Accessory cell function of airway epithelial cells

Erwin Oei,1 Thomas Kalb,1 Prarthana Beuria,2 Matthieu Allez,2 Atsushi Nakazawa,2 Miyuki Azuma,3 Michael Timony,2 Zanetta Stuart,2 Houchu Chen,2 and Kirk Sperber2

1Division of Pulmonary and Critical Care Medicine and 2Immunobiology Center, Mount Sinai School of Medicine, New York, New York 10029; and 3Department of Molecular Immunology, Graduate School, Tokyo Medical and Dental University, Tokyo, Japan

Submitted 29 May 2003 ; accepted in final form 6 February 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Oei, Erwin, Thomas Kalb, Prarthana Beuria, Matthieu Allez, Atsushi Nakazawa, Miyuki Azuma, Michael Timony, Zanetta Stuart, Houchu Chen, and Kirk Sperber. Accessory cell function of airway epithelial cells.

We previously demonstrated that airway epithelial cells (AECs) have many features of accessory cells, including expression of class II molecules CD80 and CD86 and functional Fc{gamma} receptors. We have extended these studies to show that freshly isolated AECs have mRNA for cathepsins S, V, and H [proteases important in antigen (Ag) presentation], invariant chain, human leukocyte antigen (HLA)-DM-{alpha} and HLA-DM-{beta}, and CLIP, an invariant chain breakdown product. A physiologically relevant Ag, ragweed, was colocalized with HLA-DR in AECs, and its uptake was increased by granulocyte-macrophage colony-stimulating factor and IFN-{gamma} treatments, which had no effect on CD80 and CD86 expression. We demonstrate the presence of other costimulatory molecules, including B7h and B7-H1, on AECs and the increased expression of B7-H1 on AECs after treatment with granulocyte-macrophage colony-stimulating factor and IFN-{gamma}. Finally, we compared T cell proliferation after allostimulation with AECs and dendritic cells (DCs). The precursor frequency of peripheral blood T cells responding to AECs was 0.264% compared with 0.55% for DCs. DCs stimulated CD45RO+, CD45RA+, CCR7+ and CCR7CD4+, and CD8+ T cells, whereas AECs stimulated only CD45RO+, CD45RA, CCR7, CD4+, and CD8+ T cells. There was no difference in cytokine production, type of memory T cells stimulated (effector vs. long-term memory), or apoptosis by T cells cocultured with AECs and DCs. The localization of AECs exposed to the external environment may make them important in the regulation of local immune responses.

dendritic cells; immune responses; ragweed; lymphocytes; bronchoalveolar lavage


AIRBORNE ANTIGENS (Ag) are mostly excluded or eliminated from the respiratory tract by a complex array of mechanisms, including the mucociliary escalator and the local production of antimicrobial peptides (14). Pathogens and aeroallergens that are not eliminated may elicit T and B cell immune responses. CD4+ helper T lymphocyte response to Ag requires the participation of mucosally associated dendritic cells (DCs), which provide potent Ag-presenting cell (APC) activity, including avid Ag uptake, processing, migration to local lymphoid organs, and presentation of antigenic peptide fragments complexed to class II major histocompatability complex (MHC) molecules to Ag receptors on naïve CD4+ T lymphocytes (15). Naïve T lymphocytes that are presented Ag by DCs may, on activation, recirculate and return to mucosal and submucosal sites, including the airway (16).

The predominant phenotype of CD4+ lymphocytes harvested from bronchoalveolar lavage (BAL) is consistent with a primed memory, nonnaïve phenotype, along with a smaller proportion of CD8+ lymphocytes (4). These cells are representative of those that have responded to Ag challenge. The accessory cell population that induces secondary T cell responses appears less stringent in their requirement for DCs, and, in vitro, nonnaïve T cells proliferate in response to alternative cell types bearing class II MHC molecules, including monocytes and macrophages, as well as non-bone marrow-derived cell types, which may constitutively express or be induced to express class II MHC, including some endothelial and epithelial cell types (17). Of the non-DC putative accessory cell populations in the lung, alveolar macrophages have been studied extensively, because they are actively phagocytic, express class II and CD86 and, to a lesser extent, CD80 family members, and can be harvested by BAL. In most settings, however, BAL macrophages suppress T cell activation in vitro and appear to suppress a DC-induced response (36).

We previously demonstrated that airway epithelial cells (AECs) have many features of accessory cells, including expression of class II molecules CD80 and CD86 and functional Fc{gamma} receptors (34). It was demonstrated that AECs could stimulate T lymphocyte activation and proliferation to antigenic, allogeneic, and anti-CD3 stimulation. AECs were also shown to take up soluble Ag, which was upregulated by IFN-{gamma} and granulocyte-macrophage colony-stimulating factor (GM-CSF). Furthermore, the endocytosed Ag could be colocalized with class II molecules and in different Ag-processing compartments, including early and late endosomes, acidic compartments, and lysosomes.

We have extended these earlier studies here to first demonstrate that freshly isolated AECs have mRNA for cathepsins S, V, and H (proteases important in Ag processing), invariant chain (Ii), human leukocyte antigen (HLA)-DM-{alpha}, and HLA-DM-{beta}. An Ii breakdown product, CLIP, was demonstrated in freshly isolated respiratory epithelial cells by intracytoplasmic staining. Physiologically relevant Ag [ragweed (RW)] was colocalized with HLA-DR in AECs, and its uptake was increased with GM-CSF and IFN-{gamma}. We also show that other costimulatory molecules, including B7h and B7-H1, are present on AECs and that the expression of B7-H1 is increased after treatment with GM-CSF and IFN-{gamma}. Finally, we compared T cell proliferation after allostimulation with AECs and DCs. DCs and AECs induce proliferation of CD4+ and CD8+ T cells, although DCs were much more efficient than AECs. DCs stimulated CD45R0+, CD45RA+, CCR7+ and CCR7CD4+, and CD8+ T cells, whereas AECs stimulated only CD45RO+, CD45RA, CCR7, CD4+, and CD8+ T cells. There was no difference in the pattern of cytokine production, type of memory cell stimulated (effector vs. long term), or apoptosis of T cells cocultured with AECs and DCs.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell lines. The A549 and BEAS-2B cell lines were obtained from the American Type Culture Collection (Rockville, MD) and were grown in F-12 medium (Sigma, St. Louis, MO) supplemented with 10% FCS, 1% penicillin-streptomycin, and 1% L-glutamine (Life Technologies, Grand Island, NY), henceforth called complete medium (CM). Primary cultured respiratory epithelial cells were obtained from Clonetics (Gaithersburg, MD) and grown in bronchial epithelial basal medium (Clonetics, Walkersville, MD) supplemented with bovine pituitary extract, hydrocortisone, epinephrine, insulin, epidermal growth factor, transferrin, retinoic acid, gentamicin sulfate, and triiodothyronine according to the manufacturer's specifications (16). All the cells were cultured in plastic Falcon flasks (T-25, BD Bioscieces, San Diego, CA) at a plating density of 3,500 cells/cm2 with no matrix.

Nasal epithelial cells and AECs. Nasal epithelial cells (NECs) and AECs were obtained from nasal turbinates of patients undergoing surgical resection for lung cancer and from lobar airway segments of patients undergoing thoracotomy, respectively. The patients with nasal turbinate resection had no underlying medical problems and were having the operation for cosmetic reasons. The lobar airway segments were obtained from patients undergoing thoracotomy for lung cancer. None of the patients had been previously treated, and all were ex-smokers. The patients were not clinically infected and were not taking steroids at the time of surgery. The specimens were transported on ice in sterile CM and processed immediately. For the stimulation studies, the surgical specimens were trimmed and thoroughly rinsed, and the luminal surfaces were exposed. After incubation in dispase (3 mg/ml; Boehringer-Mannheim, Indianapolis, IN) for 30 min at 37°C, the nasal and airway specimens were gently scraped with a scalpel blade. The cell suspension was washed in Ham's F-12 medium, passed over a 40-µm mesh filter, and examined by light microscopy for viability and cell composition. The viability was >95% as determined by trypan blue dye exclusion, and lymphoid cell or macrophage contamination was <1% [98% anticytokeratin positive and <1% HLe1/Leu M3+ (Becton-Dickinson, Mountain View, CA)] by immunofluorescence. Epithelial cells were identified by virtue of the beating cilia. Nonciliated columnar epithelia retained their characteristic morphological appearance and were easily distinguished by light microscopy from erythrocytes and white blood cells (18, 34). From 15 sets of respiratory epithelial cell cultures, the yield of epithelial cells varied from 105 to 2 x 106 cells. All the epithelial cells were cultured in 96-well round-bottom plates (Linbro, Oxnard, CA).

DC generation. Peripheral blood mononuclear cells (PBMCs) were isolated from buffy coats obtained from healthy blood donors by means of Ficoll-Paque (Amersham Pharmacia Biotech, Piscataway, NJ) density gradient centrifugation (45). Cells were washed three times using sterile PBS and resuspended in RPMI 1640 (Life Technologies) supplemented with 10% male AB human serum (Life Technologies) and 1% penicillin-streptomycin-glutamine. The freshly isolated PBMCs were incubated at 37°C in 5% CO2 in culture flasks and allowed to adhere for 45 min. The nonadherent cells were removed by several washes with sterile PBS. The adherent monocytes were then cultured in RPMI 1640 supplemented with 1% male AB human serum and 1% penicillin-streptomycin-glutamine and 100 U/ml of GM-CSF (BD Biosciences) and 1,000 U/ml of IL-4 (BD Biosciences). The medium was changed on day 3, yielding immature DC on day 7. On day 7, the medium was replaced and supplemented with TNF-{alpha} (BD Biosciences) at a final concentration of 5 ng/ml. Cells were cultured in TNF-{alpha}-supplemented medium for 3 days to yield mature DCs at day 10 (1).

PCR assays. mRNA was isolated from fresh AECs from bronchial sections and from A549 and BEAS-2B cells using TRIzol (GIBCO BRL, Bethesda, MD) according to the manufacturer's protocol. mRNA was reverse transcribed at 37°C for 60 min in 20 µl of buffer containing 50 mM Tris (pH 8.3), 70 mM KCl, 3 mM MgCl2, dATP, dCTP, dTTP, and dGTP at 0.5 mM each, 10 mM DTT, 20 U of RNase inhibitor, 10 µg/ml of oligo(dT), and 200 U of murine leukemia virus reverse transcriptase (GIBCO BRL). The cDNA product (0.75 µl) was used as a template for PCR amplification using AmpliTaq DNA polymerase (Perkin Elmer, Gaithersburg, MD) and the primers for cathepsins Z, L, V, B, F, W, H, C, S, and B (Table 1). The following primers were used: 5'-AAACAATTAGACCTGGCTG-3' (sense) and 5'-AGCAGCCAATGTTCACGC-3' (antisense) for B7h and 5'-AAACAATTAGACCTGGCTG-3' (sense) and 5'-TCTTACCACTCAGGACTTG-3' (antisense) for B7-H1. For all the cathepsins, thermocyclic amplification was as follows: 43 cycles of 30 s at 94°C, 1 min at 54°C, and 1 min at 72°C for 1 min. For B7h amplification, thermocyclic reactions were as follows: 42 cycles of 1 min at 94°C, 1 min at 59°C, and 1 min at 72°C. For B7-H1, the reactions were as follows: 42 cycles of 1 min at 94°C, 1 min at 55°C, and 1 min at 72°C. The products were separated by gel electrophoresis and viewed under UV light (30).


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Table 1. Primers and predicted size of PCR products of cathepsins B, C, H, K, L, S, V, E, and O

 
Fluorescence microscopy. RW Ag (a kind gift of Dr. H. Sampson, Dept. of Pediatrics, Mount Sinai School of Medicine) was labeled by dialysis with fluorescein isothiocyanate (FITC, 5 mg/ml; Sigma) in 0.5 M carbonate buffer (pH 9.5) for 16 h. The mixture was rotated overnight at 4°C, and unbound FITC was removed by passage over a Sephadex G-25 column. The protein content of the column fractions was determined from the ratio of the optical density of FITC to the optical density of FITC-labeled protein at 280 nm. An FITC-to-protein ratio of 5:1 was considered optimal for these studies. The primary cultured respiratory epithelial cells that adhere to glass coverslips were incubated with the FITC-labeled RW for various periods; then they were rinsed in PBS and fixed with 3:1 methanol-acetic acid for 5 min at 4°C. Fixed cells were blocked with PBS containing 5% goat serum and 0.2% Triton X-100 (Sigma) for 15 min before incubation with the anti-HLA-DR antibody for 1 h. The cells were rinsed three times in PBS before incubation with Texas red-labeled conjugated donkey anti-rabbit Ig (Accurate Antibodies, Westbury, NY). The coverslips containing the primary cultured respiratory epithelial cells were then rinsed in PBS three times and mounted with Immun-mount (Shandon, Pittsburgh, PA) before they were viewed with a laser confocal microscope (Fluorvert, Leica, Deerfield, IL) at a step position of 1 µm on the x-y or x-z axis. IFN-{gamma}-stimulated (100 U/ml for 48 h), GM-CSF-stimulated (100 U/ml for 48 h), and unstimulated primary respiratory epithelial cells were pulsed with FITC-labeled Ag at 37°C, harvested at various times, and analyzed with fluorescence microscopy. In some experiments, the cells were pretreated with 100 mM chloroquine for 1 h; other experiments were conducted at 4°C. Two observers routinely examined 10 separate fields. Mean fluorescence intensities of the internalized FITC-labeled Ag in the cells of the entire field examined were determined with the Leica Fluorvert laser confocal microscope using the NIH Image software package (National Institutes of Health, Bethesda, MD) (29).

Immunofluorescence. BEAS-2B, A549, and primary cultured respiratory epithelial cells, along with freshly isolated NECs and AECs, were incubated with 1, 10, 100, and 1,000 U/ml of IFN-{gamma}, IL-2, IL-4, and IL-10 for 0, 6, 12, 24, 48, and 72 h and then stained by indirect methods as previously described with various MAbs (see below) or with isotype-matched controls and then with affinity-purified F(ab')2 FITC-conjugated goat anti-mouse Ig (Tago, Burlingame, CA) and analyzed by flow cytometry, gating on live cells (37). The W6/32 (anti-class I) antibodies were obtained from the American Type Culture Collection; the anti-CD80, anti-CD86, and anti-CLIP antibodies were purchased from BD Biosciences. The B7h (Mifey 4) and B7-H1 antibodies (MIH2) were obtained from the Department of Molecular Immunology, Tokyo Medical and Dental University. Anticathepsin L, V, H, S, K, and B antibodies were kind gifts of Dr. Dieter Bromme (Dept. of Human Genetics, Mount Sinai School of Medicine); the anti-Ii (CD74) and -HLA-DM antibodies were obtained from BD Biosciences. For intracytoplasmic staining with the anticathepsin L, V, H, S, K, and B antibodies, along with anti-CLIP, -Ii, and -HLA-DM antibodies (BD Biosciences), cells were fixed and permeabilized with 70% ethanol for 30 min at 4°C. The cells were then washed three times with PBS, anti-CLIP antibodies or control antibodies were added for 30 min at 4°C followed by affinity-purified FITC-conjugated goat anti-mouse Ig antibody, and the cells were analyzed as previously described (29). Adherent epithelial cells were removed using cell dissociation solution (Invitrogen, Carlsbad, CA), which does affect surface expression of cellular proteins.

Mixed lymphocyte reaction. Freshly isolated AECs were used to stimulate unidirectional mixed lymphocyte reactions (MLRs). AECs were irradiated at 6,000 rad (cesium source). Heparinized venous blood was collected from normal donors, diluted 1:3 with PBS, layered on a Ficoll-Hypaque density gradient, and centrifuged for 30 min at 1,400 rpm. PBMCs were collected from the interface and washed three times with PBS. Cells were resuspended in RPMI 1640, and cell density was adjusted to 5 x 106 cells/ml. T cells were separated by rosetting using neuraminidase-treated sheep red blood cells and Ficoll-Hypaque density gradient centrifugation. Rosetted T cells were treated with 0.75% ammonium chloride on ice for 5–10 min to lyse sheep red blood cells; 105 T cells were cocultured with 104 freshly isolated AECs in 0.2 ml in triplicate round-bottom plates (Linbro) in serum-free culture medium (Aim V, Invitrogen) at 37°C in a 5% CO2 incubator for 5 days. In some experiments, control, anti-B7h, and anti-B7-H1 antibodies (10 µg/ml) were added to the AEC-T cell cocultures. At 18 h before the cells were harvested, 1 µCi of [3H]thymidine (ICN Pharmaceuticals, Aurora, OH) was added to each well. The cells were harvested onto glass fiber filters, and incorporated radiolabel was measured by scintillation counting (30).

Carboxyfluorescein diacetate succinimidyl ester staining. In freshly isolated T cells isolated by methods described above, we used carboxyfluorescein diacetate succinimidyl ester (CFSE) to assess T cell proliferation in AEC-T cell and DC-T cell cocultures (2). The T cell suspension was then washed three times with PBS and finally resuspended in CM. T cells were resuspended in PBS at 5 x 106/ml for staining. CFSE in the form of a 5 mM stock solution in DMSO was added at the final concentration of 1 µM for 10 min at 37°C. The CSFE-labeled cells were cultured with AECs or DCs and then stained with anti-CD3, anti-CD4, anti-CD8, anti-CD45RA, anti-CD45RO, and anti-CCR7R MAbs (BD Biosciences) labeled with phycoerythrin (PE), APC, or peridinin-chlorophyll protein (PerCP) after 7 days of coculture as described above. In other experiments, the CFSE-labeled T cells were stained with PE-labeled anti-CD62L antibodies (BD Biosciences).

Intracellular staining for cytokines. T cells were cocultured with DCs or AECs for 24, 48, and 72 h at 37°C. For the last 16 h of culture, brefeldin A (Sigma; 1 µg/ml) was added. The cells were harvested, washed with PBS + 0.1% BSA and 0.1% NaN3, fixed with 2% (vol/vol) formaldehyde in PBS for 20 min at 4°C, stained with PE-labeled anti-CD3 MAbs, and washed three times in PBS. The cells were then permeabilized with PBS + 0.5% saponin (Sigma) and stained with FITC-labeled anti-IL-2, -IL-4, -IL-10, and -IFN-{gamma} MAbs, washed three times in PBS, and analyzed by flow cytometry as described above.

Annexin V staining. FITC-labeled annexin V, a phospholipid-binding protein of the annexin family (13, 21), was used to measure apoptosis using a commercially available kit (Coulter, Hialeah, FL). T cells isolated as described above were cocultured with AECs or DCs for 16 h at 37°C, harvested, and then stained with PE-labeled anti-CD3 MAbs for 30 min. The cells were then washed in ice-cold PBS, centrifuged at 500 g at 4°C, and then incubated with annexin V-FITC at room temperature for 10 min in the dark. The cells were analyzed by flow cytometry to measure the annexin V-positive population, gating on the live cells (7).


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
In our investigations studying Ag presentation in AECs, we always tried to use freshly isolated AECs and NECs, because they are the most physiologically relevant cells and best mimic AEC function in vivo. There are two problems with using freshly isolated human AECs obtained from surgical specimens: 1) the yield of AEC is low, and 2) there is the possibility of contaminating cells, especially DCs. The small numbers of AECs make some experiments hard to perform, and results demonstrating APC activity might be difficult to interpret. For this reason, we used different AEC types, including the A549 and BEAS-2B cell lines and primary cultured epithelial cells, in our experiments to ensure that we would have adequate numbers of cells for some experiments and to demonstrate that contaminating DCs do not account for any Ag-presenting activity.

AECs have cathepsins B, L, V, C, H, and S. We previously reported that AECs can serve as accessory cells for Ag-, anti-CD3-, and mitogen-induced T cell proliferation (34). We have extended these preliminary studies to further investigate the Ag-processing capabilities of AECs, including the presence of cathepsins. Cathespins, including B, L, H, V, and S, are a family of proteases that are present in many different mammalian cell types (27). Three different cathepsins, B, L, and H, are ubiquitous in the lysosomes of mammalian cells (27). Cathepsins L and B have endopeptidase activity (27), cathepsin C acts mainly as a dipeptidyl peptidase, and cathepsin H is an aminopeptidase (27). Cathepsin V is found in the thymus, where it is important in Ag presentation of self-Ag, whereas cathepsin L is present in the intestinal epithelium and also has a role in Ag processing (42). Cathepsin S has a restricted tissue distribution, with the highest levels detected in the spleen, lung macrophages, and heart (31). Cathepsin S degrades soluble proteins and is active at pH 7.0 (31). It has a role in Ag processing, because inhibitors of cathepsin S prevent Ag processing in B cells by interfering with the proteolysis of Ii (32). To investigate whether AECs have cathepsins involved in Ag processing (cathepsins L, V, and S), we performed PCR using specific riboprobes in freshly isolated AECs. Freshly isolated AECs have amplified base-pair fragments corresponding to the presence of cathepsins L (210 bp), V (320 bp), and S (450 bp), as well as cathepsins H (320 bp), K (480 bp), B (550 bp), and F (350 bp; Fig. 1A). It is possible that injury during the isolation process of the AECs could have activated cathepsins L, V, S, H, K, and F. We performed PCR for cathepsins L, V, S, H, K, and F in the A549 and BEAS-2B cells and found that they were present in both cell lines, indicating that the isolation process did not activate the cathepsin mRNA (data not shown). To confirm that the cathepsins were present at the protein level, we performed intracytoplasmic staining. There were peak channel shifts consistent with the presence of cathepsins L, V, S, H, K, and F in the freshly isolated AECs (Fig. 1B).



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Fig. 1. Cathepsins in airway epithelial cells (AECs). A: PCR analysis for cathepsin mRNA in freshly isolated respiratory epithelial cells. RNA was extracted from AECs obtained from surgical specimens of patients undergoing resection for lung cancer, reverse transcribed, probed with primers for cathepsins L, V, H, C, S, K, B, and F and actin (CTRL), and then run on a 1.2% agarose gel. Negative control consisted of omitting RNA from the DNA amplification step. Results are representative of 1 experiment repeated 5 times. B: intracytoplasmic presence of caspases. Freshly isolated AECs were isolated and stained intracytoplasmically for the presence of CLIP, gating on live cells. Results are shown as percentage of positively staining cells and are representative of 1 experiment using 3 different donors.

 
Presence of Ii, HLA-DM-{alpha}, and HLA-DM-{beta} and Ag processing in AECs. Other nonpolymorphic HLA gene products, including Ii, HLA-DM-{alpha}, and HLA-DM-{beta}, are also essential in Ag processing (8). In the processing compartments, Ii is degraded into several peptides, including CLIP and leupeptin-induced peptide. CLIP binds to the Ag-binding groove of the class II molecules and is removed before endocytically generated antigenic peptides can bind. Leupeptin-induced peptide is a protein fragment that is derived from the COOH end of Ii (9). Removal of CLIP is catalyzed by HLA-DM. Base-pair fragments corresponding to Ii (410 bp), HLA-DM-{alpha} (341 bp), and HLA-DM-{beta} (321 bp) were present in the freshly isolated AECs (Fig. 2A). To confirm that Ii, HLA-DM-{alpha}, and HLA-DM-{beta} were present at the protein level, we performed intracytoplasmic staining. Peak channel shifts corresponding to the intracytoplasmic of presence Ii, HLA-DM-{alpha}, and HLA-DM-{beta} were detected in freshly isolated AECs (Fig. 2B). Because the AECs have cathepsins L, V, and S and possess Ii, HLA-DM-{alpha} and HLA-DM-{beta}, we next investigated whether degradation of Ii can be demonstrated in AECs from the upper and lower airways. We demonstrated the presence of the Ii breakdown product CLIP in freshly isolated NECs and AECs by intracytoplasmic staining with anti-CLIP antibodies. There are peak channel shifts corresponding to the presence of the CLIP peptide from freshly isolated NECs and AECs from the upper and lower airways, respectively (Fig. 2C).



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Fig. 2. Antigen (Ag) processing in respiratory epithelial cells. A: RNA was extracted from AECs obtained from surgical specimens of patients undergoing resection for lung cancer and A549 and BEAS-2B cells, reverse transcribed, probed with primers for invariant chain (Ii), human leukocyte antigen (HLA)-DM-{alpha}, and HLA-DM-{beta}, and then run on a 1.2% agarose gel. Negative control (actin) consisted of omitting RNA from the DNA amplification step. Results are representative of an experiment using 5 different donors and RNA from A549 and BEAS-2B cells. B: intracytoplasmic presence of Ii, HLA-DM-{alpha}, and HLA-DM-{beta}. Freshly isolated AECs were isolated and stained intracytoplasmically for the presence of Ii and HLA-DM. Results are shown as percentage of positively stained cells and are representative of 1 experiment repeated 3 times. C: freshly isolated epithelial cells from nasal turbinates (NECs) of patients undergoing cosmetic rhinoplasty and airways (AECs) from patients undergoing resection for lung cancer were isolated and stained intracytoplasmically for the presence of CLIP, gating on live cells. Results are shown as percentage of positively staining cells and are representative of 1 experiment using 3 different donors.

 
Colocalization of RW with HLA-DR and trafficking in AECs. In our previous studies, we colocalized HLA-DR with three conventional Ag of different molecular weights [ovalbumin (40,000), tetanus toxoid (70,000), and keyhole limpet hemocyanin (1,000,000)] to demonstrate that Ag uptake by AECs was an active process and was not due to passive diffusion (34). We have extended these studies to demonstrate that a physiologically relevant Ag, i.e., RW, is endocytosed by AECs and follows a class II processing pathway. We colocalized Texas red-labeled HLA-DR with FITC-labeled RW (red + green = yellow) in primary cultured respiratory epithelial cells using laser confocal microscopy (Fig. 3). Colocalization occurred optimally 60 min after pulsing. We then investigated RW trafficking in the cultured primary respiratory epithelial cells. The primary respiratory epithelial cells were pulsed with FITC-labeled RW, and extensive speckled punctuate green signals were observed, with a maximal expression at 60 min that persisted for up to 120 min (Fig. 4). RW pulsing at 4°C showed no evidence of RW uptake, suggesting that internalization of RW by the primary respiratory epithelial cells is an active process (data not shown). We also found that treatment with chloroquine (100 mM for 1 h) had no effect on RW uptake, suggesting that RW uptake occurs via non-receptor-mediated endocytosis (data not shown).



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Fig. 3. Colocalization of HLA-DR with endocyosed ragweed (RW) in respiratory epithelial cells. Primary cultured respiratory cells were pulsed with FITC-labeled RW for 0–120 min, fixed, permeabilized, and stained with Texas red-labeled anti-HLA-DR antibodies. Cells were analyzed by confocal microscopy. Yellow staining (red + green) indicates colocalization. Results are representative of 1 experiment repeated 3 times.

 


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Fig. 4. Uptake of RW by primary respiratory epithelial cells. IFN-{gamma}- and granulocyte-macrophage colony-stimulating factor (GM-CSF)-treated (100 U/ml for 48 h) and untreated primary cultured respiratory epithelial cells were pulsed with FITC-labeled RW for 0–120 min and then analyzed by confocal microscopy. Results are representative of 1 experiment repeated using 5 different donors.

 
Ag uptake in APCs can be upregulated by cytokines, including IFN-{gamma} and GM-CSF, that are present at respiratory mucosal surfaces (41). We treated the primary cultured respiratory epithelial cells with IFN-{gamma} and GM-CSF (100 U/ml) to determine whether the magnitude and kinetics of RW uptake would be affected (Fig. 4). In contrast to the lack of staining in untreated primary respiratory epithelial cells, extensive punctuate staining was observed at 30 min after pulsing with FITC-labeled RW. Staining was diminished at 120 min. To better quantify the differences in RW uptake after treatment with IFN-{gamma} and GM-CSF, we determined the mean fluorescence intensity of the endocytosed RW in the respiratory epithelial cells at all the time points tested. There was a significant increase in the mean fluorescence intensity of the entire field examined, corresponding to increased RW uptake starting at 30 min in the IFN-{gamma}- and GM-CSF-treated primary respiratory epithelial cells compared with the untreated cells (P = 0.05); this became maximal at 60 min and was diminished at 120 min (P = 0.05; Table 2).


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Table 2. Fluorescence intensities of endocytosed FITC-labeled RW in IFN-{gamma} and GM-CSF-treated and untreated primary AEC

 
Effect of cytokines on CD80 and CD86 expression in AECs. Expression of costimulatory molecules in professional APCs can be upregulated in response to stimulation with cytokines such as IFN-{gamma} and GM-CSF (26). We first investigated whether CD80 and CD86 expression in AECs was increased by IFN-{gamma}, TNF-{alpha}, and GM-CSF stimulation. There was more expression of CD80 and CD86 in the NECs, AECs, and primary cultured epithelial cells than in the A549 and BEAS-2B cell lines. Some of the cytokines downregulated CD80 and CD86 expression in the BEAS-2B and A549 cells (Fig. 5). We cultured NECs, AECs, the BEAS-2B and A549 cell lines, and the primary cultured respiratory epithelial cells with different concentrations (0, 10, 100 and 1,000 U/ml) of IFN-{gamma}, TNF-{alpha}, and GM-CSF for 16, 24, 48, 72 and 96 h. We could not determine any consistent effect of IFN-{gamma}, TNF-{alpha}, or GM-CSF on CD80 and CD86 expression at any time point or cytokine tested (data not shown). Other cytokines in the airways at the mucosal surfaces, including IL-10, IL-5, or IL-4, may also have an effect on CD80 and CD86 expression in respiratory epithelial cells. To test this, we stimulated primary freshly isolated NECs, AECs, and BEAS-2B, A549, and primary cultured respiratory epithelial cells with IL-10, IL-5, and IL-4 at 0, 10, 100, and 1,000 U/ml for 16, 24, 48, 72, and 96 h. Consistent with the data from IFN-{gamma}, TNF-{alpha}, and GM-CSF stimulation, there was no reproducible effect of any of these cytokines on CD-80 and CD-86 expression (Fig. 5).



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Fig. 5. Expression of CD80 and CD86 on AECs. AECs, NECs, primary cultured epithelial cells, and A549 and BEAS-2B cells were stimulated with IFN-{gamma}, GM-CSF, IL-4, IL-5, and IL-10 for 49 h, stained with anti-CD80 or -CD86 antibody, and then analyzed by flow cytometry, gating on live cells. Three different donors were used for NEC and AEC experiments. NC, negative control.

 
Presence of other costimulatory molecules. Although CD80 and CD86 are the best-characterized costimulatory molecules, other surface molecules, such as B7h and B7-H1, have been noted on the surface of APC (3). B7h and B7-H1 have recently been described as molecules with extracellular domains that share homology with CD80 and CD86. B7h binds to a CD28 homolog, inducible costimulator (ICOS), which is found on a specific subset of T cells. B7-H1 has as its ligand PD-1. We first investigated whether human respiratory cell lines and freshly isolated human AECs, and also the A549 and BEAS-2B cell lines (data not shown), had mRNA for B7h and B7-H1. We found that mRNA for B7h and B7-H1 was present in primary respiratory epithelial cells (Fig. 6A). It has been reported that B7h and B7-H1 expression can be upregulated by TNF-{alpha} stimulation (12, 46). We evaluated mRNA production for B7h and B7-H1 in the primary cultured respiratory cells after TNF-{alpha} stimulation (100 U/ml) for 48 h and found no increased mRNA expression (Fig. 6A).



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Fig. 6. B7h and B7-H1 expression on AECs. A: PCR analysis for B7h and B7-H1 in respiratory epithelial cells. RNA was extracted from TNF-{alpha}-treated (100 U/ml) and untreated respiratory epithelial cells and AECs isolated from patients undergoing lung resection for cancer, reverse transcribed, probed with primers for B7h and B7-H1 and actin probe, and then run on an agarose gel. Negative control consisted of omitting RNA from amplification step. Results are representative of 1 experiment repeated 3 times. B and C: immunofluorescence for B7h and B7-H1. Respiratory epithelial cells (B) and DCs (C) were treated with TNF-{alpha}, IFN-{gamma}, or GM-CSF (100 U/ml for 48 h), stained by direct immunofluorescence for B7h and B7-H1, and then analyzed by flow cytometry, gating on live cells. Results are representative of 1 experiment repeated 3 times. D: blocking of T cell proliferation with anti-B7h and -B7-H1 antibodies (Ab). Freshly isolated AECs treated with anti-B7h and -B7-H1 antibodies were cocultured with allogeneic T cells for 5 days. Cells were pulsed with [3H]thymidine for the last 16 h of culture and counted in a scintillation counter. Results are representative of 1 experiment using 3 different donors.

 
Because IFN-{gamma} and GM-CSF increased the uptake of RW in the primary cultured respiratory epithelial cells (Fig. 4) and there was no effect on CD80 and CD86, we next investigated whether surface expression of B7h and B7-H1 could be increased after TNF-{alpha}, IFN-{gamma}, or GM-CSF stimulation (100 U for 48 h). We also compared expression of B7h and B7-H1 on DCs after stimulation with TNF-{alpha}, IFN-{gamma}, and GM-CSF. Primary respiratory epithelial cells and DCs expressed B7h and B7-H1 (Fig. 6, B and C). After stimulation with IFN-{gamma} and GM-CSF, only B7-H1 expression was increased in the primary respiratory epithelial cells (Fig. 6B). In the DCs, B7h and B7-H1 were expressed on the cell surface (Fig. 6C). B7h expression could be increased after stimulation with TNF-{alpha}, IFN-{gamma}, and GM-CSF. B7-H1 was also present on the surface of DCs and could be upregulated by TNF-{alpha}, IFN-{gamma}, and GM-CSF, but to a lesser extent than on the primary respiratory epithelial cells (Fig. 6C). To determine a functional role for B7-H1 on AECs, we performed a series of blocking experiments using anti-B7h and anti-B7-H1 antibodies to block T cell proliferation induced by freshly isolated AECs in a series of coculture experiments. Anti-B7h and anti-B7-H1 did not inhibit cellular proliferation in the AEC-T cell cocultures as assessed by thymidine incorporation (Fig. 6D) (7).

Comparison of T cells stimulated by AECs and DCs. To further investigate the interaction between AECs and lymphocytes in the airways and to better define the role of AECs in Ag processing in the airways, we used CFSE to assess proliferation of T cells in AEC-T cell and DC-T cell cocultures (2). Experiments were performed using seven different specimens, and the results were examined to ensure that patient variability did not affect the results. However, in some of the experiments, differences were noted in the magnitude of the T cell response. After 5 days of culture, proliferating T cells (CD3+CFSElow) represented 5–25% of all T cells among the AECs and DCs in seven different experiments (Fig. 7A). In contrast, nonactivated T cells (T cells cultured alone) did not divide (Fig. 7B). Analysis of the CFSE staining showed that the majority of these proliferating cells had undergone more than five divisions (Fig. 7B). The precursor frequency of peripheral blood responder T cells in response to stimulation with AECs was 0.264% (Fig. 7B) (25). To more fully characterize T cell responses in the coculture experiments, we assessed whether AECs and DCs preferentially stimulated CD4+ or CD8+ T cells. In these experiments, we cocultured CSFE-labeled T cells with AECs or DCs and characterized the proliferating cells (CSFElow). AECs and DCs stimulated CD4+ and CD8+ T cells (Fig. 8). Four-color immunofluorescence demonstrated that the cells that were proliferating were CD4+ and CD8+ CD45RO+/CD45RA/CCR7, the memory cell phenotype (Fig. 8). The pool of memory T cells is heterogeneous and has been classified as resting long-lived memory or as activated effector memory T cells on the basis of the surface expression of CD62L (low in effector memory T cells and high in long-term memory T cells) (38). We further investigated whether AECs stimulated effector or long-term memory T cells. In these experiments, CSFE-labeled T cells stimulated by AECs and DCs were stained with anti-CD62L antibodies after 7 days of culture. AECs and DCs stimulated CD62L-expressing and non-CD62L-expressing T cells equally (Fig. 8). Using the same donor T cells, we performed similar studies with DCs to compare proliferation in response to allogeneic stimulation. As expected, DCs were more efficient than AECs at stimulating CD4+ and CD8+ T cells (Fig. 7A). The precursor frequency of peripheral blood responder T cells to DCs was 0.55%, i.e., double that of the peripheral blood responder T cells to AECs (Fig. 7B) (25). DCs stimulated CD4+ and CD8+ T cells and naïve CD45RA+ and CCR7+ T cells as well as CD45RO- and CCR7-expressing T cells (Fig. 8) and stimulated long-term effector and central memory T cells (Fig. 8).



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Fig. 7. T cell proliferation induced by respiratory epithelial cells and dendritic cells (DCs). A: analysis of carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled T cells 7 days after coculture. AECs isolated from surgical specimens of patients undergoing resection for lung cancer and DCs derived from peripheral blood monocytes were cocultured with the same donor T cells at a 1:10 AEC-to-T cell and DC-to-T cell ratio for 7 days. Cells were stained with peridinin-chlorophyll protein (PerCP)-labeled anti-CD3. Analysis was performed, gating on CD3+ T cells. Results are shown as percentage of proliferating cells, on the basis of decreased CSFE staining, and are representative of 1 experiment repeated using 7 different donors. B: calculation of precursor frequency of AEC- and DC-stimulated T cells in AEC-T-cell and DC-T cell cocultures. CFSE-labeled T cells were cocultured with AECs and DCs and stained with an anti-CD3 phycoerythrin (PE) on day 7. Analysis was gated on CD3+ cells. Percentage of dividing cells (cells that have undergone >1 division) is shown for AEC-T cell and DC-T cell cocultures (both at 1:10). Proportion of original population induced to divide after stimulation by AECs and DCs was determined by measuring the number of events in each cell division cohort peak (n). To calculate precursor frequency of responder T cells, percentage of events in each cohort peak n was divided by 2n. Number resulting from this calculation is referred to as undivided cohort number. Sum of these percentages (from division 2 to >7) represents total percentage of cells that started to divide after stimulation in coculture with AECs or DCs. Results are representative of 1 experiment repeated on 7 different donors.

 


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Fig. 8. Characterization of T cell response to AECs and DCs. CSFE-labeled T cells were cocultured with AECs and DCs for 7 days. For stimulation of CD4+ and CD8+ T cells, cells were stained on day 7 with combinations of PE, PerCP, and APC-labeled anti-CD3, -CD4, and -CD8 MAbs. Analysis was performed on nonproliferating (CFSEhigh) and proliferating (CSFElow) CD3+ T cells. Percentage of positively staining cells is indicated. Results are representative of 1 experiment repeated on 7 different donors. For stimulation of memory and naïve T cells, cells were stained on day 7 with combinations of PE-, PerCP-, and APC-labeled anti-CD3, -CD45RA, -CD45RO, and -CCR7 MAbs. Analysis was performed, gating on nonproliferating (CSFEhigh) and proliferating (CSFElow) CD3+ T cells. Percentages of positively staining cells are indicated. Results are representative of 1 experiment using 7 different donors. For stimulation of central and effector memory T cells, cells were stained on day 7 with combinations of PE- and APC-labeled anti-CD3 and -CD62L MAbs. Analysis was performed, gating on nonproliferating (CSFEhigh) and proliferating (CSFElow) CD3+ T cells. Percentages of positively staining cells are indicated. Results are representative of 1 experiment using 3 different donors.

 
Cytokines and apoptosis produced by T cells after stimulation with AECs and DCs. We also determined cytokine (IL-2, IL-4, IL-10, and IFN-{gamma}) production as measured by intracytoplasmic staining in T cells stimulated with AECs and DCs for 16 h. T cells cocultured with AECs and DCs produced IL-2, IFN-{gamma}, and IL-10, but not IL-4 (Fig. 9A). There were, however, variations between T cell preparations in their ability to produce cytokines (data not shown). We also assessed whether T cells stimulated by AECs underwent more apoptosis than T cells stimulated by DCs. We cocultured AECs with T cells and determined T cell apoptosis by costaining with anti-CD3 antibodies and annexin V 16 h after coculture. There was comparable T cell apoptosis (annexin V-positive staining) induced by the AECs and DCs in three different T cell preparations (Fig. 9B).



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Fig. 9. Further characterization of T cell proliferation in response to AECs and DCs. A: cytokine production. Brefeldin A-treated T cells were cocultured with allogeneic DCs and AECs for 16 h and then stained intracytoplasmically with anti-IFN-{gamma}, -GM-CSF, -IL-4, -IL-5, and -IL-10 antibodies. Analysis was performed, gating on live cells. Results are representative of 1 experiment using 4 different donors. B: induction of apoptosis by T cells stimulated with AECs and DCs. T cells were cocultured with allogeneic AECs or DCs, and apoptosis was determined by staining with FITC-labeled annexin V- or PE-labeled anti-CD3 MAbs. Percentage of positively staining cells is indicated. Results are representative of 1 experiment using 3 different donors.

 

    DISCUSSION
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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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We have extended our previous studies investigating the accessory cell function of AECs. We have demonstrated cathepsin enzymes (L, S, and V; Fig. 1), Ii, and HLA-DM-{alpha} and HLA-DM-{beta} (Fig. 2A) and breakdown of Ii in AECs (Fig. 2B). We have colocalized RW with HLA-DR in cultured primary respiratory epithelial cells (Fig. 3) and upregulated its uptake with IFN-{gamma} and GM-CSF (Fig. 4). We have demonstrated that AECs express B7h and B7H-1 (Fig. 6), the expression of which can be upregulated by IFN-{gamma} and GM-CSF (Fig. 6B). We were able to demonstrate that AECs induce proliferation of a restricted population of peripheral blood lymphocytes in an allo-MLR (Fig. 7, A and B) and that this subpopulation is enriched for a surface phenotype, CD45RO+/CCR7, which is considered characteristic of memory effector cells (Fig. 8) (38, 35). This was in contrast to the DC-driven MLRs run in parallel, where proliferative expansion of CCR7+/CD45RO and CD45RO+/CCR7 phenotypes was observed (Fig. 8). There was no difference in cytokine production, type of memory T cell stimulated (effector vs. long-term memory), or apoptosis between T cells cocultured with AECs and DCs.

Several cell types, such as B cells and macrophages, function as professional APCs (41). More recently, we demonstrated that NECs and AECs also possess the capacity to serve as accessory cells for Ag-, anti-CD3-, and mitogen-induced T cell proliferation (34). The respiratory epithelium also expressed the costimulatory molecules CD80 and CD86. In Figs. 1 and 2A, we demonstrate that AECs have mRNA for cathepsins involved in Ag presentation, particularly cathepsin S, which is associated with class II MHC-related Ii cleavage and CLIP formation (33) and HLA-DM transcription required for class II-peptide complex surface expression (9). The demonstration of CLIP, by intracytoplasmic staining of freshly isolated NECs and AECs (Fig. 2B), suggests active Ag processing in respiratory epithelial cells in vivo. The finding of RW Ag endocytosed and colocalized with labeled HLA-DR (Fig. 3) and the upregulation of RW uptake by IFN-{gamma} and GM-CSF (Fig. 4) further support the concept that AECs have the capacity to sample relevant Ag; e.g., RW in the upper and lower airways process it and represent peptide in the context of class II MHC.

The regulation of CD80 and CD86 expression on AECs appears to be complex. We previously reported that freshly isolated NECs and AECs express CD80 and CD86, as detected by RT-PCR and flow cytometry (34). However, we could not upregulate CD80 and CD86 expression on freshly isolated AECs and A549 and BEAS-2B cells or in primary cultured respiratory epithelial cells with IFN-{gamma}, GM-CSF, IL-10, IL-5, and IL-4 (Fig. 5). Our results are similar to those of Wroblweski et al. (44), who demonstrated in 31 non-small cell lung cancer lines that there was constitutive expression of CD80 and CD86 mRNA that could be upregulated by IFN-{gamma} but no increase in the cell surface expression of CD80 or CD86. In Fig. 6, A–C, we found expression of the costimulatory molecules B7h and B7-H1 on AECs. In contrast to CD80, CD86, and B7h, only B7-H1 expression was upregulated after treatment with IFN-{gamma} and GM-CSF (Fig. 6C). In the DCs (Fig. 6C), there was only a limited effect on B7-H1 expression. However, B7h expression was markedly increased by GM-CSF and IFN-{gamma} treatments. This is interesting, because both of these cytokines increased the uptake of RW by primary cultured respiratory epithelial cells at the same concentrations that induced B7-H1 expression (Fig. 4).

B7h and B7-H1 are less well characterized with regard to expression, regulation, and functional outcome of engagement with their recently described receptor counterparts, ICOS and PD-1, respectively. B7h is induced on professional APCs by TNF-{alpha} (39). Engagement of ICOS by B7h may help induce a Th2 immune phenotype and is associated with enhanced IL-10 production. ICOS expression itself is induced by initial CD28-dependent signaling, suggesting a temporal sequence of costimulatory regulation resembling that observed for induction of CD80 expression after initial CD86/CD28 engagement (12, 46). Ligation of B7-H1-costimulated T cell responses to polyclonal stimuli and allogeneic Ag can also stimulate production of IL-10 (28). Prominent B7-H1 neoexpression by lung epithelium-derived tumor lines is described and is implicated in escape from surveillance by antitumor cytotoxic lymphocytes by inducing apoptosis (11). B7-H1 is also highly expressed by syncytiotrophoblasts, which lie in direct contact with maternal blood. It has been suggested that the presence of this molecule at the maternal-fetal interface by nonprofessional APCs is such that it could participate in suppression of activated maternal leukocytes (28). In Fig. 6D, there was no inhibition of T cell proliferation in the AEC-T cell cocultures when anti-B7h or anti-B7-H1 antibodies were added. The inability of anti-B7h antibodies to block T cell proliferation suggests that it may not have a role in AEC-T cell interactions. However, involvement of B7-H1 in T cell activation is more complex: some reports demonstrate that stimulation through B7-H1 results in increased immune responses; other studies indicate a negative role for B7-H1 (43). Although PD-1 was recognized as the ligand for B7-H1, it has been demonstrated that B7-H1 mutants that abolished PD-1-binding sites could still costimulate proliferation and cytokine production of T cells, suggesting an independent costimulatory pathway other than PD-1 (43). Our antibody appears to preferentially block PD-1-B7-H1 interactions, which explains the negative results we obtained in our blocking experiments (Fig. 6D) (19). Interestingly, B7-H2, which has ICOS as its ligand, was recently found to be expressed on AECs (22).

By using in vitro AEC-T cell cocultures, we show that only a small fraction of peripheral T cells, i.e., 0.264%, responded to AECs (Fig. 7A). AECs express several Ag presentation molecules, such as class I and class II MHC, and the costimulatory molecules CD80, CD86, B7h, B7H-1, and B7-H2. We compared the phenotypes of T cells responding to allostimulation by DCs or freshly isolated AECs and found that the AECs induced proliferation of CD4+ and CD8+ T cells (Fig. 7B), which expressed the memory phenotype (CD45RO+ and CCR7+), whereas DCs stimulated naïve (CD45RA and CCR7) and memory CD4+ and CD8+ T cells (Fig. 8). Furthermore, AECs and DCs stimulated central (CD62L+) and effector (CD62L) memory T cells (Fig. 8).

In Fig. 3 and in our previous report (34), we demonstrated that AECs process Ag through a class II MHC route (Fig. 3), leading to the following question: Why are CD8+ T cells proliferating in response to AECs? We previously showed that AECs stimulate CD4+ and CD8+ lymphocytes and that immediate lck and fyn phosphotyrosine kinase activation and subsequent proliferation were reduced by anti-MHC class II antibody pretreatment, as well as by an antibody directed against an epithelial surface membrane protein present on AECs (L12) that we have identified as carcinoembryonic antigen (CEA; unpublished observations; 4, 18). CEA has been shown to be expressed on intestinal epithelial cells, where it is involved in cognate interaction with local T cell populations. Intestinal epithelial cells have been demonstrated to induce proliferation of CD8+ suppressor phenotype T lymphocytes, and CD1d restricts this induction in conjunction with a distinct CEA isotype on the intestinal epithelial cell surface that binds the CD8 molecule directly (5).

Stimulation of memory CD4+ and CD8+ T cells by AECs may in part reflect differences in the requirements for survival and proliferation that are more stringent for primary naïve T lymphocytes than for memory cells in a secondary immune response (40). AECs may be able to provide threshold levels of stimulation to induce proliferation of a pool of primed cells present in small numbers in the periphery that may be represented in the airway microenvironment. T lymphocytes recovered by BAL reveal a predominance of memory effector CD4+ T cells as well as smaller numbers of CD8+ cells (20). After Ag challenge in mice, a rapid accumulation of lymphocytes isolated by BAL consists nearly exclusively of primed, recently activated memory phenotype T lymphocytes (20). Interestingly, lymphocytes harvested after repeat particulate Ag challenge rapidly undergo activation-induced cell death, and the predominant population expresses CD45RO+, CCR7, and CD69+, the primed, memory phenotype, although it is unclear whether macrophages, DCs, or alternative APCs are involved in this phenomenon. BAL-recovered T cells continue to show activation markers, and the majority rapidly undergo FasL-mediated cell death (26). The local cell types that participate as APCs in the downregulation of immune responses and the induction of apoptosis have not been specifically determined, and it is entirely untested whether alternative costimulation, including B7-H1, plays any part in this process. In our studies, we did not detect any differences in the induction of T cell apoptosis by AECs or DCs, although the number of specimens studied was small (Fig. 9B).

Immune responses to a potential airborne Ag depends on complex interactions between accessory cells, Ag, responding T and B cell populations, and local factors in the airway that may influence the clearance of Ag and the function of these cell types. The localization of AECs exposed to the external environment may make them important in the regulation of local immune responses. An improved understanding of Ag presentation by AECs may lead to better treatment of chronic inflammatory disorders of the upper and lower respiratory tract.


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 ABSTRACT
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 RESULTS
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This work was supported by National Institutes of Health Grants AI-44236 and AI-45343 (to K. Sperber).


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Sperber, Immunobiology Center, Box 1089, 1425 Madison Ave., New York, NY 10029 (E-mail: kirk.sperber{at}mssm.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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