DC-SIGN, but not sDC-SIGN, can modulate IL-2 production from PMA- and anti-CD3-stimulated primary human CD4 T cells

Osvaldo Martinez1, Scott Brackenridge2, Mohammed El-Azami El-Idrissi3 and Bellur S. Prabhakar1

1 Department of Microbiology and Immunology (M/C 790), University of Illinois at Chicago, Room E-709, Building 935, 835 South Wolcott Avenue, Chicago, IL 60612, USA
2 Rush University Medical Center, 1650 West Harrison Street, Chicago, IL 60612, USA
3 Unité de Biologie des Régulations Immunitaires, INSERM E 0352, Institut Pasteur, Paris 75724, France

Correspondence to: B. S. Prabhakar; E-mail: bprabhak{at}uic.edu


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Dendritic cell (DC)-specific intercellular cell adhesion molecule-3 (ICAM-3)-grabbing non-integrin (DC-SIGN) is expressed on the surface of DCs and specialized macrophages and can support T cell proliferation. Antibody-mediated co-ligation of CD3 and ICAM-3, the ligand for both DC-SIGN and leukocyte function-associated antigen-1, leads to T cell activation. Therefore, we tested to see whether DC-SIGN or a splice variant of dendritic cell-specific intercellular cell adhesion molecule-3-grabbing non-integrin (sDC-SIGN) can co-stimulate primary human T cells. The sDC-SIGN lacking the transmembrane domain encoded by exon 3 localizes to the cytoplasm of cells and is not secreted. Both B7 and DC-SIGN co-stimulated phorbol myristate acetate-stimulated CD4+ cells as compared with controls. However, unlike B7, both DC-SIGN and sDC-SIGN failed to co-stimulate CD4+ T cells treated with sub-optimal amounts of anti-CD3 (2 µg ml–1) as defined by a lack of CD69 and CD25 up-regulation, cell division and cytokine secretion. Instead, DC-SIGN, and not sDC-SIGN, induced a small but consistent down-regulation of IL-2 production by these CD4+ T cells. In contrast, DC-SIGN in the presence of 30 µg ml–1 of anti-CD3 modestly up-regulated cytokine production as compared with control. These results suggest that DC-SIGN can differentially modulate T cell stimulation.

Keywords: co-stimulation, dendritic cells, T lymphocytes, tolerance/suppression/anergy


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Antigen-independent formation of T cell and dendritic cell (DC) clusters (1) is mediated by the initial interactions between adhesion and co-stimulatory molecules. These interactions occur between molecules such as CD2–leukocyte function-associated antigen-3 (LFA-3), LFA-1–intercellular cell adhesion molecule 1 and 3 (ICAM-1 and -3) and CD28–CD80/86. However, more recent studies have suggested that it is the ICAM-3–dendritic cell-specific intercellular cell adhesion molecule-3-grabbing non-integrin (DC-SIGN) interaction that is critical for facilitating TCR–peptide–MHC interactions that lead to immune synapse formation (2, 3). Other adhesion and co-stimulatory molecules are also highly expressed on most resting cells but feature lower affinities (TCR–peptide–MHC), reduced size (CD2, TCR, CD28) or require prior activation to function efficiently (LFA-1). These properties render them less important for the initial intercellular adhesion between T cells and antigen-presenting cells (APCs).

ICAM-3 belongs to the Ig superfamily and is structurally similar to ICAM-1 and ICAM-2 but is highly expressed on all resting T cells and thus can facilitate initial interaction between a T cell and an APC. The ICAM-3 is re-distributed to the interface of T cells and DCs regardless of whether the interaction is antigen independent or dependent (2). Furthermore, ICAM-3-, and not LFA-1- or ICAM-1-, blocking antibodies can inhibit antigen-independent T cell–APC conjugate formation, while both LFA-1- and ICAM-3-blocking antibodies can prevent antigen-dependent conjugate formation. These and other studies showed that ICAM-3 could play an important role in the initial T cell–APC interaction. Anti-ICAM-3 antibody treatment of Jurkat T cells can elevate the levels of intracellular calcium and cause tyrosine phosphorylation (4) and T lymphoblast homotypic aggregation in an LFA-1–ICAM-1-dependent manner (5). Further, under conditions of sub-optimal TCR stimulation, anti-ICAM-3 antibody can up-regulate CD69 and CD25 expression (6, 7) and cause increased cell proliferation and IL-2 production (8). These studies indicate that ICAM-3 can provide critical signaling required for T cell stimulation. However, these studies used stimulating or blocking anti-ICAM-3 antibodies, instead of natural ligands.

In vivo, DC-SIGN is expressed at significant levels on subsets of APCs that include immature (CD83–) DCs, mature DCs, specialized macrophages in the lungs and placenta and a subset of DC precursors in the blood. The DC-SIGN is a type II membrane protein that can bind HIV gp120 protein (9) and sequester HIV (10) and other pathogens until they are presented to T cells. Further, DC-SIGN has been shown to bind ICAM-2, which is highly expressed on endothelial cells and thus may mediate DC trafficking (11).

Geijtenbeek et al. showed that blood-derived monocytes use LFA-1 to bind ICAM-3 on a T cell but monocyte-derived DCs, which express both LFA-1 and DC-SIGN, bind ICAM-3 mostly through DC-SIGN (3). Further, they demonstrated that an anti-DC-SIGN-blocking antibody could decrease allostimulation of human T cells by monocyte-derived DCs. These studies suggested that both DC-SIGN and LFA-1 could facilitate interactions between T cells and monocyte-derived DCs. However, whether DC-SIGN can co-stimulate ICAM-3-bearing cells or only facilitate initial T cell and APC adhesion is not known. A clearer understanding of its function would provide further insights into the mechanisms involved in T cell stimulation.

In this study, we cloned and expressed DC-SIGN, a natural ligand for ICAM-3, and a splice variant of dendritic cell-specific intercellular cell adhesion molecule-grabbing non-integrin (sDC-SIGN) and tested for their ability to provide co-stimulatory signals to human T cells.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Media and antibodies
DAP-3 and 293T cells were grown in DMEM medium (Invitrogen, Carlsbad, CA, USA) supplemented with antimycotic/antibiotic, ß-mercaptoethanol and 10% FCS (Mediatech, Herndon, VA, USA). CHO cells were grown in RPMI medium (Invitrogen) containing the above-mentioned supplements. Human PBMCs and T cells were cultured ex vivo in RPMI supplemented with 5% FCS. Stable cell lines transfected with various cDNAs were further supplemented with 250 µg ml–1 of G418 (Sigma–Aldrich, St Louis, MO, USA). Polyclonal anti-DC-SIGN antiserum was produced by immunizing rabbits with a peptide containing C-terminal 20 AAs (AAVGELPEKSKQQEIYQ) of the DC-SIGN coupled to keyhole limpet hemocyanin. Antibody response against the specific peptide was monitored using an ELISA. After three immunizations, the antibody titers reached ~1/200 000 and at that time the rabbits were bled and sacrificed. Antibodies against human antigens CD45RA (HI100), CD50 (TU41), CD69 (FN50), CCR7 (2H4), CD25 (M-A251) and CD4 (RPA-T4) were purchased from BD Pharmingen (San Diego, CA, USA). Anti-human CD45RO-ECD was purchased from Beckman Coulter Immunotech (Fullerton, CA, USA). Anti-DC-SIGN (MAB1621) was obtained from R&D Systems (Minneapolis, MN, USA).

PBMC and CD4+ T cell isolation
PBMCs were isolated using lympholyte-H reagent (Cedarlane, Vancouver, Canada) following the manufacturer's protocol. Briefly, 14 ml of peripheral blood from a healthy donor was mixed with 20 ml of PBS and was under layered with 9 ml of lympholyte-H. After centrifugation at 750 x g for 20 min at room temperature, the ‘buffy layer’ was harvested and washed with PBS. PBMCs were used directly for co-stimulation assays and monocyte differentiation or further processed to isolate CD4+ T cells.

CD4+ T cells were isolated by negative selection using CD4+ T Cell Isolation Kit (Miltenyi Biotec, Auburn, CA, USA), following manufacturer's protocol. Non-CD4+ T cells that were bound to beads were subjected to an automacs separation using the ‘deplete’ program. Over 95% of the live enriched cells were CD4+.

Monocyte-derived DCs
Isolated PBMCs (5–10 x 106 cells per well) were plated into six-well dishes in RPMI containing 5% FCS and incubated at 37°C for 2 h. Wells were gently washed three times with PBS to eliminate non-adherent cells and the remaining cells were cultured in RPMI supplemented with granulocyte macrophage colony-stimulating factor (GM-CSF; 800 U ml–1; PeproTech, Rockyhill, NJ, USA) and IL-4 (400 U ml–1, PeproTech). Cultures were replenished with fresh GM-CSF- and IL-4-supplemented medium every 3 days. To induce differentiation, monocyte-derived immature DCs were treated with LPS (10 µg ml–1, L2654), human IFN-{alpha} (1000 U ml–1, I-2396) or human tumor necrosis factor {alpha} (10 µg ml–1, T-0157) (Sigma–Aldrich) for 24 h.

DC-SIGN and sDC-SIGN cloning and expression in CHO cells
cDNAs encoding DC-SIGN and sDC-SIGN were obtained by reverse transcription–PCR using RNA isolated from monocyte-derived immature DCs as a template and F1-AGAGTGGGGTGACATGAGTG and B1-GAAGTTCTGCTACGCAGGAG primers (3). Amplified products were cloned, sequenced and sub-cloned into the pIRES-2 eGFP vector (Clontech, Palo Alto, CA, USA), which was used to establish permanently transfected cells. Briefly, CHO, DAP-3 or 293 T cells were seeded (2 x 105 cells per well in a six-well plate) the day before transfection. Two micrograms of DNA was mixed with 8 µl of Superfect reagent (Qiagen, Valencia, CA, USA) at room temperature for 5 min in 150 µl of serum-free medium and added to cells in a total volume of 1 ml. After 2 h of incubation, cells were washed twice with PBS and allowed to grow for 2 days. Permanently transfected cells were established by culturing cells in the presence of G418 (1 mg ml–1) for at least 1 week. Cells expressing high levels of protein were obtained by sorting on the basis of green fluorescent protein (GFP) expression.

Co-stimulation assays
Human PBMCs or CD4+ T cells were mixed with irradiated CHO cells (8000 rads of gamma irradiation) at a ratio of 3.5 : 1 in 96-well flat-bottom plates and grown in 5% RPMI at 37°C and 5% CO2. Two separate anti-CD3 antibodies were used for T cell stimulation. For the first set of experiments, flat-bottom 96-well plates were coated with 2 µg ml–1 (sub-optimal) or 30 µg ml–1 (large dose) of anti-CD3 antibody (UCHT1, Pharmingen), incubated overnight at 4°C and gently washed once with PBS before cell mixtures were added. For the second set of experiments, soluble stimulating anti-CD3 antibody (10 ng ml–1 of HIT3a, Pharmingen) or phorbol myristate acetate (PMA) (4 ng ml–1) was added to the cell mixtures. Cells and culture supernatants harvested at 24 h were used to determine the levels of expression of CD69 and CD25 and cytokines. Cells harvested on day 3 or 4 were used to determine the cell division.

Cell division
Cells were washed twice with PBS and then incubated in the dark (107 cells ml–1) with 0.250 µM of 5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester (CFSE; Molecular Probes, Eugene, OR, USA) in PBS at 37°C for 10 min. To quench the reaction, cells were diluted with PBS containing 10% FCS and incubated for an additional 20 min. These cells were washed with PBS containing 2% FCS and used in the co-stimulation assays. Cell division was determined by the extent of CFSE dilution detected using a flow cytometer (FACSCalibur; BD, San Jose, CA, USA) followed by data analysis using Win MDI 2.8 software (http://facs.scripps.edu/software.html). Live lymphocyte-gated cells were assessed for forward scatter (FSC) and green (FL1) fluorescence properties. Quadrants were set up using live CFSE-loaded untreated T cells as negative controls for cell size (FSC) and cell division (FL1 channel for CFSE).

Cell staining
Lymphocytes were re-suspended separately from triplicate wells gently so as to harvest only the non-adherent lymphocytes. CHO cells inadvertently carried over were gated out by size exclusion. Lymphocytes were incubated for 10 min on ice with Fc-block reagent (Pharmingen) in PBS + 2% FCS. All subsequent staining was carried out on ice for 30 min in the dark using 50 µl per reaction of antibodies (5 µg ml–1) in PBS + 2% FCS. The CD4+ lymphocytes were stained with FITC-CD69 (Pharmingen) or CD25 (Pharmingen) and then with 4',6-diamidino-2-phenylindole (DAPI) (Sigma–Aldrich), Cy-chrome CD45RA and CD45RO-ECD. CCR7 staining was performed using a mouse anti-CCR7 antibody (Pharmingen) followed by the addition of biotinylated rat anti-mouse IgM (R6-60.2, Pharmingen) and streptavidin–PE (Pharmingen). Cells were washed once with 200 µl of PBS + 2% FCS. DAPI-negative cells were analyzed by FACS to determine the expression levels of various markers. The results were analyzed using Win MDI software (http://facs.scripps.edu/software.html).

ELISA for cytokines
Supernatants, harvested from co-stimulation assays performed in triplicate, were frozen at –80°C until assayed. A volume of 50 µl of supernatant was used to assay for IL-2, IL-4 and IFN-{gamma} as outlined in the manufacturer's instructions (IL-4 and IFN-{gamma}: eBioscience, San Diego, CA, USA; IL-2: OptEIA human IL-2 kit, Pharmingen). ACH8122 and biotinylated AHC7129 (Biosource, Camarillo, CA, USA) antibodies were used to capture and detect IL-12, respectively. MAXI-SORP ELISA plates (439 454; Nunc, Rochester, NY, USA) were coated with ACH8122 antibody in (50 µl per well) carbonate buffer (0.1 M Na2CO4/NaHCO4, pH = 9.0) overnight at 4°C. Next day plates were washed three times with PBS + 0.05% Tween-20 and then blocked for 1 h with 100 µl PBS + 10% FCS at room temperature. Plates were washed extensively between the following steps. Fifty microliters per well of culture supernatant was added in triplicate and incubated for 2 h, biotinylated AHC7129 antibody was added and incubated for 1 h and streptavidin–HRP was added and incubated for 30 min; the reaction was developed using 50 µl of tetramethylbenzidine (TMB) substrate (BD) for 15 min and stopped with 25 µl of 1 M HCl. Plates were read at 450 nm using a Bio-Rad 550 plate reader (Bio-Rad, Hercules, CA, USA) and then analyzed using Excel software (Microsoft, Redmond, WA, USA).

DC-SIGN ELISA
DAP-3 and 293T cells stably expressing DC-SIGN or sDC-SIGN were grown to confluence in six-well dishes containing 3 ml of medium. Culture supernatants were tested in triplicate for the presence of soluble DC-SIGN using a sandwich ELISA. A polyclonal rabbit antiserum against the C-terminal 20 AAs of DC-SIGN was used as the capture antibody. For detection, monoclonal anti-DC-SIGN antibody (MAB1621, R&D Systems) was used at 1 µg ml–1 in PBS + 10% FCS followed by the addition of HRP-labeled goat anti-mouse IgG2a polyclonal antibody (Caltag, Burlingame, CA, USA) and TMB substrate as outlined above. DAP-3 sDC-SIGN cell lysate was used as a positive control.

Western blot
293T, sDC-SIGN-293T and DC-SIGN-293T cells (0.8 x 106) were washed and re-suspended in 30 µl of 1 x boiling protein sample buffer supplemented with ß-mercaptoethanol. Samples were boiled for 5 min and subjected to SDSP on a 10% gel (Bio-Rad) using a commercial electrophoresis buffer (161-0732, Bio-Rad). Proteins were then transferred to a nitrocellulose membrane (162-0115, Bio-Rad) using a mini-gel transfer apparatus (Bio-Rad) overnight at 4°C at 30 V in transfer buffer (3 g Tris l–1, 14.4 g glycine l–1 and 20% methanol). The nitrocellulose membrane was blocked with 2% skim milk in PBS (milk/PBS) for 2 h at room temperature. Rabbit anti-DC-SIGN antibody was added (20 µg ml–1 in 1% milk/PBS) and incubated for 2 h at room temperature with gentle rocking. The membrane was washed three times with PBS + 0.05% Tween-20 for 5 min each. The secondary HRP-labeled anti-rabbit (Caltag) antibody was added at 1/500 in 1% milk/PBS and washed as before. Western blotting detection system (RPN2132; Amersham Bioscience, Piscataway, NJ, USA) was used following the manufacturer's instructions.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
sDC-SIGN is not secreted and resides in the cytoplasm of cells
We characterized sDC-SIGN which was identified on PCR amplification of DC-SIGN from human cells. Although previously identified (12), it had not been characterized. This isoform lacks the putative transmembrane region encoded by exon 3 of DC-SIGN (Fig. 1A–C) and relative to DC-SIGN it was expressed at very low levels in the IL-4- and GM-CSF-treated PBMCs of three different individuals (Fig. 1B). Both DC-SIGN and sDC-SIGN were stably expressed, and comparison of sDC-SIGN and DC-SIGN protein expression in 293T cells (Fig. 1D–F), CHO (Fig. 3) and DAP-3 cells (not shown) demonstrated that sDC-SIGN is not expressed on the cell surface. This was not due to a lack of protein expression since sDC-SIGN could be readily detected on a western blot prepared using lysates of 293T cells (Fig. 1E). Further, anti-DC-SIGN antibodies were able to stain permeabilized 293T cells expressing sDC-SIGN (Fig. 1F).



View larger version (33K):
[in this window]
[in a new window]
 
Fig. 1. sDC-SIGN, an isoform of DC-SIGN, is not expressed on the cell surface. (A) Schematic representation of sDC-SIGN compared with DC-SIGN. Both molecules are identical; however, sDC-SIGN is missing exon 3, the putative transmembrane domain of the protein. (B) Using the same DC-SIGN primers, reverse transcription–PCR was performed from GM-CSF + IL-4-treated PBMC-RNA from three different individuals (lanes 1, 2 and 3) and PCR from plasmids containing sDC-SIGN (lane 4) and DC-SIGN (lane 5). Top arrow points to DC-SIGN, while bottom arrow points to sDC-SIGN. (C) Sequence difference between sDC-SIGN and DC-SIGN is highlighted in gray (i.e. exon 3). Putative splicing signals are indicated by arrows. (D) DC-SIGN and sDC-SIGN were cloned into an MCS-IRES-GFP vector where transcripts expressing (s)DC-SIGN also expressed GFP. 293T cells transfected with DC-SIGN and sDC-SIGN express GFP (upper histograms). Staining for cell surface DC-SIGN demonstrates that sDC-SIGN is not expressed on the cell surface (bottom histograms) of transfected 293T cells gated on GFP expression. However, sDC-SIGN is expressed inside the cell as revealed by western blot (E) and by staining against DC-SIGN in permeabilized sDC-SIGN-transfected 293T cells (F). Normal 293T cells were used as control in the permeabilized 293T cell experiment (light histogram in F).

 


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 3. Stable expression of co-stimulatory molecules in CHO cells used in co-stimulation assays. CHO cells were stably transfected with the MCS-IRES-GFP vector, sDC-SIGN-IRES-GFP vector, DC-SIGN-IRES-GFP vector and with vectors expressing CD80 or CD86. Expression levels of GFP were comparable for the control, sDC-SIGN and CHO-DC-SIGN cells. However, only CHO-DC-SIGN cells expressed cell surface DC-SIGN. CHO-CD80 cells expressed CD80 but not CD86 (negative control for CD86 cells) and CHO-CD86 cells expressed CD86 but not CD80 (negative control for the CD80-expressing CHO cells).

 
Since sDC-SIGN could be found in the cytoplasm of stably expressing cells, we tested whether it was secreted into the medium. Supernatants from cultures of cells stably expressing sDC-SIGN and from both immature and stimulated monocyte-derived DCs were tested for the presence of sDC-SIGN using a newly developed DC-SIGN ELISA. No sDC-SIGN was found in the supernatant of confluent DAP-3 (Fig. 2A) or 293T cells (not shown) or CHO cells (not shown) after 3 days of culture. However, the protein was readily detected in the diluted lysates of sDC-SIGN-expressing DAP-3 cells (Fig. 2A). Similarly, sDC-SIGN was also not detected in the culture supernatants of immature and stimulated DCs (Fig. 2B). This was not due to a problem in the ability of DCs to secrete soluble factors since LPS-treated DCs secreted IL-12 (Fig. 2C). Taken together, these data suggest that sDC-SIGN is a soluble but non-secreted form of DC-SIGN localized to the cytoplasm of cells whose function remains unknown. This allowed us to use sDC-SIGN as a non-membrane-expressed control for DC-SIGN in co-stimulation assays.



View larger version (12K):
[in this window]
[in a new window]
 
Fig. 2. sDC-SIGN is a soluble protein that is not secreted by monocyte-derived DCs. (A) An ELISA for sDC-SIGN was developed and tested. DAP (mouse fibroblast) cells stably transfected with either sDC-SIGN or DC-SIGN were lysed by the freeze–thaw method. Cleared lysate or supernatant from day 3 confluent cell cultures were tested for expression of sDC-SIGN. Only lysates from cells transfected with sDC-SIGN released sDC-SIGN. DC-SIGN anchored in the membrane was not released from freeze–thawed DAP cells. (B) Supernatants from day 7 blood-derived immature DCs (no stim) or LPS-, tumor necrosis factor {alpha} (TNF-{alpha})- or IFN-{alpha}-stimulated DCs were tested for sDC-SIGN and were found to be negative. (C) The DC supernatants described above (B) were also tested for IL-12 secretion, which was found in LPS-treated immature DCs.

 
DC-SIGN co-stimulates PMA and higher concentrations of anti-CD3—but not sub-optimal concentrations of anti-CD3—induced IL-2 production by human CD4+ T cells
T cell activation requires a primary signal through the TCR and co-stimulation from accessory molecules found on the surface of APCs. The most important of the co-signaling molecules is the B7 family members CD80 and CD86 that ligate constitutively expressed CD28 on T cells. Accessory molecule, ICAM-3, expressed on all resting CD45RA+RO– (naive) and CD45RO+RA– (memory) CD4+ T cells (our unpublished observations) (13, 14) can also provide co-stimulation but to a lesser degree than CD28. Anti-CD28 (10 µg ml–1) treatment of anti-CD3 (2 µg ml–1)-stimulated CD4+ T cells increased proliferation by at least 5-fold, while anti-ICAM-3 (10 µg ml–1 of HP2/19 antibody) treatment increased T cell proliferation by 2-fold (our unpublished observations) (5) as compared with T cells treated only with this limiting amount (2 µg ml–1) of stimulating anti-CD3 antibody. However, unlike stimulating anti-ICAM-3 antibody HP2/19, other anti-ICAM-3 antibodies (2), such as 186-2G9, cannot co-stimulate T cell stimulation when used in the presence of the same anti-CD3 antibody (not shown). This suggested that simply binding to ICAM-3 on T cells is not sufficient and may require specific interaction with ICAM-3, to provide co-stimulation.

Since DC-SIGN is a physiological ligand for ICAM-3, a classical co-stimulation assay was used to test for the ability of DC-SIGN and sDC-SIGN to provide co-stimulation. DC-SIGN and sDC-SIGN were sub-cloned into the pIRES-GFP2 vector and sorted on the basis of equivalent levels of GFP expression. DC-SIGN expression levels in stably transfected CHO cells used in the co-stimulation assays are shown in Fig. 3. Human PBMCs were stimulated with a sub-optimal dose of plate-bound UCHT1 anti-human CD3 (2 µg ml–1) antibodies in the presence of CHO cells expressing either DC-SIGN or sDC-SIGN or B7 molecules, CD80 and CD86 (shown in Fig. 3). As a test for T cell stimulation we assayed the supernatants for the production of IL-2. Both DC-SIGN and sDC-SIGN could not co-stimulate IL-2 production, while B7 molecules were able to co-stimulate a 5-fold increase in IL-2 production relative to PBMCs stimulated with anti-CD3 alone (not shown). Interestingly, DC-SIGN, but not sDC-SIGN, consistently inhibited anti-CD3-induced IL-2 production by PBMCs, relative to controls (not shown). In order to better characterize the T cell response in this system, CD4+ T cells were isolated via negative selection to maintain them in resting state and used in subsequent assays.

We performed the co-stimulation assay using PMA (Fig. 4A), which activates tyrosine kinases and the high-affinity form of LFA-1. We confirmed that CD80 and CD86 could co-stimulate cytokine production from PMA-treated T cells. To a lesser extent, but consistently, DC-SIGN but not sDC-SIGN could up-regulate IL-2 production as compared with controls. The enhancement of IL-2 production by DC-SIGN in PMA-stimulated T cells represented >40% of the levels of IL-2 production by T cells when they were treated with PMA and ionomycin.



View larger version (21K):
[in this window]
[in a new window]
 
Fig. 4. DC-SIGN can modulate IL-2 production. (A) Supernatants collected from CD4 T cells stimulated with PMA in the presence of either control CHO cells or CHO cells expressing CD80, CD86, sDC-SIGN or DC-SIGN were tested for the presence of IL-2 and IFN-{gamma} 24 h after stimulation. The B7 molecules induced high levels of IL-2 and IFN-{gamma}. Similarly, DC-SIGN was able to increase IL-2 production relative to control GFP and sDC-SIGN by ~2-fold with a P value of 0.014. Cells treated with PMA and ionomycin were included as a positive control. (B) Supernatants collected from CD4+ T cells stimulated with a sub-optimal amount of anti-CD3 (2 µg ml–1) in the presence of above-mentioned co-stimulatory molecules were tested. CHO cells expressing B7 molecules were able to stimulate significant cytokine production. In contrast, CHO-DC-SIGN suppressed IL-2 production relative to control CHO cells (P = 0.0017). (C) Supernatants collected from CD4+ T cells stimulated with a non-limiting amount of anti-CD3 (30 µg ml–1) in the presence of the CHO cells-expressing vector and DC-SIGN were tested 24 h after stimulation for the presence of IL-2 and IFN-{gamma}. DC-SIGN significantly enhanced both IFN-{gamma} (P = 0.0004) and IL-2 (P = 0.0031) production as compared with GFP controls.

 
We then tested supernatants of anti-CD3 (2 µg ml–1)-stimulated CD4+ T cells for IL-2, IL-4 and IFN-{gamma} 24 h after stimulation. Only CHO cells expressing CD80 and CD86 significantly up-regulated cytokine production (minimum of 5-fold increase over GFP controls). Consistently, there was a modest but significant (P = 0.0017) inhibition of IL-2 in the presence of DC-SIGN as compared with control (100 µg ml–1 compared with 400 µg ml–1, respectively, Fig. 4B). This assay was also repeated with a different stimulatory anti-CD3 antibody (HIT3a anti-human CD3 antibody) with similar results (not shown). The level of IL-4 was very low (<10 pg ml–1) at the limit of detection and therefore is not shown.

Since DC-SIGN could provide limited co-stimulation to PMA-stimulated T cells, we wanted to determine whether a similar enhancement could be seen when T cells were stimulated with a non-limiting concentration of anti-CD3 (30 µg ml–1) antibody. Relative to the control, there was a moderate but significant increase in the production of IL-2 and IFN-{gamma} when T cells were stimulated in the presence of DC-SIGN (Fig. 4C).

We further explored the effects of DC-SIGN on weakly stimulated T cells. Previously, it was shown that anti-ICAM-3 treatment of anti-CD3-stimulated T cells could induce expression of activation markers CD69 and CD25. Therefore, we tested for the ability of DC-SIGN or sDC-SIGN to up-regulate these activation markers on T cells stimulated with sub-optimal anti-CD3 or PMA (Fig. 5). CD80- and CD86-expressing CHO cells induced higher levels of CD69 and CD25 expression on anti-CD3-stimulated T cells when compared with control CHO cells (CD69: 59 and 44%, respectively, compared with 23%; CD25: 17% each compared with 8% for the control). A modest suppression of CD69 up-regulation was seen in the presence of CHO cells expressing DC-SIGN relative to control CHO cells or cells expressing sDC-SIGN (9% compared with 23 and 22%, respectively).



View larger version (48K):
[in this window]
[in a new window]
 
Fig. 5. DC-SIGN, but not sDC-SIGN, partly inhibits the up-regulation of early activation marker CD69. Human CD4+ T cells were stimulated with PMA or sub-optimal doses of anti-CD3 and tested 24 h later for the up-regulation of CD69, CD25 (IL-2R) and expression of chemokine receptor CCR7. First row represents untreated CD4+ T cells. T cells were treated with either anti-CD3 or PMA in the presence of CHO-GFP (second row), CHO-sDC-SIGN (third row), CHO-DC-SIGN (fourth row), CHO-CD80 (fifth row) and CHO-CD86 (sixth row). Depicted in the dot plots are the percentages of cells that show CD69 or CD25 up-regulation as compared with the controls shown in the first row.

 
The levels of CD69 were higher on all T cells treated with PMA irrespective of co-stimulation. The levels of CD25, however, were up-regulated on T cells co-stimulated with CD80 and CD86 relative to controls (CD25: 77 and 75%, respectively, compared with 45%). When cells were further sorted into CD45RA+RO– (naive) and CD45R+RA– (memory), we found that CD80 and CD86 could co-stimulate both populations (not shown).

Next, we tested the abilities of DC-SIGN and sDC-SIGN to co-stimulate plate-bound anti-CD3- or PMA-induced division of Th. CD4+ T cells were loaded with CFSE dye that dilutes on cell division. T cells were analyzed by flow cytometry 3 days post-stimulation for levels of CFSE and forward scatter, which is a crude measure of cell size (Fig. 6). In the anti-CD3 stimulation assay, both CD80 and CD86 could induce blast formation (77 and 61% of T cells compared with 11% in the presence of GFP control) and cell proliferation (53 and 20%, respectively, compared with 1% for GFP control) within 3 days. However, GFP vector control, DC-SIGN and sDC-SIGN could not induce any cell division. By day 3, PMA-treated T cells underwent significant cell division in the presence of CD80 (44%) and CD86 (30%) relative to 5% in the presence of CHO-GFP. Cell division in the presence of DC-SIGN and sDC-SIGN control was 10 and 4%, respectively. Non-dividing CD4+ T cells co-stimulated with GFP, sDC-SIGN, DC-SIGN, CD80 and CD86, nonetheless showed blast formation in 55, 54, 67, 54 and 54% of cells, respectively.



View larger version (24K):
[in this window]
[in a new window]
 
Fig. 6. Anti-CD3- and PMA-mediated cell division co-stimulated by CHO cells expressing GFP, sDC-SIGN, DC-SIGN, CD80 and CD86. Purified human CD4+ cells were loaded with the dye CFSE that binds to intracellular proteins and is diluted from the cell after cell division. CD4 T cells were sub-optimally stimulated with plate-bound anti-CD3 (left panel) or PMA (right panel) alone or in the presence of CHO cells expressing GFP, sDC-SIGN, DC-SIGN, CD80 and CD86. Cells were then tested for expression levels of CFSE and forward scatter (crude measure of cell size) 3 and 4 days after anti-CD3 and PMA treatment, respectively. Division is then assessed as a decrease in CFSE. Note that PMA can induce cells to become larger (39%) by day 3 but they cannot divide, while anti-CD3 treatment alone had no effect on cells. B7 molecules provided strong co-stimulation for cell division as evidenced by CFSE dilution in 53 and 20% of anti-CD3-treated and 44 and 30% of PMA-treated cells in the presence of CD80 and CD86, respectively.

 
DC-SIGN can modulate CD80 co-stimulated T cell activation and IL-2 production
Prior experiments showed that DC-SIGN could not co-stimulate CD4+ T cells treated with sub-optimal amounts of anti-CD3. However, we found that treatment of T cells with anti-CD28 and anti-CD3 in the presence of DC-SIGN caused lower IL-2 production (not shown). Therefore, we tested the effects of DC-SIGN on CD80-mediated co-stimulation, which can occur in trans, i.e. the ligands for TCR and CD28 can be present on two different cells.

Different ratios of CHO-DC-SIGN to CHO-CD80 (10 : 1 and 1 : 1) were used to co-stimulate anti-CD3-treated Th. We tested for IL-2 secretion and activation marker up-regulation and compared them with T cells co-activated with the same ratios of CHO-vector and CHO-CD80 cells. When CHO-DC-SIGN : CHO-CD80 or CHO-GFP : CHO-CD80 cells were used at a ratio of 10 : 1, they induced 461 and 1049 pg ml–1 of IL-2 and 550 and 1100 pg ml–1 of IFN-{gamma}, respectively. The effects of DC-SIGN were somewhat subdued when they were used at a 1 : 1 ratio (Fig. 7).



View larger version (14K):
[in this window]
[in a new window]
 
Fig. 7. DC-SIGN can interfere with IL-2 and IFN-{gamma} secretion from T cells stimulated by anti-CD3 and CHO-CD80. Ratios of CHO-DC-SIGN or CHO-GFP to CHO-CD80 cells of 1 : 1 or 10 : 1 were added to T cells stimulated with sub-optimal amounts of anti-CD3. Supernatants were tested 24 h later for the amounts of (A) IL-2 and (B) IFN-{gamma}.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
In this study we expressed both sDC-SIGN and DC-SIGN and tested for their ability to provide co-stimulation to T cells stimulated with anti-CD3 and PMA. DC-SIGN increased IL-2 secretion when either 30 µg ml–1 anti-CD3 or PMA was used to stimulate T cells. Interestingly, unlike CD80 and CD86, the DC-SIGN failed to provide co-stimulation when a sub-optimal amount of anti-CD3 (2 µg ml–1) was used to treat CD4+ T cells as measured by cell division, cytokine secretion and up-regulation of CD69 and CD25. Moreover, DC-SIGN modestly suppressed anti-CD3-induced IL-2 production (Fig. 4) and CD69 expression (Fig. 5).

Unlike stimulating anti-ICAM-3 HP2/19 antibody, DC-SIGN was unable to co-stimulate human CD4+ T cells treated with the same anti-CD3 antibody at 2 µg ml–1 (not shown and Figs 4–7 GoGoGo). It is interesting to note that both the HP2/19- anti-ICAM-3-stimulating antibody and LFA-1 bind the first Ig-like domain of ICAM-3 (15), while DC-SIGN binds the second Ig-like domain (16). In our co-stimulation assay system, DC-SIGN decreased the efficiency of anti-CD3-induced T cell stimulation with respect to CD69 up-regulation, IL-2 production (Figs 4–6GoGo) and CD80-mediated T cell co-stimulation (Fig. 7). In contrast, DC-SIGN enhanced IL-2 production by T cells stimulated with PMA, which can substitute for TCR signaling and activate protein tyrosine kinases and induce the high-affinity form of LFA-1. Moreover, the observation that DC-SIGN can enhance IL-2 secretion in the presence of 30 µg ml–1 of anti-CD3 suggested that a stronger stimulus (perhaps through increased tyrosine kinase activation) might be required to elicit the positive effects of DC-SIGN. These results revealed a complex role for DC-SIGN in the initial co-stimulation required for IL-2 production by T cells.

Low-affinity interactions between accessory molecules allow T cells to probe the surface of APCs. Actin-binding proteins ezrin and moesin interact with ICAM-3, helping to localize it to the uropod of migrating lymphocytes (17, 18). Subsequently, ICAM-3 quickly re-distributes to the point of contact in an antigen-independent manner (2). This then facilitates further interactions between the T cell and APC allowing the formation of an immunological synapse. This results in re-distribution and co-localization of ICAM-3 with ICAM-1 and LFA-1 in the outer edges of the immunological synapse. TCR-mediated events within the synapse further strengthen the T cell–APC interaction by inducing the high-affinity form of LFA-1 (19). Previous studies using in vitro co-stimulation assays demonstrated that ICAM-1 signaling induced by LFA-1 could enhance T cell stimulation (20, 21) and restore co-stimulation in CD28 knockout mice (22). Further, cells lacking both pathways were unable to respond to anti-CD3-induced T cell signaling and suggested that in the absence of B7-CD28 signaling, LFA-1–ICAM-1 is the predominant co-stimulatory pathway.

Based on our current and others' findings (1922), we speculate that the DC-SIGN has two roles in stimulating a T cell. There is an initial and transient binding that occurs between APCs and T cells through DC-SIGN and ICAM-3, which allows for the engagement of TCR and the MHC–peptide complex. This initial TCR stimulation increases LFA-1 avidity for ICAM-1 (5) and decreases LFA-1–ICAM-3 interaction. Since DC-SIGN can interact with ICAM-3, and not ICAM-1, it can induce signaling through ICAM-3 and enhance IL-2 production by T cells that receive a strong TCR signal.

In vivo, both CD83+ and CD83– APCs and specialized macrophages express DC-SIGN. Monocytes differentiated into immature DCs, which induce tolerance, express higher levels of DC-SIGN as compared with matured DCs (3). Based on our observation that DC-SIGN can lower the levels of IL-2 production by T cells on sub-optimal stimulation by anti-CD3, we speculate that DC-SIGN may help maintain tolerance by inhibiting IL-2 production by weakly stimulated T cells (23, 24). How DC-SIGN brings about these differential effects under sub-optimal versus optimal T cell signaling is not fully understood but might involve negative signaling through ICAM-3 (25), interference with LFA-1 binding to ICAM-3 (26) or freeing up LFA-1 from ICAM-3 to allow homotypic ICAM-1–LFA-1 interaction (5). Further studies are required to delineate its mechanism of action both in vitro and in vivo.


    Abbreviations
 
APC   antigen-presenting cell
CFSE   5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester
DAPI   4',6-diamidino-2-phenylindole
DC   dendritic cell
DC-SIGN   dendritic cell-specific intercellular cell adhesion molecule-3-grabbing non-integrin
FSC   forward scatter
GM-CSF   granulocyte macrophage colony- stimulating factor
ICAM   intercellular cell adhesion molecule
LFA   leukocyte function-associated antigen
PMA   phorbol myristate acetate
sDC-SIGN   a splice variant of dendritic cell-specific intercellular cell adhesion molecule-3-grabbing non-integrin
TMB   tetramethylbenzidine

    Notes
 
Transmitting editor: C. Terhorst

Received 20 July 2004, accepted 13 March 2005.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

  1. Steinman, R. M., Lustig, D. S. and Cohn, Z. A. 1974. Identification of a novel cell type in peripheral lymphoid organs of mice. 3. Functional properties in vivo. J. Exp. Med. 139:1431.[Abstract/Free Full Text]
  2. Montoya, M. C., Sancho, D., Bonello, G. et al. 2002. Role of ICAM-3 in the initial interaction of T lymphocytes and APCs. Nat. Immunol. 3:159.[CrossRef][ISI][Medline]
  3. Geijtenbeek, T. B., Torensma, R., van Vliet, S. J. et al. 2000. Identification of DC-SIGN, a novel dendritic cell-specific ICAM-3 receptor that supports primary immune responses. Cell 100:575.[CrossRef][ISI][Medline]
  4. Juan, M., Vinas, O., Pino-Otin, M. R. et al. 1994. CD50 (intercellular adhesion molecule 3) stimulation induces calcium mobilization and tyrosine phosphorylation through p59fyn and p56lck in Jurkat T cell line. J. Exp. Med. 179:1747.[Abstract/Free Full Text]
  5. Campanero, M. R., del Pozo, M. A., Arroyo, A. G. et al. 1993. ICAM-3 interacts with LFA-1 and regulates the LFA-1/ICAM-1 cell adhesion pathway. J. Cell Biol. 123:1007.[Abstract]
  6. Hernandez-Caselles, T., Rubio, G., Campanero, M. R. et al. 1993. ICAM-3, the third LFA-1 counterreceptor, is a co-stimulatory molecule for both resting and activated T lymphocytes. Eur. J. Immunol. 23:2799.[ISI][Medline]
  7. Berney, S. M., Schaan, T., Alexander, J. S. et al. 1999. ICAM-3 (CD50) cross-linking augments signaling in CD3-activated peripheral human T lymphocytes. J. Leukoc. Biol. 65:867.[Abstract]
  8. Bossy, D., Buckley, C. D., Holness, C. L. et al. 1995. Epitope mapping and functional properties of anti-intercellular adhesion molecule-3 (CD50) monoclonal antibodies. Eur. J. Immunol. 25:459.[ISI][Medline]
  9. Curtis, B. M., Scharnowske, S. and Watson, A. J. 1992. Sequence and expression of a membrane-associated C-type lectin that exhibits CD4-independent binding of human immunodeficiency virus envelope glycoprotein gp120. Proc. Natl Acad. Sci. USA 89:8356.[Abstract/Free Full Text]
  10. Geijtenbeek, T. B., Kwon, D. S., Torensma, R. et al. 2000. DC-SIGN, a dendritic cell-specific HIV-1-binding protein that enhances trans-infection of T cells. Cell 100:587.[CrossRef][ISI][Medline]
  11. Geijtenbeek, T. B., Krooshoop, D. J., Bleijs, D. A. et al. 2000. DC-SIGN-ICAM-2 interaction mediates dendritic cell trafficking. Nat. Immunol. 1:353.[CrossRef][ISI][Medline]
  12. Mummidi, S., Catano, G., Lam, L. et al. 2001. Extensive repertoire of membrane-bound and soluble dendritic cell-specific ICAM-3-grabbing nonintegrin 1 (DC-SIGN1) and DC-SIGN2 isoforms. Inter-individual variation in expression of DC-SIGN transcripts. J. Biol. Chem. 276:33196.[Abstract/Free Full Text]
  13. Fawcett, J., Holness, C. L., Needham, L. A. et al. 1992. Molecular cloning of ICAM-3, a third ligand for LFA-1, constitutively expressed on resting leukocytes. Nature 360:481.[CrossRef][ISI][Medline]
  14. de Fougerolles, A. R. and Springer, T. A. 1992. Intercellular adhesion molecule 3, a third adhesion counter-receptor for lymphocyte function-associated molecule 1 on resting lymphocytes. J. Exp. Med. 175:185.[Abstract/Free Full Text]
  15. Bell, E. D., May, A. P. and Simmons, D. L. 1998. The leukocyte function-associated antigen-1 (LFA-1)-binding site on ICAM-3 comprises residues on both faces of the first immunoglobulin domain. J. Immunol. 161:1363.[Abstract/Free Full Text]
  16. Bleijs, D. A., Geijtenbeek, T. B., Figdor, C. G. and van Kooyk, Y. 2001. DC-SIGN and LFA-1: a battle for ligand. Trends Immunol. 22:457.[CrossRef][ISI][Medline]
  17. Alonso-Lebrero, J. L., Serrador, J. M., Dominguez-Jimenez, C. et al. 2000. Polarization and interaction of adhesion molecules P-selectin glycoprotein ligand 1 and intercellular adhesion molecule 3 with moesin and ezrin in myeloid cells. Blood 95:2413.[Abstract/Free Full Text]
  18. Serrador, J. M., Alonso-Lebrero, J. L., del Pozo, M. A. et al. 1997. Moesin interacts with the cytoplasmic region of intercellular adhesion molecule-3 and is redistributed to the uropod of T lymphocytes during cell polarization. J. Cell Biol. 138:1409.[Abstract/Free Full Text]
  19. van Kooyk, Y., Weder, P., Hogervorst, F. et al. 1991. Activation of LFA-1 through a Ca2(+)-dependent epitope stimulates lymphocyte adhesion. J. Cell Biol. 112:345.[Abstract]
  20. Van Seventer, G. A., Shimizu, Y., Horgan, K. J. and Shaw, S. 1990. The LFA-1 ligand ICAM-1 provides an important costimulatory signal for T cell receptor-mediated activation of resting T cells. J. Immunol. 144:4579.[Abstract/Free Full Text]
  21. Semnani, R. T., Nutman, T. B., Hochman, P., Shaw, S. and van Seventer, G. A. 1994. Costimulation by purified intercellular adhesion molecule 1 and lymphocyte function-associated antigen 3 induces distinct proliferation, cytokine and cell surface antigen profiles in human "naive" and "memory" CD4+ T cells. J. Exp. Med. 180:2125.[Abstract/Free Full Text]
  22. Gaglia, J. L., Greenfield, E. A., Mattoo, A., Sharpe, A. H., Freeman, G. J. and Kuchroo, V. K. 2000. Intercellular adhesion molecule 1 is critical for activation of CD28-deficient T cells. J. Immunol. 165:6091.[Abstract/Free Full Text]
  23. Steinman, R. M. 2000. DC-SIGN: a guide to some mysteries of dendritic cells. Cell 100:491.[CrossRef][ISI][Medline]
  24. Soilleux, E. J. 2003. DC-SIGN (dendritic cell-specific ICAM-grabbing non-integrin) and DC-SIGN-related (DC-SIGNR): friend or foe? Clin. Sci. (Lond) 104:437.[Medline]
  25. Green, J. M. and Thompson, C. B. 1996. Homotypic interactions mediated through LFA-1/ICAM-3 decrease the proliferative response of activated T cells. Cell. Immunol. 171:126.[CrossRef][ISI][Medline]
  26. Bleijs, D. A., Binnerts, M. E., van Vliet, S. J., Figdor, C. G. and van Kooyk, Y. 2000. Low-affinity LFA-1/ICAM-3 interactions augment LFA-1/ICAM-1-mediated T cell adhesion and signaling by redistribution of LFA-1. J. Cell Sci. 113(Pt 3):391.[Abstract/Free Full Text]




This Article
Abstract
Full Text (PDF)
All Versions of this Article:
17/6/769    most recent
dxh258v1
Alert me when this article is cited
Alert me if a correction is posted
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Add to My Personal Archive
Download to citation manager
Request Permissions
Google Scholar
Articles by Martinez, O.
Articles by Prabhakar, B. S.
PubMed
PubMed Citation
Articles by Martinez, O.
Articles by Prabhakar, B. S.