Matrix Metalloproteinase-9-Mediated Dendritic Cell Recruitment into the Airways Is a Critical Step in a Mouse Model of Asthma

Karim Y. Vermaelen, Didier Cataldo, Kurt Tournoy, Tania Maes, An Dhulst, Renaud Louis, Jean-Michel Foidart, Agnès Noël and Romain Pauwels
J Immunol July 15, 2003, 171 (2) 1016-1022; DOI: https://doi.org/10.4049/jimmunol.171.2.1016

Abstract

Dendritic cells (DCs) appear to be strategically implicated in allergic diseases, including asthma. Matrix metalloproteinase (MMP)-9 mediates transmigration of inflammatory leukocytes across basement membranes. This study investigated the role of MMP-9 in airway DC trafficking during allergen-induced airway inflammation. MMP-9 gene deletion affected the trafficking of pulmonary DCs in a specific way: only the inflammatory transmigration of DCs into the airway lumen was impaired, whereas DC-mediated transport of airway Ag to the thoracic lymph nodes remained unaffected. In parallel, the local production of the Th2-attracting chemokine CC chemokine ligand 17/thymus and activation-regulated chemokine, which was highly concentrated in purified lung DCs, fell short in the airways of allergen-exposed MMP-9−/− mice. This was accompanied by markedly reduced peribronchial eosinophilic infiltrates and impaired allergen-specific IgE production. We conclude that the specific absence of MMP-9 activity inhibits the development of allergic airway inflammation by impairing the recruitment of DCs into the airways and the local production of DC-derived proallergic chemokines.

Dendritic cells (DCs)3 have emerged as the most potent APCs and are now recognized as crucial initiators of immune responses in general (1). In the lung, immature DCs are continuously recruited to the airway mucosa where inhaled Ag is sampled, processed, and transported to the thoracic lymph nodes (TLNs), leading to efficient priming of Ag-specific T cells (2). The relevance of DCs in asthma was evident in studies where intratracheal transfer of Ag-pulsed DCs could elicit a Th2-skewed pulmonary immune response (3). In addition, conditional depletion of DCs during allergen challenge completely abrogated eosinophilic airway inflammation (4), suggesting that DCs are necessary APCs during the secondary immune response.

On a molecular level, the cascade of cellular and molecular events leading to the asthmatic phenotype has grown more and more complex. Next to molecules such as cytokines, chemokines, and growth factors, matrix metalloproteinases (MMPs) have recently emerged as critical mediators in this disease. MMPs form a group of proteolytic molecules drawing increasing interest in the field of lung biology (5). MMPs are known for their extracellular matrix (ECM)-degrading activity (6), but they can also modulate inflammation through proteolytic activation or inactivation of cytokines, chemokines, and growth factors (7, 8). We focused on MMP-9 (gelatinase B), a protease involved in the degradation of collagen IV, a major constituent of basement membranes (BMs): it therefore constitutes a useful tool, allowing a cell to move from one tissue compartment to another. Other functions of MMP-9 include ECM remodelling and proteolytic activation of inflammatory mediators such as IL-1β and IL-8 (9). Abnormalities in MMP-9 production have been observed in airway secretions (10), bronchial tissue (11), and blood (12) of asthmatic patients. MMP-9 is produced by many cells that accumulate in allergic airway inflammation, including eosinophils (13), neutrophils (14), and alveolar macrophages (15). Broad-spectrum pharmacologic blockade of MMP activity inhibited airway cellular infiltration caused by single aerosol challenge of allergen-sensitized animals (16) and suppressed the pathophysiology in an animal model of occupational asthma (17). In an asthma model featuring chronic allergen exposure, we have recently reported that MMP-9 gene deletion decreases peribronchial inflammation, airway lymphocyte accumulation, airway IL-13 production, and the development of bronchial hyperresponsiveness (18).

In the current study, we sought to investigate which precise cell populations and relevant chemokine networks upstream in the pulmonary allergic cascade are critically affected by the total absence of MMP-9. We show that MMP-9 gene deletion specifically impairs the inflammatory transmigration of DCs into the airways. This was accompanied by a decreased local supply of DC-derived Th2-attracting chemokines and a strongly attenuated allergic airway inflammation.

Materials and Methods

Animals

MMP-9 knockout mice and their wild-type counterparts were generated as previously described (19) and were kindly provided by Prof. Z. Werb (University of California, San Francisco, CA). Littermates obtained from breeding heterozygous brothers and sisters were used between 6 and 8 wk of age. All in vivo manipulations were approved by the local ethics committee.

Allergen sensitization and exposure

Mice were immunized by i.p. injection of 10 μg of OVA (Sigma-Aldrich, St. Louis, MO) complexed to 1 mg of aluminum hydroxide (Al(OH)3). Two weeks later, mice were exposed for 7 consecutive days to 30 min of 1% OVA aerosol or PBS for control.

Buffers and medium for preparation of single-cell suspensions and immunofluorescent labeling

Tissue culture medium (TCM) was prepared using RPMI 1640 supplemented with 5% FCS, penicillin/streptomycin, l-glutamine, and 2-ME (all from Life Technologies, Rockville, MD). Digestion medium consisted of TCM supplemented with 1 mg/ml collagenase type 2 (Worthington Biochemical, Lakewood, NJ) and 0.02 mg/ml DNase I (grade II from bovine pancreas; Boehringer Mannheim, Brussels, Belgium). FACS-EDTA buffer contained PBS (without Ca2+ or Mg2+), 0.1% azide, 1% BSA (Sigma-Aldrich), and 5 mM EDTA.

Tissue processing and cellular recovery

Twenty-four hours after the last aerosol exposure, mice were anesthetized using i.p. pentobarbital injection, and the trachea was cannulated. One milliliter of HBSS w/o Ca2+ or Mg2+, supplemented with 0.05 mM sodium EDTA, was instilled four times via the tracheal cannula and recovered by gentle manual aspiration. This bronchoalveolar lavage (BAL) fluid was centrifuged, and the cell pellet was subjected to RBC lysis and kept in FACS-EDTA buffer until cell counting and immunofluorescent labeling. Subsequently, blood was collected by cardiac puncture, and mediastinal lymph nodes (LNs) were removed. Right heart catheterization and perfusion with saline-EDTA was performed to remove the pulmonary intravascular pool of cells. One lung was clamped, removed, and kept in ice-cold TCM until enzymatic digestion, or snap-frozen in liquid nitrogen for subsequent RNA extraction. The other lung was perfused in situ with 4% paraformaldehyde through the tracheal cannula using a controlled flow-rate syringe pump and later processed for histology.

Morphometric quantification of allergic airway inflammation

Formalin-fixed, paraffin-embedded lung lobes were cut into 3-μm sections and stained with Congo red to highlight eosinophils. Quantification of inflammation was performed in a blinded fashion using a Zeiss (Oberkochen, Germany) KS400 Image Analyzer system running a custom-made morphometry program. For each mouse, digital images of five to eight bronchovascular structures were acquired and processed as follows. First, the following parameters were measured: the total area of a bronchovascular couple (ATOT), the area taken up by the airway lumen (AAW), and the area taken up by the blood vessel(s) (ABl). The bronchovascular tissue area (i.e., the tissue area prone to inflammatory infiltration) was defined as follows: ABVI = ATOT − (AAW + ABV). Within each ABVI, all mononuclear infiltrate groups were outlined and added up, yielding the total infiltrate area, AINF. The severity of inflammation was then expressed as AINF/ABVI, i.e., the percentage of bronchovascular tissue area taken up by mononuclear infiltrates. In addition, the density of eosinophil infiltration was expressed as the total eosinophil count within the ABVI, divided by ABVI.

Preparation of lung and LN single-cell suspensions

Prelevated lungs and LNs were minced, using iridectomy scissors, and incubated for 45 min in digestion medium in a humidified incubator at 37°C and 5% CO2 as described earlier (2). Subsequently, the samples were thoroughly resuspended, centrifuged, and incubated in calcium- and magnesium-free PBS containing 10 mM EDTA for 5 min at room temperature on a shaker. Finally, samples were subjected to RBC lysis, washed in FACS-EDTA, passed through a 50-μm-cell strainer and kept on ice until immunofluorescent labeling. Lung, LN, and BAL cells were counted using a Z2 Beckman Coulter (Fullerton, CA) particle count and size analyzer.

Labeling of single-cell suspensions for flow cytometry

Cells were preincubated with FcR blocking Ab (anti-CD16/CD32; clone 2.4G2) to reduce nonspecific binding. mAbs used to identify mouse DC populations were as follows: biotinylated anti-CD11c (N418) and PE-conjugated anti-IAb (AF6-120.1), followed by streptavidin-APC. As a last step before analysis, cells were incubated with 7-amino-actinomycin (Viaprobe) 10 min at room temperature for dead-cell exclusion. All reagents were obtained from BD PharMingen (Erembodegem, Belgium).

Four-color flow cytometry data acquisition was performed on a dual-laser FACSVantage SE flow cytometer running CellQuest 3.3 software (BD Biosciences, Mountain View, CA). FlowJo software (www.treestar.com/flowjo) was used for data analysis on PowerMac G3 and G4 workstations (Apple Computer, Cupertino, CA).

Morphometric quantification of airway wall DC infiltration

Sections obtained from formalin-fixed, paraffin-embedded lung lobes were subjected to the following immunohistological staining sequence: blocking reagent (Roche Diagnostics, Mannheim, Germany) plus Triton X-100; rat-anti mouse I-A/I-E (clone M5/114; American Type Culture Collection, Manassas, VA) or rat IgG2b isotype control; goat-anti-rat IgG Alexa 555 (Molecular Probes, Leiden, The Netherlands); avidin/biotin block (Vector Laboratories, Burlingame, CA); blocking reagent plus Triton X-100; biotinylated rat anti-mouse B220 (clone RA3-6B2; BD PharMingen) or biotin rat IgG2a isotype control; and streptavidin Alexa 488 (Molecular Probes). Stained sections were mounted in Gel/Mount (Biomeda, Foster City, CA). Red (MHC class II (MHCII)) and green (B220) digital images were acquired and combined using a Zeiss KS400 image analyzer platform. DCs, identified as MHCII+B220− cells, were counted in the tissue area surrounding the airway epithelium (preliminary tests with a panel of relevant mAbs indicated anti-IA/IE and anti-B220 as optimal for use on formalin-fixed lung tissue sections). Results were expressed as cell counts relative to airway perimeter, i.e., as number of cells per millimeter of BM length as outlined by digital morphometry.

Purification of pulmonary DCs

Pulmonary single-cell suspensions were obtained from lungs of which the vasculature was rinsed but no BAL was performed. Cells were first incubated with FcR block, followed by anti-CD11c microbeads (Miltenyi Biotec, Bergisch-Gladbach, Germany). CD11c+ lung cells were enriched after one passage through a VarioMACS magnetic cell separator (Miltenyi Biotec) according to the manufacturer’s instructions. Subsequently, these cells (70–80% CD11c+ on average) were labeled with PE anti-I-Ab and biotin anti-CD11c, followed by streptavidin-APC. DCs, defined as low autofluorescent/CD11c+I-Ab+, were further sorted on a FACSVantage SE to 99% purity.

Intratracheal instillation of macromolecule solutions into the trachea

Fluorescein-conjugated OVA (OVA-FITC; Molecular Probes) was diluted in sterile PBS to a final concentration of 1 mg/ml. Intratracheal instillation was performed as previously described (2) using disposable and pyrogen-free 18-gauge polyurethane catheters (Insyte-W; BD Biosciences).

Bone marrow-derived DC culture

Mouse bone marrow-derived DCs (mBMDCs) were differentiated using a standard protocol (20). Briefly, cells extracted from mouse long bones were subjected to RBC lysis and cultured for 10 days in TCM supplemented with initial doses of 200 U/ml rGM-CSF (generously provided by Prof. K. Thielemans (Vrije Universiteit Brussel, Brussels, Belgium)). Toward the end of the culture, GM-CSF supplementation was gradually decreased to reduce granulocyte contamination. The final yield and differentiation of DCs was found to be similar in MMP-9 knockout and wild-type animals.

DC Matrigel chemotaxis assay

Twenty-four-well Transwell inserts featuring 8-μm pore size and precoated with Matrigel were obtained from BD Labware (Bedford, MA). Each Transwell insert was seeded with 5 × 105 in vitro-cultured MMP-9+/+ or MMP-9−/− mBMDC in 200 μl of TCM in quadruplicate. The lower wells were filled with 500 μl of a chemokine solution (R&D Systems, Abingdon, U.K.). Recombinant murine CC chemokine ligand (CCL)5 (RANTES), CCL20 (macrophage-inflammatory protein (MIP)-3α), and CCL21 (6Ckine/secondary lymphoid organ chemokine (SLC)/Exodus) were used at an optimal concentration of 10−8, 10−7, and 10−9 M, respectively. Cells were allowed to transmigrate for 4 h in a cell culture incubator, after which the Transwells were lifted, the underside was washed with 450 μl of PBS/5 mM EDTA, and pooled with the content of the lower well. Fifty microliters (i.e., 50,000) of counting beads (Bangs Labs, Fishers, IN) was added to the transmigrated cell suspension, and this mixture was acquired on the flow cytometer. The absolute number of transmigrated cells was calculated as follows: percentage of acquired cells × (50,000/percentage of acquired beads). Results were expressed as a relative migration index (MI), i.e., the MI from chemoattractive Transwells relative to the MI of control Transwells. In some parallel experiments, MMP-9+/+ and MMP-9−/− mBMDCs were allowed to migrate across uncoated 5-μm-pore size Transwells (Costar, Badhoevedorp, The Netherlands).

In vivo migration of adoptively transferred DCs

DCs cultured from MMP-9+/+ and MMP-9−/− mouse bone marrow were incubated in TCM containing 5 μM CFSE (Molecular Probes) for 5 min at room temperature, washed extensively in excess TCM, and resuspended in PBS without Ca2+ or Mg2+. CFSE-labeled DC (106/mouse) were instilled intratracheally as described earlier (3). Thirty-six hours later, mediastinal LN were prelevated and subjected to the above-mentioned enzymatic digestion protocol. The percentage of CFSE+CD11c+ LN cells was determined by flow cytometry.

Semiquantitative RT-PCR analysis

Total lung RNA was extracted using a cesium chloride ultracentrifugation method (18). Alternatively, cellular RNA was purified using the RNeasy mini-kit (Qiagen, Hilden, Germany). Primer and TaqMan probe combinations for the different target messages were as follows: β-actin, forward, 5′-AGAGGGAAATCGTGCGTGAC-3′, reverse, 5′-CAATAGTGATGACTTGGCCGT-3′, and probe, 5′-CACTGCCGCATCCTCTTCCTCCC-3′; CCL22/monocyte-derived chemokine (MDC), forward, 5′-GGTCCCTATGGTGCCAATGT-3′, reverse, 5′-CTTGCGGCAGGATTTTGAG-3′, and probe 5′-CCTCTGCCATCACGTTTAGTGAAGGAGTTC-3′; and CCL17/thymus and activation-regulated chemokine (TARC), forward, 5′-TCCAGGGCAAGCTCATCTGT-3′, reverse, 5′-TCACGGCCTTGGGTTTTT C-3′, and probe, 5′-CCCAAAGACAAACATGTGAAGAAGGCCA-3′.

Chemokine expression was determined using TaqMan RT-PCR. Cycle conditions were as follows: 30 min at 48°C, 12 min at 95°C, and 40 cycles with 15 s at 95°C and 1 min at 60°C. Each reaction was performed in triplicate. In each PCR run, serial dilutions of a positive control sample (RNA from mouse lungs with full-blown allergic airway inflammation) were included separately for both target gene and housekeeping gene to account for possible differences in PCR efficiency between both genes. For semiquantitative analysis, Threshold cycle values were translated into relative concentrations from the standard curves. Relative concentrations of chemokine cDNA thus obtained were standardized relative to housekeeping gene in the corresponding sample. In some experiments, the results were finally expressed relative to a calibrator sample group (i.e., the nonallergic wild-type animals).

ELISA protein measurements

OVA-specific IgE was measured on serum samples using OVA-coated microtiter plates and biotinylated polyclonal rabbit anti-mouse IgE (S. Florquin; Université Libre de Bruxelles, Brussels, Belgium). One unit was defined as a 1/100 dilution of an internal standard serum pool obtained from OVA-sensitized mice.

CCL17 concentrations were determined using a commercially available ELISA kit (R&D Systems).

Statistical analysis

Statistics were performed on Graphpad Instat 3 software (www.graphpad.com). Groups were compared using parametric tests (Student’s t test or one-way ANOVA with posttest) or nonparametric tests (Mann-Whitney U or Kruskall-Wallis with posttest) following standard statistical criteria.

Results

MMP-9 deficiency inhibits allergic airway inflammation

Allergen-sensitized and exposed mice (OVA/OVA groups) demonstrate typical features of human asthma, among which are prominent cuffing of bronchi and bronchopulmonary vessels with mononuclear cells and eosinophils. These features were inhibited in MMP-9−/− mice as revealed by computer-assisted morphometry (Fig. 1⇓, a–c). In addition, synthesis of OVA-specific IgE was impaired in allergen-exposed MMP-9-deficient groups (Fig. 1⇓d).

FIGURE 1.
FIGURE 1.

Allergen-induced inflammation in OVA-sensitized MMP-9 wild-type and knockout animals after challenge with either PBS or OVA aerosol. a, Photomicrographs of Congo red-stained sections. OVA-challenged MMP-9+/+mice feature eosinophil-rich inflammation concentrated around bronchovascular structures, whereas only sporadic infiltration can be seen around the airways of allergen-exposed MMP-9−/− mice. b and c, Quantification of bronchovascular mononuclear cell infiltration and eosinophil density obtained from computer-assisted image analysis (see Materials and Methods). d, Measurement of allergen-specific IgE by ELISA. n = 7–8 mice per group. ∗, p < 0.05 compared with PBS-exposed groups. ∗∗, p < 0.05 compared with wild-typeOVA/OVA animals.

DC recruitment into and around the airways is MMP-9 dependent

Pulmonary DCs were identified by flow cytometry using three simultaneous immunofluorescent criteria: low-autofluorescence, CD11c, and MHCII positivity. Allergen sensitization and exposure caused a considerable increase in DC numbers in the BAL compartment; however, this increase was strongly impaired in MMP-9−/− animals (Fig. 2⇓a). By contrast, DCs accumulated equally in the lung interstitium of allergen-exposed MMP-9−/− compared with their allergic MMP-9+/+ counterparts (Fig. 2⇓b). Pulmonary macrophages were identified as high-autofluorescent, CD11c+ cells. In contrast to low-autofluorescent DCs, these cells have a typical macrophage morphology, are MHCII and T cell costimulatory molecule low to negative, stain strongly with MOMA-2 (21), and induce very weak T cell proliferation in MLR cultures.4 BAL macrophage numbers increased moderately after allergen exposure, but this increase was not affected by MMP-9 deficiency (Fig. 2⇓c). Within the lung interstitium, the extent of macrophage populations remained unaffected in all groups (Fig. 2⇓d). In addition, DC infiltration in the airway wall was quantified as well. DCs were identified by a combination of morphology, MHCII positivity, and the absence of B220 staining using a double immunofluorescent tissue staining protocol. As summarized in Fig. 3⇓, there was a significantly impaired accumulation of airway wall DCs in allergen-exposed MMP-9−/− animals compared with that of wild types.

FIGURE 2.
FIGURE 2.

Flow cytometrical analysis of DC (a and b) and macrophage populations (c and d) in BAL and lung tissue digests (cell populations within lung tissue were expressed in percentages, because the total cell yield after lung digest was similar in all groups). ▨ and ▪, MMP-9+/+ and MMP-9−/− groups, respectively. Only the inflammatory recruitment of DCs into the airway lumen was affected by MMP-9 deficiency. n = 6–8 mice per group. ∗, p < 0.05 compared with sham-exposed groups; ∗∗, p < 0.05 compared with wild-type OVA/OVA animals.

FIGURE 3.
FIGURE 3.

Airway wall DCs were identified as MHCII+B220− peribronchial cells using a double immunofluorescent staining technique and counted. Cell counts were expressed relative to airway caliber as measured by epithelial BM perimeter. n = 6–8 mice per group, with four to five intrapulmonary airways sampled per animal. ∗, p < 0.05 compared with sham-exposed groups; ∗∗, p < 0.05 compared with OVA-exposed wild-type animals.

Airway DC homing to mediastinal LNs is not MMP-9 dependent

In baseline conditions, intratracheal instillation of the fluorescent macromolecule OVA-FITC leads to a peak accumulation of FITC+ migratory airway-derived DCs in the TLN after 24 h (2). MMP-9 deficiency had no significant effect on this trafficking (Fig. 4⇓a). In addition, intratracheal adoptive transfer of CFSE-labeled exogenous DC yielded no difference between the number of MMP-9+/+ and MMP-9−/− CFSE+ DCs reaching the TLN (Fig. 4⇓b).

FIGURE 4.
FIGURE 4.

Homing of DCs from the airways toward local LN is not affected by MMP-9 deficiency. a, Absolute number of endogenous FITC+ airway-derived LN DCs 24 h after intratracheal instillation of OVA-FITC. b, Absolute number of exogenous CFSE+ DCs in the draining TLN 36 h after intratracheal adoptive transfer. n = 6 mice per group. n.s., Not significant (p > 0.05).

Differential requirement for MMP-9 in the transmembrane migration of immature vs mature DCs

We hypothesized that DCs attracted toward the airway lumen require MMP-9 to cross the airway epithelial BM, whereas airway DCs migrating to the draining thoracic LN do not. We substantiated our in vivo observations by studying MMP-9+/+ and MMP−/− bone marrow-derived DCs in a Transwell chemotaxis assay across a layer of Matrigel (a surrogate BM), using chemokines representative for the afferent and efferent arm of pulmonary DC trafficking. On one hand, we used CCL5 (RANTES) and CCL20 (MIP-3α). CCL5 is a CC chemokine known to attract DCs to the airways (22) and is critically involved in the development of allergic airway inflammation (23), whereas CCL20 is known to recruit immature DCs into inflamed epithelial surfaces (24). In contrast, we used CCL21 (6Ckine, SLC), a chemokine which attracts mature DCs toward lymphatic vessels and T cell areas of draining LN (25). Fig. 5⇓ shows that DC transmembrane migration along a CCL5 gradient was clearly impaired in the absence of MMP-9, a phenomenon which was even more pronounced when DCs were chemoattracted by CCL20. In contrast, DCs attracted by CCL21 were able to migrate normally through the BM in the absence of MMP-9. In addition, migration of MMP-9−/− DCs toward MIP-3α was not impaired when uncoated Transwells were used (not shown), suggesting that MMP-9−/− DCs have no inherent defects in MIP-3α responsiveness.

FIGURE 5.
FIGURE 5.

In vivo chemotaxis assay using bone marrow-derived MMP-9+/+ (▨) and MMP-9−/− (▪) DCs, Matrigel-coated porous Transwell inserts, and prototypical chemokines involved in the afferent (CCL5, CCL20) and efferent (CCL21) arm of pulmonary DC migration. ∗, p < 0.05 compared with wild-type DCs. MMP-9-deficient DCs only show an impaired transmigration across the Matrigel membrane when attracted by CCL5 and CCL20, both inflammatory CC chemokines acting on immature DCs.

Impaired production of DC-derived proallergic chemokines in the lungs of MMP-9-deficient allergen-exposed animals

As a next step, we verified whether the pulmonary expression of prototypical Th2-attracting chemokines was affected in allergen-challenged MMP-9-deficient animals. We concentrated on CCL22 (MDC) and CCL17 (TARC). Using TaqMan RT-PCR, we found that both MMP-9+/+ as well as MMP-9−/− animals could up-regulate pulmonary CCL22/MDC expression (∼2-fold) after allergen sensitization and repeated challenge (Fig. 6⇓a). In contrast, expression of CCL17/TARC in whole lungs of OVA/OVA MMP-9−/− animals was significantly impaired compared with that of allergic wild types (Fig. 6⇓b). Similarly, MMP-9 deficiency was associated with lowered levels of CCL17 protein in the BAL compartment after allergen exposure (Fig. 6⇓c). Finally, it was found that CCL17 mRNA levels were ∼30-fold more concentrated in highly purified pulmonary DCs than in whole-lung extracts (Fig. 6⇓d).

FIGURE 6.
FIGURE 6.

Effect of MMP-9 deficiency on proallergic chemokine production in the lung. a and b, Chemokine mRNA levels were expressed relative to a calibrator group (wild-type sham-exposed mice). a, CCL22/MDC expression increases ∼2-fold after allergen challenge in wild-type mice; this increase was only marginally affected by MMP-9 deficiency. b, Pulmonary CCL17/TARC expression is clearly impaired in allergen-challenged MMP-9−/− animals. c, CCL17 protein measurement in BAL reveals a marked deficit in allergen-exposed MMP-9 knockouts. ∗, p < 0.05 compared with sham-exposed groups; ∗∗, p < 0.05 compared with wild-type OVA/OVA animals. d, CCL17 expression (normalized to housekeeping gene) in naive lungs (PBS lung), lungs with allergic airway inflammation (OVA lung), and purified lung DCs (lung DC). ∗, p < 0.05 compared with naive lung. n = 6–8 mice per group.

Discussion

The decreased manifestations of allergen-induced airway inflammation in MMP-9-deficient animals led us to investigate the physiology of upstream cellular actors in this disease model. DCs are emerging as key orchestrators of allergic immune responses, including atopic dermatitis and asthma (26, 27). Therefore, we studied the specific impact of MMP-9 deficiency on DC trafficking through the different pulmonary compartments, both in baseline conditions as well as during inflammation. Our experiments suggest an important role for MMP-9 in the recruitment of airway DCs during inflammation only. Airway DC recruitment presumably follows a multistep process involving two subsequent BM barriers: first, DCs have to extravasate from bronchopulmonary venules (21); subsequently, DCs can migrate through the epithelial BM into the airway lumen or BAL compartment (28, 29). DCs within the BAL compartment (BAL-DCs) represent a unique population of cells, with no equivalent in other immunologically challenged organs such as the skin or the gut. BAL-DCs from allergen-sensitized/exposed animals show strong signs of in situ activation, with high surface expression of MHCII and T cell costimulatory molecules, a phenomenon that normally occurs only after homing into draining LN (49). This, in combination with the direct exposure to aeroallergen and the concomitant presence of BAL T cells, makes BAL-DCs prime candidates for locally maintaining the secondary immune response, perhaps even for a prolonged time after allergen exposure (30). Interestingly, whereas the peribronchial accumulation of DCs was impaired in allergen-exposed MMP-9-deficient animals, analysis of enzymatic lung digests indicated that parenchymal DC accumulation was not MMP-9 dependent (Figs. 3⇑ and 2⇑b, respectively). It could be that MMP-9 is especially required for crossing the endothelial BM barrier at the level of peribronchial blood vessels, whereas DC extravasation at the level of the lung parenchyma relies on other MMPs such as MMP-12 (our unpublished observations). Alternatively, there could be differences in BM structure around systemic vs pulmonary blood vessels, or a compartmentalized expression of DC-attracting chemokines, which in turn could be regulated differently by MMP-9 (following a mechanism described earlier (9)).

Although there is accumulating evidence implicating MMP-9 in the emigration of both human as well as murine epidermal DCs toward draining LNs (31, 32, 33), our in vivo experiments reveal that analogous DC trafficking in the pulmonary system is not MMP-9 dependent. In particular, airway Ag capture and transport toward draining mediastinal LN does not require MMP-9. It is possible that Ag-sampling airway DCs located above the epithelial BM can cross this barrier in an MMP-9-independent fashion. Indeed, MMP-9-deficient DCs deposited in the airway lumen were able to cross the epithelium and home to the TLN. Moreover, our in vitro experiments indicate that MMP-9 only participates in the transmembrane passage of immature DCs toward the inflammatory chemokines RANTES/CCL5 and CCL20/MIP-3α, but not in the transmigration toward the SLC CCL21. One explanation could be that DC maturation induces BM-degrading enzymes other than MMP-9 (MMP-12 is a possible alternative (34)). In contrast, different chemokines might induce different sets of MMPs in DCs through as-yet-unexplored molecular mechanisms. Further detailed investigations will be needed to unravel this complexity in DC-MMP-ECM interactions.

Nonetheless, the presence of DCs scattered just beneath the airway BM (as observed in mouse intrapulmonary conducting airways) might allow for an additional mechanism of MMP-9-independent aeroantigen capture and transport, similar to a recent study on the gut mucosa. Indeed, despite their subepithelial location, intestinal mucosal DCs are perfectly capable of capturing noninvasive luminal Ags by extending interepithelial processes without disrupting the epithelial barrier’s integrity (35). Likewise, aeroantigen sampling could be operated by subepithelial airway DCs in a way that does not require extensive transmigration of the cell body through the BM.

The reduced airway inflammation in MMP-9−/− mice suggested a defect in the upstream network of proallergic chemokines, i.e., chemokines that preferentially attract Th2 cells. Th2 cells typically express the chemokine receptors CCR8, CCR3, and CCR4. CCR8 gene deletion (or neutralization of its ligand) does not affect the full development of allergic airway inflammation (36, 37). CCR3 and its ligand eotaxin were shown to play a prominent role in the early phase of the pulmonary allergic response; however, after repeated challenge such as in our model, CCR4-mediated Th2 recruitment predominates (38). Therefore, we focused on CCR4 ligands, CCL22 and CCL17. CCL22/MDC is a molecule that preferentially attracts chronically activated Th2 cells (39). In addition, CCL22 is part of an amplifying cascade, whereby its expression is further stimulated by the Th2 products IL-4 and IL-13 (39). Similar to previous reports, we observed an up-regulation of CCL22 after allergen sensitization and chronic challenge, but this was only marginally affected by MMP-9 deficiency. Indeed, it has been shown in a murine model of allergic airway inflammation that CCL22 protein is predominantly found in lung myofibroblasts and macrophages (40). Although the former cells are obviously nonmigratory in nature, we now report that both tissue-bound as well as BAL macrophage mobilization is unaffected in MMP-9 knockout mice.

Therefore, we sought to examine a Th2-mobilizing chemokine that is largely produced by cells featuring an MMP-9-dependent recruitment. We focused on CCL17/TARC, another CCR4 ligand with chemoattractant specificity similar to that of CCL22 (41). In vivo inhibition of CCL17 activity suppresses bronchial hyperresponsiveness, lung eosinophilia, and BAL Th2 cytokine levels (42). In vitro studies have identified human bronchial epithelial cells, monocytes, macrophages, and DCs as sources of CCL17 (41, 43). In vivo, CCL17 was found to be expressed in human bronchial and nasal epithelial layers as well as tissue mononuclear cells, and this expression was more pronounced in allergic subjects (44, 45). Moreover, CCL17 protein levels are elevated in the BAL of asthmatic patients after allergen challenge (46). Interestingly, CCL17 expression in the mouse respiratory system was found to be concentrated in CD11c+ cells (47) that were scattered in peribronchial regions beneath the airway epithelium (although BAL-DCs were not examined) (48). In these studies, CCL17 expression was shown to be restricted to mouse DCs, with absent expression in macrophages or B cells even after stimulation. Our experiments reveal a marked deficit in pulmonary CCL17 expression and CCL17 protein levels in the airways of allergen-exposed MMP-9 knockouts. We argue that this difference is due to the impaired recruitment of DCs into and around the airways of these mice. In line with previous reports, we found pulmonary CCL17 expression to be highly concentrated in purified pulmonary DCs. Also, pulmonary DCs purified from allergic vs naive lung had equivalent CCL17 expression levels (not shown), suggesting that differences in CCL17 are due to changes in cell numbers, rather than CCL17 modulation per cell. Although we do not exclude airway epithelial cells as CCL17 producers in the allergic mouse lung, we propose that the massively recruited, activated airway DC population constitutes a more direct and relevant source of this chemokine compared with any other structural cell in the lung. Moreover, the functional impact of DC-derived CCL17/TARC far outweighs the consequences of a hypothetical in vivo interaction between epithelial cells and Th2 cells: as demonstrated earlier (4), in this model of allergic airway inflammation, the secondary immune response collapses in the specific absence of DCs.

In summary, our study suggests a mechanism through which MMP-9 gene deletion inhibits allergic airway inflammation: freshly recruited DCs cannot infiltrate the airway mucosa and do not reach the airway lumen in sufficient numbers. The crucial DC-mediated Ag presentation and costimulation to airway memory Th2 cells would come short, and DC-derived Th2-attracting chemokines would fail to accumulate as well. As a consequence, the local deficiency of Th2-derived effector cytokines would limit the amplitude of the allergic cascade. Together, these data emphasize the potential of selective MMP inhibitors in the treatment of allergic asthma, while simultaneously suggesting a critical role for airway DCs in this disease.

Acknowledgments

Many thanks to Greet Barbier, An Neessen, Indra De Borle, Katleen De Saedeleer, Philippe De Gryze, Marie-Rose Mouton, Eliane Castrique, and Christelle Snauwaert for their technical contribution to this work.

Footnotes

  • ↵1 This work was supported by the Fund for Scientific Research in Flanders (Research Project G.0393.99), the Concerted Research Initiative of the University of Ghent (Project 98-6), medical research funds of the Fonds National de la Recherche Scientifique, Fondation Leon Frédericq (University of Liège), and Fonds Spéciaux de la Recherche from the University of Liège. K.Y.V., D.C., and K.T. are research fellows of the Fund for Scientific Research. T.M. and A.D. are research fellows of the Concerted Research Initiative of the University of Ghent. A.N. is a senior research associate of the Fonds National de la Recherche Scientifique.

  • ↵2 Address correspondence and reprint requests to Dr. Karim Y. Vermaelen, Department of Respiratory Diseases, Ghent University Hospital, 7K12ie, De Pintelaan 185, B-9000 Ghent, Belgium. E-mail address: Karim.Vermaelen{at}rug.ac.be

  • ↵3 Abbreviations used in this paper: DC, dendritic cell; LN, lymph node; TLN, thoracic LN; MMP, matrix metalloproteinase; ECM, extracellular matrix; BM, basement membrane; TCM, tissue culture medium; BAL, bronchoalveolar lavage; MHCII, MHC class II; mBMDC, mouse bone marrow-derived DC; CCL, CC chemokine ligand; MIP, macrophage-inflammatory protein; MI, migration index; MDC, monocyte-derived chemokine; TARC, thymus and activation-regulated chemokine; SLC, secondary lymphoid organ chemokine.

  • ↵4 K. Y. Vermaelen and R. A. Pauwels. Accurate and simple discrimination of mouse pulmonary dendritic cell and macrophage populations by flow cytometry: methodology and new insights. Submitted for publication.

  • Received December 4, 2002.
  • Accepted May 14, 2003.

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