Many pulmonary infections elicit lymphocyte responses that lead to an accumulation of granulocytes in the lungs. A variety of lymphocytes are capable of directing eosinophils or neutrophils to the lungs, but the contribution of each subset remains enigmatic. In this study, we used a murine model to examine lymphocyte subsets that ultimately drive the eosinophil or neutrophil response to infection with the fungal pathogen Cryptococcus neoformans. We show that granulocytes are produced in the bone marrow, released into the blood stream, and accumulate in the lungs under the instruction of lung parenchymal lymphocytes. The eosinophils that populated the lungs of wild-type animals were highly dependent on Th cells or IL-5. Surprisingly, infected mice with Th cell impairment experienced a compensatory neutrophil response that required IL-17A. This unexpected swing in the response prompted us to investigate the ability of different lymphocyte subsets to produce this dichotomous eosinophilia or neutrophilia. We used mice with lymphocyte deficiencies to determine which of the remaining IL-5– or IL-17A–producing lymphocyte subsets dominated the neutrophil or eosinophil response. Finally, skewing the response toward neutrophil-inducing lymphocytes correlated with accelerated disease. Our data collectively demonstrate that the predominance of a lymphocyte subset determines the functional consequences of an immune response to pulmonary fungal infection that can ultimately affect disease.

Fungi belong to a unique class of pathogens in which the host relies on lymphocytes to confer protection from invasive disease, but the host also suffers from the unfortunate consequence of lymphocyte-mediated pathology. A selective elimination of T cells due to HIV infection results in more frequent and severe mycosis in individuals with AIDS, revealing a potent, beneficial role for lymphocytes in preventing disseminated mycosis (1). Conversely, most cases of allergic asthma experienced by otherwise healthy individuals directly result from inappropriate lymphocyte responses to fungal exposure (2). This highlights the heterogeneity among lymphocytes responding to fungal infection and further draws attention to the delicate balance of lymphocyte homeostasis. Thus, pulmonary fungal infections are an optimal model to study lymphocyte regulation.

Eosinophils and neutrophils are granulocytes that originate from the same myeloblast progenitor in the bone marrow. Upon differentiating into eosinophils or neutrophils, each of these granulocytes leaves the marrow and migrates to the inflamed tissues to enact their unique effector functions. Eosinophils expel toxic products from granules into the extracellular inflammatory milieu (3). Although this is an effective strategy to ward off helminths, it can cause host damage and, when dysregulated, contribute to allergic airway disease (4). Neutrophils also produce toxic granules, yet the additional abilities to shed extracellular traps and efficiently phagocytose microbes make neutrophils particularly useful to protect against some fungal and bacterial pathogens (5, 6).

Among other important functions, lymphocytes secrete cytokines that coordinate eosinophil and neutrophil responses. Type 2 CD4+ T (Th2) cells and type 2 innate lymphoid cells (ILCs) produce IL-5 that prompts eosinophil production under homeostatic and inflammatory conditions (7, 8). Th17 cells, CD8+ T (Tc17) cells, γδ T cells, and type 3 ILCs secrete IL-17A, leading to neutrophil maturation (911). Some pulmonary pathogens elicit type 2 responses and eosinophil accumulation (12, 13), whereas other pulmonary pathogens prompt type 3 responses and neutrophil accumulation (14, 15). Moreover, a mixture of these lymphocyte/granulocyte responses is most advantageous to resist several parasitic and fungal lung infections (16, 17). Thus, a dynamic interaction between the host and microbe fundamentally influences immunity/immunopathology, and this spurs an important question of how granulopoiesis is coordinated by an array of lymphocytes with the potential to promote or resist infectious disease.

To investigate lymphocyte and granulocyte regulation, we used a well-established murine model of experimental cryptococcosis. A highly virulent strain of the fungus Cryptococcus neoformans, KN99α (18), was instilled into the lungs of C57BL6/J mice, and we analyzed the lymphocyte and granulocyte composition in the bone marrow, blood, and lungs following infection. Eosinophils populated the lungs in a lymphocyte- and IL-5–dependent manner, and the modest accumulation of neutrophils was independent of lymphocytes and IL-17A. Surprisingly, the singular elimination of Th cells resulted in a loss of eosinophils that was replaced by IL-17A–dependent neutrophilia. Using mice with genetic deficiencies in various lymphocyte subsets, we revealed an intricate relationship between lymphocyte compensation and the downstream eosinophil and neutrophil responses. Through these genetic experiments, we were able to define a hierarchy of lymphocyte compensation that had polarizing effects on granulocyte accumulation in the lungs. Finally, perturbations of lymphocytes that skewed granulocyte balance toward neutrophilia correlated with exacerbated disease. Taken together, these findings offer unique insights into the dynamic regulation of lymphocyte and granulocyte responses to pulmonary fungal infection that could be useful for the design of immune-targeted therapies to treat human mycoses.

All mice used in this study were derived from a C57BL/6 background. B6.129P2(C)-Ccr7tm1Rfor/J, B6.129S2-H2dlAb1-Ea/J, Il17atm1.1(icre)Stck/J, B6.129S7-Rag1tm1Mom/J, B6.129S2(C)-Stat6tm1Gru/J, and B6.129S2-Tcratm1Mom/J mice were purchased from the Jackson Laboratory (Bar Harbor, ME). C57BL/6-flt3Ltm1Imx and B10;B6-Rag2tm1Fwa II2rgtm1Wjl mice were purchased from Taconic (Germantown, NY). Mice were housed in specific pathogen–free conditions and were fed ad libitum.

Six- to eight-week-old sex-matched mice were anesthetized with pentobarbital. A total of 5 × 104C. neoformans var. grubii (KN99α) cells in 25 μl of PBS was placed on the nares of each mouse, and the mice were kept upright while the inoculum was aspirated into the lower respiratory tract. For survival studies, mice were monitored for end point surrogates: 20% decline in initial weight or 1 g of weight loss over two consecutive days. CFU were determined by removing 1/50 of the lung digest and plating 10× serial dilutions on yeast peptone dextrose plates.

All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless noted otherwise.

To deplete Th cells or block IL-5, mice were injected with 500 μg of anti-CD4 (GK1.5) or 1 mg of anti–IL-5 (TRFK5; both from Bio X Cell, West Lebanon, NH) into the peritoneal cavity on days −1, 5, and 10 postinfection. Intravenous or intranasal instillation of 3 μg of fluorescently labeled anti-CD45.2 (104; BioLegend), followed by ex vivo staining with CD45 (30-F11; both from BioLegend, San Diego, CA), was used to distinguish blood-circulating (i.v. CD45.2+ CD45+), airway-resident (intranasal CD45.2+ CD45+), and parenchymal leukocytes (i.v./intranasal CD45.2 CD45+), as described previously (19).

Leukocytes were isolated from lung digests as previously described (20). For bone marrow cells, femurs were harvested from euthanized mice and pulverized in RPMI 1640. Approximately 250 μl of blood was collected from anesthetized mice via retro-orbital bleeding. RBCs were lysed in whole blood and bone marrow by incubating the samples in ACK buffer for 15 min at room temperature. Cell solutions were filtered, washed several times, and resuspended in PBS staining buffer (PBS + 2 mM EDTA + 0.5% albumin) at 5 × 106 cells per milliliter. For intracellular cytokine analyses, 106 cells were suspended in 200 μl of restimulation buffer (RPMI 1640 + 10% FBS + 1% penicillin/streptomycin + 5 μg brefeldin A) without (unstimulated) or with (stimulated) 10 ng of PMA and 50 ng of ionomycin. After 5 h, the cells were washed and immediately prepared for flow cytometry.

Samples were incubated for 15 min with CD16/32 Ab (BioLegend) and LIVE/DEAD Fixable Near Infrared stain (Invitrogen, Carlsbad, CA) to prevent nonspecific Ab binding, as well as to mark dead cells.

Granulocytes.

Samples were surface stained at 4°C for 30 min with the following Abs: CCR3 (J07E35, PE; BioLegend), Ly6G (1A8, PE-Cy7; BioLegend), CD11b (M1/70, BV650; eBioscience, San Diego, CA), CD11c (N418, PE-Cy5; eBioscience), Siglec F (E50-2440, PE; BD Biosciences, San Jose, CA), IL-5Rα (T21, FITC; BD Biosciences), CD45 (30-F11; Alexa Fluor 700, BioLegend), and erythroid cells (TER-119, Percp-Cy5.5; BioLegend).

Lymphocytes.

Samples were surface stained at 4°C for 30 min with the following Abs: CD90.2 (30-H12, allophycocyanin; BioLegend), CD4 (RM4-5, eFluor 450; eBioscience), CD8 (BV650; BioLegend), TCRβ (H57-597, PE-Cy7), TCRγδ (GL3, Alexa Fluor 488), CD11b (M1/70, Percp-Cy5.5), CD11c (N418, Percp-Cy5.5), F4/80 (BM8, Percp-Cy5.5), CD49b (DX5, Percp-Cy5.5), Ly6G (1A8, Percp-Cy5.5), CD19 (6D5, Percp-Cy5.5), FcεRIα (MAR1, Percp-Cy5.5; all from BioLegend), CD161 (PK136, Percp-Cy5.5), and B220 (RA3-6B2, Percp-Cy5.5; both from eBioscience). For intracellular cytokine detection, the cells were incubated in Foxp3 Transcription Factor Buffer (eBioscience) at 4°C for 30 min. The fixed cells were labeled with Abs against the intracellular Ags IL-5 (TRFK5) and IL-17A (BL168, BV605; both from BioLegend). For data acquisition, >500,000 events were collected on a LSR II flow cytometer (BD Biosciences), and the data were analyzed with FlowJo X (TreeStar, Ashland, OR). Cell numbers per mouse for each leukocyte lineage were calculated by multiplying hemocytometer counts of the cell suspension by the sample dilution factor (1/40) and the cell subset proportion of the total live events collected on the cytometer. The limit of detection for cell quantification is <100 cells.

Lungs were excised from naive mice or from mice 14 d postinfection, snap-frozen in liquid nitrogen, and homogenized in 3 ml of T-PER Tissue Protein Extraction Reagent (Thermo Fisher Scientific) with cOmplete Protease Inhibitor Cocktail (Roche, Indianapolis, IN). Cytokines were quantified using Luminex technology, according to the manufacturer’s instructions (Bio-Rad, Hercules, CA).

The p values for pairwise comparisons were calculated using the Mann–Whitney U test with Bonferroni adjustments for multiple comparisons. Comparison of multiple samples was by Kruskal–Wallis ANOVA. Power calculations were performed to assess the appropriate sample size for all experiments. The p values ≤ 0.05 were considered statistically significant. All statistics and graphs were processed with Prism 6 (GraphPad, La Jolla, CA).

All animal experiments were performed in concordance with the Animal Welfare Act, U.S. federal law, and National Institutes of Health guidelines. Mice were handled according to guidelines defined by the University of Minnesota Institutional Animal Care and Use Committee.

We used flow cytometry to identify eosinophils and neutrophils in the bone marrow, blood, and lungs. Siglec F is expressed by eosinophils upon commitment to the eosinophil lineage (21). CCR3 binds to eotaxin, and this interaction is required for eosinophil chemotaxis from the bone marrow to the tissues (22). Thus, mature eosinophils are defined by the expression of CCR3 and Siglec F in the bone marrow and blood. However, in the lungs, eosinophils and alveolar macrophages express CCR3 and Siglec F, yet only alveolar macrophages have CD11c on their cell surface. Consequently, lung eosinophils are denoted as Siglec F+ CD11c cells. Neutrophils are uniquely identified as Ly6G+ CD11b+ in the bone marrow, blood, and lungs (23).

Eosinophils and neutrophils develop in the bone marrow, travel through the blood stream, and enter the tissues (24, 25). More specifically, granulocytes can arrive at a site of insult along two trajectories. First, mature granulocytes are distributed throughout the body in vascular pools; upon emergency response, they can be recruited from these pools to the inflamed tissues by a process known as demargination (2628). Alternatively, immature granulocytes in the bone marrow can receive cytokine and chemokine cues from lymphocytes or other cells in the tissues that lead to de novo eosinophil and neutrophil differentiation in the marrow and recruitment to the site of inflammation (29). The kinetics of the granulocyte response is a key difference between these pathways. Demargination occurs on the order of hours to a day, whereas lymphocyte-induced production of granulocytes develops over the course of days to weeks.

To gain insight into whether granulocytes accumulated in the lungs as a result of demargination or a lymphocyte-directed process, we quantified eosinophils and neutrophils throughout the course of pulmonary infection with C. neoformans. CCR3+ Siglec F+ eosinophils progressively increased in the bone marrow (Fig. 1A), and these cells increased slightly in the bloodstream (Fig. 1B). Siglec F+ CD11c eosinophils accumulated in the lungs and accounted for an overwhelming majority of the pulmonary leukocytes at later time points of infection (Fig. 1C). In contrast to the eosinophil response, Ly6G+ CD11b+ neutrophils represented a relatively large proportion of the bone marrow cells, which remained at 30–40% of the total marrow cells (Fig. 1A). Soon after infection (i.e., 0.5 d), neutrophil proportions increased sharply in the blood and lungs, indicating a potential demargination event (Fig. 1B, 1C). Although neutrophils steadily increased in the blood, lung neutrophils waned as the infection progressed beyond 5 d postinfection (Fig. 1B, 1C). Thus, both granulocyte types were produced upon infection, but eosinophils accumulated predominantly in the lungs. The rapid, yet relatively minor, accumulation of neutrophils in the bloodstream and lungs suggests that demargination occurred early in the response. In contrast, the long duration and progressive nature of the eosinophil response to pulmonary fungal infection suggested that eosinophils are induced by a lymphocyte-dependent process and likely not as a result of demargination.

FIGURE 1.

Granulocyte response to pulmonary fungal infection. C57BL6/J mice 0–18 d postinfection with C. neoformans. Flow cytometric analyses (left panels) and line graphs (right panels) showing granulocytes as a proportion of CD45+ TER119 hematopoietic cells. (A) Siglec F+ CCR3+ eosinophils and CD11b+ Ly6G+ neutrophils contained in the bone marrow. (B) Siglec F+ eosinophils and Ly6G+ neutrophils from peripheral blood. (C) Siglec F+ CD11c eosinophils and CD11b+ Ly6G+ neutrophils collected from lung digests. All data are mean ± SEM and represent two independent experiments with ≥6 mice in each group.

FIGURE 1.

Granulocyte response to pulmonary fungal infection. C57BL6/J mice 0–18 d postinfection with C. neoformans. Flow cytometric analyses (left panels) and line graphs (right panels) showing granulocytes as a proportion of CD45+ TER119 hematopoietic cells. (A) Siglec F+ CCR3+ eosinophils and CD11b+ Ly6G+ neutrophils contained in the bone marrow. (B) Siglec F+ eosinophils and Ly6G+ neutrophils from peripheral blood. (C) Siglec F+ CD11c eosinophils and CD11b+ Ly6G+ neutrophils collected from lung digests. All data are mean ± SEM and represent two independent experiments with ≥6 mice in each group.

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The lung has three regions that can contain leukocytes: blood vasculature, airways, and lung parenchyma. To determine in which compartment the leukocytes reside, we performed a series of intravital stains in mice infected 14 d earlier with C. neoformans (19, 30). Only a minor fraction of leukocytes collected from infected mice that received i.v. anti-CD45.2 fluorescent Ab were identified upon tissue digestion and flow cytometric analyses (Fig. 2A). Likewise, cells collected from digested lungs of mice that were treated with anti-CD45.2 Ab via inhalation were mostly absent of Ab-associated fluorescence (Fig. 2B). Therefore, a majority of the leukocytes in the lungs of C. neoformans–infected mice are inaccessible to blood and airway labeling, which indicates that most of the leukocytes reside in the lung parenchyma.

FIGURE 2.

Leukocytes in the lung parenchyma mediate eosinophil and neutrophil accumulation. C57BL6/J mice infected 14 d prior and injected i.v. (A) or intranasally (I.N.) (B) with 3 μg of CD45 Ab to label cells in the blood or airway vasculature, respectively. Representative line graphs and bar graphs of total hematopoietic cells that were in the blood (i.v. stain +), airway (intranasal stain +), or lung parenchyma (i.v. −, I.N. −). (C) Eosinophils (▪) and neutrophils (□) from the lungs of wild-type, CCR7−/−, and Flt3L−/− mice 14 d postinfection with C. neoformans. All data are mean ± SEM and represent two independent experiments with ≥4 mice total in each group. *p < 0.05 versus wild-type.

FIGURE 2.

Leukocytes in the lung parenchyma mediate eosinophil and neutrophil accumulation. C57BL6/J mice infected 14 d prior and injected i.v. (A) or intranasally (I.N.) (B) with 3 μg of CD45 Ab to label cells in the blood or airway vasculature, respectively. Representative line graphs and bar graphs of total hematopoietic cells that were in the blood (i.v. stain +), airway (intranasal stain +), or lung parenchyma (i.v. −, I.N. −). (C) Eosinophils (▪) and neutrophils (□) from the lungs of wild-type, CCR7−/−, and Flt3L−/− mice 14 d postinfection with C. neoformans. All data are mean ± SEM and represent two independent experiments with ≥4 mice total in each group. *p < 0.05 versus wild-type.

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Lymph nodes are conventional locations for lymphocyte priming. However, C. neoformans can elicit a fully competent Th cell response that is not dependent on lymph node responses (20, 30). Therefore, we tested the hypothesis that granulocyte recruitment does not require lymphocyte induction in the mediastinal lymph node. Flt3 ligand (Flt3L) is required for migratory dendritic cell maturation (31), and CCR7 mediates lymphocyte entry into lymph nodes (32). Thus, genetic deficiency in each of these molecules impairs lymph node responses, yet it does not alter pulmonary lymphocyte accumulation in this model of infection (20, 30). We also found that eosinophil and neutrophil recruitment to the lungs was not decreased by Flt3L or CCR7 ablation (Fig. 2C). In fact, neutrophils increased in the lungs of infected CCR7−/− mice compared with wild-type controls (Fig. 2C). This could be due to enhanced T cell priming in the lungs as a consequence of the increased peripheral T cells observed in CCR7−/− mice (32). Therefore, lymph nodes are not a primary location for priming of granulocyte-inducing lymphocytes in this model. As a result, we focused our attention on lymphocytes residing in the lungs to understand how these cells influence granulocyte responses.

Lymphocytes are heterogeneous, so we performed multiparameter flow cytometry to distinguish the various subsets residing in the lungs. Bulk lymphocytes were identified as lineage (CD19, CD11b, CD11c, CD49b, CD161, F4/80, FcεRIα, B220) and CD90.2+. Within the bulk lymphocyte population, there are at least five lymphocyte subsets: ILCs (TCRβ TCRγδ), γδ T cells (TCRβ TCRγδ+), Th cells (TCRβ+ CD4+), CD8+ T cells (TCRβ+ CD8+), and NK cells (lineage+ CD90+). Using this flow cytometry gating strategy, we observed a significant expansion in each lymphocyte population in the lungs of infected wild-type mice compared with uninfected controls (Fig. 3A, 3B). Thus, ILCs, γδ T cells, Th cells, CD8 T cells, and NK cells respond to fungal infection; consequently, each subset could influence granulocyte responses.

FIGURE 3.

Lymphocytes control dichotomous eosinophil/neutrophil response. Naive wild-type or infected wild-type, MHCII−/−, Rag2−/−, or Rag2/IL-2Rγ−/− mice at 14 d postinfection. Flow cytometric analyses (A) and bar graphs (B) of lymphocyte subsets. CD90+ lymphocyte subsets were defined as TCRγδ TCRβ ILCs, TCRγδ+ IL-17A+ γδ T cells, TCRβ+ CD4+ Th cells, TCRβ+ CD8+ CD8 T cells, and lineage+ NK cells (see Supplemental Fig. 1A). Lineage = B220, CD11b, CD11c, CD19, CD49b, CD161, F4/80, FcεRIα. (C) Total numbers of Siglec F+ CD11c eosinophils or CD11b+ Ly6G+ neutrophils collected from whole-lung digests. All data are mean ± SEM and represent two independent experiments with ≥5 mice total in each group. *p < 0.05, Mann–Whitney U test with Bonferroni adjustment for multiple comparisons.

FIGURE 3.

Lymphocytes control dichotomous eosinophil/neutrophil response. Naive wild-type or infected wild-type, MHCII−/−, Rag2−/−, or Rag2/IL-2Rγ−/− mice at 14 d postinfection. Flow cytometric analyses (A) and bar graphs (B) of lymphocyte subsets. CD90+ lymphocyte subsets were defined as TCRγδ TCRβ ILCs, TCRγδ+ IL-17A+ γδ T cells, TCRβ+ CD4+ Th cells, TCRβ+ CD8+ CD8 T cells, and lineage+ NK cells (see Supplemental Fig. 1A). Lineage = B220, CD11b, CD11c, CD19, CD49b, CD161, F4/80, FcεRIα. (C) Total numbers of Siglec F+ CD11c eosinophils or CD11b+ Ly6G+ neutrophils collected from whole-lung digests. All data are mean ± SEM and represent two independent experiments with ≥5 mice total in each group. *p < 0.05, Mann–Whitney U test with Bonferroni adjustment for multiple comparisons.

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We used mice with genetic deficiencies in lymphocyte subsets and monitored the effect that this had on granulocyte accumulation in the lungs of infected mice. To assess whether lymphocytes are required for granulocyte induction, we used Rag2/IL-2Rγ−/− mice that lack lymphocytes (Fig. 3A, 3B). Without lymphocytes, infected mice had significantly impaired eosinophilia compared with similarly infected wild-type mice, yet neutrophil accumulation remained unimpaired (Fig. 3C). Thus, lymphocytes are required for the robust eosinophilia and are dispensable for the modest neutrophil accumulation experienced by mice with lethal pulmonary mycosis.

Three lymphocyte subsets are known to influence eosinophil differentiation: Th cells, NK cells, and ILCs. To determine whether NK cells or ILCs are sufficient to promote eosinophil accumulation, we used Rag2−/− mice, which lack all T cell subsets as a result of a genetic deficiency in TCR rearrangement, leaving ILCs and NK cells as the only lymphocytes. As expected, Rag2−/− mice had very few T cells, but they generated similar numbers of ILCs and NK cells in the lungs as did wild-type mice (Fig. 3A, 3B). In the absence of these T cell subsets, eosinophils populated the lungs in equal quantities as wild-type mice, indicating that ILCs or NK cells were sufficient to induce eosinophilia in the absence of T cells (Fig. 3C).

We also tested whether the elimination of only Th cells would affect the sufficiency of ILCs or NK cells to coordinate eosinophil responses. Th cell development/function in mice (33) and humans (34) is critically and uniquely dependent on TCR engagement of peptides bound by MHC class II molecules on APCs. Therefore, we used MHCII−/− mice to reduce Th cells, while leaving all other lymphocyte subsets intact, including ILCs and NK cells (Fig. 3A, 3B). In contrast to Rag2−/− mice, the singular loss of functional Th cell responses through MHC class II deficiency resulted in a significant decrease in eosinophil accumulation in the lungs at 14 d postinfection (Fig. 3C). Instead, MHCII−/− mice unexpectedly experienced a striking accumulation of neutrophils in the lungs (Fig. 3C).

We also analyzed the response kinetics to understand how the initial innate and later adaptive immunity phases influenced granulocyte development in the absence of Th cells. A slight increase in eosinophil production was evident in the lungs of infected MHCII−/− mice early (up to 5 d), suggesting that ILCs or NK cells were active at the beginning of the response (Fig. 4). However, eosinophil production subsided at the onset of the adaptive immune phase (>5 d) and was replaced by a large neutrophilic response (Fig. 4).

FIGURE 4.

Neutrophils dominate the granulocyte response during the adaptive immune phase in MHCII−/− mice. Pulmonary leukocytes collected from lung digests of MHCII−/− mice from 0 to 14 d of infection. Data are mean ± SEM and represent two independent experiments with a total of six mice per group.

FIGURE 4.

Neutrophils dominate the granulocyte response during the adaptive immune phase in MHCII−/− mice. Pulmonary leukocytes collected from lung digests of MHCII−/− mice from 0 to 14 d of infection. Data are mean ± SEM and represent two independent experiments with a total of six mice per group.

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Taken together, ILCs or NK cells are only sufficient to promote eosinophil accumulation when T cells are absent or during a delay in adaptive T cell priming in the context of a Th cell–deficient environment. Additionally, singular elimination of Th cells results in a preponderance of neutrophils in the lungs.

To delineate how lymphocyte subsets influence granulocyte recruitment, we sought a functional correlate that links lymphocytes and granulocytes. Lymphocytes secrete cytokines that influence a variety of immunological processes, including granulocyte development and recruitment to sites of inflammation. Therefore, we investigated how canonical lymphocyte cytokines (e.g., IFN-γ, IL-5, and IL-17A) were influenced by infection, how these cytokines were affected by lymphocyte deficiency, and/or how the depletion of these cytokines influenced the granulocyte response.

IFN-γ produced by Th1 cells, NK cells, and CD8 T cells mediates protection against cryptococcal infection (35). However, highly virulent strains of C. neoformans have evolved mechanisms to inhibit host production of IFN-γ and evade immune defense (35). The highly virulent strain KN99α that we used in this study did not induce an IFN-γ response above naive levels in the lungs of wild-type mice or any of the mutant mice tested (Fig. 5A). Thus, the alterations in granulocyte responses that we observed in the lymphocyte-depleted mice were not a consequence of fluctuations in IFN-γ signaling.

FIGURE 5.

IL-5 and IL-17A instruct pulmonary eosinophil or neutrophil accumulation. IFN-γ (A), IL-5 (B), and eotaxin (C) measured in lung homogenates from mice infected 14 d previously. (D) Wild-type mice were treated with IL-5–depleting Ab (TRFK5) or isotype control. After 14 d of infection, eosinophils were quantified in lung digests. IL-17A (E) and G-CSF (F) quantified in lung homogenates 14 d postinfection. (G) Neutrophils and eosinophils quantified from lungs of wild-type and IL-17–deficient mice treated with CD4-depleting Ab (α-CD4, GK1.5) 14 d postinfection. All data are mean ± SEM and contain ≥3 mice in each group. *p < 0.05, Mann–Whitney U test with Bonferroni adjustment for multiple comparisons. n.s., not significant.

FIGURE 5.

IL-5 and IL-17A instruct pulmonary eosinophil or neutrophil accumulation. IFN-γ (A), IL-5 (B), and eotaxin (C) measured in lung homogenates from mice infected 14 d previously. (D) Wild-type mice were treated with IL-5–depleting Ab (TRFK5) or isotype control. After 14 d of infection, eosinophils were quantified in lung digests. IL-17A (E) and G-CSF (F) quantified in lung homogenates 14 d postinfection. (G) Neutrophils and eosinophils quantified from lungs of wild-type and IL-17–deficient mice treated with CD4-depleting Ab (α-CD4, GK1.5) 14 d postinfection. All data are mean ± SEM and contain ≥3 mice in each group. *p < 0.05, Mann–Whitney U test with Bonferroni adjustment for multiple comparisons. n.s., not significant.

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IL-5 is a potent inducer of eosinophil differentiation. Large increases in IL-5 were found in lung homogenates from infected wild-type and Rag2−/− mice relative to naive mice (Fig. 5B). Conversely, Rag2/IL-2Rγ−/− and MHCII−/− mice did not mount an IL-5 response above naive controls (Fig. 5B). The elevated IL-5 in wild-type and Rag2−/− mice and the lack of IL-5 in Rag2/IL-2Rγ−/− and MHCII−/− mice directly correlated with the eosinophilia, or lack thereof, experienced by the infected wild-type and mutant animals.

Interestingly, infected wild-type and MHCII−/− mice produced similarly more eotaxin, an eosinophil chemoattractant, than did naive animals (Fig. 5C). The normal expression of eotaxin and the absence of eosinophils in MHCII−/− mice suggest that eotaxin is not sufficient to induce eosinophil accumulation in the lungs. Alternatively, treatment of infected wild-type mice with IL-5–blocking Ab severely impaired eosinophil accumulation in the lungs (Fig. 5D). These data indicate that IL-5 is necessary for the eosinophil response during infection and can be produced by Th cells and ILCs/NK cells.

IL-17A indirectly controls neutrophil maturation by cueing stromal cell production of G-CSF in models of bacterial pneumonia (36). Therefore, we examined the relationship between these cytokines and the neutrophil responses to C. neoformans infection. Infected wild-type, Rag2/IL-2Rγ−/−, and Rag2−/− mice had basal levels of IL-17A in the lungs, yet G-CSF expression in each of these animals was elevated above naive animals (Fig. 5E, 5F). These data suggest that an IL-17–independent stimulus of G-CSF supports the modest production of neutrophils experienced by these animals. In contrast, MHCII−/− mice expressed significantly more IL-17A and G-CSF in the lungs than did any of the other infected animals (Fig. 5E, 5F). These data suggest that an IL-17A–dependent induction of G-CSF may produce the severe neutrophilia that occurs in MHCII−/− mice.

To test the hypothesis that a compensatory IL-17A response by lymphocytes drives neutrophilia, we examined pulmonary neutrophil accumulation in infected wild-type or IL-17A−/− mice treated with anti-CD4 Ab to deplete Th cells. Wild-type mice that were depleted of Th cells mimicked the severe neutrophilia phenotype of MHCII−/− mice. In contrast, neutrophil levels did not increase upon Th cell depletion in IL-17A−/− mice (Fig. 5G), indicating that IL-17A is required for the severe neutrophilia observed in the absence of Th cells. Taken together, the neutrophilia experienced by mice with Th cell deficiency is due to a compensatory increase in IL-17A production by another lymphocyte population.

To understand which lymphocyte subsets are responsible for the production of IL-5 and IL-17A that influences the downstream granulocyte responses in wild-type and lymphocyte-deficient mice, we quantified IL-5 and IL-17A production by Th cells, γδ T cells, ILCs, and CD8 T cell lymphocyte subsets. Of note, IL-5 or IL-17A production by NK and NKT cells in wild-type and lymphocyte-knockout mice was equivalent to that in naive mice; thus, NK and NKT cells likely do not contribute to granulocyte responses in this model of lethal cryptococcosis (Supplemental Fig. 1).

In the wild-type infection, IL-5 was produced by Th cells and ILCs (Fig. 6A, Table I), with these two lymphocyte subsets accounting for 80% of the cytokine-producing lymphocyte response. IL-17A production by Th cells and ILCs was observed, but these cells constituted a minor proportion of the lymphocyte response. Taken together, these data suggest that the eosinophilia experienced by wild-type mice (Fig. 3C, Table I) is due to production of IL-5 by ILC2s and/or Th2 cells.

FIGURE 6.

Cytokine-producing lymphocytes compensate for the loss of other lymphocyte subsets. Wild-type (A), Rag2−/− (B), MHCII−/− (C), STAT6−/− (D), and STAT6−/− (E) mice treated with CD4-depleting Ab (GK1.5) or TCRα−/− mice 14 d postinfection with C. neoformans (F). Flow cytometric analyses (left panels) and bar graphs (right panels) showing IL-5 or IL-17A production by CD90+ lineage lymphocytes: TCRβ+ CD4+ IL-5+ Th2 cells, TCRβ+ CD4+ IL-17A+ Th17 cells, TCRγδ TCRβ IL-5+ ILC2, TCRγδ TCRβ IL-17A+ ILC3s, TCRγδ+ IL-17A+ γδ T cells, and TCRβ+ CD8+ IL-17A Tc17 cells. All data are mean ± SEM and represent at least two independent experiments with ≥5 mice total per group. Lineage = B220, CD11b, CD11c, CD19, CD49b, CD161, F4/80, FcεRIα. The asterisk (*) indicates the most abundant lymphocyte subset(s) and p < 0.05, determined by the Mann–Whitney U with Bonferroni adjustment for multiple comparisons.

FIGURE 6.

Cytokine-producing lymphocytes compensate for the loss of other lymphocyte subsets. Wild-type (A), Rag2−/− (B), MHCII−/− (C), STAT6−/− (D), and STAT6−/− (E) mice treated with CD4-depleting Ab (GK1.5) or TCRα−/− mice 14 d postinfection with C. neoformans (F). Flow cytometric analyses (left panels) and bar graphs (right panels) showing IL-5 or IL-17A production by CD90+ lineage lymphocytes: TCRβ+ CD4+ IL-5+ Th2 cells, TCRβ+ CD4+ IL-17A+ Th17 cells, TCRγδ TCRβ IL-5+ ILC2, TCRγδ TCRβ IL-17A+ ILC3s, TCRγδ+ IL-17A+ γδ T cells, and TCRβ+ CD8+ IL-17A Tc17 cells. All data are mean ± SEM and represent at least two independent experiments with ≥5 mice total per group. Lineage = B220, CD11b, CD11c, CD19, CD49b, CD161, F4/80, FcεRIα. The asterisk (*) indicates the most abundant lymphocyte subset(s) and p < 0.05, determined by the Mann–Whitney U with Bonferroni adjustment for multiple comparisons.

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Table I.
Summary of lymphocyte and granulocyte responses to pulmonary fungal infection
Mouse StrainLymphaIL-5+ Th Cells (%)IL-5+ ILCs (%)IL-17A+ Th Cells (%)IL-17A+ ILCs (%)IL-17A+ CD8 (%)IL-17A+ γδ T (%)Eos/Neub
Wild-type 878,615 30.0c 52.0 9.4 6.1 1.5 1.0 13.0 
Rag2/IL-2Rγ−/− 7,552 N/A N/A N/A N/A N/A N/A 0.12 
Rag2−/− 273,496 0.2 86.7 0.3 12.5 0.1 0.2 8.73 
MHCII−/− 601,371 0.8 18.4 8.0 9.5 56.9 6.3 0.10 
STAT6−/− 743,953 2.4 2.1 78.3 6.9 2.5 7.9 0.06 
STAT6−/− + anti-CD4 721,565 0.1 2.9 0.3 8.7 77.4 10.6 0.01 
TCRα−/− 548,224 0.8 9.5 12.5 16.5 1.3 59.4 0.03 
Mouse StrainLymphaIL-5+ Th Cells (%)IL-5+ ILCs (%)IL-17A+ Th Cells (%)IL-17A+ ILCs (%)IL-17A+ CD8 (%)IL-17A+ γδ T (%)Eos/Neub
Wild-type 878,615 30.0c 52.0 9.4 6.1 1.5 1.0 13.0 
Rag2/IL-2Rγ−/− 7,552 N/A N/A N/A N/A N/A N/A 0.12 
Rag2−/− 273,496 0.2 86.7 0.3 12.5 0.1 0.2 8.73 
MHCII−/− 601,371 0.8 18.4 8.0 9.5 56.9 6.3 0.10 
STAT6−/− 743,953 2.4 2.1 78.3 6.9 2.5 7.9 0.06 
STAT6−/− + anti-CD4 721,565 0.1 2.9 0.3 8.7 77.4 10.6 0.01 
TCRα−/− 548,224 0.8 9.5 12.5 16.5 1.3 59.4 0.03 
a

Average number of CD90+ total lymphocytes isolated from mouse lungs.

b

Eosinophil/neutrophil ratio in the lungs. Values >1 indicate the level of eosinophilia, and values <1 indicate the level of neutrophilia.

c

Percentage of total lymphocytes within the indicated phenotype.

Anti-CD4, Ab depletion of Th cells; N/A, not applicable.

T cell deficiency in Rag2−/− mice resulted in an increase in ILCs (87% of cytokine+ lymphocytes) that express IL-5 (Fig. 6B, Table I). Similar to the wild-type infection, IL-17A+ ILCs are present but account for only a minor proportion of the lymphocytes. Thus, the absence of T cells in Rag2−/− mice resulted in a compensatory increase in IL-5+ ILCs that also correlated with profound eosinophilia (Fig. 3C, Table I).

Much to our surprise, the specific loss of CD4+ Th cells in MHCII−/− mice was not compensated for by ILCs. Rather, MHCII−/− mice experienced a dramatic increase in the number of IL-17A+ CD8 T cells (Fig. 6C, Table I). This increased production of IL-17A in place of IL-5 correlated with a dichotomous switch to a granulocyte response dominated by neutrophils (Fig. 3C, Table I).

To further explore the relative potency of lymphocyte subsets, we infected mice with additional lymphocyte deficiencies and analyzed the compensatory effect that this had on the remaining lymphocytes and granulocyte recruitment. STAT6 regulates the type 2 effector phenotype of lymphocytes (37). In the absence of type 2 lymphocytes, STAT6−/− mice exhibited a compensatory response that was dominated by IL-17A–producing Th cells and neutrophilia (Fig. 6D, Table I, Supplemental Fig. 2A). To compare the relative potency of CD8 T cells, γδ T cells, and ILCs, we depleted Th cells in STAT6−/− mice with anti-CD4 Ab. IL-17+ CD8 T cells outnumbered γδ T cells and ILCs, and this also correlated with a strong neutrophil response (Fig. 6E, Table I, Supplemental Fig. 2A). Next, we restricted the compensatory response to γδ T cells and ILCs by using TCRα−/− mice. In this situation, IL-17A–producing γδ T cells compensated for the loss of Th cells and CD8 T cells (Fig. 6F, Table I), resulting in neutrophilia (Supplemental Fig. 2A, Table I).

Of note, IL-5 cytokine levels in wild-type and Rag2−/− mice with eosinophilia (Fig. 5B), as well as IL-17A cytokine levels in MHCII−/−, STAT6−/−, and TCRα−/− mice, were equivalent (Supplemental Fig. 2B). Thus, the alternate lymphocyte subsets were able to fully compensate for the loss in cytokine production by the missing lymphocyte subsets.

In summary, the dichotomous eosinophilia/neutrophilia observed in mutant mice correlated with a compensatory response by a lymphocyte subset.

We examined how the dichotomy between eosinophilia and neutrophilia affected the ability of the host to control pulmonary C. neoformans infection. All of the mutant animals tested in this experimental system had significantly elevated fungal burden in the lungs at 14 d postinfection compared with wild-type controls (Fig. 7A), suggesting that all perturbations tested had a detrimental impact on the ability to control fungal replication in the lungs. No correlation between fungal burden and eosinophilia/neutrophilia was observed. For example, the fungal burden was equivalent in Rag2−/− mice with eosinophilia and TCRα−/− mice with neutrophilia. These data show that elevated fungal burden alone does not contribute to the dichotomy between eosinophil or neutrophil accumulation in the lungs.

FIGURE 7.

Neutrophil polarization is associated with enhanced disease. (A) CFU in the lungs of wild-type and mutant animals 14 d postinfection with C. neoformans. All data are mean ± SEM and represent at least four mice per group. The p value was calculated using the Mann–Whitney U test with Bonferroni adjustment for multiple comparisons. (B) Survival curve includes 10 mice per group. The p value was calculated using the log-rank test.

FIGURE 7.

Neutrophil polarization is associated with enhanced disease. (A) CFU in the lungs of wild-type and mutant animals 14 d postinfection with C. neoformans. All data are mean ± SEM and represent at least four mice per group. The p value was calculated using the Mann–Whitney U test with Bonferroni adjustment for multiple comparisons. (B) Survival curve includes 10 mice per group. The p value was calculated using the log-rank test.

Close modal

Balanced eosinophil and neutrophil responses are more beneficial than strongly polarized responses to some fungal and parasitic infections (16, 17). In the case of Cryptococcus infection, defective IL-17A production due to IL-23 deficiency and the concomitant increase in eosinophilia are associated with accelerated death in mice (38). To determine whether the opposite was true (i.e., that neutrophilia is also associated with accelerated death), we examined survival of STAT6−/− mice with neutrophilia. STAT6−/− mice were chosen for two reasons. Compared with the other lymphocyte-deficient mice, STAT6−/− mice had only a modest increase in fungal burden (Fig. 7B); consequently, the impact of STAT6 deficiency on lethal disease is less likely to be caused by fungal overgrowth. Also, type 2 responses are considered detrimental in this model; thus, blockade of the deleterious type 2 response should ameliorate disease a priori. Upon infection, STAT6−/− mice exhibited accelerated lethal disease compared with wild-type mice (Fig. 7B), providing evidence that neutrophilia can be detrimental.

A wide array of lymphocytes influence many facets of immunity to infectious microbes. Lymphocyte subsets govern intricate and sometimes opposing responses, including eosinophil and neutrophil production. CD4+ Th cells were long believed to be the main coordinators of eosinophil and neutrophil granulopoiesis through the production of IL-5 and IL-17A, respectively (39). However, the realization that other lymphocyte subsets, like CD8 T cells, γδ T cells, and ILCs, can perform similar functions challenges this understanding (40, 41). Thus, an evaluation of the relative influence of each subset in determining downstream responses (e.g., granulocytes) is warranted.

We dissected the complex regulation of lymphocytes with regard to their ability to coordinate pulmonary granulocyte accumulation in response to C. neoformans infection. Through systematic genetic disruptions of lymphocyte subsets, we identified a hierarchy in lymphocyte compensation that impacts granulocyte polarization. In wild-type mice infected with highly virulent C. neoformans strain KN99α, Th2 cells and ILCs produce IL-5 that is required for the large influx of eosinophils into the lungs. Mice depleted of type 2 responses (i.e., STAT6−/−) exhibited a compensatory IL-17A+ Th cell response that correlated with a predominance of neutrophils. Elimination of all Th cells (MHCII−/−, anti-CD4 Ab treatment) resulted in compensation by IL-17A+ CD8 T cells to induce neutrophil accumulation in the lungs. Depletion of Th cells and CD8 T cells (TCRα−/−) resulted in compensation by an overproduction of γδ T cells, leading to neutrophilia. Finally, ILCs were only able to restore the eosinophil response in the absence of all T cell subsets, as observed in the infection of Rag2−/− animals. Using this readout, we show that the Th2→Th17→Tc17→γδ T→ILC2 compensation results in dramatic shifts in the granulocyte response that depend on lymphocyte availability.

A major impediment to evaluating lymphocyte subset regulation is the identification of an independent correlate of lymphocyte function. We leveraged the potent ability of lymphocytes to direct granulocyte accumulation in the lungs upon fungal infection. We were able to use cytokine production by lymphocytes as a means to identify particular lymphocyte subsets and correlate their presence with a functional outcome (i.e., eosinophil or neutrophil accumulation in the lungs). As a result, we avoided the common circular argument that attempts to mark lymphocyte lineage, as well as assumes function, based on the production of cytokines by lymphocytes. Moreover, the striking dichotomy of either eosinophil or neutrophil responses in this model, without much of an intermediate response in most cases, permitted us to draw compelling correlations between lymphocyte presence and granulocyte induction.

Fei et al. (42) observed a similar phenomenon whereby BALB/c or C57BL/6 mice mounted disparate neutrophil or eosinophil responses with repeated exposure to Aspergillus conidia, and the impairment of TNF-α in BALB/c mice caused a neutrophil-to-eosinophil transition upon Aspergillus infection. This observation of granulocyte switching was further corroborated in a model system of pulmonary inoculation with purified chitin beads (43). Chitin is a polysaccharide that elicits a CD4/8 T cell–independent, ILC-mediated allergic response (44). Depletion of IL-5+ ILCs in chitin-treated mice resulted in a compensatory increase in γδ T cell–driven neutrophil recruitment (43). Our results are consistent with these prior observations, and we also unveiled a mechanism underlying eosinophil/neutrophil reciprocity in the lungs of mice with lethal fungal infections. Methodical examination of lymphocyte competition in the context of bacterial and viral infections could reveal an equally interesting lymphocyte dynamic.

Szymczak et al. (38) reported the presence of NKT cells that express IL-17A in the lungs of C. neoformans–infected mice. It is not entirely clear why we did not find similar IL-17A–producing NKT cells. One key difference between the two models is the strain of Cryptococcus and subsequent disease severity that results from infection with the different strains. In the current study, we used a highly virulent serotype A strain that rapidly kills infected mice, whereas the aforementioned study used a nonlethal serotype D strain that is controlled by an intact immune system. Because NK cells participate in protective immunity to Cryptococcus (45), it is possible that the more virulent strain has adapted an evasion strategy to subvert NKT cell responses.

In recent years, ILCs have emerged as main players in a variety of processes, including maintenance of tissue homeostasis, inflammation, and tissue damage repair (41). The standard model for investigating the sufficiency of ILCs to coordinate these processes involves the use of Rag−/− mice compared with ILC elimination by CD90 Ab treatment or Rag/IL-2Rγ double-knockout mice (43, 4648). Using a panoply of knockout mice, we showed that T cells, not ILCs, are the main propagators of granulocyte responses to C. neoformans infection. Perhaps, a lack of competition with other lymphocytes for shared growth factors, like IL-2 and IL-7, exaggerates the effect of ILCs on driving eosinophilia in Rag2−/− models. With the development of new methods to specifically ablate ILCs (49, 50), a more accurate description of the roles of ILCs during inflammation may follow.

IL-17–induced neutrophil responses are paradigmatically credited for protection against fungal infections (51). Although true in some cases, as evidenced by enhanced susceptibility to mucocutaneous candidiasis in humans with IL-17 deficiencies (52), there are instances where IL-17 responses to fungal infection are deleterious, particularly in the lungs (53). We showed that IL-17–producing lymphocytes and profound neutrophilia correlate with exacerbated cryptococcal disease in type 2–deficient, STAT6−/− mice compared with wild-type animals. Therefore, the protection afforded by IL-17 against some pathogens should be weighed in the context of potential IL-17/neutrophil–associated immunopathology.

The importance of C. neoformans extends beyond a model experimental organism. In fact, cryptococcosis is the leading cause of death in people stricken with AIDS in sub-Saharan Africa (54). The emergence of drug resistance in fungi and the inherent difficulty in targeting eukaryotic pathogens with chemotherapeutics have heightened the urgency to design novel treatment approaches (35). One such strategy under development is IFN-γ adjunct therapy to combat cryptococcosis by augmenting host immunity (55). However, this approach has not demonstrated drastic improvement in patient survival, in part because of the challenges associated with balancing protective immunity and immunopathology. Lymphocytes are gatekeepers of health and disease in this scenario. As a result, our findings about individual contributions of lymphocyte subsets informs important therapeutic targets, predicts problematic lymphocyte compensation, and thereby, affords rational design of clinical interventions. Our data will aid in the development of custom therapies that hold the potential to modulate lymphocytes to promote microbial clearance while also maintaining granulocyte balance and patient health.

We thank Marc Jenkins and Peter Southern for helpful discussions. We are also grateful to the University of Minnesota Flow Cytometry Core Facility for instrumentation.

This work was supported by National Institutes of Health Grants AI080275 and AI122352 (to K.N.), National Institutes of Health T32 Training Grant AI007313, a University of Minnesota Doctoral Dissertation Fellowship, and a Dennis W. Watson Fellowship (to D.L.W.).

The online version of this article contains supplemental material.

Abbreviations used in this article:

Flt3L

Flt3 ligand

ILC

innate lymphoid cell

Th2

type 2 CD4+ T.

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The authors have no financial conflicts of interest.

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