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Human Nasal Mucosa Contains Antigen-Presenting Cells of Strikingly Different Functional Phenotypes

Laboratory for Immunohistochemistry and Immunopathology (LIIPAT), Institute of Pathology, and Department of Ear Nose and Throat, University of Oslo, Rikshospitalet, Oslo, Norway

Address correspondence to: Frode L. Jahnsen, LIIPAT, Institute of Pathology, Rikshospitalet, N-0027 Oslo, Norway. E-mail: f.l.jahnsen{at}labmed.uio.no

	Abstract

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Professional antigen-presenting cells (APCs) constitute a heterogeneous leukocyte population that controls T cell induction. Experimental animal studies have delineated the principal APCs of the airway mucosa as a network of intraepithelial dendritic cells (DCs). Whether the situation is comparable in the human airways is unknown. Here we performed a detailed characterization of putative APCs residing in the normal upper airway mucosa, employing confocal microscopy of whole-mount preparations combined with immunophenotyping. A dense network of human leukocyte antigen–DR+ cells with dendritic morphology was found not only in the epithelium (median number, 573/mm²), but also in the lamina propria. In both compartments these cells could be divided into two main populations based on their phenotypic characteristics: the majority expressed a macrophage-like phenotype (CD11b+CD14+CD64+CD68+RFD7+), whereas the smaller population was predominantly constituted by CD1c+CD11c+ immature DCs intermingled with the former. These immature DCs corresponded to the lineage-negative human leukocyte antigen–DR+CD11c+ DC subset present in peripheral blood. Thus, the human upper airway mucosa, in contrast to the rodent counterpart, contains a heterogeneous dense network of dendritic APCs consisting of spatially closely related macrophages and DCs. How these two cell populations regulate the tone of the local adaptive immune system should be the focus of further studies.

Abbreviations: antigen-presenting cell, APC • dendritic cell, DC • lineage-negative, lin^neg • myeloid DC, M-DC • phosphate-buffered saline, PBS • plasmacytoid DC, P-DC • room temperature, RT

	Introduction

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The nasal mucosa is constantly bombarded by a wide variety of exogenous antigens. This situation is a severe challenge to the maintenance of local immunologic homeostasis. To this end, local T cells are believed to play a key role in being able to respond vigorously to microbial pathogens while ignoring harmless ubiquitous environmental antigens. The T cell system is under strict control of professional antigen-presenting cells (APCs), such as macrophages and various dendritic cell (DC) subsets (1, 2). The heterogeneity of DCs exists at several levels, such as anatomic location, phenotype, and function. In peripheral blood, three distinct DC precursor subsets have been identified: CD14+CD11c+ monocytes, lineage-negative (lin^neg) human leukocyte antigen (HLA)-DR+CD11c+ cells (myeloid DCs, M-DCs), and lin^neg HLA-DR+CD123+CD11c- cells (plasmacytoid DCs [P-DCs]) (3, 4).

DCs are widely distributed in most tissues, and different subsets may reside in the same organ (e.g., CD1a+ epidermal Langerhans' cells and CD1a-CD1c+ dermal DCs), but it remains mostly enigmatic how such tissue DCs are related to their precursor counterparts in the circulation. In the normal situation, DCs in peripheral tissue express an immature phenotype with low levels of costimulatory molecules and are functionally characterized by high antigen-capturing but low antigen-presenting capacity (2). After antigen uptake, these immature DCs migrate to draining lymph nodes, where they mature and present major histocompatibility complex (MHC)–peptide complexes to naive T cells and initiate efficiently primary immune responses. The DCs regulate the type of the immune response elicited, and recent in vitro data suggest that this control depends on the diversity of the DC subsets involved. Thus, whereas monocyte-derived DCs induce naive T cells to produce Th1 cytokines, interleukin (IL)-3–stimulated P-DCs initiate a Th2 response (5). Moreover, emerging evidence is accumulating to suggest that DCs also play a critical role in the induction of peripheral immunologic tolerance (6).

Macrophages are also widely distributed in most tissues, but in contrast to DCs that transfer information from the periphery to the regional lymph nodes, tissue macrophages appear to exert their function in situ. These highly phagocytic cells play an important role in innate immunity by engulfing infectious agents and killing them in lysosomes (7). Similar to DCs, they constitutively express MHC class II molecules and are thus able to instruct the adaptive immune system by presenting pathogen-derived peptides to the local T cells (1). Therefore, to understand how the local T cell system is able to successfully maintain immunologic homeostasis in the upper airways, a detailed analysis of local APC subpopulations is required.

Studies in experimental animals have identified DCs as the principal resident APC in the airway mucosa and have shown that these cells form a continuous network of intraepithelial MHC class II+ cells, comparable to the Langerhans' cell network in the epidermis (8). Similarly, a network of intraepithelial HLA-DR+ cells with dendritic morphology has been identified in the human upper airways (9). Only a small proportion of these cells were shown to express DC markers such as CD1a (9, 10), and the same was the case in the lower airways (11). Thus, at present it remains unknown whether DCs or macrophages are the prinicipal APCs in the human upper airways.

To identify APCs in situ, a combination of selected phenotypic characteristics and specific morphologic criteria have to be applied. Tissue-residing immature DCs and macrophages are large cells with extensive projections. Such morphologic characteristics are often difficult to appreciate in conventional thin tissue sections. To overcome this problem, we have established a method employing thick mucosal tissue samples immunostained and prepared for confocal microscopy. This enabled three-dimensional visualization of all positive cells within a depth of 70–100 µm from the surface epithelium. Combined with multicolor immunophenotyping of thin sections, we were able to obtain detailed information about the functional phenotype of mucosal APCs, as well as their morphology and spatial location within the tissue. Our study thus provided crucial information contributing to the understanding of how the respiratory mucosal immune system operates, not only as a defense system against infectious invaders, but also in the pathophysiology of various respiratory immune-mediated disorders.

	Materials and Methods

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Subjects
Mucosal biopsy specimens from the lower turbinate were obtained from 14 patients. For ethical reasons, the sampling was restricted to patients undergoing nasal surgery (primarily for septum reconstruction) but otherwise being healthy. Those with a previous nasal trauma were not subjected to surgery until at least 6 mo after the accident. The specimens were always obtained from a nostril with macroscopically normal mucosa and free airflow. Histologic evaluation examined by an experienced pathologist revealed no abnormalities. The patients had no known allergies (confirmed by a negative skin prick test to a panel of eight common aeroallergens), no recent upper-respiratory tract illness, received no medication at the time of the study, and were without heavy exposure to industrial or otherwise pollutant air. The study was approved by the National Ethics Committee, and informed consent was obtained from each participating subject.

Preparation of Tissue Specimens
Mucosal biopsy specimens from three randomly chosen patients were prepared for whole mounts, whereas the 11 remaining samples were prepared for cryosectioning. The latter specimens were either frozen bedside or pre-fixed in 100% ethanol at 4°C overnight and then washed for 4 h in phosphate-buffered saline (PBS) at room temperature (RT) before freezing. The samples were placed on a thin slice of carrot for appropriate orientation and handling, embedded in OCT (Tissue-Tek; Miles Laboratories, Elkhart, IN), snap-frozen in liquid nitrogen, and stored at -70°C. Cryosections were cut serially at 4–10 µm. Sections of ethanol-fixed samples were air-dried for 2 h, whereas sections of samples frozen bedside were air-dried overnight at RT and then post-fixed in acetone for 10 min. All sections were wrapped in aluminum foil, and stored at -20°C until used. Mucosal samples prepared for whole mounts were immediately placed in 100% ethanol and kept at 4°C overnight. The samples were trimmed with scissors to obtain a tissue thickness of less than < 0.5 mm, and then washed for 5 h in PBS at RT before staining.

Multicolor Immunofluorescence Staining
Whole-mount preparations were incubated for 24 h at RT with mAbs directed against HLA-DR (clone L243, IgG2a, 1/50; Becton Dickinson Immunocytometry Systems [BDIS], San Jose, CA) or CD11c (clone KP90, IgG1, 1/10; gift from K. Pulford, Oxford, UK), or a mixture of anti–HLA-DR and anti-CD68 (clone PG-M1, IgG3, 1/20; DAKO, Glostrup, Denmark) or mAb RFD-7 directed against tissue macrophages (IgG1, 1/100; gift from L.W. Poulter, London, UK). All these mAbs were applied in combination with rabbit antiserum to human laminin (1/100; DAKO). The specimens were washed for 5 h in PBS, and then incubated for 20 h with either Cy3-labeled goat anti-mouse IgG (0.8 µg/ml; Jackson ImmunoResearch Laboratories, West Grove, PA) or FITC-labeled goat anti-mouse IgG2a (10 µg/ml) mixed with Cy3-labeled goat anti-mouse IgG3 (2 µg/ml) or Cy-3 labeled goat anti-mouse IgG1 (2 µg/ml; all from Southern Biotechnology Associates, Inc., Birmingham, AL), always in combination with Cy5-labeled goat anti-rabbit IgG (1/200; Amersham Pharmacia Biotech, Oslo, Norway). The samples were then washed for another 5 h and mounted in custom-made plastic slides.

The immunostaining protocol applied for cryosections has been detailed elsewhere (12, 13). Briefly, to determine the density and phenotype of APCs, an mAb directed against HLA-DR (clone L243, 1/50) was combined with either of the following mAbs: CD1c (clone M241, IgG1,1/5000; gift from R. Blumberg, Boston, MA); CD11b (clone 2LMP19c, IgG1, 1/10; gift from K. Pulford, Oxford, UK), CD11c (clone KP90, 1/10), CD32 (clone IV.3, IgG2b, 1/50; gift from L. Bjørge, Bergen, Norway), CD64 (clone 10.1, IgG1, 1/500; Pharmingen, San Diego, CA), CD68 (clone PG-M1, 1/20), CD80 (clone L307.4, IgG1, 1/100; Pharmingen), CD86 (clone FUN-1, IgG1, 1/100; Pharmingen), CD123 (clone 9F5, IgG1, 2 µg/ml; Pharmingen), rabbit antiserum to FcRI (1/500; Upstate Biotechnology, Lake Placid, NY), and mAb RFD-7 (1/100), which was also mixed with CD14 (clone RM052, IgG2a, 1/300; Biosys, Compeigne, France). To further phenotype CD1c+ cells, we combined anti-CD1c (clone 241) with mAbs to either: CD1a (Clone NA1/34, IgG2a, 1/10; DAKO), CD4 (clone SK3+SK4, IgG1, 10 µg/ml; IBIS), CD14, CD20 (IgG2a, 1/10; BDIS) CD32, CD45RO (clone UCHL-1, IgG2a, 1/10, gift from P. C. L. Beverly, London, UK), CD68, or FcRI. In some staining experiments antiserum to laminin (1/100) was added in the first incubation step.

P-DCs were identified by CD123 (clone 7G3, IgG2a, 2 µg/ml; Pharmingen) combined with either CD4, CD45RA (clone L48, IgG1, 1/10; BDIS), or HLA-DR as previously described (12). All these antibody reagent mixtures were applied for 1 h at RT on serial sections followed by combinations of either Cy3-labeled goat anti-mouse IgG1 (1.5 µg/ml) or IgG3 (2 µg/ml), mixed with FITC-labeled goat anti-mouse IgG2a (10 µg/ml) and Cy5-labeled goat anti-rabbit IgG (1/200; Amersham), or rabbit antiserum to human laminin or human cytokeratin combined with biotinylated goat anti-mouse IgG2a, IgG2b, or IgG3 (all 10 µg/ml from Southern Biotechnology), followed by Cy-3-labeled goat anti-mouse IgG1 together with Cy2-streptavidin (1 µg/ml; Amersham) or 7-amino-4-methylcoumarin-3-acetic acid (AMCA)-labeled goat anti-rabbit IgG (7.5 µg/ml; Vector Laboratories, Burlingame, CA) for 30 min. The laminin and cytokeratin staining was included to delineate theepithelial basement membrane and surface epithelium, respectively.

All staining experiments were controlled for unwanted signals by both omission of primary Abs and by incubation with irrelevant isotype- and concentration-matched primary mAbs.

Immunohistochemical Evaluation
Fluorescent images of immunostained whole-mount preparations were obtained with a confocal laser scanning microscope (Leica, TCS SP, Heidelberg, Germany). The preparations were optically sectioned by scanning at increasing focus depths (typically in steps of 1 µm) from the surface epithelial side. Identical series of optical images at increasing depths (image stacks) from one field were obtained for each wavelength. These image stacks were then merged and three-dimensional information (xyz-axis) obtained was analyzed with Object-Image 2.08 software (an extended version of NIH image). Images were presented as single optical sections or as projections of superimposed optical sections from individual or merged image stacks. Immunostained cryosections were examined blindly at x400 magnification, and cell enumeration was performed as previously described (12, 13). Briefly, to determine the density and phenotype of different APC subsets, cell counts were performed by superimposing a grid (242 x 242 µm) parallel to the basement membrane. The included area of the surface epithelium was determined by measuring the average epithelial height for every grid length and multiplying it by the number of grids examined. At least 1 mm² of tissue section area was examined for each pair of antibodies applied. Data are reported as medians with interquartile ranges.

Statistical Analysis
Spearman's rank correlation test was performed to evaluate the relationship between the density of APC subsets in the epithelium and lamina propria. Data are reported as medians with interquartile ranges.

	Results

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Whole-mount preparations of normal nasal mucosa examined by confocal microscopy, revealed a dense network of HLA-DR+ cells in the surface epithelium (Figure 1a); three-dimensional reconstruction of the image stacks showed that the largest surface area of these cells was parallel to the basement membrane (not shown). The cell density (median number per mm²: 573; interquartile range: 480–743; n = 11) was similar to that previously reported for intraepithelial MHC class II+ cells in the tracheal mucosa of rats (8). Notably, a similar dense network of HLA-DR+ cells was also observed in the upper parts of the lamina propria, displaying an even more impressive dendritic morphology with cellular processes extending >20 µm (Figure 1b). These characteristic intraepithelial and lamina propria APCs were positive for CD11c (Figure 1c), a marker known to be expressed on myeloid-derived APCs (14).

View larger version (70K): [in this window] [in a new window]   Figure 1. Immunofluoresence staining for human leukocyte antigen–DR (HLA-DR; a and b) and CD11c (c) in whole-mount preparations of normal nasal mucosa. The images represent superimposed optical sections of image stacks along the z axis of epithelium (a, total depth 15 µm) and lamina propria (b, depth 15 µm; c, depth 10 µm), respectively.

View larger version (132K): [in this window] [in a new window]   Figure 2. In situ phenotypic characterization of human mucosal APC subsets. Multicolor immunofluorscence staining for: (a–c, k) HLA-DR (green) and CD68 (a), RFD-7 (b), CD64 (c), or CD1c (k) (red), and laminin (blue in a and c); (g) RFD-7 (green) and CD14 (red); (h–j) Cd1c (red) and CD68, CD1a, or CD11c (green) in cryosections of normal nasal mucosa. (d–f) Paired immunostaining for HLA-DR (green) and CD68 (red) in whole-mount preparation of normal nasal mucosa; the image is a Z-projection of 18 superimposed optical sections (total depth 15 µm) from a field of lamina propria. A CD68-HLA-DR+ cell (f, arrow). (l–n) Paired immunostaining for CD11c (red) and CD2 (green) in cryosection of normal nasal mucosa. (m and n) Two consecutive optical sections (0.8-µm steps) of intraepithelial cells. (l) A superimposed image of five consecutive optical sections including m and n. Coexpression of markers is shown by yellow (some indicated by arrows). (a–c, g–i) Note similar distribution on epithelium and lamina propria of double positive cells for most markers except for less CD14+ cells in the epithelium (g) and lack of CD1a+ cells in the lamina propria (i). Broken line indicates the epithelial basement membrane and asterisk indicates the luminal side. Scale bars: a–c, g–k, 30 µm; e, 15 µm; and l, 10 µm.

The other main APC subset was identified by its expression of the DC marker CD1c (Figure 2h) that decorated 33% of the HLA-DR+ cells in the epithelium (interquartile range: 29–36%; n = 11). Notably, of all intraepithelial HLA-DR+ cells, the proportions of CD68+HLA-DR+ and CD1c+HLA-DR+ cells added up to a median of 102% (interquartile range: 94–107%), which strongly suggested that these two subsets in fact comprised the entire intraepithelial APC population. The HLA-DR+CD1c+ population was clearly distinct from macrophages as it was virtually negative for CD68 (Figure 2h). The Langerhans' cell marker CD1a, often used to identify airway DCs (16–18), was virtually confined to CD1c+ cells and decorated 40% of this subset in the epithelium (Figure 2i). Notably, those situated in the lamina propria were almost invariably negative for CD1a (Figure 2i).

As shown in Figure 3, the compartmental density of CD1c+HLA-DR+ cells in the lamina propria was quite similar to that found in the epithelium (median number per mm²: 108; interquartile range: 103–156 versus 166; 143–216, respectively; n = 11). Conversely, the density of CD68+RFD7+HLA-DR+ cells in the lamina propria was lower than that found in the epithelium (median number per mm²: 195; interquartile range: 165–227 versus 429; 328–526, respectively; n = 11).

View larger version (21K): [in this window] [in a new window]   Figure 3. Relationship between numbers of CD1c+HLA-DR+ (squares) and CD68+HLA-DR+ (triangles) cells in the epithelial versus the lamina propria compartment in cryosections of nasal mucosa (n = 11). The subjects are indicated by numbers to enable individual comparison of the compartmental density of the two cell subsets. There was no significant correlation between the number of CD1c+HLA-DR+ cells (rs = -0.27, P = 0.42) and CD68+HLA-DR+ cells (rs = 0.21, P = 0.54) in the epithelium and lamina propria, when analyzed separately.

A prerequisite for T cell/APC interactions is physical contact. Therefore, we examined the spatial relationship between APCs and T cells. By combining T cell markers (CD2 or CD3) with two pan APC markers (CD11c or HLA-DR), we identified many resident T cells in close contact with dendritic projections extending from APCs (Figures 2l–2n). All HLA-DR+ APCs, both in the epithelium and lamina propria, were negative for the costimulatory molecule CD80, and only some expressed CD86 at very low levels (not shown).

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
We show here for the first time that the upper airway mucosa of humans contains a dense network of dendritic APCs both in the epithelium and the lamina propria. In the surface epithelium of rodent airways an APC population of similar morphologic appearance and cell density has been reported, although showing a strikingly different phenotype (8). Whereas intraepithelial MHC class II+ cells in the rat trachea by morphologic and phenotypic criteria were deemed to be almost exclusively DCs (8), the dendritic HLA-DR+ cells in human nasal mucosa could be divided into two main populations based on their phenotypic characteristics: macrophages constituted the majority, whereas immature DCs were fewer and intermingled with the former.

By conventional immunophenotyping performed on thin tissue sections, airway APCs were previously believed to comprise a morphologically heterogeneous population of cells (9). More recently, Holt and coworkers (8) established a novel sectioning technique cutting the trachea tangentially along the epithelium, and were able to demonstrate that all intraepithelial MHC class II+ cells in the trachea could be accounted for by a network of cells with classical DC morphology. However, that technique allowed only visualization of APCs within the epithelium. Therefore, little is currently known about the morphologic and phenotypic characteristics of APCs in the airway lamina propria. To perform a detailed analysis of APC subsets both in the epithelium and the lamina propria of human airways, we examined immunostained whole mounts of mucosal tissue by confocal microscopy. With this technique all reactive epitopes within a thick tissue sample (70–100 µm) could be visualized, thereby rendering a three-dimensional reconstruction of immunostained cells and other tissue elements possible. Viewing the mucosal tissue from the surface side, we identified a dense network of evenly distributed large HLA-DR+ cells in the epithelium, very similar to that described in rat trachea.

Notably, however, an even more impressive network of highly dendritic HLA-DR+ cells was observed in the lamina propria, although the compartmental density was almost two times higher in the epithelium. These cells displayed distinct characteristics by projecting numerous long processes (> 20 µm) with total cell lengths up to 80 µm. Surprisingly, therefore, despite their highly dendritic morphology the majority of the HLA-DR+ cells in the epithelium as well as in the lamina propria expressed a macrophage phenotype (CD11b+ CD11c+CD14+CD32+CD64+CD68+RFD-7+). Macrophages are often considered as pleomorphic, large rounded cells with few dendrites, but this presumption is mainly based on microscopy of thin tissue sections cut perpendicular to the surface epithelium. Our results showed that by such a conventional approach only an arbitrary part of the cell structure is displayed and dendritic processes are difficult to identify (Figures 2a–2c). Indeed, in accordance with our results, similar studies applying whole-mount preparations of brain and eye tissue from rats have clearly demonstrated a network of macrophages with morphologic characteristics comparable to DCs (19, 20). Although immature DCs have been shown to express CD68 and some of the other macrophage markers described above (21), the unique phenotype displayed by the nasal CD68+ cells, including their reactivity to RFD7, strongly suggested that these cells indeed are mature macrophages and not DCs. However, the functional capacity of these cells remains to be elucidated.

The other main HLA-DR+ cell population in human nasal mucosa was phenotypically distinct from macrophages as they did not express significant levels of CD68 but reacted with a mAb to the nonclassical MHC receptor CD1c. A recent study (14) showed that most M-DC precursors in peripheral blood likewise express high levels of this marker, and we therefore wanted to investigate whether the nasal CD1c+ APC was derived from these circulating precursors. Indeed, the vast majority of these nonmacrophage APCs displayed a phenotype (CD4+ CD11c+ CD14- CD32± CD45RO+CD80-CD86-/lo CD123- FcERI+) similar to their counterparts in peripheral blood (14). This strongly suggested that the nasal population of immature DCs was derived from the circulating M-DC precursor subset. Interestingly, in keeping with this possibility, we recently showed that the same DC phenotype accumulated rapidly (within 4–5 h) in the bronchial mucosa after allergen challenge of patients with allergic asthmatic (22). We are currently investigating whether this DC subset accelerates its migration also into the upper airways during allergic inflammation.

Originally, a marker for Langerhans' cells in the epidermis, CD1a has been widely used to identify DCs in human airways (16–18). In agreement with others, we found that CD1a+ cells in normal nasal mucosa were preferentially localized in the epithelium. Immunologic costaining experiments showed that CD1a was almost completely confined to the CD1c+CD11c+HLA-DR+ intraepithelial population and stained 40% of those cells. It has been shown that M-DCs isolated from blood upregulate CD1a in vitro (23). Thus, it is possible that CD1a+ DCs in the airways originate from the M-DC precursor subset in circulation, and express CD1a after migration into the surface epithelium.

Recent information on DC biology suggests high levels of heterogeneity and plasticity (24). In vitro, CD14+ monocytes can give rise to either tissue macrophages or DCs. How this differentiation is regulated in the tissue is not understood, but a recent in vitro report showed that IL-6 produced by fibroblasts or epithelial cells interacted with M-CSF and switched monocyte differentiation to macrophages rather than DCs (25). A similar mechanism appears to operate in nasal mucosa in the normal state, as suggested by the fact that we found most APCs to be mature macrophages and that the main fraction of immature DCs most likely was derived from blood M-DCs. It has, moreover, been shown that macrophages can be redirected into DC differentiation with a shift in culture conditions (26). Thus, it is conceivable that in a situation where tissue damage is beyond the capability of first-line innate immunity and local repair, resident macrophages may convert into immature DCs that travel to the draining lymph nodes to stimulate adaptive immunity. Based on the detailed information derived from this study elucidating the phenotype, morphology, and localization of upper airway macrophages, it would be possible to examine whether redirection of macrophage differentiation occurs in various inflammatory reactions in vivo.

The human nose is the entry site for a large variety of airborne antigens, including a range of infectious microorganisms, especially viruses; it thus provide an extremely challenging environment for the maintenance of immunologic homeostasis. Within this mucosal environment APCs play a key regulatory role via sampling and presenting peptides in an immunogenic form to the adaptive immune system. The dense nasal network of APCs is located in an ideal position to perform a variety of functions as reflected by its content of both macrophages and different immature DC subsets. This complementary cellular composition of the network constitutes an efficient defense against pathogenic intruders apparently while maintaining immunologic homeostasis.

We have previously shown that a small number of P-DCs emigrate to the nasal mucosa independent of inflammation (12). These cells were previously believed to traffic directly from bone marrow via blood to secondary lymphoid organs. In vitro studies have shown that these cells produce large amounts of type I interferons in response to viruses (27). It is possible, therefore, that these cells play an important role in both innate and adaptive immune defense against various viruses. The upper airways, which are a primary site for viral infections, apparently constitute a microenvironment that preferentially attracts these cells also in the normal situation. We have recently shown that P-DCs accumulate in skin lesions of type I interferon–related disorders (28, 29) and Th2 cell–dominated allergic reactions in the upper airways (12). Whether P-DCs are recruited to the nasal mucosa during viral infections is currently not known.

Evidence strongly suggests that APCs, especially DCs, are decisive for the induction of productive immune responses as well as tolerization (6). Our observation that APCs also in the normal situation are in direct contact with T cells indicated that a constant immunologic control by resident APCs are performed locally. How the APC populations identified here regulate the tone of the local adaptive immune system should be the focus of further studies.

It is important to understand mechanisms underlying abrogation of the fine-tuned balance between rapid responses against pathogens and hyporesponsivess toward harmless proteins (such as pollen) or self antigens to avoid hypersensitivity reactions or autoimmunity. Comparisons of the density, localization, and functional phenotype of local macrophages and DCs in inflamed tissue with the normal situation are needed to understand the role played by these regulatory cells in upper respiratory tract disorders.

	Acknowledgments

 
Studies in the authors' laboratories are supported by the University of Oslo, the Research Council of Norway, the Norwegian Cancer Society, and the Norwegian Society for Asthma and Allergy.

Received in original form October 30, 2002

Received in final form June 23, 2003

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