American Journal of Respiratory and Critical Care Medicine

Lymphocyte migration from the blood into the lung has been suggested as being responsible for the increase of lymphocytes, in particular CD4 T cells, in the bronchoalveolar lavage (BAL) and bronchial mucosa in human asthma, but so far there has been no direct proof. We studied lymphocyte immigration and lymphocyte subpopulations in three lung compartments in ovalbumin (OVA)-sensitized and -challenged brown Norway (BN) rats. Increased numbers of CD4 and interleukin 2 (IL-2) receptor–positive T cells were found in the BAL and lung parenchyma in treated animals, but also increased numbers of CD8 T cells, B cells, and natural killer (NK) cells. For direct proof of lymphocyte migration from the blood into the lung, leukocytes were labeled with a fluorescent dye, 5- (and 6-) carboxyfluorescein-diacetate-succinimidyl-ester (CFSE), and injected intravenously immediately prior to OVA aerosol challenge. One day after challenge the number of CFSE+, i.e., newly immigrated lymphocytes, was determined by flow cytometry gated on the lymphocyte cluster. A 15 times (1.5 times) higher number of CFSE+ lymphocytes was found in the BAL (the lung parenchyma) of treated animals in comparison with control rats. In the BAL 51.8% of CFSE+ cells were CD4-positive (parenchyma 72.7%) and 29.4% IL-2 receptor–positive (parenchyma 34.2%). There was no difference whether the leukocytes for labeling and injection were obtained from untreated or from OVA-sensitized donor animals. Our data show that lymphocyte immigration is at least in part responsible for the increase in lymphocyte numbers in the BAL and lung parenchyma in this animal asthma model. Schuster M, Tschernig T, Krug N, Pabst R. Lymphocytes migrate from the blood into the bronchoalveolar lavage and lung parenchyma in the asthma model of the brown Norway rat.

The importance of eosinophils in the pathogenesis of asthma has been known for a long time. Recent research has discovered that lymphocytes play a crucial role as well, particularly in orchestrating the cellular immune response. Increased numbers of lymphocytes, especially interleukin-2 (IL-2) receptor– positive (i.e., activated) CD4 T cells, are present in bronchial biopsies and in the bronchoalveolar lavage (BAL) of asthmatic patients (1, 2), but also in the blood of patients during acute asthma attacks (3). The expression of several cytokines is increased in BAL lymphocytes of asthmatics, including interleukin-4 (IL-4), IL-5, granulocyte macrophage colony– stimulating factor (GM-CSF), and in some studies also IL-2, IL-3 and interferon gamma (IFN-γ) (4-7). Most attention has been given to IL-5 which is a potent chemoattractant and survival factor of eosinophils (8, 9). CD4 T cells are thought to be the main source of IL-5 in the BAL of asthmatics (2, 4). Further evidence for the importance of lymphocytes during the immune response in asthma has been derived from animal models of asthma. Lymphocytes of ovalbumin (OVA)-sensitized brown Norway (BN) rats were injected into unsensitized animals which were challenged with the antigen afterwards. Transferring primed CD4 lymphocytes, but not CD8 lymphocytes, resulted in bronchial hyperresponsiveness and cellular infiltration of the lung (10) and was accompanied by an increased expression of IL-4 and IL-5 in BAL lymphocytes (11). A depletion of CD8 lymphocytes through specific antibodies led to an increased immune response in the lung of sensitized and challenged Sprague-Dawley rats (12).

In recent years adhesion molecules in the lung have attracted a great deal of attention and several investigators have used the BN asthma model for their studies (13-17). The underlying idea is that blocking specific endothelial adhesion molecules would stop the immigration of lymphocytes (and other immune cells) into the lung. This could prove to be an important new therapeutic option in asthma. Using this asthma model some researchers found that blocking adhesion molecules reduces the airway hyperresponsiveness (AHR), but does not affect the cellular immune response (13-15). Other investigators found that both AHR and cellular immune response were reduced after blocking certain adhesion molecules (16, 17). Because eosinophils only divide in the bone marrow, but not in the periphery, it is clear that lung eosinophilia in asthma is caused by immigrating eosinophils. For lymphocytes the situation is more complex. Because lymphocytes also proliferate, recirculate, and die in the lung (18), the increased numbers of lymphocytes after antigen challenge is not only influenced by immigration. Up to now there has been no direct proof that, and to what extent, lymphocytes migrate from the blood into the lung during an asthma attack (19).

In this article we present data on the migration of lymphocytes from the blood into the marginal vascular pool, the lung parenchyma, and the BAL after allergen challenge using the BN asthma model. These compartments have been chosen for certain reasons. The bronchoalveolar space is the only lung compartment that is accessible without major risks for the patients. Therefore the BAL is often used for diagnostic purposes in lung disease.

To what extent the BAL reflects the immune response in the lung parenchyma, the second compartment studied, is not well understood. The marginal vascular pool of the blood vessels in the lung was examined as a third lung compartment. It has been previously characterized as a distinct lymphocyte compartment (20) and it consists of leukocytes adhering to the blood vessel endothelium in the lung. This compartment is especially interesting for migration studies, because the multistep immigration process of leukocytes is thought to begin with the adherence of circulating leukocytes to the vascular endothelium. So far there have been no studies of the lung marginal leukocyte pool during an asthmatic response. To obtain a better understanding of the migration process, we studied various lymphocyte subpopulations (T cells, B cells, natural killer [NK] cells, CD4 T cells, CD8 T cells, IL-2 receptor–positive T cells) in the three lung compartments and blood of the animals. For direct proof of lymphocyte migration from the blood into the lung, peripheral lymphocytes were labeled with a fluorescent dye and the appearance of the intravenously injected labeled lymphocytes was determined in the three compartments after allergen challenge.

Sensitization and Allergen Challenge

Male BN rats (mean weight 228 g ± 4.8 g) were kept under specific pathogen-free conditions and had ad libitum access to food and water. Eighteen animals were used for studying lymphocyte subsets in different lung compartments and 32 animals were used for examining lymphocyte migration.

The animals were divided into three groups. The OVA-OVA group was sensitized with 1 mg of OVA (Sigma, Deisenhofen, Germany) and 200 mg of Al(OH)3 (Sigma) in 1 ml 0.9% (sterile, pyrogen-free) NaCl (Braun, Melsungen, Germany) applied subcutaneously. As second adjuvant, concentrated preparations of 5 × 109 heat-killed Bordetella pertussis bacilli (kindly donated by the manufacturers, Chiron Behring, Marburg, Germany) in 0.4 ml 0.9% NaCl were given intraperitoneally at the same time. All steps of sensitization were repeated 6 to 7 d after the first sensitization. After another 6 to 7 d the animals were placed 15 min in a Plexiglas tube which was tight enough to restrict movement. A flexible latex mask was placed over the nose and connected to a nebulizer (Pariboy; Ritzau, Starnberg, Germany) which delivered an aerosol of 5% OVA/saline solution at 0.25 L/min. To study the effect of OVA inhalation without prior sensitization the second group of animals (NaCl-OVA group) received equal amounts of 0.9% sterile, pyrogen-free NaCl instead of OVA, Al(OH)3 and Bordetella bacilli during the two sensitizations. The challenge was carried out as described previously with OVA. To exclude any effect of the adjuvants, three animals of this group received the adjuvants Al(OH)3 and Bordetella, but no OVA during the sensitization. Because no differences were seen in the leukocyte subpopulations in the BAL in the animals with or without adjuvant, the results of these animals were pooled. The third group of animals (untreated) was neither sensitized nor challenged.

Dissection of the Animals and Isolation of Blood Leukocytes

For dissection the animals were anesthetized with ether (Baker, Deventer, Netherlands) 22 ± 2 h after challenge. The abdominal wall was opened and the animals were killed by aortic exsanguination. The blood was collected in heparinized tubes and the erythrocytes were removed by incubation with lysing reagent (8.3 g NH4Cl, Serva, Heidelberg, Germany; + 0.1 g ethylenediaminetetraacetic acid [EDTA], Serva; + 1.0 g KHCO3, Merck, Darmstadt, Germany; in 1 L distilled water) for 10 min at room temperature. After centrifugation (400 × g, 10 min) the cell pellet was resuspended in 1 ml phosphate-buffered saline (PBS) (Seromed, Berlin, Germany; containing 1% bovine serum albumin [BSA], Merck; and 0.1% sodium azide [NaN3], Sigma).

BAL

A cannula was inserted in the trachea in situ and the lungs were lavaged with portions of 5 ml cold (4° C) 0.9% NaCl. The fluid was retrieved by gentle aspiration and this procedure was repeated 10 times. The recovery of fluid was over 90% in all animals. The BAL was pooled, centrifuged (400 × g, 10 min) and the cell pellet was resuspended in 1 ml PBS (containing 1% BSA and 0. 1% sodium azide).

Lung Perfusion

The cells from the marginal pool were extracted by extensive perfusion of the main pulmonary artery as described previously (20, 21). The heart and lungs were dissected from the chest and rinsed with 10 ml 0.9% NaCl to remove adhering blood. The outflow tract of the right ventricle was cannulated and the left ventricle was cut open to allow the effluent to be collected. The lung vascular bed was perfused 12 times with 10 ml RPMI (with 35 g/L dextran; Seromed). The first two fractions were discarded, representing cells from the blood pool (21), and the following 10 fractions were pooled for analysis. The cells were centrifuged (400 × g, 10 min) and the cell pellet was resuspended in 1 ml PBS (containing 1% BSA and 0.1% sodium azide).

Lung Cell Extraction

For lung cell extraction a mechanical disaggregation method was used. Compared with enzymatic digestion this yields higher cell numbers (21). The trachea, main bronchi, and hilar lymph nodes were removed from the rest of the lung tissue. The left lungs were used for analysis of the lung cells. The complete lung tissue was disaggregated by passing through a metal sieve with two rounded tweeezers and rinsed with 40 ml PBS. Cells were separated from debris by passing through a 75-μm nylon mesh and centrifuged (400 × g, 10 min). The cell pellet was resuspended in 1 ml PBS (containing 1% BSA and 0.1% sodium azide).

Staining of Eosinophils, Lymphocytes, Granulocytes, and Monocytes/Macrophages

Leukocyte numbers were determined for all three compartments and the blood using standard staining with Tuerk's solution (Merck) and a Neubauer counting chamber. The number of erythrocytes in the BAL was also determined with a Neubauer counting chamber. The differential cell count was assessed on slides prepared by centrifuging 1 × 105 cells in anticoagulant (8.8 g NaCl, Merck; + 0.99 g EDTA + 50 g BSA; in 1 L distilled water) for 8 min at 800 × g on a cytospin centrifuge (Shandon, Pittsburgh, PA). After May-Grünwald/Giemsa staining (Riedel de Haen, Seelze, Germany) eosinophils and granulocytes were identified under the light microscope at ×1,000 magnification using the oil immersion technique. At least 400 cells were differentiated on each slide. Because lymphocytes and monocytes are difficult to differentiate in the blood, marginal pool, and lung parenchyma in rats solely based on morphologic criteria, immunohistochemistry was performed on cytospin slides. Three different antibodies were used for lymphocyte staining: R73 for T cells, MARD 3 for B cells, and 3.2.3. for NK cells (22). All three antibodies were used together as a mixture, after preliminary studies had shown that there was no difference between using a mixture of the three antibodies and preparing three different cytospin slides and adding the numbers. The slides were fixed in acetone (Baker) for 10 min and washed with TRIS-buffered saline (TBS)–Tween (Serva). Of the antibodies (R73 1:1,000; MARD-3 1:500; 3.2.3. 1:10,000; monoclonal mouse anti-rat, Serotec, Oxford, UK; in PBS with 1% BSA and 0.1% NaN3), 100 μl were added to the slides and incubated for 30 min. This (and all following) incubation was performed in a humid chamber. The slides were washed with TBS–Tween and the bridging antibody (100 μl DAKO Z 0259, 1:50; rabbit anti-mouse, Dako, Hamburg, Germany; in PBS) was added and again incubated. After another rinse the alkaline phosphatase–antialkaline phosphatase (APAAP) complex (100 μl DAKO D 065; 1:50; mouse, Dako; in TBS–Tween) was added and incubated. The second and third steps were repeated. Fast blue was used as chromogen. The cytospins were washed with TBS–Tween and counterstained with Haemalaun (Merck) for 2 min. The lymphocytes were counted using phase contrast light microscopy at a ×250 magnification. At least 400 cells were counted on each slide. The percentage of monocytes was determined by subtracting the percentage of eosinophils, granulocytes, and lymphocytes from the total cell number. For the lung parenchyma macrophages, fibroblasts, etc. were grouped together as “other cells.”

Immunostaining of Lymphocyte Subsets and Fluorescent-activated Cell Sorter (FACS) Analysis

Cells of the cell solutions were transferred to a microtiter plate (Greiner, Solingen, Germany; with approximately 1 × 106 cells in each well) and washed with 100 μl PBS (containing 1% BSA and 0.1% NaN3). Previous to a second rinse 50 μl human serum (1:10) was added and incubated for 5 min. The following antibodies (monoclonal mouse anti-rat; Serotec Inc., Raleigh, NC) were used: R73 for T cells, Ox12 for B cells, 3.2.3. for NK cells, W3/25high for CD4-positive lymphocytes, and W3/25low for CD4-positive monocytes, Ox8 for CD8-positive cells, Ox39 for IL-2 receptor–positive cells, and RP1 for granulocytes (22). After the second washing 50 μl primary antibody (Serotec Inc.; in PBS with 1% BSA and 0.1% NaN3) was added to the wells and incubated at 4° C for 30 min. Isotype-matched antibodies served as controls. All antibodies were detected by phycoerythrin (PE)-conjugated goat anti-mouse antibody (Becton Dickinson, Erembodegem Aalst, Belgium; 50 μl, 1:50 in PBS). IL-2 receptor–positive T cells were determined by a double staining technique. Ox39 was used as primary antibody detected by PE-conjugated secondary antibody. As second primary antibody fluorescein isothiocyanate (FITC)-conjugated R73 antibody was used. After the incubation with antibodies the cells were washed three times, resuspended in 200 μl PBS (containing 1% BSA and 0.1% NaN3) and analyzed on a FACscan flow cytometer (Becton Dickinson) focusing on the lymphocyte cluster. Viability was determined by adding 25 μl propidium iodide (1 μl/ml; Sigma) to 200 μl unstained cell solution. At least 10,000 cells were counted and the percentage was calculated for each lymphocyte subset.

Lymphocyte Labeling with CFSE

The fluorescein derivative 5- (and 6-) carboxyfluorescein-diacetate-succinimidyl-ester (CFSE; Molecular Probes, Eugene, OR) is nonfluorescent until it enters the cell by crossing the cell membrane. It is activated by unspecific esterases and forms non–membrane-passing conjugates (23). In most experiments leukocytes used for staining were obtained from the blood of untreated BN rats. Additionally, in six animals of the OVA-OVA group leukocytes were obtained from donor animals which were sensitized with OVA and adjuvants. The aim of this experiment was to compare the migration of “sensitized” versus “nonsensitized” lymphocytes in the asthmatic lung. After lysing erythrocytes the leukocyte number was determined in a counting chamber. From these cells 30 to 40 × 106 were suspended in 500 μl RPMI and 4 μl of a solution prepared from 50 μl of a 5 mmol CFSE stock solution and 450 μl dimethylsulfoxide (DMSO; Merck, Darmstadt, Germany) was added. The cell solution was incubated for 30 min at 37° C and washed twice with 10 ml PBS (+ 0.1% BSA) afterwards. The cells were resuspended in 500 μl 0.9% NaCl and again counted in a counting chamber. For injection 25 × 106 cells were used. The staining was assessed by FACS analysis of excess cells prior to injection.

Cell Injection and Dissection of the Animals

The suspension of labeled cells was injected into the animals before the antigen challenge. After the induction of anesthesia with ether, the tail vein of the animals was punctured with a 24-gauge butterfly needle and the cell suspension injected into the vein. After this the animals were placed in the tube for challenge as described previously. The animals were killed 22 ± 2 h after the challenge and dissected as described previously for subset analysis. The nonsensitized, nonchallenged (untreated) animals were injected 22 ± 2 h before dissection. The leukocytes from the three lung compartments and the blood were analyzed directly with the flow cytometer gated on the lymphocyte cluster, counting 20,000 lymphocytes for each compartment. The percentage of CFSE+ lymphocytes as detected in fluorescence 1 was calculated.

Only the animals from the OVA-OVA group had a sufficient number of immigrated, CFSE+ lymphocytes in the BAL for further subtyping. Six animals of this group were used. Before flow cytometric analyses BAL and lung parenchyma were counterstained with PE-conjugated antibodies against CD4 and IL-2 receptor as described previously for the subset analyses. With this double staining technique it was possible to analyze the percentage of CD4 and IL-2 receptor– positive lymphocytes of all immigrated, CFSE+ lymphocytes. In these experiments 50,000 lymphocytes were counted for each compartment in the lymphocyte gate.

Because postmortem BAL in rats leads to small leakages in the vascular bed with blood spilling over into the BAL, the number of CFSE+ blood lymphocytes contaminating the BAL was estimated based on the numbers of erythrocytes in the BAL and the lymphocyte/erythrocyte ratio in the blood.

The right lung was instilled with 5 ml optimal cutting temperature (OCT) embedding medium (Sakura, Tokyo, Japan), frozen in liquid nitrogen and stored at −70° C. For preparation of cryostat sections, 5-μm-thick sections were cut and placed on glass slides. Immunostaining with DE1 (Boehringer, Mannheim, Germany; monoclonal mouse, 1:1,000, serving as primary antibody detecting CFSE+ cells) was performed on these cryostat sections and on BAL cytospins as described previously for cytospin preparations.

Data Analysis

All data are given as arithmetic mean ± 1 SEM. Error bars in the figures also represent 1 SEM. Means of the three groups were compared using the Mann-Whitney U test, preceded by the Kruskal-Wallis test using SPSS for Windows 6.0.1. (SPSS Inc., Chicago, IL). A p value < 0.05 was considered to be significant.

Effects of OVA Challenge on Leukocyte Populations and Lymphocyte Subpopulations

Compared with untreated animals the allergen challenge in sensitized animals led to a drastic increase in all leukocyte populations in the lung parenchyma and BAL. This increase was not observed in the NaCl-OVA group (Figures 1C and 1D). In the blood neutrophils, and in the marginal pool eosinophils and neutrophils were increased significantly compared with the controls (Figures 1A and 1B). The subset composition of the lymphocytes was also studied and the data for T cells, B cells, and NK cells are shown in Figure 2. No significant differences between the three groups were found in the blood or the marginal pool (Figures 2A and 2B). In the BAL, however, the number of T cells was 2.9 (3.8) times higher in the OVA-OVA group compared with the NaCl-OVA group (in parentheses: untreated group). The number of B cells in the BAL was 2.9 (4.3) times higher. In the parenchyma the number of T cells was 2.7 (3.0) times, and the B cells 2.4 (1.9) times higher. In the parenchyma the number of NK cells was also increased significantly (Figures 2C and 2D). Because of the importance of T cells in the pathogenesis of asthma the CD4 and CD8 T cell populations were determined. Compared with both control groups no significant differences were found in the blood or the marginal pool of the OVA-OVA group (Figures 3A and 3B). The CD4 T cells were increased 3.8- (4.2-) fold in the BAL and 2.9- (2.8-) fold in the parenchyma. CD8 T cells were increased significantly only in the lung parenchyma, but not in the BAL, even though there was a tendency toward higher numbers in the OVA-OVA group (Figures 3C and 3D). Finally, we studied the number of IL-2 receptor–positive T cells in the three lung compartments and the blood. Again there were no significant differences between the three groups in the blood or the marginal pool. In the BAL, however, the number of activated lymphocytes was 4.4 (7.3) times higher than in the controls and in the parenchyma 5.4 (8.4) times higher (Figure 4).

CFSE+ Lymphocytes in the Three Lung Compartments and the Blood

The immigrated CFSE+ lymphocytes were readily identifiable with FACS analysis of the different compartments (Figure 5) but also using immunohistochemistry in the lung parenchyma (Figure 6).

Very few CFSE+ lymphocytes were found in the BAL of untreated control animals (absolute number in whole lung lavage: 41 ± 15) and the NaCl-OVA group (135 ± 65), but 2,063 (± 581) CFSE+ lymphocytes were found in the BAL of the OVA-OVA group. A significantly higher number of immigrated CFSE+ lymphocytes was also present in the asthmatic lung parenchyma as compared with that of untreated animals (absolute number in the left lung: 6,161 ± 1,064 versus 1,307 ± 213). No differences were found between control rats and asthmatic animals in the blood or in the marginal vascular lymphocyte pool (Figure 5). Further subtyping of immigrated CFSE+ lymphocytes in the OVA-OVA animals revealed that in the BAL 51.8 ± 1.5% were CD4-positive cells and 29.4 ± 3.8% were IL-2 receptor–positive. In the parenchyma 72.7 ± 1.7% of immigrated CFSE+ lymphocytes were CD4-positive and 34.2 ± 3.3% were IL-2 receptor–positive.

In an additional experiment the migration of “sensitized” lymphocytes was studied in animals from the OVA-OVA group and the results compared with the migration of “unsensitized” lymphocytes as described previously. “Sensitized” lymphocytes were obtained from the blood of animals that had undergone OVA sensitization. The absolute number of immigrated CFSE+ lymphocytes was 2,326 ± 514 in the BAL and 6,108 ± 1,049 for the left lung parenchyma. Neither in the BAL nor in the lung parenchyma was there a significant difference between the migration of “sensitized” and “nonsensitized” lymphocytes in animals of the OVA-OVA group.

In this study we investigated changes in leukocytes and lymphocyte subsets in three different lung compartments after OVA inhalation in the asthma model of sensitized BN rats. In accordance with previous investigators we found increased numbers of eosinophils and lymphocytes 22 h after allergen challenge in the lung (14, 24). Furthermore, increased numbers of CD4 and activated, IL-2 receptor–positive T cells were detectable in BAL and lung parenchyma, demonstrating that activated T-helper cells are involved in the allergic response in the lung in this animal asthma model, similar to the findings in human asthma (1, 2). Additionally, we found increased numbers of CD8 T cells, B cells, and NK cells in lung parenchyma and B cells in the BAL. Whereas an elevated number of CD8 T cells has also been demonstrated in the bronchial mucosa of allergen-challenged BN rats (25), the role of B cells and NK cells has not been investigated so far.

The overall picture of lymphocyte subpopulations in the allergic lung shows some similarities between the BAL and the lung parenchyma, especially in the predominance of CD4 T cells, but the lymphocytic responses in the BAL and the lung parenchyma do not match completely. For example, the T cell/B-cell ratio is 5.1 times higher in the BAL compared with the lung parenchyma, and the percentage of IL-2 receptor– positive T cells in the BAL is 2.5 times higher than in the lung parenchyma. Furthermore, CD8 T cells and NK cells are only elevated in the lung parenchyma. The subset composition of the marginal pool, which has been described previously as a distinct pool of lung lymphocytes in pigs and rats (20, 21) but which has not been investigated in asthma, is very different from that of the BAL or the lung parenchyma. Regarding lymphocytes and lymphocyte subsets we could not find any changes after allergen challenge in this compartment. These data provide clear evidence that the immune response in the lung is compartmentalized and that observations in one compartment do not necessarily reflect the situation in another compartment. In clinical practice the most widespread tool to examine the cellular immune response in the lung is the BAL. It seems to be important to keep in mind that the lymphocytes in the BAL do not provide direct information about the lymphocytes in the lung parenchyma. The same is valid for peripheral blood. As can be seen in this animal asthma model, the blood does not reflect the immune response in the lung (19).

The increased numbers of lymphocytes in the lung are thought to be the result of lymphocyte recruitment from the blood into the lung. However, up to now there were only indirect signs for this migration. One of these was that the number of blood CD4 T cells decreases and the number of BAL CD4 T cells increases after antigen challenge of asthmatics (26). Because there has been no direct proof of this migration so far, we investigated the appearance of intravenously injected, fluorescent (CFSE)-labeled peripheral lymphocytes in the lung after allergen challenge. Significant increased numbers of CFSE+ lymphocytes were found in both the BAL and the parenchyma of the asthmatic lungs 1 d after injection and challenge, whereas in the BAL of the control animals almost no newly immigrated lymphocytes were detectable. In the BAL of the asthmatic lungs over half of these CFSE+ lymphocytes were CD4-positive and an even higher percentage in the lung parenchyma. Approximately a third were IL-2 receptor–positive in both BAL and parenchyma. These data provide strong evidence that the newly immigrated cells are taking part in the allergic immune response which is dominated by activated CD4 T cells. We further investigated whether “allergen-sensitized” lymphocytes (i.e., lymphocytes obtained from the blood of OVA-sensitized animals) would facilitate the migration of these cells into the lung after allergen inhalation. However, we were unable to demonstrate differences in the number of CFSE+ cells in the lung when comparing “sensitized” with “nonsensitized” lymphocytes. This raises the question whether the recruitment of lymphocytes during an allergic response is allergen-specific, as has been suggested (27), or whether it is nonspecific triggered by inflammatory cytokines.

In a previous study in rats intravenously injected, FITC- labeled thoracic duct lymphocytes could not be found in the BAL of normal lungs within 4 d after injection (28). Even though our data demonstrate the immigration of fluorescent labeled lymphocytes into the asthmatic lung, the fraction of injected cells found 22 h after allergen challenge in the lung was surprisingly low. One reason for this is the relatively small number of lymphocytes in the lung, even the inflamed lung, compared with the huge number of lymphocytes in the lymphatic organs, such as lymph nodes and spleen.

Some methodological comments are also necessary regarding our in vivo model of lymphocyte migration into the asthmatic lung. First, we investigated the appearance of labeled cells in the lung only at 22 h after the injection, which might not be the optimal timepoint. However, since the increase of eosinophils and lymphocytes in BAL and parenchyma is prominent at 22 h after the challenge, it seems to be a reasonable timepoint. Second, although we injected a total of 25 × 106 CFSE+ leukocytes, only about 60% of these cells were in fact lymphocytes. The other CFSE-stained leukocytes were excluded at the time of analysis by gating on the lymphocyte cluster. We did not purify the lymphocytes prior to injection in order to reduce in vitro manipulation. Furthermore, we did not try to maximize the number of migrating cells, as would have been possible by using large numbers of lymphocytes from lymph nodes. To imitate the physiological situation as closely as possible, we used a reasonably low number of blood leukocytes for staining—approximately a third of the lymphocyte pool in the blood. As the low index of stained lymphocytes in the blood shows, most of the injected lymphocytes had left the blood 1 d after injection. Instead they were found in large numbers in lymph nodes (data not shown). One further possible problem using labeled cells in studying migration could be the influence of labeling on the migration pattern or a loss of staining in an in vivo model. However, previous studies have shown that staining lymphocytes with immunofluorescent dyes does not alter the viability or migration pattern in vivo (29). Furthermore, it is known that CFSE does not influence the capacity of stained lymphocytes to divide in vivo and in vitro (23) and that CFSE-stained lymphocytes can be obtained from the spleen up to 8 wk after injection and are still readily identifiable by FACS analysis (30). Finally, extensive postmortem BAL in rats almost inevitably leads to small leakages in the vascular bed with blood spilling over into the BAL. Therefore CFSE+ lymphocytes found in the BAL could be the result of this contamination rather than reflecting cell migration.

We therefore calculated the number of CFSE+ lymphocytes from the blood in the BAL based on the number of erythrocytes in the BAL, as previously described. The calculated range was between 4 and 200 CFSE+ lymphocytes. Because the number of CFSE+ cells in the BAL of the untreated group was in that range, the majority of these lymphocytes were a result of this contamination. In the OVA-OVA group the number of contaminating lymphocytes was in the same range, demonstrating that contamination is not of relevance for the increase of labeled lymphocytes after allergen challenge.

Although immigration of lymphocytes from the peripheral circulation to the lung is an important factor to increase the cell numbers in the lung after allergen challenge, other factors, e.g., local proliferation, changed apoptosis, and altered recirculation of lymphocytes, might influence the number of lymphocytes in the BAL and lung parenchyma in asthma. Proliferation of lymphocytes has been studied in the blood (27) and the BAL (31, 32) of asthmatics, but the results are contradictory in part because of the necessary in vitro stimulation which varies from researcher to researcher. The best available data for in vivo proliferation of BAL lymphocytes come from a mouse model of pulmonary immune response to intratracheal sheep erythrocytes. Only very low numbers of proliferating BAL lymphocytes were found (18). The regulation of apoptosis could be another factor influencing lymphocyte numbers. In healthy humans over 80% of the BAL T cells carry the Fas antigen (CD95) (33, 34), which is involved in cellular mechanisms of apoptosis. In asthmatics, but not in healthy volunteers, the physiological ligand of CD95, Fas ligand (CD95L), is increased on T cells after segmental allergen challenge (34), which suggests increased apoptosis of lymphocytes in the BAL in asthmatics. In a mouse model of lung inflammation increased apoptosis of BAL lymphocytes was also found after intratracheal instillation of sheep erythrocytes (35). Very few data exist about the third possible factor influencing BAL cell numbers: recirculation. But it has been shown in pigs (36) and in rats (37) that lymphocytes leave the BAL via the afferent lymphatics and migrate to the mediastinal lymph nodes. Whether any of these three processes (proliferation, apoptosis, recirculation) is extensive enough to influence the total number of lymphocytes in the BAL is not known. Further studies are necessary to determine the importance of these factors for lymphocyte homeostasis in the lung in asthma. Furthermore, the allergen specificity of the immigrating lymphocytes is one of the important yet unknown facts which need further investigation.

The technical assistance of A. Weiß and K. Westermann and the correction of the English by S. Fryk are gratefully acknowledged.

Supported by grants from the Studienstiftung des Deutschen Volkes (M.S.) and the Deutsche Forschungsgemeinschaft (Pa 240/7-2, Kr 1405/2-1).

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Correspondence and requests for reprints should be addressed to Dr. R. Pabst, Department of Functional and Applied Anatomy, Medical School of Hanover, 30623 Hanover, Germany. E-mail:

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