A subset of neutrophils in human systemic inflammation inhibits T cell responses through Mac-1

Abstract

Suppression of immune responses is necessary to limit damage to host tissue during inflammation, but it can be detrimental in specific immune responses, such as sepsis and antitumor immunity. Recently, immature myeloid cells have been implicated in the suppression of immune responses in mouse models of cancer, infectious disease, bone marrow transplantation, and autoimmune disease. Here, we report the identification of a subset of mature human neutrophils (CD11cbright/CD62Ldim/CD11bbright/CD16bright) as what we believe to be a unique circulating population of myeloid cells, capable of suppressing human T cell proliferation. These cells were observed in humans in vivo during acute systemic inflammation induced by endotoxin challenge or by severe injury. Local release of hydrogen peroxide from the neutrophils into the immunological synapse between the neutrophils and T cells mediated the suppression of T cell proliferation and required neutrophil expression of the integrin Mac-1 (αMβ2). Our data demonstrate that suppression of T cell function can be accomplished by a subset of human neutrophils that can be systemically induced in response to acute inflammation. Identification of the pivotal role of neutrophil Mac-1 and ROS in this process provides a potential target for modulating immune responses in humans.

Figures

Figure 1. Neutrophil subsets after LPS administration.

(A) Neutrophils were stained for CD16 and CD62L before and 180 minutes after LPS administration. Before LPS administration, neutrophils formed one population; the CD16low cells are eosinophils. At 180 minutes after LPS administration, 2 distinct neutrophil subsets were identified. Subsets were FACS sorted, and cytospins were made and stained with May-Grunwald Giemsa. Original magnification, ×100. (B) Neutrophils were stained for CD16 and CD62L at different time points; time point 0 is before LPS challenge. Percentages of neutrophil subsets were calculated from flow cytometry data. Data are expressed as mean ± SEM; n = 7. (C) Absolute counts of neutrophil subsets were calculated using the percentage of the subsets and the absolute neutrophil counts (mean ± SEM; n = 7).

Figure 2. Neutrophil phenotype after LPS administration (A) Neutrophils were stained for CD16 and CD62L to discriminate between the subsets.

Additionally, they were stained with CD11b, CD11c, CD54, and CD88. Mean fluorescence intensity is depicted. Red lines depict CD16bright/CD62Ldim; gray lines depict CD16bright/CD62Lbright; green lines depict CD16dim/CD62Lbright. Graphs are representative of 5 experiments. (B) Neutrophil survival after 24 hours. Neutrophils subsets were FACS sorted, and cells were incubated for 24 hours at 37°C, 5% CO2; after incubation, they were stained with annexin V PE for 15 minutes and measured by flow cytometry. Data are expressed as the percentage of annexin V–negative, living cells (mean ± SEM; n = 5). (C) Relative increase in H2O2 in unsorted neutrophils measured by flow cytometry with DHR. Neutrophil subsets are visualized with CD16 and CD62L staining. Cells were stimulated with fMLF (10–6 M) and PAF (10–6 M) for 15 minutes. Neutrophils from healthy controls were used as a control. Graph shows relative increase in H2O2 release (mean ± SEM; n = 7). **P < 0.005; ***P < 0.001.

Figure 3. T cell proliferation after incubation with neutrophil subsets.

Blood was drawn 180 minutes after LPS administration, neutrophils were stained for CD16 and CD62L, and subsets were sorted. PBMCs were isolated from blood drawn before LPS administration and isolated by ficoll gradient separation. T cell proliferation was measured by [3H]thymidine incorporation. (A) PBMCs were stimulated with PHA (10 μg/ml) and incubated with different concentrations of the neutrophil subsets for 4 days. Data are depicted as the percentage inhibition of proliferation (mean ± SEM; n = 7). (B) PBMCs were stimulated with CD3 at 0.15 μg/m/CD28 at 1 μg/ml and incubated with different concentrations of the neutrophil subsets for 4 days. Data are presented as mean ± SEM; n = 7. (C) PBMCs from healthy volunteers who had received a tetanus booster vaccination less than 5 years ago were stimulated with tetanus toxoid, incubated with CD62Ldim or CD16dim neutrophils, and added in a 2:1 neutrophil-to-lymphocyte ratio for 4 days. Data are depicted as the percentage inhibition of proliferation (mean ± SEM; n = 10). (D) T cell apoptosis in neutrophil cocultures. Neutrophils were added in a 2:1 ratio in culture conditions described for the T cell proliferation cultures. At various time points, cells were stained with CD3 FITC and annexin V PE, and apoptosis was measured by gating the CD3-positive cells by flow cytometry. Data are depicted as the percentage of living cells (mean ± SEM; n = 4). *P < 0.05; ***P < 0.001.

Figure 4. Lymphocyte phenotype after neutrophil coculture.

PBMCs were incubated with or without CD62Ldim and CD16dim neutrophils added in a 2:1 ratio. The cells were stimulated with PHA. (A and B) After 2 days of coculture, cells were stained for flow cytometry, and the percentages of (A) CD4-positive and (B) CD8-positive T cells was determined. Data are presented as mean ± SEM; n = 3. (C and D) After 4 days, (C) IFN-γ and (D) IL-13 production was measured in the supernatant of the coculture using a sandwich ELISA. Data are presented as relative production of IFN-γ or IL-13 compared with that of PBMCs only (mean ± SEM; n = 6). (E–G) The percentage IFN-γ– and IL-4–producing T cells was measured by intracellular cytokine staining. PBMCs were stained for CD4/CD8 and intracellular IFN-γ and IL-4 after 2 days of coculture Depicted are (E) the percentage IFN-γ– or (F) IL-4–producing lymphocytes. Data are presented as mean ± SEM; n = 4. An example of the staining is shown in G. Numbers indicate the percentages of cells in each gate. *P < 0.05.

Figure 5. Mechanism of neutrophil-induced suppression of T cell proliferation.

(A) Soluble mediators of T cell inhibition. CD62Ldim neutrophils were added in a 2:1 ratio to PHA-stimulated (10 μg/ml) PBMCs in the presence or absence of anti–IL-10 (5 μg/ml), TGF-β inhibitor LAP (1 μg/ml), or l-arginine (1 mM). Data are presented as mean ± SEM of 5 independent experiments. (B) Separation of CD62Ldim neutrophils and PBMCs. Neutrophils and T cells were separated by a membrane using a Thinsert system (MS). Neutrophils were placed in the upper compartment in a 2:1 ratio, and PBMCs were placed in the lower. These data were compared with nonseparated neutrophils and PBMCs (–). Data are expressed as mean ± SEM of 4 individual experiments. (C) CD62Ldim neutrophils were added in a 2:1 ratio to PHA- stimulated (10 μg/ml) PBMCs in the presence or absence of Mac-1–blocking antibody 44a (10 μg/ml) and/or catalase (4,000 U/ml). As a control, 44a and catalase were also added to PBMCs without adding neutrophils; data were corrected for this control. Data are presented as mean ± SEM of 7 independent experiments. (D) Cocultures contained PBMCs with or without FACS-sorted neutrophils subsets or neutrophils from before LPS (t = 0). Cocultures were stimulated with PHA (10 μg/ml) and were incubated in culture medium containing Amplex Red (25 μM) and HRP (0.25 U/ml). Fluorescence after reduction of Amplex Red was measured over a period of 8 hours. As a positive control, PAF (10–6 M) and fMLF (10–6 M) were added to the culture. One representative experiment of 3 is depicted. *P < 0.05.

Figure 6. Visualization of neutrophil-lymphocyte interactions.

CD62Ldim-sorted neutrophils stained with CD16 FITC were added in a 2:1 ratio to unlabeled PBMCs and stimulated with PHA (10 μg/ml). Cells were incubated in culture medium containing Amplex Red (50 μM) and HRP (0.5 U/ml). Images were acquired using a Zeiss LSM510 Meta microscope. (A) Neutrophil-lymphocyte interaction with a localized Amplex Red signal indicative for local production of H2O2. (B) Same interaction as in A, showing that the Amplex Red signal colocalized with the CD16-labeled neutrophil membrane. The green line represents CD16; the red line represents H2O2. (C) Time-lapse image of a neutrophil-lymphocyte interaction. See Supplemental Video 1 for the full video. Representative examples are shown of 4 independent experiments. Original magnification, ×40 (A–C). (D) Quantification of local production of H2O2 spots in time-lapse images. Time-lapse images were made by deconvolution microscopy (Applied Precision DeltaVision Core Imaging System; the camera used was a Cascade EMCCD). CD62Ldim- and CD16dim-sorted neutrophils stained with calcein were added in a 2:1 ratio to unlabeled lymphocytes and stimulated with PHA (10 μg/ml). Cells were incubated in culture medium containing Amplex UltraRed (50 μM) and HRP (0.5 U/ml). In one of the conditions, Mac-1–blocking antibody 44a was added. The number of H2O2-positive spots were counted. Data are presented as mean ± SEM of 3 independent experiments. *P < 0.05.

Figure 7. Suppression of T cell proliferation in severely injured patients.

(A) Blood was drawn from severely injured patients immediately after admission in the hospital. Total leukocytes were stained for CD16 and CD62L. A CD62Ldim subset of neutrophils was seen similar to the subset found 3 hours after LPS. Numbers indicate the percentages of cells in each gate. (B) Total leukocytes from severely injured patients were stimulated with PHA (10 μg/ml) and incubated in the presence or absence (–) of blocking antibodies to Mac-1 (44a) or CD11c (clone 3.9) for 5 days. After 5 days of incubation, total leukocytes were counted. Groups were compared using a paired samples t test. Data are presented as mean ± SEM of 4 independent experiments. (C) IFN-γ concentration was measured by ELISA in the supernatants of the 5-day cultures. Data are depicted as relative increase compared with that of PHA only. Groups were compared using a paired samples t test. Data are presented as mean ± SEM of 5 independent experiments. *P < 0.05.