Research ArticleASTHMA

Neutrophils restrain allergic airway inflammation by limiting ILC2 function and monocyte–dendritic cell antigen presentation

See allHide authors and affiliations

Science Immunology  08 Nov 2019:
Vol. 4, Issue 41, eaax7006
DOI: 10.1126/sciimmunol.aax7006

Neutrophils keep the peace

Allergic airway inflammation is a complex disease, and multiple immune and nonimmune contribute to development and progression. Here, using a house dust mite–induced mouse model of allergic inflammation, Patel et al. report that depletion of neutrophils worsens airway inflammation. Neutrophil depletion, as expected, led to enhanced myelopoiesis that was driven by granulocyte colony-stimulating factor (G-CSF). By carrying out experiments to understand how neutrophil depletion and systemic increase in G-CSF levels drive airway inflammation, the authors have uncovered a previously unsuspected link between G-CSF and type 2 innate lymphoid cells (ILC2s). They report that G-CSF acts directly on both human and mouse ILC2s to promote production of TH2 cytokines interleukin-5 (IL-5) and IL-13.

Abstract

Neutrophil mobilization, recruitment, and clearance must be tightly regulated as overexuberant neutrophilic inflammation is implicated in the pathology of chronic diseases, including asthma. Efforts to target neutrophils therapeutically have failed to consider their pleiotropic functions and the implications of disrupting fundamental regulatory pathways that govern their turnover during homeostasis and inflammation. Using the house dust mite (HDM) model of allergic airway disease, we demonstrate that neutrophil depletion unexpectedly resulted in exacerbated T helper 2 (TH2) inflammation, epithelial remodeling, and airway resistance. Mechanistically, this was attributable to a marked increase in systemic granulocyte colony-stimulating factor (G-CSF) concentrations, which are ordinarily negatively regulated in the periphery by transmigrated lung neutrophils. Intriguingly, we found that increased G-CSF augmented allergic sensitization in HDM-exposed animals by directly acting on airway type 2 innate lymphoid cells (ILC2s) to elicit cytokine production. Moreover, increased systemic G-CSF promoted expansion of bone marrow monocyte progenitor populations, which resulted in enhanced antigen presentation by an augmented peripheral monocyte-derived dendritic cell pool. By modeling the effects of neutrophil depletion, our studies have uncovered previously unappreciated roles for G-CSF in modulating ILC2 function and antigen presentation. More broadly, they highlight an unexpected regulatory role for neutrophils in limiting TH2 allergic airway inflammation.

INTRODUCTION

Neutrophils are essential components of the body’s immune surveillance and host defense, owing to their capacity to readily eliminate invading pathogens. However, because of their considerable destructive capacity and potential to cause damage to healthy tissue, it is critical that neutrophil homeostasis is tightly regulated. Neutrophil homeostasis is maintained by a fine balance between granulopoiesis, bone marrow storage and release, intravascular margination, and ensuing clearance and destruction. The principal regulator of granulopoiesis at steady state is granulocyte colony-stimulating factor (G-CSF)/CSF3, with mice lacking the G-CSF receptor shown to be severely neutropenic (14). After stress or an inflammatory insult, G-CSF and an array of other mediators, such as ELR+ CXC chemokines, are increased and function to further promote granulopoiesis and neutrophil recruitment to a tissue (5, 6). However, it is critical that this neutrophilic inflammation is tightly regulated and efficiently resolved and a homeostatic state is restored. Increasingly, there is a growing comprehension for the prominent role neutrophils play in regulating their own turnover both during homeostasis and inflammation (7, 8).

Overexuberant and persistent neutrophilic responses have been implicated in the pathology of an array of chronic diseases, including chronic obstructive pulmonary disease, cystic fibrosis, and asthma. In the context of asthma, elevated neutrophil numbers are associated with enhanced severity of disease, impaired lung function, diminished responsiveness to corticosteroids, exacerbations, and fatality (814). However, manipulation of neutrophilic inflammation in a clinical setting has been disappointing and failed to ameliorate disease pathology (1521). One potential explanation for this is that strategies seeking to reduce neutrophilic inflammation may inadvertently disrupt regulatory functions performed by neutrophils.

In this study, we demonstrate that chronic, systemic depletion of neutrophils in a house dust mite (HDM) murine model of allergic airway disease unexpectedly resulted in exacerbated T helper 2 (TH2) inflammation, augmented mucus production, and increased airway resistance. Central to this augmented inflammation in neutrophil-depleted animals was a marked increase in G-CSF concentrations, which arose due to a failure of apoptotic neutrophils to trigger a negative feedback interleukin-23 (IL-23)–IL-17–G-CSF regulatory axis in the periphery, classically designed to limit neutrophil production and mobilization from the bone marrow. In the context of our allergic airway disease model, the accumulated G-CSF directly potentiated allergen sensitization at early time points by promoting TH2 cytokine production by type 2 innate lymphoid cells (ILC2s) and acting on bone marrow progenitors to drive monocytosis. This monocytosis consequently resulted in augmented antigen presentation by monocyte-derived dendritic cells (moDCs). Thus, we highlight unappreciated roles for G-CSF in promoting ILC2 function and antigen presentation and more broadly for neutrophils in negatively regulating type 2 allergic airway inflammation.

RESULTS

HDM administration elicits a neutrophilic inflammation in the lung and airways of mice

Our well-established HDM model of allergic airway disease (fig. S1A) (22) provokes a mixed granulocytic inflammation with prominent neutrophilia. In this model, a robust increase in neutrophil numbers and percentages in the lung (fig. S1, B and C, respectively) and airways (fig. S1, D and E, respectively) were observed from 1 week of HDM exposure and persisted until 3 weeks, by which time eosinophils were the prominent granulocyte (fig. S1, B to E). The elevated neutrophilic inflammation observed in HDM-treated animals coincided with increased expression levels and protein concentrations of classical ELR+ CXC chemokine CXCL1/KC (fig. S1, F to H) and neutrophil granulopoiesis regulator G-CSF (fig. S1, I to K). Neutrophil-derived proteases myeloperoxidase (MPO; fig. S2, A and B) and matrix metalloproteinase-9 (MMP-9; fig. S2, C and D) are often used as a clinical surrogate for neutrophilic inflammation, and levels were found to be elevated in the lung and airways of HDM-exposed mice. Whereas MPO levels temporally correlated with neutrophilic infiltrate, MMP-9 levels in the airways continued to increase when neutrophilia had stabilized, likely reflective of the multitude of cellular sources of this protease.

Neutrophil depletion exacerbates TH2 inflammation and airway remodeling in HDM-exposed mice

To interrogate the role of neutrophils in our model of allergic airway disease, mice were administered an anti-Ly6G (1A8) neutrophil-depleting antibody (23). Differential 1A8 dosing regimens were used to assess the optimal strategy to deplete neutrophils in mice exposed to HDM (fig. S3A), and a previously reported (24) flow cytometry gating protocol was used and modified to identify neutrophils independently of their Ly6G expression (fig. S3B), with neutrophils defined as CD11b+ CD11clo, F480, and Ly6Cintermediate. A single intraperitoneal administration of 100 μg of 1A8 was sufficient to completely ablate neutrophil numbers in the lung (fig. S3, C and E), airways (fig. S3, D and E), blood (fig. S3F), and spleen (fig. S3G) 24 hours after a single HDM exposure, with neutrophil numbers in the bone marrow (fig. S3H) also partially reduced. When this dosing regimen was used throughout the 3-week duration of the HDM allergic airway disease model (Fig. 1A and fig. S3I), a consistent reduction in neutrophil numbers was observed at all time points in the lung (Fig. 1B), airways (Fig. 1C), and blood (fig. S3J) relative to mice treated with an immunoglobulin G2a (IgG2a) isotype control antibody (2A3). However, depletion of bone marrow neutrophils was not apparent at later time points (fig. S3K). The anti-Ly6G 1A8 antibody has become the preferential strategy to deplete neutrophils (23), owing to its purported selective expression by these cells (25, 26). The neutrophil specificity of Ly6G expression is further supported by the Catchup Ly6G reporter mice, although low transgene activity was reported in a small number of eosinophils within this study (27). In our hands, Ly6G expression was largely absent on lymphocytes and monocytes, but low levels were detectable on a subset of eosinophils in the bone marrow, blood, and lungs of phosphate-buffered saline (PBS)– and HDM-treated mice (fig. S4, A to C). However, 1A8 administration selectively depleted neutrophils in HDM-treated mice, with no effect on numbers of eosinophils and lymphocyte numbers, whereas monocytes were unexpectedly increased in HDM/1A8-treated animals (fig. S4, D to F). A marginal reduction in lung eosinophil numbers was, however, observed in PBS mice administered 1A8 (fig. S4F).

Fig. 1 Neutrophil-depleted mice display augmented type 2 inflammation after 3 weeks of HDM exposure.

(A) Balb/c mice were administered HDM or PBS intranasally three times per week for up to 3 weeks, and at 24 hours before each HDM/PBS administration, mice were intraperitoneally dosed with either 100 μg of neutrophil-depleting antibody, 1A8, or isotype control antibody, 2A3. At 24-hour, 1-week, and 3-week time points (in each instance 24 hours after the final HDM/PBS exposure), lung tissue and BALF were collected (Ŧ). Total numbers of neutrophils in the lungs (B) and airways (C) were determined by flow cytometry. (D) Total cell numbers in the lung were assessed after 3 weeks of HDM exposure by trypan blue exclusion. (E) Representative H&E-stained lung sections from mice exposed to PBS or HDM for 3 weeks and treated with 2A3 or 1A8. (F) The number of eosinophils in the lung was quantified by flow cytometry at 3 weeks. The number of CD4+ T cells expressing T1ST2 (G) or IL-13 (H) in the lung was assessed by flow cytometry after 3 weeks of PBS/HDM exposure. (I) After 3 weeks of HDM/PBS exposure, concentrations of HDM-specific IgE and IgG1 in the serum were determined by ELISA. Figures present combined data from two independent experiments with four to six mice per group in each experiment. Results depicted as means ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 using Mann-Whitney statistical test.

The hallmark clinical features of allergic airway disease were established within our model by 3 weeks of HDM exposure (22). At this time point, despite the substantial reduction in neutrophil numbers in 1A8-treated, HDM-exposed mice, these animals actually exhibited an increase in total cell numbers in their lungs (Fig. 1D), whereas total numbers in the airways were modestly elevated, albeit not significantly relative to control animals (fig. S5A). The increase in pulmonary inflammation in HDM-exposed neutrophil-depleted mice was visible by observation of hematoxylin and eosin (H&E)–stained lung sections (Fig. 1E). We next sought to determine the nature of the populations that must be elevated in HDM-treated neutrophil-depleted mice to account for the increase in total inflammation despite the loss of neutrophils. HDM-exposed neutrophil-depleted mice displayed increased eosinophils in their lungs (Fig. 1F) and airways (fig. S5B) relative to control treated mice administered HDM, although this again failed to reach significance in the airways. Furthermore, the number of CD4+ TH2 cells in both the lungs (Fig. 1, G and H) and airways (fig. S5C) was significantly elevated in neutrophil-depleted mice relative to control animals after 3 weeks of HDM administration. In keeping with greater TH2 inflammation in neutrophil-depleted animals after 3 weeks of HDM exposure, a significant increase in classical type 2 antibodies was shown, with serum levels of both total (fig. S5D) and HDM-specific (Fig. 1I) IgE and IgG1 being elevated compared to 2A3 control animals. Observation of H&E-stained lung sections (Fig. 1E) revealed that many of the airways of HDM-exposed neutrophil-depleted mice were plugged with mucus. Accordingly, 1A8-treated, HDM-exposed mice exhibited increased lung tissue expression of the major airway mucin Muc5ac (fig. S5E), with an associated increase in airway MUC5AC protein (fig. S5F) relative to 2A3/HDM animals. IL-13 is a major instigator of epithelial MUC5AC production within our model of allergic airway disease, and accordingly, IL-13 concentrations were elevated in the bronchoalveolar lavage fluid (BALF) of neutrophil-depleted mice exposed to HDM (fig. S5G), in keeping with the augmented TH2 response within this group. Furthermore, BALF IL-13 concentrations showed a significant correlation with MUC5AC protein levels across HDM groups (fig. S5H). In keeping with airway obstruction and plugging by mucus, the neutrophil-depleted HDM-treated animals exhibited an increase in baseline airway resistance relative to 2A3/HDM controls (fig. S5I), with airway resistance correlating with BALF MUC5AC concentrations (fig. S5J). Changes in lung function of HDM-exposed mice after neutrophil depletion were, however, restricted to baseline airway resistance, with no differences detectable in airway hyperresponsiveness to increasing doses of methacholine relative to 2A3/HDM mice (fig. S6).

An early increase in ILC2-derived TH2 cytokines supports the augmented type 2 inflammation observed in neutrophil-depleted mice administered HDM

Given the prominent effect of neutrophil depletion on antigen-specific T cell responses and type 2 antibody generation, we questioned the potential role of neutrophils in regulating allergen sensitization at early time points after HDM exposure when neutrophil numbers were at their peak (Fig. 2A). TH2 cytokines are of fundamental importance in driving the type 2 inflammation and associated antibody responses in allergen models of allergic airway disease (28). Accordingly, levels of prototypic TH2 cytokines IL-4, IL-5, and IL-13 were significantly increased in the airways of neutrophil-depleted mice relative to controls after 1 week of HDM exposure, in terms of both protein levels in BALF (Fig. 2B) and Il4, Il5, and Il13 messages in cells derived from the airways (fig. S7A). This phenotype was conserved in lung tissue, albeit less pronounced, at the level of protein (fig. S7B) and transcript (fig. S7C). We next sought to ascertain the cellular source of the elevated TH2 cytokines in neutrophil-depleted animals at 1 week of HDM exposure. Lung epithelial (CD45 EpCAM+), endothelial (CD45 EpCAM CD31+), and leukocytes (CD45+) were isolated via fluorescence-activated cell sorting (FACS) from mice administered PBS or HDM for 1 week (fig. S8A) to assess the gene expression of TH2 cytokines. Il4, Il5, and Il13 messages were only detected in the leukocyte population and were increased upon HDM challenge (fig. S8B). ILC2s and CD4+ TH2 cells are classically acknowledged as primary sources of TH2 cytokines in allergen models of allergic airway disease (28), and assessment of IL-13 gfp-reporter mice (29) administered HDM for 1 week demonstrated that ILC2s were the prominent source of this cytokine at this early time point (fig. S8C).

Fig. 2 Augmented TH2 cytokine production by ILC2s in neutrophil-depleted mice administered HDM for 1 week.

(A) Balb/c mice were intranasally administered HDM or PBS three times per week for 1 week, and at 24 hours before each HDM treatment, mice were intraperitoneally dosed with either 100 μg of neutrophil-depleting antibody, 1A8, or isotype control antibody, 2A3. At 24 hours after the final HDM administration, BALF was collected (Ŧ). (B) Concentrations of IL-4, IL-5, and IL-13 protein were assessed using BALF by ELISA. In some experiments, CD4+ T cells and ILC2s (pooled from three mice per data point) were isolated from the airways by FACS, at 24 hours after the final HDM exposure, for subsequent mRNA gene expression analysis. (C) Relative expression of Il-4, Il-5, Il-13, and Gata3 in T cells and ILC2s derived from BAL of HDM-treated mice, as determined by qPCR. At the same time point, the number of IL-13+ ILC2s and IL-5+ ILC2s (D) and geometric expression of IL-13 and IL-5 in ILC2s (E) in the BAL were assessed by flow cytometry. Figures present combined data from two independent experiments with four to six mice per group in each experiment (B, D, and E) or from one experiment whereby each data point represents cells pooled from three independent mice (C). Results depicted as means ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 using Mann-Whitney statistical test.

Subsequently, ILC2s and CD4+ T cells were isolated by FACS (fig. S9) from the airways (Fig. 2C) and lungs (fig. S10) of control and neutrophil-depleted mice challenged with HDM for 1 week and were assessed for expression of genes encoding TH2 cytokines. At this early time point, CD4+ T cells were the prominent source of IL-4, and expression was significantly elevated in cells derived from the lungs of neutrophil-depleted animals (fig. S10). Conversely, ILC2s were the primary source of IL-5 and IL-13 (Fig. 2C and fig. S10), and their levels were markedly elevated in airway ILC2s derived from neutrophil-depleted animals (Fig. 2C), in keeping with the prominent increase in levels of these cytokines in BALF of these mice. Supportive of these findings, the master transcriptional regulator of TH2 cytokine production, GATA-3, showed elevated mRNA expression in lung CD4+ T cells (fig. S10) and airway ILC2s (Fig. 2C) derived from neutrophil-depleted mice. Intracellular cytokine staining and flow cytometry analysis supported these assertions because, whereas the total number of lung and airway ILC2s (fig. S11A) and the proportion (fig. S11, B and C) and number (Fig. 2D and fig. S11, D to F) of ILC2-producing IL-13 or IL-5 were not consistently elevated in neutrophil-depleted animals administered HDM, the amount of cytokines the ILC2s produced on a per-cell basis (as adjudged by assessment of geometric mean) was significantly elevated (Fig. 2E and fig. S11, D to F).

Elevated moDCs and ensuing antigen presentation in HDM-exposed neutrophil-depleted mice as a consequence of perturbations in bone marrow progenitor pools

In addition to a TH2 milieu, antigen sampling and presentation by dendritic cells (DCs) is central to allergen sensitization and establishment of a type 2 adaptive response in models of allergic airway disease. Concomitant with an increase in ILC2-derived TH2 cytokines in neutrophil-depleted mice administered HDM for 1 week was a significant increase in lung Ly6Clow and Ly6Chigh monocytes (Fig. 3A). Although this increase was most pronounced in neutrophil-depleted mice given HDM, it was also apparent in neutrophil-depleted PBS-treated animals. This increase in monocyte populations in the lungs of HDM-administered neutrophil-depleted animals was apparent in precision-cut lung slices (PCLSs) stained for monocyte marker CD115 (fig. S12A). Subsequently, neutrophil-depleted animals also presented with an increase in lung moDCs that was again most pronounced in those animals administered HDM for 1 week (Fig. 3B). Unexpectedly, this phenotype was not restricted to moDCs, with a significant increase in lung CD11b+ conventional DCs (cDCs) and CD103+ cDCs (fig. S12B) also observed in neutrophil-depleted mice.

Fig. 3 HDM-exposed neutrophil-depleted mice exhibit augmented monocytosis, DC numbers, and antigen presentation.

Balb/c mice were intranasally administered HDM or PBS three times per week for 1 week. At 24 hours before each HDM/PBS administration, mice were intraperitoneally treated with either 100 μg of neutrophil-depleting antibody, 1A8, or isotype control antibody, 2A3. At 24 hours after the final HDM/PBS exposure, lung tissue was collected. (A) Lung monocyte subsets identified as Ly6Clow and Ly6Chigh were enumerated by flow cytometry. (B) Numbers of lung moDCs were also determined by flow cytometry. The number of moDCs (C) and CD86+ moDCs (D) in the MLNs was enumerated by flow cytometry. In some experiments, the final dose of HDM was admixed with 100 μg of Alexa Fluor 488–labeled OVA (E), and MLNs were collected (Ŧ) at 24 hours after the final HDM/PBS exposure. (F) The number of OVA+ moDCs within the MLNs was enumerated by flow cytometry. Figures present data combined from two independent experiments with four to six mice per group for each experiment (A to D) or from one experiment with five mice per group (F). Results depicted as means ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 using Mann-Whitney statistical test.

Activation and migration of DCs to draining lymph nodes and ensuing priming of allergen-specific T cell responses are integral to allergen sensitization. In the HDM model, moDCs and CD11b+ cDCs are integral to transporting antigen to draining mediastinal lymph nodes (MLNs) (28, 30). Numbers of total and activated moDCs (Fig. 3, C and D) and CD11b+ cDCs (fig. S12, C and D) were increased in the MLNs of neutrophil-depleted animals administered HDM. To ascertain whether antigen presentation was augmented in neutrophil-depleted animals exposed to HDM, 2A3- and 1A8-treated mice were administered HDM for 1 week, with Alexa Fluor 488–tagged ovalbumin (OVA) administered concomitantly with the final dose of allergen (Fig. 3E). Numbers of OVA+ moDCs in neutrophil-depleted mice were also significantly elevated in MLNs (Fig. 3F), demonstrating that they were effectively transporting antigen to localized lymph nodes. Although the same trend was observed with OVA+ CD11b+ cDCs (fig. S12E), the increase in neutrophil-depleted animals did not reach statistical significance. Supportive of the assertion of increased antigen presentation in HDM-treated neutrophil-depleted mice, MLN T cells from these animals expressed higher levels of activation markers ICOS (fig. S12F) and PD1 (fig. S12G) and showed greater proliferation (fig. S12H). Thus, enhanced antigen presentation in the context of an augmented TH2 cytokine environment observed in neutrophil-depleted animals during allergen sensitization at 1 week of HDM exposure is conducive to the elevated TH2 inflammation seen at later time points.

To rationalize the accumulation of monocyte populations and ensuing moDCs in the lungs of neutrophil-depleted mice, we assessed lung concentrations of classical monocyte chemokines CCL2 and CX3CL1 after 1 week of HDM exposure. Although HDM administration resulted in an increase in concentrations of CCL2 and CX3CL1, these chemokines were not further elevated in mice depleted of neutrophils and could not therefore account for the tissue monocytosis observed in these animals (fig. S13A). Analysis of PCLS revealed increased intravascular pools of monocytes in neutrophil-depleted HDM-exposed animals (fig. S12A). Accordingly, flow cytometry analysis demonstrated that neutrophil depletion resulted in increased numbers of blood Ly6Clow and Ly6Chigh monocytes—in both PBS- and HDM-treated animals (Fig. 4A)—thus providing a greater circulating pool of monocytes to extravasate into the lung in response to localized chemokine gradients. Subsequently, we questioned whether alteration in bone marrow progenitor pools could account for systemic changes in monocyte numbers after neutrophil depletion. Accordingly, the percentage of monocyte–dendritic cell progenitors (MDPs) (31, 32), defined as CD115+, Lineage, c-Kit+, and FLT-3+ (fig. S13B), was shown to be elevated in the bone marrow of neutrophil-depleted mice after 1 week of HDM exposure (Fig. 4B). Recently, it has been suggested that MDPs differentiate into common–dendritic cell progenitor cells (CDPs) (32), which are the first committed DC progenitor to give rise to the cDC subsets that were also shown to be universally elevated in neutrophil-depleted mice. Consequently, the percentage of CDPs, defined as CD115+, Lineage, c-Kit, FLT-3+, CD11b, and CD11c (fig. S13B), was also shown to be elevated in neutrophil-depleted animals (Fig. 4B). Thus, neutrophil depletion gives rise to increased MDPs and CDPs in the bone marrow, which ultimately drives elevated monocyte and DC populations in the lung.

Fig. 4 Neutrophil-depleted mice display increased numbers of monocyte and DC progenitors within their bone marrow owing to a dysregulated IL-23–IL-17–G-CSF axis.

Balb/c mice were intranasally administered HDM or PBS three times per week for 1 week. At 24 hours before each HDM/PBS administration, mice were intraperitoneally treated with either 100 μg of neutrophil-depleting antibody, 1A8, or isotype control antibody, 2A3. At 24 hours after the final HDM/PBS exposure, BALF, lung tissue, blood, and bone marrow were collected. (A) The number of Ly6Clow and Ly6Chigh monocytes in the blood was quantified by flow cytometry. (B) The percentage of MDPs and CDPs within the bone marrow was assessed by flow cytometry. (C) The concentration of G-CSF in the serum, lung homogenate, and BALF was determined by ELISA. Expression of Csf3 (G-CSF gene; D), Il17 (E), and Il23 (F) was assessed in whole lung by qPCR. (G) Schematic depicting the negative feedback pathway by which tissue neutrophils limit granulopoiesis by modulation of the IL-23–IL-17–G-CSF axis. Figures present data from two independent experiments with four to six mice per group in each experiment. Results depicted as means ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 using Mann-Whitney statistical test.

A dysregulated IL-23–IL-17–G-CSF axis in neutrophil-depleted mice

We next sought to ascertain what mediator(s) was driving the augmented progenitor pools in the bone marrow of neutrophil-depleted animals and the ensuing increase in lung monocytes and DCs. A proteome profiler array was used to semiquantitatively assess broad changes in cytokines, chemokines, and proteases in serum of control and neutrophil-depleted mice after 1 week of HDM administration. Although differential levels of various mediators were observed via proteome profiler analysis, a marked increase in G-CSF in neutrophil-depleted animals was particularly noteworthy (fig. S14). This was deemed to be especially pertinent given that CSF3R, the G-CSF receptor, is expressed on MDPs and CDPs (33) and that an increase in monocytes has been reported in neutropenic patients administered G-CSF to promote granulopoiesis (34). Subsequently, serum, lung homogenate, and BALF G-CSF levels were assessed by enzyme-linked immunosorbent assay (ELISA) and demonstrated to be markedly elevated in both PBS- and HDM-treated mice depleted of neutrophils (Fig. 4C). The increase in G-CSF concentration did not just reflect a reduced internalization of the G-CSF after ablation of neutrophils, because lung Csf3 gene expression was elevated in neutrophil-depleted mice (Fig. 4D).

Previous studies have demonstrated that G-CSF expression is prominently controlled by IL-17 (3537), with IL-17 derived from tissue-resident T cell populations central to G-CSF regulation during homeostasis (38, 39). It was noteworthy therefore that the proteome profiler revealed an increase in IL-17 levels in HDM-exposed neutrophil-depleted animals (fig. S14). Accordingly, it was demonstrated that Il17a expression was significantly elevated in the lungs of neutrophil-depleted mice, with the increase potentiated by HDM exposure (Fig. 4E). Expression analysis in T cells and ILC2s isolated from the lungs of HDM-treated animals at this time point demonstrated that T cells were the primary source of IL-17 and that those derived from neutrophil-depleted mice produced more of this cytokine (fig. S15A). This was supported by flow cytometry, with both CD4+ αβ T cells and γδ T cells derived from neutrophil-depleted mice producing more IL-17 after 1 week of HDM administration (fig. S15, B and C). IL-23 is a potent regulator of IL-17 expression, and it has previously been demonstrated during homeostasis that phagocytosis of transmigrated, apoptotic neutrophils by resident macrophages and DCs suppresses their intrinsic IL-23 production (38, 39). We thus questioned whether the failure of neutrophils to reach the lung in HDM-challenged neutrophil-depleted mice would lead to an increase in IL-23 concentrations, which would rationalize an ensuing increase in IL-17 and ultimately G-CSF. Consequently, Il23 expression was shown to be elevated in the lungs of neutrophil-depleted animals administered HDM for 1 week (Fig. 4F). Flow cytometry analysis demonstrated that resident CD11c+ macrophages and moDCs were the primary sources of IL-23 in HDM-treated mice, with increased numbers of IL-23+ moDCs present in neutrophil-depleted animals (fig. S15D). Furthermore, both the CD11c+ macrophages and moDCs derived from neutrophil-depleted HDM-exposed mice produced more IL-23 on a per-cell basis than those from HDM/control antibody–treated animals (fig. S15, E and F). Thus, disruption of a peripheral feedback system, normally regulated by lung-infiltrating, apoptotic neutrophils, resulted in the increased G-CSF levels observed in our 1A8-treated mice (Fig. 4G).

Neutralization of G-CSF reduces the monocytosis and elevated TH2 cytokine levels observed in neutrophil-depleted HDM-exposed mice

We next sought to determine whether neutralization of G-CSF in our HDM model would abrogate the augmentation of bone marrow progenitors and tissue monocytes seen in neutrophil-depleted animals. Mice exposed to HDM for 1 week were concomitantly treated with 2A3/1A8 and anti–G-CSF neutralizing antibody/control (Fig. 5A). Neutralization of G-CSF resulted in a significant reduction in neutrophils in HDM-exposed animals (Fig. 5B), highlighting a prominent role for this mediator in defining neutrophilia in this model. The increase in lung Ly6Clow and Ly6Chigh (Fig. 5C) monocytes observed in HDM-treated mice depleted of neutrophils was completely negated after anti–G-CSF treatment. Furthermore, anti–G-CSF administration also reduced the percentage of bone marrow MDPs and CDPs (Fig. 5D) in neutrophil-depleted mice exposed to HDM for 1 week to a level observed in 2A3-treated animals. Thus, G-CSF, elevated as a consequence of neutrophil depletion, is driving the augmentation in bone marrow progenitors and ensuing increase in circulating and tissue monocytes. Reassuringly, the phenotype was largely recapitulated when IL-17 was neutralized in HDM-exposed neutrophil-depleted mice, validating the importance of this cytokine in defining G-CSF levels (fig. S16A) and consequential changes in tissue neutrophils (fig. S16B) and monocytes (fig. S16C) and bone marrow progenitors (fig. S16D).

Fig. 5 Neutralization of G-CSF abrogates the augmented monocytes, bone marrow progenitors, and TH2 cytokines observed in neutrophil-depleted HDM-exposed mice.

(A) Balb/c mice were intranasally administered HDM three times per week for 1 week. At 24 hours before each HDM administration, mice were treated with either 100 μg of neutrophil-depleting antibody, 1A8, or isotype control antibody, 2A3. To neutralize G-CSF, mice were also intraperitoneally administered 100 μg of anti–G-CSF or isotype control, 2A3, at 24 hours before each HDM dose. At 24 hours after the final HDM administration, BAL, lung tissue, and bone marrow were collected (Ŧ). The number of lung and airway neutrophils (B) and lung Ly6Clow and Ly6Chigh monocytes (C) was determined by flow cytometry. (D) The percentage of MDPs and CDPs within the bone marrow was assessed by flow cytometry. (E) The concentrations of BALF IL-4, IL-5, and IL-13 were assessed by ELISA. (F) Geometric mean of IL-13 by lung ILC2s was assessed by flow cytometry. Figures present data combined from two independent experiments with four to six mice per group in each experiment. Results depicted as means ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001 using Mann-Whitney statistical test.

Previously, we demonstrated that HDM-exposed neutrophil-depleted mice exhibited a prominent increase in TH2 cytokine levels at 1 week of HDM exposure, which was attributable to augmented production on a per-cell basis by ILC2s. Remarkably, the increase in BALF concentrations of IL-4, IL-5, and IL-13 observed in neutrophil-depleted mice after 1 week of HDM administration was significantly reduced upon anti–G-CSF administration (Fig. 5E). Similarly, the amount of TH2 cytokines produced by ILC2s on a per-cell basis, as exemplified by IL-13 and assessed by flow cytometry, was significantly reduced after anti–G-CSF administration to neutrophil-depleted animals exposed to HDM (Fig. 5F)—a phenotype that was again conserved with anti–IL-17 treatment (fig. S16E).

Recombinant G-CSF can augment ILC2 TH2 cytokine production

We next questioned whether the profound capacity of G-CSF to modulate ILC2 TH2 cytokine production and the expansion of bone marrow progenitor populations in HDM-exposed animals was unique to mice depleted of neutrophils or whether it could also operate comparably in fully immunocompetent animals. HDM-exposed mice were therefore administered recombinant G-CSF for 1 week (Fig. 6A). Administration of recombinant G-CSF to HDM-exposed mice further increased not only tissue neutrophils (Fig. 6B) but also augmented tissue monocytes (Fig. 6C) and bone marrow progenitors (Fig. 6D), as seen after neutrophil depletion. Furthermore, HDM-driven increases in BALF IL-4, IL-5, and IL-13 (Fig. 6E) and ILC2 TH2 cytokine production (Fig. 6F) were also further accentuated by recombinant G-CSF administration. Thus, G-CSF is a physiologically relevant regulator of ILC2 function and monocyte numbers.

Fig. 6 Coadministration of recombinant G-CSF augments numbers of HDM-induced neutrophils, monocytes, and bone marrow progenitors and ILC2 cytokine responses.

(A) Balb/c mice were intranasally administered PBS or HDM three times per week for 1 week. Mice were also intranasally administered 100 ng of recombinant G-CSF (in 50 μl of PBS) and intraperitoneally administered 2 μg of recombinant G-CSF (in 200 μl of PBS), or respective PBS controls, daily throughout the study. At 24 hours after the final HDM administration, BAL, lung tissue, and bone marrow were collected (Ŧ). The number of lung and airway neutrophils (B) and lung Ly6Clow and Ly6Chigh monocytes (C) was determined by flow cytometry. (D) The percentage of MDPs and CDPs within the bone marrow was assessed by flow cytometry. (E) The concentrations of BALF IL-4, IL-5, and IL-13 were assessed by ELISA. (F) Geometric mean of IL-13 by BAL ILC2s was assessed by flow cytometry. Figures present data from one experiment with four to six mice per group in each experiment. Results depicted as means ± SEM. *P < 0.05 and **P < 0.01 using Mann-Whitney statistical test.

We next questioned whether G-CSF could directly act on ILC2s to potentiate cytokine production. Data mining of previous RNA sequencing (RNA-seq) datasets indicated that ILC2s express the G-CSF receptor csf3r (40, 41), and this was confirmed in our HDM model with ILC2s isolated from the airways by FACS expressing Csf3r (Fig. 7A). Csf3r expression was completely absent in T cells isolated from airways (Fig. 7A), rationalizing earlier observations that the primary source of IL-5 and IL-13 in neutrophil-depleted animals was ILC2s and not CD4+ T cells. IL-33 is a potent regulator of ILC2s, and a week-long model of intranasal recombinant IL-33 administration (Fig. 7B) induced substantial ILC2 numbers in the absence of an antigen-specific TH2 cell response. Once again, ILC2s isolated from the airways of recombinant IL-33–treated mice expressed Csf3r, which was again absent from T cells derived from these animals (Fig. 7C). When recombinant G-CSF was coadministered with recombinant IL-33 into the airways of mice (Fig. 7B), it augmented Il5 and Il13 expression within isolated airway ILC2s but not T cells, as adjudged by quantitative polymerase chain reaction (qPCR) (Fig. 7D). Furthermore, coadministration of recombinant G-CSF with IL-33 increased BALF concentrations of IL-5 and IL-13 (Fig. 7E).

Fig. 7 G-CSF augments TH2 cytokine production from IL-33–expanded airway Csf3r-expressing ILC2s.

Balb/c mice were intranasally administered HDM three times per week for 1 week. At 24 hours after the final HDM administration, BAL was collected. (A) Relative expression of mRNA Csf3r in T cells and ILC2s isolated by FACS from the BAL of HDM-exposed mice, as determined by qPCR. (B) Balb/c mice were intranasally administered 1 μg of recombinant IL-33 three times per week for 1 week. At 24 hours after the final IL-33 administration, BAL was collected. (C) Relative expression of mRNA Csf3r in T cells and ILC2s isolated by FACS from the BAL of IL-33–exposed mice, as determined by qPCR. In some experiments, 100 ng of recombinant G-CSF was co-administered with IL-33 (B), and CD4+ T cells and ILC2s were isolated from the airways by FACS, at 24 hours after the final IL-33 administration, for subsequent mRNA gene expression analysis. (D) Relative expression of IL-4 and IL-13 in T cells and ILC2s derived from BAL, as determined by qPCR. (E) From the same experiments, BALF concentrations of IL-5 and IL-13 were determined by ELISA. Figures present data from one experiment with four to six mice per group in each experiment. Results depicted as means ± SEM. *P < 0.05 and **P < 0.01 using Mann-Whitney statistical test.

To validate that G-CSF was able to directly augment TH2 cytokine production from ILC2s, ILC2s were isolated from the airways of recombinant IL-33–treated mice (fig. S17A) or from peripheral blood of healthy donors (Fig. 8A) and then stimulated with or without G-CSF. As observed with mouse ILC2s, human ILC2s exhibited robust expression of G-CSF receptor, CSF3R (Fig. 8B). Mouse and human ILC2s stimulated with G-CSF released significantly more IL-5 and IL-13 than medium-treated control cells (fig. S17B and Fig. 8C, respectively), whereas levels of IL-4, IL-17, interferon-γ (IFN-γ), and IL-12 were undetectable and levels of IL-6 and tumor necrosis factor–α (TNF-α) were extremely low and comparable. Accordingly, IL5 and IL13 messages were significantly elevated in human ILC2s stimulated with G-CSF, as were levels of TH2 master transcriptional regulator GATA3 (Fig. 8D). Expression of IL4 and IL17 messages was undetectable. Similarly, Il4, Il5, and Gata3 transcripts were dose-dependently augmented in mouse ILC2s stimulated with G-CSF (fig. S17C). Thus, G-CSF is a previously unrecognized potentiator of TH2 cytokine production by ILC2s, which, together with the augmented G-CSF–driven moDC antigen presentation, facilitates the greater type 2 inflammation observed in HDM-exposed neutrophil-depleted mice at 3 weeks.

Fig. 8 G-CSF directly augments TH2 cytokine production from human ILC2s.

(A) Human ILC2s were isolated from peripheral blood and expanded in bulk culture with IL-7 (5 ng/ml) and IL-33 (15 ng/ml) before the addition of medium or medium containing recombinant G-CSF (100 ng/ml) for 72 hours. (B) Relative expression of CSF3R was assessed by qPCR after 72 hours. (C) The levels of IL-5 and IL-13 in the ILC2 supernatant were assessed by a multiplex cytokine assay and expressed as a donor-specific fold change after G-CSF treatment. (D) Expression of IL5, IL13, and GATA3 was assessed by qPCR after 72 hours and presented as a donor-specific fold change after G-CSF treatment. Results depicted as means ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001 using a paired t test.

DISCUSSION

An overexuberant or persistent neutrophilic response is implicated in the pathology of an array of inflammatory diseases, and neutrophils have consequently represented an attractive therapeutic target. However, therapeutic strategies that seek to ameliorate neutrophilic inflammation have failed to fully consider the potential regulatory roles fulfilled by these cells. In this study, we demonstrate that depletion of neutrophils in an HDM model of allergic airway disease led to perturbations of an IL-23–IL-17–G-CSF regulatory feedback pathway. The accumulated G-CSF subsequently promoted allergen sensitization and exacerbated type 2 inflammation, epithelial remodeling, and lung function. This was mediated by the capacity of G-CSF to promote TH2 cytokine production by airway ILC2s and to drive monocytosis and ensuing moDC-mediated antigen presentation (fig. S18).

We demonstrate that G-CSF acts on Csf3r+ ILC2s to potentiate their production of IL-5 and IL-13 on a per-cell basis. IL-4 levels were also elevated in vivo in a G-CSF–dependent manner but derived from Csf3r CD4+ T cells potentially attributable to the reported capacity of ILC2s to promote IL-4 production by CD4+ T cells (4244). ILC2s are early effectors in type 2 inflammation that sense and respond to cytokines and stress signals evoked from the proximal environment upon disruption of tissue homeostasis (45). It is, thus, rational that an innate mediator such as G-CSF can function to potentiate ILC2 function. We also reveal the capacity to G-CSF to drive the expansion of bone marrow MDPs and CDPs and an ensuing monocytosis. Rationalizing this effect, G-CSF has previously been demonstrated to drive the expansion of granulocyte macrophage progenitor compartments (GMPs) within the bone marrow (46). GMPs lie developmentally upstream of MDPs and CDPs and may thus account for the accumulation of the latter in neutrophil-depleted animals. Data mining of historic RNA-seq datasets (47) revealed that GMPs, MDPs, and CDPs all express csf3r, supportive of the notion that G-CSF is able to drive the proliferation of one or more of these progenitors. Although G-CSF is classically recognized as a potent regulator of granulopoiesis, G-CSF administration has been reported to stimulate monocyte production and release in neutropenic patients (4850). In our allergen-sensitized mice, these expanded monocyte populations gave rise to augmented moDC numbers, which, in keeping with previous literature (28, 30), transported antigen to draining MLNs and activated T cells—potentially further licensed by the elevated ILC2-derived IL-13 (42). Thus, enhanced antigen presentation, in the context of elevated ILC2-derived type 2 cytokines, in neutrophil-depleted mice facilitated the increase in CD4+ TH2 cells after 3 weeks of allergen exposure. Similarly, the elevated levels of IL-4 would function to promote B cell responses and class switching that underlie the augmented levels of HDM-specific IgE and IgG1.

Given the previously unidentified roles ascribed to G-CSF in this study, it is worth reevaluating the broader implications of G-CSF in the context of asthma. Increased levels of G-CSF have been reported in the BALF of asthmatic patients relative to healthy controls (51) and in serum of patients with unstable asthma (52). Pertinent to our current findings, G-CSF has been shown to increase concomitantly with IL-5 in unstable asthma (52), whereas treatment of people with G-CSF augmented the number of circulating DCs with the capacity to prime T cells to produce TH2 cytokines (53). Consequently, it is also appropriate to contextualize our findings with regard to the perceived role of neutrophils in asthma. The general consensus, based primarily on circumstantial clinical data, is that neutrophils are detrimental in the context of asthma and indicate a worse prognosis (8, 1014). Based on our current findings, it could be argued that neutrophils also represent a compensatory mechanism to indirectly restrain TH2 inflammation in the context of allergic disease by negatively regulating the bioavailability of G-CSF. In keeping with this hypothesis, adoptive transfer of neutrophils has recently been shown to attenuate TH2 responses in a mouse model of allergic airway disease (54). Our data in mice that neutrophils negatively regulate type 2 inflammation are supported by a previously reported case of a patient with cyclic neutropenia that suffered episodic acute asthma, whereby the cyclical depreciation in neutrophils concomitantly correlated with a substantial increase in eosinophils, serum IgE levels, and asthma exacerbation (55). In addition, eosinophilia is a well-recognized feature of a number of primary immunodeficiency disorders associated with neutropenia (56).

Our study demonstrates the complex regulatory feedback pathways that define neutrophil turnover and the intrinsic challenges of targeting neutrophils as a therapeutic modality in the context of asthma. Any intervention that affects the ability of neutrophils to transmigrate into tissues at steady state will likely modify G-CSF levels. Thus, mice deficient in CXCR2 (39) or specific adhesion molecules (38) display increased baseline levels of G-CSF. Accordingly, therapeutic strategies used to ameliorate neutrophilic inflammation in asthma may inadvertently augment G-CSF levels and potentiate TH2 inflammation. It is noteworthy, therefore, that the CXCR2 antagonist AZD5069 not only reduced peripheral blood neutrophils in healthy volunteers but also caused a significant increase in serum G-CSF concentrations (57). Similarly, AZD5069 reduced lung, sputum, and blood neutrophil numbers in patients with persistent asthma but again significantly increased G-CSF expression (57, 58). Furthermore, the CXCR2 antagonist SCH526123 reduced sputum neutrophils in patients with severe asthma but concomitantly increased the percentage sputum eosinophils (15). It is feasible, therefore, that augmented G-CSF levels leading to increased TH2 inflammation could be a confounding factor that has contributed to the disappointing results of clinical trials with CXCR2 antagonists. Alternative approaches, which target neutrophilic inflammation in combination with reducing the compensatory increase in G-CSF, could potentially be more clinically beneficial. Alternatively, would we be better equipped with strategies that seek to promote neutrophil apoptosis in tissues (5961) so as to reduce neutrophil numbers but simultaneously suppress the IL-23–IL-17–G-CSF axis? More broadly, our study highlights the complexities of targeting neutrophils within the clinic and implies that broad-sword approaches to block neutrophils may be suboptimal given neutrophils’ heterogeneity, pleiotropic functionality, and regulatory capacity. It would be prudent to gain a fuller understanding of pathways by which neutrophils instigate pathology in asthma so that these facets of their biology may be targeted more specifically. Our findings may also, of course, have implications outside the remit of asthma, whereby a multitude of animal studies have investigated depletion or manipulation of neutrophil numbers in diverse inflammatory models, and numerous clinical trials have assessed therapies aimed at attenuating neutrophilic inflammation. Although we have demonstrated the consequence of G-CSF accumulation in the context of antigen sensitization and type 2 inflammation of the airways, it will be intriguing to ascertain the implications in other experimental systems.

In conclusion, we demonstrate that depletion of neutrophils in the context of a murine model of allergic airway disease results in a dysregulated IL-23–IL-17–G-CSF feedback loop. Ensuing accumulation of G-CSF functioned to augment allergen sensitization by directly potentiating ILC2 TH2 cytokine production and acting on bone marrow progenitors to drive a monocytosis, which, in turn, resulted in augmented antigen presentation by moDCs. These studies are of basic biological significance in that they demonstrate previously unrecognized roles for G-CSF and of translational significance in that they highlight the capacity of neutrophils to indirectly restrain allergic airway inflammation, potentially rationalizing the failure of previous neutrophil-targeting therapeutic strategies.

MATERIALS AND METHODS

Study design

The primary objective of this study was to define the importance of neutrophils in governing inflammation and pathology in allergic airway disease. In all experiments, appropriate control groups were used, and mice were housed under the same environmental conditions and were age-matched. Adult female mice were randomly placed in distinct experimental groups. Authors were blinded for cell counts and histology analysis. The number of mice in each group was determined by power calculations based on extensive previous experience with the model system and is defined in the respective figure legends. The number of independent replicates for each experiment is defined within the respective figure legends. No samples or animals were excluded from data analyses.

Human-isolated ILC2s were used to determine the direct effects of G-CSF on ILC2 functionality. Blood was obtained from three male and two female donors. The number of donors was determined by previous experience with blood-derived ILC2 culturing methods. Authors were blinded from analysis of supernatant cytokine and mRNA expression analysis. All research ethics and patient consent were in place and were further detailed in the “Human ILC2 cultures” section.

Experimental animals

Eight- to 12-week-old female Balb/c mice were purchased from Envigo (Huntingdon, UK). All mice were randomly assigned to experimental groups. IL13-eGFP mice were provided by A. N. McKenzie (Medical Research Council Laboratory of Molecular Biology, Cambridge) and subsequently bred in house (29). Mice were kept in specific pathogen–free conditions and provided autoclaved food, water, and bedding. All mouse experiments were performed in accordance with the recommendations in the Guide for the Use of Laboratory Animals of Imperial College London, with the ARRIVE (Animal Research: Reporting of In Vivo Experiments) guidelines. All animal procedures and care conformed strictly to the UK Home Office Guidelines under the Animals (Scientific Procedures) Act 1986, and the protocols were approved by the Home Office of Great Britain.

Allergic airway disease murine model

Balb/c mice were administered 25 μg of HDM extract [DerP1 (10.17 μg/liter), 9 endotoxin units per endotoxin content; Greer Laboratories] in 50 μl of sterile PBS intranasally three times per week for up to 3 weeks. Control mice were administered 50 μl of sterile PBS intranasally at analogous time points. All mice were culled 24 hours after the final dose of HDM or PBS.

Recombinant IL-33 and G-CSF administration

Mice were intranasally administered 1 μg of recombinant IL-33 protein (Thermo Fisher Scientific) in 50 μl of PBS three times over a period of 1 week. In some experiments, mice were concomitantly intranasally administered 100 ng of recombinant G-CSF (PeproTech, New Jersey, USA) in 50 μl of PBS (or PBS vehicle control) on a daily basis. When necessary, IL-33 was admixed with G-CSF for the treatment of mice. Harvests were performed 24 hours after the final dose of IL-33.

In other experiments, mice administered HDM (intranasally) for 1 week (as detailed above) were also treated daily with 2.5 μg of recombinant G-CSF intraperitoneally in 200 μl of PBS (or PBS vehicle control) and 100 ng of G-CSF intranasally in 50 μl of PBS (or PBS vehicle control). Harvests were performed 24 hours after the final dose of G-CSF/HDM.

In vivo neutrophil depletion

To deplete neutrophils, Balb/c mice received intraperitoneal administration of 100 μg of anti-Ly6G (clone 1A8, BioXCell) in 200 μl of PBS. Control mice received 100 μg of isotype control antibody (clone 2A3, BioXCell) in 200 μl of PBS. During the allergic airway disease model, mice were administered the respective antibodies 24 hours before each HDM or PBS treatment.

In vivo G-CSF neutralization

Neutralization of G-CSF was achieved by intraperitoneal administration of 100 μg of anti-mouse G-CSF antibody (R&D Systems). Respective control mice for G-CSF neutralization received intraperitoneal administration of 100 μg of isotype control antibody (clone 2A3, R&D Systems). Where appropriate, the respective antibodies were admixed with anti-Ly6G antibody (clone 1A8, BioXCell) for neutrophil-depleted mice or isotype control antibody (clone 2A3, BioXCell) for control mice. Antibodies were administered 24 hours before each HDM or PBS intranasal exposure.

In-vivo IL-17 neutralization

Neutralization of IL-17 was achieved by intraperitoneal administration of 20 μg of anti-mouse IL-17A antibody (R&D Systems). Respective control mice for IL-17 neutralization received intraperitoneal administration of 20 μg of isotype control antibody (clone 2A3, R&D Systems). Where appropriate, the respective antibodies were admixed with anti-Ly6G antibody (clone 1A8, BioXCell) for neutrophil-depleted mice or isotype control antibody (clone 2A3, BioXCell) for control mice. Antibodies were administered 24 hours before each HDM or PBS intranasal exposure.

Administration of fluorescently tagged OVA

OVA (Invivogen) was tagged with Alexa Fluor 488 fluorophore using an Alexa Fluor 488 Protein Labeling kit as per the manufacturer’s guidelines (Thermo Fisher Scientific). Mice were administered HDM intranasally for 1 week, and upon the last HDM dose, mice received HDM mixed with 100 μg of Alexa Fluor 488–tagged OVA intranasally.

Lung function measurements

For studies where mice were administered HDM for 3 weeks, measurements of dynamic resistance, elastance, and compliance were performed in anesthetized and tracheotomized mice using a Flexi-vent system (SCIREQ) in response to increasing concentrations (0, 3, 10, 30, and 100 mg/ml) of methacholine (Sigma-Aldrich), as described previously (62).

Cell recovery and isolation

Mice were exsanguinated via cardiac puncture, and 200 μl of blood was immediately lysed in ACK buffer [0.15 M ammonium chloride, 1 M potassium hydrogen carbonate, and 0.01 mM EDTA (pH 7.2)] for 5 min and subsequently centrifuged at 800g for 5 min before the cells were resuspended in 0.5 ml of complete medium (R10F; RPMI supplemented with 10% heat-inactivated fetal bovine serum). Serum was isolated from excess clotted blood by centrifugation (8 min at 5000g). Bronchoalveolar lavage (BAL) was performed by inflating the lungs three times each with 0.4 ml of PBS via a tracheal cannula. The BALF was then pooled and centrifuged at 800g for 5 min, and the BAL supernatant was collected and frozen at −80°C until required. The remaining cell pellet was resuspended in 0.5 ml of R10F.

The accessory, middle, and superior lobes of the right lung were snap-frozen in liquid nitrogen and subsequently stored at −80°C until required. The left lobe was chopped finely and incubated at 37°C for 30 min in complete medium containing liberase (0.15 mg/ml; Sigma-Aldrich) and deoxyribonuclease (DNase) (25 μg/ml; type 1, Sigma-Aldrich). The cells were recovered by disruption through a 70-μm sieve before being centrifuged at 800g for 5 min. Lysis of red blood cells in ACK buffer was performed for 3 min at room temperature before a final centrifugation at 800g for 5 min and resuspension of the remaining cell pellet in 2 ml of R10F.

Both femurs from each mouse were removed and freed of soft tissue attachments, and the extreme distal tip of each femur was cut off. Both ends of the femurs were flushed with 1 ml of PBS containing 0.1% (w/v) sodium azide and 1% (w/v) bovine serum albumin (BSA). The cells were carefully dispersed by filtration through a 70-μm sieve before being centrifuged at 800g for 5 min. Lysis of red blood cells in ACK buffer was performed for 3 min at room temperature before a final centrifugation at 800g for 5 min and resuspension of the remaining cell pellet in 2 ml of R10F.

The spleens of respective mice were disrupted through a 70-μm sieve, and the suspension was centrifuged at 800g for 5 min. Red blood cell lysis was performed for 3 min using ACK buffer, and the cells were centrifuged at 800g for 5 min before the final resuspension of the cell pellet in 2 ml of R10F.

MLNs were digested in liberase (0.15 mg/ml; Sigma-Aldrich) for 30 min. The cells were recovered by disruption through a 70-μm sieve before being centrifuged at 800g for 5 min. The cell pellet was subsequently resuspended in 2 ml of R10F.

FACS of leukocytes and stromal cells

Lungs of mice were inflated with 1.5 ml of dispase II (5 mg/ml; Sigma-Aldrich) and then allowed to collapse naturally. Low–melting point agarose [0.5 ml of 1% (w/v)] was then slowly injected into the lungs and was immediately solidified by packing of the lungs in ice. Lungs were then removed and incubated for 40 min in dispase solution. Lung tissue was subsequently transferred to Dulbecco’s modified Eagle’s medium (DMEM) containing Hepes (25 mM; Thermo Fisher Scientific) and DNase I (50 μg/ml; Sigma-Aldrich), and the digested tissue was “teased away” from the upper airways and incubated with gentle agitation for a further 10 min. Digested lung tissue was disrupted into single-cell suspensions by passage through a 70-μm sieve (BD Labware) before being centrifuged at 800g for 5 min. Lysis of red blood cells in ACK buffer was performed for 3 min before a final centrifugation at 800g for 5 min and resuspension of the remaining cell pellet in 2 ml of R10F. Cell suspensions were stained with anti-mouse CD45-PerCP (peridinin chlorophyll protein), anti-mouse EpCAM-PE (phycoerythrin), and anti-mouse CD31-APC (allophycocyanin) as detailed below, and populations (endothelial cells: CD45 CD31+ EpCAM; epithelial cells: CD45 CD31 EpCAM+; hematopoietic cells CD45+ CD31 EpCAM) were isolated by FACS on a BD FACS LSR Aria III sorter. Isolated cells were centrifuged 800g for 5 min and resuspended in 350 μl of RLT buffer (Qiagen) and stored at −80°C for real-time PCR (see below).

FACS of T cells and ILC2s

Single-cell suspensions were obtained from lung tissue and BAL as described above in the “Cell recovery and isolation” section. For interrogation of airway T cells and ILC2s, it was necessary to pool the BAL from three mice for each data point. Cell suspensions were stained with anti-mouse CD4-FITC (fluorescein isothiocyanate), anti-mouse CD45-PerCP, anti-mouse ICOS-PE-Cy7, and an internally made lineage (Lin) cocktail [consisting of T cell receptor β (TCRβ)–APC, CD5-APC, CD19-APC, TCRγδ-APC, CD11b-APC, CD11c-APC, NKp46-APC, FCεR1-APC, GR-1-APC, F4/80-APC, TER-119–APC] as detailed in table S2, and populations (T cells: Lin+ CD45+ CD4+; ILC2s: Lin CD45+ CD4 ICOS+ KLRG1+) were isolated by FACS on a BD FACS LSR Aria III sorter. Isolated cells were centrifuged 800g for 5 min and resuspended in 350 μl of RLT buffer (Qiagen) and stored at −80°C for real-time PCR (see below).

Human ILC2 cultures

Blood from healthy nonallergic male (n = 3) and female (n = 2) volunteers, with an age range of 24 to 40 years, was collected with written informed consent approved by the Brompton, Harefield, and National Heart and Lung Institute ethics committee. All experiments were carried out in accordance with the approved guidelines. Blood (50 ml) was collected from each donor via venipuncture and placed in 1 ml of EDTA (0.5 M; Thermo Fisher Scientific). Whole-blood ILC2s were enriched using the RosetteSep Human ILC2 Enrichment Kit (STEMCELL Technologies, UK) as per the manufacturer’s instructions. The enriched ILC2s were then stained with anti-human lineage cocktail-FITC (Thermo Fisher Scientific), anti-human CD45- PerCP (Thermo Fisher Scientific), and anti-human CRTH2-BV421 (BioLegend) antibodies for 30 min before being sorted using a BD LSR Aria III cell sorter (BD Biosystems). The ILC2s were identified as Lin CD45+ CRTH2+. The isolated ILC2s were cultured in medium containing DMEM and 10% fetal calf serum (FCS), IL-7 (5 ng/ml; PeproTech), and IL-33 (10 ng/ml; PeproTech) for 2 weeks. The cells were subsequently split and cultured in control medium or medium supplemented with G-CSF (100 ng/ml; PeproTech) for 72 hours. After stimulation, the cell supernatant was collected for protein analysis, and the cells were lysed with 350 μl of RLT buffer (Qiagen).

Mouse ILC2 culture

Mice were administered 1 μg of recombinant IL-33 (Thermo Fisher Scientific) for 1 week as detailed in the “Recombinant IL-33 and G-CSF administration” section. At 24 hours after the final IL-33 dose, lung tissue was extracted and single-cell suspensions were obtained as described in the “Cell recovery and isolation” section. ILC2s were sorted as described in the “FACS of T cells and ILC2s” section. After sorting, the isolated ILC2s were immediately plated in medium containing DMEM + 10% FCS and IL-7 (5 ng/ml; PeproTech) containing IL-33 (10 ng/ml; PeproTech) or lacking IL-33. Respective wells were subsequently supplemented with either G-CSF (10 ng/ml) or G-CSF (100 ng/ml) for 72 hours. After 72 hours, the supernatant was collected for the measurement of IL-5 and IL-13 by ELISA, and the cells were lysed in 350 μl of RLT buffer (Qiagen) for RNA analysis. The lysed cells were assessed for mRNA expression of Il5, Il13, and Gata3.

Flow cytometry

Cells were stained with the LIVE/DEAD Fixable Near-IR-Dead Cell Staining Kit (Molecular Probes, Invitrogen) for 10 min in PBS before being blocked with anti-CD16/CD32 Fc receptor block (BD Pharmingen) for 20 min. Cells were then washed in PBS, stained for surface markers for 30 min at 4°C in PBS that contained 0.1% (w/v) sodium azide and 1% (w/v) BSA, and fixed with 2% (v/v) paraformaldehyde. All samples were acquired immediately on a BD LSRFortessa cell analyzer (BD Biosystems) and analyzed using BD FACSDiva (BD Biosystems). Cells were defined by markers, as described in table S2.

For intracellular cytokine staining, single-cell suspensions from lung tissue and BAL were stimulated with phorbol 12-myristate 13-acetate (PMA; 40 ng/ml; Sigma-Aldrich) and ionomycin (3 μg/ml; Merck Millipore) in complete medium supplemented with brefeldin A (10 μg/ml; Sigma-Aldrich) at 37°C and 0.5% CO2 for 3 hours. The cells were then stained for extracellular markers and fixed as stated above. Cells were subsequently permeabilized with saponin buffer [PBS with 0.05% (w/v) sodium azide, 1% (w/v) BSA, and 1% (w/v) saponin (Sigma-Aldrich)] containing anti-mouse IL-13–PE, anti-mouse IL-5–BV421, anti-mouse IL-4–FITC, anti-mouse IL-17A–Alexa Fluor 700, and/or anti-mouse IL-23–PE-Cy7. For Ki-67 staining, the cells were incubated with anti-mouse Ki-67–Alexa Fluor 488 antibodies. Then, 30 min later, cells were washed once in saponin buffer and once in PBS containing 0.1% (w/v) sodium azide and 1% (w/v) BSA, and data were acquired immediately on a BD LSRFortessa cell analyzer.

Cytokine analysis

Murine superior and middle lung lobes were homogenized at a concentration of 50 mg/ml in PBS. Lung homogenates were centrifuged for 10 min (800g), and supernatant was harvested for cytokine analysis. Lung homogenates, BALF, serum, and mouse ILC2 supernatants were assessed for concentrations of MMP-9, MPO, CCL2, CX3CL1, G-CSF (DuoSet, R&D Systems), IL-13 (Thermo Fisher Scientific), IL-4, and IL-5 (BD Pharmingen), where relevant, by ELISA.

Human ILC2 cell supernatants were assessed for cytokines using the LEGENDPlex Human TH Cytokine Kit. The preparation of samples was performed as per the manufacturer’s instructions and recorded on a BD LSRFortessa flow cytometer (BD Biosystems).

Measurement of immunoglobulin concentrations

Total IgG1 and IgE (BD Biosciences, UK) were measured in diluted serum using standardized sandwich ELISAs according to the manufacturer’s protocol. Allergen-specific IgE and IgG1 were measured in diluted serum as previously described (63).

Measurement of airway mucin concentrations

BALF MUC5AC protein was determined using a previously described protocol (64). Briefly, BALF was placed on a 96-well plate (Corning, UK) and allowed to evaporate overnight at 37°C. The plate was subsequently washed and blocked with 2% BSA for 2 hours. Protein Muc5ac was measured using a biotinylated anti-Muc5ac detection antibody (400 ng/ml; Thermo Fisher Scientific).

Quantitative reverse transcription PCR

For whole lung, tissue was homogenized in RLT buffer (Qiagen) and total RNA was extracted using the RNeasy Plus Mini Kit (Qiagen). Reverse transcription (RT) was subsequently performed using the High-Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). For suspensions of isolated cell populations [in RLT buffer (Qiagen)], total RNA was extracted using the RNeasy Plus Micro Kit (Qiagen) and RT was performed using the GoScript Reverse Transcriptase Kit (Promega). TaqMan gene expression assays (Thermo Fisher Scientific) were used to assess relative expression of murine Csf3, Csf3r, Cxcl1, Il4, Il5, Il13, Il17, Il23, Gata3, and Muc5ac mRNA expression and normalized to Gapdh. Similarly, relative expression of human IL5, IL13, and GATA3 expression was normalized to GAPDH. Quantitative RT-PCR was performed on the ViiA 7 Real-Time PCR System (Thermo Fisher Scientific). Data were obtained from two technical replicates and expressed as fold change in ΔΔCT from respective control groups or relative expression of gene (calculated as 1000 × ΔCT2).

Precision-cut lung slicing

Control and neutrophil-depleted mice were intranasally administered HDM for 1 week. Twenty-four hours after the final HDM dose, anti-mouse CD31–Alexa Fluor 647 (Thermo Fisher Scientific), anti-mouse CD115-PE (BioLegend), and anti-mouse Ly6G–Alexa Fluor 488 (BioLegend) antibodies (at a concentration of 50 μg/ml) were intravenously injected to stain blood cells approximately 5 min before harvesting. Anti-mouse CD115-PE (BioLegend), anti-mouse Ly6G–Alexa Fluor 488 (BioLegend), and anti-mouse CD11c-BV421 (BioLegend) antibodies (at a concentration of 50 μg/ml) were also intranasally administered to stain cells within the airways. Mice were harvested, the tracheas were exposed, and the lungs were inflated with 1 ml of 2% low–melting point agarose (Thermo Fisher Scientific). Once the agarose was solidified, the right lung lobe was isolated and sliced to 300 μl thickness using a Bio-Rad H1200 vibratome (Bio-Rad, France). The lung slices were immediately washed twice in R10F. To ensure staining of tissue-resident cells, the lung slices were incubated with anti-mouse CD115-PE (BioLegend), anti-mouse Ly6G–Alexa Fluor 488 (BioLegend), anti-mouse Ly6G–Alexa Fluor 488 (BioLegend), and anti-mouse CD11c-BV421 (BioLegend) antibodies (at a concentration of 50 μg/ml) for 30 min at room temperature. The fully stained lung slices were washed three times with R10F before being mounted onto an imaging μ-plate (ibidi). Images were obtained with an inverted SP5 confocal microscope using a 20× objective (Leica, UK). The images were subsequently analyzed and prepared using Imaris software version 8.1 (Bitplane, Oxford Instruments, UK).

Lung sectioning for H&E staining

After 3 weeks of HDM/PBS exposure, histology assessment was performed as previously described (62). Briefly, the inferior lobe of the right lung was fixed with 10% neutral-buffered formalin for 24 hours. The lungs were then paraffin wax–embedded and cut to 4 μm thickness. The sections were subsequently stained with H&E to assess general inflammation and airway morphology.

Statistical analysis

Statistical significance was calculated with a nonparametric Mann-Whitney test (two-sided) and Prism software (GraphPad Software Inc.). Statistical significance for human ILC2 experiments was calculated using a paired t test. Statistical significance for mouse ILC2 experiments was calculated using an analysis of variance (ANOVA) test with Bonferroni correction. Results are depicted as means ± SEM unless stated otherwise. Statistical significance for correlations was calculated using a Spearman rank test with P values and r values noted on the respective graphs. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 were considered significant and are referred to as such in the text.

SUPPLEMENTARY MATERIALS

immunology.sciencemag.org/cgi/content/full/4/41/eaax7006/DC1

Fig. S1. Intranasal administration of HDM induces neutrophilic inflammation in the lungs and airways of mice.

Fig. S2. Intranasal administration of HDM induces an increase in neutrophil proteases in the lungs and airways of mice.

Fig. S3. Intraperitoneal administration of anti-Ly6G antibody, 1A8, systemically depletes neutrophils in HDM-treated mice.

Fig. S4. Intraperitoneal administration of anti-Ly6G antibody, 1A8, specifically depletes neutrophils.

Fig. S5. Neutrophil-depleted mice display augmented type 2 inflammation, epithelial remodeling, and airway resistance after 3 weeks of HDM exposure.

Fig. S6. Neutrophil-depleted mice display comparable airway hyperresponsiveness after 3 weeks of HDM exposure.

Fig. S7. Augmented TH2 cytokine levels in neutrophil-depleted mice administered HDM for 1 week.

Fig. S8. ILC2s are the predominant source of IL-13 after 1 week of HDM administration.

Fig. S9. Gating strategy for isolation of ILC2s and CD4+ T cells by FACS.

Fig. S10. Augmented IL-5 and IL-13 levels in lungs of neutrophil-depleted mice administered HDM for 1 week are primarily derived from ILC2s.

Fig. S11. ILC2s from neutrophil-depleted mice administered HDM for 1 week produce more IL-5 and IL-13 on a per-cell basis.

Fig. S12. HDM-exposed neutrophil-depleted mice exhibit augmented DC numbers and antigen presentation.

Fig. S13. Lung concentrations of classical monocyte chemokines and flow cytometry gating strategy to identify MDPs and CDPs.

Fig. S14. G-CSF is elevated in the serum of neutrophil-depleted mice administered HDM.

Fig. S15. Neutrophil-depleted mice administered HDM produce elevated levels of IL-17 and IL-23.

Fig. S16. Neutralization of IL-17 reverts the effects of neutrophil depletion in HDM-administered mice.

Fig. S17. G-CSF directly augments TH2 cytokine production from mouse ILC2s.

Fig. S18. Neutrophil depletion alters the IL-23–IL-17–G-CSF regulatory feedback pathway to exacerbate TH2 cytokine production and allergen sensitization.

Table S1. Raw data file.

Table S2. List of antibodies used for flow cytometry staining.

REFERENCES AND NOTES

Acknowledgments: We thank L. Lawrence for histological sectioning and staining. Funding: R.J.S. is a Wellcome Trust Senior Research Fellow in Basic Biomedical Sciences (209458/Z/17/Z). C.M.L. is a Wellcome Trust Senior Fellow in Basic Biomedical Sciences (107059/Z/15/Z). T.P. is supported by a Marie Curie Intra European Fellowship within the Seventh European Community Framework Programme (FP7-PEOPLE-2013-IEF No627374). A.S. is supported by a pump priming grant from the British Lung Foundation (PPRG15-9) and a research grant from the British Medical Association (HC Roscoe 2015 grant). Aspects of the work were funded by an award from the Rosetrees Trust (M612) to R.J.S., L.G.G., and C.M.L. L.M.C. was funded by core support from Cancer Research UK (A23983 and A17196), the MRC (MR/M01245X/1), and the National Heart and Lung Institute Foundation. Author contributions: D.F.P. and R.J.S. designed and interpreted the experiments, performed statistical analysis, and prepared the manuscript. D.F.P. performed most of the experiments with assistance from T.P., N.B., J.V., S.A., F.P., C.J.P., K.S., A.S., S.A.W., L.G.G., and R.J.S. L.M.C. and C.M.L. provided key reagents and contributed discussions throughout the work. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
View Abstract

Navigate This Article