INTRODUCTION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

A significant number of asthmatic individuals are atopic and have respiratory allergies that may act as stimuli for asthma attacks (1). In such individuals, inhalation of allergens such as mold spores, pollens, animal dander, and house dust can elicit wheezing, dypsnea, eosinophil influx into the lung, and airway hyperresponsiveness (AHR) (2, 3).

IgE antibody and activated T lymphocytes are thought to play fundamental yet separate roles in the temporal phases of responses known to exist in allergic asthma (4). Antigen challenge causes an immediate hypersensitivity reaction (early asthmatic response [EAR]) via the crosslinking of IgE molecules on the surface of mast cells and subsequent release of histamine, prostaglandin-D₂, leukotriene-C₄, and other mediators responsible for inflammation and bronchoconstriction (5, 6). Between 2 and 8 h after this event, there often occurs a more severe and prolonged reaction known as the late asthmatic response (LAR), which is characterized by increased mucus production, prolonged bronchoconstriction, AHR to nonspecific agonists, and airway inflammation (7).

Recent studies have shown increased numbers of antigen-specific CD4⁺ T lymphocytes in the bronchoalveolar lavage fluid (BALF) of allergic asthmatic individuals (8, 9), which produce cytokines typical of the Th2 lymphocyte phenotype (10). Th2 cells promote immediate-type allergic responses through the production of interleukin-4 (IL-4), which stimulates IgE antibody production (4, 11), as well as IL-3 and IL-5, which facilitate the growth, maturation, and recruitment of eosinophils to the lung (12).

Adoptive transfer techniques have been used in a number of studies to investigate the mechanisms of pulmonary allergy (17, 18). Experimental hypersensitivity pneumonitis was successfully transferred to naive guinea pigs (19) and mice (20) by activated T cells. Intraperitoneal injection of antigen-primed lymphocytes promoted cholinergic AHR and late airway responses in a Brown Norway (BN) rat model of allergic pulmonary disease (21, 22). In addition, experiments involving the passive transfer of monoclonal IgE antibody resulted in immediate, but not late-phase, responses to ovalbumin challenge in rats (23). To date, no studies have addressed the transfer of both immediate and delayed airway responses, pulmonary inflammation, and histologic changes to the same group of animals.

In a rodent model of pulmonary allergy, (BN) rats sensitized with house dust mite (HDM) and later challenged with antigen show EAR, increases in pulmonary inflammatory cells, and AHR as compared with adjuvant-treated control animals (24). The inflammation peaks between 1 and 2 d after challenge with HDM and AHR peaks at 1 d, but persists for up to 7 d after challenge. In the present study, we hypothesized that activated T lymphocytes potentiate the LAR, whereas serum from actively sensitized donors elicits the EAR. Serum or lymphocytes were taken from HDM-sensitized rats to determine whether they could transfer allergic responses (antigen-induced immediate bronchoconstriction, pulmonary inflammation, histopathologic changes, and AHR) to recipient rats after challenge with HDM antigen. Use of the adoptive transfer technique allowed us to evaluate separately the contributions of both humoral and cell-mediated immunity in the pathophysiology of allergic lung disease.

	METHODS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Animals

Inbred female BN rats (strain BN/Ss NHsd), 8 to 10 wk old and ranging in weight from 160 g to 170 g, were purchased from Harlan Sprague- Dawley, Inc. (Indianapolis, IN). Male Sprague-Dawley rats 11 to 12 mo old and ranging in weight from 300 g to 400 g were obtained from Charles River, Inc. (Raleigh, NC) for use as recipients in the passive cutaneous anaphylaxis (PCA) assay. All animals were housed in animal facilities with air filtered by high-efficiency particulate accumulators, were fed rat chow and water ad libitum, and were maintained according to the animal care and use committee guidelines of our institutions. Randomly selected animals, tested serologically upon arrival, and sentinels monitored throughout the study, were free of sendai virus, pneumonia virus, mouse hepatitis virus, and a variety of other rodent viruses, as well as Mycoplasma sp. Rats were also monitored for and found to be free of ectoparasites and endoparasites.

Antigen Preparation

Semipurified dust mite antigen from Dermatophagoides farinae and D. pteronyssinuss (Greer Laboratories, Lenoir, NC) was obtained from ground, whole-bodied mites after defatting, extraction in 0.125 M ammonium bicarbonate, and dialysis against distilled water. Antigen fractions from each species of mite were mixed equally and stored at 80° C until use. Total protein concentration was 1.76 mg/ml by Bradford assay. The amount of group I allergen as assessed by the vendor through immunoassay was 126.7 µg/mg protein.

Immunization

Rats were immunized subcutaneously with 88 µg HDM antigen suspended in aluminum hydroxide gel adjuvant (Alhydrogel; Accurate Antibodies Inc., St. Louis, MO). Simultaneously, 0.5 ml of Bordetella pertussis vaccine (IAF Biovac Inc., Montreal, Canada), containing 1 × 10⁹ heat-killed organisms, was injected intraperitoneally. Control animals received injections of both adjuvants without the antigen extract. Two weeks after-sensitization, donor animals (antigen-sensitized and adjuvant controls) were lightly anesthetized by halothane inhalation (Aldrich Chemical Co., Milwaukee, WI) and challenged via the trachea with 300 µl of 0.90% sterile saline solution (Fujisawa USA Inc., Deerfield, IL) containing 17.6 µg HDM antigen.

Experimental Design

Preliminary experiments had shown that although antigen challenge caused significant airway inflammation and AHR, which peaked at 1 d after challenge, the secondary immune response to HDM antigen, as assessed by serum antibody and lymphocyte proliferation, was maximal at 3 d after antigen challenge. We therefore chose 3 d after challenge as the time at which to obtain tissue from the donor animals, but measured pathophysiologic responses in the recipient animals 1 d after challenge. Additional experiments examined the optimal time between transfer and challenge in recipient animals. In these experiments we waited 1,3,7, and 10 d between transfer and challenge, and found that responses after cell transfer were maximized at 3 d. The serum-transfer responses were also maximal at this time point, although the effect remained for at least 7 d. Hence, we chose the 3-d time point after transfer to measure immediate responses to antigen challenge.

Adoptive Transfer of Cells and Serum

Three days after challenge, donor animals (six antigen-sensitized and six adjuvant-treated controls) were tested in the airway reactivity system prior to being euthanized with 150 mg/kg of sodium pentobarbital (Abbott Laboratories, Chicago, IL). Rats were bled by cardiac puncture and serum was obtained by centrifugation at 4° C for 15 min at 2,500 rpm. Peripheral lymph nodes (axillary, brachial, popliteal, inguinal, gluteal, and posterior mediastinal) were removed.

Lymph-node-cell suspensions were prepared with a Stomacher 80 laboratory blender (Seward Inc., London, UK). Red blood cells were lysed with 0.89% ammonium chloride, and cell suspensions were pooled within each group. Lymphocytes were washed three times in complete medium (RPMI containing 10% fetal calf serum [FCS], 5% penicillin/ streptomycin, and 10⁵ M -mercaptoethanol), and cells from HDM-sensitized rats were incubated overnight at 37° C in complete medium containing 1.76 µg/ml HDM, while cells from adjuvant-treated control rats were incubated in complete medium containing 1.76 µg/ml sterile bovine serum albumin (BSA; Sigma Chemical Co., St. Louis, MO). The following day, lymphocytes were washed twice with complete medium. Viability was assessed by trypan blue dye exclusion, and cells were resuspended to 4.0 × 10⁷ cells/ml in sterile saline. Naive recipients received an intraperitoneal injection of 1.0 ml of the lymphocyte cell suspension from actively sensitized donors or adjuvant-treated controls.

Serum was collected by centrifugation of blood from donor rats, and was pooled within groups. Naive recipient rats received an intravenous injection of 0.5 ml serum from actively sensitized donors or adjuvant controls.

Total IgE and HDM-Specific IgE/IgG ELISA

Levels of total and HDM-specific IgE were determined by enzyme-linked immunosorbent assay (ELISA) of serum specimens obtained from both actively sensitized donor animals and adjuvant-treated control donor animals. For the assay, 96-well polystyrene plates (Costar, Cambridge, MA) were coated with 5 µg/ml mouse monoclonal antibody to rat IgE (Harlan Bioproducts, Inc., Indianapolis, IN) or with 17.6 µg/ml HDM, and were incubated overnight at 4° C. Plates were washed five times with 0.1 M phosphate-buffered saline containing 0.05% Tween-20 (PBS-T), and were treated successively with: (1) 150 µl/well PBS + 1.0% BSA (blocking buffer); (2) 100 µl/well rat serum diluted 1:10 in coating buffer, and rat IgE myeloma following a standard concentration curve ranging from 162.5 to 20,000 ng/ml (Harlan Bioproducts); (3) biotin-labeled mouse monoclonal antibody to rat IgE (Harlan Bioproducts) diluted 1:1,000; and (4) phosphatase-labeled streptavidin diluted 1:1,500 (Kirkegaard and Perry Laboratories, Gaithersburg, MD). Plates were washed five times with PBS-T and were incubated at 37° C for 1 h between each step. p-Nitrophenyl phosphate disodium (Kirkegaard and Perry Laboratories) was added as a substrate for streptavidin, and plates were developed for 15 min at room temperature and read at 405 nm with a Thermomax ELISA plate reader (Molecular Devices, Menlo Park, CA).

PCA Assay

HDM-specific cytophilic antibody in donor and recipient rats was also measured through a PCA assay. Sprague-Dawley rats were anesthetized with 35 ml/kg sodium pentobarbital, were shaved, and were passively sensitized with a subcutaneous injection of 100 µl of diluted, pooled serum (1:10, 1:50, and 1:100 dilutions) obtained from sensitized and control donors as well as recipients of cells and serum. Two days later the animals were injected intravenously with 0.5 ml of a solution of 88 µg/ml HDM in 10 mg/ml Evans Blue dye (Sigma Chemical Co.). Extravasation of the dye was measured 30 min after challenge.

Flow Cytometry

After overnight incubation, and prior to transfer to naive recipients, pellets containing 1 × 10⁶ lymph-node cells were incubated at 4° C with optimized concentrations of fluorescent antibodies to the following cell-surface markers: CD4 (phycoerythrin [PE] conjugated), CD8 (fluorescein isothiocyanate [FITC] conjugated), CD3 (PE-conjugated) (PharMingen, San Diego, CA), and OX12 (B220, FITC-conjugated) (Serotec Inc., Indianapolis, IN). Thirty minutes later, cells were washed twice with complete medium and resuspended in 1.0 ml of flow-cytometry buffer (1% BSA, 0.1% sodium azide in 1 × PBS). Flow-cytometric analyses were performed with a Coulter-Epics XL-MXL flow cytometer (Coulter Electronics, Hialeah, FL). Dead cells and debris were excluded by gating on forward and 90° light-scatter measurements representing the lymphocyte population. Control cells were labeled with either IgG_2a-PE-conjugated, or IgG₁-FITC-conjugated antibodies to gate out nonspecific binding.

Immediate Bronchoconstriction Responses to HDM

Three days after transfer, recipients of either cells or serum were placed in a whole-body plethysmograph (Buxco Electronics, Troy, NY) equipped with a pneumotachograph and pressure transducer to monitor pulmonary-function responses. The index of bronchoconstriction that was measured was based on the wave shape of the box flow signal. The box flow signal reflected the difference between the animal's nasal flow and the flow due to chest movement (thoracic flow). Prior to antigen challenge, the thoracic flow (air into the box) and nasal flow (air out of the box) were equal and in phase. When obstructions in the airways had to be overcome because of bronchoconstriction, a difference in magnitude and phase appeared between the nasal flow and the thoracic flow. The time delay (enhanced pause) between nasal and thoracic flows during response to challenge was determined to be proportional to specific airway resistance (SGaw) (25).

In the experiments described here, baseline readings were measured for 15 min prior to HDM challenge. Animals were then removed from the plethysmograph, lightly anesthetized with halothane, intratracheally instilled with 17.6 µg HDM in 300 µl saline, and placed in the plethysmograph for a further 20 min to measure immediate bronchoconstriction in response to HDM challenge. Immediate responses to instilled antigen were compared among cell and serum recipients and the donor animals.

AHR

Because antigen challenge in sensitized animals resulted in pulmonary inflammation and AHR that peaked at 24 h after antigen challenge, this time point was selected to measure pathophysiologic endpoints in recipient animals. One day after challenge, recipient rats were anesthetized with an intraperitoneal injection of urethane (1.0 g/kg) (Sigma Chemical Co.). A transoral cannula was placed in the trachea, and a saline-filled polyethylene PE-50 catheter was inserted into the jugular vein. The animal was then paralyzed with an intramuscular injection of 2.5 mg/kg succinylcholine chloride (Glaxo-Burroughs Wellcome, Research Triangle Park, NC), placed supine into a whole-body pressure plethysmograph, and ventilated with a small-animal respirator at a frequency of 90 breaths/minute (bpm). After an initial baseline measurement of airway pressure during periods of no infusion followed by intravenous infusion of saline (0.04 ml/min), rats were infused with acetylcholine chloride (Sigma Chemical Co.) at a dose adjusted for each animal (0.25 mg/g body weight per ml) via the jugular catheter for 10 min, with a total delivered volume of 2.48 ml. During the infusion, the flow rate of the peristaltic pump doubled every 2 min, starting at 0.04 ml/min and ending at 0.64 ml/min, and airway pressure was measured with a pressure transducer located distal to the transoral catheter.

Bronchoalveolar Lavage

Bronchoalveolar lavage (BAL) was performed immediately after airway reactivity testing. Animals were bled by cardiac puncture, the trachea was cut, and a cannula was inserted and tied off with suture. The left lung was cannulated and inflated with 10% formalin (Fisher Scientific, Fair Lawn, NJ). The remaining lobes were lavaged three times via the tracheal cannula, using 29.75 ml/kg warmed saline. Total cell counts were made through trypan blue dye exclusion, with a hemocytometer. Differential cell counts were made on a glass slide that was prepared with a Cytospin Model II (Shandon, Pittsburgh, PA) and stained with Diff-Quik (American Scientific Inc., Sewickley, PA). At least 100 cells were counted in duplicate for each specimen.

Histology Assessment

Tissues were embedded in paraffin, sliced in the midsagittal plane in 4-µm-thick sections, and stained with hematoxylin-eosin (H&E) for light microscopy. Histology slides from each animal were read by a veterinary pathologist working under contract, and were scored according to the relative severity of inflammatory, degenerative, and proliferative changes. Specifically, lung sections were scored in nine separate categories for the degree of alveolitis, edema, hemorrhage, lymphoid hyperplasia, and peribronchial and perivascular infiltration of mononuclear cells and leukocytes, with a score of 1 representing a mild and a score of 5 a severe degree of change. The total scores for each animal were calculated, and the histologic changes were expressed as the mean ± SE for each treatment group.

Statistical Analysis

Immediate-bronchoconstriction and airway-pressure data were analyzed by comparing adjuvant-treated control groups with antigen-treated experimental groups, using repeated-measures analysis of variance (ANOVA). Group differences were considered significant if the test statistic type I error was < 0.05. 

Statistical comparison of histology scores and total lavageable eosinophil, neutrophil, macrophage, and lymphocyte numbers in antigen versus adjuvant (control) groups was done with Student's t test. The form of the t test used was determined by the results of an F test for the homogeneity of variances. ELISA data were analyzed by comparing serum antibody concentrations of antigen groups and adjuvant groups, using Student's t test for equal variances. Group differences were considered significant if the statistic type I error was < 0.05.

	RESULTS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Immunoglobulin ELISAs and PCA Titers

Antigen-specific IgG and IgE titers in the serum of HDM- sensitized donors were significantly higher than titers in serum from adjuvant controls (Table 1). Total IgE in the serum from HDM-sensitized donors was also significantly higher than that of adjuvant donors. In addition, passive cutaneous anaphylaxis revealed dye extravasation at sites of injection of serum from HDM-sensitized donors at serum dilutions > 1:100. No PCA reactions were observed in the sera from adjuvant-treated donors or in the sera from recipients of serum and cells.

Flow Cytometry

Flow-cytometric analysis after overnight culture revealed that the majority of transferred cells were T cells; the cell percentages were CD4⁺ = 67.96 ± 0.58%, CD8⁺ = 5.7 ± 0.61%, CD3⁺ = 75.14 ± 1.9%, and OX12 (B cell) 23.62 ± 0.61%.

Immediate Bronchoconstriction

Donor rats sensitized to HDM antigen experienced significant bronchoconstriction following intratracheal instillation of HDM as compared with adjuvant-treated control rats (Figure 1A). Enhanced pauses of sensitized rats were significantly greater than those of adjuvant-treated controls following intratracheal antigen provocation. An average enhanced pause of 2.0 ± 0.06 was observed for sensitized rats and 1.0 ± 0.03 for adjuvant controls. Recipients of serum from donor rats sensitized to HDM experienced significant increases in bronchoconstriction responses following intratracheal challenge with HDM, as compared with recipients of serum from adjuvant-treated controls (Figure 1B). An average enhanced pause of 1.7 ± 0.63 was observed for recipients of serum from sensitized donors, whereas the average enhanced pause of recipients of serum from adjuvant-treated controls was 1.0 ± 0.27. When serum was heated to 57° C prior to transfer to recipients, this effect was lost (data not shown).

View larger version (31K): [in this window] [in a new window]   Figure 1.   Antigen-induced immediate responses in sensitized and adjuvant-treated control donors (A), serum recipients (B), and cell recipients (C ). Animals were placed in a plethysmograph for 15 min to assess baseline pulmonary function, and were monitored for 20 min after intratracheal challenge with 17.6 µg HDM. *p < 0.05, adjuvant versus antigen groups as determined by ANOVA. Data are presented as means ± SE, n = 5 to 9 animals per treatment group.

Recipients of lymphocytes from HDM-sensitized donor rats or adjuvant controls did not experience significant bronchoconstriction in response to intratracheal HDM provocation (Figure 1C). Mean enhanced pauses were not statistically different for the different cell-recipient groups following antigen challenge (1.2 + 0.49 for recipients of lymphocytes from adjuvant treated-donors; 1.3 ± 0.72 for recipients of lymphocytes from antigen-treated donors).

AHR

To illustrate that the donor animals had hyperreactive airways at 3 d after antigen challenge, airway pressure was measured during infusion of acetylcholine (ACh) prior to killing and extraction of lymph nodes and serum. As the ACh dose increased, airway pressure increased to a greater extent in sensitized donors than in adjuvant-treated control donors treated in exactly the same way, including antigen challenge (Figure 2A). Airway pressure measured during intravenous infusion of ACh 1 d after HDM challenge was not significantly different in recipients of serum from sensitized donor rats and recipients of serum from adjuvant-treated control donors (Figure 2B). Furthermore, the airway pressures measured during infusion of Ach in both recipient groups did not differ significantly from average baseline airway pressure measured during infusion of saline.

View larger version (23K): [in this window] [in a new window]   Figure 2.   Airway responsiveness to ACh infusion in sensitized and adjuvant-treated control donors (A), serum recipients (B), and cell recipients (C ). Airway pressure was measured in rats during intravenous infusion of ACh. *p < 0.05, adjuvant versus antigen groups as determined by ANOVA. Data are presented as means ± SE, n = 5 to 9 animals per treatment group.

In contrast, recipients of lymphocytes from HDM-sensitized rats experienced significant AHR (increased airway pressure) upon ACh infusion at 1 d after challenge with HDM, as compared with recipients of lymphocytes from adjuvant-treated control rats (Figure 2C).

Airway Inflammation

HDM-sensitized donor rats had significant increases in alveolar macrophages (AM), lymphocytes, and eosinophils in their BALF as compared with that of adjuvant-treated control donor rats (Figure 3A). Eosinophil numbers were 6.6 × 10⁵ in the BALF of sensitized donors and 2.4 × 10⁴ in the BALF of adjuvant-treated control donors, which represented 67% and 14%, respectively, of the total lavageable cells. Recipients of serum from both adjuvant-treated and HDM-sensitized donors had some inflammatory infiltrate (eosinophils and neutrophils) in the BALF that appeared to be a result of the serum injection as opposed to antigen-specific inflammation, since there was no significant difference in macrophage, neutrophil, lymphocyte, or eosinophil numbers between recipients of sensitized or adjuvant-control serum (Figure 3B).

View larger version (28K): [in this window] [in a new window]   Figure 3.   Differential cell counts in BALF of sensitized and adjuvant-treated control donors (A), serum recipients (B), and cell recipients (C ). BALF cells were cytospin-prepared and stained for differential cell counts. *p < 0.05, and p < 0.01 in (A). *p < 0.01 and p < 0.05 in (C ). AM = alveolar macrophage; EOS = eosinophil; LYM = lymphocyte; PMN = neutrophil. Data are presented as means ± SE, n = 5 to 9 animals per treatment group.

Recipients of lymphocytes from HDM-sensitized donor rats had significantly higher numbers of eosinophils and lymphocytes in their BALF at 1 d after challenge with HDM than did recipients of lymphocytes from adjuvant-treated control donors (Figure 3C).

Histopathology

Histology specimens from all animals were scored for the degree of alveolitis, edema, hemorrhage, and peribronchial and perivascular infiltrate, with a score of 1 indicating mild and a score of 5 indicating severe change. Recipients of adjuvant-control serum and adjuvant cells displayed mild pathologic changes related to the adoptive transfer injection and antigen challenge, with a total score for all animals in each group of 27 and 25, respectively. In contrast, histology specimens from recipients receiving serum and cells from antigen-sensitized donors had total scores of 32 and 41, respectively. The mean scores for each treatment group are depicted in Figure 4, and show that the cell recipients had statistically greater histologic changes than did any other group. Figures 5A through D depict examples of small-airway histology findings in one animal from each treatment group. All histology sections showed a modest level of alveolitis and edema (Figure 5A through D), which were due to a combination of the adoptive transfer injections and antigen challenge, since lung sections from nontreated animals had no discernable lesions. In general however, the degree of alveolitis, edema, and inflammatory infiltrate around the airways was most severe in animals that received cells from sensitized donors (Figure 5D).

View larger version (14K): [in this window] [in a new window]   Figure 4.   Histopathologic changes in recipients of serum or cells from antigen-sensitized or adjuvant-treated control rats. Lung sections were scored for degree of alveolitis, edema, hemorrhage, lymphoid hyperplasia, and peribronchial and perivascular infiltration of mononuclear cells and leukocytes, with a score of 1 indicating mild and a score of 5 indicating severe change. The total scores (out of nine categories) for each animal were calculated and the histologic changes expressed as the mean ± SE for each treatment group. Data were analyzed by ANOVA. *Significant differences at p < 0.05, n = 5.

View larger version (105K): [in this window] [in a new window]   View larger version (103K): [in this window] [in a new window]   View larger version (101K): [in this window] [in a new window]   View larger version (124K): [in this window] [in a new window]   Figure 5.   Histology sections from lungs of rats 24 h after antigen challenge. (A) Recipient of serum from adjuvant-treated control donors (B) Recipient of serum from sensitized donors (C ) recipient of cells from adjuvant control donors, and (D) recipient of cells from sensitized donors H&E: original magnification = ×400.

	DISCUSSION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Activation of mast cells via crosslinking of IgE on the cell surface is accompanied by the release of numerous mediators that cause erythema, vasodilation, and smooth-muscle constriction in surrounding tissue (1, 5). Although this anaphylactic reaction (characterized by early bronchoconstriction) does occur in allergic asthmatic individuals (7, 11), recent evidence suggests that subsequent airway inflammation and bronchial hyperresponsiveness can also be generated by products of T lymphocytes and eosinophils (4, 12, 21, 22, 26). The experiments described here were designed to quantify the pathophysiologic changes seen after an allergen challenge in rats sensitized to HDM, and to assess allergen-induced changes in recipient animals injected with cells or serum from sensitized or adjuvant control donors.

Recipients of serum from HDM-sensitized donors experienced immediate bronchoconstriction after antigen challenge, whereas animals injected given serum from adjuvant controls or heated serum from sensitized rats did not respond. The magnitude of the immediate responses observed in serum recipients following antigen challenge was comparable to that seen in sensitized donor rats, but was not followed by the vigorous inflammatory response or AHR that occurred in the latter. Since the donor serum was rich in antigen-specific IgE antibody, the immediate bronchoconstriction in recipients probably occurred via mast-cell degranulation. The lack of significant airway inflammation or AHR suggested, however, that there were either insufficient concentrations of bound IgE to elicit these later responses, or that another mediator or cell was involved that was not transferred in serum. These results confirm previous physiologic observations by Sorkness and associates (23), who showed that passive transfer of monoclonal IgE antibody specific for dinitrophenol (DNP) elicited immediate bronchospasm but not late changes in the lung mechanics of Sprague-Dawley rats after challenge with DNP-BSA. Similarly, AHR was observed in IgE-deficient mice after sensitization to Aspergillus fumigatus (27).

Although the foregoing studies have shown that AHR and pulmonary inflammation can occur in the absence of IgE antibody, a number of studies have shown the converse, and have especially implicated mast cells in the development of asthmatic symptoms. Mast cells produce a number of cytokines, including IL-4, IL-5, IL-6, and tryptase, which are involved in allergic inflammation and AHR (28, 29). In addition, mast-cell activation with anti-IgE antibody enhanced methacholine responsiveness in normal mice but not in mast-cell-deficient animals (30). Taken together, these findings illustrate the multiple mechanisms by which asthmatic symptoms can develop, and provide a starting point for identifying the principal pathways of allergic lung disease.

In contrast to the serum results in our study, lymphoid cells from sensitized rats did not transfer immediate bronchoconstriction responses to recipients challenged with antigen, but conferred the ability to mount a substantial immune-mediated inflammatory response and AHR whose magnitude equalled that in the donor animals. The absence of any immediate responses was probably due to a lack of IgE antibody, since no reaginic activity was present in the serum of these animals. The pronounced inflammation after antigen challenge occurred in the recipients of T lymphocytes from sensitized donors, and may have resulted from the release of inflammatory mediators such as IL-5 after antigen challenge (13, 14). This increased inflammation was also associated with increased airway reactivity to ACh. AHR to nonspecific stimuli such as cholinergics agents, cold air, and exercise is a hallmark of chronic asthma (9, 31). Although the mechanism for the development of AHR in asthma is unknown, there is a strong association between the degree of eosinophilia and the severity of hyperresponsiveness (32, 33). Recent studies have also shown that eosinophil-derived major basic protein (MBP), and in some cases IL-5, can induce AHR, whereas antibodies against these products negate the AHR response (34, 35).

Previous work by Schuyler and colleagues (19) demonstrated that inflammatory responses of delayed-type hypersensitivity (DTH) could be transferred to guinea pigs by T cells but not by serum from donors sensitized to M. fanii. Since publication of their study, a number of reports have also implicated T lymphocytes in the transfer of pulmonary inflammation, late-phase responses, and AHR. In a murine model of DTH to picryl chloride, Garssen and colleagues (26) observed hyperreactivity and pulmonary inflammation after challenge of mice that were passively sensitized with lymphoid cells from sensitized donor mice. Watanabe and coworkers (21) injected rats with purified T-cell suspensions and later with CD4⁺ T cells (22) from donors sensitized to ovalbumin, and found that some of the animals experienced LARs and AHR to methacholine, with no detectable immediate bronchoconstriction.

The results of the present study confirm these findings and extend them by illustrating differences in immediate responses, pulmonary inflammation, and bronchial reactivity in rats passively sensitized with cells or serum from actively sensitized animals. Lymphoid cells from HDM-sensitized donors transferred AHR and eosinophilic inflammation after antigen challenge, but conferred no immediate bronchoconstriction responses. Serum from HDM-sensitized donors elicited an immediate response to antigen challenge but no AHR and only modest airway inflammation. These observations support the hypothesis that immediate-type hypersensitivity responses are mediated via mechanism involving IgE, whereas the more severe pathophysiology associated with LARs develops as a result of antigen-specific T-cell activation.