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. 2010 Mar 15;7(3):211–220. doi: 10.1038/cmi.2010.9

Th1 cytokines promote T-cell binding to antigen-presenting cells via enhanced hyaluronan production and accumulation at the immune synapse

Paul L Bollyky 1, Stephen P Evanko 1, Rebecca P Wu 1, Susan Potter-Perigo 1, S Alice Long 1, Brian Kinsella 1, Helena Reijonen 1, Kelly Guebtner 1, Brandon Teng 1, Christina K Chan 1, Kathy R Braun 1, John A Gebe 1, Gerald T Nepom 1, Thomas N Wight 1
PMCID: PMC3027489  NIHMSID: NIHMS261328  PMID: 20228832

Abstract

Hyaluronan (HA) production by dendritic cells (DCs) is known to promote antigen presentation and to augment T-cell activation and proliferation. We hypothesized that pericellular HA can function as intercellular ‘glue' directly mediating T cell–DC binding. Using primary human cells, we observed HA-dependent binding between T cells and DCs, which was abrogated upon pre-treatment of the DCs with 4-methylumbelliferone (4-MU), an agent which blocks HA synthesis. Furthermore, T cells regulate HA production by DCs via T cell-derived cytokines in a T helper (Th) subset-specific manner, as demonstrated by the observation that cell-culture supernatants from Th1 but not Th2 clones promote HA production. Similar effects were seen upon the addition of exogenous Th1 cytokines, IL-2, interferon γ (IFN-γ) and tumor necrosis factor α (TNF-α). The critical factors which determined the extent of DC–T cell binding in this system were the nature of the pre-treatment the DCs received and their capacity to synthesize HA, as T-cell clones which were pre-treated with monensin, added to block cytokine secretion, bound equivalently irrespective of their Th subset. These data support the existence of a feedforward loop wherein T-cell cytokines influence DC production of HA, which in turn affects the extent of DC–T cell binding. We also document the presence of focal deposits of HA at the immune synapse between T-cells and APC and on dendritic processes thought to be important in antigen presentation. These data point to a pivotal role for HA in DC–T cell interactions at the IS.

Keywords: dendritic cell, hyaluronan, immune synapse, pericellular matrix, Th1

Introduction

The pericellular matrix or glycocalyx is a ‘coat' of hyaluronan (HA), proteoglycans and protein-associated glycans which extends outwards from the cell membrane of many cell types including all immune cells.1, 2, 3, 4 The precise composition and volume of the pericellular matrix varies substantially between cell types according to the state of activation.2

The primary structural component of the pericellular matrix is HA,1, 5 a repeating disaccharide of N-acetylglucosamine and D-glucuronic acid. The diameter of the pericellular matrix has been found to correlate with the level of HA secretion.6 Conversely, the pericellular matrix can be made to disappear from most cells by treating cells with 4-methylumbelliferone (4-MU), an inhibitor for HA synthesis.7, 8 Three HA synthases, termed HAS1, HAS2 and HAS3, extrude HA directly through the plasma membrane. Pericellular matrix-associated HA is anchored to the cell surface by an HAS or cell-surface receptors such as CD44.9 Additionally, a number of proteins and proteoglycans crosslink HA on the cell surface9, 10, 11 and create higher order levels of structure that may be significant to the formation of this coat.1 HA is a highly dynamic molecule and is constantly turned over via the activity of several hyaluronidases.12

While pericellular HA has well-established roles in osmotic balance, cell adhesion, control of proliferation and migration, and diffusion of nutrients and growth factors,1 little is known about the importance of HA at the immune synapse (IS) between T cells and antigen-presenting cells (APCs). Mummert et al.13 reported that HA produced and displayed on the surface of dendritic cells (DCs) contributed to both polyclonal and antigen-specific T-cell activation. They found that inhibition of HA production by DCs via 4-MU treatment or blockade with the HA-specific binding peptide, Pep-1, led to impaired T-cell proliferation, and diminished IL-2 and interferon γ (IFN-γ) production. These data suggest that pericellular HA facilitates T-cell activation in a paracrine manner. In addition to a well-documented role in adhesion in the context of migration and extravasation,1 HA is known to promote DC maturation and upregulation of costimulatory molecules14, 15 and can stimulate both T cells and DCs individually via the receptors CD4416, 17 and TLR4.18, 19

HA production is known to be positively regulated by proinflammatory cues and negatively regulated by anti-inflammatory agents, a paradigm which is consistent with a putative role for HA in promoting antigen presentation. Interestingly, the majority of cytokines, such as IFN-γ, tumor-necrosis factor (TNF)-α and IL-1β, which promote HA production in a variety of cell types tend to be associated with the T helper 1 (Th1) subset of T cells;20, 21, 22, 23 cytokines associated with the Th2 subset of T cells, such as IL-4, on the contrary, generally do not,24, 25, 26 though there are exceptions.27 This observation led us to investigate whether T cells might not directly influence cell-surface HA production by virtue of their cytokine production profiles.

We have evaluated the hypothesis that pericellular HA directly moderates DC–T-cell interactions. We have evaluated whether HA production by DC promotes the formation and stability of T-cell-DC binding. We have then asked whether individual Th1 cytokines or cultured media from Th1 clones have the capacity to positively influence T-cell binding to DC in an HA-dependant manner. Our results represent a novel, pivotal role for HA and the pericellular matrix in regulating DC–T-cell interactions.

Materials and methods

Reagents

HA with a molecular weight of 1.5×106 kDa was provided by Genzyme (Cambridge, MA, USA). 4-MU and Streptomyces hyaluronidase were obtained from Sigma-Aldrich (St Louis, MO, USA).

Human blood samples

Human peripheral blood mononuclear cell (PBMC) samples were obtained from healthy volunteers with informed consent, participating in a research protocol approved by the institutional review board of the Benaroya Research Institute at Virginia Mason (BRI, Seattle, WA, USA).

Isolation of leukocyte populations

Human PBMCs were prepared by centrifugation of peripheral blood over Ficoll–Hypaque gradients. CD4+ T cells were isolated using the Dynal CD4 Positive Isolation Kit (Invitrogen, Carlsbad, CA, USA) as per the manufacturer's instructions. Purity of the resulting cell fractions was reliably >98% CD4+ by flow cytometry; anti-CD4 Ab (RPA-T4), from BD-Biosciences (San Jose, CA) was used for this purpose. Cells were cultured in RPMI 1640 (Invitrogen) supplemented with 10% pooled human serum, 100 µg/ml penicillin, 100 U/ml streptomycin and 1 mM Na pyruvate (Invitrogen). Monocytes were isolated from the CD4− population as per the manufacturer's instructions.

Generation of monocyte-derived DCs

CD4−CD14+ cells (monocytes) were cultured in 24 well plates, 3×106 cells/well. Cells were cultured 4–6 days in the presence of IL-4 (50 ng/ml; R&D, Minneapolis, MN, USA) and granulocyte/macrophage colony-stimulating factor (5 ng/ml; BD Pharmingen, BD Biosciences). Cells were stained before and after this protocol for CD14, CD80 and CD86 to document their development into DCs. A representative staining example is shown in Supplementary Figure 1.

Generation and characterization of Th1 and Th2 clones

PBMCs from human leukocyte antigen (HLA) DRB0404+ patients were stimulated as described earlier.28 Briefly, cells were cultured with RPMI 1640 containing 10% (v/v) pooled human serum at the density of 5×106/ml, in the presence of a GAD65 555-567 (557I; NFIRMVISNPAAT) peptide at a concentration of 10 µg/ml. On day 10, the cells were transferred at a density of 4×106/ml onto 48-well plate that had been adsorbed with 8 µg/ml DRB0404 monomer containing GAD65 557I peptide major histocompatibility complex (MHC) in 1X phosphate-buffered saline (PBS) for 3 h at 37 °C. One microgram per milliliter of anti-CD28 antibody (BD Pharmingen, BD Biosciences) was added to the media and the cells incubated additional 5–6 days and stained using 10 µg/ml phycoerythrin-labeled 0404 tetramer for 2 h at 37 °C, and subsequently with fluorochrome-labeled anti-CD25 and anti-CD4 (BD Pharmingen, BD Biosciences) for 30 min on ice. Cells were then washed with PBS containing 1% fetal bovine serum and analyzed using a BD FACSCalibur flow cytometer (BD Biosciences). Data analysis was performed using CellQuest (BD Biosciences) software. CD4highCD25+ tetramer-binding cells were single-cell sorted into 96-well plates using a FACSVantage cell sorter (BD Biosciences). Sorted clones were expanded for 10–12 days by stimulation with irradiated unmatched PBMCs (1.5×105/well), 5 µg/ml phytohemagglutinin and 10 U/ml IL-2 for two cycles, followed by stimulation with HLA-DRB404-matched PBMCs pulsed with 10 µg/ml GAD65 557I peptide and 10 U/ml IL-2. On days 10–12, clones were selected based on growth for further expansion. Resting T cells (5×104) were tested for antigen specificity by stimulation with irradiated HLA-DR404-matched PBMCs (1×105/well) with and without a specific peptide GAD65 555-567 (NFFRMVISNPAAT) in the culture (0.01–10 µg/ml). Proliferation as measured by 3H-thymidine incorporation was tested after 72 h in culture. The restriction elements of the T-cell clones were confirmed by testing proliferation induced by DR0404-transfected type 1 bare-lymphocyte-syndome cell lines (BLS-1) pulsed with GAD65 peptide. Cytokine secretion by the cells was measured at 48 h after stimulation by Cytometric Beads Array assay (BD Pharmingen, BD Biosciences) according to the manufacturer's instructions. Clones were classified as Th1 or Th2 based on their cytokine production profiles (Supplementary Table 1). T-cell clones were tested for tetramer binding by staining with 10 µg/ml GAD65 or control tetramer for 1 h at 37 °C followed by fluorochrome-conjugated antibody on ice for 30 min. The T-cell clones were expanded by stimulation at 2-week intervals with either irradiated non-HLA-matched PBMCs, phytohemagglutinin (5 µg/ml) and IL-2 (10 U/ml), or GAD peptide-pulsed DR0401+ PBMCs and IL-2 (10 U/ml). After 3–4 cycles of stimulation the cells were frozen and aliquots were subsequently thawed for the experiments described in this work.

Generation and characterization of stock Th1- and Th2-conditioned media solutions

Three pairs of thawed Th1 and Th2 clones were stimulated with irradiated non-HLA-matched PBMCs, phytohemagglutinin and IL-2, as described above. Cells were maintained for three weeks with periodic readministration with IL-2 and replacement of media. Whenever this was done the conditioned media was harvested and stored at 4 °C. Conditioned media was collected from the three Th1 clones in approximately equal proportions and pooled. The same was done for the Th2 cell cultures. In this manner a single stock of Th1 conditioned media and a single stock of Th2 conditioned media were generated over the course of 2 weeks. These two stocks underwent cytokine profiling as described above and the results are shown in Table 1.

Table 1. Cytokine profiles for the Th1- and Th2-conditioned media used as stock solutions for these experiments. These stock solutions were derived from pooled Th1- and Th2-clone culture supernatants.

Conditioned media IFN-γ TNF-α IL-10 IL-4 IL-2
Th1 1183.5 84.7 1452.6 10.6 38.9
Th2 264.4 51.2 1176.7 9571.3 0
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Abbreviations: IFN, interferon; Th, T helper; TNF, tumor-necrosis factor.

Treatment of DCs with cytokines or Th1- or Th2-conditioned media

DCs generated from monocytes were collected, washed in complete media, and replated at 3×106 cells/well in a 6-well plate. Th1- or Th2-conditioned media were added in 2∶1 ratio of fresh complete media to stock conditioned media. Fresh media alone were used as a control. Alternatively, where indicated fresh media was supplemented with recombinant cytokines individually and in combination at the following concentrations: 100 IU/ml IL-2 from (Chiron, Emeryville, CA, USA); 10 ng/ml recombinant human IFN-γ (R&D Systems, Minneapolis, MN, USA); 10 ng/ml recombinant human TNF-α (BD Pharmingen, BD Biosciences), 10 ng/ml recombinant human IL-1β (BD Pharmingen, BD Biosciences). DCs were treated in this manner for 4 h prior to mRNA harvest.

Quantitiative PCR

Total RNA was harvested from T cells and DCs using the RNeasy Mini Kit from Qiagen (Valencia, CA, USA). cDNA was prepared from 350 ng total RNA reverse transcribed in a 40 µl reaction mix with random primers using the High-Capacity cDNA Archive Kit according to the manufacturer's instructions. Relative quantification of transforming growth factor-β1 gene expression was performed using Taqman Gene Expression Assay Mm03024053_m1 and eukaryotic 18S rRNA Endogenous Control part no.4333760. Briefly, 1.2 µl cDNA was amplified in 1XTaqman Fast Universal PCR Mix with 250 nM Taqman probe in a 20 µl reaction using the Fast program for 50 cycles on an ABI7900HT thermocycler. All qPCR reagents were from Applied Biosystems (Foster City, CA, USA). All samples were done in duplicate and data were analyzed using the Comparative Ct Method with software from Applied Biosystems. Estimated copy numbers were generated from a standard curve created by using a selected reference cDNA template and Taqman probe.29

Quantification of ha synthesis

H3-glucosamine was added at a concentration of 40 µCi/ml to DC cultures. After 24 h the supernatant was removed, thereby separating each condition into soluble and cellular fractions. These fractions were then digested with pronase (100 µg/ml) in 0.5 M Tris pH 6.5 overnight at 37 °C. Following digestion, the pronase was inactivated by heating to 100 °C for 20 min. Radiolabeled macromolecules were then recovered and separated from unincorporated precursor by precipitation on nitrocellulose membranes using slot blot analysis as described previously.30 Briefly, 200 µl of the sample was added to an equal volume of 2% cetylpyridiniumchloride, 50 mM NaCl buffer and the solution blotted onto 0.45 µm nitrocellulose membrane (Schleicher and Schuell, Keene, NH, USA). The membrane was washed for six times in 2% cetylpyridiniumchloride, 50 mM NaCl buffer and once in deionized water before air drying at room temperature overnight. Incorporation of 3H-glucosamine into HA was measured by digesting an equivalent radiolabeled aliquot with Streptomyces hyaluronidase (2 U/ml) for 24 h at 37 °C before slot blotting. HA was measured as the amount of hyaluronidase sensitive material precipitated to the nitrocellulose membrane. To determine the amount of chondriotin sulfate and dermatan sulfate present in the sample, an equal aliquot of sample was adjusted to pH 8.0 before digesting with Chondroitin ABC lyase (0.03 U/ml; North Star BioProducts, East Falmouth, MA, USA). All scintillation counting was done on Beckman LS 6500 (Beckman Instruments, Fullerton, CA, USA).

T cell–DC binding experiments

These were modeled on previous work by Do et al.,14 with modifications. DCs induced from monocytes were stained using SNARF-1 (Invitrogen). After this 1×105 DCs per condition were cultured on a coverslip in a 6-well plate (Corning, Corning, NY, USA) overnight. Th1- or Th2-conditioned media was added in 2∶1 ratio of fresh complete media to conditioned media. Fresh media alone was used as a control. Alternatively, fresh media was supplemented with 100 IU/ml IL-2 and 10 ng/ml IFN-γ. DCs were maintained under these culture conditions overnight. The plates were gently washed twice to remove any conditioned media.

Primary T cells were activated for 4 days with anti-CD3/28 beads (Invitrogen) at a ratio of 1 bead to 10 cells without exogenous IL-2. Activated T cells were then labeled with 50 µM, carboxyfluorescein succinimidyl ester (CFSE; Invitrogen). CD4+ T cells which were labeled by 1×105 CFSE were added to the plates. After incubation for 2 h, the plates were washed in PBS; the coverslips were removed and mounted on slides for analysis.

Analysis of binding was performed as follows. For each condition, at least 10 non-overlapping fields were photographed using a digital camera (Diagnostic Instruments, Sterling Height, MI, USA) attached to a Leica DM-IRB microscope (Leica Microsystems, Wetzlar, Germany). Spot Software 4.5 (Diagnostic Instruments) was used for analysis. For each field photographs were taken using the excitation laser at 488 and 568 in order to capture binding of both CFSE-labeled T cells as well as SNARF-1-labeled DCs. For each field the two images were then merged using software in order to provide an assessment of clustering involving both T cells and DCs. Using this image, for each field the number of DCs was counted as well as the number of clusters. A cluster was defined as ≥2 T cells bound to one or more DCs.

Binding experiments were also performed using CFSE-labeled Th1- or Th2-cell clones activated in an identical manner to the primary CD4+ T cells described above. The cytokine production profiles for these clones are shown in Supplementary Table 1. These T-cell clones were pre-incubated for 1 h with BD GolgiStop (BD Pharmingen, BD Biosciences). BD GolgiStop was added to prevent any further cytokine contribution from T cells to the binding assay. The T-cell clones were then washed twice and incorporated into the same binding assay protocol as above.

Immunocytochemistry

For visualization of HA, cells were fixed in an acid alcohol formalin buffer as described by Lin et al.31, thereby allowing maximum retention of the HA and associated proteins. The coverslips were blocked at room temperature with 1% bovine serum albumin/5% normal donkey serum in PBS for at least 1 h. Subsequently, they were incubated with bPG (the N- terminal HA binding region of aggrecan which has been biotinylated) as previously described.32 Following three washes, cells wereincubated with Alexa Fluor 488 streptavidin (2 µg/ml), Alexa Fluor 647 anti-HLA-DR (L243; Biolegend, San Diego, CA, USA) (8 µg/ml) and DAPI (1 µg/ml) for 1 h. Following three washes in PBS, coverslips were mounted in Gel/Mount (Biomeda, Foster City, CA, USA), and photographed using the apparatus described above.

Statistical analysis

Statistical comparisons were made using a Student's t-test. Standard error is shown unless otherwise noted. Values of P<0.05 were considered significant.

Results

Biosynthesis of HA by DCs

We first sought to quantify and characterize HA production by monocyte-derived human DCs. This was accomplished using a method by which the incorporation of radiolabeled glucosamine into HA is assessed.33 Because glucosamine is converted into HA as well as other molecules, the proportion of HA was calculated from parallel aliquots digested with and without Streptomyces hyaluronidase, an enzyme specifically degrading HA;34 chondroitinase is used as a control in these studies to confirm the specificity of the assay for HA. Using this method we found that monocyte-derived human DCs but not primary T cells produced HA (Figure 1a). The majority of this HA production was cell associated as HA was observed in the cellular fraction but not the media fraction from DC cultures (Figure 1b).

Figure 1.

Figure 1

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DCs produce cell-associated HA. (a) Radiolabeled glucosamine incorporation into HA by DCs and T cells. Since glucosamine is converted into HA as well as other molecules, the proportion of HA was calculated from parallel aliquots digested with and without Streptomyces hyaluronidase. Chondroitin ABC lyase is used as a control in these studies to confirm the specificity of the assay for HA. (b) Radiolabeled glucosamine incorporation into HA for the media and cell fractions of DC cultures. DCs were cultured with and without 4-MU treatment at the time of glucosamine addition. Error bars are for replicates in triplicate. Data are representative of four experiments. (c) mRNA expression for the three hyaluronan synthases normalized to 18S mRNA. Results include data from four independent experiments each using DCs from different individuals. CPM, counts per min; DC, dendritic cell; HA, hyaluronan; HAS, hyaluronan synthases; 4-MU, 4-methylumbelliferone.

Of the three HAS genes, HAS3 was most highly expressed by monocyte-derived human DCs with minimal expression of HAS1 and no detectable expression of HAS2. Activated T cells produced negligible quantities of cell-associated or soluble HA (Figure 1a). Of note, this was also the case for unactivated T cells as well as the T-cell clones used in later experiments (data not shown). We therefore subsequently focused only on HA production by DCs.

DC production of HA was substantially abrogated upon treatment with 4-MU used at 50 µg/ml (Figure 1b). This dose of 4-MU was demonstrated to be non-toxic to DCs as ascertained by staining with 7-AAD and Annexin V (Supplementary Figure 2). This corroborates reports with other cell types that 4-MU at comparable concentrations is not directly cytotoxic.35, 36, 37 Hyaluronidase treatment had the paradoxical effect of increasing DC production of HA (data not shown) as has been shown in other systems.38, 39

DCs produce HA in response to Th1-conditioned media and to IL-2 and IFN-γ

We were interested in what factors might govern HA production by DCs and hypothesized that soluble factors produced by T cells might play a role, given the importance of DC–T cell cross-talk in antigen presentation. Conditioned media from Th1-clone cultures but not Th2-clone cultures had the capacity to upregulate HA production by DCs (Figure 2a). This was not the case for chondroitin sulfate, demonstrating that this effect is specific to HA.

Figure 2.

Figure 2

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DCs produce cell-associated HA in response to Th1-conditioned media and to specific Th1 cytokines. (a) Radiolabeled glucosamine incorporation into HA or chondroitin sulfate upon coculture in the setting of Th1- or Th2-cell culture supernatants. CPM shown are for the amount of radiolabel lost upon treatment of the radiolabeled cell lysate with either Streptomyces hyaluronidase or chondroitin ABC lyase. (b) Radiolabeled glucosamine incorporation into HA upon coculture in the setting of Th1-cell culture supernatants or specific cytokines as shown. CPM shown are for the amount of radiolabel lost upon treatment of the radiolabeled cell lysate with Streptomyces hyaluronidase. Error bars are for replicates in triplicate; data are representative of four experiments. CPM, counts per min; DC, dendritic cell; HA, hyaluronan; IFN, interferon; Th, T helper; TNF, tumor-necrosis factor; 4-MU, 4-methylumbelliferone.

The role of cytokines in regulating HA production by other cell types is well documented. Therefore, we suspected that a cytokine or several cytokines might be responsible for the effects seen with Th1-cultured media. To this end, we evaluated the effects of various Th1 cytokines alone or in combination on DC HA production (Figure 2b). We found that TNF-α and IL-2 alone promoted HA production while IL-1β and IFN-γ did not. However, the combination of IL-2 plus IFN-γ produced more HA than IL-2 alone while the combination of IL-2 and TNF-α performed the best. As reported above for DCs which did not receive further cytokine or cultured media treatment, the entirety of this HA production was cell associated with no detectable HA in the media fraction (data not shown). The finding of a role for IFN-γ and TNF-α in HA production is consistent with previous reports using fibroblast cell lines.20, 21, 22, 23 We opted to focus our further work on the contribution of IL-2 and IFN-γ to DC production of HA because these cytokines are well characterized as being produced primarily by T cells. Therefore, any IL-2 and IFN-γ in the mixed DC–T cell cultures could be expected to be of T-cell origin. TNF-α, conversely, can also be produced by DCs themselves.40

Interestingly, these data suggest that DCs have a previously unreported capacity to respond to IL-2. DCs do not express the high-affinity IL-2 receptor CD25 (data not shown), but we did find that DCs do express the low-affinity receptor CD122 (Supplementary Figure 1). However, neither antibodies directed at CD122 nor at IL-2 itself negated the enhanced production of HA observed upon treatment with Th1-conditioned media (data not shown). This suggests that IL-2 is not a strict requirement for enhanced DC production of HA under this condition.

The accumulation of cell-surface HA of DCs is likely to reflect the balance of production as well as degradation of HA. We therefore evaluated the effects of Th1-conditioned media and controls on the genes responsible for HA synthesis and degradation. Interestingly, Th1-conditioned media, and IL-2 and IFN-γ supplementation did not engender any increase in HAS3 mRNA (Figure 3a). Neither was there any increase in HAS1 nor HAS2 mRNA as these both remained at negligible levels under all conditions tested (data not shown). However, we did note a significant decrease in the expression of mRNA for the hyaluronidase HYAL1 with either Th1-conditioned media, or IL-2 and IFN-γ supplementation (Figure 3b). This suggests that Th1-conditioned media, and IL-2 and IFN-γ may affect the HA content of DCs primarily by impacting the rate of degradation rather than production. Similar complex effects on HA production and degradation were reported for IFN-γ and TNF-α in lung fibroblasts.41

Figure 3.

Figure 3

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DCs downregulate HYAL1 in response to IL-2 and IFN-γ. (a) mRNA expression of HAS3 4 h following addition of Th1- and Th2-cell culture supernatants or supplementation with IL-2 and IFN-γ. (b) mRNA expression of the hyaluronidase genes HYAL1 and HYAL2 under the same conditions. Results include data from four independent experiments each using DCs from different individuals. * connotes conditions where there was a significant (P≤0.05) difference between that sample and the media-only condition. DC, dendritic cell; HAS, hyaluronan synthases; HYAL, hyaluronidase; IFN, interferon; Th, T helper.

Conditioned media from Th1 cytokines, and IL-2 and IFN-γ supplementation promote DC spreading

Induced DCs had the typical morphological appearance of immature DCs: they were predominantly non-adherent and had few DC processes (Figure 4a). Upon treatment with Th1-conditioned media, these cells became more adherent, formed long dendrites and tended to cluster (Figure 4b). These are features associated with DC maturation and heightened capacity to present antigen.42, 43. These morphologic changes could be abrogated by treatment with 4-MU (Figure 4c). In contrast, DCs incubated with Th2-conditioned media were only slightly altered in appearance (Figure 4d). The same changes observed with Th1-conditioned media could be seen upon treatment with IL-2 and IFN-γ (Figure 4e). As previously reported, these treatments impacted expression of CD80 and CD86 in tandem with their effects on dendritic processes.44, 45, 46 (Data not shown).

Figure 4.

Figure 4

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Conditioned media from Th1 cells promote DC spreading in an HA-dependent manner. DCs treated with (a) control media, (b) Th1-conditioned media, (c) Th1-conditioned media with the addition of 4-MU, (d) Th2-conditioned media, or (e) control media supplemented with IL-2 and IFN-γ. Images are shown at ×10 magnification. Data are representative of three separate experiments. DC, dendritic cell; HA, hyaluronan; IFN, interferon; Th, T helper; 4-MU, 4-methylumbelliferone.

Treatment of DCs with Th1-conditioned media as well as IFN-γ and IL-2 supplementation promote T-cell binding in an HA-dependent manner

We sought to ascertain whether HA production by DCs contributed to T-cell binding. To this end, we evaluated cluster formation between SNARF-labeled DCs and CSFE-labeled T cells. DCs pre-treated with Th1-cultured media demonstrated a significantly greater propensity to bind to primary T cells than controls (Figure 5a–e). This enhanced binding could be abrogated by inclusion of 4-MU during the pre-treatment phase (Figures 5c and e). A similar response to that seen with Th1-cultured media pre-treatment was seen upon pre-treatment of DCs with media supplemented with IFN-γ and IL-2 (Figure 5e). The addition of exogenous HA to the DC–T cell cultures did not significantly improve cluster formation (Figure 5e). These data support the conclusion that Th1 soluble factors act on DCs in a paracrine manner to promote HA-dependent DC–T cell binding.

Figure 5.

Figure 5

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Treatment of DCs with Th1 cytokines promotes T-cell binding. Binding of SNARF-labeled DCs with CSFE-labeled T cells was ascertained by evaluating DC–T cell clusters. A cluster was defined as ≥2 T cells bound to one or more DCs. The DCs were pre-treated overnight with a stock of Th1-cell culture supernatants or controls and then washed prior to the addition of the T cells. Representative images are shown for DCs pre-treated with (a) media alone, (b) Th1-cell culture supernatants and (c) Th1-cell culture supernatants together with 4-MU. Random fields (10–15) were evaluated per condition. The number of conjugates was normalized to the number of DCs for each field. A close-up of DC–T cell clusters present upon Th1 supernatant treatment is shown (d). Brightness and contrast were identical for each color in single and merged images. (e) The percentage of DCs pre-treated with Th1-cell culture supernatants or controls bound to activated primary T cells in a cluster. Data are representative of three separate experiments made using DCs and autologous T cells from different individuals. (f) Fold change in the percentage of DCs bound to T-cell clones (two Th1 and two Th2) taken from a pair of individuals (termed a and b). For each clone comparison the left bracket is for the test of significance for the Th1 supernatant condition versus media alone while the right bracket is for the Th1 supernatants comparison versus Th2 supernatants. * indicates a P value of <0.05. DC, dendritic cell; HA, hyaluronan; IFN, interferon; Th, T helper; 4-MU, 4-methylumbelliferone.

These experiments were repeated using Th1 and Th2 clones in order to better determine whether the effects on DC–T cell binding described here were restricted by T-cell subtype. The T-cell clones used were the same clones from which the conditioned media were derived and activated in an identical manner to the aforementioned primary T cells. These T cells were CFSE labeled and pre-treated with GolgiStop to prevent any further cytokine production by these cells. We found that in all cases T cells exhibited significantly increased binding to DCs pre-treated with Th1-conditioned media (Figure 5f). For three of the four clones analyzed, the clone bound significantly better to DCs pre-treated with Th1 cytokines, irrespective of whether the clone was a Th1 or a Th2 clone. The fourth clone, a Th1 clone, likewise demonstrated increased binding to Th1-conditioned media-treated DCs over those treated by Th2-conditioned media, but this difference was not significant (Figure 5f). These data support the conclusion that the crucial determinant in these HA-dependent binding interactions is the nature of the cytokine milieu which DCs encounter and that subsequent binding to T cells is not subtype restricted.

IFN-γ and IL-2 promote HA accumulation on dendritic processes and at the IS

Pseudopodia are cellular extensions known to be enriched for MHC class II molecules and thought to function in antigen presentation. We observed that HA tended to be concentrated on such structures upon treatment of CD4 APCs with IFN-γ and IL-2 (Figure 6a) and that these could be removed with hyaluronidase treatment (Figure 6b) and pre-treatment with 4-MU (data not shown). These structures and the HA deposits are seen clearly in cross-sectional views (Figure 6d and e).

Figure 6.

Figure 6

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IFN-γ and IL-2cytokines promote HA accumulation on dendrites. (a–d) bPG staining of CD4 cells cultured overnight in the presence of media alone (a) or with the addition of IL-2 and IFN-γ (b). (c) bPG binding of IL-2/IFN-γ conditioned cells subsequently treated with HA'ase prior to staining with HABP. (d–e) A single cell from an IL-2 and IFN-γ conditioned culture; images are shown for both bPG staining (d) and phase contrast (e). The white arrow in both D and E points to the same pseudopodia in both views. bPG, bovine proteoglycan; DC, dendritic cell; HA, hyaluronan; HABP, hyaluronic acid binding protein; HA'ase, hyaluronidase; IFN, interferon.

Upon addition of activated autologous CD4 T cells to these same CD4 APCs which had been pre-treated with IFN-γ and IL-2, we observed the colocalization of HA (green) and MHC class II (red) at sites of union between cells (nuclei stained in blue) (Figure 7). MHC class II is enriched upon APCs and is a key component of the IS. The colocalization of HA and MHC class II supports the conclusion that HA is present at the IS.

Figure 7.

Figure 7

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IFN-γ and IL-2cytokines promote focal clusters of HA accumulation at the immunological synapse. PBMCs were activated with soluble anti-CD3 and anti-CD28 Ab in the setting of exogenous IFN-γ and IL-2. Images are shown for two clusters of cells for bPG (green) alone (a and c) and for costaining for MHC class II (red) and DAPI (blue) (b and d). Treatment with 4-MU abrogated clustering as well as diminished the amount of visible hyaluronan (not shown). Brightness and contrast were identical for each color in single and merged images. This experiment was performed twice with similar results. Ab, antibodies; DAPI, 4,6-diamidino-2-phenylindole; DC, dendritic cell; HA, hyaluronan; IFN, interferon; MHC, major histocompatibility complex; PBMC, peripheral blood mononuclear cell; 4-MU, 4-methylumbelliferone.

Discussion

We have evaluated the hypothesis that pericellular HA can function as intercellular ‘glue' directly mediating T cell–DC binding. Using primary human cells, we observed HA-dependent binding between T cells and DCs. Moreover, we describe the existence of a feedforward loop wherein T-cell cytokines influence DC production of HA in a Th1-dependent manner which in turn affects the extent of DC–T cell binding in a Th1-independent manner. These data suggest that HA plays a dynamic pivotal role in DC–T cell interactions.

Human monocyte-derived DCs produce substantial cell surface-associated HA, which is associated with HAS3 expression. These results are consistent with previously published data for mouse DC lines and mouse bone marrow-derived DCs.13 In contrast, we observed negligible production of HA by either freshly isolated or activated T cells. While mouse T cells are reported to produce physiologically relevant quantities of HA,35 these data suggest that the bulk of HA present in human DC–T cell interactions is of DC origin.

Pericellular HA facilitates DC–T cell binding. 4-MU pre-treatment abrogated this binding while treatments were shown to increase HA-promoted binding. Interestingly, we find that the addition of exogenous HA to the DC–T cell cocultures did not significantly improve binding, suggesting that exogenous soluble HA was not equivalent to cell-associated HA. This binding could be mediated by interactions between HA on the surface of DCs and membrane-bound CD44 on the surface of T cells, as suggested by work with CD44−/− mice where CD44 expression was required on T cells but not DCs for optimal antigen presentation.47 However, the mechanisms by which HA promotes DC–T cell binding are unlikely to be exclusively CD44 mediated. Indeed one group reported that CD44 expression on T cells is not necessary for antigen presentation by DC.14 A variable role for charge-based interactions or the binding of other receptors to HA-associated proteoglycans could explain these conflicting data. Such alternative mechanisms could also account for data showing that the addition of an HA-binding peptide, Pep-1, did not significantly impair DC–T cell conjugate formation, whereas this treatment inhibited most other HA receptor-mediated interactions.13 Indirect effects of HA on DC maturation and expression of adhesion molecules are also likely to contribute to binding.

Production of HA by DCs was promoted by conditioned media from Th1 clones but not Th2 clones. Similar effects were seen upon treatment with the Th1 cytokines, TNF-α and IFN-γ, in combination with IL-2. IFN-γ and TNF-α are both well known to promote antigen presentation by DCs; their effects on HA are consistent with this role. These findings square well with reports of these same cytokines alone and in concert promoting HA production by other cell types.20, 21, 22, 23

These data suggest that Th1 cells may directly influence HA production by APCs by virtue of their cytokine production profiles. In the context of our aforementioned binding data, this represents a novel feedforward mechanism whereby T cells influence their own binding to DCs in an HA-dependent manner. While similar feedforward mechanisms exist elsewhere in immunology, none, to our knowledge, involve the extra-cellular matrix. Moreover, these data raise the intriguing possibility that HA production is an integral part of the Th1 program. A putative role for HA in Th1 responses is supported by reports that the viral mimetic Poly(IC)32 and infection with the Epstein–Barr virus48 both promote HA production.

We were interested to find that IL-2 promotes HA production by DCs because these cells do not express the high-affinity receptor for IL-2, CD25. DCs presumably respond to IL-2 through the low-affinity IL-2 receptor, CD122. We demonstrate here that monocyte-derived DCs express this receptor. CD122 expression has also been reported for bone marrow-derived DCs.46

We observed focal deposits of HA at sites where T cells and APCs are thought to interface. Specifically, HA deposits colocalized with MHC class II at the IS. This may help explain published data on the role of HA in antigen presentation.11 HA deposits were also present atop cellular protrusions known to play a role in antigen presentation. Furthermore, 4-MU treatment abrogated formation of dendrites on DCs; this is consistent with reports that HA may play a role in maintaining similar structures on other cell types.5, 49, 50

IS-associated HA could contribute to antigen presentation via several mechanisms, potentially acting in concert. HA has osmotic properties and could promote gradients of molecules that signal at the IS.51 HA could act as a costimulatory molecule through paracrine and/or autocrine interactions with its primary extracellular HA receptor, CD44. This molecule has been shown to complex with the T-cell antigen receptor52 and to colocalize with lipid rafts at the IS.47 Indeed, mice deficient for CD44 demonstrated decreased phosphotyrosine and protein kinase C-enrichment at the synapse and impaired antigen presentation.14, 47 An intriguing possibility is that HA might also directly promote the formation and stability of the IS by acting as ‘intercellular glue' joining cells in apposition and impacting their engagement and activation. It is certainly the case that cells must have mechanisms for interdigitating the pericellular matrix on their surfaces, otherwise it is hard to imagine that they could achieve the close contact necessary for MHC–T-cell antigen receptor complex formation.53, 54, 55 It seems likely that pericellular matrix interactions represent a heretofore unappreciated and potentially important level at which the IS is regulated.

Acknowledgments

This work was supported by grants from the NIH (DK46635, HL18645 and DK53004) and the JDRF (The Center for Translational Research at BRI). PLB is supported by NIH K-08 grant DK080178-01 and an NIH LRP grant. The authors would like to thank Nathan Standifer and Michael Kinsella for their helpful comments and Tuan Nguyen for tissue processing.

Footnotes

Note: Supplementary information is available on the Cellular & Molecular Immunology website (http://www.nature.com/cmi/).

Supplementary Information

Supplementary information
Click here for additional data file. (114KB, pdf)

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