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Aged polymorphonuclear leukocytes cause fibrotic interstitial lung disease in the absence of regulation by B cells

Nature Immunologyvolume 19pages192201 (2018) | Download Citation

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Abstract

Pulmonary immunity requires tight regulation, as interstitial inflammation can compromise gas exchange and lead to respiratory failure. Here we found a greater number of aged CD11bhiL-selectinloCXCR4+ polymorphonuclear leukocytes (PMNs) in lung vasculature than in the peripheral circulation. Using pulmonary intravital microscopy, we observed lung PMNs physically interacting with B cells via β2 integrins; this initiated neutrophil apoptosis, which led to macrophage-mediated clearance. Genetic deletion of B cells led to the accumulation of aged PMNs in the lungs without systemic inflammation, which caused pathological fibrotic interstitial lung disease that was attenuated by the adoptive transfer of B cells or depletion of PMNs. Thus, the lungs are an intermediary niche in the PMN lifecycle wherein aged PMNs are regulated by B cells, which restrains their potential to cause pulmonary pathology.

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References

  1. 1.

    Bagnato, G. & Harari, S. Cellular interactions in the pathogenesis of interstitial lung diseases. Eur. Respir. Rev. 24, 102–114 (2015).

  2. 2.

    Shum, A. K. et al. Identification of an autoantigen demonstrates a link between interstitial lung disease and a defect in central tolerance. Sci. Transl. Med. 1, 9ra20 (2009).

  3. 3.

    Wells, A. U. & Denton, C. P. Interstitial lung disease in connective tissue disease–mechanisms and management. Nat. Rev. Rheumatol. 10, 728–739 (2014).

  4. 4.

    Travis, W. D. et al. An official American Thoracic Society/European Respiratory Society statement: update of the international multidisciplinary classification of the idiopathic interstitial pneumonias. Am. J. Respir. Crit. Care. Med. 188, 733–748 (2013).

  5. 5.

    Hasan, S. A. et al. Role of IL-17A and neutrophils in fibrosis in experimental hypersensitivity pneumonitis. J. Allergy. Clin. Immunol. 131, 1663–1673 (2013).

  6. 6.

    Kuebler, W. M. & Goetz, A. E. The marginated pool. Eur. Surg. Res. 34, 92–100 (2002).

  7. 7.

    Doerschuk, C. M. et al. Marginated pool of neutrophils in rabbit lungs. J. Appl. Physiol. 63, 1806–1815 (1987).

  8. 8.

    Yipp, B. G. et al. The lung is a host defense niche for immediate neutrophil-mediated vascular protection. Science Immunol. 2, 1–13 (2017).

  9. 9.

    Zhang, D. et al. Neutrophil ageing is regulated by the microbiome. Nature 525, 528–532 (2015).

  10. 10.

    Uhl, B. et al. Aged neutrophils contribute to the first line of defense in the acute inflammatory response. Blood 128, 2327–2337 (2016).

  11. 11.

    Colom, B. et al. Leukotriene B4-neutrophil elastase axis drives neutrophil reverse transendothelial cell migration in vivo. Immunity 42, 1075–1086 (2015).

  12. 12.

    Woodfin, A. et al. The junctional adhesion molecule JAM-C regulates polarized transendothelial migration of neutrophils in vivo. Nat. Immunol. 12, 761–769 (2011).

  13. 13.

    Cerutti, A., Puga, I. & Magri, G. The B cell helper side of neutrophils. J. Leukoc. Biol. 94, 677–682 (2013).

  14. 14.

    Kamenyeva, O. et al. Neutrophil recruitment to lymph nodes limits local humoral response to Staphylococcus aureus. PLoS. Pathog. 11, e1004827 (2015).

  15. 15.

    Puga, I. et al. B cell-helper neutrophils stimulate the diversification and production of immunoglobulin in the marginal zone of the spleen. Nat. Immunol. 13, 170–180 (2011).

  16. 16.

    Looney, M. R. et al. Stabilized imaging of immune surveillance in the mouse lung. Nat. Methods 8, 91–96 (2011).

  17. 17.

    Evans, R. et al. Integrins in immunity. J. Cell. Sci. 122, 215–225 (2009).

  18. 18.

    Matsushima, H. et al. Neutrophil differentiation into a unique hybrid population exhibiting dual phenotype and functionality of neutrophils and dendritic cells. Blood 121, 1677–1689 (2013).

  19. 19.

    Gosselin, E. J., Wardwell, K., Rigby, W. F. & Guyre, P. M. Induction of MHC class II on human polymorphonuclear neutrophils by granulocyte/macrophage colony-stimulating factor, IFN-γ, and IL-3. J. Immunol. 151, 1482–1490 (1993).

  20. 20.

    Casanova-Acebes, M. et al. Rhythmic modulation of the hematopoietic niche through neutrophil clearance. Cell 153, 1025–1035 (2013).

  21. 21.

    Coxon, A. et al. A novel role for the β2 integrin CD11b/CD18 in neutrophil apoptosis: a homeostatic mechanism in inflammation. Immunity 5, 653–666 (1996).

  22. 22.

    Weinmann, P., Scharffetter-Kochanek, K., Forlow, S. B., Peters, T. & Walzog, B. A role for apoptosis in the control of neutrophil homeostasis in the circulation: insights from CD18-deficient mice. Blood 101, 739–746 (2003).

  23. 23.

    Buch, T. et al. A Cre-inducible diphtheria toxin receptor mediates cell lineage ablation after toxin administration. Nat. Methods 2, 419–426 (2005).

  24. 24.

    Demircik, F., Buch, T. & Waisman, A. Efficient B cell depletion via diphtheria toxin in CD19-Cre/iDTR mice. PLoS. ONE. 8, e60643 (2013).

  25. 25.

    Eash, K. J., Means, J. M., White, D. W. & Link, D. C. CXCR4 is a key regulator of neutrophil release from the bone marrow under basal and stress granulopoiesis conditions. Blood. 113, 4711–4719 (2009).

  26. 26.

    Soehnlein, O. & Lindbom, L. Phagocyte partnership during the onset and resolution of inflammation. Nat. Rev. Immunol. 10, 427–439 (2010).

  27. 27.

    Brain, J. D., Molina, R. M., DeCamp, M. M. & Warner, A. E. Pulmonary intravascular macrophages: their contribution to the mononuclear phagocyte system in 13 species. Am. J. Physiol. Lung. Cell. Mol. Physiol. 276, L146–L154 (1999).

  28. 28.

    Smith, M. L. Update on pulmonary fibrosis: not all fibrosis is created equally. Arch. Pathol. Lab. Med. 140, 221–229 (2016).

  29. 29.

    Watanabe, R. et al. Regulatory B cells (B10 cells) have a suppressive role in murine lupus: CD19 and B10 cell deficiency exacerbates systemic autoimmunity. J. Immunol. 184, 4801–4809 (2010).

  30. 30.

    Yanaba, K. et al. A regulatory B cell subset with a unique CD1dhiCD5+ phenotype controls T cell-dependent inflammatory responses. Immunity 28, 639–650 (2008).

  31. 31.

    Haas, K. M., Poe, J. C., Steeber, D. A. & Tedder, T. F. B-1a and B-1b cells exhibit distinct developmental requirements and have unique functional roles in innate and adaptive immunity to S. pneumoniae. Immunity 23, 7–18 (2005).

  32. 32.

    Bosma, G. C. et al. Evidence of functional lymphocytes in some (leaky) scid mice. J. Exp. Med. 167, 1016–1033 (1988).

  33. 33.

    Enomoto, K. et al. Bronchoalveolar lavage fluid cells in mixed connective tissue disease. Respirology 8, 149–156 (2003).

  34. 34.

    Martin, C. et al. Chemokines acting via CXCR2 and CXCR4 control the release of neutrophils from the bone marrow and their return following senescence. Immunity 19, 583–593 (2003).

  35. 35.

    Devi, S. et al. Neutrophil mobilization via plerixafor-mediated CXCR4 inhibition arises from lung demargination and blockade of neutrophil homing to the bone marrow. J. Exp. Med. 210, 2321–2336 (2013).

  36. 36.

    Mayadas, T. N. & Cullere, X. Neutrophil β2 integrins: moderators of life or death decisions. Trends. Immunol. 26, 388–395 (2005).

  37. 37.

    Geering, B. & Simon, H. U. Peculiarities of cell death mechanisms in neutrophils. Cell. Death. Differ. 18, 1457–1469 (2011).

  38. 38.

    Duffin, R., Leitch, A. E., Fox, S., Haslett, C. & Rossi, A. G. Targeting granulocyte apoptosis: mechanisms, models, and therapies. Immunol. Rev. 236, 28–40 (2010).

  39. 39.

    Luo, H. R. & Loison, F. Constitutive neutrophil apoptosis: mechanisms and regulation. Am. J. Hematol. 83, 288–295 (2008).

  40. 40.

    Wu, D. et al. Reverse-migrated neutrophils regulated by JAM-C are involved in acute pancreatitis-associated lung injury. Sci. Rep. 6, 20545 (2016).

  41. 41.

    Wang, J. et al. Visualizing the function and fate of neutrophils in sterile injury and repair. Science 358, 111–116 (2017).

  42. 42.

    Wallace, B., Vummidi, D. & Khanna, D. Management of connective tissue diseases associated interstitial lung disease: a review of the published literature. Curr. Opin. Rheumatol. 28, 236–245 (2016).

  43. 43.

    Roubille, C. & Haraoui, B. Interstitial lung diseases induced or exacerbated by DMARDS and biologic agents in rheumatoid arthritis: a systematic literature review. Semin. Arthritis. Rheum. 43, 613–626 (2014).

  44. 44.

    Lioté, H., Lioté, F., Séroussi, B., Mayaud, C. & Cadranel, J. Rituximab-induced lung disease: A systematic literature review. Eur. Respir. J. 35, 681–687 (2010).

  45. 45.

    Maarschalk-Ellerbroek, L. J. et al. CT screening for pulmonary pathology in common variable immunodeficiency disorders and the correlation with clinical and immunological parameters. J. Clin. Immunol. 34, 642–654 (2014).

  46. 46.

    Verma, N., Grimbacher, B. & Hurst, J. R. Lung disease in primary antibody deficiency. Lancet Respir. Med. 3, 651–660 (2015).

  47. 47.

    Shull, M. M. et al. Targeted disruption of the mouse transforming growth factor-β1 gene results in multifocal inflammatory disease. Nature 359, 693–699 (1992).

  48. 48.

    Thomas, B. J., Kan-O, K., Loveland, K. L., Elias, J. A. & Bardin, P. G. In the shadow of fibrosis: innate immune suppression mediated by transforming growth factor-β. Am. J. Respir. Cell. Mol. Biol. 55, 759–766 (2016).

  49. 49.

    Aschner, Y. & Downey, G. P. Transforming growth factor-β: master regulator of the respiratory system in health and disease. Am. J. Respir. Cell. Mol. Biol. 54, 647–655 (2016).

  50. 50.

    Martinod, K. et al. Peptidylarginine deiminase 4 promotes age-related organ fibrosis. J. Exp. Med. 214, 439–458 (2017).

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Acknowledgements

We thank P. Kubes for mentorship and lab members; L. Zbytnuik, T. Nussbaumer and M. Willson for assisting with reagents and animals; and E. De Heuvel for assistance in processing the histology; and acknowledge the use of the Snyder Mouse Phenomics Resources Laboratory and Live Cell imaging facility (funded by the Snyder Institute) and the flow cytometry core facility. Supported by the Canadian Foundation for Innovation–John R. Evans Leaders fund with matching support from the Alberta Enterprise and Advanced Education Research Capacity Program (infrastructure funding), the Canadian Institutes of Health Research (operating grant RS-342013), the Department of Critical Care Medicine of the University of Calgary (startup funds), the University of Calgary Medical Group (bridge funding), the Cumming School of Medicine Research Enhancement Program and a tier II Canada Research Chair in Pulmonary Immunology, Inflammation and Host Defense (B.G.Y.).

Author information

Affiliations

  1. Calvin, Phoebe and Joan Snyder Institute for Chronic Diseases, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Jung Hwan Kim
    • , John Podstawka
    • , Yuefei Lou
    • , Lu Li
    • , Esther K. S. Lee
    • , Margaret M. Kelly
    •  & Bryan G. Yipp
  2. Department of Critical Care Medicine, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Jung Hwan Kim
    • , John Podstawka
    • , Yuefei Lou
    • , Lu Li
    • , Esther K. S. Lee
    •  & Bryan G. Yipp
  3. Meakins-Christie Laboratories, Department of Medicine, Department of Microbiology and Immunology, Department of Pathology, McGill International TB Centre, McGill University Montreal, Montreal, QC, Canada

    • Maziar Divangahi
  4. Mouse Phenomics Resource Laboratory, Calvin, Phoebe and Joan Snyder Institute for Chronic Diseases, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Björn Petri
  5. Department of Microbiology, Immunology and Infectious Diseases, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Björn Petri
  6. Department of Biochemistry and Molecular Biology, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Frank R. Jirik
  7. Department of Pathology and Laboratory Medicine, Cumming School of Medicine, University of Calgary, Calgary, AB, Canada

    • Margaret M. Kelly

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Contributions

Conceptualization, J.H.K., M.D., F.R.J., M.M.K. and B.G.Y.; methodology, J.H.K., J.P., Y.L., L.L., E.K.S.L., B.P., F.R.J., M.M.K. and B.G.Y.; investigation, J.H.K., J.P., Y.L., L.L., E.K.S.L. and B.G.Y.; writing (original draft), J.H.K., E.K.S.L. and B.G.Y.; writing (review and editing), J.H.K., M.D., M.M.K. and B.G.Y.; funding acquisition, B.G.Y.; resources, M.M.K, F.R.J. and B.G.Y.; and supervision, B.G.Y.

Competing interests

The authors declare no competing financial interests.

Corresponding author

Correspondence to Bryan G. Yipp.

Integrated supplementary information

  1. Supplementary Figure 1 Intravital imaging model and flow cytometry gating strategy.

    (a) Pulmonary intravital imaging was used to assess CD19+ (fluorochrome conjugated anti-CD19 antibody), Ly6G+ PMN (fluorochrome conjugated anti-Ly6G antibody) and MHCII+ cells (fluorochrome conjugated anti-MHCII antibody). CD19+ MHCII+ cells were quantified per field of view. Experiments were repeated three times (n = 3). Data is mean ± s.e.m. (b) Flow cytometry was used to assess leukocyte populations. Leukocytes were identified by forward and side scatter. PMN were identified as Ly6G+ and CD19-, while B cells were identified by Ly6G- CD19+. Dead cells were excluded using propidium iodide (PI) staining.

  2. Supplementary Figure 2 In vitro MHCII transfer experiments.

    (a) Flow cytometry was used to assess B cells and PMN that were co-incubated in wells that allowed physical contact. MHCII cell surface levels were measured on PMN after B cell co-incubation in wells treated with an isotype control antibody, an irrelevant monoclonal antibody that binds PMN (anti-CD29 monoclonal antibody, clone Hmb1-1, 10 µg/mL) or an anti-CD18 blocking monoclonal antibody (clone GAME-46, 10 µg/ml). Experiments were repeated twice (n = 2). (b) PMN and B cell co-culture experiments were repeated in standard well that allow physical contact. B cells were pre-treated with Eα peptide prior to PMN incubation. MHCII (I-Ab) with properly configured Eα peptide can be detected with the Y-Ae monoclonal antibody. Ova peptide was used as a loading control and will not be detected by Y-Ae. PMN were verified to have acquired MHCII from B cells using a monoclonal antibody that detects I-Ab. Y-Ae antibody stains for B cells and PMN after peptide-loaded B cells are co-cultured with PMN are shown. (n = 3). (c) Single cell suspensions from blood and lung were treated with and without collagenase then cultured in vitro for 4-hours. After the culture, the percentage of MHCII+ annexinV+ early apoptotic PMN were analyzed. Each data point represents an individual experiment. Bars represent s.e.m. One way ANOVA with adjusted p values. * p = 0.02, ** p = 0.009.

  3. Supplementary Figure 3 B cell depletion in vivo.

    (a) Flow cytometry assessed B cell depletion in lung and blood of Rosa-DTR and CD19-DTR mice following DT (25 ng/g of body weight) injected intraperitoneal for four consecutive days. Blood and lung were analyzed for the presence of CD19+ B cells and Ly6G+ PMN. n = 3. (b) B cells were quantified in blood and lung in Rosa-DTR and CD19-DTR mice that received 4 days of DT. Unpaired Student’s t test, ** p = 0.0035 for blood and p = 0.0013 for lung. (c) CD19-DTR mice received DT (25 ng/g of body weight, Monday-Wednesday-Friday) for 3 weeks and received intravenous adoptively transferred C57BL/6 B cells weekly. The numbers of B cells in the lung were quantified using flow cytometry. Each data point represents an individual experiment (n). Bars represent s.e.m.

  4. Supplementary Figure 4 Interstitial PMN localization in mice depleted of B cells.

    Circulating PMN were identified by injecting mice with intravenous fluorochrome conjugated anti-CD45 antibody prior to tissue harvesting. Interstitial PMN were identified as Ly6G+ CD45-, while circulating PMN were identified as Ly6G+ CD45+. Alveolar cells, which are not in the circulation, were used to verify that the intravenous anti-CD45 antibody was not leaking from the blood into the tissue. Representative histogram is shown on the left for vascular and tissue pulmonary neutrophils. Bar graph displaying the percentage of CD45- interstitial neutrophils is shown on the right. Each data point represents an individual experiment (n). Bars represent s.e.m.

  5. Supplementary Figure 5 Co-depletion of B cells and PMNs.

    CD19-DTR or control (Rosa-DTR) mice received DT (25 ng/g of body weight, Monday-Wednesday-Friday) for 3 weeks. Additionally, mice received the PMN depleting antibody (1A8, 250 µg every Monday and Friday) intermittently during the 3-week B cells depletion. Flow cytometry shows PMN counts comparing control mice and B cell depleted mice with or without PMN depletion.

  6. Supplementary Figure 6 Macrophages clear MHCII+ aged PMNs.

    Clodronate (200 μg) was injected into mice intravenously for 24-hours and the percentage of F4/80+ cells were analyzed in bone marrow. The representative dot plots are shown on left and bar graphs showing the percentage are on the right. Unpaired Student’s t test, p = 0.0129, n = 3. (b) The percentage of MHCII+ and (c) MHCII+ annexin V+ early apoptotic PMN circulating in peripheral blood are shown after the clodronate injection. n=3. Bars represent s.e.m. * p = 0.048, ** p = 0.0072.

  7. Supplementary Figure 7 Assessments of immunodeficient mice.

    (a) B cells were quantified in the blood and lung of lymphocyte deficient mice (Rag1-/-). PMN were quantified in the blood and lung of Rag1-/-. Data represent the mean ± s.e.m. For panel a, each data point represents an individual experiment (n). * p = 0.02, ** p = 0.005. (b) CD19-DTR or Rosa-DTR mice received DT (25ng/g of body weight, Monday-Wednesday-Friday) for 3 weeks. Serum was collected at the end of 3 weeks and assayed for total IgG using standard ELISA. n = 4. Bars represent s.e.m. ***p < 0.0005.

  8. Supplementary Figure 8 Gating strategy for Ly6Chi and Ly6Clo monocytes.

    Facs assessed monocyte populations from DT-treated Rosa-DTR (Control) and CD19-DTR animals. Lung cells were gated from a CD45+, F4/80-, Ly6G-, CD115+ population. Ly6G+SSC+ neutrophils were gated from CD45+ leukocytes.

  9. Supplementary Figure 9 B cell deletion does not induce the systemic production of inflammatory mediators.

    ELISA assessed IFN-γ, GM-CSF, IL-1β, IL-2, IL-4, IL-6, IL-10, IL-12, MCP-1, TNF-α and KC from (a) lung tissue homogenates, (b) bronchoalveolar lavage (BAL) and (c) serum of Rosa26-DTR and CD19-DTR mice treated with 3 weeks of DT. All mediators were determined using multiplex except KC, which was assessed using a standard ELISA. Data represent the mean ± s.e.m. For panel a-f, each data point represents an individual experiment. Unpaired Student’s t test, *** p < 0.0006.

Supplementary information

Videos

  1. Supplementary Video 1

    Lung PMN and B cell interactions in vivo.

  2. Supplementary Video 2

    Lung PMN acquire MHCII from B cells.

  3. Supplementary Video 3

    Lung PMN are activated following B cell depletion.

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https://doi.org/10.1038/s41590-017-0030-x