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Microdomains of the C-type lectin DC-SIGN are portals for virus entry into dendritic cells

Microdomains of the C-type lectin DC-SIGN are portals for virus entry into dendritic cells JCB Article Microdomains of the C-type lectin DC-SIGN are portals for virus entry into dendritic cells 1 1 4 4 1 Alessandra Cambi, Frank de Lange, Noortje M. van Maarseveen, Monique Nijhuis, Ben Joosten, 5 5 2 3 Erik M.H.P. van Dijk, Bärbel I. de Bakker, Jack A.M. Fransen, Petra H.M. Bovee-Geurts, 1 5 1 Frank N. van Leeuwen, Niek F. Van Hulst, and Carl G. Figdor 1 2 3 Department of Tumor Immunology, Department of Cell Biology, and Department of Medical Biochemistry, Nijmegen Center for Molecular Life Sciences, University Medical Center Nijmegen, 6500 HB Nijmegen, Netherlands Department of Virology, University Medical Center Utrecht, 3508 GA Utrecht, Netherlands Applied Optics Group, Department of Science and Technology, Molecular Engineering Sensors and Actuators Research Institute for Nanotechnology, University of Twente, 7500 AE Enschede, Netherlands he C-type lectin dendritic cell (DC)–specific inter- becomes organized in well-defined microdomains, with an cellular adhesion molecule grabbing non-integrin average diameter of 200 nm. Biochemical experiments (DC-SIGN; CD209) facilitates binding and internal- and confocal microscopy indicate that DC-SIGN micro- ization of several viruses, including HIV-1, on DCs, but the domains reside within lipid rafts. Finally, we show that the underlying mechanism for being such an efficient phagocytic organization of DC-SIGN in microdomains on the plasma pathogen-recognition receptor is poorly understood. By high membrane is important for binding and internalization resolution electron microscopy, we demonstrate a direct of virus particles, suggesting that these multimolecular relation between DC-SIGN function as viral receptor and assemblies of DC-SIGN act as a docking site for pathogens its microlocalization on the plasma membrane. During like HIV-1 to invade the host. development of human monocyte-derived DCs, DC-SIGN Introduction In the past two years, several new genes have been identified and Steinman, 1998). DCs are equipped with a variety of encoding leukocyte-specific carbohydrate binding proteins dynamically regulated pathogen-recognition receptors. Al- that belong to the lectin-like receptors family (Kogelberg though some of them are members of the toll-like receptor and Feizi, 2001; Figdor et al., 2002). Many of these lectins (TLR) family, signaling molecules specialized in sensing are members of the calcium-dependent C-type lectin family pathogens (Akira, 2003), others belong to the C-type lectin and recognize their ligands through the structurally related family and mediate pathogen binding and uptake (Stahl and Ca -dependent carbohydrate-recognition domains (C-type Ezekowitz, 1998; Mahnke et al., 2000). CRDs; Drickamer, 1999). Many C-type lectins act as cell DC-specific intercellular adhesion molecule (ICAM) grab- adhesion receptors (Vestweber and Blanks, 1999), whereas bing non-integrin (DC-SIGN; CD209) is a C-type lectin others are specialized in antigen recognition (Stahl and specifically expressed by DCs and has a dual function. As an Ezekowitz, 1998; Mahnke et al., 2000). adhesion receptor, DC-SIGN supports initial DC–T cell in- Dendritic cells (DCs) constitute a specific group of pro- teraction by binding to ICAM-3 (Geijtenbeek et al., 2000a), fessional antigen presenting leukocytes, constantly patrolling and mediates tethering and rolling of DCs on the endo- the body for foreign intruders (Steinman 1991; Banchereau thelium by interacting with ICAM-2 (Geijtenbeek et al., 2000c). As a pathogen-recognition receptor, DC-SIGN The online version of this article contains supplemental material. binds HIV gp120 thus facilitating the transport of HIV Address correspondence to Carl G. Figdor, Dept. of Tumor Immunology, from mucosal sites to draining lymph nodes where infection Nijmegen Center for Molecular Life Sciences, University Medical Center of T lymphocytes occurs (Geijtenbeek et al., 2000b). Recently, Nijmegen, P.O. Box 9101, 6500 HB Nijmegen, Netherlands. Tel.: 31- DC-SIGN was also shown to bind other viruses like CMV 24-361-7600. Fax: 31-24-354-0339. (Halary et al., 2002), Ebola (Alvarez et al., 2002), Dengue email: c.figdor@ncmls.kun.nl (Tassaneetrithep et al., 2003), and hepatitis C (Lozach et al., Key words: pathogen recognition receptor; lectin; electron microscopy; multiprotein assembly; lipid rafts 2003; Pöhlmann et al., 2003), as well as microorganisms  The Rockefeller University Press, 0021-9525/2004/01/145/11 $8.00 The Journal of Cell Biology, Volume 164, Number 1, January 5, 2004 145–155 http://www.jcb.org/cgi/doi/10.1083/jcb.200306112 145 The Journal of Cell Biology 146 The Journal of Cell Biology | Volume 164, Number 1, 2004 such as Leishmania (Colmenares et al., 2002), Candida albi- cans (Cambi et al., 2003), Mycobacterium (Geijtenbeek et al., 2003; Maeda et al., 2003; Tailleux et al., 2003), and Schisto- soma (van Die et al., 2003). Recent works have demonstrated that some microbial pathogens exploit cholesterol-enriched lipid microdomains as essential docking sites to enter host cells (Gatfield and Pieters, 2000; Rosenberger et al., 2000; Lafont et al., 2002). These microdomains, also known as lipid rafts, are localized regions with elevated cholesterol and glycosphingolipid con- tent that can be found on the plasma- and endosomal-mem- brane of eukaryotic cells (London and Brown, 2000; Simons and Toomre, 2000; Simons and Ehehalt, 2002). Some vi- ruses, such as HIV-1, appear to target lipid raft micro- domains during viral entry into cells, as well as during viral Figure 1. DC-SIGN is organized in microdomains on the cell surface of K-DC-SIGN. (A) The expression levels of DC-SIGN on assembly before budding from cells (Dimitrov, 1997; Mañes untransfected K562 and on K-DC-SIGN were assessed by FACS et al., 2000). Other works suggest that cholesterol-depen- analysis. The open histogram represents the isotype control, and dent membrane properties, rather than lipid rafts per se, are shaded histogram indicates the specific staining with anti–DC-SIGN responsible to promote efficient HIV-1 infection in T cells (AZN-D1). (B) DC-SIGN expressed by K562 transfectants strongly (Percherancier et al., 2003). binds to ICAM-3 and gp120. The adhesion was determined using DC-SIGN, like other C-type lectins, recognizes pathogens 1-m ligand-coated fluorescent beads. Specificity was determined by measuring binding in presence of AZN-D1. No blocking was by binding to carbohydrate moieties in a Ca -dependent observed in presence of isotype control (not depicted). Blocking manner, through a conserved CRD (Drickamer, 1999). exerted by EGTA indicates that DC-SIGN binds to the ligands in a This CRD has a high specificity for complex mannose resi- Ca -dependent manner. The average of five independent experi- dues and is located at the distal end of the extracellular do- ments is shown (P  0.001). (C) K-DC-SIGN cells were specifically main (ECD), which consists of several amino acid repeats labeled with 10-nm gold particles, as described in Materials and (Soilleux et al., 2000). Recently, purified truncated forms of methods. Cells were allowed to adhere onto poly-L-lysine–coated formvar film and photographed in an electron microscope. Gold DC-SIGN containing either the complete ECD or only the particles were detected on the periphery on the thinner less electron CRD were used to analyze the quaternary structure as well as dense parts of cells, where good contrast could be achieved. One the affinity of DC-SIGN for its ligands. Biochemical experi- representative picture is shown. Bar, 200 nm. ments indicated that in vitro ECDs aggregate to form tetra- mers, thus enhancing DC-SIGN capacity to bind multiva- lent ligands, such as pathogen sugar arrays (Mitchell et al., 1-m fluorescent beads coated with ICAM-3 or GP120 (Fig. 2001). Surface plasmon resonance experiments showed that 1 B). The binding of K-DC-SIGN to these ligands was spe- whereas the ECD readily binds hepatitis C virus glycopro- cific, as shown by the inhibition exerted by anti–DC-SIGN tein E2, distant monomeric CRDs do not, unless closely mAbs. Moreover, the lack of adhesion in presence of EGTA seeded (Lozach et al., 2003). confirmed that DC-SIGN bound to these ligands in a Ca - These findings suggest that the organization of DC-SIGN dependent manner, typical of C-type lectin-like receptors. molecules on the plasma membrane may be critical for To determine the organization of DC-SIGN on the cell pathogen binding. This prompted us to investigate the cell membrane at high resolution, TEM was used on whole- surface distribution pattern of DC-SIGN and its possible as- mount samples of K-DC-SIGN, after specific labeling with sociation with lipid rafts. Using transmission EM (TEM) on anti–DC-SIGN mAb and 10-nm gold particles. It should be whole-mount samples of transfected cells and monocyte- noted that the specimens were not sectioned thus making the derived DCs, we have mapped the microlocalization of whole cell surface available for gold labeling and subsequent DC-SIGN at high resolution. Subsequently, spatial-point TEM analysis. This method has been successfully applied to pattern analysis showed that DC-SIGN is organized in detect spatial distribution of other membrane proteins such well-defined microdomains. Moreover, biochemical experi- as the potassium channels Kv1.3 (Panyi et al., 2003) and the ments and confocal microscopy analysis demonstrated that IL-2 receptor -subunit (Vereb et al., 2000). As shown in DC-SIGN colocalizes with lipid rafts. Finally, we show that Fig. 1 C, DC-SIGN showed a clear distribution in well- these microdomains of DC-SIGN act as a docking site for defined microdomains on K-DC-SIGN plasma membrane. HIV-1 particles, facilitating entry of the virus into DCs. These microdomains had an average diameter of 200 nm and appeared to be randomly localized on the cell surface. To ex- clude that the gold labeling pattern observed could be due to Results internalized DC-SIGN molecules; also, sections of these cell DC-SIGN is organized in microdomains samples were analyzed by TEM. The results demonstrated on the cell surface that the gold particles were exclusively detected on the out- To study the correlation between functional state and cell side of the cell membrane (unpublished data), confirming the surface organization of DC-SIGN, we used K562 transfec- presence of DC-SIGN in microdomains on the cell surface. tants stably expressing DC-SIGN. K-DC-SIGN expressed To show that the clustering of DC-SIGN is not an artifact high levels of DC-SIGN (Fig. 1 A) and bound strongly to due to the procedure, we also analyzed by TEM other trans- DC-SIGN microdomains | Cambi et al. 147 membrane receptors transfected into K562, among which the 2-integrin LFA-1. Unlike DC-SIGN, LFA-1 molecules showed a random distribution pattern (unpublished data). It should be noted that the number of gold particles observed per micrometer squared was generally comparable with that observed for K-DC-SIGN, which should exclude difference in distribution due to major difference in the expression lev- els of the two receptors. DC-SIGN predominantly resides in lipid rafts To determine whether lipid rafts are important for DC- SIGN function, K-DC-SIGN cells were treated with methyl--cyclodextrin (MCD) to extract membrane choles- terol, and its effect on DC-SIGN–mediated ligand binding was tested by the fluorescent beads adhesion assay. As shown in Fig. 2 A, MCD treatment partially inhibits binding to gp120-coated beads, indicating that cholesterol extraction partially affected DC-SIGN ligand binding capacity. To further demonstrate the association of DC-SIGN with lipid rafts, K-DC-SIGN cells were solubilized in Triton X-100 on ice and fractionated by centrifugation on a sucrose gradient at 4C. With this procedure, lipid rafts, which can be isolated as detergent-resistant membranes (DRM), were recovered in low density fractions, whereas any other deter- gent-soluble material was concentrated in the high density fraction (Fig. 2 B). The GPI-anchored protein CD55 was used as the raft marker and was found in raft fractions, whereas the negative control (nonraft-associated protein), CD46, was almost completely detectable in the nonraft frac- tions. Similarly to CD55, a significant portion of DC-SIGN was recovered in the low density fraction, indicating that DC-SIGN is localized in DRM on the plasma membrane of K-DC-SIGN cells. To provide further evidence that DC-SIGN colocalizes with lipid rafts, we examined the co-distribution of DC- SIGN and the lipid raft marker ganglioside (GM1), on K-DC-SIGN by antibody patching and confocal micros- copy. As shown in Fig. 2 C, when capping was induced on K-DC-SIGN cells, GM1 clearly colocalizes with DC-SIGN and to the same extent as for CD55. However, some smaller patches of DC-SIGN were also detectable outside the lipid raft area, indicating that DC-SIGN might not permanently reside in lipid rafts, as also observed in Fig. 2 B. As expected, no colocalization of CD46 with GM1 was detected. Based on these observations, we conclude that DC-SIGN resides Figure 2. DC-SIGN colocalizes with lipid rafts on K-DC-SIGN. within a lipid raft environment on the plasma membrane of (A) To investigate the effect of cholesterol depletion on DC-SIGN– K-DC-SIGN cells. mediated adhesion, K-DC-SIGN cells were incubated in serum-free medium with or without 20 mM MCD for 30 min at 37C. Subse- DC-SIGN distribution changes during DC development quently, gp120-coated fluorescent beads (1-m diam) were added and the mixture was incubated for an additional 30 min at 37C. During the differentiation of DCs from monocyte precursors, Binding was measured by flow cytometry. After MCD treatment, cell the expression of DC-SIGN on the cell surface gradually in- viability was assessed by trypan blue staining. The values represent creases (Geijtenbeek et al., 2000a). However, as shown by the mean of three independent experiments SD. (B) K-DC-SIGN flow cytometry (Fig. 3 A), no significant increases in DC- were solubilized with 1% Triton X-100, subjected to sucrose gradient SIGN expression levels are seen between cells harvested after centrifugation and analyzed by Western blotting for the indicated molecules. The numbers indicate the gradient fractions. Fractions 9 3 d of culture (designated intermediate DCs) and immature and 10 are low density fractions containing DRM and are referred to as raft fractions. (C) Confocal microscopy analysis of copatching of DC-SIGN and GM1. K-DC-SIGN cells were stained at 4C with 10 g/ml anti–DC-SIGN (or anti-CD55 or anti-CD46) and 10 g/ml by confocal microscopy. Merged images are shown in the right panel. FITC-CTxB. Co-patching was induced by adding secondary Ab Results are representatives of multiple cells in three independent (Materials and methods), and, after fixation in PFA, cells were analyzed experiments. Bar, 5 m. 148 The Journal of Cell Biology | Volume 164, Number 1, 2004 Figure 3. DC-SIGN cell surface distribution during monocyte- derived DC development. DC-SIGN binding activity was monitored during development of monocyte-derived DCs. As shown in the box, intermediate DCs indicate cells harvested after 3 d of monocytes differentiation. (A) The expression levels of DC-SIGN on monocytes, intermediate and immature DCs were assessed by FACS analysis. The dotted line histogram represents the isotype control, and the thick line histogram indicates the specific staining with anti–DC-SIGN (AZN-D1). Mean fluorescence intensity is indicated. One represen- tative donor is shown. (B) The adhesion to ICAM-3 and gp120 was determined using 1 m ligand-coated fluorescent beads. Specificity was determined by measuring binding in presence of AZN-D1. No blocking was observed in presence of isotype control (not depicted). One representative experiments out of three is shown. (C) Intermediate and immature DCs were let adhere onto fibronectin-coated formvar film, specifically labeled for DC-SIGN with 10-nm gold particles (Materials and methods), and analyzed by TEM. Results are representatives of multiple cells in several independent experiments. Bar, 200 nm. DCs. Maximum DC-SIGN–mediated adhesion to ICAM-3 gold particles are evenly distributed over the cell surface, on as well as GP120 was observed on immature DCs, although immature DCs, there is a clear organization of DC-SIGN in already on intermediate DC DC-SIGN was capable of com- spatially well-defined microdomains. To exclude the possible pletely mediating the binding to ICAM-3 (Fig. 3 B, top), influence of the fibronectin substrate on DC-SIGN distribu- which on monocytes is LFA-1 dependent (unpublished data). tion, TEM analysis was also performed on DCs that were gold Comparably, while on monocytes, binding to gp120 is medi- labeled in suspension and mounted onto poly-L-lysine. No ated by CD4 (Kedzierska and Crowe, 2002; Kohler et al., differences were seen between cells stretched on fibronectin or 2003), on intermediate DCs, as well as on immature DCs, cells adhering to poly-L-lysine (unpublished data). Moreover, DC-SIGN is almost entirely responsible for binding to gp120 thin sections of resin-embedded immature DCs were gold la- (Fig. 3 B, bottom). To examine DC-SIGN cell surface distri- beled and analyzed by TEM. Clusters of DC-SIGN molecules bution, both intermediate and immature DCs were allowed to could be observed exclusively at the plasma membrane (un- adhere to fibronectin, and DC-SIGN molecules were labeled published data). We also analyzed by TEM the cell surface dis- with gold particles. Subsequently, the distribution on the tribution of other transmembrane receptors expressed on plasma membrane was analyzed by TEM (Fig. 3 C). Given DCs, including LFA-1. Unlike DC-SIGN, LFA-1 did not the high capacity of DCs to widely spread on the used sub- show any changes in cell surface distribution pattern on inter- strates, very large membrane areas (often up to 60–70% of the mediate and immature DCs (unpublished data). whole visible plasma membrane) were available for gold parti- To quantitatively describe the DC-SIGN distribution pat- cles analysis, ensuring that the areas used for quantitation were tern, the nearest neighbor distance values among the gold truly representative (Fig. S1, available at http://www.jcb. particles were calculated applying a spatial-point pattern org/cgi/content/full/jcb.200306112/DC1). Surprisingly, we analysis. A quantitative comparison of DC-SIGN clustering found that the distribution of DC-SIGN changes dramatically among intermediate DCs, immature DCs, and K-DC-SIGN during DC development. Although on intermediate DCs, the is shown in Fig. 4 A. On immature DCs, as well as on DC-SIGN microdomains | Cambi et al. 149 Figure 4. Quantitative analysis of the distribution of gold particles labeling DC-SIGN. The digital images of electron micrographs were processed by a custom-written software based on Labview. Gold labels were counted, and coordinates were assigned to each feature. Subsequently, interparticles distances were calculated using a nearest neighbor distance algorithm. Nearest neighbor distance values were calculated for each image, and the data of several independent experiments were pooled. Subse- quently, the nearest neighbor distances were divided into three classes 0–50 nm (gray bar), 50–150 nm (black bar), and 150 nm (white bar), and the percentage of nearest neighbor distance values falling into each class was plotted (A). The partitioning of gold labels in clusters of various size (i.e.: number of particles/cluster) was also quantified. Clusters were defined when gold particles were 50 nm apart from a neighboring particle. The percentage of gold particles involved in the formation of a certain cluster size was calculated for (B) K-DC-SIGN, (C) immature DC, and (D) intermediate DC. The insets are three representative processed digital images, where each type of cluster is shown in a different color. For K-DC-SIGN, one represen- tative experiment out of two is shown; for immature DC, one representative experiment out of six is shown; and for intermediate DCs, one representative experiment out of three is shown. K-DC-SIGN, almost 70 and 80% of the gold particles reside erage diameter of 200 nm and were separated from each within a 50-nm distance from its nearest neighbor, respec- other by an average distance of 400 nm. Spatial analysis of the tively. In contrast, on intermediate DCs, only 30% of the microdomains distribution over the cell surface did not reveal nearest neighbor distance values are found in the same cate- any preferential localization in specific surface areas of the ob- gory, and the majority of interparticle distances rather fall served cells. These data were also confirmed by using high res- within the 50–150 nm range. It has to be noted that the in- olution near-field fluorescence imaging (unpublished data). termediate and the immature DCs compared by spatial-point The relative partitioning of gold particles in clusters of pattern analysis had a similar amount of gold particles per various sizes (i.e., number of particles/cluster) was also quan- micrometer squared, confirming the comparable expression tified. A value of 50 nm was set to define the involvement of levels of DC-SIGN on the two cell types. a gold particle in a cluster. Fig. 4 D shows that on interme- Similarly to what was observed for K-DC-SIGN, on imma- diate DCs up to 80% of gold particles occur as single fea- ture DCs, these microdomains were also found to have an av- tures. In contrast, on immature DCs (Fig. 4 C), as well as on 150 The Journal of Cell Biology | Volume 164, Number 1, 2004 K-DC-SIGN (Fig. 4 B), DC-SIGN shows a much broader served on K-DC-SIGN, small patches of DC-SIGN were distribution of cluster sizes, with an average of 10–20 gold also found not colocalizing with GM1. To further prove the particles per cluster on immature DCs. colocalization of GM1 with DC-SIGN, we performed dou- ble gold labeling for DC-SIGN and GM1 on immature DCs, and analyzed the samples by TEM. As shown in Fig. 5 DC-SIGN resides in lipid rafts on DCs C, GM1 (10 nm gold) was found within DC-SIGN micro- Because, on K-DC-SIGN, we found a clear association of domains (5 nm gold). Consistent with observations by con- DC-SIGN with lipid rafts, we investigated whether DC- focal microscopy, some microdomains of DC-SIGN that SIGN also colocalized with lipid rafts on immature DCs. lacked GM1 were also detected. Together, these results Extraction of cholesterol by MCD resulted in a significant demonstrate that on immature DC DC-SIGN resides in mi- block in binding of both ICAM-3– and gp120-coated beads crodomains that to a significant extent are associated with a (Fig. 5 A). In contrast, DC-SIGN–mediated binding on in- lipid rafts environment. termediate DCs was not affected by cholesterol depletion (unpublished data). DC-SIGN in microdomains facilitates binding To further demonstrate an interaction of DC-SIGN with to virus-sized particles lipid rafts on immature DCs, we investigated the co-distri- bution of DC-SIGN and GM1 by confocal microscopy. Fig. We showed that fluorescent beads coated with DC-SIGN– 5 B shows that, when capping was induced, DC-SIGN specific ligands were bound by immature as well as interme- clearly colocalized with GM1, almost to the same extent as diate DCs, apparently independently of DC-SIGN organi- the raft marker CD55. However, similarly to what was ob- zation on the cell membrane (Fig. 3, A and B). Figure 5. DC-SIGN resides in lipid rafts on immature DCs. (A) DC-SIGN–mediated adhesion to ligand-coated fluorescent beads (1-m diam) on immature DCs was measured after cholesterol depletion by 20 mM MCD. The assay was performed as described in Fig. 2 A. Data shown are means  SD of one representative experiment performed in triplicate out of three. One representative experiment out of three is shown. (B) Confocal microscopy analysis of copatching of DC-SIGN and GM1 on immature DCs. Staining was performed as described in Fig. 2 C. Cells were analyzed by confocal microscopy. Merged images are shown in the right panel. Results are representative of multiple cells in two independent experiments. Bar, 5 m. (C) Whole-mount samples of immature DCs were double labeled for DC-SIGN (5 nm gold) and the raft marker GM1 (10 nm gold) and analyzed by TEM. Thin white arrows indicate GM1 colocalizing in DC-SIGN microdomains. Thick white arrows indicate DC-SIGN microdomains with no GM1. Black arrow indicates GM1 alone. Results are representatives of multiple cells in two independent experiments. Bar, 50 nm. DC-SIGN microdomains | Cambi et al. 151 However, because DC-SIGN on immature DCs is an ex- any microbeads, comparable to background levels or when quisite virus receptor and the fluorescent beads are at least DCs were pretreated with the carbohydrate mannan to one order of magnitude larger than viral particles, we inves- block DC-SIGN (Fig. 6, C and F). Similar results were ob- tigated whether clustering of DC-SIGN into higher order tained when microbeads coated with ICAM-3 were used assemblies might be needed to stabilize interactions with vi- (unpublished data). Moreover, DC-SIGN on intermediate rus-sized particles. To detect possible differences in “avidity” as well as on immature DCs was found to be equally capable for multimeric ligands between scattered and clustered DC- of binding to soluble recombinant gp120 (Fig. 6 G), sug- SIGN molecules, and to mimic virus binding to DC-SIGN, gesting that the intrinsic binding capacity of DC-SIGN we used fluorescent virus-sized microbeads of 40-nm diam, molecule on the two different DC types is comparable. Sim- coated with gp120, which are now referred to as virus-sized ilar results were obtained when soluble ICAM-3-Fc chime- particles. The extremely broad fluorescence emission peak of ras were used (unpublished data). All these observations sup- these microbeads currently precludes accurate quantification port our hypothesis that DC-SIGN molecules acquire a by flow cytometry, therefore binding of DCs to these virus- higher avidity for multimeric ligands when organized in sized particles was visualized by confocal microscopy. multimolecular assemblies. When comparing the capacity of intermediate and imma- DC-SIGN microdomains enhances HIV-1 infection ture DCs to bind these virus-sized particles (Fig. 6), a very high binding was observed on immature DCs. Many virus- Having established that DC-SIGN microdomains bind sized particles were bound to the plasma membrane and a virus-sized particles more efficiently in comparison to significant amount was phagocytosed (Fig. 6, D and E). In randomly distributed receptor molecules, we investigated contrast, intermediate DCs (Fig. 6, A and B) hardly bound whether the uptake of real virus was also enhanced by a clus- Figure 6. DC-SIGN microdomains on immature DCs bind virus-sized particles. Green fluorescent microbeads (40-nm diam) were coated with gp120 and added to the cells in a ratio of 20 beads/cell. The cells were incubated for 30 min at 37C, washed, and fixed in PFA. DC-SIGN was stained with AZN-D1 and Alexa 647– conjugated goat anti–mouse Ab (red). Subsequently, the cells were mounted onto poly-L-lysine–coated glass coverslips and analyzed by confocal microscopy. The overview of binding to gp120 micro- beads of (A) intermediate DCs and (D) immature DCs is shown. Two represen- tative cells of (B) intermediate and (E) immature DCs are shown. Specific block with 100 g/ml mannan was also per- formed by preincubating the cells at RT for 10 min before adding the microbeads (C and F); bars, 5 m. Similar results were obtained with ICAM-3–coated microbeads (not depicted). (G) Binding of K-DC-SIGN, intermediate DCs, and immature DCs to soluble gp120 was also performed: 50,000 cells were incubated with 50 mM biotinylated gp120 for 30 min on ice, in presence or absence of 20 g/ml anti–DC-SIGN blocking mAb (AZN-D1). A subsequent incubation with Alexa 488–conjugated streptavidin for 30 min on ice followed, and gp120 molecules bound to the cells were detected by flow cytometry. The values represent the mean of three independent experiments SD. 152 The Journal of Cell Biology | Volume 164, Number 1, 2004 lipid raft environment on DCs that forms highly organized well-defined (200 nm) multiprotein assemblies on the sur- face of a cell. To establish the association of proteins with lipid rafts is complicated and often controversial, which underscores the complexity of cell membranes. These lipid domains differ in their composition, physical properties, and biological func- tions (Schuck et al., 2003). Moreover, lipid raft composi- tions differ between cell types. Finally, the dynamics of the chemical membrane composition adds a further level of complexity. Therefore, the association of proteins with lipid Figure 7. Clustered DC-SIGN molecules efficiently bind HIV-1 rafts can only be analyzed using different techniques simul- particles and infect PBMC. DC-SIGN on immature DCs enhances HIV-1 infection as measured in a DC-PBMC coculture. Either inter- taneously. Our observation that DC-SIGN resides within mediate or immature DCs (1.5 10 ) were preincubated for 20 min lipid rafts is based on several well-established raft analysis at RT with or without blocking mAb against 20 g/ml DC-SIGN techniques. First, we showed by biochemical experiments (AZN-D1 and AZN-D2). Preincubated intermediate or immature DCs the partitioning of DC-SIGN in DRM. Second, co-patch- were pulsed for 2 h with HIV-1 (M-tropic HIV-1Ba-L strain), and ing of DC-SIGN with the lipid raft marker GM1 was unbound virus particles and mAb were washed away. Subsequently, observed by confocal microscopy. Third, this association DCs were cocultured with activated PBMC (1.5 10 ) for 7 d. Coculture supernatants were collected, and p24 antigen levels were between DC-SIGN and GM1 was confirmed by TEM. measured by ELISA. Black histogram represents PBMC infected in the Fourth, cholesterol extraction by MCD inhibited DC- absence of DCs. One representative experiment out of two is shown. SIGN–mediated binding. Surprisingly, on immature DCs, the same MCD treatment did not have any effect on the or- ganization of DC-SIGN microdomains, as observed by tered distribution of DC-SIGN, using p24 ELISA. There- TEM (unpublished data). This may be explained by the fact fore, intermediate and immature DCs were pulsed for 2 h that the remaining cholesterol may be sufficient to keep such with HIV-1 (M-tropic HIV-1Ba-L strain), washed, and cul- domains intact (Schuck et al., 2003). Moreover, besides tured in the presence of activated peripheral blood mononu- cholesterol, also glycosphingolipids, another lipid raft com- clear cells (PBMC). As shown in Fig. 7, virus replication was ponent, which is not depleted by MCD treatment, may in significantly higher when PBMC were cocultured with ei- part maintain the integrity of DC-SIGN microdomains. ther intermediate or immature DCs with respect to PBMC The fact that cholesterol extraction by MCD partially affects alone challenged with the same amount of infectious virus. DC-SIGN–mediated binding could be explained by consid- However, infection of PBMC cultured with virus-pulsed ering the pleiotropic effects that MCD can have on cell immature DCs was much higher than the infection exhib- functions, besides disrupting lipid rafts integrity (Brown and ited by PBMC cultured with intermediate DCs. In addition, London 2000; Edidin 2001; Schuck et al., 2003). There- the infection of PBMC with immature DCs was signifi- fore, MCD treatment can only be a preliminary indicator cantly blocked by anti–DC-SIGN mAbs that were added to for a possible association of a protein with a lipid rafts envi- DCs before incubation with HIV-1. In contrast, DC-SIGN ronment. Also, it should be added that, unlike for GPI- contribution to PBMC infection was clearly much lower anchored proteins, very little is known about how trans- when intermediate DCs were used. The incomplete block- membrane proteins are recruited into lipid rafts, how stable ing observed in the presence of anti–DC-SIGN Abs suggests these interactions are and what exactly the role of cholesterol the involvement of other HIV receptors also expressed on is. Further experiments are needed to better identify the mo- DCs (Turville et al., 2002). The comparable expression lev- lecular determinants that control the association of trans- els of DC-SIGN, as shown in Fig. 3 A, as well as several co- membrane proteins with lipid rafts. stimulatory molecules (such as CD40 and CD80) on inter- In vitro experiments with isolated recombinant C-type lec- mediate and immature DCs suggest that these two DC types tins suggested that these can oligomerize providing multiple have similar antigen presentation capacity (unpublished surfaces to bind (multivalent) ligands (Drickamer, 1999). data). Altogether, our data demonstrate that DC-SIGN or- DC-SIGN has been shown to form tetramers stabilized by an ganized in microdomains rather than randomly distributed -helical stalk (Mitchell et al., 2001), and purified truncated is more efficient in mediating binding and uptake of virus- forms of DC-SIGN containing the complete ECD were also sized microbeads as well as real HIV-1 particles. shown to be able to bind ligands by surface plasmon reso- nance (Lozach et al., 2003). In these works, the affinity of Discussion isolated CRDs or complete ECDs of DC-SIGN for the ligand was measured and compared with the affinity of the DC-SIGN is a DC-specific C-type lectin that acts both as adhesion molecule and as pathogen-recognition receptor. membrane-bound form. Although no interactions with the ligand could be found using isolated CRDs, significant bind- Here, we demonstrate that DC-SIGN can form micro- domains on the plasma membrane, and that there is a direct ing was detected if single CRDs were closely seeded (K nM) or if CRDs were part of a complete oligomeric-soluble correlation between the distribution of DC-SIGN in micro- domains and its capacity to bind and internalize virus-sized ECD of DC-SIGN (K 30 nM). The highest affinity was seen with membrane-bound DC-SIGN expressed on the sur- ligand-coated particles. Moreover, to the best of our knowl- edge, this is the first report describing a C-type lectin in a face of transfected cells (K 3 nM), suggesting that the nat- DC-SIGN microdomains | Cambi et al. 153 ural plasma membrane environment strongly influences the function of this receptor. To some extent, this mirrors our observations: DC-SIGN on the cell surface of intermediate DCs is randomly distributed and hardly binds virus-sized particles, whereas on immature DCs, the protein is organized in well-defined microdomains, which allows binding of 40- nm virus-sized particles (Fig. 6, A–F). In apparent contrast, we showed that on intermediate DCs, randomly distributed DC-SIGN was able to bind 1-m beads, coated with ICAM-3 or gp120 (Fig. 3 B). This apparent discrepancy between 40-nm virus-sized particles and 1-m beads can be explained Figure 8. DC-SIGN microdomains enhances binding of virus-sized by the fact that beads of 1-m diam have 600-fold larger particles with respect to isolated DC-SIGN molecules. Beads of interaction surface saturated with numerous ligand molecules 1-m diam are saturated with numerous coated ligand molecules that can engage simultaneous interactions with several indi- that can engage simultaneous interactions with several individual vidual DC-SIGN molecules. These multiple interactions DC-SIGN molecules. These multiple interactions may strengthen the binding both with random and clustered DC-SIGN. In contrast, when may signal into the cell and lead to a rapid recruitment of virus-sized particles are used, the contact surface and therefore, the new DC-SIGN molecules, thus strengthening the binding. number of ligand molecules is much smaller. Consequently, only In contrast, when virus-sized particles are used, the contact interactions with DC-SIGN molecules in highly organized multipro- surface and therefore, the number of ligand molecules is tein assemblies may result in stable binding of virus particles. much smaller. Consequently, only interactions with DC- SIGN molecules in highly organized multiprotein assemblies pathogen-associated molecular patterns displayed at the can result in stable binding of virus-sized particles (Fig. 8). cell surface of microorganisms. Although TLRs have been When a computer-aided simulation was performed to predict shown to induce differential gene expression upon recogni- the capacity of round objects of different sizes to establish in- tion of distinct pathogen structural components (Akira, teractions with either random or clustered DC-SIGN mole- 2003), for the C-type lectins, and particularly for DC- cules, we found that objects with a diameter in the range of SIGN, no direct signaling pathways has been demonstrated virus sizes (40–200-nm diam) preferentially bind to clusters so far. Therefore, it will be interesting to investigate the in- of DC-SIGN having the same size (unpublished data). In teractions that might occur between these two families of agreement with this model, no significant differences were pathogen-recognition receptors. detected in binding of intermediate and immature DCs to In conclusion, our findings emphasize the importance of soluble gp120 or to ICAM-3-Fc chimeras, indicating that the relating the function with the cell surface organization of capacity of each single DC-SIGN molecule to recognize and DC-SIGN. Clustered distribution is essential to enhance the bind the ligand is comparable on both DC types (Fig. 6 G). interaction as well as the internalization efficiency of DC- The biological relevance of this clustering phenomenon is SIGN–pathogen complexes. In addition, our data highlight shown by the fact that immature DCs compared with inter- the importance of the plasma membrane as a specialized mediate DCs showed an enhanced DC-SIGN–mediated microenvironment, where finely orchestrated interactions binding and internalization of virus particles, and subse- among proteins, carbohydrates, and lipids take place. These quent infection of PBMC in trans (Fig. 7). Therefore, we complex interactions mediate many fundamental processes propose that engagement of viruses with DC-SIGN occurs that occur at the contact site between cells (cell–cell or cell– much more productively when this receptor is organized in pathogen), such as assembly of signaling platforms and for- microdomains and resides in a lipid raft or cholesterol- mation of entry portals for invasive pathogens. enriched environment. Insight in the cell surface organization of pathogen It remains to be established what controls DC-SIGN ran- scavenger receptors like DC-SIGN will contribute to the dom distribution on intermediate DCs and what mecha- development of novel strategies that specifically inhibit nism is responsible for the change into a clustered organi- interactions with life-threatening pathogens like HIV-1 or zation on immature DCs. We cannot exclude that, on Mycobacterium. intermediate DCs, DC-SIGN may interact with an un- known membrane associated protein that prevents the for- mation of such microdomains. Alternatively, cytoskeletal Materials and methods constraints may differ between these cells, and this issue is Antibodies and reagents currently under investigation. Preliminary observations sug- For labeling as well as blocking of DC-SIGN, the mAb AZN-D1 was used gest that releasing DC-SIGN molecules from the cortical ac- (Geijtenbeek et al., 2000a). For detection of CD46 and CD55 the Abs E4.3 (BD Biosciences) and 143–30 (CLB) were used, respectively. Alexa 647– tin cytoskeleton by cytochalasin D treatment results in an conjugated goat anti–mouse IgG was purchased from Molecular Probes. enhanced ligand binding, particularly on intermediate DCs FITC-conjugated cholera toxin B subunit (FITC-CTxB), Triton X-100, and (unpublished data). Further experiments are needed to ob- MCD were purchased from Sigma-Aldrich. Goat anti-CTxB antibody was purchased from Calbiochem. tain insight into the formation of DC-SIGN multiprotein assemblies on DCs. Cells DC-SIGN belongs to the C-type lectin family that, to- Monocytes were obtained from buffy coats of healthy individuals and were gether with the TLR family, forms the first barrier against purified using Ficoll density centrifugation. Immature DCs were obtained invading pathogens. This occurs through recognition of as reported elsewhere (Geijtenbeek et al., 2000a). In brief, DCs were cul- 154 The Journal of Cell Biology | Volume 164, Number 1, 2004 tured from monocytes in presence of IL-4 and GM-CSF (500 and 800 defined when gold particles were less than a set distance apart from a U/ml, respectively) for 3 d to obtain intermediate DCs and for 6 d to obtain neighboring particle. immature DCs. Stable K562 transfectants expressing DC-SIGN (K-DC- SIGN) were generated as published previously (Geijtenbeek et al., 2000a). Confocal microscopy Cells were stained at 4C with 10 g/ml anti–DC-SIGN antibody (AZN- Adhesion assays D1), anti-CD55 antibody, anti-CD46 antibody, and 10 g/ml FITC-CTxB. Carboxylate-modified TransFluorSpheres (488/645 nm, 1-m diam; Mo- Isotype-specific controls were always included. Secondary staining for lecular Probes) were coated with ICAM-3-Fc or GP120, and the fluores- DC-SIGN, CD55, and CD46 was with Alexa 647–conjugated goat anti– cent beads adhesion assay was performed as described previously (Geij- mouse IgG, and for FITC-CTxB with goat anti-CTxB. Patching was induced tenbeek et al., 1999). When the lipid raft-disrupting agent MCD was used, by incubation at 12C for 1 h, followed by fixation with 1% PFA. Cells the cells were resuspended in serum-free medium containing the appropri- were mounted onto poly-L-lysine–coated glass coverslips. Signals were ate concentration of MCD and preincubated for 30 min at 37C. When collected sequentially to avoid bleed through. necessary, 50,000 cells were incubated with 20 g/ml blocking mAb for 10 min at RT. The ligand-coated fluorescent beads (20 beads/cell) were Detergent extraction and flotation assay added and the suspension was incubated for 30 min at 37C. Adhesion For detergent resistant membrane isolation, K-DC-SIGN cells (10 10 ) was determined by measuring the percentage of cells, which have bound were lysed on ice in 0.5 ml of lysis buffer (50 mM Tris, pH 7.4, 150 mM fluorescent beads, by flow cytometry using the FACSCalibur™ (Becton NaCl, 5 mM EDTA, and 1% Triton X-100, plus a cocktail of protease inhibi- Dickinson). Streptavidin-modified TransFluorSpheres (505/515 nm, 40-nm tors: 1 mM PMSF, 10 g/ml aprotinin, and 10 g/ml leupeptin). After 30 min diam; Molecular Probes) were coated with ICAM-3-Fc or GP120 as de- of incubation, the lysate was made 40% with respect to sucrose. Next, the scribed previously for 1-m beads (Geijtenbeek et al., 1999) and were lysate-sucrose mixture was overlaid with 2 ml of 30% sucrose in lysis buffer added to DCs in a ratio of 20 beads/cell. Bound beads were detected by and finally with 1 ml of 4% sucrose in lysis buffer. The mixture was centri- confocal microscopy analysis of the cells. fuged at 200,000 g for 14–16 h in an SW60Ti rotor (Beckman Coulter). The When binding to soluble GP120 was performed, 50 mM biotinylated gradient was fractionated in 0.5-ml fractions from the bottom of the tube. GP120 was added to the cells (10 ) for 30 min on ice, in presence or ab- sence of 20 g/ml anti–DC-SIGN blocking Ab (AZN-D1). After the incuba- Immunoblot analysis tion, cells were extensively washed and incubated with Alexa 488–con- Proteins from the sucrose fractions were separated by SDS-PAGE and trans- jugated streptavidin for 30 min on ice. Subsequently, bound GP120 ferred to PROTRAN nitrocellulose transfer membranes (Schleicher & molecules were detected by flow cytometry. Schuell). Membranes were blocked for 1 h at RT using 2% nonfat dried milk (Campina) and 1% BSA (Calbiochem) in 0.1% PBS/Tween 20. Blots were Labeling procedures subsequently incubated with specific antibodies (polyclonal anti-CD55 and For flow cytometry analysis, cells were incubated (30 min; 4C) in PBA CD46 from Santa Cruz Biotechnology, Inc., and a polyclonal anti–DC- (PBS, containing 0.5% BSA and 0.01% sodium azide), with different mAb SIGN antibody [CSRD] that was raised in our laboratory), followed by the (5 g/ml), followed by incubation with FITC-labeled goat anti–mouse IgG appropriate HRP-conjugated secondary antibodies (DakoCytomation). Fi- antibody (GAM-FITC; Zymed Laboratories; 1:50 dilution in PBA) for 30 nally, proteins were detected using ECL (Amersham Biosciences). min at 4C. The relative fluorescence intensity was measured on a FACS- Calibur™. Isotype–specific controls were included. For TEM, K-DC-SIGN HIV-1 infection of DCs L-lysine–coated formvar, whereas DCs were allowed were fixed onto poly- Infection of intermediate and immature DCs with the M-tropic strain HIV- to spread on glass coverslips covered by a thin layer of fibronectin-coated 1Ba-L was performed as described previously (Geijtenbeek et al., 2000b). formvar for 1 h at 37C and immediately fixed with 1% PFA for 15 min. Af- 6 ) were incubated with In brief, intermediate and immature DCs (1.5 10 ter two washing steps with PBS and a subsequent incubation (60 min at mAb against 20 g/ml DC-SIGN (AZN-D1 and 20 g/ml AZN-D2 for 20 RT) with I-buffer (PBS, 0.1% glycine, 1% BSA, and 0.25% gelatin) to re- min at RT. Subsequently, cells were incubated with the M-tropic strain duce a specific background, the specimens were incubated for 30 min HIV-1Ba-L for 2 h with a multiplicity of infection of 0.004. After incuba- with the primary antibodies in I-buffer on ice, rinsed in PBS, and fixed in tion, cells were washed and cocultured with activated PBMC in the pres- 1% PFA and 0.1% glutaraldehyde for 15 min. After two washing steps with ence of 5 U/ml human recombinant IL-2 (Roche Diagnostics). PBMC were PBS and a block in I-buffer, the samples were incubated with rabbit anti– activated by culturing them before infection for 3 d in the presence of 2 mouse IgG (to detect mAb) for 30 min on ice. A final incubation with 10- g/ml of phyto-HA (Sigma-Aldrich). After 7 d, culture supernatants were nm-diam gold-labeled Protein A (to detect polyclonal antibodies) was per- collected, and p24 antigen levels were determined by a p24 antigen ELISA formed, followed by final fixation in 1% glutaraldehyde in phosphate TM ; DakoCytomation). (AMPAK buffer for 20 min at RT. When double labeling was performed, the cells were treated as described above, except that anti–DC-SIGN (AZN-D1), Online supplemental material and biotinylated-CTxB were added simultaneously; and finally, goat anti– Fig. S1 is an overview of DC whole-mount samples by TEM. To determine mouse-conjugated 5-nm gold and streptavidin-conjugated 10-nm gold the organization of DC-SIGN on the cell membrane at high resolution, TEM were used to label DC-SIGN and GM1, respectively. Isotype-specific con- was used on whole-mount samples of intermediate as well as immature trols were always included. The 10-nm-diam gold-labeled protein A was a DCs, after specific labeling with anti–DC-SIGN mAb and 10-nm gold parti- gift of M. Wijers and H. Croes (Nijmegen Center for Molecular Life Sci- cles. It should be noted that the specimens were not sectioned, thus, mak- ences). The goat anti–mouse-conjugated 5-nm gold and streptavidin-con- ing the whole cell surface available for gold labeling and subsequent TEM jugated 10-nm gold were purchased from Aurion. analysis. DCs were adhered onto fibronectin-coated formvar film, fixed, and labeled as described in Materials and methods, and finally analyzed by EM and analysis of the distribution patterns of gold particles TEM. Given the high capacity of both intermediate and immature DCs to After gold labeling and fixation, the specimens were dehydrated by se- widely spread on the substrate, very large membrane areas (often up to 60– quential passages through 30, 50, 70, 90%, and absolute ethanol. Next, 70% of the whole visible plasma membrane) were available for gold parti- the ethanol was substituted by liquid CO , and the samples were critical cles analysis, ensuring that the areas used for quantitation were truly rep- point dried. The formvar films were transferred from the glass onto copper resentative. One representative overview of DCs on fibronectin-coated grids, and the specimens were observed in a transmission electron micro- formvar is shown. Bar, 6 m. Online supplemental material is available at scope (model 1010; JEOL), operating at 60–80 kV. Gold particles were de- http://www.jcb.org/cgi/content/full/jcb.200306112/DC1. tected on the periphery and thinner parts of cells, where a good contrast could be achieved. In case of DCs, which widely spread (Fig. S1), the This work is dedicated to the loving memory of Hans Smits. membrane area available for analysis represented up to 60–70% of the whole labeled plasma membrane. For each cell at least four to six areas We thank Mietske Wijers and Huib Croes for providing gold beads and were analyzed at random. The digital images of electron micrographs were for their help with electron microscopy and W.J. de Grip for biochemistry processed by custom-written software based on Labview (National Instru- facility. We also acknowledge Rob Schuurman for his help with the p24 ments). The distribution pattern of DC-SIGN (i.e., the degree of clustering) ELISA. We are also grateful to Maria Garcia-Parajo, Gosse Adema, and was analyzed by counting the number of gold particles found on the Ruurd Torensma for critical reading of the manuscript. plasma membrane in a semi-automatic fashion. Subsequently, coordinates A. Cambi is supported by a grant SLW 33.302P from the Netherlands were assigned to the observed beads and interparticles distance statistics Organization of Scientific Research, Earth and Life Sciences. F. de Lange is were obtained using a nearest neighbor distance algorithm. Clusters were supported by a grant FB-N/T-1a from the Netherlands Foundation for Fun- DC-SIGN microdomains | Cambi et al. 155 damental Research of Matter. C.G. Figdor is supported by grant NWO A. Amara, C. Houles, F. Fieschi, O. Schwartz, J.L. Virelizier, et al. 2003. 901-10-092. 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Microdomains of the C-type lectin DC-SIGN are portals for virus entry into dendritic cells

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Copyright © 2004, The Rockefeller University Press
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10.1083/jcb.200306112
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Abstract

JCB Article Microdomains of the C-type lectin DC-SIGN are portals for virus entry into dendritic cells 1 1 4 4 1 Alessandra Cambi, Frank de Lange, Noortje M. van Maarseveen, Monique Nijhuis, Ben Joosten, 5 5 2 3 Erik M.H.P. van Dijk, Bärbel I. de Bakker, Jack A.M. Fransen, Petra H.M. Bovee-Geurts, 1 5 1 Frank N. van Leeuwen, Niek F. Van Hulst, and Carl G. Figdor 1 2 3 Department of Tumor Immunology, Department of Cell Biology, and Department of Medical Biochemistry, Nijmegen Center for Molecular Life Sciences, University Medical Center Nijmegen, 6500 HB Nijmegen, Netherlands Department of Virology, University Medical Center Utrecht, 3508 GA Utrecht, Netherlands Applied Optics Group, Department of Science and Technology, Molecular Engineering Sensors and Actuators Research Institute for Nanotechnology, University of Twente, 7500 AE Enschede, Netherlands he C-type lectin dendritic cell (DC)–specific inter- becomes organized in well-defined microdomains, with an cellular adhesion molecule grabbing non-integrin average diameter of 200 nm. Biochemical experiments (DC-SIGN; CD209) facilitates binding and internal- and confocal microscopy indicate that DC-SIGN micro- ization of several viruses, including HIV-1, on DCs, but the domains reside within lipid rafts. Finally, we show that the underlying mechanism for being such an efficient phagocytic organization of DC-SIGN in microdomains on the plasma pathogen-recognition receptor is poorly understood. By high membrane is important for binding and internalization resolution electron microscopy, we demonstrate a direct of virus particles, suggesting that these multimolecular relation between DC-SIGN function as viral receptor and assemblies of DC-SIGN act as a docking site for pathogens its microlocalization on the plasma membrane. During like HIV-1 to invade the host. development of human monocyte-derived DCs, DC-SIGN Introduction In the past two years, several new genes have been identified and Steinman, 1998). DCs are equipped with a variety of encoding leukocyte-specific carbohydrate binding proteins dynamically regulated pathogen-recognition receptors. Al- that belong to the lectin-like receptors family (Kogelberg though some of them are members of the toll-like receptor and Feizi, 2001; Figdor et al., 2002). Many of these lectins (TLR) family, signaling molecules specialized in sensing are members of the calcium-dependent C-type lectin family pathogens (Akira, 2003), others belong to the C-type lectin and recognize their ligands through the structurally related family and mediate pathogen binding and uptake (Stahl and Ca -dependent carbohydrate-recognition domains (C-type Ezekowitz, 1998; Mahnke et al., 2000). CRDs; Drickamer, 1999). Many C-type lectins act as cell DC-specific intercellular adhesion molecule (ICAM) grab- adhesion receptors (Vestweber and Blanks, 1999), whereas bing non-integrin (DC-SIGN; CD209) is a C-type lectin others are specialized in antigen recognition (Stahl and specifically expressed by DCs and has a dual function. As an Ezekowitz, 1998; Mahnke et al., 2000). adhesion receptor, DC-SIGN supports initial DC–T cell in- Dendritic cells (DCs) constitute a specific group of pro- teraction by binding to ICAM-3 (Geijtenbeek et al., 2000a), fessional antigen presenting leukocytes, constantly patrolling and mediates tethering and rolling of DCs on the endo- the body for foreign intruders (Steinman 1991; Banchereau thelium by interacting with ICAM-2 (Geijtenbeek et al., 2000c). As a pathogen-recognition receptor, DC-SIGN The online version of this article contains supplemental material. binds HIV gp120 thus facilitating the transport of HIV Address correspondence to Carl G. Figdor, Dept. of Tumor Immunology, from mucosal sites to draining lymph nodes where infection Nijmegen Center for Molecular Life Sciences, University Medical Center of T lymphocytes occurs (Geijtenbeek et al., 2000b). Recently, Nijmegen, P.O. Box 9101, 6500 HB Nijmegen, Netherlands. Tel.: 31- DC-SIGN was also shown to bind other viruses like CMV 24-361-7600. Fax: 31-24-354-0339. (Halary et al., 2002), Ebola (Alvarez et al., 2002), Dengue email: c.figdor@ncmls.kun.nl (Tassaneetrithep et al., 2003), and hepatitis C (Lozach et al., Key words: pathogen recognition receptor; lectin; electron microscopy; multiprotein assembly; lipid rafts 2003; Pöhlmann et al., 2003), as well as microorganisms  The Rockefeller University Press, 0021-9525/2004/01/145/11 $8.00 The Journal of Cell Biology, Volume 164, Number 1, January 5, 2004 145–155 http://www.jcb.org/cgi/doi/10.1083/jcb.200306112 145 The Journal of Cell Biology 146 The Journal of Cell Biology | Volume 164, Number 1, 2004 such as Leishmania (Colmenares et al., 2002), Candida albi- cans (Cambi et al., 2003), Mycobacterium (Geijtenbeek et al., 2003; Maeda et al., 2003; Tailleux et al., 2003), and Schisto- soma (van Die et al., 2003). Recent works have demonstrated that some microbial pathogens exploit cholesterol-enriched lipid microdomains as essential docking sites to enter host cells (Gatfield and Pieters, 2000; Rosenberger et al., 2000; Lafont et al., 2002). These microdomains, also known as lipid rafts, are localized regions with elevated cholesterol and glycosphingolipid con- tent that can be found on the plasma- and endosomal-mem- brane of eukaryotic cells (London and Brown, 2000; Simons and Toomre, 2000; Simons and Ehehalt, 2002). Some vi- ruses, such as HIV-1, appear to target lipid raft micro- domains during viral entry into cells, as well as during viral Figure 1. DC-SIGN is organized in microdomains on the cell surface of K-DC-SIGN. (A) The expression levels of DC-SIGN on assembly before budding from cells (Dimitrov, 1997; Mañes untransfected K562 and on K-DC-SIGN were assessed by FACS et al., 2000). Other works suggest that cholesterol-depen- analysis. The open histogram represents the isotype control, and dent membrane properties, rather than lipid rafts per se, are shaded histogram indicates the specific staining with anti–DC-SIGN responsible to promote efficient HIV-1 infection in T cells (AZN-D1). (B) DC-SIGN expressed by K562 transfectants strongly (Percherancier et al., 2003). binds to ICAM-3 and gp120. The adhesion was determined using DC-SIGN, like other C-type lectins, recognizes pathogens 1-m ligand-coated fluorescent beads. Specificity was determined by measuring binding in presence of AZN-D1. No blocking was by binding to carbohydrate moieties in a Ca -dependent observed in presence of isotype control (not depicted). Blocking manner, through a conserved CRD (Drickamer, 1999). exerted by EGTA indicates that DC-SIGN binds to the ligands in a This CRD has a high specificity for complex mannose resi- Ca -dependent manner. The average of five independent experi- dues and is located at the distal end of the extracellular do- ments is shown (P  0.001). (C) K-DC-SIGN cells were specifically main (ECD), which consists of several amino acid repeats labeled with 10-nm gold particles, as described in Materials and (Soilleux et al., 2000). Recently, purified truncated forms of methods. Cells were allowed to adhere onto poly-L-lysine–coated formvar film and photographed in an electron microscope. Gold DC-SIGN containing either the complete ECD or only the particles were detected on the periphery on the thinner less electron CRD were used to analyze the quaternary structure as well as dense parts of cells, where good contrast could be achieved. One the affinity of DC-SIGN for its ligands. Biochemical experi- representative picture is shown. Bar, 200 nm. ments indicated that in vitro ECDs aggregate to form tetra- mers, thus enhancing DC-SIGN capacity to bind multiva- lent ligands, such as pathogen sugar arrays (Mitchell et al., 1-m fluorescent beads coated with ICAM-3 or GP120 (Fig. 2001). Surface plasmon resonance experiments showed that 1 B). The binding of K-DC-SIGN to these ligands was spe- whereas the ECD readily binds hepatitis C virus glycopro- cific, as shown by the inhibition exerted by anti–DC-SIGN tein E2, distant monomeric CRDs do not, unless closely mAbs. Moreover, the lack of adhesion in presence of EGTA seeded (Lozach et al., 2003). confirmed that DC-SIGN bound to these ligands in a Ca - These findings suggest that the organization of DC-SIGN dependent manner, typical of C-type lectin-like receptors. molecules on the plasma membrane may be critical for To determine the organization of DC-SIGN on the cell pathogen binding. This prompted us to investigate the cell membrane at high resolution, TEM was used on whole- surface distribution pattern of DC-SIGN and its possible as- mount samples of K-DC-SIGN, after specific labeling with sociation with lipid rafts. Using transmission EM (TEM) on anti–DC-SIGN mAb and 10-nm gold particles. It should be whole-mount samples of transfected cells and monocyte- noted that the specimens were not sectioned thus making the derived DCs, we have mapped the microlocalization of whole cell surface available for gold labeling and subsequent DC-SIGN at high resolution. Subsequently, spatial-point TEM analysis. This method has been successfully applied to pattern analysis showed that DC-SIGN is organized in detect spatial distribution of other membrane proteins such well-defined microdomains. Moreover, biochemical experi- as the potassium channels Kv1.3 (Panyi et al., 2003) and the ments and confocal microscopy analysis demonstrated that IL-2 receptor -subunit (Vereb et al., 2000). As shown in DC-SIGN colocalizes with lipid rafts. Finally, we show that Fig. 1 C, DC-SIGN showed a clear distribution in well- these microdomains of DC-SIGN act as a docking site for defined microdomains on K-DC-SIGN plasma membrane. HIV-1 particles, facilitating entry of the virus into DCs. These microdomains had an average diameter of 200 nm and appeared to be randomly localized on the cell surface. To ex- clude that the gold labeling pattern observed could be due to Results internalized DC-SIGN molecules; also, sections of these cell DC-SIGN is organized in microdomains samples were analyzed by TEM. The results demonstrated on the cell surface that the gold particles were exclusively detected on the out- To study the correlation between functional state and cell side of the cell membrane (unpublished data), confirming the surface organization of DC-SIGN, we used K562 transfec- presence of DC-SIGN in microdomains on the cell surface. tants stably expressing DC-SIGN. K-DC-SIGN expressed To show that the clustering of DC-SIGN is not an artifact high levels of DC-SIGN (Fig. 1 A) and bound strongly to due to the procedure, we also analyzed by TEM other trans- DC-SIGN microdomains | Cambi et al. 147 membrane receptors transfected into K562, among which the 2-integrin LFA-1. Unlike DC-SIGN, LFA-1 molecules showed a random distribution pattern (unpublished data). It should be noted that the number of gold particles observed per micrometer squared was generally comparable with that observed for K-DC-SIGN, which should exclude difference in distribution due to major difference in the expression lev- els of the two receptors. DC-SIGN predominantly resides in lipid rafts To determine whether lipid rafts are important for DC- SIGN function, K-DC-SIGN cells were treated with methyl--cyclodextrin (MCD) to extract membrane choles- terol, and its effect on DC-SIGN–mediated ligand binding was tested by the fluorescent beads adhesion assay. As shown in Fig. 2 A, MCD treatment partially inhibits binding to gp120-coated beads, indicating that cholesterol extraction partially affected DC-SIGN ligand binding capacity. To further demonstrate the association of DC-SIGN with lipid rafts, K-DC-SIGN cells were solubilized in Triton X-100 on ice and fractionated by centrifugation on a sucrose gradient at 4C. With this procedure, lipid rafts, which can be isolated as detergent-resistant membranes (DRM), were recovered in low density fractions, whereas any other deter- gent-soluble material was concentrated in the high density fraction (Fig. 2 B). The GPI-anchored protein CD55 was used as the raft marker and was found in raft fractions, whereas the negative control (nonraft-associated protein), CD46, was almost completely detectable in the nonraft frac- tions. Similarly to CD55, a significant portion of DC-SIGN was recovered in the low density fraction, indicating that DC-SIGN is localized in DRM on the plasma membrane of K-DC-SIGN cells. To provide further evidence that DC-SIGN colocalizes with lipid rafts, we examined the co-distribution of DC- SIGN and the lipid raft marker ganglioside (GM1), on K-DC-SIGN by antibody patching and confocal micros- copy. As shown in Fig. 2 C, when capping was induced on K-DC-SIGN cells, GM1 clearly colocalizes with DC-SIGN and to the same extent as for CD55. However, some smaller patches of DC-SIGN were also detectable outside the lipid raft area, indicating that DC-SIGN might not permanently reside in lipid rafts, as also observed in Fig. 2 B. As expected, no colocalization of CD46 with GM1 was detected. Based on these observations, we conclude that DC-SIGN resides Figure 2. DC-SIGN colocalizes with lipid rafts on K-DC-SIGN. within a lipid raft environment on the plasma membrane of (A) To investigate the effect of cholesterol depletion on DC-SIGN– K-DC-SIGN cells. mediated adhesion, K-DC-SIGN cells were incubated in serum-free medium with or without 20 mM MCD for 30 min at 37C. Subse- DC-SIGN distribution changes during DC development quently, gp120-coated fluorescent beads (1-m diam) were added and the mixture was incubated for an additional 30 min at 37C. During the differentiation of DCs from monocyte precursors, Binding was measured by flow cytometry. After MCD treatment, cell the expression of DC-SIGN on the cell surface gradually in- viability was assessed by trypan blue staining. The values represent creases (Geijtenbeek et al., 2000a). However, as shown by the mean of three independent experiments SD. (B) K-DC-SIGN flow cytometry (Fig. 3 A), no significant increases in DC- were solubilized with 1% Triton X-100, subjected to sucrose gradient SIGN expression levels are seen between cells harvested after centrifugation and analyzed by Western blotting for the indicated molecules. The numbers indicate the gradient fractions. Fractions 9 3 d of culture (designated intermediate DCs) and immature and 10 are low density fractions containing DRM and are referred to as raft fractions. (C) Confocal microscopy analysis of copatching of DC-SIGN and GM1. K-DC-SIGN cells were stained at 4C with 10 g/ml anti–DC-SIGN (or anti-CD55 or anti-CD46) and 10 g/ml by confocal microscopy. Merged images are shown in the right panel. FITC-CTxB. Co-patching was induced by adding secondary Ab Results are representatives of multiple cells in three independent (Materials and methods), and, after fixation in PFA, cells were analyzed experiments. Bar, 5 m. 148 The Journal of Cell Biology | Volume 164, Number 1, 2004 Figure 3. DC-SIGN cell surface distribution during monocyte- derived DC development. DC-SIGN binding activity was monitored during development of monocyte-derived DCs. As shown in the box, intermediate DCs indicate cells harvested after 3 d of monocytes differentiation. (A) The expression levels of DC-SIGN on monocytes, intermediate and immature DCs were assessed by FACS analysis. The dotted line histogram represents the isotype control, and the thick line histogram indicates the specific staining with anti–DC-SIGN (AZN-D1). Mean fluorescence intensity is indicated. One represen- tative donor is shown. (B) The adhesion to ICAM-3 and gp120 was determined using 1 m ligand-coated fluorescent beads. Specificity was determined by measuring binding in presence of AZN-D1. No blocking was observed in presence of isotype control (not depicted). One representative experiments out of three is shown. (C) Intermediate and immature DCs were let adhere onto fibronectin-coated formvar film, specifically labeled for DC-SIGN with 10-nm gold particles (Materials and methods), and analyzed by TEM. Results are representatives of multiple cells in several independent experiments. Bar, 200 nm. DCs. Maximum DC-SIGN–mediated adhesion to ICAM-3 gold particles are evenly distributed over the cell surface, on as well as GP120 was observed on immature DCs, although immature DCs, there is a clear organization of DC-SIGN in already on intermediate DC DC-SIGN was capable of com- spatially well-defined microdomains. To exclude the possible pletely mediating the binding to ICAM-3 (Fig. 3 B, top), influence of the fibronectin substrate on DC-SIGN distribu- which on monocytes is LFA-1 dependent (unpublished data). tion, TEM analysis was also performed on DCs that were gold Comparably, while on monocytes, binding to gp120 is medi- labeled in suspension and mounted onto poly-L-lysine. No ated by CD4 (Kedzierska and Crowe, 2002; Kohler et al., differences were seen between cells stretched on fibronectin or 2003), on intermediate DCs, as well as on immature DCs, cells adhering to poly-L-lysine (unpublished data). Moreover, DC-SIGN is almost entirely responsible for binding to gp120 thin sections of resin-embedded immature DCs were gold la- (Fig. 3 B, bottom). To examine DC-SIGN cell surface distri- beled and analyzed by TEM. Clusters of DC-SIGN molecules bution, both intermediate and immature DCs were allowed to could be observed exclusively at the plasma membrane (un- adhere to fibronectin, and DC-SIGN molecules were labeled published data). We also analyzed by TEM the cell surface dis- with gold particles. Subsequently, the distribution on the tribution of other transmembrane receptors expressed on plasma membrane was analyzed by TEM (Fig. 3 C). Given DCs, including LFA-1. Unlike DC-SIGN, LFA-1 did not the high capacity of DCs to widely spread on the used sub- show any changes in cell surface distribution pattern on inter- strates, very large membrane areas (often up to 60–70% of the mediate and immature DCs (unpublished data). whole visible plasma membrane) were available for gold parti- To quantitatively describe the DC-SIGN distribution pat- cles analysis, ensuring that the areas used for quantitation were tern, the nearest neighbor distance values among the gold truly representative (Fig. S1, available at http://www.jcb. particles were calculated applying a spatial-point pattern org/cgi/content/full/jcb.200306112/DC1). Surprisingly, we analysis. A quantitative comparison of DC-SIGN clustering found that the distribution of DC-SIGN changes dramatically among intermediate DCs, immature DCs, and K-DC-SIGN during DC development. Although on intermediate DCs, the is shown in Fig. 4 A. On immature DCs, as well as on DC-SIGN microdomains | Cambi et al. 149 Figure 4. Quantitative analysis of the distribution of gold particles labeling DC-SIGN. The digital images of electron micrographs were processed by a custom-written software based on Labview. Gold labels were counted, and coordinates were assigned to each feature. Subsequently, interparticles distances were calculated using a nearest neighbor distance algorithm. Nearest neighbor distance values were calculated for each image, and the data of several independent experiments were pooled. Subse- quently, the nearest neighbor distances were divided into three classes 0–50 nm (gray bar), 50–150 nm (black bar), and 150 nm (white bar), and the percentage of nearest neighbor distance values falling into each class was plotted (A). The partitioning of gold labels in clusters of various size (i.e.: number of particles/cluster) was also quantified. Clusters were defined when gold particles were 50 nm apart from a neighboring particle. The percentage of gold particles involved in the formation of a certain cluster size was calculated for (B) K-DC-SIGN, (C) immature DC, and (D) intermediate DC. The insets are three representative processed digital images, where each type of cluster is shown in a different color. For K-DC-SIGN, one represen- tative experiment out of two is shown; for immature DC, one representative experiment out of six is shown; and for intermediate DCs, one representative experiment out of three is shown. K-DC-SIGN, almost 70 and 80% of the gold particles reside erage diameter of 200 nm and were separated from each within a 50-nm distance from its nearest neighbor, respec- other by an average distance of 400 nm. Spatial analysis of the tively. In contrast, on intermediate DCs, only 30% of the microdomains distribution over the cell surface did not reveal nearest neighbor distance values are found in the same cate- any preferential localization in specific surface areas of the ob- gory, and the majority of interparticle distances rather fall served cells. These data were also confirmed by using high res- within the 50–150 nm range. It has to be noted that the in- olution near-field fluorescence imaging (unpublished data). termediate and the immature DCs compared by spatial-point The relative partitioning of gold particles in clusters of pattern analysis had a similar amount of gold particles per various sizes (i.e., number of particles/cluster) was also quan- micrometer squared, confirming the comparable expression tified. A value of 50 nm was set to define the involvement of levels of DC-SIGN on the two cell types. a gold particle in a cluster. Fig. 4 D shows that on interme- Similarly to what was observed for K-DC-SIGN, on imma- diate DCs up to 80% of gold particles occur as single fea- ture DCs, these microdomains were also found to have an av- tures. In contrast, on immature DCs (Fig. 4 C), as well as on 150 The Journal of Cell Biology | Volume 164, Number 1, 2004 K-DC-SIGN (Fig. 4 B), DC-SIGN shows a much broader served on K-DC-SIGN, small patches of DC-SIGN were distribution of cluster sizes, with an average of 10–20 gold also found not colocalizing with GM1. To further prove the particles per cluster on immature DCs. colocalization of GM1 with DC-SIGN, we performed dou- ble gold labeling for DC-SIGN and GM1 on immature DCs, and analyzed the samples by TEM. As shown in Fig. 5 DC-SIGN resides in lipid rafts on DCs C, GM1 (10 nm gold) was found within DC-SIGN micro- Because, on K-DC-SIGN, we found a clear association of domains (5 nm gold). Consistent with observations by con- DC-SIGN with lipid rafts, we investigated whether DC- focal microscopy, some microdomains of DC-SIGN that SIGN also colocalized with lipid rafts on immature DCs. lacked GM1 were also detected. Together, these results Extraction of cholesterol by MCD resulted in a significant demonstrate that on immature DC DC-SIGN resides in mi- block in binding of both ICAM-3– and gp120-coated beads crodomains that to a significant extent are associated with a (Fig. 5 A). In contrast, DC-SIGN–mediated binding on in- lipid rafts environment. termediate DCs was not affected by cholesterol depletion (unpublished data). DC-SIGN in microdomains facilitates binding To further demonstrate an interaction of DC-SIGN with to virus-sized particles lipid rafts on immature DCs, we investigated the co-distri- bution of DC-SIGN and GM1 by confocal microscopy. Fig. We showed that fluorescent beads coated with DC-SIGN– 5 B shows that, when capping was induced, DC-SIGN specific ligands were bound by immature as well as interme- clearly colocalized with GM1, almost to the same extent as diate DCs, apparently independently of DC-SIGN organi- the raft marker CD55. However, similarly to what was ob- zation on the cell membrane (Fig. 3, A and B). Figure 5. DC-SIGN resides in lipid rafts on immature DCs. (A) DC-SIGN–mediated adhesion to ligand-coated fluorescent beads (1-m diam) on immature DCs was measured after cholesterol depletion by 20 mM MCD. The assay was performed as described in Fig. 2 A. Data shown are means  SD of one representative experiment performed in triplicate out of three. One representative experiment out of three is shown. (B) Confocal microscopy analysis of copatching of DC-SIGN and GM1 on immature DCs. Staining was performed as described in Fig. 2 C. Cells were analyzed by confocal microscopy. Merged images are shown in the right panel. Results are representative of multiple cells in two independent experiments. Bar, 5 m. (C) Whole-mount samples of immature DCs were double labeled for DC-SIGN (5 nm gold) and the raft marker GM1 (10 nm gold) and analyzed by TEM. Thin white arrows indicate GM1 colocalizing in DC-SIGN microdomains. Thick white arrows indicate DC-SIGN microdomains with no GM1. Black arrow indicates GM1 alone. Results are representatives of multiple cells in two independent experiments. Bar, 50 nm. DC-SIGN microdomains | Cambi et al. 151 However, because DC-SIGN on immature DCs is an ex- any microbeads, comparable to background levels or when quisite virus receptor and the fluorescent beads are at least DCs were pretreated with the carbohydrate mannan to one order of magnitude larger than viral particles, we inves- block DC-SIGN (Fig. 6, C and F). Similar results were ob- tigated whether clustering of DC-SIGN into higher order tained when microbeads coated with ICAM-3 were used assemblies might be needed to stabilize interactions with vi- (unpublished data). Moreover, DC-SIGN on intermediate rus-sized particles. To detect possible differences in “avidity” as well as on immature DCs was found to be equally capable for multimeric ligands between scattered and clustered DC- of binding to soluble recombinant gp120 (Fig. 6 G), sug- SIGN molecules, and to mimic virus binding to DC-SIGN, gesting that the intrinsic binding capacity of DC-SIGN we used fluorescent virus-sized microbeads of 40-nm diam, molecule on the two different DC types is comparable. Sim- coated with gp120, which are now referred to as virus-sized ilar results were obtained when soluble ICAM-3-Fc chime- particles. The extremely broad fluorescence emission peak of ras were used (unpublished data). All these observations sup- these microbeads currently precludes accurate quantification port our hypothesis that DC-SIGN molecules acquire a by flow cytometry, therefore binding of DCs to these virus- higher avidity for multimeric ligands when organized in sized particles was visualized by confocal microscopy. multimolecular assemblies. When comparing the capacity of intermediate and imma- DC-SIGN microdomains enhances HIV-1 infection ture DCs to bind these virus-sized particles (Fig. 6), a very high binding was observed on immature DCs. Many virus- Having established that DC-SIGN microdomains bind sized particles were bound to the plasma membrane and a virus-sized particles more efficiently in comparison to significant amount was phagocytosed (Fig. 6, D and E). In randomly distributed receptor molecules, we investigated contrast, intermediate DCs (Fig. 6, A and B) hardly bound whether the uptake of real virus was also enhanced by a clus- Figure 6. DC-SIGN microdomains on immature DCs bind virus-sized particles. Green fluorescent microbeads (40-nm diam) were coated with gp120 and added to the cells in a ratio of 20 beads/cell. The cells were incubated for 30 min at 37C, washed, and fixed in PFA. DC-SIGN was stained with AZN-D1 and Alexa 647– conjugated goat anti–mouse Ab (red). Subsequently, the cells were mounted onto poly-L-lysine–coated glass coverslips and analyzed by confocal microscopy. The overview of binding to gp120 micro- beads of (A) intermediate DCs and (D) immature DCs is shown. Two represen- tative cells of (B) intermediate and (E) immature DCs are shown. Specific block with 100 g/ml mannan was also per- formed by preincubating the cells at RT for 10 min before adding the microbeads (C and F); bars, 5 m. Similar results were obtained with ICAM-3–coated microbeads (not depicted). (G) Binding of K-DC-SIGN, intermediate DCs, and immature DCs to soluble gp120 was also performed: 50,000 cells were incubated with 50 mM biotinylated gp120 for 30 min on ice, in presence or absence of 20 g/ml anti–DC-SIGN blocking mAb (AZN-D1). A subsequent incubation with Alexa 488–conjugated streptavidin for 30 min on ice followed, and gp120 molecules bound to the cells were detected by flow cytometry. The values represent the mean of three independent experiments SD. 152 The Journal of Cell Biology | Volume 164, Number 1, 2004 lipid raft environment on DCs that forms highly organized well-defined (200 nm) multiprotein assemblies on the sur- face of a cell. To establish the association of proteins with lipid rafts is complicated and often controversial, which underscores the complexity of cell membranes. These lipid domains differ in their composition, physical properties, and biological func- tions (Schuck et al., 2003). Moreover, lipid raft composi- tions differ between cell types. Finally, the dynamics of the chemical membrane composition adds a further level of complexity. Therefore, the association of proteins with lipid Figure 7. Clustered DC-SIGN molecules efficiently bind HIV-1 rafts can only be analyzed using different techniques simul- particles and infect PBMC. DC-SIGN on immature DCs enhances HIV-1 infection as measured in a DC-PBMC coculture. Either inter- taneously. Our observation that DC-SIGN resides within mediate or immature DCs (1.5 10 ) were preincubated for 20 min lipid rafts is based on several well-established raft analysis at RT with or without blocking mAb against 20 g/ml DC-SIGN techniques. First, we showed by biochemical experiments (AZN-D1 and AZN-D2). Preincubated intermediate or immature DCs the partitioning of DC-SIGN in DRM. Second, co-patch- were pulsed for 2 h with HIV-1 (M-tropic HIV-1Ba-L strain), and ing of DC-SIGN with the lipid raft marker GM1 was unbound virus particles and mAb were washed away. Subsequently, observed by confocal microscopy. Third, this association DCs were cocultured with activated PBMC (1.5 10 ) for 7 d. Coculture supernatants were collected, and p24 antigen levels were between DC-SIGN and GM1 was confirmed by TEM. measured by ELISA. Black histogram represents PBMC infected in the Fourth, cholesterol extraction by MCD inhibited DC- absence of DCs. One representative experiment out of two is shown. SIGN–mediated binding. Surprisingly, on immature DCs, the same MCD treatment did not have any effect on the or- ganization of DC-SIGN microdomains, as observed by tered distribution of DC-SIGN, using p24 ELISA. There- TEM (unpublished data). This may be explained by the fact fore, intermediate and immature DCs were pulsed for 2 h that the remaining cholesterol may be sufficient to keep such with HIV-1 (M-tropic HIV-1Ba-L strain), washed, and cul- domains intact (Schuck et al., 2003). Moreover, besides tured in the presence of activated peripheral blood mononu- cholesterol, also glycosphingolipids, another lipid raft com- clear cells (PBMC). As shown in Fig. 7, virus replication was ponent, which is not depleted by MCD treatment, may in significantly higher when PBMC were cocultured with ei- part maintain the integrity of DC-SIGN microdomains. ther intermediate or immature DCs with respect to PBMC The fact that cholesterol extraction by MCD partially affects alone challenged with the same amount of infectious virus. DC-SIGN–mediated binding could be explained by consid- However, infection of PBMC cultured with virus-pulsed ering the pleiotropic effects that MCD can have on cell immature DCs was much higher than the infection exhib- functions, besides disrupting lipid rafts integrity (Brown and ited by PBMC cultured with intermediate DCs. In addition, London 2000; Edidin 2001; Schuck et al., 2003). There- the infection of PBMC with immature DCs was signifi- fore, MCD treatment can only be a preliminary indicator cantly blocked by anti–DC-SIGN mAbs that were added to for a possible association of a protein with a lipid rafts envi- DCs before incubation with HIV-1. In contrast, DC-SIGN ronment. Also, it should be added that, unlike for GPI- contribution to PBMC infection was clearly much lower anchored proteins, very little is known about how trans- when intermediate DCs were used. The incomplete block- membrane proteins are recruited into lipid rafts, how stable ing observed in the presence of anti–DC-SIGN Abs suggests these interactions are and what exactly the role of cholesterol the involvement of other HIV receptors also expressed on is. Further experiments are needed to better identify the mo- DCs (Turville et al., 2002). The comparable expression lev- lecular determinants that control the association of trans- els of DC-SIGN, as shown in Fig. 3 A, as well as several co- membrane proteins with lipid rafts. stimulatory molecules (such as CD40 and CD80) on inter- In vitro experiments with isolated recombinant C-type lec- mediate and immature DCs suggest that these two DC types tins suggested that these can oligomerize providing multiple have similar antigen presentation capacity (unpublished surfaces to bind (multivalent) ligands (Drickamer, 1999). data). Altogether, our data demonstrate that DC-SIGN or- DC-SIGN has been shown to form tetramers stabilized by an ganized in microdomains rather than randomly distributed -helical stalk (Mitchell et al., 2001), and purified truncated is more efficient in mediating binding and uptake of virus- forms of DC-SIGN containing the complete ECD were also sized microbeads as well as real HIV-1 particles. shown to be able to bind ligands by surface plasmon reso- nance (Lozach et al., 2003). In these works, the affinity of Discussion isolated CRDs or complete ECDs of DC-SIGN for the ligand was measured and compared with the affinity of the DC-SIGN is a DC-specific C-type lectin that acts both as adhesion molecule and as pathogen-recognition receptor. membrane-bound form. Although no interactions with the ligand could be found using isolated CRDs, significant bind- Here, we demonstrate that DC-SIGN can form micro- domains on the plasma membrane, and that there is a direct ing was detected if single CRDs were closely seeded (K nM) or if CRDs were part of a complete oligomeric-soluble correlation between the distribution of DC-SIGN in micro- domains and its capacity to bind and internalize virus-sized ECD of DC-SIGN (K 30 nM). The highest affinity was seen with membrane-bound DC-SIGN expressed on the sur- ligand-coated particles. Moreover, to the best of our knowl- edge, this is the first report describing a C-type lectin in a face of transfected cells (K 3 nM), suggesting that the nat- DC-SIGN microdomains | Cambi et al. 153 ural plasma membrane environment strongly influences the function of this receptor. To some extent, this mirrors our observations: DC-SIGN on the cell surface of intermediate DCs is randomly distributed and hardly binds virus-sized particles, whereas on immature DCs, the protein is organized in well-defined microdomains, which allows binding of 40- nm virus-sized particles (Fig. 6, A–F). In apparent contrast, we showed that on intermediate DCs, randomly distributed DC-SIGN was able to bind 1-m beads, coated with ICAM-3 or gp120 (Fig. 3 B). This apparent discrepancy between 40-nm virus-sized particles and 1-m beads can be explained Figure 8. DC-SIGN microdomains enhances binding of virus-sized by the fact that beads of 1-m diam have 600-fold larger particles with respect to isolated DC-SIGN molecules. Beads of interaction surface saturated with numerous ligand molecules 1-m diam are saturated with numerous coated ligand molecules that can engage simultaneous interactions with several indi- that can engage simultaneous interactions with several individual vidual DC-SIGN molecules. These multiple interactions DC-SIGN molecules. These multiple interactions may strengthen the binding both with random and clustered DC-SIGN. In contrast, when may signal into the cell and lead to a rapid recruitment of virus-sized particles are used, the contact surface and therefore, the new DC-SIGN molecules, thus strengthening the binding. number of ligand molecules is much smaller. Consequently, only In contrast, when virus-sized particles are used, the contact interactions with DC-SIGN molecules in highly organized multipro- surface and therefore, the number of ligand molecules is tein assemblies may result in stable binding of virus particles. much smaller. Consequently, only interactions with DC- SIGN molecules in highly organized multiprotein assemblies pathogen-associated molecular patterns displayed at the can result in stable binding of virus-sized particles (Fig. 8). cell surface of microorganisms. Although TLRs have been When a computer-aided simulation was performed to predict shown to induce differential gene expression upon recogni- the capacity of round objects of different sizes to establish in- tion of distinct pathogen structural components (Akira, teractions with either random or clustered DC-SIGN mole- 2003), for the C-type lectins, and particularly for DC- cules, we found that objects with a diameter in the range of SIGN, no direct signaling pathways has been demonstrated virus sizes (40–200-nm diam) preferentially bind to clusters so far. Therefore, it will be interesting to investigate the in- of DC-SIGN having the same size (unpublished data). In teractions that might occur between these two families of agreement with this model, no significant differences were pathogen-recognition receptors. detected in binding of intermediate and immature DCs to In conclusion, our findings emphasize the importance of soluble gp120 or to ICAM-3-Fc chimeras, indicating that the relating the function with the cell surface organization of capacity of each single DC-SIGN molecule to recognize and DC-SIGN. Clustered distribution is essential to enhance the bind the ligand is comparable on both DC types (Fig. 6 G). interaction as well as the internalization efficiency of DC- The biological relevance of this clustering phenomenon is SIGN–pathogen complexes. In addition, our data highlight shown by the fact that immature DCs compared with inter- the importance of the plasma membrane as a specialized mediate DCs showed an enhanced DC-SIGN–mediated microenvironment, where finely orchestrated interactions binding and internalization of virus particles, and subse- among proteins, carbohydrates, and lipids take place. These quent infection of PBMC in trans (Fig. 7). Therefore, we complex interactions mediate many fundamental processes propose that engagement of viruses with DC-SIGN occurs that occur at the contact site between cells (cell–cell or cell– much more productively when this receptor is organized in pathogen), such as assembly of signaling platforms and for- microdomains and resides in a lipid raft or cholesterol- mation of entry portals for invasive pathogens. enriched environment. Insight in the cell surface organization of pathogen It remains to be established what controls DC-SIGN ran- scavenger receptors like DC-SIGN will contribute to the dom distribution on intermediate DCs and what mecha- development of novel strategies that specifically inhibit nism is responsible for the change into a clustered organi- interactions with life-threatening pathogens like HIV-1 or zation on immature DCs. We cannot exclude that, on Mycobacterium. intermediate DCs, DC-SIGN may interact with an un- known membrane associated protein that prevents the for- mation of such microdomains. Alternatively, cytoskeletal Materials and methods constraints may differ between these cells, and this issue is Antibodies and reagents currently under investigation. Preliminary observations sug- For labeling as well as blocking of DC-SIGN, the mAb AZN-D1 was used gest that releasing DC-SIGN molecules from the cortical ac- (Geijtenbeek et al., 2000a). For detection of CD46 and CD55 the Abs E4.3 (BD Biosciences) and 143–30 (CLB) were used, respectively. Alexa 647– tin cytoskeleton by cytochalasin D treatment results in an conjugated goat anti–mouse IgG was purchased from Molecular Probes. enhanced ligand binding, particularly on intermediate DCs FITC-conjugated cholera toxin B subunit (FITC-CTxB), Triton X-100, and (unpublished data). Further experiments are needed to ob- MCD were purchased from Sigma-Aldrich. Goat anti-CTxB antibody was purchased from Calbiochem. tain insight into the formation of DC-SIGN multiprotein assemblies on DCs. Cells DC-SIGN belongs to the C-type lectin family that, to- Monocytes were obtained from buffy coats of healthy individuals and were gether with the TLR family, forms the first barrier against purified using Ficoll density centrifugation. Immature DCs were obtained invading pathogens. This occurs through recognition of as reported elsewhere (Geijtenbeek et al., 2000a). In brief, DCs were cul- 154 The Journal of Cell Biology | Volume 164, Number 1, 2004 tured from monocytes in presence of IL-4 and GM-CSF (500 and 800 defined when gold particles were less than a set distance apart from a U/ml, respectively) for 3 d to obtain intermediate DCs and for 6 d to obtain neighboring particle. immature DCs. Stable K562 transfectants expressing DC-SIGN (K-DC- SIGN) were generated as published previously (Geijtenbeek et al., 2000a). Confocal microscopy Cells were stained at 4C with 10 g/ml anti–DC-SIGN antibody (AZN- Adhesion assays D1), anti-CD55 antibody, anti-CD46 antibody, and 10 g/ml FITC-CTxB. Carboxylate-modified TransFluorSpheres (488/645 nm, 1-m diam; Mo- Isotype-specific controls were always included. Secondary staining for lecular Probes) were coated with ICAM-3-Fc or GP120, and the fluores- DC-SIGN, CD55, and CD46 was with Alexa 647–conjugated goat anti– cent beads adhesion assay was performed as described previously (Geij- mouse IgG, and for FITC-CTxB with goat anti-CTxB. Patching was induced tenbeek et al., 1999). When the lipid raft-disrupting agent MCD was used, by incubation at 12C for 1 h, followed by fixation with 1% PFA. Cells the cells were resuspended in serum-free medium containing the appropri- were mounted onto poly-L-lysine–coated glass coverslips. Signals were ate concentration of MCD and preincubated for 30 min at 37C. When collected sequentially to avoid bleed through. necessary, 50,000 cells were incubated with 20 g/ml blocking mAb for 10 min at RT. The ligand-coated fluorescent beads (20 beads/cell) were Detergent extraction and flotation assay added and the suspension was incubated for 30 min at 37C. Adhesion For detergent resistant membrane isolation, K-DC-SIGN cells (10 10 ) was determined by measuring the percentage of cells, which have bound were lysed on ice in 0.5 ml of lysis buffer (50 mM Tris, pH 7.4, 150 mM fluorescent beads, by flow cytometry using the FACSCalibur™ (Becton NaCl, 5 mM EDTA, and 1% Triton X-100, plus a cocktail of protease inhibi- Dickinson). Streptavidin-modified TransFluorSpheres (505/515 nm, 40-nm tors: 1 mM PMSF, 10 g/ml aprotinin, and 10 g/ml leupeptin). After 30 min diam; Molecular Probes) were coated with ICAM-3-Fc or GP120 as de- of incubation, the lysate was made 40% with respect to sucrose. Next, the scribed previously for 1-m beads (Geijtenbeek et al., 1999) and were lysate-sucrose mixture was overlaid with 2 ml of 30% sucrose in lysis buffer added to DCs in a ratio of 20 beads/cell. Bound beads were detected by and finally with 1 ml of 4% sucrose in lysis buffer. The mixture was centri- confocal microscopy analysis of the cells. fuged at 200,000 g for 14–16 h in an SW60Ti rotor (Beckman Coulter). The When binding to soluble GP120 was performed, 50 mM biotinylated gradient was fractionated in 0.5-ml fractions from the bottom of the tube. GP120 was added to the cells (10 ) for 30 min on ice, in presence or ab- sence of 20 g/ml anti–DC-SIGN blocking Ab (AZN-D1). After the incuba- Immunoblot analysis tion, cells were extensively washed and incubated with Alexa 488–con- Proteins from the sucrose fractions were separated by SDS-PAGE and trans- jugated streptavidin for 30 min on ice. Subsequently, bound GP120 ferred to PROTRAN nitrocellulose transfer membranes (Schleicher & molecules were detected by flow cytometry. Schuell). Membranes were blocked for 1 h at RT using 2% nonfat dried milk (Campina) and 1% BSA (Calbiochem) in 0.1% PBS/Tween 20. Blots were Labeling procedures subsequently incubated with specific antibodies (polyclonal anti-CD55 and For flow cytometry analysis, cells were incubated (30 min; 4C) in PBA CD46 from Santa Cruz Biotechnology, Inc., and a polyclonal anti–DC- (PBS, containing 0.5% BSA and 0.01% sodium azide), with different mAb SIGN antibody [CSRD] that was raised in our laboratory), followed by the (5 g/ml), followed by incubation with FITC-labeled goat anti–mouse IgG appropriate HRP-conjugated secondary antibodies (DakoCytomation). Fi- antibody (GAM-FITC; Zymed Laboratories; 1:50 dilution in PBA) for 30 nally, proteins were detected using ECL (Amersham Biosciences). min at 4C. The relative fluorescence intensity was measured on a FACS- Calibur™. Isotype–specific controls were included. For TEM, K-DC-SIGN HIV-1 infection of DCs L-lysine–coated formvar, whereas DCs were allowed were fixed onto poly- Infection of intermediate and immature DCs with the M-tropic strain HIV- to spread on glass coverslips covered by a thin layer of fibronectin-coated 1Ba-L was performed as described previously (Geijtenbeek et al., 2000b). formvar for 1 h at 37C and immediately fixed with 1% PFA for 15 min. Af- 6 ) were incubated with In brief, intermediate and immature DCs (1.5 10 ter two washing steps with PBS and a subsequent incubation (60 min at mAb against 20 g/ml DC-SIGN (AZN-D1 and 20 g/ml AZN-D2 for 20 RT) with I-buffer (PBS, 0.1% glycine, 1% BSA, and 0.25% gelatin) to re- min at RT. Subsequently, cells were incubated with the M-tropic strain duce a specific background, the specimens were incubated for 30 min HIV-1Ba-L for 2 h with a multiplicity of infection of 0.004. After incuba- with the primary antibodies in I-buffer on ice, rinsed in PBS, and fixed in tion, cells were washed and cocultured with activated PBMC in the pres- 1% PFA and 0.1% glutaraldehyde for 15 min. After two washing steps with ence of 5 U/ml human recombinant IL-2 (Roche Diagnostics). PBMC were PBS and a block in I-buffer, the samples were incubated with rabbit anti– activated by culturing them before infection for 3 d in the presence of 2 mouse IgG (to detect mAb) for 30 min on ice. A final incubation with 10- g/ml of phyto-HA (Sigma-Aldrich). After 7 d, culture supernatants were nm-diam gold-labeled Protein A (to detect polyclonal antibodies) was per- collected, and p24 antigen levels were determined by a p24 antigen ELISA formed, followed by final fixation in 1% glutaraldehyde in phosphate TM ; DakoCytomation). (AMPAK buffer for 20 min at RT. When double labeling was performed, the cells were treated as described above, except that anti–DC-SIGN (AZN-D1), Online supplemental material and biotinylated-CTxB were added simultaneously; and finally, goat anti– Fig. S1 is an overview of DC whole-mount samples by TEM. To determine mouse-conjugated 5-nm gold and streptavidin-conjugated 10-nm gold the organization of DC-SIGN on the cell membrane at high resolution, TEM were used to label DC-SIGN and GM1, respectively. Isotype-specific con- was used on whole-mount samples of intermediate as well as immature trols were always included. The 10-nm-diam gold-labeled protein A was a DCs, after specific labeling with anti–DC-SIGN mAb and 10-nm gold parti- gift of M. Wijers and H. Croes (Nijmegen Center for Molecular Life Sci- cles. It should be noted that the specimens were not sectioned, thus, mak- ences). The goat anti–mouse-conjugated 5-nm gold and streptavidin-con- ing the whole cell surface available for gold labeling and subsequent TEM jugated 10-nm gold were purchased from Aurion. analysis. DCs were adhered onto fibronectin-coated formvar film, fixed, and labeled as described in Materials and methods, and finally analyzed by EM and analysis of the distribution patterns of gold particles TEM. Given the high capacity of both intermediate and immature DCs to After gold labeling and fixation, the specimens were dehydrated by se- widely spread on the substrate, very large membrane areas (often up to 60– quential passages through 30, 50, 70, 90%, and absolute ethanol. Next, 70% of the whole visible plasma membrane) were available for gold parti- the ethanol was substituted by liquid CO , and the samples were critical cles analysis, ensuring that the areas used for quantitation were truly rep- point dried. The formvar films were transferred from the glass onto copper resentative. One representative overview of DCs on fibronectin-coated grids, and the specimens were observed in a transmission electron micro- formvar is shown. Bar, 6 m. Online supplemental material is available at scope (model 1010; JEOL), operating at 60–80 kV. Gold particles were de- http://www.jcb.org/cgi/content/full/jcb.200306112/DC1. tected on the periphery and thinner parts of cells, where a good contrast could be achieved. In case of DCs, which widely spread (Fig. S1), the This work is dedicated to the loving memory of Hans Smits. membrane area available for analysis represented up to 60–70% of the whole labeled plasma membrane. For each cell at least four to six areas We thank Mietske Wijers and Huib Croes for providing gold beads and were analyzed at random. The digital images of electron micrographs were for their help with electron microscopy and W.J. de Grip for biochemistry processed by custom-written software based on Labview (National Instru- facility. We also acknowledge Rob Schuurman for his help with the p24 ments). The distribution pattern of DC-SIGN (i.e., the degree of clustering) ELISA. We are also grateful to Maria Garcia-Parajo, Gosse Adema, and was analyzed by counting the number of gold particles found on the Ruurd Torensma for critical reading of the manuscript. plasma membrane in a semi-automatic fashion. Subsequently, coordinates A. Cambi is supported by a grant SLW 33.302P from the Netherlands were assigned to the observed beads and interparticles distance statistics Organization of Scientific Research, Earth and Life Sciences. F. de Lange is were obtained using a nearest neighbor distance algorithm. Clusters were supported by a grant FB-N/T-1a from the Netherlands Foundation for Fun- DC-SIGN microdomains | Cambi et al. 155 damental Research of Matter. C.G. Figdor is supported by grant NWO A. Amara, C. Houles, F. Fieschi, O. Schwartz, J.L. Virelizier, et al. 2003. 901-10-092. 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Published: Jan 5, 2004

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