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Heterogeneity of Human Nasal Vascular and Sinusoidal Endothelial Cells from the Inferior Turbinate

Divisions of Clinical Immunology, Otorhinolaryngology, and Transplantation Surgery, Karolinska University Hospital-Huddinge, Karolinska Institutet, Stockholm, Sweden

Correspondence and requests for reprints should be addressed to Carolina Holmén, Department of Clinical Immunology, F79, Karolinska University Hospital-Huddinge, S-141 86 Stockholm, Sweden. E-mail: carolina.holmen{at}kus.se

	Abstract

Top Abstract Introduction MATERIALS AND METHODS Results DISCUSSION References

 
The vast heterogeneity of endothelial cells (EC) in various organs necessitates isolation of EC from the relevant organs when defining mechanisms of site-specific pathologies. We report a novel finding that describes the presence of two heterogeneous populations of human nasal microvascular EC isolated from the inferior turbinate. Light and electron microscopy, flow cytometric analysis, and immunocytochemistry analysis demonstrated that one EC population exhibited the classic vascular endothelial markers with cobblestone-like morphology, whereas the other was sinusoidal with fusiform morphology. The sinusoidal EC (SEC) lacked surface expression of the endothelial markers CD31 and E-selectin, were discontinuous, showed fenestrae and pinocytic vesicles, and did not form tight junctions. Gene expression analysis using microarray revealed significant but limited heterogeneity between the two cell types. Immunohistochemical staining of normal nasal biopsies confirmed the presence of two distinct populations of EC. We found that CD31 was exclusively expressed on vascular EC (VEC), whereas the molecule L-SIGN was mainly expressed on SEC. Both cell types formed capillary-like tubules in matrigel in vitro. The two heterogeneous EC populations provide a unique in vitro system to study the biology of nasal VEC and SEC in normal conditions and in inflammatory processes in various nasal disorders.

Key Words: endothelial cell • inferior turbinate • vascular endothelial cell • sinusoidal endothelial cell • L-SIGN

	Introduction

Top Abstract Introduction MATERIALS AND METHODS Results DISCUSSION References

 
Endothelial cells (EC) are key players in various biologic processes, including leukocyte trafficking and homing, inflammation, wound healing, tumor metastasis, and angiogenesis (1–2). EC are the "gatekeepers" of the tissues from the perspective of the blood stream, and therefore it is likely that this site suffers the first major insult by the immune system (3). Thus, EC are important targets for studying mechanisms underlying the pathogenesis and progression of various organ disorders. Investigations of these processes have traditionally used EC isolated from easily accessible tissues such as large vessels of the human umbilical cord. Today, with more recent knowledge in the field of EC heterogeneity, there are many fundamental differences between different organs within the vascular loop of a given organ and between neighboring EC of a single blood vessel (1–4). For example, the liver, spleen, and bone marrow sinusoids are lined by discontinuous EC that allow cellular trafficking between intracellular gaps, whereas tight junctions are present between EC in the brain (1). The endocrine glands and kidneys are lined by fenestrated EC that facilitate selective permeability required for sufficient absorption, secretion, and filtering (5). EC heterogeneity is also evident in individual organs. For example, the kidney contains fenestrated EC in its peritubular capillaries, discontinuous EC in its glomerular capillaries, and continuous EC in other regions (6). The great morphologic, biochemical, and functional heterogeneity that EC express necessitates the development of techniques to isolate microvascular EC from different tissues to create relevant in vitro models for guidance of what an in vivo situation might resemble.

The microvasculature of the nasal mucosa is unique from other parts of the respiratory tract in many ways. It has a copious system of capacitance vessels or sinuses and many arteriovenous anastomoses, which are absent or far less frequent in the tracheobronchial tree. The nasal vasculature shows cyclic changes of congestion not observed in the lower airways. Furthermore, capillaries with and without fenestrae allowing rapid escape of water into the lumen for evaporation and heat loss are present in the nasal mucosa (7, 8). These factors make the nasal EC important and interesting targets for studying their role in thermal regulation, humidification, filtering, and conditioning of inspired air. Furthermore, microvascular EC play a central role in the development of immune responses by regulating leukocyte recirculation and antigen presentation to T lymphocytes (9). Thus, studying the interaction of the immune cells with nasal EC in normal and pathologic conditions is important in understanding mechanisms underlying the pathogenesis of various nasal disorders.

The morphologic, phenotypic, and molecular heterogeneity of the different EC types in the nasal tissue has not been studied. The aim of this study was to establish a simple and reproducible method for isolation and cultivation of human nasal microvascular endothelial cells (HNMEC) from the inferior turbinate. For comparison, we used vascular endothelial cells (VEC) from human umbilical cord vein and sinusoidal endothelial cells (SEC) isolated from a normal healthy liver as control cells. Our interest in establishing such cell lines is primarily to elucidate the mechanisms underlying the pathogenesis of Wegener granulomatosis (a vasculitis disease with inflammatory complications in the nasal region) by studying the interaction of immune cells with HNMEC. However, we believe these cells will also provide a unique in vitro system to study the biology of these cells in various nasal disorders such as allergy rhinitis, nasal polyposis, and other inflammatory diseases of the nasal mucosa.

	MATERIALS AND METHODS

Top Abstract Introduction MATERIALS AND METHODS Results DISCUSSION References

 
Monoclonal Antibodies
Monoclonal antibodies used are listed in Table 1.

For magnetic isolation, Mini MACS magnetic beads were used (Miltenyi Biotec, GmbH, Bergisch, Germany). Matrigel and multiwell eight-chamber culture slides were purchased from Becton Dickinson (Franklin Lakes, NJ) and 4,6-Diamidino-2-phenylindole (DAPI) from Roche Diagnostics Scandinavia AB (Bromma, Sweden). Normal goat serum, Vectastain Elite ABC kit, and avidin/biotin blocking kit were purchased from Vector Laboratories (Burlingame, CA). As positive controls, a skin fibroblast primary cell line (a generous gift from Prof. Karl-Gösta Sundquist, Huddinge Hospital, Sweden), biliary epithelial cells isolated by us as described previously (10), and an aortic smooth muscle cell line purchased from Clonetics (BioWhittaker) were used.

Human Nasal Tissue Specimen
Human nasal tissue specimen (1–2 cm) was obtained from patients scheduled for surgery due to structural nasal deformities, such as septal deviation. Apart from their nasal deformity, patients were healthy and did not exhibit any pathophysiologic condition that might have changed the isolated in vitro EC phenotype. Informed consent was obtained from each patient, and approval was obtained from the ethics committee. During the surgical procedure, tissue biopsies from the inferior turbinate were resected using a scalpel with an atraumatic technique to reduce the damage of the tissue specimen. The tissue was placed into RPMI 1640 medium, and the isolation of HNMEC was performed within 3–4 h after the operation. Six nose biopsies were obtained from different individuals; four were used for isolation of cells and two for immunohistochemical staining.

Isolation and Cultivation of HNMEC and HLSECs
The nasal tissue specimen was cut into 2-mm² sections and enzymatically digested using 0.2% collagenase type IV solution for 5–7 min in a 37°C waterbath, washed with MCBD 131 medium containing 5% FCS (GIBCO), and placed in 0.2% gelatin-coated culture plates. Each 2-mm² section was placed in individual wells and incubated with the EC-selective medium MCBD 131 containing 2 ng/ml of vascular endothelial growth factor. After 24 h, the medium was discarded, and the culture plate was washed twice to remove floating cells. Fresh medium supplemented with 10% heat-inactivated human AB serum and EGM-2 singlequots was added. The tissue pieces were removed from the culture wells after 48 h, and cells were allowed to grow until they reached confluence.

Confluent cells from all wells were detached by trypsinization, pooled, and washed once with phosphate-buffered saline (PBS). EC were isolated with CD144-coated Mini MACS beads according to the manufacturer's instructions. The number of CD144-positive HMNEC obtained from each biopsy piece was counted and viability tested using ethidium bromide. HNMEC were cultured in 0.2% gelatin-coated culture wells at 37°C in a humidified atmosphere of 95% air and 5% CO₂. The medium was replaced every 2–3 d.

During cultivation, we noticed two morphologically distinct populations of HNMEC: one with cobblestone and the other with fusiform and elongated morphology. Colonies of each cell type were trypsinized individually by the addition of a small drop of trypsin-EDTA (1x) on each colony. After 20 s, the detached cells were manually picked by a thin (20 µl) microtiter pipette and transferred to separate culture dishes. Both cells types were manually picked and cultured separately under identical conditions for further analysis. Both cell types maintained their identity throughout their cultivation time. When cross-contamination was evident, the cell fraction was not used for further analysis.

In our laboratory, we have previously isolated and cultivated well characterized HLSEC (11). HLSEC were isolated according to a method described previously (11) and cultivated in complete MCBD 131 medium. HUVEC were purchased and cultivated in recommended medium from Clonetics.

The cultured cells were phenotypically and morphologically characterized by flow cytometry, immunocytochemistry, and electron microscopy. In addition, molecular differences between the cobblestone and fusiform HNMEC were detected by microarray analysis. All cells were used in passages 3–6 for the present study.

Phenotyping of HNMEC, HUVEC, and HLSEC by Flow Cytometry
Single-color fluorescence was used to phenotypically characterize (see Table 1) unstimulated and cytokine stimulated (overnight with 20 ng/ml TNF- and 200 ng/ml IFN-) (R&D Systems) cobblestone and fusi-form HNMEC as described previously (11). HUVEC and HLSEC were used as control cells. Briefly, cells were trypsinized, washed twice in PBS, and incubated with the primary antibody for 30 min at room temperature (rt). Primary antibodies used for staining were anti-CD141, -CD142, -CD144, -Ac-LDL, -CD106, -CD62E, -CD31, -vWF, -CD105, -L-SIGN (a molecule shown to be specifically expressed on lymph node and liver SEC), -EPCAM, -fibroblast, and --actin. Cells were washed twice, and a secondary goat-anti mouse IgG antibody (FITC) was added and incubated at 4°C on ice for 30 min.

Intracellular staining for -actin was performed after cell permeabilization using 5% saponin in PBS (11). The stained cells were suspended in 200 µl PBS and further analyzed by a flow cytometer (FACSsorter; Becton Dickinson, San José, CA). Fluorescence signals from 10,000 cells were recorded and analyzed by the CellQuest software.

Immunocytochemistry
Cobblestone and fusiform HNMEC were grown separately on 0.2% gelatin-coated, eight-well chamber slides. Cells were rinsed with PBS, fixed in 30% acetone/methanol for 10 min at rt, washed twice with PBS, and incubated with 3% bovine goat serum for 20 min at rt. The slides were rinsed twice with PBS and incubated for 30 min at rt (dark) with primary monoclonal antibodies (see Table 1) against CD31, CD144, vWF, and L-SIGN. Cells were stained with a secondary FITC-conjugated goat anti-mouse antibody for 30 min at 4°C (dark). After two washes with PBS, cells were incubated with DAPI (a blue fluorescence DNA stain) for 40 s. Finally, the cells were washed with PBS, mounted with glycerol, and visualized under a fluorescence microscope. Cells stained only with the secondary antibody served as controls.

Scanning Electron Microscopy
HNMEC were grown to confluence on membrane filters in 24-well plates. The membranes were prepared for scanning electron microscopy (SEM) as described elsewhere (11). Briefly, membranes were fixed in 2% glutaraldehyde, briefly rinsed in distilled water, placed in 70% ethanol for 10 min, placed in 99.5% ethanol for 15 min (all at 4°C), and dried. After drying, the membranes were cut off, mounted on an aluminium stub, and coated with 15-nm platinum (Polaron Components Group, Watford, Herts, UK). The samples were analyzed in a Jeol JSM-820 scanning electron microscope at 15 kV.

Transmission Electron Microscopy
Cells were fixed as described previously, and a further procedure was followed as described elsewhere (11). In short, after fixation, the membrane was cut free and fixed for 1 h at 4°C in a buffer containing 0.15-mol/l sodium cacodylate, 1% osmium tetraoxide, and 3 mmol/l CaCl₂ (pH 7.4). The wells were rinsed briefly in 0.15 mol/l sodium cacodylate buffer, dehydrated in ethanol as described previously, and imbedded in Spurr resin (Agar Scientific Ltd., Essex, UK). The sections were contrasted with uranyl acetate followed by lead citrate and examined at 80 kV in a Leo 906 (Oberkochen, Germany) transmission electron microscope.

In Vitro Angiogenesis Assay
Formation of capillary tube-like structures of untreated CD31⁺ and CD31^– HNMEC was assessed in a solubilized basement membrane preparation extracted from the Engelbreth-Holm-Swarm mouse sarcoma, frequently used for the evaluation of in vitro angiogenesis. Twenty-four well plates were coated with 200 µl of Matrigel (pre-gelled for 30 min at 37°C in 5% CO₂), and the cells were seeded on the polymerized matrix at a density of 5 x 10⁴ cells/well. After 18 h at 37°C in 5% CO₂, the resulting tube-like structures were examined using a phase-contrast light microscope. HUVEC were used as positive control cells.

Gene Microarray and Data Analysis
Total RNA from unstimulated CD31⁺ and CD31^– HNMEC was prepared with the RNeasy kit (QIAGEN). The total RNA quality was checked using the Agilent Bioanalyser. Probe synthesis from total RNA samples, hybridization, detection, and scanning were performed according to standard protocols from Affymetrix, Inc. (Santa Clara, CA). Fifteen micrograms of labeled cDNA were hybridized to each Human Genome U133 Plus 2.0 gene chip (Affymetrix). Gene chips were scanned (GeneChip Scanner 3,000), and signals obtained by scanning were processed by the GeneChip Operating Software (Affymetrix).

Immunohistochemistry
To confirm the presence of the two distinct types of EC in vivo, we stained normal nasal biopsies with antibodies to various endothelial and epithelial markers (see Table 1). Normal liver biopsies were used for comparison. Nasal and liver biopsy tissues were cut in cross-sections of 5 µm using a cryostat microtome (Leica CM3050S). The sections were mounted on glass coverslips, air-dried at rt, and fixed in 30% acetone/methanol for 10 min. The sections were washed with PBS and pre-treated with 3% H₂O₂ for 5 min to block any endogenous peroxidase activity. Sections were washed with PBS again and incubated with 2% normal serum, avidin, and later biotin (avidin/biotin blocking kit, Vector Laboratories) each for 15 min at rt. The sections were washed again with PBS and incubated with the primary antibody for 30 min at rt. As a negative control, the primary antibody was replaced with 2% normal serum. Visualization of the primary antibodies was performed using Vectastain Elite ABC kit reagents, and diaminobenzidine tetrahydrochloride, giving a brown staining, was used as peroxidase color developer. Sections were counterstained with hematoxylin, mounted with glycerol, and visualized by light microscopy.

The nasal and liver biopsy sections were single or double immunofluorescence stained according to the Vectastain Elite ABC protocol described previously. Briefly, sections were stained with antibodies against -L-SIGN, -CD31, -CD144, -CD141, -vWF, CK18, and -EPCAM, followed by staining with secondary anti-mouse specific FITC or Texas red conjugated antibodies. After three washes with PBS, sections were incubated with DAPI for 40 s. Cells stained only with the secondary antibody served as controls. The sections were mounted with glycerol and visualized by fluorescence microscopy.

	Results

Top Abstract Introduction MATERIALS AND METHODS Results DISCUSSION References

 
Culture and Morphologic Characterization of HNMEC, HUVEC, and HLSEC
After isolation, the number of magnetically isolated CD144⁺ cells was counted, and cells were viability tested with ethidium bromide. The size of the nasal specimen influenced the number of isolated CD144⁺ HNMEC, whereas the viability and number of passages obtained did not differ significantly between different isolates (Table 2A). The HNMEC grew in several clusters that formed monolayers within 1 wk. During cultivation, we observed that the cell cultures contained two morphologically distinct populations, one cobblestone and one fusiform. When each cell type was grown separately, the cells grew in monolayers and showed contact inhibition. At confluence, both cell types could be maintained with stable morphology for 11 passages, after which the cells ceased to divide. In general, the number of cobblestone-like cells obtained from CD144⁺ cell cultures was higher as compared with the fusiform cells (Table 2B). Figure 1 shows phase-contrast micrographs of the two populations of HNMEC in comparison with HUVEC and HLSEC.

View larger version (44K): [in this window] [in a new window]   Figure 1. Phase-contrast micrographs of endothelial monolayers. Confluent layers of CD31+ cobblestone human nasal microvascular endothelial cells (HNMEC), CD31– fusiform HNMEC, human umbilical vein endothelial cells, and human liver sinusoidal endothelial cells (original magnification: x20).

HUVEC were positive for all tested EC markers but not for fibroblast, epithelial, and smooth muscle cell markers, used as controls.

HLSEC expressed same EC markers as the fusiform CD31^– HNMEC and did not express vWF, EPCAM, and markers for -actin and fibroblasts. The skin fibroblast cell line, epithelial cell line, and the smooth-muscle cell line stained positive with their respective antibodies (the anti-fibroblast, -EPCAM, and - actin antibody) (Table 3).

Table 3 shows the phenotypic characteristics of cobblestone CD31⁺ and fusiform CD31^– HNMEC, HUVEC, and HLSEC. CD31^– were found in the fusiform cell population, whereas all cells with cobblestone morphology expressed CD31 (Table 3). The purity of each cell type was 93–95%, as determined by flow cytometric analysis with anti-CD31 antibodies. Furthermore, phenotypic analysis of cells in various passages (up to 11 passages) showed stable phenotype as determined by the panel of markers given in Table 3.

In vitro Cultured Fusiform CD31^– but not Cobblestone CD31⁺ HNMEC Stain Positively for L-SIGN
To further confirm the in vitro EC characteristics of cobblestone CD31⁺ and fusiform CD31^– HNMEC in culture, we analyzed the cells using immunocytochemistry. Slightly different staining patterns were observed with the two HNMEC types. Fusiform CD31^– HNMEC stained for CD144, vWF, and L-SIGN, with a staining pattern uniformly distributed throughout the cell surface (Figure 2A), but showed no staining with anti-CD31 antibodies (Figure 2A). On the other hand, cobblestone CD31⁺ HNMEC cells did not stain for L-SIGN but stained positively for CD31 and CD144 at junctional locations, and a patchy/granular staining was observed with anti-vWF antibodies (Figure 2B). The cells were counterstained with nuclear DAPI staining.

View larger version (30K): [in this window] [in a new window]   Figure 2. Immunofluorescence staining of human nasal microvascular endothelial cells. (A) Fusiform CD31– human nasal microvascular endothelial cells (HNMEC) stained positively for CD144, vWF, and L-SIGN but not for CD31. (B) Cobblestone CD31+ HNMEC stained positive for CD31, CD144, and vWF but not L-SIGN. The cells were counterstained with DAPI staining the nucleus blue. No staining is seen with control cells (original magnification: x20).

View larger version (83K): [in this window] [in a new window]   Figure 3. Scanning and transmission electron micrographs and in vitro micrographs of capillary tube formation. Scanning electron micrograph showing fenestrae (arrows) in the cytoplasm (A) and a large amount of pinocytic vesicles (arrows) just beneath the plasma membrane are visible (B). (C) Transmission electron micrograph showing an endothelial cell (c) attached to the membrane (m) without any basement membrane (*) and no tight junctions are observed. When plated on Matrigel, a network of capillary tube formation was observed with (D) cobblestone CD31+ human nasal microvascular endothelial cells and (E) human umbilical vein endothelial cells, whereas fusiform CD31– HNMEC showed tubule-like formation but no network of branching tubules (F).

Gene Expression Analysis of CD31⁺ Versus CD31^– HNMEC
From one representative individual, we analyzed the gene expression profile in untreated CD31⁺ and fusiform CD31^– HMNEC at passage 1 isolated from the nasal inferior turbinate. A broad expression profile was obtained with the Human Genome U133 Plus 2.0 array (around 38,500 genes, where some genes were represented by more than one probe set in the microarray). For simplicity, we selected our analysis to genes considered to be involved in EC biology as specified in the GEArray Q Series Human Endothelial Cell biology Gene Array kit (SuperArray Bioscience Corporation, MedProbe, Oslo, Norway).

From 262 identified transcripts, 43 genes were found to be regulated (e.g., having a significant ±2-fold change in transcript levels) between the two cell types giving call signals (i.e., "increase" or "decrease"). In Figure 4, each sample signal value is introduced into a 100% stacked bar format, which compares the percentage each value contributes to a total across categories. Among the regulated genes involved in EC biology, five genes in the sinusoidal HNMEC showed increased expression as compared with the other genes. These genes encode matrix metalloproteinase-1 (MMP1), chemokine ligand 2 (monocyte chemotactic protein-1 [MCP-1]), CD106, collagen type 1, and the TNF receptor superfamily member 11b (osteoprotegerin). Vascular HNMEC showed increased expression of genes encoding CD58, CD142, fibronectin 1, vascular adhesion protein-1, and low-density lipoprotein receptor. The FACS and the array data for the cells correlated in most cases. Immunohistochemistry showed two distinct patterns of EC staining in nasal biopsies.

View larger version (102K): [in this window] [in a new window]   Figure 4. Microarray gene expression analysis of cobblestone CD31+ and fusiform CD31– human nasal microvascular endothelial cells (HMNEC). In this representative 100% stacked bar figure, signal values are shown from the significantly regulated 43 genes involved in endothelial cell biology. In general, the cobblestone CD31+ and fusiform CD31– HMNEC had a similar expression profile. However, a few genes in the fusiform CD31– HNMEC showed higher expression for MMP-1, MCP-1, VCAM-1/CD106, collagen type 1, and osteoprotegerin, whereas the cobblestone CD31+ showed a higher expression for the LDL receptor, fibronectin 1, vascular adhesion protein-1, CD58, and CD142. Light gray bars, vascular CD31+ HNMEC; dark gray bars, sinusiodal CD31– HNMEC.

View larger version (56K): [in this window] [in a new window]   Figure 5. Immunofluorescence staining of normal human nose and liver biopsy cryosections. Nasal biopsy sections stained by immunofluorescence with antibodies to CD144 and CD31 showed coexpression of the two markers (merged yellow) on nasal endothelial cells (EC) (A). Similarly, EC showed coexpression of CD144 and L-SIGN (merged yellow) (B). However, CD31 expression (green) was found on EC in capillaries lying interdispersed between the CK18 positive epithelial cells (red) (C), whereas L-SIGN (green) stained EC in capillaries directly beneath the CK18 positive epithelium (red) (D). Control biopsy section, stained with only secondary antibodies, showed staining of only nucleus with DAPI (E). Double staining of nasal biopsy with CD31/CK18 (F) and L-SIGN/CK18 (G) at lower magnification give an overview of the distinct pattern of staining observed with anti-CD31 and anti-L-SIGN antibodies. A similar finding is observed in the liver (H). Expression of CK18 was found on all hepatocytes (epithelial cells, green)(I), whereas anti-L-SIGN antibodies stained sinusoidal endothelial cells (red) in close contact with hepatocytes, and CD31 (green) expression was found mainly on EC around blood vessels (J) (original magnification: A, B: x20; C, D, E: x40; F, G: x10; H, I, J: x20).

View larger version (123K): [in this window] [in a new window]   Figure 6. Single-color immunohistochemical staining of normal human nasal biopsies with antibodies to various endothelial markers. Nasal biopsy sections showing positive staining (brown) for (A) CD144, (B) CD31, (C) L-SIGN (note the brown staining of endothelial cells in close contact with epithelial cells), (D) CD141, (E) von Willebrand Factor, and (F) epithelial cell adhesion molecule. Liver biopsy sections showing positive staining for (G) hepatocytes expressing CK18 (red) and sinusoidal endothelial staining with L-SIGN (black), (H) CK18 (red) but not CD31, and (I) CD31 (black) in blood vessels. All sections were counterstained with hematoxylin (original magnification: x20).

	DISCUSSION

Top Abstract Introduction MATERIALS AND METHODS Results DISCUSSION References

Morphologically, phenotypically, and molecularly, there existed differences between the two HNMEC populations. One population showed classical vascular endothelial characteristics such as cobblestone morphology and expression of well known, EC-specific surface molecules. The other population had sinusoidal characteristics, had a fusiform cell shape, lacked expression of EC-specific cell surface molecules CD31 and CD62E, had no tight junctions, showed presence of fenestrae, and was discontinuous.

SEC in the liver lack surface expression of CD31 and CD62E (10, 12). The reason for this is not clear. Previous studies on specialized EC types, such as high endothelial venules in lymph nodes, suggest that different properties are induced by the local milieu and can be modified depending on stimuli (4). Thus, it is reasonable to assume that the described regional expression patterns are influenced by the local micromilieu. The lack of CD31 and CD62E expression on nasal SEC may indicate the presence of specific local suppressive factor or that sinusoidal lining cells may inherently lack the ability to respond to inducers of these molecules. Keeping in mind that the nasal sinusoid is a unique, specialized unit different from the microvasculature of the other parts of the respiratory tract, the described specific constellation of adhesion ligand molecules may reflect a functional requirement. On the other hand, the lack of expression of these adhesion molecules important for leukocyte–EC interactions may have a major effect on the mode of leukocyte adherence and cell interaction at this anatomic site.

A deficiency or lack of these molecules might indicate an increased risk of recurrent infections (13, 14). It is not known how the immune cells interact with nasal sinusoids and the consequences of these interactions in inflammation and in various nasal disorders.

To further provide novel insights into the molecular differences, the gene expression patterns between the vascular endothelial cells (VEC) and nasal SECs were analyzed. Due to the large amount of data generated from such an analysis, we restricted our analysis to the expression of genes considered to be involved in human EC biology. Of the reported genes, the array generated a similar transcriptional profile between the two cell types, except for higher expression of five genes in the nasal SEC. These genes encode for MMP-1, osteoprotegerin, MCP-1, collagen type 1, and CD106/VCAM-1.

The clinical or physiologic implications of the differences in this gene expression pattern between the two cell types are not known and need further investigation. The reported changes in expression represent one individual at one single time point, and it is possible that the gene expression pattern might alter between individuals.

Morphologic and phenotypical heterogeneity has been shown in other vascular beds. One example is the liver, where EC display vascular and sinusoidal morphology. A similar lack of surface expression of CD31 and CD62E and a lack of tight junctions is also evident in the SEC of the liver (13–15). Liver sinusoids are regarded as capillaries that differ from other capillaries in the body because of the presence of open pores or fenestrae. The unique arrangement of the normal sinusoidal endothelium is likely to facilitate the large exchanges that take place between hepatocytes and the blood (13–15).

The clinical importance of a structural integrity of the fenestrated liver SEC is well recognized in several diseases where capillarization and loss of fenestrae result in diseases such as atherosclerosis, cirrhosis, fibrosis, hyperlipoproteinemia, and cancer (16, 17). We have recently shown that autoantibodies to HLSEC can transform SEC to vascular type and that this transformation may play an important role in the development of hepatocellular failure and portal hypertension in patients with primary biliary cirrhosis and autoimmune hepatitis (11). Furthermore, evidence indicates that SEC of the liver can act as antigen-presenting cells (APC) and may play a central role in immune responses (18, 19).

As in the liver, the microvasculature of the nose presents a similar filtering effect whereby a dense subepithelial network of capillaries with fenestrations between the EC allows water to escape into the airway lumen and allows evaporation to take place, enabling conditioning of the inspired air (7). It would be interesting to study whether capillarization of the nasal sinusoids occurs in various nasal disorders and to evaluate whether nasal sinusoids EC may act as APC to initiate local specific immune responses in patients with nasal disorders.

Dissimilarities can be found between the SEC of the liver and the nose, indicating that heterogeneity in biochemical, phenotypic, and antigenic expression dictated by microenvironment of the organ exists. This is clearly demonstrated in our recent study (20), where we reported that a significantly high fraction of patients with Wegener's granulomatosis had antibodies that recognize and bind to molecules expressed on nasal VEC but not liver SEC (P < 0.0001). Our unpublished results demonstrate that patients with Wegener's granulomatosis have autoantibodies that bind to nasal vascular EC and SEC, indicating that the antibodies may recognize an antigen commonly expressed on both cell types.

This finding emphasizes the importance of identifying the role of such tissue/organ-specific antigens in diseases. In addition, we found that inflammatory cytokines such as TNF- and IFN- were cytotoxic to nasal SEC but not to liver SEC (P < 0.0001). These findings further justify the importance of using relevant target EC in studies of site-specific lesions.

The existence of VEC and SEC in the nose was confirmed by in vivo staining of nasal biopsy sections. Two distinct staining patterns were observed whereby SEC were found mainly in the vicinity of epithelial cells, whereas VEC were found in blood vessels—a finding similar to that observed in the liver (21). We also found that nasal SEC expressed the lymph node/liver specific marker L-SIGN in vivo and in vitro. This finding implies that one may use antibodies to L-SIGN to isolate nasal SEC and anti-CD31 antibodies for VEC.

In conclusion, we describe a simple and novel protocol for the isolation and cultivation of two heterogeneous populations of HNMEC that will be of great advantage for in vitro research in illnesses such as asthma, non-allergic inflammation, hyper-reactivity, allergic rhinitis (22, 23), vasculitides such as Wegener's granulomatosis (24), and other nasal diseases. The use of these two distinct, well-characterized populations of HNMEC as targets in such studies may provide important clinical knowledge regarding the role of each EC type to respond during inflammation and various types of damage in nasal disorders.

	Acknowledgments

 
The authors thank Professor Erna Möller for financial support, Silvia Nava for technical help with immunohistochemistry and immunocytochemistry stainings, Dr. Kjell Hultenby for help with electron microscopy, and Dr. Makiko Kumagi-Braesch for help with the fluorescence microscopy.

	Footnotes

 
This study was financed by grant no. 00793 to E.M. and grant no. K2002-06X-14004-02B to S.S.-H. from the Medical Research Council; the Konung Gustaf V's 80 Year, the Reumatikerförbundet, and the Sunds Foundations.

Conflict of Interest Statement: C.H. has no declared conflicts of interest; P.S. has no declared conflicts of interest; and S.S-H. has no declared conflicts of interest.

Received in original form August 9, 2004

Received in final form October 10, 2004

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