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Intratracheal Priming with Ovalbumin- and Ovalbumin 323–339 Peptide-Pulsed Dendritic Cells Induces Airway Hyperresponsiveness, Lung Eosinophilia, Goblet Cell Hyperplasia, and Inflammation¹

Sun-sang J. Sung^2,*,,, C. Edward Rose, Jr. and Shu Man Fu^*,,,¶

^* Division of Rheumatology and Immunology, and Pulmonary and Critical Care Division, Department of Internal Medicine, University of Virginia Specialized Center of Research on Systemic Lupus Erythematosus, University of Virginia Cancer Center, and ^¶ Department of Microbiology, University of Virginia Health Sciences Center, Charlottesville, VA 22908

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Dendritic cells (DC) are the primary APC responsible for the capture of allergens in the airways and the shuttling of processed allergens to the draining lymph nodes where Ag presentation and T cell activation take place. The mechanism of this Ag handling and presentation in asthma is poorly understood. In addition, the feasibility of asthma induction by DC priming has not been directly tested. In this report an asthma model using intratracheally (i.t.) injected splenic DC was used to address these issues. DC pulsed with a model Ag OVA or the major MHC class II-restricted OVA T epitope peptide OVA_323–339 and instilled i.t. primed mice to exhibit asthma-like diseases. With OVA as the Ag, mice exhibit airway hyperresponsiveness (AHR), lung eosinophilia and inflammation, and pulmonary goblet cell hyperplasia. In OVA_323–339-immunized mice, AHR and goblet cell hyperplasia were noted, with little eosinophilia and parenchymal inflammation. The latter finding provides evidence for dissociating AHR from eosinophilia. In both cases mediastinal node hypertrophy occurred, and high levels of Th2 cytokines were produced by the lung and mediastinal lymph node cells (LNC). Interestingly, mediastinal LNC also produced high levels of Th1 cytokines. Lung cells produced much less Th1 cytokines than these LNC. These results demonstrate that DC when introduced i.t. are potent in inducing asthma-like diseases by recruiting lymphocytes to the lung-draining lymph nodes and by stimulating Th2 responses and also suggest that the lung environment strongly biases T cell responses to Th2.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Allergic asthma is characterized by reversible airway obstruction and airway hyperresponsiveness (AHR),³ pulmonary inflammation, airway eosinophilia, and the elevation of total and allergen-specific IgE (1, 2, 3). The immunological basis of this disease has been a topic of intense investigation. T cell responses to allergens have been found to be critical in the pathogenesis of asthma. In addition, a large number of cytokines, chemokines, adhesion molecules, and costimulatory molecules have been shown to be involved (reviewed in Ref. 3).

The T cell response in allergic asthma is considered to be a Th2-biased response to the inciting allergen. This is supported by both clinical and experimental data (2, 3, 4, 5). Despite this paradigm, the mechanism by which allergens induce Th2 responses in the lungs remains to be clarified. In rats it has been shown that Ag-pulsed lung dendritic cells (DC) prime the rat to produce Ag-specific Th2-mediated IgG1 with little Th1-directed Ag-specific IgG2b generation (6). This and other studies place lung DC in a central role in the Th2-biased response to inhalant allergens (7).

In the mouse it is not feasible to use isolated lung DC for studying Ag priming in the lungs and for asthma induction because of the low DC numbers that can be purified from mouse lungs (8, 9). Recently, mouse bone marrow-derived DC instead of lung DC have been used successfully in intratracheal (i.t.) injections for studying DC migration and T cell priming in an adoptive transfer model with peptide Ag (10). These DC were found to present T epitope peptides to lung-draining lymph node T cells and induce these T cells to exhibit an activated phenotype and divide rapidly. However, specific peptide induction of asthma-like symptoms and pathology has not been studied in these mice. To date, few studies using T epitope peptides to induce asthma have been reported. This may be related to reports of tolerance induction by intranasally injected peptides (11, 12, 13) and the weak AHR responses in mice induced by aerosolized peptide detectable only by sensitive measurements such as tracheal smooth muscle constriction determinations (14). There has also been a lack of studies of asthma induction by Ag-pulsed DC. The reports that show homing of i.t. injected DC to the lung-draining lymph nodes (10, 15) provide further impetus for using i.t injected DC for asthma induction.

In this report DC functions and T cell priming in asthma were studied with splenic DC pulsed in vitro with either OVA or the major I-A^d-restricted OVA peptide 323–339 (OVA₃₂₃). When injected i.t., these DC were efficient in priming mice to both protein and peptide Ag. The immunized mice exhibited AHR and lung eosinophilia, inflammation, and goblet cell hyperplasia. Ag-pulsed DC when injected i.t. also induced hypertrophy of the lung-draining nodes and stimulated both Th1 and Th2 cells in these nodes. In contrast, in DC-primed and Ag-restimulated mice, lower levels of Th1 cytokines were produced by lung cells, although Th2 cytokine production by these cells was similar to that by lymph node cells (LNC). In the circulation, Ag-specific IgE was detected in OVA- but not OVA₃₂₃ peptide-immunized mice. These results showed that i.t. injected DC were efficient in inducing asthma-like diseases in mice and stimulated both LN and lung Th2 cells, which may be responsible for the manifestation of asthma-related symptoms. The experiments with OVA₃₂₃ peptides also dissociated AHR from eosinophilia, severe airway inflammation, and Ag-specific IgE production. These studies reveal important cellular functions during asthma induction and provide a versatile model for studying asthma pathogenesis.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Mice

Female BALB/cByJ mice (4–6 wk old) were purchased from The Jackson Laboratory (Bar Harbor, ME) and housed in a specific pathogen-free facility. Mice were fed rodent diet 7012 (Harlan, Madison, WI), which is free from animal protein products. Experiments were performed with a protocol approved by the University of Virginia animal use and care committee.

Reagents

PE-conjugated anti-CD11c and anti-B220, FITC-conjugated anti-CD8 and anti-Thy1.2, and chromophore-conjugated rat isotype control mAb were purchased from PharMingen (La Jolla, CA). Anti-F4/80-FITC mAb was purchased from Caltag (Burlingame, CA). Anti-pan I-A-FITC mAb was obtained from Cappel (Aurora, OH). OVA (type V) and acetyl--methylcholine chloride (methacholine) were purchased from Sigma (St. Louis, MO). Collagenase D was obtained from Roche (Indianapolis, IN). The OVA T epitope peptides were synthesized at the University of Virginia Biomolecular Research Facility with F-moc chemistry on a Symphony peptide synthesizer (Rainin, Woburn, MA) and purified by reversed-phase HPLC to 98% purity. The molecular masses of the purified putative OVA peptides were confirmed by matrix-assisted laser desorption ionization-time of flight mass spectrometry on a Voyager-DE PRO mass spectrometer (PerSeptive Biosystems, Framingham, MA).

Cells

Cells were cultured in RPMI 1640 supplemented with 5% heat-inactivated (56°C, 30 min) FCS (HyClone, Logan, UT), 50 µM 2-ME, 100 U/ml penicillin, 100 µg/ml streptomycin sulfate, and 2 mM L-glutamine unless noted otherwise. Splenic DC were isolated essentially as previously described (16) and were further purified by magnetic bead depletion. Briefly, low density cells from collagenase-dispersed splenocytes were plated at 40 x 10⁶ in 4 ml of medium in 60-mm dishes. After gentle pipetting and washing, the remaining adherent cells were Ag pulsed overnight in 2 ml of complete medium containing 5% heat-inactivated mouse serum (Pel-Freez, Little Rock, AK) instead of FCS, 10 ng/ml mouse rGM-CSF (5 x 10⁶ U/mg; PeproTech, Rocky Hill, NJ), and 20 µM OVA peptide or 500 µg/ml OVA. DC incubated overnight with no Ag addition were designated control DC. After overnight incubation, the nonadherent cells were treated with a mixture of magnetic microbeads conjugated with anti-B220, anti-Thy1.2, or DX5 anti-NK cell mAb, and the non-DC populations were depleted by a VarioMACS assembly fitted with a VS column (Miltenyi, Auburn, CA). The purified DC with yields of 0.5–1 x 10⁶ cells/spleen were washed, counted, and resuspended in RPMI 1640 for i.t. or s.c. injection. For purified DC characterization, cells were stained essentially as previously described (17) and analyzed on a FACScan. In some experiments, DC were pulsed overnight with 500 µg/ml FITC-labeled OVA (Molecular Probes, Eugene, OR) for estimation of the amount of OVA coinjected with DC. The OVA-pulsed and purified DC were resuspended at 2 x 10⁷ cells/ml in PBS, and graded volumes of cells were diluted to 2 ml with H₂O. A parallel culture with DC incubated with unlabeled OVA was used as a control. The fluorescence of the lysed DC samples was compared with a standard curve of FITC-OVA with and without addition of control DC lysate, and the amount of DC-associated OVA was calculated from the standard curve.

For lung cell isolation, mice were injected i.p. with 10 mg/kg xylazine, 60 mg/kg ketamine, and 150 U heparin. The heart was exposed in the anesthetized mouse, the aorta was severed, and the lungs were perfused gently with 5 ml of cold PBS through the right ventricle using a syringe fitted with a 25-gauge needle. The lungs were excised, minced, and digested for 30 min at 37°C with collagenase D (200 U/ml). The cells were further dispersed by pipetting with Pasteur pipettes, and the suspensions were passed through sterile cotton plugs. The flow-through was further sedimented over a Ficoll-Hypaque (Sigma; density, 1.083) cushion. The interface mononuclear cells were washed with cold PBS, and the cell numbers were counted.

For LNC isolation, the mediastinal lymph nodes were excised under a dissecting scope, and the cells were dispersed in medium. Cells from all mediastinal lymph nodes in the same mice were pooled and counted.

Mouse immunization

When alum was used as adjuvant, priming was performed twice i.p., 1 wk apart, followed by eight consecutive i.t. daily injections of 100 µg of OVA or 50 nmol of peptide beginning on the 15th day (day 14). Proteins (100 µg) or peptides (50 nmol) were alum (Imject, Pierce, Rockford, IL; 2.3 mg)-precipitated overnight in 100 µl before injection. For i.t and footpad injections, mice were anesthetized with methoxyflurane inhalation. Fifty microliters of DC in medium or Ag in IFA was used in footpad injections. For i.t. immunizations, Ag or DC in 50 µl of solution were injected with a catheter fitted with 0.61-mm (OD) polyethylene tubing. Two i.t. injection protocols were used. Protocol I was used to induce AHR by OVA. Mice were primed twice with DC or Ag i.t. on days 0 and 6 followed by i.t. injections of saline or 100 µg of OVA on days 12, 14, 16, and 18. Protocol II was used to induce AHR by peptides. Mice were primed once with DC or soluble Ag (100 µg of OVA or 50 nmol of OVA peptides) on day 0 followed by consecutive daily i.t. injections starting with day 6 for a total of eight boosting doses of 100 µg of OVA or 50 nmol of peptides. AHR was measured 12–24 h after the final i.t. injection. Surgery for obtaining blood and bronchoalveolar lavage (BAL) and for lung histology preparation was performed 1 day after the last injection.

AHR measurements

AHR was measured by whole-body plethysmography as previously described (18) using a Buxco (Troy, NY) plethysmography assembly consisting of unrestrained plethymographs, a MAXII signal analyzer, and Biosystems XA data-handling software. Immunized mice were placed individually in plethysmographs. An air supply with an inward flow of 600 ml/min/box was connected to the box for air exchange and aerosol delivery. For AHR measurements, mice were aerosolized for 3 min with increasing concentrations of methacholine solution (6.25–50 mg/ml in saline) nebulized in an Ultra-Neb 99 (DeVilbiss, Somerset, PA), which generates 0.5- to 5-µm diameter aerosol droplets. Enhanced pause (Penh) values were used as indicators of AHR. This is an index of airway resistance calculated by software from box (plethysmograph) pressure recorded during the inhalation and exhalation of the animals. Penh is defined as [(expiratory time/relaxation time) - 1][peak expiratory flow/peak inspiratory flow], and it reflects both the characteristics of pronounced changes in box pressure during early expiration and the increased interval between breaths during broncho-constriction (18, 19). The ratios of the Penh values at a given methacholine concentration to the basal Penh values of the animals were used in AHR plots to adjust for differences in individual animals. Mice were aerosolized with saline before methacholine treatment. No difference in Penh values between nonaerosolized and saline-aerosolized mice was observed.

BAL and lung histology

Mice were anesthetized with 10 mg/kg xylazine and 60 mg/kg ketamine i.p., injected with 150 U heparin, and sacrificed by exsanguination. Blood was collected for Ig measurements by incision of the axillary artery. BAL collection and lung fixation were performed essentially as previously described (20). Briefly, a 22-gauge catheter was inserted into the exposed trachea of exsanguinated mice and secured with ligatures. The ribcage was opened, and the left bronchus was clamped with a plastic-shod hemostat. The right lung was lavaged with 6 ml of saline at 25 cm hydrostatic pressure. The BAL cells were spun down, counted, and stained with Diff-Quik for differential counting. After lavage of the right lung, the left lung was fix-inflated for 30 min with 10% buffered formalin (Fisher Scientific, Pittsburgh, PA) at 25 cm hydrostatic pressure. The fixed left lung was then excised, fixed in formalin overnight, and sectioned across the bronchus transversely into upper and lower halves. Paraffin blocks with the incision plane facing upward were made, and serial lung sections near the bronchus were stained with hematoxylin and eosin and periodic acid-Schiff (PAS) reagent. Stained BAL cells and lung histology were photographed with an Olympus BH-2 microscope equipped with a digital MTI 3CCD camera (Dage-MTI, Michigan City, IN), and the images were recorded with Image-Pro software (Media Cybernetics, Silver Spring, MD).

Scoring of lung leukocyte infiltration and goblet cell hyperplasia

Lung leukocyte infiltration was scored by two alternative methods. A graded pathological scale of 0–4 was used as an inflammation indicator with the criteria: 0, no infiltration; 1, sparse perivascular and peribronchial infiltration; 2, large patches of perivascular and peribronchial infiltration spanning <20% of the bronchial or large arteriole walls; 3, extensive perivascular and peribronchial infiltration spanning 20–70% of the large bronchial and arteriole walls; and 4, massive perivascular and peribronchial infiltration surrounding 70–100% of the large bronchial and arteriole walls. A quantitative scale measuring the total area of peribronchial and perivascular infiltration in whole lung section was also used. In this measurement, the infiltration areas of two sections close to the bronchus per lung were quantitated with the software Image-Pro from the digital images of the sections using x100 magnification, and the infiltration areas per section were reported.

Goblet cell hyperplasia was determined by counting the number of PAS⁺ cells surrounding the large bronchioles on a digital image under x400 magnification. Two sections were scored per lung. Because tangential sections of the bronchial walls overestimate the number of PAS⁺ cells along the basement membrane, only PAS⁺ cells with nuclei within the section were scored. The circumference of the bronchi at the basement membrane was measured by the polygon function of Image Pro. The results are expressed as the number of PAS⁺ cells per unit length of basement membrane.

Cytokine production by lung cells and LNC

For cytokine production determination by ELISA, culture supernatants at appropriate dilutions were assayed in duplicate. Lung cells or LNC were resuspended at 4 x 10⁶ cells/ml. To these cells were added irradiated (10 Gy) splenocytes as APC and Ag to give final concentrations of 2 x 10⁶/ml lung cells or LNC, 1 x 10⁶/ml splenocytes, and 500 µg/ml OVA or 20 µM OVA₃₂₃ peptide. Cells were incubated for 2 days, and the supernatants were collected for IFN-, IL-2, IL-4, IL-5, and IL-13 determinations by ELISA. Preliminary time-course experiments showed that 2 days were optimal for the supernatant collection. Splenocytes alone produced no detectable cytokines.

Cytokine concentrations in cell supernatants were determined by sandwich ELISAs using paired mAb against murine IFN- (Endogen, Woburn, MA), IL-2 (PharMingen), IL-4 (Endogen), IL-5 (Endogen), and IL-13 (R&D Systems, Minneapolis, MN; affinity-purified goat detecting Ab) and protocols provided by the manufacturers. Streptavidin-HRP and o-phenylenediamine were used for color development. Standard curves were constructed for each cytokine over the range of 0.1–3 ng/ml. Standard recombinant mouse cytokines were obtained from the following sources: rIFN-, Endogen; IL-2 and IL-4, PeproTech (Rocky Hill, NJ); and IL-5 and IL-13, PharMingen. The detection limits of the assays were: IFN-, 100 pg/ml; IL-2, 50 pg/ml; IL-4, 20 pg/ml; IL-5, 25 pg/ml; and IL-13, 30 pg/ml.

Ag-specific Ig subclass quantitation

Anti-OVA and anti-OVA peptide Ab were quantitated by ELISA. Protein and peptides (2 µg/well) were adsorbed overnight on Immulon 1 and Immulon 4 plates, respectively (Dynatech, Chantilly, VA). After blocking the wells with 5% BSA, appropriate dilutions of serum samples in duplicate were added to the wells. Ig subclasses were detected with HRP-conjugated rat anti-mouse IgG1 or anti-mouse IgG2a mAb (PharMingen). IgE was measured by biotinylated rat anti-mouse IgE mAb followed by streptavidin-conjugated HRP (PharMingen). Plate-bound HRP was measured by colorimetric assays with o-phenylenediamine (Sigma) as substrate. A pooled mouse serum sample with Ab against various Ag was used as a reference standard, and the detected amount of Ig subclass Ab in the reference serum was defined as 1000 units.

Statistics

Results were expressed as the mean ± SD. The statistical significance of the differences between experimental and control groups was analyzed by Student’s t test with the graphic software Slidewrite Plus (Advanced Graphics Software, Carlsbad, CA).

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Intratracheal injection of OVA induces AHR

The protocol described by Krinzman et al. (21) with OVA as the protein Ag and two weekly i.p. primings with alum as adjuvant was first employed to establish the efficacy of i.t. boosting for AHR induction. The results showed that with alum as adjuvant for priming, significant AHR was induced in mice injected i.t. with OVA (Fig. 1A). However, no AHR was observed in BALB/c mice immunized with either of the T cell epitope peptides OVA₃₂₃ or peptide 273–288 (OVA₂₇₃) restricted by I-A^d. Thus, for peptide Ag more efficient adjuvants than alum are needed for AHR induction. Other control groups in these experiments showed that no AHR was induced in mice with sham immunization (Fig. 1A) or in mice with i.t. OVA boosting but with no Ag priming (not shown). These experiments showed that i.t. injection induced AHR in properly primed mice.

View larger version (22K): [in this window] [in a new window]   FIGURE 1. Mice immunized i.t. exhibited AHR. A, Priming i.p. with alum as adjuvant. Four mice per group were immunized i.p. with alum alone twice followed by saline (Sal), or OVA, OVA323, or OVA273 in alum followed by i.t. injection of OVA (Ova), OVA323 (323), or OVA273 (273), respectively, as described in Materials and Methods. B, DC dose response and AHR in mice primed with OVA-pulsed DC. Mice (four per group) were primed i.t. with control DC (1 x 106/mouse) followed by saline (Sal) or were primed with 0.1 x 106 (0.1, DC(Ov)), 0.3 x 106 (0.3, DC(Ov)), or 1 x 106 (1, DC(Ov)) OVA-pulsed DC followed by OVA injection according to protocol I in Materials and Methods. C, AHR in mice primed with OVA-pulsed DC. Six mice per group were injected i.t. twice with 0.5 x 106/injection of control DC followed by saline (Sal) or with soluble OVA or 0.5 x 106/injection of OVA-pulsed DC (DC(Ov)) followed by OVA injection as described for protocol I. AHR was measured as described in Materials and Methods. Results are expressed as the ratio of Penh value of animals at the indicated challenging methacholine concentration vs the Penh value of the same unchallenged animals. Differences in Penh values between experimental and control mice were tested for statistical significance by Student’s t test and are indicated by asterisks over the symbols (*, p < 0.05; **, p < 0.01; ***, p < 0.005; ****, p < 0.001). C, Statistical significance in Penh values between animals primed with OVA-pulsed DC and soluble OVA is indicated by stars (, p < 0.05; , p < 0.01). The experiments in A and B were performed twice, and those in C were performed three times.

Splenic DC were chosen for Ag priming in mouse AHR induction because they are the most well-studied DC type. The purities of the DC preparations were characterized by flow cytometry (Fig. 2). There was no detectable T cells in the purified DC populations, as shown by the absence of Thy1.2 staining cells (Fig. 2C). Low levels of B220⁺ cells (2–3%) were observed (Fig. 2C). Macrophages, identified as F4/80⁺ and CD11c^low cells, were low in number in these preparations (<2%; Fig. 2F). No DX5⁺ NK cells were detected in these preparations (not shown). Based on these estimates and the percentage of CD11c⁺I-A^high cells (Fig. 2D) normalized to negative control staining (Fig. 2B), DC comprised 94–95% of the total cellular populations. The CD11c⁺ and I-A^high DC population could be divided into two subpopulations based on CD8 expression, with about a 5:1 ratio for the CD8^-:CD8⁺ populations (Fig. 2E). This is comparable to the ratio observed by other investigators (22).

View larger version (44K): [in this window] [in a new window]   FIGURE 2. Flow cytometric analysis of purified splenic DC. Splenic DC from BALB/c mice were purified as described in Materials and Methods using BSA density gradient centrifugation, adherence, and magnetic bead depletion. Cells in the gated area in A were analyzed for B and T cell (C), DC (D), DC subpopulations (E), and macrophage (F) surface marker expression. Control mAb staining is shown in B. Low side and forward scatter events (A) outside the gated area are comprised of noncellular material with background fluorescence signals.

Priming with OVA-pulsed DC induced airway eosinophilia and lung inflammation

A significantly higher number of BAL cells was found in mice primed with OVA-pulsed DC than in mice primed with either control DC or soluble OVA (Fig. 3A). The majority of this increase in BAL cell number could be attributed to the increase in eosinophils (Fig. 3B). The percentage of eosinophils in mice primed with OVA-pulsed DC was manyfold higher than that in mice primed with control DC and was 4- to 5-fold higher than that in mice primed with soluble OVA (Fig. 3B). The total eosinophil numbers in the BAL of mice primed with OVA-pulsed DC was 10-fold higher than that of mice primed with soluble OVA. The increased eosinophil numbers in BAL of mice primed with OVA-pulsed DC is evident in Fig. 4. BAL cells from control mice consisted predominantly of macrophages (Fig. 4A). A small percentage of eosinophils was observed in the BAL of soluble OVA-primed mice (Fig. 4B). In BAL of mice primed with OVA-pulsed DC (Fig. 4C), many more eosinophils were found. Thus, with the priming by OVA-pulsed DC, severe airway eosinophilia was induced.

View larger version (28K): [in this window] [in a new window]   FIGURE 3. Total cell numbers and differential counts of BAL in mice primed with OVA-pulsed DC. Mice were primed twice with control DC followed by i.t. saline (Sal) or OVA (Sal/Ova) injection or were primed twice with soluble OVA (Ova) or OVA-pulsed DC (Ova-DC) followed by i.t. injections of OVA as described for protocol I in Materials and Methods. Total BAL cell numbers from the right lung (A) and the differential of the BAL cells (B) are shown. The statistical significance of differences in cell number and percentage of eosinophils between Sal- and OVA-DC-treated mice is indicated (*, p < 0.05). The results are representative of two similar experiments. In B, the cell types were: Eos, eosinophils; M, macrophages; N, neutrophils; and L, lymphocytes.

View larger version (115K): [in this window] [in a new window]   FIGURE 4. Cells in BAL of mice primed twice with OVA-pulsed DC. Mice were immunized as described for protocol I in Materials and Methods. Cells were spun on slides and stained with Diff-Quik. BAL cells from mice immunized with control DC followed by saline (A) or with soluble OVA (B) or OVA-pulsed-DC (C) followed by OVA were photographed as described in Materials and Methods at x1000. The bar in A equals 20 µm. Arrows indicate selected eosinophils. M, Alveolar macrophages; N, neutrophils.

View larger version (133K): [in this window] [in a new window]   FIGURE 5. Lung leukocyte infiltration and goblet cell hyperplasia in mice primed with soluble OVA or OVA-pulsed DC. Immunization was performed according to protocol I in Materials and Methods. Mice were injected i.t. with control DC followed by saline (A, D, and G), soluble OVA followed by OVA (B, E, and H), or OVA-pulsed DC followed by OVA (C, F, and I). The left lungs were fixed, and 3-µm sections were stained by hematoxylin-eosin (A–F) or PAS (G–I). The magnifications were: A–C, x100; and D—I, x400. Bars in C and I equal 200 and 50 µm, respectively. Small arrows, Infiltrating leukocytes; large arrowheads, goblet cells with strong PAS staining; b, bronchus; a, arteriole.

As stated previously, peptide priming with alum as adjuvant was ineffective, indicating that a different protocol for peptide-induced asthma is required. Several preliminary experiments were performed. It was found that IFA was required as an adjuvant in the priming via the footpad with OVA₃₂₃ as the Ag. As shown in Fig. 6A, significant AHR was induced. The control peptide OVA₂₇₃ was ineffective in inducing AHR in this protocol.

View larger version (28K): [in this window] [in a new window]   FIGURE 6. AHR of mice primed with OVA323-pulsed DC. BALB/c mice were primed with 100 µg of peptide emulsified in IFA in the footpad and base of tail (A) or with 3 x 105 control DC, OVA323-pulsed DC (323), or OVA273-pulsed DC (273), either i.t. (B) or in the footpad (C), as described in Materials and Methods (protocol II). Control mice (Sal) were primed with IFA alone (A) or control DC (B and C) followed by saline boosting. D, DC dose-response studies are shown. Mice were primed with 5 x 104 (DC, 5), 15 x 104 (DC, 15), or 50 x 104 (DC, 50) OVA323-pulsed DC and boosted with OVA323. Mice injected with varying numbers of control DC exhibited no changes in Penh values. Thus, the Penh values from all control DC-treated mice were pooled (DC, Sal). Six days later mice were injected i.t. with saline or 100 µg of peptide for 8 consecutive days. AHR was measured as described in Materials and Methods. The results were expressed as the mean ± SD. The statistical significance between the Penh values in OVA323- and saline-immunized mice of each methacholine dose were: **, p < 0.01;***, p < 0.005. In all experiments four to six animals were used per group. The experiment in B was repeated four times and the experiments in A, C, and D were repeated twice.

The number of OVA₃₂₃-pulsed DC required for effective i.t. priming of mice for AHR was determined (Fig. 6D). Mice injected with 5–50 x 10⁴ control DC showed no significant difference in Penh values compared with mice with no DC priming (not shown). Significant AHR was observed in mice primed with 5 x 10⁵ DC (Fig. 6D). Combined with the results shown in Fig. 6B, 3–5 x 10⁵ DC were found to be required for effective priming of mice to OVA₃₂₃ in AHR induction.

BAL composition and lung inflammation in mice immunized i.t. with OVA₃₂₃ peptide

In mice primed with OVA₃₂₃-pulsed DC and restimulated with the OVA peptide, no significant change in total BAL numbers was found (Fig. 7A). The great majority of the cells in the BAL of these peptide-immunized mice as well as those of control mice were macrophages (>90%; Fig. 7B). Few eosinophils were found in BAL of control mice or in mice primed with soluble peptide (1–2%). There was a low, but statistically significant, increase in eosinophil numbers in the lungs of OVA₃₂₃-stimulated mice (6%; Fig. 7A).

View larger version (31K): [in this window] [in a new window]   FIGURE 7. BAL cell numbers and cellular composition in OVA323-immunized mice. Mice were immunized i.t. with control DC followed by saline (Sal), soluble OVA323 peptide followed by OVA323 (323), or OVA323-loaded DC followed by OVA323 (DC/323) as described in Materials and Methods (protocol II). The total numbers of BAL cells and the number of eosinophils in BAL from the right lung of mice are shown in A. The differential counts of cell populations in BAL of mice enumerated in A are shown in B. The percentages of mast cells and basophils were low (<1%) and therefore are not shown. DC were used at 5 x 105/mouse in this experiment. The results are representative of three experiments. Statistical significance: *, p < 0.01 in A between OVA323-treated and control mice.

View larger version (95K): [in this window] [in a new window]   FIGURE 8. Goblet cells staining in lungs from mice primed with OVA323-pulsed DC. Fixed lungs from mice immunized according to protocol II with one-time priming using control DC or OVA323-pulsed DC followed by saline (A) or OVA323 (B), respectively, were sectioned and stained by PAS as described in Materials and Methods. The staining of lung epithelia in mice primed with soluble OVA (C) or OVA-pulsed DC (D) followed by OVA using the same immunization schedule is shown for comparison. The epithelia adjacent to the large bronchioles are shown. b, Bronchiolar space; a, arteriolar space. Arrowheads point to PAS+ goblet cells. The images were captured at x400 magnification. The bar in A is 40 µm in length.

Goblet cell hyperplasia in mice immunized with OVA and OVA₃₂₃

Mucin production in goblet cells in lung epithelia were identified by PAS staining. In control mice, lung epithelia showed few PAS⁺ cells (Fig. 5G). Mice primed with soluble OVA showed increased PAS staining of goblet cells in their lung epithelia (Fig. 5H and Table I). Lungs in mice primed with OVA-pulsed DC showed marked goblet cell hyperplasia, characterized by higher goblet cells density, larger cell size, and more intense PAS staining of goblet cells than control lungs (Fig. 5I). The density of goblet cells in the lung epithelia of mice primed with OVA-pulsed DC was twice as high as that in soluble OVA-primed mice and 13 times higher than that in control mice (Table I). These results demonstrate the effectiveness of priming by OVA-pulsed DC in goblet cell hyperplasia induction.

Compared with control mice (Fig. 8A), PAS staining of goblet cells in lung epithelia of mice primed with OVA₃₂₃-pulsed DC increased 6-fold over that in control mice (Fig. 8B and Table II). Using the same immunization protocol, soluble OVA induced comparable goblet cell staining (Fig. 8C). However, lung epithelia of mice primed once with OVA-pulsed DC showed higher goblet cell staining in both intensity and density (Fig. 8D) and showed a further 2-fold increase in the density of PAS⁺ cells over that in mice primed with OVA₃₂₃-pulsed DC (Table II). Control mice primed with peptide alone followed by i.t. peptide immunization exhibited no significant lung epithelia PAS staining (not shown).

Ag-specific Ab in OVA-immunized mice

In mice primed with soluble OVA or OVA-pulsed DC, anti-OVA IgG1, IgG2a, and IgE were detected. Although variable among individual mice, the IgG1 and IgE levels were higher in the sera of mice primed with OVA-pulsed DC than in those primed with soluble OVA (Fig. 9). The mean anti-OVA-specific IgG1 and IgE concentrations in mice primed with OVA-pulsed DC were 2- and 12-fold higher than those in mice primed with soluble OVA, respectively. These two groups of mice had comparable levels of serum anti-OVA-specific IgG2a. Consistent with AHR and lung inflammation results, i.t. DC priming was more effective than soluble protein priming in inducing Ag-specific IgG1 and IgE, the two Ig subclasses relevant in asthma.

View larger version (20K): [in this window] [in a new window]   FIGURE 9. Anti-OVA-specific plasma Ig levels in mouse primed with soluble OVA or OVA-pulsed DC. Mice were primed twice i.t with soluble OVA (Ova) or OVA-pulsed DC (DC) followed by OVA restimulation (protocol I). The anti-OVA-specific IgG1, IgG2a, and IgE levels in the plasma of these mice were determined by ELISA and expressed as units. The Ig levels of individual mice are shown in circles, and the mean Ig level of each group is shown as a bar. The results are pooled from two identical experiments, and there were seven mice in the soluble OVA-primed group and eight mice in the group primed with OVA-pulsed DC. The difference in IgE levels between mice primed with soluble OVA and OVA-pulsed DC was statistically significant: *, p < 0.05.

T cell migration in lungs and lung-draining lymph nodes in OVA- and OVA₃₂₃-immunized mice

Examination of the peribronchial lymph nodes showed marked mediastinal lymph node hypertrophy in mice primed with OVA-pulsed DC and boosted with OVA or primed with OVA₃₂₃-pulsed DC and boosted with OVA₃₂₃. A total of 45.9 ± 2.3 x 10⁶ and 16.0 ± 4.1 x 10⁶ cells/mouse were recovered from their mediastinal lymph nodes, respectively. In control mice, only 4–6 x 10⁶ mediastinal LNC per mouse were isolated. Thus, there is an active recruitment of cells to the lung-draining lymph nodes in mice primed with Ag-pulsed DC, resulting in 4- to 7-fold increases in cell numbers over control values. The mediastinal LNC number in mice primed with soluble OVA followed by OVA restimulation was only 14 ± 4.1 x 10⁶ cells/mouse, which equals only 30% of the cell number in the mediastinal nodes in mice primed with OVA-pulsed DC. Similarly, in mice primed and restimulated i.t. with soluble OVA₃₂₃ peptide, mediastinal LNC numbers were low (2.7 ± 1.0 x 10⁶ cells/mouse). These results demonstrate the potency of Ag-pulsed DC in recruiting lymphocytes to the lung-draining lymph nodes.

The lung mononuclear cell numbers were similar in control groups and groups immunized with OVA or OVA₃₂₃ in the presence or the absence of DC during priming. The mean cell number was 3.7 x 10⁶ cells/mouse.

Th1 and Th2 cytokines produced by lung cells and lung-draining LNC

Th1 and Th2 cytokines produced by mediastinal LNC and lung cells were measured using IFN- and IL-2 as indicators for Th1 cells and IL-4, IL-5, and IL-13 as indicators for Th2 cells. Substantial amounts of Th2 cytokines IL-4, IL-5, and IL-13 were produced by mediastinal LNC (Fig. 10A) and lung cells (Fig. 10B) from mice primed with OVA- or OVA₃₂₃-pulsed DC followed by restimulation with the priming Ag. In contrast, LNC and lung cells from control mice or mice primed and restimulated with soluble Ag produced little or no Th2 cytokine. These cytokine production results correlated with AHR and lung inflammation results, showing that mice primed by Ag-pulsed DC exhibit asthma-like disease and produce high levels of Th2 cytokines. It should be noted that in mice primed with OVA-pulsed DC, which have more severe disease, the LNC and lung cells produced more Th2 cytokines than those cells from mice primed with OVA₃₂₃-pulsed DC.

View larger version (30K): [in this window] [in a new window]   FIGURE 10. Th1 and Th2 cytokine production by mediastinal LNC and lung cells in mice primed with Ag-pulsed DC. Mice (three or four mice per group) were primed i.t. with control DC followed by saline (Sal); primed with soluble OVA or OVA-pulsed DC (DC(OVA)) followed by OVA according to protocol I; or primed i.t. with soluble OVA323 peptide (323) or OVA323 peptide-pulsed DC (DC (323)) followed by OVA323 peptide according to protocol II. LNC (A) and lung cells (B) were isolated from the mice as described in Materials and Methods. Cells were incubated for 2 days in medium with or without Ag. The culture supernatants were used for cytokine measurement by ELISA. The results show mean cytokine concentrations ± SD in culture supernatants of cells with the appropriate Ag. The results are representative of three experiments.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
DC are widely considered the major APC in the processing and presentation of foreign airway Ag and are an important participant in the pathogenesis of asthma (7). However, the abilities of DC to induce asthma in vivo have not been tested. In this report we have used purified Ag-pulsed splenic DC to prime mice to a model allergen OVA. These mice exhibited the major symptoms of asthma, which include lung eosinophilia, inflammation, goblet cell hyperplasia, AHR, and anti-Ag-specific IgE production. In addition, splenic DC are capable of inducing a Th2 response under certain experimental conditions. Priming with soluble Ag failed to induce these responses. These experiments demonstrate for the first time that DC are potent as an adjuvant in priming mice for asthma induction and support the critical role of DC as the major APC in asthma pathogenesis.

Allergic asthma is generally considered a Th2-dependent disease. In mice exhibiting asthma-like diseases, large amounts of Th2 cytokines were produced by both lung cells and draining LNC (Fig. 10). However, mediastinal LNC in mice primed with Ag-pulsed DC also produced high levels of Th1 cytokines. This Th1 stimulation may, in fact, be important for Th2 development. In T cell adoptive transfer experiments, Th1 cells not only failed to abrogate Th2-induced eosinophilia (26) and asthma (27, 28), but also exacerbated the disease when coinjected with Th2 cells (27, 28). Compared with LNC, lungs cells of mice primed with Ag-pulsed DC produced far less Th1 cytokines (Fig. 10), suggesting that there may be either a selective migration of the Th2 cells to the lungs or the presence of Th1-suppressing factors in the lung environment. The former possibility is supported by findings that Th2 cells express CCR3 and CCR4, which bind eotaxin and monocyte-derived chemokine, respectively, and that these two chemokines are produced at high levels in inflamed lungs (29). Th2 cytokines such as IL-4 have also been shown to suppress Th1 development (4). This cross-regulation of Th subsets may be operating in the lungs during inflammation, resulting in reduced Th1 numbers in the lungs. In mice primed with Ag-pulsed DC, the level of Th1 cytokines produced by lung cells was markedly different between mice immunized with OVA₃₂₃ peptide and OVA protein (Fig. 10B). A similar observation has been made by Randolph et al. in a TCR transgenic model in which Th1 cells in lungs dominate the early phase of immunization with less severe disease, and Th2 cells are predominant during the late phase of immunization with more severe disease (27). In the current study mice immunized with peptides exhibited less severe disease and thus may correspond to mice at the early phase of immunization in the adoptive transfer model of Randolph et al. and mice primed with OVA-pulsed DC correspond to the late phase of immunization in the adoptively transferred mice. The difference in the amounts of Th1 cytokine produced by lung cells in mice primed with peptide-pulsed DC vs protein-pulsed DC (Fig. 10B) can thus be explained by the difference in disease stage between these mice.

Although AHR is an important hallmark for asthma, the mechanism of AHR induction is poorly understood. In patients the severity of asthma does not correlate with the degree of esoinophilia (30). In this report AHR was observed in mice primed with OVA₃₂₃-pulsed DC, although these mice only have a low grade of eosinophilia and leukocyte infiltration in their lungs ( Figs. 6–8). This may be due to the use of OVA peptide, which is a weaker immunogen with a single T epitope and without dominant B cell epitopes, thus limiting the participation of both T and B cells and the formation of immune complexes. Wilder et al. (31) and more recently Tournoy et al. (32) reported similar findings in mice immunized with limited frequency or Ag dose. In all cases relatively weak immune responses have been elicited. The results suggest that AHR is among the most readily inducible symptom in asthma, and its occurrence is independent of eosinophilia and severe lung inflammation.

In contrast to human asthma, asthma-like diseases in the mouse do not require the participation of IgE (2, 3, 31, 32, 33). The presence of B cells is also not a prerequisite for the induction of asthma-related symptoms (34). The finding that AHR, eosinophilia, and lung inflammation were induced in mice primed with OVA₃₂₃ while no detectable Ag-specific IgE was detected is consistent with the literature. However, AHR in mice have been shown to be enhanced by the passive transfer of Ag-specific IgG1 and IgE (35). The report suggests that the presence of high levels of anti-OVA IgG1 and IgE in OVA-immunized mice may be a significant factor for the higher lung inflammatory response and AHR in OVA-compared with OVA₃₂₃-immunized mice (Figs. 1C and 3–5; Table I and II).

Peptides that have a single or limited number of T epitopes are useful for studying T cell stimulation. T epitope peptides or altered peptide ligands have been used to modulate Th subset responses (5, 36, 37, 38). This cannot be achieved with intact proteins, which usually contain multiple T epitopes. Furthermore, peptides are important in studying T cell responses in TCR transgenic mice (10, 36, 37, 38, 39). In the past, peptides have been of limited use in asthma studies. Renz et al. used aerosolized OVA₃₂₃ to immunize mice. A weak induction of AHR measurable only by electrical field stimulation of tracheal smooth muscle preparations, but not by whole body plethysmography was observed (14, 40). To amplify the T cell response to peptides, DC have been used in this study because they have been shown to be potent in priming T cells to peptides (5, 10, 41) in part due to the presence of large number of empty MHC class II molecules on their surfaces ready for peptide loading (42). In addition, peptides have been delivered i.t. instead of using aerosolization, thus rendering airway peptide immunization economically feasible. The results showed that peptides when presented by i.t. injected DC are efficient in stimulating asthma-related symptoms such as AHR and inflammation. This successful induction of asthma-related symptoms by peptide-pulsed DC provides a simple method for studying the functions of T epitope peptides and altered peptide ligands in asthma.

Direct studies of asthma induction with mouse lung DC are difficult because of the low numbers of lung DC (8, 9). In lung digests, DC numbers were estimated to be 4 x 10⁴/mouse (8), and the yield of purified DC was only 2.5 x 10³/mouse (9). These DC numbers fall far short of the numbers required for priming mice for AHR induction (3–5 x 10⁵/mouse; Figs. 1 and 6) or for footpad priming of mice to protein Ag (43). In this study splenic DC were efficient in priming mice to manifest an asthma-like disease state. The results suggest that the lung environment modulate DC to stimulate Th2-biased responses. Splenic DC have been shown to be composed of two phenotypically and functionally distinct populations, a smaller CD8⁺CD11c⁺ population and a larger CD8^-CD11c⁺ population, which stimulate biased Th1 and biased Th2 responses, respectively (22) (Fig. 2E). The CD8⁺ population has further been shown to produce much more IL-12 than the CD8^- population (44). Despite the presence of strongly Th1-directing DC, our results showed that these splenic DC preparations were capable of inducing asthma-like diseases and the induction of Th2 cell development in both draining lymph nodes and lungs, as shown by cytokine production measurements (Fig. 10). The presence of IL-12-producing Th1-directing CD8⁺CD11c⁺ DC during Ag priming may further stimulate, rather than inhibit, Th2 development in the lungs and lung-draining lymph nodes. In Th2-dependent schistosome egg-induced granuloma formation, IL-12 accentuates rather than abrogates the inflammation, indicating that Th1-stimulating cytokines also enhance Th2-dependent diseases (45).

In i.t. priming for both OVA₃₂₃ and OVA protein responses and for asthma induction, DC are clearly more efficient than soluble peptide or protein despite the much lower amount of Ag epitopes introduced by DC injection. Based on the estimate of 5 x 10⁵ surface I-A molecules/DC (42), the total amount of peptide bound by I-A molecules on 5 x 10⁵ injected DC is 0.4 pmol, which is far less than the 50 nmol of OVA₃₂₃ or 100 µg of OVA protein (2.3 nmol) used in soluble Ag priming. Furthermore, our results showed that very small amounts of fluorescent OVA are associated with the injected DC (<2.2 µg/10⁶ DC). The poor priming with soluble Ag suggests either that the Ag uptake, processing, or loading into the MHC class II molecules on lung DC are inefficient or that the great majority of injected Ag are degraded by the alveolar macrophages and proteases in the lungs.

The functional potencies of i.t. injected, Ag-pulsed DC in inducing asthma-related symptoms reported in this study further support the thesis that i.t. injected splenic DC possess the machinery for crossing the lung epithelia and home to the T cell zone of the draining lymph for T cell priming (10, 15, 46). This DC property of migration to the draining lymph nodes will be useful in studying T cell stimulation in pulmonary immune responses and will allow the identification of cytokines, chemokines, adhesion molecules, and accessory molecules involved in primary and secondary responses in the lung and the elucidation of mechanisms of asthma development. The model system described in this report can also be used to identify the role of T cell subsets in lung inflammation and to examine the correlation between biased Th2 stimulation and asthma. Furthermore, the ability of injected DC to act as a shuttle between the target organs and their draining lymph nodes will be useful for the delivery a variety of factors to the lung draining lymph nodes for functional studies. In gene-gun DNA immunization studies, DC have been shown to be the primary transfected population responsible for Ag presentation, and they are efficient in eliciting long-lasting Ag-specific immune responses (47, 48, 49, 50). These findings suggest the usefulness of in vitro transfected DC in modifying DC Ag presentation or T cell responses in pulmonary inflammation. Ag priming with i.t. injected in vitro-transfected DC can be a powerful fool for studying the functions of receptors, costimulatory and adhesion molecules, cytokines, or chemokines in asthma.

	Acknowledgments

 
We thank Kulsoom Ghias, Julie M. Weed, Prashanth Mally, and Ji Xian Zhang for technical assistance, and Dr. Y. B. Kim for advice.

	Footnotes

 
¹ This work was supported in part by American Heart Association Grant 9950268N, a grant from the American Heart Association Virginia Affiliate, and National Institutes of Health Grants P50AR45222 and RO1CA34546.

² Address correspondence and reprint requests to Dr. Sun-sang J. Sung, Department of Internal Medicine, Box 800412, University of Virginia Health Sciences Center, Charlottesville, VA 22908.

³ Abbreviations used in this paper: AHR, airway hyperresponsiveness; BAL, bronchoalveolar lavage; DC, dendritic cells; i.t., intratracheal(ly); LNC, lymph node cell(s); OVA₂₇₃, OVA peptide 273–288; OVA₃₂₃, OVA peptide 323–339; PAS, periodic acid-Schiff; Penh, enhanced pause.

Received for publication April 7, 2000. Accepted for publication October 12, 2000.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Mannino, D. M., D. M. Homa, C. A. Pertowski, A. Ashizawa, L. L. Nixon, C. A. Johnson, L. B. Ball, E. Jack, D. S. Kang. 1998. Surveillance for asthma, United States, 1960–1995. Morbid. Mortal. Wkly. Rep. 47:1.[Medline]
2. Bochner, B. S., B. J. Undem, L. M. Lichtenstein. 1994. Immunological aspects of allergic asthma. Annu. Rev. Immunol. 12:295.[Medline]
3. Wills-Karp, M.. 1999. Immunologic basis of antigen-induced airway hyperresponsiveness. Annu. Rev. Immunol. 17:255.[Medline]
4. Seder, R. A., W. E. Paul. 1994. Acquisition of lymphokine producing phenotype by CD4⁺ T cells. Annu. Rev. Immunol. 12:635.[Medline]
5. Constant, S. L., K. Bottomly. 1997. Induction of Th1 and Th2 CD4⁺ T cell responses: the alternative response. Annu. Rev. Immunol. 15:297.[Medline]
6. Stumbles, P. A., J. A. Thomas, C. L. Pimm, P. T. Lee, T. J. Venaille, S. Proksch, P. G. Holt. 1998. Resting respiratory tract dendritic cells preferentially stimulate T helper cell type 2 (Th2) responses and require obligatory cytokine signals for induction of Th1 immunity. J. Exp. Med. 188:2019.[Abstract/Free Full Text]
7. McWilliam, A. S., D. J. Nelson, P. G. Holt. 1995. The biology of airway dendritic cells. Immunol. Cell Biol. 73:405.[Medline]
8. Pollard, A., M. F. Lipscomb. 1990. Characterization of murine lung dendritic cells: similarities to Langerhans cells and thymic dendritic cells. J. Exp. Med. 172:159.[Abstract/Free Full Text]
9. Masten, B. J., M. F. Lipscomb. 1999. Comparison of lung dendritic cells and B cells in stimulating naive antigen-specific T cells. J. Immunol. 162:1310.[Abstract/Free Full Text]
10. Lambrecht, B. N., R. A. Pauwels, B. F. de St. Groth.. 2000. Induction of rapid T cell activation, division, and recirculation by intratracheal injection of dendritic cells in a TCR transgenic model. J. Immunol. 164:2937.[Abstract/Free Full Text]
11. Hoyne, G. F., R. E. O’Hehir, D. C. Wraith, W. R. Thomas, and J. R. Lamb. 1993. Inhibition of T cell and antibody responses to house dust mite allergen by inhalation of the dominant T cell epitope in naive and sensitized mice. J. Exp. Med. 1783.
12. Tian, J., M. A. Atkinson, M. Clare-Salzler, A. Herschenfeld, T. Forsthuber, P. V. Lehmann, D. L. Kaufman. 1996. Nasal administration of glutamate decarboxylase (GAD65) peptides induces Th2 responses and prevents murine insulin-dependent diabetes. J. Exp. Med. 183:1561.[Abstract/Free Full Text]
13. Wolvers, D. A. W., C. J. J. Coenen-de Roo, R. E. Mebius, M. J. F. van der Cammen, F. Tirion, A. M. M. Miltenburg, G. Kraal. 1999. Intranasal induced immunological tolerance is determined by characteristics of the draining lymph nodes: studies with OVA and human cartilage gp-39. J. Immunol. 162:1994.[Abstract/Free Full Text]
14. Renz, H., K. Bradley, G. L. Larsen, C. McCall, E. W. Gelfand. 1993. Comparison of the allergenicity of ovalbumin and ovalbumin peptide 323–339: differential expansion of V-expressing T cell populations. J. Immunol. 151:7206.[Abstract]
15. Havenith, C. E. G., P. P. M. C. van Miert, A. J. Breedijk, R. H. J. Beelen, E. C. M. Hoefsmit. 1993. Migration of dendritic cells into the draining lymph nodes of the lung after intratracheal instillation. Am. J. Respir. Cell Mol. Biol. 9:484.
16. Swiggard, R. M., R. M. Nonacs, M. D. Witmer-Pack, R. M. Steinman, and K. Inaba. 1992. Enrichment of dendritic cells by plastic adherence and EA rosetting. In Current Protocols in Immunology. J. E. Coligan, A. M. Kruisbeek, D. H. Margulies, E. M. Shevach, and W. Strober, eds. John Wiley & Sons, New York, p. 3.7.1.
17. Sung, S. J., C. Y. Guo, J. M. Weed. 1997. Monoclonal antibodies against human dendritic cell-like peripheral blood monocytes activated by granulocyte/monocyte-colony-stimulating factor plus interleukin 4. Cell. Immunol. 182:113.[Medline]
18. Hamelmann, E., J. Schwarze, K. Takeda, A. Oshiba, G. L. Larsen, C. G. Irvin, E. W. Gelfand. 1997. Noninvasive measurement of airway hyperresponsivenss in allergic mice using barometric plethysmography. Am. J. Respir. Crit. Care. Med. 156:766.[Abstract/Free Full Text]
19. Vijayaraghavan, R., M. Schaper, R. Thompson, M. F. Stock, Y. Alarie. 1993. Characteristic modifications of the breathing pattern of mice to evaluate the effects of airborne chemicals on the respiratory tract. Arch. Toxicol. 67:478.[Medline]
20. Kamochi, M., F. Kamochi, Y. B. Kim, S. Sawh, J. M. Sanders, I. Sarembock, S. Green, J. S. Young, K. Ley, S. M. Fu, et al 1999. P-selectin and ICAM-1 mediate endotoxin-induced neutrophil recruitment and injury to the lung and liver. Am. J. Physiol. 277:L310.[Abstract/Free Full Text]
21. Krinzman, S. J., G. T. de Sanctis, M. Cernadas, D. Mark, Y. Wang, J. Listman, L. Kobzik, C. Donovan, K. Nassr, I. Katona, et al 1996. Inhibition of T cell costimulation abrogates airway hyperresponsiveness in a murine model. J. Clin. Invest. 98:2693.[Medline]
22. Maldonado-Lopez, R., T. de Smedt, P. Michel, J. Godfroid, B. Pajak, C. Heirman, K. Thielemans, O. Leo, J. Urbain, M. Moser. 1999. CD8⁺ and CD8^- subclasses of dendritic cells direct the development of distinct T helper cells in vitro. J. Exp. Med. 189:587.[Abstract/Free Full Text]
23. Kips, J. C., G. J. Bruselle, G. F. Joos, R. A. Peleman, J. H. Tavernier, R. R. Devos, R. A. Pauwels. 1996. Interleukin-12 inhibits antigen-induced airway hyperresponsiveness in mice. Am. J. Respir. Crit. Med. 153:535.[Abstract]
24. Shimonkevitz, R., J. Kappler, P. Marrack, H. Grey. 1983. Antigen recognition by H-2-restricted T cells. I. Cell-free antigen processing. J. Exp. Med. 158:303.[Abstract/Free Full Text]
25. Vidard, L., K. L. Rock, B. Benacerraf. 1992. Diversity in MHC class II ovalbumin T cell epitopes generated by distinct proteases. J. Immunol. 149:498.[Abstract]
26. Li, L., Y. Xia, A. Nguyen, L. Feng, D. Lo. 1998. Th2-induced eotaxin expression and eosinophilia coexist with Th1 responses at the effector stage of lung inflammation. J. Immunol. 161:3128.[Abstract/Free Full Text]
27. Randolph, D. A., C. J. L. Carruthers, S. J. Szabo, K. M. Murphy, D. D. Chaplin. 1999. Modulation of airway inflammation by passive transfer of allergen-specific Th1 and Th2 cells in a mouse model of asthma. J. Immunol. 162:2375.[Abstract/Free Full Text]
28. Hansen, G., G. Berry, R. H. DeKruyff, D. T. Umetsu. 1999. Allergen-specific Th1 cells fail to counterbalance Th2 cell-induced airway hyperreactivity but cause severe airway inflammation. J. Clin. Invest. 103:175.[Medline]
29. Lloyd, C. M., T. Delaney, T. Nguyen, J. Tian, C. Martinez-A, A. J. Coyle, J.-C. Gutierrez-Ramos. 2000. CC chemokine receptor (CCR) 3/eotaxin is followed by CCR4/monocyte-derived chemokine in mediating pulmonary T helper lymphocyte type 2 recruitment after serial antigen challenge in vivo. J. Exp. Med. 191:265.[Abstract/Free Full Text]
30. McFadden, E. R.. 1994. Asthma: morphologic-physiologic interactions. Am. J. Respir. Crit. Care Med. 150:S23.
31. Wilder, J. A., D. D. S. Co