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The Journal of Immunology, 2000, 165: 5428-5434.
Copyright © 2000 by The American Association of Immunologists

Dermal Microvascular Endothelial Cells Express the 180-kDa Macrophage Mannose Receptor In Situ and In Vitro1

Marion Gröger*, Wolfgang Holnthoner*, Dieter Maurer{dagger}, Sonja Lechleitner*, Klaus Wolff*, Bettina Beate Mayr{ddagger}, Werner Lubitz{ddagger} and Peter Petzelbauer2,*

Department of Dermatology, Divisions of * General Dermatology and {dagger} Immunology, Allergy, and Infectious Diseases, and {ddagger} Institute for Microbiology and Genetics, University of Vienna, Vienna, Austria


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Expression of the 180-kDa mannose receptor (MR) is mainly found on cells of the macrophage lineage. MR mediates the uptake of micro-organisms and host-derived glycoproteins. We demonstrate that endothelium of the human skin in situ and dermal microvascular endothelial cells (DMEC) in vitro expressed MR at both the protein and mRNA levels. In contrast, HUVEC were consistently negative for MR expression. DMEC internalized dextran as well as Escherichia coli by the way of MR into acidic phagosomes, only a few of which fused with CD63- and lysosomal-associated membrane glycoprotein-2-positive lysosomes. This contrasts with the situation in monocyte-derived dendritic cells, where almost all of the MR-Ag complexes reached CD63- and lysosomal-associated membrane glycoprotein-2-positive compartments, indicating differences in the phagolysosomal fusion rate between DMEC and dendritic cells. In conclusion, DMEC express functional MR, a finding that corroborates a role of skin endothelium in Ag capture/clearing.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The 180-kDa mannose receptor (MR)3 is a prototype member of a family of multilectin receptors that function as pattern recognition receptors and effectuate innate immune responses. The eight membrane-proximal carbohydrate recognition domains (CRDs) of MR bind mannose-, N-acetylglucosamine-, and fucose-terminating oligosaccharides, which are commonly found on the cell wall of bacteria, yeast, and parasites, but are rarely seen in sufficient densities in terminal positions of mammalian oligosaccharides (1). For the NH2-terminal cystein-rich domain of MR (Cys-MR), only endogenous ligands have described to date, which terminate in 4-sulfated N-acetylgalactoseamine and include sulfated carbohydrates on pituitary hormones, chondroitin sulfate, and sulfated blood group chains (2, 3). Whereas the role of Cys-MR ligand binding in innate immunity is not yet well defined, the fate of CRD ligands is well established in macrophages and dendritic cells. In macrophages, CRD ligation may lead to MR recycling to and from phagosomes (4). Alternatively, internalized MR-Ag complexes are found in acidic compartments fused with CD63+ (lysosomal-associated membrane glycoprotein-3)- and CD107b+ (lysosomal-associated membrane glycoprotein-2) lysosomes, where the Ags are degraded (1, 5, 6, 7, 8). In dendritic cells, MR can deliver Ags into compartments for MHC class II loading (8), but certain Ags are also transported into late endosomes for loading onto the MHC class I-like molecule CD1b (9). Thus, in dendritic cells this MR-mediated pathway links recognition of microbial Ags to the induction of adaptive T cell responses.

Apart from macrophages and subtypes of dendritic cells, MR expression has been described on kidney mesangium, tracheal smooth muscle, and retinal pigment epithelium (10, 11, 12, 13). Interestingly, in endothelial cells, MR expression appears to be restricted to certain vascular beds, such as the sinus-lining cells of the liver, spleen, and lymph nodes (10, 14, 15, 16). Sinusoidal liver endothelial cells are unique in several ways. They express another scavenger receptor, CD32 (Fc{gamma}RIIa), and their role in Ag capture and clearing is well characterized (17, 18). They constitutively express MHC class II molecules, which suggests that they are involved not only in Ag uptake but also in Ag presentation (19, 20). Human dermal microvascular endothelial cells (DMEC) also constitutively express CD32 and MHC class II molecules and thus appear to share some of the properties of sinusoidal liver endothelium in playing a role in Ag capture/clearing and presentation (21, 22, 23, 24). We therefore analyzed DMEC for MR expression in vivo and in vitro.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Abs and reagents

Anti-mannose receptor (anti-MR) mAbs were clone 19 (PharMingen, Uppsala, Sweden) and clone PAM-1 (16). CD36-FITC and anti-HLA-DR (clone L243) were obtained from Becton Dickinson (San Jose, CA). CD31 (clone 7E4) was a gift from Dr. Otto Majdic (Institute of Immunology, University of Vienna Medical School, Vienna, Austria). CD63 was purchased from Immunotech (Marseilles, France), and CD107b from PharMingen. Rabbit anti-human von Willebrand factor was obtained from Dako (Glostrup, Denmark). FITC- and tetramethylrhodamine isothiocyanate (TRITC)-labeled second-step Abs were purchased from Jackson (West Grove, PA). Isotype controls were obtained from PharMingen. FITC-, Oregon Green-, and TRITC-labeled dextran (Mr, 70.000) and Lucifer Yellow CH potassium salt were obtained from Molecular Probes (Leiden, The Netherlands). Dextrans were spun in a microfuge to remove aggregates. Mannan from Saccharomyces cerevisiae was obtained from Sigma (St. Louis, MO). Recombinant human IL-4, IL-10, IL-13, and IFN-{gamma} were obtained from Stratagene (La Jolla, CA), and TNF was obtained from Sigma. Nondenatured bacterial cell envelopes (ghosts) were created from Escherichia coli K12 strain pop2135 (provided by O. Raibaud, Institute Pasteur, Paris, France) as described (25, 26) and labeled with Alexa488 (Molecular Probes) according to the manufacturer’s instruction. Staphylococcus aureus BioParticles-FITC were obtained from Molecular Probes.

Cells

DMEC were isolated from human foreskins as previously described (27). In addition, DMEC were purchased from Promo Cell (Heidelberg, Germany). DMEC were grown in Endothelial Cell Growth Medium MV (Promo Cell) on fibronectin-coated dishes (10 µg/ml; Life Technologies, Gaithersburg, MD). Cells were used between passages 1 through 5. DMEC uniformly expressed VE-cadherin, CD31, and CD34, and following 4-h TNF stimulation the entire cell population expressed CD62E, confirming their origin from blood vessel endothelium (data not shown). HUVEC were isolated and subcultured as previously described (27) and used between passages 1 through 5. Human monocyte-derived dendritic cells were generated as previously described (8). Briefly, monocytes were cultured in RPMI 1640 culture medium (Life Technologies) supplemented with GM-CSF (800 U/ml; Novartis, Basel, Switzerland), IL-4 (1000 U/ml; Stratagene), and 10% FCS for 10 days.

Immunofluorescence of human skin

Five-micron cryostat sections were prepared from snap-frozen normal skin and from skin with metastatic melanoma and processed as described previously (21). First-step reagents were mouse anti-MR mAb (2.5 µg/ml) and rabbit anti-von Willebrand factor serum (1/400) diluted in 1% PBS/BSA; second-step reagents were TRITC-labeled goat anti-mouse (2.5 µg/ml) and TRITC-labeled goat anti-rabbit (2.5 µg/ml) Abs. Specimens were examined by a confocal laser scan microscope (LSM 410, Zeiss, Oberkochen, Germany).

Quantification of mannose receptor surface expression and quantification of dextran uptake by FACS analysis

DMEC and HUVEC were suspended in trypsin/EDTA (Life Technologies) and washed in PBS. To analyze surface Ag expression, cells were incubated with PE-labeled anti-MR, PE-labeled CD31 and FITC-labeled CD36 (1 µg/ml each) in PBS for 30 min on 4°C. Isotype-matched Abs were used as a negative control. Surface-bound fluorescence was analyzed by FACScan (Becton Dickinson, San Jose, CA).

To quantify dextran uptake, DMEC or HUVEC were washed in PBS/1% FCS twice, followed by incubation with 1 mg/ml Oregon Green-labeled dextran in PBS/1% FCS for the indicated time points at 37°C and, as a negative control, at 4°C. Fluorescence emission of Oregon Green is pH insensitive. Dextran uptake was blocked by preincubation of cells with mannan (2 mg/ml) or anti-MR mAbs (2 µg/ml) for 20 min followed by incubation with 1 mg/ml Oregon Green-labeled dextran in the continuous presence of the respective blocking reagent. Cells were then washed five times with cold PBS/1% FCS and analyzed by FACScan (Becton Dickinson). MR-dependent dextran uptake was calculated by two methods. First, the geometric mean fluorescence of Oregon Green dextran-positive cells minus the geometric mean fluorescence of Oregon Green dextran-positive cells pretreated with mannan. Second, the geometric mean fluorescence of Oregon green dextran-positive cells minus the geometric mean fluorescence of Oregon Green dextran-positive cells pretreated with anti-MR mAb. The mean ± SEM of five independent experiments are given. Lucifer Yellow (1 mg/ml), with or without preincubation with mannan, was used as a control for fluid phase uptake.

Western blot

DMEC and HUVEC were lysed in Tris lysis buffer, loaded onto a 7% polyacrylamide gel, electrophoresed, and blotted as described previously (21, 28). After blocking with 1% low fat milk (Bio-Rad, Hercules, CA) for 12 h, membranes were incubated with anti-MR mAb, CD31, or an isotype control Ab (1 µg/ml each) diluted in 0.5% Tween in TBS for 1 h. For detection, an HRP-labeled goat anti-mouse Ab (1/50,000; Bio-Rad) in 0.5% Tween/TBS was used, and bound Abs were visualized by chemiluminescence (ECL system; Amersham, Arlington Heights, IL) and recorded on film.

RT-PCR

Total RNA was isolated and reverse transcribed as previously described (29). Primer sequences for the MR amplify cDNA only and were described by Lu et al. (30). They amplify a 400-bp product. Thirty PCR cycles were performed under the following conditions: 94°C for 30 s, 53°C for 30 s, 72°C for 30 s, and a final extension at 72°C for 7 min. GAPDH primers were obtained from Clontech (Palo Alto, CA) and run for 25 cycles: 94°C for 5 min, 94°C for 30 s, 60°C for 30 s, and 72°C for 1 min. The identities of the respective PCR products were identified by their expected sizes. RNA without RT was used as a control.

Confocal laser scan microscopy

DMEC and HUVEC cultured to confluence on chamber slides (Nunc, Roskilde, DK) were incubated with equal amounts of FITC- and TRITC-labeled dextran (1 mg/ml each) for the indicated times and at the indicated temperatures. Cells were then fixed in 3% paraformaldehyde/PBS for 15 min on ice followed by incubation with 50 mM NH4Cl for 10 min on ice. FITC and TRITC emissions were analyzed by laser scan microscopy with standardized laser brightness, and contrast with the scanning level set through the center of the cell.

To analyze the antigenic phenotype of the subcellular organelles, cell suspensions of DMEC or monocyte-derived dendritic cells were pulsed with TRITC-dextran as described above or alternatively with Alexa488-labeled E. coli cell envelopes or Staphylococcus aureus BioParticles. After fixation with paraformaldehyde as described above, permeabilization with 0.01% saponin/PBS/1% FCS for 2 min at room temperature, cells were incubated with first-step mAbs for 20 min on ice, followed by a FITC- or TRITC-labeled goat anti-mouse second-step Ab, and were examined by confocal laser scan microscopy.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
DMEC express MR

In normal skin specimens (n = 4), MR expression was found on ~30–50% of small and medium-sized blood vessels of the deep vascular plexus of the dermis (Fig. 1GoA). Vessels forming the papillary loops within the tips of the rete ridges beneath the epidermis were negative for MR, but were surrounded by MR-positive macrophages (Fig. 1GoB). All MR-positive vessels coexpressed CD36 molecules (data not shown). To analyze MR expression under conditions of pathologic neovascularization, skin samples with metastatic melanoma (n = 3) were analyzed; 60–80% of the small tumor vessels expressed MR (Fig. 1GoC). Stainings with isotype-matched control mAb were negative (data not shown).



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  		FIGURE 1. Immunohistochemistry on cryostat sections of normal skin (A and B). The deep vascular plexus of the dermis coursing around adnexal structures is shown in A, the capillary loops within the tips of the rete just below the epidermis in B (dermo-epidermal junction is marked by the dotted line), and a cutaneous metastasis of a melanoma is shown in C. Sections were double-stained with anti-MR mAbs (three left panels) and with anti-von Willebrand factor (anti-vWF; three middle panels). The overlays are shown in the three right panels. Endothelial cells within the deep dermis of normal skin (A) or endothelial cells within metastatic cutaneous melanoma (C) stain yellow due to the overlay of red (anti-MR) and green (anti-vWF). B, Vessels within the rete are negative for MR and thus appear green only (anti-vWF). Scale, 0.1 mm.

 
We next analyzed DMEC in culture (n = 8) and, for comparison, HUVEC by flow cytometry. DMEC at passage 1 expressed MR (ranging from 50 to 95% of cells) as determined by FACS analysis (Fig. 2GoA). Upon further subculture, MR surface expression decreased and was virtually absent at passage 5. MR expression could not be maintained or reinduced with cytokines such as IL-4, IL-10, IL-13, IFN, and TNF, which are known to enhance MR expression on macrophages and sinusoidal liver endothelium (20, 31, 32) (data not shown). The correct m.w. of MR expressed by DMEC was confirmed by Western blotting (Fig. 2GoB). mRNA expression in DMEC was analyzed by RT-PCR and was detectable at passages 1 through 3 (Fig. 2GoC).



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  		FIGURE 2. A, FACS analysis of DMEC and HUVEC at passage 3. B, Immunoblots of HUVEC (lane 1), DMEC (lane 2), and monocyte-derived dendritic cells (lane 3). The 180-kDa MR protein is absent in HUVEC (lane 1) and is strongly expressed in DMEC (lane 2) and monocyte-derived dendritic cells (lane 3). As a positive control, immunoblots using CD31 mAbs are shown in the middle panel. It should be noted that after longer exposure CD31 proteins are also detected in lane 3. As a negative control, immunoblots using an isotype control mAb are shown in the right panel. C, RT-PCR for the MR mRNA expression. An amplification product of 400 bp is detectable after RT into cDNA in DMEC, but not HUVEC. The RNA controls without RT are negative. mRNA for GAPDH is found in both DMEC and HUVEC.

 
HUVEC, analyzed for comparison, did not express MR protein or mRNA at early or late passages (Fig. 2Go) or upon stimulation with any of the above mentioned cytokines (data not shown).

DMEC internalize the occupied MR

To test the functionality of endothelial MR, the uptake of Oregon Green-labeled dextran was measured in the presence or the absence of mannan or in the presence or the absence of anti-MR mAbs. Fluorescence uptake was quantified by FACS analysis (Fig. 3Go). This technique has been successfully employed, e.g., to characterize MR function in dendritic cells (8).



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  		FIGURE 3. FACS analysis of DMEC (A) or HUVEC (B), which were incubated with Oregon Green-labeled dextran in the absence (bold black lines) or the presence of mannan (thin lines) at the indicated times and the indicated temperatures. Mannan reduces dextran uptake in DMEC, but not in HUVEC. At 4°C neither DMEC nor HUVEC internalize dextran. As a control for fluid phase uptake, Lucifer yellow is shown, which enters into DMEC in the presence or the absence of mannan. C, Calculated MR-dependent dextran uptake. The left panel shows geometric mean fluorescence of Oregon Green dextran-positive cells minus geometric mean fluorescence of Oregon Green dextran-positive cells pretreated with mannan. The right panel shows geometric mean fluorescence of Oregon Green dextran-positive cells minus geometric mean fluorescence of Oregon Green dextran-positive cells pretreated with anti-MR mAb. Both calculations revealed a more or less equal increase in MR-dependent dextran uptake in DMEC (difference not significant). In contrast, in HUVEC it remained virtually zero. Each curve represents the mean of five independent experiments.

 
Dextran uptake was detectable as early as 5 min after the dextran pulse and continuously increased thereafter (Fig. 3Go). The MR-dependent dextran uptake was determined by two methods. First, by subtracting the dextran uptake in the presence of mannan from the total dextran uptake (left panel) and, second, by subtracting the dextran uptake in the presence of anti-MR mAbs from the total dextran uptake (right panel). Both calculations revealed a more or less equal increase in MR-dependent dextran uptake in DMEC, whereas in HUVEC it remained virtually zero (Fig. 3GoC). It should be noted that in DMEC the MR-independent dextran uptake (dextran uptake in the presence of the respective blocking reagent) accounted for <30% of the total dextran uptake (data not shown). Moreover, this MR-independent dextran uptake was comparable in HUVEC and DMEC (Fig. 3Go). As controls, cells were incubated with dextran at 4°C, revealing no dextran uptake in DMEC or in HUVEC. As an additional control, cells were pulsed with Lucifer Yellow, which is internalized by fluid phase uptake (8). As expected, Lucifer Yellow uptake was not blocked by mannan (Fig. 3GoA).

To confirm that dextran was indeed internalized and not just fixed to the cell membrane, cells were analyzed by laser scan microscopy, which clearly showed the fluorescence located within the cytoplasm (Fig. 4Go). Moreover, these experiments allowed determination of the pH of dextran-positive organelles by taking advantage of the fact that fluorescence emission of FITC is significantly reduced at pH 6.0 and is absent at pH <5.0, whereas fluorescence emission of TRITC remains stable even at pH <5 (33, 34). DMEC incubated with equal amounts of FITC- and TRITC-dextran showed numerous FITC- and TRITC-positive intracellular organelles following a 1-min dextran pulse, indicating that the pH in these vesicles was >6 (Fig. 4Go). After 10 min, FITC fluorescence was reduced (not shown) and had disappeared almost completely after 60 min (Fig. 4Go). In contrast, TRITC-fluorescence persisted within the cytoplasm of the cell, which allowed the conclusion that dextran had entered acidic compartments (Fig. 4Go).



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  		FIGURE 4. Laser scan images of DMEC grown on tissue culture plastic and pulsed with equal amounts of TRITC-labeled (images on the left) and FITC-labeled (images in the middle) dextran. Following a 1-min incubation, both TRITC- and FITC-labeled dextran entered the cell. The upper right image gives the overlay; equal amounts of green and red fluorescence colocalize in cytoplasmic organelles, which therefore appear yellow. In contrast, following 60 min of coincubation, the cytoplasm of DMEC is filled with TRITC fluorescence, whereas the pH-sensitive fluorescence of FITC is not detectable anymore; the overlay appears thus red only (lower right image).

 
Phenotype of organelles targeted by the MR

DMEC were pulsed for 30 min with TRITC-labeled dextran (pH insensitive) and chased for up to 24 h. The subsequent immunolabeling revealed most of the dextran-positive vesicles as being negative for CD63 and CD107b (Fig. 5Go, A and C), indicating that the majority of dextran-positive phagosomes did not fuse with lysosomes. Representative examples following a 30-min chase of dextran are shown in Fig. 5GoA, but it should be noted that the phenotype of dextran-positive organelles in DMEC did not change even after a chase period of up to 24 h. These compartments were also negative for HLA class I Ags (data not shown). Because DMEC in culture (in contrast to the in vivo situation) do not constitutively express MHC class II molecules, we analyzed DMEC pretreated with IFN-{gamma} for 48 h. Only 20–30% of dextran-positive compartments reacted with anti-HLA-DR mAbs (Fig. 5GoA, right panel, and Fig. 5GoC). To determine whether this inefficient phagolysosomal fusion rate is a special feature of DMEC, we investigated the dextran uptake by monocyte-derived dendritic cells under the same experimental set-up. Following a 30-min chase period, 70–80% of dextran-loaded vesicles were positive for CD63, CD107b, as well as HLA-DR molecules (Fig. 5Go, B and C).



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  		FIGURE 5. Laser scan images of cells in suspension pulsed with TRITC-labeled dextran (pH insensitive) for 30 min at 37°C, fixed, permeabilized, and stained with the indicated Abs and a FITC-labeled second-step Ab. A, In DMEC, only a few of the dextran-positive organelles react with CD63 or CD107b mAbs, which is indicative of lysosomal fusion (lysosomes appear yellow due to the overlay of red and green fluorescence; cell borders are marked by the dotted line). In the right upper panel, IFN-treated DMEC are shown. Only a small number of dextran-positive compartments react with anti-HLA-DR mAbs. B, In contrast, in monocyte-derived dendritic cells almost all the internalized dextran colocalizes with CD63, CD107b, and HLA-DR molecules. C, Absolute numbers of dextran-positive vesicles per cell as well as numbers of dextran-positive vesicles reactive with the indicated mAbs were counted in three independent experiments, and the mean ± SEM values are given.

 
Finally, we analyzed whether bacteria can serve as natural ligands for endothelial MR. DMEC were pulsed with E. coli cell envelopes (ghosts). After 1 h E. coli uptake was seen in 78% of cells, and this E. coli uptake was sensitive to preincubation with dextran; only 26% of cells internalized E. coli in the presence of dextran (p < 0.05; example shown in Fig. 6Go, A and B). As a control, we used Staphylococcus aureus particles, which enter the cell in the absence as well as the presence of dextran (Fig. 6Go, C and D). As seen with dextran, DMEC internalized E. coli mainly into CD63- and CD107b-negative compartments (Fig. 6Go, E and F).



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  		FIGURE 6. Laser scan images of DMEC pulsed with Alexa488-labeled E. coli ghosts for 60 min at 37°C in the absence (A) or the presence (B) of dextran-TRITC. As a control, DMEC were incubated with S. aureus BioParticles in the absence (C) or the presence (D) of dextran-TRITC. E. coli, but not S. aureus, uptake was blocked by preincubation with TRITC-labeled dextran. E and F, Immunofluorescence stainings of DMEC after a 60-min pulse with E. coli ghosts. As seen with dextran in Fig. 5Go, only a few of the E. coli ghosts colocalized with CD63 (E) and CD107b (F) molecules.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Here we demonstrate that endothelial cells of the skin express macrophage MR. Ligands bound to endothelial MR were rapidly internalized, with kinetics roughly comparable to those seen in monocyte-derived dendritic cells (8). Endothelial MR delivered dextran or E. coli mainly into CD63- and CD107b-negative phagosomes, which were rapidly acidified. This contrasts the situation in monocyte-derived dendritic cells or macrophages, where most phagosomes fused with lysosomes (7, 8) (see also Fig. 5GoB), indicating that phagolysosomal fusion rates were low in DMEC. Mechanisms promoting or attenuating phagolysosomal fusion are as yet ill defined. Most of the knowledge comes from the observation that certain bacterial strains reduce or prevent phagolysosomal fusion, which is thought to be part of their survival strategy (6, 35, 36, 37, 38, 39). Based on these studies, phagolysosomal fusion appears to be Ca2+ dependent, involves the redistribution of small Ras-like GTP binding proteins rab5 and rab7, involves the vesicular proton-ATPase that is responsible for phagosomal acidification and is regulated by proinflammatory and anti-inflammatory cytokines (35, 36, 37, 40). We currently have no explanation for the low phagolysosomal fusion deficit seen in DMEC. It should be noted that phagolysosomal fusion could not be enhanced by pretreating DMEC with TNF or IFN or by prolonging chase periods for up 24 h (data not shown). The latter was not unexpected, because phagosomes turn into a low fusion state 4–13 h after internalization of a target (41). It has thus to be assumed that as yet undefined cell-specific differences between DMEC and dendritic cells account for the observed different phagolysosomal fusion rates.

We next analyzed whether internalized dextran or bacteria colocalized with MHC class II-positive organelles. Because DMEC in cell culture do not express HLA-DR, which contrasts with the in vivo situation, we pretreated DMEC with IFN-{gamma} to induce HLA-DR expression (23, 42). Only a small fraction of dextran-positive organelles reacted with anti-HLA-DR mAbs, whereas in monocyte-derived dendritic cells most of the MR-Ag complexes were found in MHC class II-positive endosomes (8) (see also Fig. 5GoB). This correlated well with the low phagolysosomal fusion rate in DMEC discussed above. Unfortunately, we were unable to directly analyze whether this observed low delivery rate of Ags into MHC class II compartments is sufficient to cause Ag-specific response of sensitized T cells, because T cells syngeneic to our neonatal foreskin-derived DMEC were not available. Moreover, due to the rather rapid loss of MR expression of DMEC during cell culture, sufficient cell numbers were not available to allow the creation of MHC-matched T cell lines. The loss of endothelial MR expression was due to the loss of mRNA transcription at passage 3 as well as to the shedding of MR protein into the culture medium (data not shown).

MR is a pattern recognition receptor expressed mainly on cells of the myeloid lineage, with only a few exceptions. Apart from some specialized cell types in the kidney, trachea, and retina (10, 11, 12, 13), MR has to date only been described on endothelial cells of organs, which are specialized in Ag uptake/clearing and/or presentation such as the liver, spleen, and lymph nodes (10, 14, 15, 16). Here we show that endothelial cells of a peripheral organ, the skin, also express MR. DMEC are unique in several ways. In addition to pattern recognition receptors broadly expressed on most endothelium, e.g., proteins of the Toll-like receptor family (43) or receptors of the scavenger cell pathway of low density lipoprotein metabolism (44), DMEC express other scavenger receptors, which have a very restricted expression pattern. For example, DMEC express CD36 (27), which is otherwise found on endothelium of brain and heart only (45, 46), and DMEC express CD32 (21), which is otherwise found on endothelium of liver only (17). This combined expression of multiple pattern recognition receptors is to date unique for DMEC and raises important questions about their roles in innate immunity. Specifically, what is the fate of the MR-Ag complexes internalized into endothelial phagosomes that are not fused with lysosomes? It has been shown previously that Ags from nonfused phagosomes can return to the cell membrane (47). Following the concept for skin-associated lymphoid tissues (48), it is tempting to speculate that Ags modified or degraded within the endothelial acidic phagosomes are recycled and then delivered into the cutaneous environment. The skin is filled with dendritic cells, which mature upon uptake of bacterial Ags and give rise to systemic anti-microbial immunity (49).

Finally, it should be noted that MR expression is mainly expressed at the luminal surface of endothelium. Because circulating hemopoietic cells do not express MR (50), endothelial cells are thus the only cell type that have MR exposed to the bloodstream. Endothelial MR could therefore be used for MR-dependent gene transfer, which was shown to function for tissue macrophages (50) and with bacterial cell envelopes being a potential carrier (25).


   Footnotes
 
1 This work was supported by grants from the Austrian Science Foundation (P12240-MED), the Niarchos Foundation, and from the ICP Program. Back

2 Address correspondence and reprint requests to Dr. Peter Petzelbauer, Department of Dermatology, Division of General Dermatology, University of Vienna Medical School, Waehringer Guertel 18-20, A-1090 Vienna, Austria. Back

3 Abbreviations used in this paper: MR, mannose receptor; CRD, carbohydrate recognition domain; DMEC, dermal microvascular endothelial cells; TRITC, tetramethylrhodamine isothiocyanate. Back

Received for publication June 12, 2000. Accepted for publication August 15, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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