To test the hypothesis that CD8+ T cells may suppress the allergen-induced late airway response (LAR) and airway eosinophilia, we examined the effect of administration of Ag-primed CD8+ T cells on allergic airway responses, bronchoalveolar lavage (BAL) leukocytes, and mRNA expression for cytokines (IL-4, IL-5, and IFN-γ) in OVA-sensitized Brown Norway rats. On day 12 postsensitization to OVA, test rats were administered 2 million CD8+ T cells i.p. isolated from either the cervical lymph nodes (LN group; n = 8) or the spleen (Spl group; n = 6) of sensitized donors. On day 14, test rats were challenged with aerosolized OVA. Control rats were administered PBS i.p. on day 12, and challenged with OVA (n = 10) or BSA (n = 6) on day 14. The lung resistance was measured for 8 h after challenge. BAL was performed at 8 h. Cytospin slides of BAL were analyzed for major basic protein by immunostaining and for cytokine mRNA by in situ hybridization. The LAR was significantly less in the LN group (1.8 ± 0.5 U; p < 0.01) and BSA controls (1.4 ± 0.7; p < 0.01), but not in the Spl group (6.7 ± 2.2), compared with that in OVA controls (8.1 ± 1.8). In BAL, the number of major basic protein-positive cells was lower in the LN and Spl groups compared with OVA controls (p < 0.05 and p < 0.01). IL-4- and IL-5-positive cells were decreased in the LN group compared with the OVA controls (p < 0.01). INF-γ-positive cells were increased in the LN and Spl groups compared with the OVA controls (p < 0.01). Serum OVA-specific IgE levels were unaffected by CD8+ T cell transfers. These results indicate that Ag-primed CD8+ T cells have a potent suppressive effect on LAR.

In human atopic asthma, early airway responses (EAR)3 provoked by Ag challenge of sensitized subjects are often followed by late airway responses (LAR). The LAR appears to be relevant to chronic asthma because it is associated with airway hyperresponsiveness and airway inflammation, two defining features of the disease (1). There is substantial evidence that T cells are pivotal in the initiation of the LAR. In human subjects with atopic asthma, an increased number of activated T cells has been shown in the bronchial mucosa and bronchoalveolar lavage (BAL) fluid (2, 3, 4). Furthermore, it has been demonstrated that these activated T cells predominantly express Th2-type cytokine mRNA, which is characteristic of allergic inflammation (5, 6). It is likely that through the secretion of Th2-type cytokines such as IL-4 and IL-5, these cells are responsible for promoting IgE synthesis (7) and the recruitment and activation of eosinophils (8, 9, 10, 11). Although it has yet to be established whether these cytokines originate from CD4+ or CD8+ T cells, in a previous study we demonstrated direct evidence for the involvement of CD4+ T cells in LAR (12). Administration of Ag-primed CD4+ T cells, but not CD8+ T cells, transferred the LAR to unsensitized BN rats (12). Allergen challenge of these passively sensitized recipients was associated with increased expression of Th2 cytokines in BAL cells (13). Recently, successful transfers of allergen-induced airway eosinophilia and airway hyperresponsiveness were reported in rats using Ag-primed CD4+ T cells (14) and in mice using Th2 cell clones (15, 16).

Although the role of the CD4+ T cells in allergic airway processes has been extensively studied, there are relatively few studies that have focussed on the involvement of CD8+ T cells in LAR. In human subjects undergoing allergen challenge, relative increases in CD8+ T cells in BAL fluid have been reported for subjects showing an isolated early response compared with those with both EAR and LAR (17). These observations were postulated to indicate a suppressive function of CD8+ T cells in the LAR. Depletion of CD8+ T cells with an mAb enhanced the LAR after OVA challenge in Sprague Dawley rats, a nonatopic strain that rarely develops LAR (18). These data also suggested a suppressive role of CD8+ T cells in LAR. However, the pathway by which CD8+ T cells may be activated after allergen exposure is uncertain, because exogenous proteins such as OVA are usually present in association with MHC class II, but not MHC class I, molecules, which are usually required for activation of CD8+ T cells. Furthermore the mechanism by which CD8+ T cells may modulate the LAR is unclear, although CD8+ T cells are capable of producing IFN-γ (19), which is known to counterregulate the Th2 response (20).

Observations in the existing literature prompted us to examine more directly than in previous studies the possibility that CD8+ T cells may modulate the LAR. We hypothesized that OVA-primed CD8+ T cells have a suppressive effect on LAR in OVA-sensitized BN rats. To test this hypothesis, we examined the effect of administration of CD8+ T cells derived from OVA-sensitized donors on the LAR of actively sensitized recipients. We also determined the effect of T cell transfers on the cytokine profiles of the cells in the BAL fluid by the technique of in situ hybridization as well as serum OVA-specific Ig E levels.

Highly inbred male Brown Norway (BN) rats between 7 and 9 wk of age were purchased from Harlan Sprague Dawley U.K. (Blackthorn, U.K.) and maintained in a conventional animal facility at McGill University (Montreal, Canada). Rats were actively sensitized to OVA (grade V, Sigma, St. Louis, MO) with a s.c. injection in the dorsal aspect of the neck of 1 mg of OVA precipitated in 4.28 mg of aluminum hydroxide gel (Anachemia Chemicals, Montreal, Canada) in 1 ml of normal saline. Bordetella pertussis vaccine (0.5 ml) containing 6 × 109 heat-killed bacilli/ml (IAF, Laval-Des-Rapides, Montreal, Canada) was injected i.p. as an adjuvant.

All rats were sensitized to OVA, and airway responses to inhaled aerosolized Ag were measured 14 days later. CD8+ T cell transfers were performed 2 days before Ag challenge. Rats were divided into four groups: 1) eight rats were transferred CD8+ T cells harvested from cervical lymph nodes (LN; the drainage nodes for the site of sensitization) of OVA-sensitized rats and challenged with OVA( LN group); 2) six rats were transferred CD8+ T cells derived from the spleen of OVA-sensitized rats and challenged with OVA (Spl group); 3) 10 rats were administered medium only and challenged with OVA (OVA control); and 4) six rats were administered medium and challenged with BSA (BSA control).

Two additional groups of four rats each were sensitized to OVA and were transferred CD8+ T cells isolated from unsensitized or BSA-sensitized donors to test the necessity of donors having been sensitized to OVA for CD8+ T cell transfers to affect the response to OVA challenge.

W3/25 (CL003AP, mouse anti-rat CD4 mAb, IgG1), MRC OX8 (MCA48G, mouse anti-rat CD8 mAb, IgG1), MRC OX-33 (CL0033A, mouse anti-rat CD45 mAb, IgG1), ED9 (MCA620, mouse anti-rat myeloid differentiation Ag, IgG1), NKR-PI (mouse anti-rat CD161, IgG1), and anti-γδ TCR (mouse anti-rat γδ TCR, IgG1) were purchased from Cedarlane Laboratories (Hornby, Canada); anti-rat MACS rat anti-mouse IgG1 microbeads were purchased from Miltenyi Biotec (Bergisch Gladbach, Germany); goat anti-mouse FITC-labeled IgG was purchased from Life Technologies (Gaithersburg, MD). BMK13, a mouse anti-human major basic protein (MBP) Ab, was provided by Dr. R. Moqbel (University of Alberta, Edmonton, Canada).

Fourteen days after sensitization, donor rats were sacrificed, and the cervical LN and spleen were removed. Cells were harvested by mincing the tissues and subsequently passaging them through a stainless steel sieve. Cells were then washed and suspended in PBS/BSA (0.5%). Purification of CD8+ T cells was conducted by negative selection using immunomagnetic cell sorting (MACS). In the case of spleen cells, mononuclear cells were first isolated by Ficoll-Paque and depleted of B cells by passing them through a nylon wool column before magnetic separation. Magnetic separation was performed first by a 15-min incubation with primary Abs (W3/25; anti-CD4, OX-33; anti-B cell, ED9; anti-myeloid cell). After washing, cells were incubated with MACS beads for 30 min. The labeled cells were then passed through the MACS column, and the cells that were not retained in the column were collected. The purity of the CD8+ T cells was analyzed by flow cytometry (FACScan, Becton Dickinson, Mountain View, CA). We also examined the purified CD8+ T cell preparation for its content of NK cells and γ δ cells using the Abs, anti-CD161 and anti-γδ TCR.

Two million CD8+ T cells were administered to recipient rats 12 days after their sensitization. Following purification the CD8+ T cells were resuspended in 1 ml of sterile PBS and injected i.p. Control rats were injected with PBS only. The cells were harvested from three or four donors and were pooled for administration to two rats.

On day 14 after sensitization general anesthesia was induced with an i.p. injection of urethane (1.25 g/kg). Animals were intubated endotracheally with a 6-cm length of polyethylene tube (PE240, Commercial Plastics, Montreal, Canada) and placed in the left lateral decubitus position on a heating pad. Rectal temperature was continuously monitored with an electronic thermometer. Air flow was measured by placing the tip of the endotracheal tube inside a small Plexiglas box (∼250 ml in volume). A pneumotachograph (Fleisch no. 0, Bionetics, Montreal, Canada) coupled to a differential transducer (PX 170–14DV, Omega Engineering, Stamford, CT) was connected to the other end of the box to measure air flow. A water-filled catheter connected to a pressure transducer (Transpac II, Sorenson, Abbott, IL) was advanced into the lower third of the esophagus to measure pleural pressure changes. Pulmonary resistance (RL) was determined by the technique of multiple linear regression from pleural pressure and air flow using a commercial software package (RHT Infodat, Montreal, Canada) (21).

After determination of baseline RL, animals were challenged with aerosolized OVA or BSA (5%, w/v) using a disposable nebulizer (model 1400, Hudson, Temecula, CA) with an air flow of 10 l/min for 5 min. RL was measured every 5 min in the initial 30 min after challenge and subsequently at 15-min intervals for a total period of 8 h. The EAR was defined as the maximal value of RL, expressed as a percentage of the baseline RL in the first 30 min after challenge. The LAR was calculated as the area under the curve of RL against time from 3–8 h after challenge, after correction of RL for the baseline value.

BAL was performed 8 h after Ag challenge using five consecutive instillations of 5 ml of saline at room temperature. The total cell count was determined using a hemacytometer. Cytospin slides were prepared using a Cytospin model II (Shandon, Pittsburgh, PA). Glass slides were coated with poly-l-lysine (Sigma). Cellular differential was assessed on May-Grünwald-Giemsa-stained slides.

BAL cytospin preparations were fixed in acetone-methanol for 5 min and stored at −80°C until analysis. Cells were stained with mouse anti-human MBP mAb (BMK-13), using the alkaline phosphatase anti-alkaline phosphatase method. MBP-positive cells were assessed by microscopy by an investigator blinded to group status. The results were expressed as the absolute number of MBP-positive cells in BAL.

Cytospin slides were fixed in 4% paraformaldehyde for 30 min, then washed in PBS, baked at 37°C overnight, and stored at −80°C until analysis. In situ hybridization was performed as previously described (22, 23). Antisense and sense riboprobes were prepared from cDNAs coding for rat IL-4, IL-5, and IFN-γ mRNA (5). The probes for IL-4, IL-5, and IFN-γ were gifts from Drs. A. Neil Barclay (Oxford, U.K.), T. Blankenstein (Berlin, Germany), and P. H. van der Meide (Rijswijk, The Netherlands), respectively. cDNAs were inserted into a pGEM vector and linearized. In vitro transcription was conducted in the presence of [35S]UTP and the T7 or SP6 RNA polymerases. For detection of cytokine mRNAs, cytospin preparations were permeabilized with Triton X-100 and proteinase K (1 μg/ml) in 0.1 M Tris containing 50 mM EDTA for 20 min at 37°C. To prevent nonspecific binding of 35S-labeled RNA probes, the preparations were incubated with 10 mM N-ethylmaleimide and 10 mM iodoacetamide for 30 min at 37°C, followed by incubation in 0.5% acetic anhydride and 0.1 M triethanolamide for 10 min at 37°C. Prehybridization was performed with 50% formamide and 2× standard saline citrate for 15 min at 40°C. For hybridization, antisense or sense probes (106 cpm/section) were diluted in hybridization buffer. DTT (100 mM) was present in the hybridization mixture to ensure blocking of any nonspecific binding of the 35S-labeled probes. Posthybridization washing was performed in decreasing concentrations of SCC at 45°C. Unhybridized single-strand RNAs were removed by RNase A (20 μg/ml). After dehydration, the slides were immersed in NBT2 emulsion and exposed for 10 days. The autoradiographs were developed in Kodak D-19 (Eastman Kodak, Rochester, NY), fixed, and counterstained with hematoxylin.

The percentage of cytokine mRNA-positive cells was determined using light microscopy by an investigator blinded to group status. The results were subsequently expressed as the absolute number of positive cells per BAL.

OVA-specific IgE was measured by ELISA as previously described (12) on serum samples obtained at the end of the 8-h period of monitoring of pulmonary function after allergen challenge. Briefly, 96-well assay plates (Corning Glass Works, Corning, NY) were coated overnight at 4°C with 200 μl of the mouse anti-rat IgE mAb (Zymed, San Francisco, CA) in carbonate-bicarbonate buffer (2 μg/ml). The plates were washed four times with PBS-Tween-azide (0.05% Tween 20 and 0.01% NaN3 in PBS at pH 7.4) and blocked with 0.5% casein and 0.1% Tween 20 in PBS. Then, plates were successively treated with 100 μl of diluted rat serum samples (1/10) at 37°C for 1 h, 100 μl of biotinylated OVA (0.02 mg/ml) (27) at 37°C for 1 h, and alkaline phosphatase-conjugated streptavidin (1/500 dilution; Zymed) at 20°C for 30 min. After addition of p-nitrophenyl phosphate disodium (Sigma) as substrate, plates were developed at 20°C for 5–15 min and spectrophotometrically read at 405 nm with an ELISA plate reader (400 ATC, SLT Lab Instruments, Pittsburgh, PA).

The data are presented as the mean ± SE. Statistical comparisons were performed using one-way ANOVA followed by Fischer’s least significant difference test. Statistical significance was accepted at the 5% level of confidence.

Table I shows the lymphocyte subsets of mononuclear cell preparations derived from spleen or cervical lymph nodes of sensitized and nonsensitized animals. In sensitized animals, the absolute numbers of CD4+, CD8+, and B cells in lymph node preparations increased significantly compared with those in naive animals. In the spleen only the number of CD8+ cells increased with sensitization. The CD4/8 ratio was decreased in both lymph node and spleen preparations compared with that in nonsensitized animals. The purity of transferred CD8+ T cells was 90 ± 1.1% (mean ± SE). Contamination of the CD8+ cells by CD4+ (W3/25) cells, B cells (OX-33), and myeloid cells (ED9) was <1%. γδ cells represented 5.4% of the cells in lymph node preparations from sensitized animals, whereas their numbers were greater in spleen preparations (13.2%). NK cells in sensitized lymph node preparations numbered 7.7% and 33.6% in spleen preparations.

Table I.

Lymphocyte subset analysis in lymph nodes and spleen of BN ratsa

LN (Sensitized)LN (Naive)Spl (Sensitized)Spl (Naive)
CD4 64.5 ± 6.8* 41.0 ± 10.0 29.8 ± 1.7 27.8 ± 1.4 
 (65.6 ± 1.1%) (79.7 ± 1.9%) (38.2 ± 2.1%) (42.9 ± 3.3%) 
CD8 10.4 ± 2.9*** 4.6 ± 0.4 13.8 ± 0.8*** 8.3 ± 0.4 
 (10.6 ± 1.3%) (8.7 ± 2.1%) (16.1 ± 1.3%) (12.5 ± 1.5%) 
CD4/CD8 6.4 ± 1.0* 10.2 ± 2.4 2.4 ± 0.3** 3.6 ± 0.5 
B cell 23.2 ± 6.5*** 4.7 ± 0.4 36.3 ± 2.1 33.8 ± 1.7 
 (23.7 ± 0.3%) (11.6 ± 2.9%) (45.7 ± 1.4%) (44.6 ± 3.7%) 
LN (Sensitized)LN (Naive)Spl (Sensitized)Spl (Naive)
CD4 64.5 ± 6.8* 41.0 ± 10.0 29.8 ± 1.7 27.8 ± 1.4 
 (65.6 ± 1.1%) (79.7 ± 1.9%) (38.2 ± 2.1%) (42.9 ± 3.3%) 
CD8 10.4 ± 2.9*** 4.6 ± 0.4 13.8 ± 0.8*** 8.3 ± 0.4 
 (10.6 ± 1.3%) (8.7 ± 2.1%) (16.1 ± 1.3%) (12.5 ± 1.5%) 
CD4/CD8 6.4 ± 1.0* 10.2 ± 2.4 2.4 ± 0.3** 3.6 ± 0.5 
B cell 23.2 ± 6.5*** 4.7 ± 0.4 36.3 ± 2.1 33.8 ± 1.7 
 (23.7 ± 0.3%) (11.6 ± 2.9%) (45.7 ± 1.4%) (44.6 ± 3.7%) 
a

Data are expressed as mean ± SE from three different experiments. Lymphocytes were pooled from two or three rats in each experiment. Cell numbers are expressed as 106 cells per animal, and statistical significance was tested by Student’s t test for unpaired data. The cell counts expressed in percentages are shown in parentheses after removal of myeloid cells.

b

, p < 0.05; ∗∗, p < 0.01; and ∗∗∗, p < 0.001 compared with the naive group.

To test the effects of CD8+ T cells on the pulmonary function of allergen-challenged rats, we measured pulmonary resistance for up to 8 h after challenge in four study groups. The cells were harvested from either the cervical lymph nodes or the spleen of sensitized donors, and the recipients underwent airway challenge with OVA. Sensitized animals challenged with OVA or BSA served as controls.

There was no significant difference in mean baseline RL among groups before challenge. Fig. 1 shows the mean RL at each time point after OVA or BSA challenge. The EAR was not altered significantly by administration of CD8+ T cells. In the BSA-challenged control group the EAR was significantly lower than in the OVA controls (p < 0.05; Fig. 2). The LAR, which was measured as the area under the curve of RL against time, as expected was greater in the OVA controls compared with the BSA controls (p < 0.01; Fig. 3). The LAR was significantly less in the recipients of CD8+ T cells from lymph nodes (LN group) compared with the OVA controls (p < 0.01; Fig. 3), and although the LAR was also less in recipients of spleen CD8+ T cells (Spl group), the difference between this group and the OVA controls was not significant.

FIGURE 1.

The change in pulmonary resistance (percentage of baseline value) of each group is shown as a function of time after Ag challenge. The symbols represent the mean data, and the vertical lines are 1 SEM. All animals were sensitized with OVA. The LN group (n = 8) received CD8+ T cells derived from cervical lymph nodes of sensitized donors. The Spl group (n = 6) received CD8+ T cells derived from spleen of sensitized donors. The OVA control group (n = 10) consisted of OVA-challenged controls; PBS was injected, but no cells were transferred. The BSA control group (n = 7) consisted of BSA-challenged controls; PBS was injected, but no cells were transferred. There was no significant difference in mean baseline RL among groups.

FIGURE 1.

The change in pulmonary resistance (percentage of baseline value) of each group is shown as a function of time after Ag challenge. The symbols represent the mean data, and the vertical lines are 1 SEM. All animals were sensitized with OVA. The LN group (n = 8) received CD8+ T cells derived from cervical lymph nodes of sensitized donors. The Spl group (n = 6) received CD8+ T cells derived from spleen of sensitized donors. The OVA control group (n = 10) consisted of OVA-challenged controls; PBS was injected, but no cells were transferred. The BSA control group (n = 7) consisted of BSA-challenged controls; PBS was injected, but no cells were transferred. There was no significant difference in mean baseline RL among groups.

Close modal
FIGURE 2.

The EAR were calculated as the peak resistance in the first 30 min after challenge. The bars represent the mean values, and the vertical lines indicate 1 SEM. There was a small EAR following OVA challenge that was not altered significantly by administration of CD8+ T cells. In the BSA control the EAR was significantly lower than in the OVA control (p < 0.05).

FIGURE 2.

The EAR were calculated as the peak resistance in the first 30 min after challenge. The bars represent the mean values, and the vertical lines indicate 1 SEM. There was a small EAR following OVA challenge that was not altered significantly by administration of CD8+ T cells. In the BSA control the EAR was significantly lower than in the OVA control (p < 0.05).

Close modal
FIGURE 3.

The LAR were calculated as the area under the curve of RL against time from 3–8 h after OVA challenge. The bars represent the mean values, and the vertical lines indicate 1 SEM. The LAR was significantly reduced in the LN group compared with that in the OVA control group (p < 0.01).

FIGURE 3.

The LAR were calculated as the area under the curve of RL against time from 3–8 h after OVA challenge. The bars represent the mean values, and the vertical lines indicate 1 SEM. The LAR was significantly reduced in the LN group compared with that in the OVA control group (p < 0.01).

Close modal

The inhibitory effect of CD8+ T cell transfers was dependent on prior sensitization of donor animals (Table II). The rats that were challenged with OVA after administration of CD8+ T cells harvested from BSA-sensitized donors (n = 4) or naive donors (n = 4) showed LAR of equivalent magnitude to those in control animals that did not receive any T cells (Table II). As the control OVA-challenged animals were not studied concurrently with the CD8+ T cell recipients, we tested the reproducibility of the LAR in a second group of four rats studied on a separate occasion; the LAR in these animals was 8.0 ± 1.7 cm H2O/ml/s · min, which was not significantly different from the values obtained in the first group studied (8.1 ± 1.8).

Table II.

Effect of transfers of CD8+ T cells derived from cervical lymph nodes of naive or BSA-sensitized donor rats on airway responses of OVA-sensitized and challenged recipient BN ratsa

EAR (% baseline RL)LAR (AUC of RL vs time)
OVA control (n = 10) 130.2 ± 6.3 8.1 ± 1.8 
   
Recipients of CD8+ T cells from naive rats (n = 4) 122.8 ± 9.1 6.9 ± 4.0 
   
Recipients of CD8+ T cells from BSA sensitized rats (n = 4) 140.8 ± 36.1 9.7 ± 6.1 
EAR (% baseline RL)LAR (AUC of RL vs time)
OVA control (n = 10) 130.2 ± 6.3 8.1 ± 1.8 
   
Recipients of CD8+ T cells from naive rats (n = 4) 122.8 ± 9.1 6.9 ± 4.0 
   
Recipients of CD8+ T cells from BSA sensitized rats (n = 4) 140.8 ± 36.1 9.7 ± 6.1 
a

Data are expressed as mean ± SE. Two million CD8+ T cells from cervical lymph nodes of naive or BSA-sensitized rats were transferred to OVA-sensitized rats. There was no significant difference in either EAR or LAR among the three study groups.

Among the various study groups, no statistically significant difference was observed in the numbers of macrophages, neutrophils, and lymphocytes in the BAL fluid (data not shown). Eosinophil numbers (MBP-positive cells) were significantly decreased in both LN and Spl groups compared with those in the OVA control group (p < 0.01 and p < 0.05; Fig. 4). Neither the LN nor the Spl group was significantly different from the BSA controls.

FIGURE 4.

The eosinophils in BAL fluid were identified by immunocytochemical staining using BMK13, a mAb against MBP. MBP-positive cells were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in BAL fluid. The bars represent the mean values, and the vertical lines indicate 1 SEM. MBP-positive cells were significantly decreased in both LN (p < 0.01) and Spl (p < 0.05) groups compared with those in the OVA control.

FIGURE 4.

The eosinophils in BAL fluid were identified by immunocytochemical staining using BMK13, a mAb against MBP. MBP-positive cells were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in BAL fluid. The bars represent the mean values, and the vertical lines indicate 1 SEM. MBP-positive cells were significantly decreased in both LN (p < 0.01) and Spl (p < 0.05) groups compared with those in the OVA control.

Close modal

There was a significant increase in the percentage of cells that expressed IFN-γ mRNA among the CD8+ cells harvested from lymph nodes of sensitized animals compared with those derived from lymph nodes of naive animals (p < 0.05) and from spleen of sensitized (p < 0.05) or naive (p < 0.01; Fig. 5) animals. Approximately 3% of cells expressed IFN-γ mRNA in the lymph nodes of sensitized animals, about twice the level of expression in cells from lymph nodes from naive animals. Spleen cells from naive and sensitized animals showed similar levels of expression of IFN-γ mRNA and were also not different from lymph nodes from naive animals. IL-4 levels were similar among the four groups (Fig. 5).

FIGURE 5.

Cytokine expression in lymph node and spleen CD8+ T cells were analyzed by in situ hybridization. Cytospin preparations of purified CD8+ T cells were used for this analysis. Cytokine mRNA-positive cells were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in BAL. The bars represent the mean values from three different experiments, and the vertical lines indicate 1 SEM. There was a significant increase in the percentage of cells that expressed IFN-γ mRNA in the CD8+ T cells derived from lymph nodes of sensitized animals compared with those from lymph nodes of naive animals (p < 0.05), spleen of sensitized animals (p < 0.05), and spleen of naive animals (p < 0.01). IL-4 levels were similar among the four groups.

FIGURE 5.

Cytokine expression in lymph node and spleen CD8+ T cells were analyzed by in situ hybridization. Cytospin preparations of purified CD8+ T cells were used for this analysis. Cytokine mRNA-positive cells were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in BAL. The bars represent the mean values from three different experiments, and the vertical lines indicate 1 SEM. There was a significant increase in the percentage of cells that expressed IFN-γ mRNA in the CD8+ T cells derived from lymph nodes of sensitized animals compared with those from lymph nodes of naive animals (p < 0.05), spleen of sensitized animals (p < 0.05), and spleen of naive animals (p < 0.01). IL-4 levels were similar among the four groups.

Close modal

Expression of the cytokines IL-4, IL-5, and IFN-γ mRNA in BAL leukocytes is shown in Fig. 6. The number of mRNA-positive cells for IL-4 was significantly lower in LN and Spl groups compared with the OVA control group (p < 0.01). The number of mRNA-positive cells for IL-5 was also significantly lower in the LN group, but not in the Spl group, compared with that in the OVA control group (p < 0.01). The number of IFN-γ-positive cells was significantly higher in both LN and Spl groups compared with that in the OVA control (p < 0.01). In the LN group the number of IFN-γ-positive cells was also higher than in the BSA control group (p < 0.05).

FIGURE 6.

Cytokine mRNA-positive cells in BAL fluid were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in total BAL fluid. The bars represent the mean values, and the vertical lines indicate 1 SEM. The number of mRNA-positive cells for IL-4 was significantly lower in LN and Spl groups compared with that in the OVA control (p < 0.01). The number of mRNA-positive cells for IL-5 was significantly lower in LN group, but not in Spl group, compared with that in the OVA control (p < 0.01). The number of IFN-γ-positive cells was significantly higher in the LN and Spl groups compared with that in the OVA control (p < 0.01).

FIGURE 6.

Cytokine mRNA-positive cells in BAL fluid were assessed by microscopy by an investigator blinded to group status. The results were expressed as the total number of positive cells in total BAL fluid. The bars represent the mean values, and the vertical lines indicate 1 SEM. The number of mRNA-positive cells for IL-4 was significantly lower in LN and Spl groups compared with that in the OVA control (p < 0.01). The number of mRNA-positive cells for IL-5 was significantly lower in LN group, but not in Spl group, compared with that in the OVA control (p < 0.01). The number of IFN-γ-positive cells was significantly higher in the LN and Spl groups compared with that in the OVA control (p < 0.01).

Close modal

OVA-specific IgE levels were significantly increased in OVA-sensitized rats compared with those in negative control unsensitized animals. There was no detectable effect of CD8+ T cell transfers, harvested from cervical lymph nodes from either sensitized or unsensitized donors or from the spleen, on OVA-specific IgE levels in any of the test groups (Fig. 7).

FIGURE 7.

Serum OVA-specific IgE levels were determined by ELISA. The results are shown as units of OD (O. D.) and are corrected for the values of the blank wells. The columns indicate the mean OD for each group, and the bars represent 1 SEM. All the study groups were significantly greater than negative controls. A positive control is shown, representing banked serum from sensitized BN rats (n = 7) that were not otherwise part of the current study. There was no significant difference among the control rats (OVA or BSA challenged; n = 12) that did not receive CD8+ T cells and the groups that received CD8+ T cells from lymph nodes (LN; n = 6) and spleen (Spl; n = 4) from OVA-sensitized donors. IgE levels in the rats that were sensitized to OVA and administered CD8 + T cells from the lymph nodes of naive rats are also shown (n = 4) and were not significantly different from those in the control groups or the other recipients of CD8+ T cells.

FIGURE 7.

Serum OVA-specific IgE levels were determined by ELISA. The results are shown as units of OD (O. D.) and are corrected for the values of the blank wells. The columns indicate the mean OD for each group, and the bars represent 1 SEM. All the study groups were significantly greater than negative controls. A positive control is shown, representing banked serum from sensitized BN rats (n = 7) that were not otherwise part of the current study. There was no significant difference among the control rats (OVA or BSA challenged; n = 12) that did not receive CD8+ T cells and the groups that received CD8+ T cells from lymph nodes (LN; n = 6) and spleen (Spl; n = 4) from OVA-sensitized donors. IgE levels in the rats that were sensitized to OVA and administered CD8 + T cells from the lymph nodes of naive rats are also shown (n = 4) and were not significantly different from those in the control groups or the other recipients of CD8+ T cells.

Close modal

The results of the present study demonstrate a potent suppressive effect of Ag-primed CD8+ T cells on the LAR as well as modulation of the cellular cytokine profiles responsible for allergic airway inflammation. The transfer of purified CD8+T cells harvested from draining lymph nodes, but not from spleens, of actively sensitized donors significantly suppressed the LAR in actively sensitized recipients. This suppression was associated with significant decreases in the numbers of IL-4 and IL-5 mRNA-positive cells and increases in the numbers of IFN-γ-positive cells in BAL fluid that was retrieved 8 h after Ag challenge. Serum OVA-specific IgE levels were unaffected by the T cell transfers. Likewise, the EAR was not altered significantly.

The LAR in the rat appears to be a CD4+ T cell-driven response to allergen challenge (12) and is dependent on cysteinyl-leukotrienes (25). In human studies and animal models, the LAR is usually associated with airway eosinophilia (24, 26, 27), suggesting an important place for this cell in the response. In the current study the inhibition of the LAR by CD8+ T cell transfers was also accompanied by a reduction in the degree of allergen-induced airway eosinophilia. However, the suppression of eosinophilia was of comparable extent following transfer of CD8+ T cells from both the lymph nodes and spleen, whereas the suppression of the LAR did not quite attain a significant level in the group transferred CD8+ T cells derived from spleen. This discrepancy suggests a possible dissociation between the degree of eosinophilia and the LAR and is consistent with data from several studies showing that airway hyperresponsiveness, another consequence of allergen exposure, is independent of airway eosinophilia (28, 29). Further uncertainty concerning the contribution of the eosinophil to the LAR is raised by the finding that in rats the eosinophil does not appear to be a significant source of cysteinyl-leukotrienes (30). Therefore, it is possible that the recruitment of eosinophils may occur concurrently with the LAR, but may not be a pivotal effector of airway narrowing in this model of the LAR.

The mechanism of allergen-induced airway eosinophilia is complex and multifactorial, involving the concerted action of cytokines (5), chemokines (31), and lipid mediators (32). IL-5 is generally thought to be one of the critical factors responsible for induction of airway eosinophilia (33). Although airway eosinophilia was suppressed by CD8+ cell transfers, the number of IL-5-positive cells in BAL fluid was only slightly and not significantly suppressed in the group given CD8+ T cells derived from the spleen. This suggests that the mechanism of the CD8+ T cell-suppressive effects may have been independent of IL-5. Perhaps IL-5 has a greater role in the accumulation of eosinophils in BAL fluid at later time points after single allergen challenge; eosinophilia usually peaks in most animal models at 24 h or more after challenge (24, 34). We speculate that factors other than IL-5 are important in the early phase of eosinophil recruitment. C-C chemokines such as eotaxin and lipid mediators such as the leukotrienes and 5-oxo-eicosatetraenoate are plausible candidates (31, 32), but whether the synthesis and secretion of these substances are interfered with by CD8+ T cells has not been shown.

Generally, CD8+ T cells recognize endogenous Ags associated with MHC class I molecules, whereas exogenous proteins such as OVA are presented in association with MHC class II molecules and are recognized by CD4+ T cells. Hence, CD8+ T cells are not expected to be activated by OVA challenge. Recently, however, there is a growing body of evidence that suggests a leak between class I and class II pathways in APC (35, 36). There are several reports showing that OVA-specific, MHC I-restricted CD8+ T cells are inducible in vivo by OVA sensitization in conjunction with certain adjuvants (36, 37). Consistent with these results are the current findings of substantial inhibitory effects of CD8+ T cells on allergen-induced airway responses in the rat and the lack of an effect of CD8+ T cells transferred from unsensitized donors or from donors sensitized to an irrelevant Ag.

Modulation of allergic airway inflammation by CD8+ T cells has been shown in several animal studies by both depletion and transfer experiments. In vivo depletion of CD8+ T cells by mAb enhanced the LAR, OVA-specific IgE, and airway eosinophilia (18), although not airway responsiveness to methacholine (38), in Sprague Dawley rats. Administration of ricin, which depletes a subpopulation of CD8+ T cells, enhanced IgE production and airway eosinophilia, but not airway hyperresponsiveness, in sensitized BN rats (39). Consistent with suppressive effects of the CD8+ T cells it has been shown that the transfer of spleen CD8+ T cells from OVA-sensitized mice to sensitized syngeneic recipients suppressed the IgE production and prevented the increase in responsiveness of excised tracheal muscle to electrical field stimulation ex vivo (40). Our results are somewhat at variance with these latter findings, because we failed to find any effect of the CD8+ T cells on IgE levels despite marked inhibition of the LAR. Direct comparison of the results of the two studies is difficult given the differences in sensitization protocols and the numbers of T cells transferred. Inhibition of IgE levels in the mouse required a minimum of 107 CD8+ T cells (40).

One of the possible mechanisms by which CD8+ T cells modulate LAR and airway eosinophilia is through the production of inhibitory cytokines. It has been well documented that IFN-γ can suppress the proliferation of Th2-type CD4+ T cells and favor the development of Th1 cells (41, 42). Administration of recombinant murine IFN-γ inhibited allergen-induced eosinophil infiltration of the trachea of sensitized mice, whereas the converse was observed after pretreatment with anti-IFN-γ, which promoted eosinophil and CD4+ T cell infiltration (43). Mucosal IFN-γ gene transfer inhibited pulmonary eosinophilia and airway hyper-reactivity in mice induced by both Ag and a Th2 cell clone (44). These results suggest an important role of IFN-γ in the regulation of pulmonary eosinophilia, and CD8+ T cells are known to be capable of producing large amounts of IFN-γ (45). Although most reports of CD8+ T cell involvement in allergic airway responses have focussed on inhibitory or suppressive functions, quasi-complete depletion of CD8+ T cells inhibited the development of airway hyperresponsiveness and eosinophilia after allergen challenge of mice, indicating a complex interaction of subpopulations of CD8+ T cells in these processes (46).

The CD8+ T cells involved in mediating the effects we have observed have not been further characterized. However, γδ cells are potentially important contributors (47, 48). Although their numbers were small in the lymph nodes and spleen of our sensitized rats, their potency is substantial. These cells can produce high levels of IFN-γ in the rat, and when these cells are harvested from animals in which tolerance has been induced, their administration to naive syngeneic animals inhibits IgE synthesis. As few as 103 cells are sufficient for these effects (48).

The CD8 Ag is expressed on NK cells as well as on T lymphocytes. The proportion of NK cells in the transferred CD8+ cells in the lymph node preparations from sensitized animals was small (7.7%), but was greater in the splenic preparations (33.6%). These results are consistent with published reports that indicate that NK cells are rare in lymph nodes and more frequent in spleen (49). In this study, the suppressive effect on LAR and airway eosinophilia was less prominent in the spleen-derived CD8+ cells. Although this information does not directly address the issue of NK cell involvement, it does suggest that NK cells are unlikely to be responsible for the observed suppressive effects of the lymph node-derived CD8+ cells. Indeed, NK cells have been recently reported to promote, rather than inhibit, allergic airways inflammation in mice (50).

The frequency of cytokine-expressing cells in lymphoid tissue is variable in different animal models in which different methods have been applied. Most studies indicate that the frequency of cytokine-expressing cells is very low or undetectable in vitro unless the cells undergo stimulation (51, 52, 53). Our observations in CD8+ T cells harvested and studied without stimulation are not inconsistent with these studies. Although the number of cells expressing IFN-γ was small, the larger number (3%) of IFN-γ-positive cells in the CD8+ T cell populations isolated from cervical lymph nodes from sensitized animals is consistent with successful in vivo priming.

Whether the IFN-γ-expressing cells found in the BAL of recipient animals are the transferred CD8+ T cells or reflect the influence of these cells on the cytokine profiles of other cells needs to be established. We transferred CD8+ T cells 12 days after the sensitization of recipients. At this time point, we presume that the Th2 response has already been determined in the recipients. It has been suggested that highly polarized Th2 cell clones are irreversible in terms of cytokine-secreting profile (54, 55). However, in vivo there is evidence of a wide spectrum of Th cells in various stages of differentiation (56). Therefore, it is possible that the transferred CD8+ T cells have the potential to induce an alteration in T cell differentiation toward a Th1 dominance through the alteration of the cytokine milieu.

In conclusion, Ag-primed CD8+ T cells have a potent suppressive effect on LAR. This is associated with a significant decrease in eosinophil accumulation and down-regulation of expression of Th2-type cytokine mRNA in BAL cells. Although it seems likely that the suppressive effect of the CD8+ T cells is mediated by IFN-γ, additional studies will be required to definitively establish its role in CD8+ T cell inhibition of allergic airway responses.

We thank Elsa Schotman for her technical assistance.

1

This work was supported by Medical Research Council of Canada Grant 10381.

3

Abbreviations used in this paper: EAR, early airway response; LAR, late airway response; BN rat, Brown Norway rat; BAL, bronchoalveolar lavage; LN, lymph nodes; Spl, spleen; MBP, major basic protein; RL, lung resistance.

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