Allergen-induced CD4⁺ T-cell cytokine production within airway mucosal DC-T-cell clusters drives the local recruitment of myeloid effector cells

Abstract

Allergic asthma develops in the mucosal tissue of small bronchi. At these sites, local cytokine production by T_H2/T_H17 cells is believed to be critical for the development of tissue eosinophilia/neutrophilia. Using the mouse trachea as a relevant model of human small airways, we performed advanced in vivo dynamic and in situ static imaging to visualize individual, cytokine-producing T cells in the airway mucosa and to define their immediate cellular environment. Upon allergen sensitization, newly recruited CD4⁺ T cells formed discrete antigen-driven clusters with dendritic cells (DCs). Within T-DC clusters, a small fraction of CD4⁺ T cells produced IL-13 or IL-17 following prolonged, antigen-specific interactions with DCs. As a result of local T_H2 cytokine signaling, eosinophils were recruited into these clusters. Neutrophils also infiltrated these clusters in a T-cell dependent manner but their mucosal distribution was more diffuse. Our findings reveal the focal nature of allergen-driven responses in the airways and define multiple steps with potential for interference with the progression of asthmatic pathology.

Introduction

Allergic asthma primarily affects the mucosal tissue layers of conducting airways. The morphological and functional changes leading to pathological broncho-obstruction in asthmatics occur in a well-defined micro-anatomical compartment of small-diameter bronchi. Classically, cytokine production (e.g., IL-4, IL-5 and IL-13) by effector T_H2 cells has been considered primarily responsible for the pathological changes in asthma, such as airway hyper-reactivity (AHR), airway remodeling and increased mucus production, as well as the infiltration of the airway mucosa with eosinophils, basophils and mast cells. However, recent studies also suggest the involvement of type-2 innate lymphoid cells (ILC2s) as a source of effector cytokines (1, 2). Additionally, some severe forms of asthma are dominated by a T_H17-type response with IL-17 producing CD4⁺ T cells, whose activation in the tissue leads to neutrophilia (3). Indeed, it is becoming increasingly clear that especially in chronic inflammation, several T_H subsets (T_H1, T_H2, T_H17, T_FH and T_regs) simultaneously participate in the inflammatory allergic response (4, 5).

Although the key cellular players and mediators of asthma are well defined (6, 7), we have only limited knowledge on the micro-anatomical organization of the various mediator-producing cells and the specific cellular targets of each mediator within the airway mucosal compartment. This is mainly due to technical difficulties in accessing the airway mucosa of mice for microscopic studies as well as visualizing signaling molecules in histological samples. It is also difficult to analyze inflammation in the airway mucosa separately from the lung parenchyma: commonly measured immunological parameters (e.g., inflammatory cells and cytokines in the broncho-alveolar lavage fluid) reflect changes at the level of the whole bronchoalveolar space that might not be representative of the bronchial mucosa. Furthermore, there is a pronounced difference between the mouse and human intrapulmonary airways: in contrast to human bronchi, mouse intrapulmonary airways have no cartilage and lack several layers of the mucosa that are present in humans. Moreover, mice do not have bronchial arteries, and their blood supply relies on the pulmonary circulation (8, 9).

Overcoming these problems would allow us to better understand how each cell type finds its “designated” micro-anatomical position in the airway wall and how specific interactions among different cell types (e.g. DCs and effector CD4⁺ T cells) result in cellular activation, mediator production and local inflammatory cell (e.g., eosinophil) recruitment. Such information would also provide a markedly improved understanding of the cellular and molecular events underlying asthma pathogenesis.

Insight into the spatiotemporal aspects of immune effector function in a tissue context has been gained by the utilization of dynamic multiphoton intravital microscopy (MP-IVM) (10), multiplex immunohistochemistry (11, 12) and their combinations with functional read-outs (13-16). For example, such analyses in infected liver and antigen-challenged ear skin have shown that during T_H1-type responses, effector cytokine production by CD4⁺ cells depends on specific antigen recognition and TCR signaling that results in T-cell migration arrest and localized cytokine production (14, 17-19). In the context of asthma, the respective question is whether such a tight spatiotemporal control of T-cell cytokine production by local antigen presentation also applies for T_H2 or T_H17-type responses of the airway mucosa.

Here we developed a new approach to address the role of effector CD4⁺ T cells in organizing the allergic response of the airway mucosa. We combined in situ cytokine staining and confocal microscopy with IVM to accurately localize cytokine-producing CD4⁺ T cells, then studied the dynamics of their interaction with airway mucosal DCs. Furthermore, we examined the local consequences of effector cytokine production, ultimately leading to tissue eosinophilia and neutrophilia. An important feature of our approach was imaging the mouse trachea, because its structure and cellular composition more closely resembles human small airways involved in asthma, compared to the mouse small airways (20). Our detailed spatiotemporal analyses reveal that upon allergen challenge, there is a very localized, low-frequency activation of antigen-specific T cells to cytokine production in DC-T-cell clusters, and subsequent focal recruitment of myeloid effectors such as eosinophils to these focal sites of T-cell activation. These findings suggest that the inflammation driving asthmatic remodeling may occur in an episodic manner in restricted locations that over time comes to involve larger tissue areas, with possible therapeutic implications.

Materials and Methods

Mice

C57Bl/6J, CAG-ECFP, CAG-DsRed and TCRα^−/− mice were purchased from Jackson Laboratories. OT-II Rag-1^−/− mice were purchased from Taconic. CD11c-EYFP mice were a kind gift from Dr. Michel C. Nussenzweig (The Rockefeller University, NY) (21). NJ.1638 mice were obtained from the Laboratories of Drs. Nancy and Jamie Lee (Mayo Clinic, AZ) (22). IL-13 DsRed mice were kindly provided by Dr. William Paul (NIAID, NIH) (23). Both male and female mice were used at 6-12 weeks of age; mice in different experimental groups were matched for age and sex. Animals were maintained at the Central Animal Laboratory of the University of Turku or at an Association for Assessment and Accreditation of Laboratory Animal Care- accredited animal facility at NIAID. All animal procedures were approved by the Ethical Committee for Animal Experimentation in Finland and/or the Animal Care and Use Committee, NIAID, NIH. They were done in adherence with the rules and regulations of the Finnish Act on Animal Experimentation (62/2006) and were performed according to the 3R-principle (animal license number 5588/04.10.07/2014).

Allergen sensitization and challenge

For sensitization, mice were treated with 100 μg (dry weight) House Dust Mite (D. pteronyssinus, Greer Labs) and 1 μg cholera toxin (Sigma) in a volume of 20 μl via the oropharyngeal (o.ph.) route under isoflurane anesthesia (“HDM/CT-model”). In contrast to the commonly used volume of 50 μl, this smaller volume ensured that only a small amount reaches the lungs, while the rest is evenly distributed along the tracheal wall/mainstem bronchi. In some experiments, mice were challenged with 100 μg HDM o.ph. one week later. Alternatively, mice were sensitized with 100 μg HD M on d0 and challenged between days 7-11 with 10 μg HDM daily, all in volume of 20 μl o.ph., as described earlier (24) (“repetitive HDM-model”). Mice were analyzed on day 14.

Cell isolation and adoptive transfer experiments

For the generation of polyclonal “HDM-primed” T cells, CAG-ECFP or CAG-DsRed mice were o.ph. sensitized on d0 with HDM/CT. On d6, the lung-draining bronchial lymph nodes were harvested and CD4⁺ T cells were immunomagnetically isolated using negative selection (CD4⁺ T Cell Isolation Kit, mouse, Miltenyi Biotec). Typically, the purity of isolated cells was >97%. After washing twice with PBS, cells were i.v. injected into the tail vein of recipient mice. For the generation of OT-II effector cells, OT-II Rag-1^−/− mice were o.ph. sensitized with HDM/CT containing also 100 μg Endograde Ovalbumin (Hyglos) in a volume of 30 μl. On d6, a single-cell suspension of tracheas and lungs was prepared and leukocytes were enriched by density gradient centrifugation with Lympholyte M (Cedarlane Labs). CD4⁺ T cells were isolated via positive selection using CD4 (L3T4) MicroBeads (Miltenyi Biotec), yielding a purity of CD4⁺ effector T-cells typically >95%. T cells were subsequently labeled with 0.5 μM CellTracker Deep Red Dye (Life Techologies) in HBSS at 37°C for 10 min. Labeled cells were washed twice with cRPMI and twice with PBS. Finally, 1-2 × 10⁷ cells were i.v. injected into the tail vein of recipient mice 12-16h before in vivo imaging. Eosinophils were immuno-magnetically isolated from the spleens of NJ.1638 mice via negative selection. After incubation with CD90.2 and CD45R/B220 Microbeads, splenocytes were passed through a CS column (all from Miltenyi Biotec), resulting a typical purity of >90% Siglec-F⁺ SSC^high cells. Eosinophils were then labeled with 1 μM CellTracker Deep Red (or 2 μM TAMRA) and 2-5 × 10⁷ cells were injected via a tail vein catheter just before imaging.

Tissue collection and flow cytometry

Mice were i.p. injected with an overdose of pentobarbital sodium and once in deep anesthesia, 15 μg CD45-APC-Cy7 antibody (30-F11, BD Biosciences) were i.v. injected into the retro-orbital plexus to label intravascular leukocytes. Single-cell suspensions of tracheas and lung were prepared using Liberase TM, DNAse I, Collagenase XI and Hyaluronidase, as adapted from a previous protocol (25). The tissue digest was passed through a 100 μm cell strainer and washed twice with FACS buffer (PBS, 2% FCS, 0.01% NaN₃). Samples were blocked in the presence of anti CD16/32 (clone 2.4 G2) for 10 min, followed by labeling for 20 min with various conjugates of the following antibodies: CD3 (17A2, 1:100), CD4 (RM4-5, 1:400), CD11c (HL3, 1:200), CD44 (IM7, 1:200), CD127 (SB/199, 1:100), KLRG-1 (2F1, 1:100), Ly-6G (1A8, 1:200), Siglec-F (E50-2440, 1:200), TCR β (H57-597, 1:200) TCR γ:δ (GL3, 1:200), (all from BD Biosciences or BioLegend). Lineage cocktail contained CD3 (17A2, 1:100), CD5 (53-7.3, 1:400), CD8 (53-6.7, 1:200), CD11b (M1/70, 1:200), CD11c (HL3, 1:200), CD19 (1D3, 1:200), B220 (RA3-6B2, 1:200), NK1.1 (PK136, 1:100), TER-119 (1:200) and Gr-1 (RB6-8C5, 1:200), (all from BD Biosciences, eBioscience or BioLegend). PE-labeled CD1d Dextramer was obtained from Immudex. Just before analysis, a viability dye (7-AAD, 1:40, eBioscience) was added to each sample. Samples were analyzed using a BD Fortessa flow cytometer running FACS Diva 8.0.1, which was also used for data analysis. Dead cells and intravascular leukocytes were excluded from the final analysis.

Analysis of intracellular cytokines by flow cytometry

Direct ex vivo analysis of cytokines was performed as described previously (18). In vitro stimulation with antigen or PMA/ionomycin was done using a published protocol (26). After staining for surface antigens, samples were washed twice with FACS buffer and fixed with 4% paraformaldehyde for 20 min at 4°C. After two wash steps with FACS buffer, intracellular cytokine staining with antibodies against IL -13 (eBio13A, 1:50, eBioscience) and IL-17A (TC11-18H10, 1:50, BD Biosciences) or respective isotype controls was done using BD Perm/Wash buffer according to the manufacturer's instructions.

Tissue fixation and immunofluorescent staining

For the collection of tracheas, mice were i.p. injected with an overdose of pentobarbital sodium and once in deep anesthesia, perfused with 10 ml of ice-cold periodate-lysine-paraformaldehyde (PLP) buffer. In separate mice used for the collection of lung tissue, lungs were inflated with PLP-buffer via a tracheal cannula. Tracheas were “tethered” to a small piece of silicone (Sylgard, Dow Corning) with two insect pins and fixed overnight in PLP buffer. Thereafter, tracheas were washed and cut open on the ventral side while still pinned to Sylgard.

Whole-mount immunostaining and mounting for microscopy were performed as described earlier (27). Tracheas (or lungs) for frozen-section immunostaining were immersed in 30% sucrose in phosphate buffer overnight before snap-freezing in OCT (Tissue-Tek, Sakura). 20 μm-thick sections were air-dried and fixed with ice-cold acetone for 10 min. Immunostaining was performed using Shandon Sequenza immunostaining racks and coverplates (Thermo Fisher). Sections were blocked with 5% mouse serum (before primary antibodies) or 5% donkey serum (before secondary antibodies) in PBS/0.3% Triton-X-100 (blocking buffer) for 15 min, followed by incubation with directly conjugated or primary/secondary antibodies (diluted in blocking buffer), each for 1h at RT. Washes were performed using PBS/Triton-X-100.

For indirect immunostainings, primary antibodies were chicken anti-GFP (polyclonal, 1:500, Abcam), rabbit anti-RFP (polyclonal, 1:200, Rockland), rabbit anti-phospho-STAT6 (pTyr641) (polyclonal, 1:200, Sigma-Aldrich), rat anti-MHC-II (M5/114.15.2, BD Pharmingen). Secondary antibodies were species-selective F(ab’)₂-fragments raised in donkey, conjugated with Alexa Fluor 488, TRITC or Alexa Fluor 647 (all from Jackson ImmunoResearch). For in situ cytokine stainings, Alexa Fluor 488 conjugates of antibodies for IL-13 or IL-17A (the same as those used for flow cytometry) were used as primary antibodies at 1:50 dilution. These were followed by rabbit anti-Alexa Fluor 488 (polyclonal, 1:400, Life Technologies) as a second step label. In the third step, samples were labeled with a donkey anti-rabbit Alexa Fluor 488 F(ab’)₂-fragment.

Confocal microscopy, data visualization and image processing

Multi-channel XYZ image acquisition was performed on a Zeiss LSM 780 confocal microscope using 20x (air) or 40x (water immersion) objectives. Z-projections or combined XY, XZ and YZ representations were prepared using Imaris 8.1.2 (Bitplane). Confocal data involving p-STAT6 immunostains were processed as shown in Supplemental Figure 1.

Intravital two-photon microscopy and motility analysis

Mice were anesthetized with an i.p injection of 75 mg/kg Ketamine and 1 mg/kg Medetomidine. After shaving the ventral neck area, a 30G cannula, connected to a piece of PE 10 polyethylene tubing (Becton Dickinson) was inserted into the tail vein for the i.v. administration of reagents during imaging. Mice were then placed on a custom-built microscope stage in a supine position (Supplemental Figure 2A). The head was immobilized via slightly pulling the front teeth with an adjustable device. The trachea was surgically exposed and a custom-modified steel intubation cannula (Harvard Apparatus) was orotracheally inserted (with the exception of the experiment shown in Supplementary Video 2, which was performed without intubation). The cannula was secured in the trachea with an adjustable holding device. Finally, the exposed area was sealed with a custom-made “imaging window” (Supplemental Figure 2, B and C). This window was carefully positioned using a UM-3C coarse manipulator (Narishige) so that the coverslip mounted on it would touch the trachea, allowing the observation of 2 or 3 cartilage-free areas. A ring of vacuum grease was used to seal the gap between the skin and window. Importantly, in vivo imaging was performed on spontaneously breathing mice, and the tracheal wall was mechanically stabilized for imaging via holding it between the intubation cannula and the coverslip of the imaging window. Just before microscopic analysis, a drop of Immersol W immersion medium (Carl Zeiss) was applied on the window.

Multi-dimensional (XYZT) images were acquired using a Leica TCS SP5 MP two-photon microscope equipped with a Chameleon Vision II and a Chameleon Ultra II femtosecond laser (Coherent), using excitation at 865nm (for ECFP and CellTracker Deep Red) and 920 nm (for EYFP and TdTomato). An HCX IRAPO L 25x/0.95 W objective with a WD=2.5 mm was used to bridge the >2 mm level difference between the skin surface and the tracheal wall. For the simultaneous detection of ECFP, EYFP, DsRed/TdTomato and CellTracker Deep Red, the following band-pass filters were used: 483/32, 535/30, 585/40 and 650/50. Image processing for visualization (axial projections) and quantitative analysis were done using Imaris 8.1.2. Quantitation of cell motility was done using automated spot detection and track analysis, where the result of the automated detection was manually verified and corrected. Arrest coefficient was defined as the fraction of time points in which instantaneous velocity was < 2 μm/min, as described earlier (18). Exported movies were annotated with text and drawing elements using ImageJ Fiji.

Statistical analysis

Prism 6 (GraphPad Software) was used for the visualization and statistical analysis of quantitative data. The statistical significance of differences between the mean values of two groups was determined using Mann-Whitney test.

Results

The mouse trachea as a more relevant model to study allergic immune responses of the airway mucosa

It has been reported earlier that unlike the mouse intrapulmonary airways, the mouse trachea shares both micro-anatomical and physiological similarities with human small bronchi, the anatomical sites primarily affected by asthma (28). Although the trachea of mice is clearly very different from human small peripheral airways (e.g. while the mouse trachea has cartilage rings as shown in Figure 1A, human bronchi have cartilage plates/islands), there are several similarities, such as a comparable internal diameter (~1.2 mm), a complex pseudostratified epithelium (Figure 1, B, C) with basal cells, submucosal glands (both of which are lacked by mouse bronchi), as well as “real” arterial circulation (9, 20, 28). The trachea has been used earlier to visualize DC and mast cell dynamics in response to allergen challenge ex vivo (29), and it was previously demonstrated that this is achievable also in vivo (30). Therefore we have established a combined histological and intravital microscopic approach to study immune responses at this anatomical site. Using MP-IVM, we typically observed the anterior wall between two cartilage rings (Figure 1, A, D, E, F and Supplemental Video 1), allowing us to visualize in living, anesthetized mice such events as the dynamic probing movement of DCs within and beneath the epithelium (Supplemental Video 2) as well as blood flow in mucosal post-capillary venules (Supplemental Video 3).

Figure 1.

The mouse trachea as a model to study immune responses of the airway mucosa. (A) Stereomicroscopic image of an explanted trachea. The box shows a region between cartilage rings typically accessed by intravital microscopy (bar=200μm). (B, C) Confocal microscopic images of a tracheal cross-section of a CAG-DsRed x CD11c-EYFP mouse showing the epithelium (block arrow) and underlying DCs (arrows). Nuclei are visualized by To-Pro-3 staining. Bars=100μm. (D-F) Two-photon microscopic Z-stack showing the epithelium (E) and subepithelial (F) tissue. (E, F) lower panels: cross-sectional views. Regions between the dashed lines are shown in upper panels as Z-projections (bars=100μm). (D) Enlargement of the boxed region shown in E. Arrows indicate epithelial DCs (bar=40μm). Tracheas from two mice were used in a single experiment.

With this imaging capacity in hand, we searched for an appropriate model of airway mucosal inflammation in which exposure to a clinically-relevant allergen would produce a similar type of cellular infiltrate in the tracheal mucosa as is observed in humans, i.e., one dominated by CD4⁺ T cells and eosinophils/neutrophils. House dust mite (HDM) has been an increasingly well-characterized model allergen due to its clinical relevance and the properties it shares with several other environmental allergens such as protease activity (31), agonist activity for pattern recognition receptors (PRRs) (32) and an ability to provoke a positive skin-prick test in asthmatics (33). In prior studies using HDM as a model allergen, multiple exposures were necessary to produce an allergic phenotype (5, 24). However, repetitive exposures “blur” the line between primary sensitization and challenge (34), and although such models can be used to induce an asthma-like inflammatory response, they do not allow studying the temporal sequence of cellular recruitment and activation as it occurs after single allergen encounter. To overcome this problem, we developed a system that is based on the single-bolus administration of HDM in combination with the mucosal adjuvant cholera toxin (CT), which was used in earlier studies to induce an airway mucosal T_H2-type response (35). This approach allowed us to perform a kinetic study on cellular recruitment to the trachea and select optimal time points for imaging analysis. Sensitization of C57Bl/6 mice with HDM/CT via the oropharyngeal (o.ph.) route induced a tracheal (and lung) T-cell response that developed from d4 post-sensitization, and was followed by an eosinophil response (Figure 2A). By d8 we observed tracheal mucosal T-cell clusters, infiltrated with eosinophils (Figure 2B). A careful analysis of tracheas (and lungs) showed that CD4⁺ cells dominated the CD3⁺ T-cell response (Figure 2, C and D), without the presence of CD8⁺ cells. However, a few γδ T cells and some NKT cells were present in the CD3⁺CD4⁻ population (data not shown). We also observed that the eosinophilia (Figure 2, E and F) co-emerged with a DC and neutrophil response (Figure 2, E, G, and H). The key components of the tracheal and lung response at d8 post-sensitization with HDM/CT (e.g. the recruitment and co-clustering of DCs, CD4⁺ T cells and eosinophils) were similar to the response that developed in a more standard, adjuvant-free model after repetitive HDM administration, although neutrophilia did not develop in that standard model (Supplemental Figure 3, A-F). However, in contrast to the standard model, the new model also enabled us to capture the early phases of discrete inflammatory cluster formation in the tracheal mucosa, which was necessary for establishing the setup shown in Figure 7, as discussed later.

Figure 2.

Induction of a tracheal mucosal inflammatory response to HDM allergen. (A) Percentages of T cells (blue) and eosinophils (red) from tracheal and lung tissue digests on days 2-12 following a single o.ph. treatment of C57Bl/6 mice with HDM/CT. Gated on live extravascular cells. Lung eosinophils were distinguished from alveolar macrophages based on their low autofluorescence in the green (AF 488) channel. (B) Confocal images showing T-cell clusters and associated eosinophils in a tracheal whole-mount at d9 after HDM/CT. Boxed region in left panel (bar=200μm) was enlarged (right panel, bar=20μm). (C, E, G) Comparison of lung and tracheal tissue digests from HDM/CT and PBS-treated mice. Gates indicate the CD4⁺ T-cell response at d8 (C), eosinophil and DC response at d10 (E) and neutrophil response at d8 (G). Numbers show percentages from the total live extravascular population. (D, F, H) Analysis of the tracheal inflammatory response developing between d6-d10. Total numbers of CD4⁺ T cells (D), eosinophils (F) and neutrophils (H) per trachea were determined using the gates shown in C, E and G. Graphs indicate mean ± SEM. n=2-3 mice per time point (in a single experiment) were analyzed.

Figure 7.

STAT6 phosphorylation and myeloid effector cell recruitment in the tracheal mucosa are associated with CD4⁺ T-cell clusters. “HDM-primed” polyclonal CD4⁺ T cells, isolated from the bronchial lymph node of CAG-DsRed mice, were adoptively transferred into TCRα^−/− mice on d-7. At d0, recipients received HDM/CT o.ph. and at different time points, perfusion-fixed tracheas were collected for whole-mount confocal analysis. (A-C): Analysis of p-STAT6 distribution at d3. (A) Co-localization of a p-STAT6⁺ cell cluster with a cluster of DsRed⁺ polyclonal effector T cells (block arrows indicate the clusters). (B) Lack of bright p-STAT6⁺ cells in mice without transferred T cells (arrows indicate a few dim p-STAT6⁺ cells). Images shown in A and B were acquired using the same laser intensity and detector settings (bars=50 μm). (C): XY, XZ and YZ representations of a subepithelial effector cluster (slicer positions are indicated by tick marks). The tracheal epithelium is shown between dashed lines. Arrows show bright p-STAT6⁺ cells. DsRed⁺ p-STAT6⁺ cells were excluded from the final images.

(D-F): Mice were challenged on d7 and on d9, perfusion-fixed tracheas were collected for whole-mount confocal analysis of eosinophil and neutrophil accumulation. (D): Localization of Siglec-F⁺ eosinophils in a DsRed⁺ CD4⁺ effector cluster. Boxed region in the low-magnification image (left panels, bar=100 μm) has been enlarged (right panel) and shown as XY, XZ and YZ representations (a range of slices is shown, as indicated by tick marks). (E): Upper panels: low-magnification image of two DsRed⁺ CD4⁺ effector clusters (dashed lines) showing the partial or complete co-localization of CD11b⁺ neutrophils and Siglec-F⁺ eosinophils, respectively (bar=100μm). Lower panels: high-magnification image of a DsRed⁺ CD4⁺ effector cluster showing the positioning of both eosinophils and neutrophils within the clusters (bar=20 μm). (F): Quantitation of eosinophil vs. neutrophil localization within T-cell clusters. Six clusters (n=6) were analyzed in 3 tracheas (**:p < 0.01 as determined using Mann-Whitney test). All images are representative of at least two independent experiments.

Visualization of effector CD4⁺ T-cell cytokine responses

Classically, the asthmatic response is dominated by T_H2-type CD4⁺ cells and tissue eosinophilia. However, recent evidence suggests that more severe, corticosteroid-resistant forms of asthma are associated with a T_H17 response and neutrophils (3). Our model system had components of both disease types: we observed a mixed eosinophil/neutrophil response in the mouse airways and the effector CD4⁺ T cells in the tracheal mucosa showed a mixed T_H2/T_H17 phenotype, as indicated by a prominent IL-13 or IL-17 response of these cells upon in vitro re-stimulation (Figure 3A). When compared with the standard, adjuvant-free model, we found a similar IL-13 but increased IL-17 response, together with lower IFN-γ production (Supplemental Figure 3G). Although both models showed a mixed T_H1/T_H2/T_H17 response, HDM/CT sensitization resulted in a decreased T_H1 and increased T_H17 component, possibly due to the effect of CT (36). This increased T_H17 response was presumably responsible for the development of neutrophilia, which allowed us to compare the mucosal positioning of neutrophils and eosinophils during cluster formation (see below). Although a T_H1 response was present in both models, previous research suggested a key role of T_H2 and T_H17 cells in asthma, therefore, we focused on IL-13 and IL-17 to localize individual cytokine-producing CD4⁺ T cells in tissue.

Figure 3.

Visualization of effector cytokine production by CD4⁺ T cells in the trachea. (A) C57Bl/6 mice were treated with HDM/CT o.ph. and 8 days later, effector cytokine production by CD4⁺ T cells from digested tracheas was analyzed after re-stimulation with PMA/Ionomycin or treatment with Brefeldin A (BFA) only. Numbers indicate percentages. (B) C57Bl/6 mice were treated with HDM/CT o.ph. on d0 and received HDM o.ph. challenge on d7. Tracheas were analyzed on d9. Confocal microscopic analysis of cytokine-producing cells in tracheal frozen sections. Boxed regions were enlarged to show individual cytokine-positive cells (left panels, bars=50μm, right enlarged panels, bars=20μm, asterisk shows the tracheal lumen). Data are representative of at least two independent experiments. (C, D) Analysis of T-cell cytokine production directly after tissue digest (C) or after in vitro re-stimulation (D). 3 tracheas were pooled for each measurement; numbers indicate percentages (isotype control vs. specific anti-cytokine staining was done using the same digest). Plots were gated on extravascular, CD4⁺CD44^hi cells. (E, F) Quantitative representation of T-cell cytokine production measured directly ex vivo 12h after HDM challenge (E) or after in vitro re-stimulation (F). Data were pooled from two independent experiments; each dot indicates a single measurement. n=5 measurements for E and n=4 measurements for F; three tracheas were pooled and used for each measurement (*:p < 0.05; **:p < 0.01 as determined using Mann-Whitney test).

Initially, we visualized cytokine-producing cells using tracheal and lung frozen sections from HDM/CT-sensitized and HDM-challenged mice. A secondary allergen challenge was performed to ensure that antigen is abundantly available at the time of analysis. Staining for IL-17 and IL-13 revealed several clearly identifiable T cells in the subepithelial lamina propria of the trachea with the presence of these cytokines within delimited cytoplasmic structures (Figure 3B). In the lungs, we found such prominent cytokine-positive cells both in the peribronchial areas as well as in the lung parenchyma (data not shown).

Next, we performed a detailed phenotypic analysis of in situ cytokine-producing cells. It has been reported that in an HDM-induced pulmonary response, ILC2s significantly contribute to T_H2-cytokine production (37). Since ILCs appear as CD3⁻cytokine⁺ cells, we first measured the frequency of such cells among IL-13 or IL-17 positive cells in tissue sections. The vast majority of cytokine-positive cells were CD3⁺; the frequency of non-T-cell cytokine producers ranged between 0-2% both in the trachea and lung (Table 1). We also performed a more detailed flow cytometric analysis to compare the overall numbers and intracellular cytokine levels of CD4⁺ T cells vs. ILC2s in the trachea and lung (Supplemental Figure 4, A-D). Our findings suggest that in this experimental model, the major producers of IL-13 and IL-17 are CD3⁺ T cells, whereas the in vivo contribution of ILCs to cytokine production is quite limited. Although conventional CD4⁺ T cells are the most likely cytokine producers within the CD3⁺ population, earlier reports also suggest an important role for IL-13 producing NKT cells (38) and IL-17 producing γδ T cells (39) during the asthmatic response. We found that the frequency of both NKT cells and γδ T cells was very low, compared to conventional CD4⁺ T cells (data not shown) and the contribution of CD1d-restricted iNKT cells to the IL-13 response was minimal (Supplemental Figure 4, E-G). A detailed analysis of prominent IL-13⁺ and IL-17⁺ cells in tracheal sections also confirmed that these cells were mainly conventional, CD4⁺ αβ T cells (Supplemental Figure 4, H and I). We verified this finding also using the adjuvant-free, repetitive HDM model. Although in that model we could not visualize IL-17⁺ cells, the vast majority of IL-13⁺ cells were CD3⁺CD4⁺ conventional T cells (Supplemental Figure 3H).

Table 1.

Quantitative analysis of CD3⁺ and CD3⁻ cytokine-producing cells in tracheal and lung tissue sections¹

n=3 mice	% of IL-13+ cells (out of total CD3+ T cells) ± SEM	% of IL-17+ cells (out of total CD3+ T cells) ± SEM	# of CD3− cells / total IL-13+ cells (percent)	# of CD3− cells / total IL-17+ cells (percent)
Trachea	1.59 ± 0.11%	1.06 ± 0.24%	1/53 (1,89%)	0/48 (0%)
Lung	0.98 ± 0.30%	1.93 ± 0.51%	4/285 (1.4%)	8/565 (1.42%)

¹C57Bl/6 mice were treated with HDM/CT o.ph. on d0 and received HDM o.ph. challenge on d7. Tracheas and lungs were analyzed on d9. IL-17⁺ or IL-13⁺ cells were manually identified and co-staining for CD3 was determined. Tissue samples from 3 mice were analyzed.

It is important to note that the frequency of cytokine-positive cells among all T cells was altogether very low (typically 1-2%) both in the trachea and lung. These numbers are clearly below the values commonly measured when T-cell IL-13 and IL-17 responses are studied using in vitro re-stimulation (typically varying between 10-30 %) (40, 41), and they suggest that during an airway mucosal effector CD4⁺ T-cell response to inhaled allergen, only a few percent of all effector cells produce cytokines at a given time. This same phenomenon has been previously reported in liver and lung mycobacterial infection models (18, 42). To address this issue in more detail, we performed flow-cytometric analysis of tracheal T-cell effector cytokine production (Figure 3, C-F). Direct ex vivo analysis at d8 after initial HDM/CT sensitization confirmed the low in vivo frequency of cytokine-producing cells both in the trachea and lung. In response to a secondary in vivo HDM challenge, as well as after in vitro re-stimulation with HDM, we measured a ~3-fold increase in the frequency of IL-17⁺ cells, whereas the increase of IL-13⁺ cells was less pronounced. However, all these values were clearly below those measured after a maximal stimulus, represented by PMA/Ionomycin (P/I) re-stimulation. Altogether, these data suggest that in our model, the allergic response to HDM is mainly driven by conventional CD4⁺ T cells, with only a small fraction of them being activated to cytokine production at any given time.

Cytokine-producing CD4⁺ T cells form conjugates with APCs in the airway mucosa

Previous studies on T_H1 effector CD4⁺ responses in the liver and ear skin using a TCR-transgenic system suggest that cytokine production by CD4⁺ cells depends on prolonged interaction with APCs (18, 19). Therefore, we asked whether the production of IL-17 and IL-13 by endogenous, polyclonal effector T cells in the tracheal mucosa also depends on local antigen presentation. Whole-mount immunostains for IL-17 enabled us to carefully localize IL-17 producing T cells within T-cell clusters within the tracheal mucosa (Figure 4A). In contrast, IL-13 staining only worked well on thick frozen sections, and therefore we could only study the local environment of individual cells, without a proper view on the larger tissue context.

Figure 4.

Cytokine-producing T cells are engaged in contact with APCs. C57Bl/6 mice were treated with HDM/CT on d0 and re-challenged with HDM on d7. On d9, perfusion-fixed tracheas were collected for confocal microscopic analysis. (A, B) Visualization of an effector T-cell/DC cluster in a tracheal whole mount. Boxed regions in A (upper panels, bars=50μm) were enlarged (lower panels, bars=10μm) and displayed as XY, XZ and YZ representations (B) to show an IL-17⁺ T-cell in direct contact with an APC (arrow). (C, D) Visualization of an effector T-cell/DC cluster in a frozen cross-section of the tracheal mucosa. Boxed region in C (left panel, bar=50μm) is shown also as separate channels (right panels, bars=10μm) and displayed as XY, XZ and YZ representations (D) to indicate an IL-13⁺ T cell in direct contact with an APC (arrow). B and D show single slices, the position of slicers is indicated by tick marks. (E, F) Quantitative analysis showing % of cytokine-positive vs. cytokine-negative cells in contact with an APC. Between 74-104 tracheal cytokine-positive or randomly selected cytokine-negative T cells were analyzed; n=7-8 mice for E, n=6 mice for F (*:p < 0.05; ***:p < 0.001 as determined using Mann-Whitney test). Results of a single experiment are shown.

During an HDM/CT-induced tracheal mucosal response, CD3⁺ T-cell clusters appeared together with APC clusters. Within these cellular clusters, a few IL-17⁺ T cells were localized in the immediate proximity of MHC-II⁺ cells (Figure 4, A and B). Similarly, IL-13⁺ T cells appeared in close contact with MHC-II⁺ structures, as seen on tracheal cross-sections (Figure 4, C and D), an observation that we could confirm also in the adjuvant-free, repetitive HDM model (Supplemental Figure 3, I and J). Quantitatively, we found that the vast majority of both IL-17⁺ and IL-13⁺ T cells were in direct contact with MHC-II⁺ cells (Figure 4, E and F). However, we could also identify a low number of cytokine-producing cells outside of APC clusters, clearly without contact to MHC-II⁺ cells (data not shown). These results suggest that direct cell-cell contact with local MHC-II bearing cells of the airway mucosa is largely responsible for the elicitation of effector cytokine production by T_H2 and T_H17 cells.

CD11b⁺ DCs represent the major DC population serving as APCs to induce local effector cytokine production in airway mucosal CD4⁺ T cells

Like in most non-lymphoid organs, myeloid cDCs of the airway mucosa belong to one of the two major DC subtypes: CD11b⁺CD103⁻ or CD11b⁻CD103⁺ DCs (here referred to briefly as CD11b⁺ and CD11b⁻ cDCs respectively) (43). In HDM-induced allergic responses, large numbers of CD11b⁺ monocyte-derived DCs are recruited to the airways (44). Previous work revealed that CD11b⁺ cDCs and monocyte-derived DCs are mainly involved in the initiation and maintenance, respectively, of T_H2-immunity to HDM allergen (44). Therefore we tried to determine which DC subtypes are involved in local antigen presentation (defined here as direct contact with cytokine-positive T cells). Although we were histologically unable to differentiate between CD11b⁺ cDCs and CD11b⁺ monocyte-derived DCs, we could clearly separate CD11b⁺ DCs from CD103⁺ DCs by whole-mount immunostaining for CD11b in CD11c-EYFP mice. We observed that within large subepithelial DC clusters that are formed during the inflammatory response, the majority of CD11c-EYFP⁺ DCs co-stained for CD11b. Cytokine-producing T cells were interlaced within clusters of amoeboid DCs possessing a few dendritic extensions, in direct contact with CD11b⁺ CD11c-EYFP⁺ cells (Figure 5, A and B). However, mainly within the epithelium, we could also observe individual CD11b⁻ (and presumably CD103⁺) CD11c-EYFP⁺ DCs with a dendritic morphology in contact with cytokine-positive T cells (Figure 5, C and D). Quantitatively, we found that the majority of both IL-17 and IL-13 positive T cells formed contacts with CD11b⁺ DCs (Figure 5E). However, this probably reflects the increased frequency of this DC subtype in the inflamed tracheal mucosa (Figure 5F), rather than their increased propensity to activate T cells. Therefore, our histological findings are in accordance with earlier reports that suggest that CD11b⁺ cDCs and monocyte-derived DCs are the major antigen-presenting DC subtypes in the airways (44).

Figure 5.

Both CD11b⁺ and CD11b⁻ DCs form contact with cytokine-producing effector T cells in the tracheal mucosa. CD11c-EYFP mice were treated with HDM/CT on d0 and re-challenged with HDM on d7. On d9, perfusion-fixed tracheas were collected for whole-mount immunostaining. (A-D) Confocal microscopic analysis of interactions between IL-17⁺ T cells and CD11b⁺ (A, B) or CD11b⁻ (C, D) DCs. Bars=20μm, arrows indicate IL-17⁺ T cells in contact with DCs. In C, double arrow shows a CD11b⁻ DC, block arrow shows a CD11b⁺ DC. A and C show separate channels, B and D show the respective merged images as XY, XZ and YZ representations. In B and D, a range of slices is shown, as indicated by tick marks. (E) Quantitative analysis showing the contribution of CD11b⁺ vs. CD11b⁻ DCs to forming conjugates with cytokine-positive T cells in the tracheal mucosa. A total of 68 IL-17⁺ T cells in contact with a CD11c-EYFP⁺ DC in tracheal whole mounts of 5 mice were analyzed. A total of 49 IL-13⁺ T cells in DC-contact in tracheal frozen sections of 5 mice were analyzed. (F) Relative frequency of CD11b⁻ vs. CD11b⁺ DCs in the tracheal mucosa. Tracheal whole mounts of 5 mice were analyzed. Results of a single experiment are shown.

Effector CD4⁺ T cells form stable contact with DCs in the airway mucosa

Given the previously demonstrated link between antigen recognition, motility and cytokine production by TCR-transgenic cells during T_H1-responses in the liver and skin (18, 19), we sought to determine if prolonged T-DC interactions also precede cytokine production in the airway mucosa in course of a polyclonal T_H2/ T_H17 response to HDM allergen.

To visualize the polyclonal effector CD4⁺ T-cell response in the trachea in vivo, we transferred bronchial lymph node polyclonal CD4⁺ cells from HDM/CT-sensitized, ubiquitously DsRed or ECFP-expressing mice into CD11c-EYFP Rag-1^−/− recipients (Figure 6A). Treatment of recipient mice with HDM/CT resulted in the recruitment of the transferred cells to the tracheal mucosa (Figure 6B). At d7 post-sensitization, another HDM challenge ensured abundant antigen availability. Using MP-IVM two days later, we were able to observe cellular interactions within DC-T-cell clusters in real time (Supplemental Video 4). Within these clusters, DCs formed stable interactions with the transferred “HDM-primed” polyclonal effector T cells (Supplemental Video 5).

Figure 6.

Polyclonal “HDM-primed” T cells, but not OT-II cells arrest on DCs and produce effector cytokine. (A) Adoptive transfer and sensitization protocol. (B) Flow-cytometric analysis of tracheal tissue digests to compare the tracheal polyclonal CD4⁺ response in HDM/CT sensitized vs. control (PBS-treated) mice. Analysis of recipient mice was performed at d8 post-sensitization. Numbers indicate percentages. (C) Snapshot from Supplemental Video 6 (left panel, bar=100 μm) and representation of corresponding cell motility tracks (right panel). (D) Quantitative comparison of mean velocity and arrest coefficient of “HDM-primed” polyclonal vs. OT-II effector cells. Circles indicate individual cell tracks; horizontal white bars show mean values. Data were pooled from the analysis of three mice, which were performed as three independent experiments. (***:p < 0.001 and ****:p < 0.0001 as determined using Mann-Whitney test). (E) Whole-mount IL-17 staining of tracheas collected after MP-IVM analysis. Boxed region in left panel (bar=50μm) was enlarged (right panel, bar=10μm) to indicate IL-17 production by polyclonal T cells (arrow) but not by OT-II cells (double arrow). 20 IL-17⁺ cells were analyzed in the tracheas of two mice (from two independent experiments).

Next we examined whether these stable T-DC interactions form in an antigen-specific manner. To this end, we i.v. injected in vivo-generated OT-II Rag-1^−/− CD4⁺ effector cells the day before analysis (Figure 6A). These cells served as a control population with known specificity to an irrelevant antigen (OVA). It has been reported earlier that effector T cells enter inflamed tissues even in the lack of specific antigen (18). Similarly to previous observations in the liver and skin, injected OT-II effector cells were recruited to the inflamed tracheal mucosa and showed a pattern of random migration (Figure 6C and Supplemental Video 6). However, in the absence of OVA, these cells did not stop on DCs and showed a greater mean velocity as well as a smaller arrest coefficient as compared to the polyclonal “HDM-primed” population (Figure 6D). In situ cytokine staining of tracheas fixed after MP-IVM demonstrated that only the polyclonal cells, but not OT-IIs produced effector cytokines (Figure 6E). These results suggest that the in vivo observed stable interactions of polyclonal effector T cells with airway mucosal DCs and local cytokine production are the result of specific antigen recognition and not cytokine-induced cytokine production by bystander lymphocytes.

T_H2 cytokine signaling and eosinophil recruitment are localized to effector CD4⁺ T-cell clusters

As the majority of T-DC interactions leading to effector cytokine production occur within discrete clusters, our data suggest that such “allergic lesions” might function as “hot-spots” driving the allergic response. As a next step, we asked 1) to what extent are the local effects of T_H2 (and T_H17) cytokines restricted to these cell clusters and 2) whether the recruitment of myeloid effector cells (eosinophils and neutrophils) associated with a T_H2/T_H17 response is locally restricted to these clusters. To address these issues, we used the adoptive cell transfer approach described previously (see Figure 6A), involving the transfer of polyclonal “HDM-primed” CD4⁺ T cells into Rag1^−/− mice. In this setting, tracheal (and lung) eosinophilia was completely dependent on the presence of the “HDM-primed” polyclonal T-cell population (Supplemental Figure 5, A and B). Neutrophilia also greatly depended on T cells; however (at least in the trachea), neutrophils were present in low numbers even in the absence of T cells. We obtained similar results when TCRα^−/− mice were used as recipients instead of Rag1^−/− mice (results not shown). Therefore, in the following experiments, we transferred DsRed⁺ “HDM-primed” polyclonal CD4⁺ T cells into TCRα^−/− mice to visualize the local consequences of effector T-cell cytokine production.

IL-13 (together with IL-4) is known to induce STAT6 phosphorylation in target cells (45), so we first performed immunostaining for STAT6 phosphorylated on Y641 (p-STAT6) to reveal locally responding cells. In response to HDM/CT, DsRed⁺ polyclonal T cells formed discrete small clusters in the tracheal mucosa on d2-d4 post-sensitization. By day 7, the numbers of recruited cells increased and cell clusters became larger, with scattered cells distributed between these clusters. To determine if the effect of IL-13 is locally restricted, we visualized early, small effector DsRed⁺ CD4⁺ clusters together with p-STAT6 staining (Figure 7, A-C). We observed that the DsRed⁺ effector cells were among the brightest p-STAT6⁺ cells and therefore, via image processing, we removed all p-STAT6 signals that originated from DsRed⁺ cells (as shown in Supplemental Figure 1). We found that clusters of bright p-STAT6⁺ cells completely overlapped with DsRed⁺ effector clusters (Figure 7A). However, in control tracheas from HDM/CT sensitized TCRα^−/− animals that did not receive polyclonal T cells, only very few, dim p-STAT6⁺ cells were present and these did not form clusters (Figure 7B). A closer examination of DsRed⁺ effector clusters revealed individual, bright p-STAT6⁺ “locally responding” cells within and around subepithelial T cell clusters (Figure 7C).

Since our results suggested that the effect of CD4⁺ T-cell cytokines is concentrated within the clusters they form, we next asked whether the T-cell dependent recruitment of eosinophils and neutrophils occurs in a local manner. We observed earlier that eosinophilia develops at later time points (typically d8-d12 after HDM/CT). To ensure the detection of small, separate clusters at these late time points, we initially injected a smaller number of CD4⁺ polyclonal effectors (1×10⁵/mouse instead of 0.5 – 1 × 10⁶ that was used for analysis at early time points). In line with our expectations, eosinophil recruitment was locally restricted to these late effector clusters (Figure 7D). We further verified this finding by injecting a bolus of fluorescently labeled eosinophils and visualizing their tracheal localization on the next day. We found that the injected cells also localized into both early (d3, see Supplemental Figure 5C) and late (d9, see Supplemental Figure 5D and Supplemental Video 7) effector clusters.

In contrast to the local accumulation of eosinophils, neutrophils were more evenly distributed in the tracheal mucosa and did not show such prominent clustering with T cells (Figure 7, E and F). However, CD4⁺ effector clusters also contained a large number of neutrophils, which suggests that both T_H2 and T_H17 cells participate in their formation.

Discussion

In asthmatics, allergic inflammation mainly develops within the mucosal layer of small-diameter bronchi. Upon repeated allergen challenge, the prolonged activation of T_H2 and T_H17 cells causes, via the release of effector cytokines, tissue eosinophilia and neutrophilia, and ultimately leads to tissue damage, airway re-modeling and bronchial obstruction. To explore how this complex sequence of cellular activation and migration events is spatially and dynamically organized, we established an in vivo dynamic / ex vivo static imaging-based model of small-airway allergic disease. The key features of our ‘disease-relevant’ approach include 1) focusing on the mouse trachea, due to its similarities to small human bronchi (from which mouse small airways are very different) and 2) characterizing a polyclonal (and not TCR-transgenic) T-cell response to a clinically relevant allergen (HDM). These features represent an important advance and distinguish our work from prior studies exploring T-DC interactions in the peripheral lung/intrapulmonary airways of mice by using ex vivo slice preparations with TCR-transgenic OTII-cells in response to OVA (46).

The notion that effector CD4⁺ responses to inhaled antigen occur in discrete cellular foci has been suggested earlier (47). Our results clearly demonstrate that the initial recruitment of effector T cells and the formation of clusters in the airway mucosa is accompanied by the local accumulation of antigen-presenting DCs (Figure 4A). At later time points, we also observed the local recruitment of eosinophils and neutrophils, which complete the “loop” of a typical adaptive T_H2/T_H17 immune response. The formation of clusters that contain all key cellular players of an allergic response might represent a general feature of the immune system, which has evolved to localize the cellular response for the containment of helminthes and parasites. Similarly, T_H1-responses may also develop in the form of discrete cellular clusters, as observed e.g. in models of mycobacterial infection (granuloma formation) (17, 18).

The relatively low (1-2%) in vivo frequency of cytokine-positive T cells (as detected both by immunohistology and flow cytometry) reflects the rate at which effector T cells normally become activated in course of a polyclonal, effector CD4⁺ T_H response. This is in apparent contrast to values that have been commonly measured in T_H cells ex vivo, typically after re-stimulation with P/I, anti-CD3 or antigen (frequencies up to 30-40%). Although the fraction of cytokine⁺ cells increased after secondary antigen administration both in vivo and in vitro, it remained clearly below those values detected following P/I. This could be explained by the asynchronous activation of polyclonal effector T cells in vivo, which is subject to the availability of specific antigen, the previously described process of rapid desensitization following activation (19) and other tissue factors yet to be explored.

In contrast, antigen non-specific stimuli (e.g. with P/I) induce a synchronized activation of all cells, which does not necessarily reflect the rate at which this happens in vivo. The importance of such direct in situ/ex vivo analyses of intracellular cytokine content becomes obvious especially in the accurate identification of the major cellular source of effector cytokines. It has been reported that ILC2s represent a major source of IL-13 producing cells in an HDM-induced allergic response, as measured after P/I stimulation (37). However their contribution to IL-13 production in our model was minimal. This was also supported by the finding that eosinophilia developed in a T-cell dependent manner, as observed here as well as by others (48).

Our finding that CD4⁺ T cells are the major source of local cytokine production, with only a minor involvement of ILC2s slightly differs from previous observations made in a papain model of asthma (49, 50) or another study using repetitive HDM exposure (51), both of which suggest an important role for ILC2s in terms of cytokine production and the initiation of a T_H2-type adaptive response. In this respect, our study might not “accurately” model how HDM normally induces initial sensitization, i.e. via the epithelial induction of IL-33, IL-25 or TSLP, which induce ILC2 cytokine production and promotes adaptive T_H2-responses. Instead, we might bypass this “epithelium-ILC2 pathway” by the use of CT (as discussed later) and also induce a strong T_H17 (and T_H1) adaptive response, for which ILC2s are dispensable (51). Due to this limitation, our study is not directly comparable to previous work on how protease allergens elicit an allergic response; instead we provide an alternative, imaging-based approach to visualize the effector function of polyclonal, mixed T_H2/T_H17 CD4⁺ T cells in the complex airway mucosal tissue in response to HDM challenge, using a protocol that has been developed and adjusted for our specific imaging purposes.

The vast majority of cytokine⁺ T cells were engaged in direct cell-cell contact with MHC-II⁺ cells, which suggests that cytokine gene activation and cytokine production in effector T cells is triggered by the recognition of specific antigen presented by local DCs , mainly of the CD103⁻ CD11b⁺ subtype. This notion is also supported by our in vivo imaging experiments showing prolonged interactions between “HDM-primed” polyclonal T cells with DCs, whereas control, OVA-specific OT-II effector cells that were initially primed under the same conditions as the polyclonal effectors, fail to form stable interactions with DCs or produce cytokine. In light of the previous observations made in liver (18) and skin (19) models of T_H1 effector responses, our results suggest that the requirement of specific antigen recognition and prolonged interaction with antigen-presenting DCs for local cytokine production also applies to polyclonal T_H2/T_H17 responses of the airway mucosa. After dissociation from DCs, increasing T-cell motility correlates with an immediate decrease in cytokine production (19). We also observed a very low number of cytokine⁺ T cells without direct contact to DCs, however these were always within a few 10 microns away from a neighboring MHC-II⁺ cell (data not shown), suggesting that for a limited time period, T cells are capable of producing cytokines even after dissociating from a DC.

Lately, it has been a matter of intense debate whether effector cytokines are just locally secreted or widely distributed in complex tissues. Within lymph nodes, locally produced T_H1 and T_H2 cytokines have been reported to reach the majority of cells (52). However, in peripheral tissues, cytokine effects might be more locally restricted. Previous studies showing an inverse correlation between motility and cytokine production (18, 19) clearly favor the model according to which the ‘radius of action’ for T-cell cytokines is limited to about 80-100 μm, (14, 53) thereby reaching only neighboring cells. Our model did not allow an exact measurement of the distances at which locally released T_H2 cytokines exert their effect (detected as STAT-6 phosphorylation), since there were always a few scattered CD4⁺ cells, as well as p-STAT6⁺ cells present in areas between the prominent clusters. However, local accumulations of p-STAT6⁺ cells perfectly overlapped effector clusters, suggesting that the locally restricted model of cytokine effects also applies for airway mucosal T_H2 responses.

IL-13 is a key mediator of allergic asthma, and its effects through the IL-13/IL-4/STAT-6 signaling pathway have been well characterized (54). One of the main functions of IL-13 is the induction of tissue eosinophilia via induction of eotaxin production, which promotes the recruitment of eosinophils from the bloodstream (55). In our model, eosinophil accumulation followed the pattern of STAT-6 phosphorylation, i.e. the initial clusters of newly arriving eosinophils preferentially localized to CD4⁺ effector clusters. This finding suggests that the effector arm of the allergic response is concentrated around the site of initial antigen encounter and T-cell activation.

It is important to note that the use of CT as an adjuvant greatly helped us to perform imaging analysis by inducing a robust primary T_H2/T_H17 CD4⁺ (but not CD8⁺) response to HDM, revealing the kinetics of cluster formation and allowing the side-by-side analysis of eosinophil and neutrophil positioning in the tracheal mucosa, which occurred in a T-cell dependent manner. However, CT is known to engage relatively unique pathways (56), potently activate CD11b⁺ DCs (36) and skew the immune response to HDM towards T_H17 cells (57). While we could confirm some of the main findings (e.g. T-DC cluster formation with local eosinophil recruitment and low-frequency cytokine production by T cells while in contact with DCs) also using repetitive HDM administration without CT (Supplemental Figure 3), both models reflect changes during a very acute primary or secondary response to allergen. However, asthma is a chronic disorder and to better understand the airway mucosal response in chronic settings, it would be useful to direct future imaging studies at the chronic allergic response involving airway remodeling induced by long-term repetitive allergen administration. Our combined approach of in vivo tracheal imaging and in situ staining for signaling events provide an ideal platform for such future studies as well.

In conclusion, our results provide a deeper insight into the spatial and funct ional organization of an airway mucosal T_H2/T_H17 allergic response. Key features of this response are 1) the formation of discrete cellular clusters; 2) antigen-specific T-DC interactions within these clusters leading to local T-cell activation and cytokine production and 3) the resulting recruitment of myeloid effector cells into these clusters, possibly in response to local cytokine effects.

These observations also imply that the episodic formation of inflammatory cell clusters might represent a mechanism by which acute inflammatory foci produce diffuse bronchial disease over time, which could be equivalent to the allergen-induced acute exacerbations of chronic inflammation, as observed in clinical settings. This underlines the importance of the early detection and appropriate, specific therapeutic targeting of acute inflammatory episodes in order to preclude the development of chronic airway disease.

Abbreviations used in this article

CT

cholera toxin

HDM

house dust mite

ILC

innate lymphoid cell

MP-IVM

multi-photon intravital microscopy

o.ph.

oropharingeal

PLP

periodate-lysine-paraformaldehyde