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Perivascular macrophages mediate neutrophil recruitment during bacterial skin infection

Nature Immunology volume 15, pages 45–53 (2014)Cite this article

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Abstract

Transendothelial migration of neutrophils in postcapillary venules is a key event in the inflammatory response against pathogens and tissue damage. The precise regulation of this process is incompletely understood. We report that perivascular macrophages are critical for neutrophil migration into skin infected with the pathogen Staphylococcus aureus. Using multiphoton intravital microscopy we showed that neutrophils extravasate from inflamed dermal venules in close proximity to perivascular macrophages, which are a major source of neutrophil chemoattractants. The virulence factor α-hemolysin produced by S. aureus lyses perivascular macrophages, which leads to decreased neutrophil transmigration. Our data illustrate a previously unrecognized role for perivascular macrophages in neutrophil recruitment to inflamed skin and indicate that S. aureus uses hemolysin-dependent killing of these cells as an immune evasion strategy.

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Figure 1: Hla reduces neutrophil influx in S. aureus–infected skin, resulting in increased bacterial survival.
Figure 2: Hla-mediated prevention of neutrophil extravasation during cutaneous S. aureus infection.
Figure 3: Perivascular macrophages can be identified in transgenic DPE-GFP mice.
Figure 4: Neutrophils extravasate adjacent to perivascular macrophages in inflamed dermal vessels.
Figure 5: Hla specifically lyses ADAM10+ macrophages.
Figure 6: Specific lysis of DPE-GFP+ PVM by ΔHla S. aureus in vivo.

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Acknowledgements

We thank M. Willcox (University of New South Wales, Sydney) for S. aureus strains, E. Harry (University of Technology Sydney) for S. aureus USA300, T. Graf (Center for Genomic Regulation, Barcelona, Spain) for providing Lyz2gfp/+ mice, D. Hume (University of Edinburgh) for Csf1r-EGFP mice, M.J. Fraunholz (University of Würzburg Biocenter) for the pmRFPmars plasmid, A. Smith, S. Allen and S. Dervish for their technical assistance, and M. Rizk and J. Qin for animal husbandry. This work was supported by grant 1030147 from the National Health and Medical Research Council of Australia (to A.A., R.J., N.F. and W.W.).

Author information

Author notes

  1. Arby Abtin, Rohit Jain and Andrew J Mitchell: These authors contributed equally to this work.

Authors and Affiliations

  1. The Centenary Institute, Newtown, New South Wales, Australia

    Arby Abtin, Rohit Jain, Andrew J Mitchell, Ben Roediger, Shweta Tikoo, Lois L Cavanagh & Wolfgang Weninger

  2. School of Biological Sciences, University of Sydney, New South Wales, Australia

    Anthony J Brzoska & Neville Firth

  3. Monash University Department of Medicine, Centre for Inflammatory Diseases, Monash Medical Centre, Clayton, Victoria, Australia

    Qiang Cheng & Michael J Hickey

  4. Singapore Immunology Network (SIgN), Agency for Science, Technology and Research (A*STAR), Biopolis, Singapore

    Lai Guan Ng

  5. Sydney Medical School, New South Wales, Australia

    Lois L Cavanagh

  6. Immune Disease Institute and Division of Immunology, Department of Microbiology and Immunobiology, Harvard Medical School, Boston, Massachusetts, USA

    Ulrich H von Andrian

  7. Discipline of Dermatology, Sydney Medical School, New South Wales, Australia

    Wolfgang Weninger

  8. Department of Dermatology, Royal Prince Alfred Hospital, Camperdown, New South Wales, Australia

    Wolfgang Weninger

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Contributions

A.A. and W.W. conceived the idea for this project. A.A., R.J., A.J.M. and W.W. wrote the paper. A.A., A.J.M. and B.R. performed immunological and flow cytometry experiments and assisted with imaging experiments. R.J. designed and conducted intravital and confocal imaging experiments. B.R. conducted flow cytometry and imaging experiments. R.J. and A.J.M. performed sorting experiments for perivascular macrophages. A.J.M. performed quantitative PCR experiments and cytometric bead arrays. R.J., A.J.M., B.R. and S.T. performed all other experiments. A.J.B. and N.F. generated fluorescent protein–expressing bacteria. All authors discussed the results and commented on the paper.

Corresponding author

Correspondence to Wolfgang Weninger.

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Integrated supplementary information

Supplementary Figure 1 Effect of Hla on bacterial infection parameters.

(a) Clinical manifestations, (b) Ear thickness, (c) RBC count following infection of C57BL/6 mice with S. aureus (WT or ΔHla) at various time points. (d) Ear swelling response and (e) RBC count of concomitant injection of purified Hla together with S. aureus (ΔHla) 12h p.i. (f) Neutrophil myeloperoxidase activity in back skin of mice infected with S. aureus (WT or ΔHla) at 12h p.i. MPO activity was normalized to tissue weight. (g) Neutrophil influx into ears infected with S. aureus strain USA300 in the presence or absence of anti-Hla serum. Bars represent mean±SD. Student's two-tailed unpaired t-test was used to determine the P-value. Data are representative of at least two independent experiments. Error bars indicate SD. **P<0.01; ***P<0.001.

Supplementary Figure 2 Hla suppresses the homing of adoptively transferred Lyz2gfp/+ neutrophils to the site of infection.

(a) Confocal imaging of extravasated Lyz2gfp/+ neutrophils in dermis of mice infected with S. aureus (WT or ΔHla) 6h p.i. and 1h following Lyz2gfp/+ bone marrow cell transfer. Blood vessels are visualized by CD31 staining. Scale bar represents 100μm. (b) Flow cytometric analysis of S. aureus (ΔHla) infected tissue 6h p.i. and 1h following Lyz2gfp/+ bone marrow cell transfer showing percentage of extravasated neutrophils (Ly6G+Ly6Cint) and monocytes (Ly6G-Ly6Chi). (c) CCL2, CCL3 and CCL4 protein levels in ears of mice infected with S. aureus (WT or ΔHla). Bars represent mean±SD of five to six mice/group/time point. One-way ANOVA with Tukey's post-test was used to determine the P-value. ***P<0.001.

Supplementary Figure 3 Phenotypic analysis of PVM.

(a) Flow cytometric analysis of GFP+ and GFP- macrophages from skin of DPE-GFP mice in comparison to red pulp and peritoneal macrophages. (b) Confocal imaging of cremaster muscle showing the localization of PVM (DPE-GFP, green) in relation to pericytes (anti-smooth muscle actin, red) and endothelial cells (CD31, blue). (c) Expression of Csf1r mRNA transcripts in GFP+ macrophages from DPE-GFP mice. Bars represent mean±SD of 3 independent experiments. (d) Comparative mRNA expression of indicated genes in GFP+ vs. GFP- dermal macrophages from DPE-GFP mice. Bars represent mean±SD of 3 independent experiments. (e) Intravital imaging of ears from Csf1r-EGFP mice showing presence of GFP+ PVM (asterisk). Student's two-tailed unpaired t-test was used to determine the P-value. *P<0.05

Supplementary Figure 4 Neutrophils extravasate adjacent to perivascular macrophages during E. coli infection.

(a) Figure depicts the methodology employed to calculate predicted neutrophil to macrophage correlation (detailed in Materials and Methods section). The total observed extravasation tracks for the blood vessel imaged are overlayed in comparison to the predicted tracks. The insets depict one such event of neutrophil extravasation (red) from blood vessel (blue, vessel wall marked in cyan) in relation to PVM (green, asterisk in main figure). 00:00, min:sec. (b) Time-lapse intravital multi-photon imaging of ear dermis of DPE-GFP mice infected with E. coli depicting the adherence and extravasation of adoptively transferred cells isolated from mT/mG mice (red, numbered 1-4) in close proximity to PVM (DPE-GFP, green). Lines (white) represent migration tracks of selected neutrophils (red) 00:00:00, hr:min:sec. (c) Statistical analysis of neutrophil extravasation site with respect to perivascular macrophages. Numbers of neutrophils extravasating at theoretical/random (white bars) or observed (black bars) distances from GFP+ PVM within the same vessels. Data is cumulative of 50 events of extravasation from 3 independent experiments. (d) Neutrophil influx, (e) ear thickness and (f) RBC counts of ears concomitantly injected with E. coli and Hla (10μg) or only E. coli alone for 12h p.i. Data shown are mean±SEM of 3 independent experiments. P-values were calculated using two-way ANOVA or two-tailed unpaired Student's t-test, **P<0.01; ***P<0.001; ****P<0.0001; NS, not significant.

Supplementary Figure 5 Vascular integrity is unaffected by S. aureus infection.

(a) Confocal imaging of ear whole mounts from untreated DPE-GFP mice (left panels) or animals infected with S. aureus (WT) (right panels). Basement membranes are delineated by laminin and endothelium by CD31 labeling. (b) Intravital imaging of Evans blue (red; conjugated to BSA) leakage from S. aureus (ΔHla or WT) infected tissue 6h p.i. Adoptively transferred neutrophils (green) and SHG (blue) are also depicted. The images are tiles of 1mmx1mm region of ear tissue. (c) Confocal imaging of Thy-1 stained ear whole mounts from untreated DPE-GFP mice (left panels) or animals infected with S. aureus (WT) (right panels) showing no appreciable loss of Thy-1+ cells.

Supplementary Figure 6 Comparable uptake of wild-type and ΔHla S. aureus by neutrophils.

(a) Percent Ly6G+ neutrophils containing RFP-expressing S. aureus strains (WT or ΔHla) were determined by flow cytometry at 12h p.i. (b) Representative dotplots illustrating fluorescence intensity of neutrophils containing RFP-expressing S. aureus (WT or ΔHla). (c) Fluorescence intensity of RFP expressing S. aureus strains as determined by flow cytometry. Data are representative of two independent experiments (four to five mice/group). Error bars indicate SD.

Supplementary information

Supplementary Text and Figures

Supplementary Figures 1–6 (PDF 13883 kb)

Supplementary Movie 1.

Time-lapse intravital imaging of mice infected with either WT or ΔHla S. aureus depicting neutrophil rolling and sticking in dermal blood vessels (red). The mice were infected 6 h before imaging and neutrophils (green) were adoptively transferred immediately before imaging. Time represents min:s. Increased red background in S. aureus (ΔHla) infection is observed due inflammation induced vascular leakage of Evans blue. (AVI 7381 kb)

Supplementary Movie 2.

Time-lapse intravital imaging of ear dermis of mice infected with S. aureus (ΔHla) depicting hotspots for neutrophil sticking and extravasation from the blood vessel (red). The mice were infected 6 h before imaging, and neutrophils (green) were adoptively transferred immediately prior to imaging. Time represents min:s. (AVI 4718 kb)

Supplementary Movie 3.

Z stack of ear dermis of DPE-GFP mice showing the localisation of GFP+ macrophages (green) along dermal blood vessel (red). Second harmonic generation is depicted in blue. (AVI 454 kb)

Supplementary Movie 4.

Three-dimensional time-lapse reconstruction of Z stacks of ear dermis of DPE-GFP mice showing the homeostatic scanning behaviour of GFP+ macrophages. Time represents h:min:s. (MOV 455 kb)

Supplementary Movie 5.

Time-lapse intravital imaging of ear dermis of mice infected with S. aureus (ΔHla) depicting neutrophil (red) extravasation in relation to GFP+ perivascular macrophages (green). The mice were infected 6 h before imaging, and neutrophils were adoptively transferred immediately before imaging. Time represents h:min:s. (MOV 4240 kb)

Supplementary Movie 6.

In vitro time-lapse imaging of macrophage (green) death in the presence of Hla (7.5μg/ml). Time represents min:s after addition of Hla. (AVI 632 kb)

Supplementary Movie 7.

Three-dimensional time lapse reconstruction of Z stacks of ear dermis of DPE-GFP mice infected with S. aureus (WT) (red) showing the behaviour of GFP+ macrophages in presence of S. aureus (WT). Imaging was performed immediately after injection of the bacterium. Time represents h:min:s. (MOV 1022 kb)

Supplementary Movie 8.

Three-dimensional time lapse reconstruction of Z stacks of ear dermis of DPE-GFP mice infected with S. aureus (ΔHla) (red) showing the behavior of GFP+ macrophages in presence of ΔHla S. aureus (ΔHla). Imaging was performed immediately after injection of the bacterium. Time represents h:min:s. (MOV 2097 kb)

Supplementary Movie 9.

Time-lapse intravital imaging of ear dermis of mice injected with RFP-expressing S. aureus (WT) (red) in relation to perivascular macrophages (green). The mice were injected just before imaging. Time represents h:min:s. (MOV 3846 kb)

Supplementary Movie 10.

Time-lapse intravital imaging of ear dermis of mice injected with fluorescent beads (red) in relation to perivascular macrophages (green). The mice were injected just before imaging. Time represents h:min:s. (MOV 4825 kb)

Supplementary Movie 11.

Time-lapse intravital imaging of ear dermis of mice injected with RFP-expressing S. aureus (ΔHla) (red) in relation to perivascular macrophages (green). The mice were injected just before imaging. Time represents h:min:sec. (MOV 7552 kb)

Supplementary Movie 12.

Time-lapse intravital imaging of ear dermis of dermal dendritic cells (yellow) in CD11c-YFP mice injected with RFP-expressing S. aureus (WT) (red). The mice were injected just before imaging. Time represents h:min: (MOV 3597 kb)

Supplementary Movie 13.

Time-lapse intravital imaging of ear dermis of γδ T cells (green) in CXCR6-GFP mice injected with RFP-expressing S. aureus (WT) (red). The mice were injected just before imaging. Time represents h:min:s. (MOV 4477 kb)

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Abtin, A., Jain, R., Mitchell, A. et al. Perivascular macrophages mediate neutrophil recruitment during bacterial skin infection. Nat Immunol 15, 45–53 (2014). https://doi.org/10.1038/ni.2769

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