25. 25.

    Burton, N. A. et al. Disparate impact of oxidative host defenses determines the fate of Salmonella during systemic infection in mice. Cell Host Microbe 15, 72–83 (2014).

  26. 26.

    Figueira, R. & Holden, D. W. Functions of the Salmonella pathogenicity island 2 (SPI-2) type III secretion system effectors. Microbiology 158, 1147–1161 (2012).

  27. 27.

    Bumann, D. & Schothorst, J. Intracellular Salmonella metabolism. Cell. Microbiol. 19, e12766 (2017).

  28. 28.

    Steeb, B. et al. Parallel exploitation of diverse host nutrients enhances Salmonella virulence. PLOS Pathog. 9, e1003301 (2013). This is an extremely comprehensive analysis of the major metabolic pathways that are required to support the survival and growth of Salmonella spp. inside mammalian cells using proteomics, genetics and computational modelling.

  29. 29.

    Isaac, D. T. & Isberg, R. Master manipulators: an update on Legionella pneumophila Icm/Dot translocated substrates and their host targets. Future Microbiol. 9, 343–359 (2014).

  30. 30.

    Eylert, E. et al. Isotopologue profiling of Legionella pneumophila: role of serine and glucose as carbon substrates. J. Biol. Chem. 285, 22232–22243 (2010).

  31. 31.

    Oliva, G., Sahr, T. & Buchrieser, C. The life cycle of L. pneumophila: cellular differentiation is linked to virulence and metabolism. Front. Cell. Infect. Microbiol. 8, 3 (2018).

  32. 32.

    Hauslein, I. et al. Legionella pneumophila CsrA regulates a metabolic switch from amino acid to glycerolipid metabolism. Open Biol. 7, 170149 (2017).

  33. 33.

    Lang, C. & Flieger, A. Characterisation of Legionella pneumophila phospholipases and their impact on host cells. Eur. J. Cell. Biol. 90, 903–912 (2011).

  34. 34.

    Lerner, T. R. et al. Mycobacterium tuberculosis replicates within necrotic human macrophages. J. Cell Biol. 216, 583–594 (2017).

  35. 35.

    Mahamed, D. et al. Intracellular growth of Mycobacterium tuberculosis after macrophage cell death leads to serial killing of host cells. eLife 6, e28205 (2017).

  36. 36.

    van der Wel, N. et al. M. tuberculosis and M. leprae translocate from the phagolysosome to the cytosol in myeloid cells. Cell 129, 1287–1298 (2007).

  37. 37.

    Stanley, S. A., Johndrow, J. E., Manzanillo, P. & Cox, J. S. The type I IFN response to infection with Mycobacterium tuberculosis requires ESX-1-mediated secretion and contributes to pathogenesis. J. Immunol. 178, 3143–3152 (2007).

  38. 38.

    McKinney, J. D. et al. Persistence of Mycobacterium tuberculosis in macrophages and mice requires the glyoxylate shunt enzyme isocitrate lyase. Nature 406, 735–738 (2000).

  39. 39.

    Pandey, A. K. & Sassetti, C. M. Mycobacterial persistence requires the utilization of host cholesterol. Proc. Natl Acad. Sci. USA 105, 4376–4380 (2008). This study identifies the Mce4 membrane protein complex as the primary transporter of cholesterol in M. tuberculosis and shows that the activity of this transporter is necessary for maintenance of M. tuberculosis infection in vivo.

  40. 40.

    VanderVen, B. C. et al. Novel inhibitors of cholesterol degradation in Mycobacterium tuberculosis reveal how the bacterium’s metabolism is constrained by the intracellular environment. PLOS Pathog. 11, e1004679 (2015). This paper presents the first identification of chemical inhibitors of M. tuberculosis enzymes that are involved in the degradation of host-derived cholesterol. It shows that chemical inhibition of this pathway in the bacteria limits their intracellular growth.

  41. 41.

    Lee, W., VanderVen, B. C., Fahey, R. J. & Russell, D. G. Intracellular Mycobacterium tuberculosis exploits host-derived fatty acids to limit metabolic stress. J. Biol. Chem. 288, 6788–6800 (2013).

  42. 42.

    Nazarova, E. V. et al. Rv3723/LucA coordinates fatty acid and cholesterol uptake in Mycobacterium tuberculosis. eLife 6, e26969 (2017).

  43. 43.

    Marrero, J., Trujillo, C., Rhee, K. Y. & Ehrt, S. Glucose phosphorylation is required for Mycobacterium tuberculosis persistence in mice. PLOS Pathog. 9, e1003116 (2013).

  44. 44.

    Beste, D. J. et al. 13C-flux spectral analysis of host-pathogen metabolism reveals a mixed diet for intracellular Mycobacterium tuberculosis. Chem. Biol. 20, 1012–1021 (2013).

  45. 45.

    O’Riordan, M., Moors, M. A. & Portnoy, D. A. Listeria intracellular growth and virulence require host-derived lipoic acid. Science 302, 462–464 (2003).

  46. 46.

    Grubmuller, S., Schauer, K., Goebel, W., Fuchs, T. M. & Eisenreich, W. Analysis of carbon substrates used by Listeria monocytogenes during growth in J774A.1 macrophages suggests a bipartite intracellular metabolism. Front. Cell. Infect. Microbiol. 4, 156 (2014).

  47. 47.

    Kochan, I. The role of iron in bacterial infections, with special consideration of host-tubercle bacillus interaction. Curr. Top. Microbiol. Immunol. 60, 1–30 (1973).

  48. 48.

    Botella, H. et al. Mycobacterial p1-type ATPases mediate resistance to zinc poisoning in human macrophages. Cell Host Microbe 10, 248–259 (2011).

  49. 49.

    Crouch, M. L., Castor, M., Karlinsey, J. E., Kalhorn, T. & Fang, F. C. Biosynthesis and IroC-dependent export of the siderophore salmochelin are essential for virulence of Salmonella enterica serovar Typhimurium. Mol. Microbiol. 67, 971–983 (2008).

  50. 50.

    Liu, J. Z. et al. Zinc sequestration by the neutrophil protein calprotectin enhances Salmonella growth in the inflamed gut. Cell Host Microbe 11, 227–239 (2012).

  51. 51.

    Wolschendorf, F. et al. Copper resistance is essential for virulence of Mycobacterium tuberculosis. Proc. Natl Acad. Sci. USA 108, 1621–1626 (2011).

  52. 52.

    Bange, F. C., Brown, A. M. & Jacobs, W. R. Jr. Leucine auxotrophy restricts growth of Mycobacterium bovis BCG in macrophages. Infect. Immun. 64, 1794–1799 (1996).

  53. 53.

    Clark-Curtiss, J. E. & Curtiss, R. 3rd. Salmonella vaccines: conduits for protective antigens. J. Immunol. 200, 39–48 (2018).

  54. 54.

    Ensminger, A. W., Yassin, Y., Miron, A. & Isberg, R. R. Experimental evolution of Legionella pneumophila in mouse macrophages leads to strains with altered determinants of environmental survival. PLOS Pathog. 8, e1002731 (2012).

  55. 55.

    Vilcheze, C. et al. Rational design of biosafety level 2-approved, multidrug-resistant strains of Mycobacterium tuberculosis through nutrient auxotrophy. mBio 9, e00938 (2018).

  56. 56.

    Murray, P. J. Amino acid auxotrophy as a system of immunological control nodes. Nat. Immunol. 17, 132–139 (2016).

  57. 57.

    Pfefferkorn, E. R. Interferon gamma blocks the growth of Toxoplasma gondii in human fibroblasts by inducing the host cells to degrade tryptophan. Proc. Natl Acad. Sci. USA 81, 908–912 (1984).

  58. 58.

    Schmidt, S. V. & Schultze, J. L. New insights into IDO biology in bacterial and viral infections. Front. Immunol. 5, 384 (2014).

  59. 59.

    O’Neill, L. A. J. & Artyomov, M. N. Itaconate: the poster child of metabolic reprogramming in macrophage function. Nat. Rev. Immunol. (in the press).

  60. 60.

    Lampropoulou, V. et al. Itaconate links inhibition of succinate dehydrogenase with macrophage metabolic remodeling and regulation of inflammation. Cell Metab. 24, 158–166 (2016).

  61. 61.

    Mills, E. L. et al. Itaconate is an anti-inflammatory metabolite that activates Nrf2 via alkylation of KEAP1. Nature 556, 113–117 (2018).

  62. 62.

    Weiss, J. M. et al. Itaconic acid mediates crosstalk between macrophage metabolism and peritoneal tumors. J. Clin. Invest. 128, 3794–3805 (2018).

  63. 63.

    Honer Zu Bentrup, K., Miczak, A., Swenson, D. L. & Russell, D. G. Characterization of activity and expression of isocitrate lyase in Mycobacterium avium and Mycobacterium tuberculosis. J. Bacteriol. 181, 7161–7167 (1999).

  64. 64.

    Munoz-Elias, E. J. & McKinney, J. D. Mycobacterium tuberculosis isocitrate lyases 1 and 2 are jointly required for in vivo growth and virulence. Nat. Med. 11, 638–644 (2005).

  65. 65.

    Savvi, S. et al. Functional characterization of a vitamin B12-dependent methylmalonyl pathway in Mycobacterium tuberculosis: implications for propionate metabolism during growth on fatty acids. J. Bacteriol. 190, 3886–3895 (2008).

  66. 66.

    Naujoks, J. et al. IFNs modify the proteome of legionella-containing vacuoles and restrict infection via IRG1-derived itaconic acid. PLOS Pathog. 12, e1005408 (2016).

  67. 67.

    Nair, S. et al. Irg1 expression in myeloid cells prevents immunopathology during M. tuberculosis infection. J. Exp. Med. 215, 1035–1045 (2018).

  68. 68.

    Sasikaran, J., Ziemski, M., Zadora, P. K., Fleig, A. & Berg, I. A. Bacterial itaconate degradation promotes pathogenicity. Nat. Chem. Biol. 10, 371–377 (2014).

  69. 69.

    Ito, K. & Suda, T. Metabolic requirements for the maintenance of self-renewing stem cells. Nat. Rev. Mol. Cell. Biol. 15, 243–256 (2014).

  70. 70.

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

  71. 71.

    Everts, B. et al. TLR-driven early glycolytic reprogramming via the kinases TBK1-IKKε supports the anabolic demands of dendritic cell activation. Nat. Immunol. 15, 323–332 (2014).

  72. 72.

    Rodriguez-Prados, J. C. et al. Substrate fate in activated macrophages: a comparison between innate, classic, and alternative activation. J. Immunol. 185, 605–614 (2010).

  73. 73.

    Tan, Z. et al. The monocarboxylate transporter 4 is required for glycolytic reprogramming and inflammatory response in macrophages. J. Bio. Chem. 290, 46–55 (2015).

  74. 74.

    Lachmandas, E. et al. Microbial stimulation of different Toll-like receptor signalling pathways induces diverse metabolic programmes in human monocytes. Nat. Microbiol. 2, 16246 (2016).

  75. 75.

    Gillmaier, N., Gotz, A., Schulz, A., Eisenreich, W. & Goebel, W. Metabolic responses of primary and transformed cells to intracellular Listeria monocytogenes. PLOS ONE 7, e52378 (2012).

  76. 76.

    Czyz, D. M., Willett, J. W. & Crosson, S. Brucella abortus induces a Warburg shift in host metabolism that is linked to enhanced intracellular survival of the pathogen. J. Bacteriol. 199, e00227 (2017).

  77. 77.

    Cheng, S. C. et al. mTOR- and HIF-1α-mediated aerobic glycolysis as metabolic basis for trained immunity. Science 345, 1250684 (2014).

  78. 78.

    Netea, M. G. et al. Trained immunity: a program of innate immune memory in health and disease. Science 352, aaf1098 (2016).

  79. 79.

    Escoll, P. et al. Legionella pneumophila modulates mitochondrial dynamics to trigger metabolic repurposing of infected macrophages. Cell Host Microbe 22, 302–316 (2017). This study shows that L. pneumophila modulates the metabolism of its host cell through driving fission of the mitochondria in a DNM1L-dependent manner.

  80. 80.

    Stavru, F., Bouillaud, F., Sartori, A., Ricquier, D. & Cossart, P. Listeria monocytogenes transiently alters mitochondrial dynamics during infection. Proc. Natl Acad. Sci. USA 108, 3612–3617 (2011).

  81. 81.

    Abramovitch, R. B. Mycobacterium tuberculosis reporter strains as tools for drug discovery and development. IUBMB Life 70, 818–825 (2018).

  82. 82.

    Boot, M. et al. Accelerating early antituberculosis drug discovery by creating mycobacterial indicator strains that predict mode of action. Antimicrob. Agents Chemother. 62, e00083 (2018).

  83. 83.

    Claudi, B. et al. Phenotypic variation of Salmonella in host tissues delays eradication by antimicrobial chemotherapy. Cell 158, 722–733 (2014). This study uses fluorescent TIMER-expressing Salmonella spp. in an in vivo infection model to analyse the rate of bacterial division, identify a subpopulation of non-replicating bacteria and demonstrate that these bacteria have acquired a drug-tolerant phenotype.

  84. 84.

    Gill, W. P. et al. A replication clock for Mycobacterium tuberculosis. Nat. Med. 15, 211–214 (2009).

  85. 85.

    Helaine, S. et al. Internalization of Salmonella by macrophages induces formation of nonreplicating persisters. Science 343, 204–208 (2014). In this study, a green fluorescent protein-dilution reporter strain is used to examine non-replicating Salmonella spp. bacteria in vivo and to show that, in vitro, IFNγ-activated macrophages induce markedly higher levels of drug tolerance in Salmonella spp. than do resting macrophages.

  86. 86.

    Helaine, S. & Holden, D. W. Heterogeneity of intracellular replication of bacterial pathogens. Curr. Opin. Microbiol. 16, 184–191 (2013).

  87. 87.

    MacGilvary, N. J. & Tan, S. Fluorescent Mycobacterium tuberculosis reporters: illuminating host — pathogen interactions. Pathog. Dis. 76, fty017 (2018).

  88. 88.

    Sukumar, N., Tan, S., Aldridge, B. B. & Russell, D. G. Exploitation of Mycobacterium tuberculosis reporter strains to probe the impact of vaccination at sites of infection. PLOS Pathog. 10, e1004394 (2014).

  89. 89.

    Saliba, A. E. et al. Single-cell RNA-seq ties macrophage polarization to growth rate of intracellular Salmonella. Nat. Microbiol. 2, 16206 (2016). This study uses single-cell RNA-seq to show that macrophages that preferentially support the growth of Salmonella spp. in vitro express polarization markers consistent with an M2-like macrophage phenotype.

  90. 90.

    Westermann, A. J., Barquist, L. & Vogel, J. Resolving host-pathogen interactions by dual RNA-seq. PLOS Pathog. 13, e1006033 (2017).

  91. 91.

    Westermann, A. J. et al. Dual RNA-seq unveils noncoding RNA functions in host-pathogen interactions. Nature 529, 496–501 (2016).

  92. 92.

    Bumann, D. & Cunrath, O. Heterogeneity of Salmonella-host interactions in infected host tissues. Curr. Opin. Microbiol. 39, 57–63 (2017).

  93. 93.

    Cadena, A. M., Fortune, S. M. & Flynn, J. L. Heterogeneity in tuberculosis. Nat. Rev. Immunol. 17, 691–702 (2017).

  94. 94.

    Helaine, S. & Kugelberg, E. Bacterial persisters: formation, eradication, and experimental systems. Trends Microbiol. 22, 417–424 (2014).

  95. 95.

    Liu, Y. et al. Immune activation of the host cell induces drug tolerance in Mycobacterium tuberculosis both in vitro and in vivo. J. Exp. Med. 213, 809–825 (2016).

  96. 96.

    Manina, G., Dhar, N. & McKinney, J. D. Stress and host immunity amplify Mycobacterium tuberculosis phenotypic heterogeneity and induce nongrowing metabolically active forms. Cell Host Microbe 17, 32–46 (2015).

  97. 97.

    Stapels, D. A. C. et al. Salmonella persisters undermine host immune defenses during antibiotic treatment. Science 362, 1156–1160 (2018).

  98. 98.

    Perez-Morales, D. & Bustamante, V. H. The global regulatory system Csr senses glucose through the phosphoenolpyruvate: carbohydrate phosphotransferase system. Mol. Microbiol. 99, 623–626 (2016).

  99. 99.

    Aldridge, B. B. et al. Asymmetry and aging of mycobacterial cells lead to variable growth and antibiotic susceptibility. Science 335, 100–104 (2012).

  100. 100.

    Rego, E. H., Audette, R. E. & Rubin, E. J. Deletion of a mycobacterial divisome factor collapses single-cell phenotypic heterogeneity. Nature 546, 153–157 (2017).

  101. 101.

    Cohen, S. B. et al. Alveolar macrophages provide an early Mycobacterium tuberculosis niche and initiate dissemination. Cell Host Microbe 24, 439–446 (2018).

  102. 102.

    Srivastava, S., Ernst, J. D. & Desvignes, L. Beyond macrophages: the diversity of mononuclear cells in tuberculosis. Immunol. Rev. 262, 179–192 (2014).

  103. 103.

    Srivastava, S., Grace, P. S. & Ernst, J. D. Antigen export reduces antigen presentation and limits T cell control of M. tuberculosis. Cell Host Microbe 19, 44–54 (2016).

  104. 104.

    Bodnar, K. A., Serbina, N. V. & Flynn, J. L. Fate of Mycobacterium tuberculosis within murine dendritic cells. Infect. Immun. 69, 800–809 (2001).

  105. 105.

    Mattila, J. T. et al. Microenvironments in tuberculous granulomas are delineated by distinct populations of macrophage subsets and expression of nitric oxide synthase and arginase isoforms. J. Immunol. 191, 773–784 (2013).

  106. 106.

    Marino, S. et al. Macrophage polarization drives granuloma outcome during Mycobacterium tuberculosis infection. Infect. Immun. 83, 324–338 (2015).

  107. 107.

    Huang, L., Nazarova, E. V., Tan, S., Liu, Y. & Russell, D. G. Growth of Mycobacterium tuberculosis in vivo segregates with host macrophage metabolism and ontogeny. J. Exp. Med. 215, 1135–1152 (2018). This is a recent study showing that macrophages of different developmental lineages differentially support the growth of M. tuberculosis in vivo and that alveolar macrophages are more permissive for bacterial growth than are interstitial macrophages through metabolism-dependent mechanisms.

  108. 108.

    Rohde, K. H., Veiga, D. F., Caldwell, S., Balazsi, G. & Russell, D. G. Linking the transcriptional profiles and the physiological states of Mycobacterium tuberculosis during an extended intracellular infection. PLOS Pathog. 8, e1002769 (2012).

  109. 109.

    Tan, S., Sukumar, N., Abramovitch, R. B., Parish, T. & Russell, D. G. Mycobacterium tuberculosis responds to chloride and pH as synergistic cues to the immune status of its host cell. PLOS Pathog. 9, e1003282 (2013).

  110. 110.

    Dunlap, M. D. et al. A novel role for C-C motif chemokine receptor 2 during infection with hypervirulent Mycobacterium tuberculosis. Mucosal Immunol. 11, 1727–1742 (2018).

  111. 111.

    Gleeson, L. E. et al. Cigarette smoking impairs the bioenergetic immune response to mycobacterium tuberculosis infection. Am. J. Respir. Cell. Mol. Biol. 59, 572–579 (2018).

  112. 112.

    Goletti, D., Petruccioli, E., Joosten, S. A. & Ottenhoff, T. H. Tuberculosis biomarkers: from diagnosis to protection. Infect. Dis. Rep. 8, 6568 (2016).

  113. 113.

    Lee, S. H. et al. Mannose receptor high, M2 dermal macrophages mediate nonhealing Leishmania major infection in a Th1 immune environment. J. Exp. Med. 215, 357–375 (2018).

  114. 114.

    Gibbings, S. L. et al. Transcriptome analysis highlights the conserved difference between embryonic and postnatal-derived alveolar macrophages. Blood 126, 1357–1366 (2015).

  115. 115.

    Gibbings, S. L. et al. Three unique interstitial macrophages in the murine lung at steady state. Am. J. Respir. Cell. Mol. Biol. 57, 66–76 (2017).

  116. 116.

    Mould, K. J. et al. Cell origin dictates programming of resident versus recruited macrophages during acute lung injury. Am. J. Respir. Cell. Mol. Biol. 57, 294–306 (2017). This study shows that resident and recruited macrophages respond differently to LPS challenge in the lung airways, which indicates that the developmental origin of macrophages, rather than their environment, is decisive in determining their response to the same immune stimulus.

  117. 117.

    Jenkins, S. J. et al. Local macrophage proliferation, rather than recruitment from the blood, is a signature of TH2 inflammation. Science 332, 1284–1288 (2011).

  118. 118.

    Jenkins, S. J. et al. IL-4 directly signals tissue-resident macrophages to proliferate beyond homeostatic levels controlled by CSF-1. J. Exp. Med. 210, 2477–2491 (2013).

  119. 119.

    Ruckerl, D. et al. Macrophage origin limits functional plasticity in helminth-bacterial co-infection. PLOS Pathog. 13, e1006233 (2017).

  120. 120.

    Devchand, P. R. et al. The PPARα-leukotriene B4 pathway to inflammation control. Nature 384, 39–43 (1996).

  121. 121.

    Nobs, S. P. & Kopf, M. PPAR-γ in innate and adaptive lung immunity. J. Leukoc. Biol. 104, 737–741 (2018).

  122. 122.

    Bedi, B. et al. Enhanced clearance of Pseudomonas aeruginosa by peroxisome proliferator-activated receptor gamma. Infect. Immun. 84, 1975–1985 (2016).

  123. 123.

    Almeida, P. E., Carneiro, A. B., Silva, A. R. & Bozza, P. T. PPARγ expression and function in mycobacterial infection: roles in lipid metabolism, immunity, and bacterial killing. PPAR Res. 2012, 383829 (2012).

  124. 124.

    Guirado, E. et al. Deletion of PPARγ in lung macrophages provides an immunoprotective response against M. tuberculosis infection in mice. Tuberculosis (Edinb.) 111, 170–177 (2018).

  125. 125.

    Kim, Y. S. et al. PPAR-α activation mediates innate host defense through induction of TFEB and lipid catabolism. J. Immunol. 198, 3283–3295 (2017).

  126. 126.

    Salamon, H. et al. Cutting edge: vitamin D regulates lipid metabolism in Mycobacterium tuberculosis infection. J. Immunol. 193, 30–34 (2014).

  127. 127.

    Arts, R. J. W. et al. Immunometabolic pathways in BCG-induced trained immunity. Cell Rep. 17, 2562–2571 (2016).

  128. 128.

    Bekkering, S. et al. Metabolic induction of trained immunity through the mevalonate pathway. Cell 172, 135–146 (2018).

  129. 129.

    Mitroulis, I. et al. Modulation of myelopoiesis progenitors is an integral component of trained immunity. Cell 172, 147–161 (2018).

  130. 130.

    Kaufmann, E. et al. BCG educates hematopoietic stem cells to generate protective innate immunity against tuberculosis. Cell 172, 176–190 (2018).

  131. 131.

    Roostalu, J., Joers, A., Luidalepp, H., Kaldalu, N. & Tenson, T. Cell division in Escherichia coli cultures monitored at single cell resolution. BMC Microbiol. 8, 68 (2008).

  132. 132.

    Terskikh, A. et al. “Fluorescent timer”: protein that changes color with time. Science 290, 1585–1588 (2000).

  133. 133.

    Reyes-Lamothe, R., Sherratt, D. J. & Leake, M. C. Stoichiometry and architecture of active DNA replication machinery in Escherichia coli. Science 328, 498–501 (2010).

  134. 134.

    Abramovitch, R. B., Rohde, K. H., Hsu, F. F. & Russell, D. G. aprABC: a Mycobacterium tuberculosis complex-specific locus that modulates pH-driven adaptation to the macrophage phagosome. Mol. Microbiol. 80, 678–694 (2011).

  135. 135.

    Miesenbock, G., De Angelis, D. A. & Rothman, J. E. Visualizing secretion and synaptic transmission with pH-sensitive green fluorescent proteins. Nature 394, 192–195 (1998).

  136. 136.

    Vandal, O. H., Pierini, L. M., Schnappinger, D., Nathan, C. F. & Ehrt, S. A membrane protein preserves intrabacterial pH in intraphagosomal Mycobacterium tuberculosis. Nat. Med. 14, 849–854 (2008).

  137. 137.

    Gierahn, T. M. et al. Seq-Well: portable, low-cost RNA sequencing of single cells at high throughput. Nat. Methods 14, 395–398 (2017).

  138. 138.

    Shah-Simpson, S., Pereira, C. F., Dumoulin, P. C., Caradonna, K. L. & Burleigh, B. A. Bioenergetic profiling of Trypanosoma cruzi life stages using Seahorse extracellular flux technology. Mol. Biochem. Parasitol. 208, 91–95 (2016).

  139. 139.

    Kloehn, J. et al. Using metabolomics to dissect host-parasite interactions. Curr. Opin. Microbiol. 32, 59–65 (2016).

  140. 140.

    Saunders, E. C., Naderer, T., Chambers, J., Landfear, S. M. & McConville, M. J. Leishmania mexicana can utilize amino acids as major carbon sources in macrophages but not in animal models. Mol. Microbiol. 108, 143–158 (2018).

  141. 141.

    Goldman-Pinkovich, A. et al. An arginine deprivation response pathway is induced in leishmania during macrophage invasion. PLOS Pathog. 12, e1005494 (2016).

  142. 142.

    Li, Y. et al. Transcriptome remodeling in Trypanosoma cruzi and human cells during intracellular infection. PLOS Pathog. 12, e1005511 (2016).

  143. 143.

    Shah-Simpson, S., Lentini, G., Dumoulin, P. C. & Burleigh, B. A. Modulation of host central carbon metabolism and in situ glucose uptake by intracellular Trypanosoma cruzi amastigotes. PLOS Pathog. 13, e1006747 (2017).

  144. 144.

    Blume, M. et al. A Toxoplasma gondii gluconeogenic enzyme contributes to robust central carbon metabolism and is essential for replication and virulence. Cell Host Microbe 18, 210–220 (2015).

  145. 145.

    Jacot, D., Waller, R. F., Soldati-Favre, D., MacPherson, D. A. & MacRae, J. I. Apicomplexan energy metabolism: carbon source promiscuity and the quiescence hyperbole. Trends Parasitol. 32, 56–70 (2016).

  146. 146.

    Jensen, K. D. et al. Toxoplasma polymorphic effectors determine macrophage polarization and intestinal inflammation. Cell Host Microbe 9, 472–483 (2011).

  147. 147.

    Leroux, L. P. et al. The protozoan parasite Toxoplasma gondii selectively reprograms the host cell translatome. Infect. Immun. 86, e00244 (2018).

  148. 148.

    Barelle, C. J. et al. Niche-specific regulation of central metabolic pathways in a fungal pathogen. Cell. Microbiol. 8, 961–971 (2006).

  149. 149.

    Lorenz, M. C. & Fink, G. R. The glyoxylate cycle is required for fungal virulence. Nature 412, 83–86 (2001).

  150. 150.

    Osborne, S. E. et al. Type I interferon promotes cell-to-cell spread of Listeria monocytogenes. Cell. Microbiol. 19, e12660 (2017).

  151. 151.

    Harouz, H. et al. Shigella flexneri targets the HP1gamma subcode through the phosphothreonine lyase OspF. EMBO J. 33, 2606–2622 (2014).

  152. 152.

    Kentner, D. et al. Shigella reroutes host cell central metabolism to obtain high-flux nutrient supply for vigorous intracellular growth. Proc. Natl Acad. Sci. USA 111, 9929–9934 (2014).

  153. 153.

    Waligora, E. A. et al. Role of intracellular carbon metabolism pathways in Shigella flexneri virulence. Infect. Immun. 82, 2746–2755 (2014).

  154. 154.

    Calverley, M., Erickson, S., Read, A. J. & Harmsen, A. G. Resident alveolar macrophages are susceptible to and permissive of Coxiella burnetii infection. PLOS ONE 7, e51941 (2012).

  155. 155.

    Graham, J. G. et al. Virulent Coxiella burnetii pathotypes productively infect primary human alveolar macrophages. Cell. Microbiol. 15, 1012–1025 (2013).

  156. 156.

    Hauslein, I. et al. Multiple substrate usage of Coxiella burnetii to feed a bipartite metabolic network. Front. Cell. Infect. Microbiol. 7, 285 (2017).

  157. 157.

    Mulye, M., Zapata, B. & Gilk, S. D. Altering lipid droplet homeostasis affects Coxiella burnetii intracellular growth. PLOS ONE 13, e0192215 (2018).

  158. 158.

    Hauslein, I., Manske, C., Goebel, W., Eisenreich, W. & Hilbi, H. Pathway analysis using 13C-glycerol and other carbon tracers reveals a bipartite metabolism of Legionella pneumophila. Mol. Microbiol. 100, 229–246 (2016).

Download references


D.G.R., L.H. and B.C.V. acknowledge the support of the National Institutes of Health, USA, and the Bill and Melinda Gates Foundation.

Reviewer information

Nature Reviews Immunology thanks K. Fitzgerald, C. Sassetti and other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information


  1. Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA

    • David G. Russell
    • , Lu Huang
    •  & Brian C. VanderVen


  1. Search for David G. Russell in:

  2. Search for Lu Huang in:

  3. Search for Brian C. VanderVen in:


D.G.R. researched data for the article. All authors contributed to discussing content and writing and editing the manuscript.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to David G. Russell.


Foamy macrophages

Macrophages with cytosolic lipid droplets containing cholesterol, cholesterol ester and triacylglycerol, which are frequently induced by chronic pro-inflammatory stimuli.

M1 and M2 macrophages

M1 and M2 are classifications historically used to define macrophages activated in vitro as pro-inflammatory (when classically activated with IFNγ and lipopolysaccharide) or as anti-inflammatory (when alternatively activated with IL-4 or IL-10), respectively. However, in vivo, macrophages are highly specialized, transcriptomically dynamic and extremely heterogeneous with regards to their phenotypes and functions, which are continuously shaped by their tissue microenvironment. Therefore, the M1 or M2 classification is too simplistic to explain the true nature of in vivo macrophages, although these terms are still often used to indicate whether the macrophages in question are more pro-inflammatory or anti-inflammatory.

Dual RNA sequencing

(Dual RNA-seq). A transcriptional profiling technique that enables simultaneous acquisition of the expression levels of mRNA transcripts in both the host cell and the pathogen.

Tricarboxylic acid cycle

(TCA cycle). Also known as the citric acid cycle or Krebs cycle. This is a series of enzymatic reactions used in aerobic metabolism to release energy through the oxidation of acetyl-CoA to yield ATP and carbon dioxide.

Succinate dehydrogenase complex

(SDH complex). An enzyme complex found in bacterial cells and in the inner mitochondrial membrane of eukaryotic mitochondria that is active in both the tricarboxylic acid cycle and the electron transport chain.

Bipartite metabolism

A metabolic programme whereby one carbon source is used exclusively as an energy supply while another carbon source, or sources, is used for anabolic processes.


Bacteria that are unable to synthesize all of the compounds required for growth are auxotrophic, meaning that they are dependent on their hosts to supply those compounds they cannot synthesize.

Oxygen consumption rate

(OCR). The total oxygen utilization capacity of a biological system under examination against time.

Spare respiratory capacity

(SRC). The difference in the amount of ATP generated by oxidative phosphorylation at basal rate and at maximal respiratory capacity.

Aerobic glycolysis

The conversion of glucose to lactate under conditions where oxygen is present at non-limiting concentrations.

Type IV secretion system

An ATP-dependent bacterial transporter complex that is frequently used to inject bacterial effector proteins or bacterial DNA into eukaryotic and prokaryotic target cells.

Mitochondrial fusion and fission

The fusion or fragmentation of mitochondria in a highly controlled manner, which can regulate the oxidative phosphorylation capacity of eukaryotic cells.

Fitness reporter organisms

Bacterial reporter strains, usually encoding a fluorescent protein-based readout, that are used to assess bacterial fitness with respect to responsiveness to noxious stimuli and replicative capacity.

Ribosomal RNA correlates

Sequences encoding a destabilized or short-lived green fluorescent protein are inserted into ribosomal RNA loci to provide a correlate of ribosomal RNA activity and bacterial replication.

pH-sensitive green fluorescent protein

A green fluorescent protein derivative that exhibits a shift in fluorescence emission wavelength in a pH-dependent manner.

Fluorescence dilution reporter strain

A bacterial strain that can be induced to transiently express a fluorescent protein, which can then be quantified as it becomes diluted when the bacteria divide to infer bacterial replication rates.


A re-engineered red fluorescent protein that undergoes a conformational shift, and hence a change in fluorescence emission wavelength, as it ages, thus providing a correlate of replication rates.

‘Clock’ plasmid

An episomal plasmid encoding an antibiotic-resistance marker that is lost from a bacterial population at a fixed rate directly proportional to the rate of replication.

Chromosomal replication complex reporter

A single-stranded DNA-binding protein–green fluorescent protein fusion complex that persists for the duration of chromosomal replication and can be used to assess the replication status of a bacterial population.

Environmentally responsive promoter reporter

A dual-fluorescent bacterial reporter strain that expresses one fluorescent protein constitutively and the other fluorescent protein under the control of promoters that are responsive to specific environmental stimuli.

About this article

Publication history