Article | Published:

Resistance of HIV-infected macrophages to CD8+ T lymphocyte–mediated killing drives activation of the immune system

Nature Immunologyvolume 19pages475486 (2018) | Download Citation



CD4+ T lymphocytes are the principal target of human immunodeficiency virus (HIV), but infected macrophages also contribute to viral pathogenesis. The killing of infected cells by CD8+ cytotoxic T lymphocytes (CTLs) leads to control of viral replication. Here we found that the killing of macrophages by CTLs was impaired relative to the killing of CD4+ T cells by CTLs, and this resulted in inefficient suppression of HIV. The killing of macrophages depended on caspase-3 and granzyme B, whereas the rapid killing of CD4+ T cells was caspase independent and did not require granzyme B. Moreover, the impaired killing of macrophages was associated with prolonged effector cell–target cell contact time and higher expression of interferon-γ by CTLs, which induced macrophage production of pro-inflammatory chemokines that recruited monocytes and T cells. Similar results were obtained when macrophages presented other viral antigens, suggestive of a general mechanism for macrophage persistence as antigen-presenting cells that enhance inflammation and adaptive immunity. Inefficient killing of macrophages by CTLs might contribute to chronic inflammation, a hallmark of chronic disease caused by HIV.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.


  1. 1.

    Swingler, S., Mann, A. M., Zhou, J., Swingler, C. & Stevenson, M. Apoptotic killing of HIV-1-infected macrophages is subverted by the viral envelope glycoprotein. PLoS Pathog. 3, 1281–1290 (2007).

  2. 2.

    Groot, F., Welsch, S. & Sattentau, Q. J. Efficient HIV-1 transmission from macrophages to T cells across transient virological synapses. Blood 111, 4660–4663 (2008).

  3. 3.

    Duncan, C. J. et al. High-multiplicity HIV-1 infection and neutralizing antibody evasion mediated by the macrophage-T cell virological synapse. J. Virol. 88, 2025–2034 (2014).

  4. 4.

    Collins, D. R., Lubow, J., Lukic, Z., Mashiba, M. & Collins, K. L. Vpr promotes macrophage-dependent HIV-1 infection of CD4+ T lymphocytes. PLoS Pathog. 11, e1005054 (2015).

  5. 5.

    Honeycutt, J. B. et al. Macrophages sustain HIV replication in vivo independently of T cells. J. Clin. Invest. 126, 1353–1366 (2016).

  6. 6.

    Honeycutt, J. B. et al. HIV persistence in tissue macrophages of humanized myeloid-only mice during antiretroviral therapy. Nat. Med. 23, 638–643 (2017).

  7. 7.

    Avalos, C. R. et al. Quantitation of productively infected monocytes and macrophages of simian immunodeficiency virus-infected macaques. J. Virol. 90, 5643–5656 (2016).

  8. 8.

    Avalos, C. R. et al. Brain Macrophages in simian immunodeficiency virus-infected, antiretroviral-suppressed macaques: a functional latent reservoir. mBio 8, e01186–e01117 (2017).

  9. 9.

    Nowlin, B. T. et al. SIV encephalitis lesions are composed of CD163+ macrophages present in the central nervous system during early SIV infection and SIV-positive macrophages recruited terminally with AIDS. Am. J. Pathol. 185, 1649–1665 (2015).

  10. 10.

    Sattentau, Q. J. & Stevenson, M. Macrophages and HIV-1: an unhealthy constellation. Cell Host Microbe 19, 304–310 (2016).

  11. 11.

    DiNapoli, S. R., Hirsch, V. M. & Brenchley, J. M. Macrophages in progressive human immunodeficiency virus/simian immunodeficiency virus infections. J. Virol. 90, 7596–7606 (2016).

  12. 12.

    Lamers, S. L. et al. HIV-1 Nef in macrophage-mediated disease pathogenesis. Int. Rev. Immunol. 31, 432–450 (2012).

  13. 13.

    Hsu, D. C., Sereti, I. & Ananworanich, J. Serious non-AIDS events: immunopathogenesis and interventional strategies. AIDS Res. Ther. 10, 29 (2013).

  14. 14.

    Streeck, H. & Nixon, D. F. T cell immunity in acute HIV-1 infection. J. Infect. Dis. 202(Suppl 2), S302–S308 (2010).

  15. 15.

    Goulder, P. J. & Walker, B. D. The great escape — AIDS viruses and immune control. Nat. Med. 5, 1233–1235 (1999).

  16. 16.

    Fujiwara, M. & Takiguchi, M. HIV-1-specific CTLs effectively suppress replication of HIV-1 in HIV-1-infected macrophages. Blood 109, 4832–4838 (2007).

  17. 17.

    Severino, M. E. et al. Inhibition of human immunodeficiency virus type 1 replication in primary CD4+ T lymphocytes, monocytes, and dendritic cells by cytotoxic T lymphocytes. J. Virol. 74, 6695–6699 (2000).

  18. 18.

    Walker-Sperling, V. E., Buckheit, R. W. III & Blankson, J. N. Comparative analysis of the capacity of elite suppressor CD4+ and CD8+ T cells to inhibit HIV-1 replication in monocyte-derived macrophages. J. Virol. 88, 9789–9798 (2014).

  19. 19.

    Walker-Sperling, V. E. et al. Short communication: HIV controller T cells effectively inhibit viral replication in alveolar macrophages. AIDS Res. Hum. Retroviruses 32, 1097–1099 (2016).

  20. 20.

    Rainho, J. N. et al. Nef Is dispensable for resistance of simian immunodeficiency virus-infected macrophages to CD8+ T cell killing. J. Virol. 89, 10625–10636 (2015).

  21. 21.

    Vojnov, L. et al. The majority of freshly sorted simian immunodeficiency virus (SIV)-specific CD8. T cells cannot suppress viral replication in SIV-infected macrophages. J. Virol. 86, 4682–4687 (2012).

  22. 22.

    Walker, B. D. & Yu, X. G. Unravelling the mechanisms of durable control of HIV-1. Nat. Rev. Immunol. 13, 487–498 (2013).

  23. 23.

    Slee, E. A., Adrain, C. & Martin, S. J. Executioner caspase-3, -6, and -7 perform distinct, non-redundant roles during the demolition phase of apoptosis. J. Biol. Chem. 276, 7320–7326 (2001).

  24. 24.

    Lieberman, J. The ABCs of granule-mediated cytotoxicity: new weapons in the arsenal. Nat. Rev. Immunol. 3, 361–370 (2003).

  25. 25.

    Belizario, J., Vieira-Cordeiro, L. & Enns, S. Necroptotic cell death signaling and execution pathway: lessons from knockout mice. Mediators Inflamm. 2015, 128076 (2015).

  26. 26.

    Lieberman, J. Cell-mediated cytotoxicity. in Fundamental Immunology (ed. Paul, W.E.) 7th edn (Lippincott Williams & Wilkins, Philadelphia, 2013).

  27. 27.

    Kaiserman, D. & Bird, P. I. Control of granzymes by serpins. Cell Death Differ. 17, 586–595 (2010).

  28. 28.

    Kataoka, T. et al. Concanamycin A, a powerful tool for characterization and estimation of contribution of perforin- and Fas-based lytic pathways in cell-mediated cytotoxicity. J. Immunol. 156, 3678–3686 (1996).

  29. 29.

    Sutton, V. R. et al. Granzyme B triggers a prolonged pressure to die in Bcl-2 overexpressing cells, defining a window of opportunity for effective treatment with ABT-737. Cell Death Dis. 3, e344 (2012).

  30. 30.

    Jenkins, M. R. et al. Failed CTL/NK cell killing and cytokine hypersecretion are directly linked through prolonged synapse time. J. Exp. Med. 212, 307–317 (2015).

  31. 31.

    Stepp, S. E. et al. Perforin gene defects in familial hemophagocytic lymphohistiocytosis. Science 286, 1957–1959 (1999).

  32. 32.

    Calabia-Linares, C. et al. Endosomal clathrin drives actin accumulation at the immunological synapse. J. Cell Sci. 124, 820–830 (2011).

  33. 33.

    Corbera-Bellalta, M. et al. Blocking interferon γ reduces expression of chemokines CXCL9, CXCL10 and CXCL11 and decreases macrophage infiltration in ex vivo cultured arteries from patients with giant cell arteritis. Ann. Rheum. Dis. 75, 1177–1186 (2016).

  34. 34.

    Foley, J. F. et al. Roles for CXC chemokine ligands 10 and 11 in recruiting CD4+ T cells to HIV-1-infected monocyte-derived macrophages, dendritic cells, and lymph nodes. J. Immunol. 174, 4892–4900 (2005).

  35. 35.

    Reinhart, T. A. et al. Increased expression of the inflammatory chemokine CXC chemokine ligand 9/monokine induced by interferon-? in lymphoid tissues of rhesus macaques during simian immunodeficiency virus infection and acquired immunodeficiency syndrome. Blood 99, 3119–3128 (2002).

  36. 36.

    de Poot, S. A. et al. Granzyme M targets topoisomerase II alpha to trigger cell cycle arrest and caspase-dependent apoptosis. Cell Death Differ. 21, 416–426 (2014).

  37. 37.

    Ewen, C. L., Kane, K. P. & Bleackley, R. C. Granzyme H induces cell death primarily via a Bcl-2-sensitive mitochondrial cell death pathway that does not require direct Bid activation. Mol. Immunol. 54, 309–318 (2013).

  38. 38.

    Liu, J. & Roederer, M. Differential susceptibility of leukocyte subsets to cytotoxic T cell killing: implications for HIV immunopathogenesis. Cytometry A 71, 94–104 (2007).

  39. 39.

    Medema, J. P. et al. Expression of the serpin serine protease inhibitor 6 protects dendritic cells from cytotoxic T lymphocyte-induced apoptosis: differential modulation by T helper type 1 and type 2 cells. J. Exp. Med. 194, 657–667 (2001).

  40. 40.

    Migueles, S. A. et al. HIV-specific CD8+ T cell proliferation is coupled to perforin expression and is maintained in nonprogressors. Nat. Immunol. 3, 1061–1068 (2002).

  41. 41.

    Wherry, E. J. T cell exhaustion. Nat. Immunol. 12, 492–499 (2011).

  42. 42.

    Hersperger, A. R. et al. Perforin expression directly ex vivo by HIV-specific CD8 T-cells is a correlate of HIV elite control. PLoS Pathog. 6, e1000917 (2010).

  43. 43.

    Trimble, L. A. & Lieberman, J. Circulating CD8 T lymphocytes in human immunodeficiency virus-infected individuals have impaired function and downmodulate CD3 zeta, the signaling chain of the T-cell receptor complex. Blood 91, 585–594 (1998).

  44. 44.

    Draenert, R. et al. Persistent recognition of autologous virus by high-avidity CD8 T cells in chronic, progressive human immunodeficiency virus type 1 infection. J. Virol. 78, 630–641 (2004).

  45. 45.

    Appay, V. et al. HIV-specific CD8+ T cells produce antiviral cytokines but are impaired in cytolytic function. J. Exp. Med. 192, 63–75 (2000).

  46. 46.

    Migueles, S. A. et al. Lytic granule loading of CD8+ T cells is required for HIV-infected cell elimination associated with immune control. Immunity 29, 1009–1021 (2008).

  47. 47.

    Migueles, S. A. et al. Defective human immunodeficiency virus-specific CD8+ T-cell polyfunctionality, proliferation, and cytotoxicity are not restored by antiretroviral therapy. J. Virol. 83, 11876–11889 (2009).

  48. 48.

    Pachlopnik Schmid, J. et al. Neutralization of IFNγ defeats haemophagocytosis in LCMV-infected perforin- and Rab27a-deficient mice. EMBO Mol. Med. 1, 112–124 (2009).

  49. 49.

    Critchfield, J. W. et al. Magnitude and complexity of rectal mucosa HIV-1-specific CD8+ T-cell responses during chronic infection reflect clinical status. PLoS One 3, e3577 (2008).

  50. 50.

    Pereyra, F. et al. Increased coronary atherosclerosis and immune activation in HIV-1 elite controllers. AIDS 26, 2409–2412 (2012).

  51. 51.

    Pereyra, F. et al. HIV control is mediated in part by CD8+ T-cell targeting of specific epitopes. J. Virol. 88, 12937–12948 (2014).

  52. 52.

    Salerno-Goncalves, R., Fernandez-Vina, M., Lewinsohn, D. M. & Sztein, M. B. Identification of a human HLA-E-restricted CD8+ T cell subset in volunteers immunized with Salmonella enterica serovar Typhi strain Ty21a typhoid vaccine. J. Immunol. 173, 5852–5862 (2004).

  53. 53.

    Metkar, S. S. et al. Granzyme B activates procaspase-3 which signals a mitochondrial amplification loop for maximal apoptosis. J. Cell Biol. 160, 875–885 (2003).

  54. 54.

    Bellucci, R. et al. Tyrosine kinase pathways modulate tumor susceptibility to natural killer cells. J. Clin. Invest. 122, 2369–2383 (2012).

Download references


We thank S. Buus (University of Copenhagen) for monomers; G. Mylvaganam, G. Gaiha, T. Diefenbach and A. Balazs for comments; J. Trapani for discussions about granzymes; A. Piechocka-Trocha for experimental help; and the Flow Cytometry and Sample Processing Cores at the Ragon Institute for help with instrumentation and processing of the samples. Supported by the Howard Hughes Medical Institute (D.R.C. and B.D.W.), the Ragon Institute of MGH, MIT and Harvard (B.D.W.), the Canadian Institute of Health Research (K.L.C.), the US National Institutes of Health (R01 AI118544 to B.D.W.), amfAR (109326-59-RGRL to K.L.C. and B.D.W.) and the Harvard University Center for AIDS Research (P30 AI060354 to B.D.W.), which is supported by the following institutes and centers co-funded by and participating with the US National Institutes of Health: NIAID, NCI, NICHD, NHLBI, NIDA, NIMH, NIA, FIC and OAR.

Author information


  1. Ragon Institute of MGH, MIT and Harvard, Cambridge, MA, USA

    • Kiera L. Clayton
    • , David R. Collins
    • , Josh Lengieza
    • , Musie Ghebremichael
    •  & Bruce D. Walker
  2. Howard Hughes Medical Institute, Chevy Chase, MD, USA

    • David R. Collins
    •  & Bruce D. Walker
  3. Program in Cellular and Molecular Medicine, Boston Children’s Hospital, Boston, MA, USA

    • Farokh Dotiwala
    •  & Judy Lieberman
  4. Department of Pediatrics, Harvard Medical School, Boston, MA, USA

    • Farokh Dotiwala
    •  & Judy Lieberman
  5. Institute of Medical Engineering and Sciences, Massachusetts Institute of Technology, Cambridge, MA, USA

    • Bruce D. Walker


  1. Search for Kiera L. Clayton in:

  2. Search for David R. Collins in:

  3. Search for Josh Lengieza in:

  4. Search for Musie Ghebremichael in:

  5. Search for Farokh Dotiwala in:

  6. Search for Judy Lieberman in:

  7. Search for Bruce D. Walker in:


K.L.C. designed and performed the experiments and wrote the manuscript; D.R.C and B.D.W. contributed to the design of the experiments and writing of the manuscript; J. Lengieza performed experiments and optimized macrophage infection conditions; M.G provided advice for statistical analysis; F.D contributed to the discussions of granzyme-induced cell-death experiments; J. Lieberman provided guidance and design for apoptosis, granzyme and perforin experiments and for editing of the manuscript; and B.D.W. provided overall supervision of the project.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to Bruce D. Walker.

Integrated supplementary information

  1. Supplementary Figure 1 Experimental setups related to Fig. 1.

    Schematic of the experimental process to coordinate maturation, activation, and infection of target cells with the creation of HIV-specific CTL lines and setup of ex vivo CTLs. Details are described in the Methods.

  2. Supplementary Figure 2 Elimination assay gating strategy related to Fig. 1.

    Following a 4-hour co-culture between CellTrace Violet-stained CTLs and either infected autologous CD4+ T cells or macrophages, cells were stained with antibodies for CD3 (T cell only cultures), CD14 (macrophage cultures) and CD4, with a LIVE/DEAD fixable stain, and an antibody for intracellular Gag p24. Flow cytometry analysis for all elimination assay samples was performed as shown in these plots.

  3. Supplementary Figure 3 Ex vivo CTL elimination of human-serum differentiated macrophages, peptide-loaded targets, and resting CD4+ T cells related to Fig. 1.

    (a) Human-serum derived macrophages are relatively resistant to CTL-mediated killing. Monocytes were isolated from PBMCs as described in the Materials and Methods and differentiated into macrophages using 10% human serum in RPMI-1640. CD4+ T cells were activated in parallel followed by HIV infection (strain JRSCF) of both cell types five days after activation/differentiation. Two days after infection, HIV-infected macrophages and CD4+ T cells were co-cultured with autologous CellTrace Violet-stained ex vivo CTLs for 4 hours at multiple effector to target (E:T) ratios and elimination of infected (HIV Gag-p24+) populations was assessed via flow cytometry. Shown is the summary of elimination assays from HIV-infected patients (n = 18 individual samples from six independent experiments). Elimination assays using CTLs from an HIV- donor was used as a control (dotted lines). Shown are the means +/- SEM. Statistical analysis, two-sided unpaired t-test, *p<0.0001. (b) Ex vivo CTL elimination of peptide-loaded targets. CellTrace Violet-stained ex vivo CTLs were co-cultured with 50% peptide-loaded CellTrace FarRed-stained CD4+ T cells or macrophages for 4 hours followed by flow cytometry analysis of live targets. Shown is the summary from HIV+ patients (n = 4 individual samples from one experiment). Shown are the means +/− SEM. Statistical analysis, two-sided unpaired t-test; *p = 0.001. (c) CellTrace Violet-stained expanded CTLs were co-cultured with 50% peptide-loaded CellTrace FarRed-stained resting (ex vivo) CD4+ T cells, activated CD4+ T cells, or macrophages for 4 hours followed by flow cytometry analysis of live targets by flow cytometry. Shown is the summary of CEF responses from HIV- donors for activated CD4+ T cells and macrophages (n = 6 individual samples from three independent experiments) and for resting CD4+ T cells (n = 3 distinct samples from one experiment). Shown are the means +/− SEM. Statistical analysis, two-sided unpaired t-test; * p = 0.0317, ** p = 0.0011.

  4. Supplementary Figure 4 Elimination assay time course related to Fig. 2.

    CellTrace Violet-stained expanded CTLs were co-cultured with peptide-loaded or unloaded CellTrace FarRed-stained CD4+ T cells or macrophages for 15 min, 1 h, 4 h or 12 h followed by flow cytometry analysis of live targets. Shown is the summary from HIV+ patients (n = 8 individual samples from two independent experiments). Shown are the means +/− SEM. Statistical analysis, two-sided unpaired t test; *p = 0.0286, ** p = 0.0004, ***p<0.0001.

  5. Supplementary Figure 5 Staining controls and cytolytic responses related to Fig. 4.

    (a) FMO for tetramer staining of CD8+ T cells. (b) Perforin, granzyme A, granzyme B, granzyme H, granzyme K, and granzyme M staining of naïve CD8+ T cells as negative controls for staining. (c) Perforin and granzyme staining of CD8+ T cell, CMV-tetramer+ cells as a comparison for Fig. 4a. (d) Perforin and granzyme FMO staining controls for the antigen-specific CD8+ T cells in Fig. 4d. (e) Cytolytic capacity of HIV-specific CD8+ T cells responding to infected CD4+ T cells and macrophages. Ex vivo or expanded CTL effector cells were incubated with HIV-infected CD4+ T cells or macrophages at an E:T of 2 for 6 hours followed by flow cytometry-based analysis of degranulation (surface CD107a expression) and perforin and granzyme expression. Shown are the summaries of the perforin and granzyme phenotyping of CTLs responding to infected CD4+ T cells or macrophages for HIV+ patients (n = 8 individual samples for two independent experiments). Box elements, center line, limits and whiskers are the median, 25th-75th percentiles and min-max, respectively. Statistical analysis, two-sided paired t-test.

  6. Supplementary Figure 6 ImageStream-based assessment of MHC-I surface density and control recognition assay related to Fig. 5.

    (a) Representative image of CD4+ T cell and macrophage samples acquired on the ImageStream. CD4+ T cells and macrophages were stained for surface MHC-I (clone W6/32) and LIVE/DEAD Near-IR followed by fixing and staining for actin. (b) Representative dot plots of the gating strategy used to gate on live cells followed by assessment of cell size (diameter) versus MHC-I intensity. (c) HIV- donor recognition assay responses. Ex vivo CTL effector cells were incubated with mock or HIV-infected CD4+ T cells or macrophages at an E:T of 2 for 6 hours followed by flow cytometry-based analysis of degranulation (surface CD107a expression). Shown is a representative analysis of CTL effector cells degranulation responses from an HIV- donor.

  7. Supplementary Figure 7 Chemokine receptor expression related to Fig. 7.

    Cells used for chemotaxis assays were phenotyped for chemokine receptor expression. CXCR3 is the receptor for CXCL9, CXCL10 and CXCL11. CCR2 is the receptor for CCL2. CCR5 is the receptor for MIP-1α, MIP-1β, and RANTES.

Supplementary information

About this article

Publication history





Further reading