International Immunology Advance Access originally published online on January 13, 2006
International Immunology 2006 18(3):445-452; doi:10.1093/intimm/dxh384
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Purification of splenic dendritic cells induces maturation and capacity to stimulate Th1 response in vivo
1 Institut Pasteur, Unité de Biologie des Régulations Immunitaires, Paris F-75015, France
2 Inserm, E 352, Paris F-75015, France
Correspondence to: G. Dadaglio; E-mail: gdadag{at}pasteur.fr
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Abstract |
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Dendritic cell (DC) maturation state is a key parameter for the issue of DC–T cell cognate interaction, which determines the outcome of T cell activation. Indeed, immature DCs induce tolerance while fully mature DCs generate immunity. Here we show that, in the absence of any deliberate activation signal, DCs freshly isolated from mouse spleen spontaneously produce IL-12 and tumor necrosis factor-
and up-regulate co-stimulation molecules, even when directly re-injected into their natural environment. Furthermore, after their isolation, these cells acquire the capacity to induce specific Th1 responses in vivo. These results demonstrate that the sole isolation of spleen DCs leads to the full maturation of these cells, which therefore cannot be considered as immature DCs. Moreover, we also show that the kinetics of DC activation do not influence the polarization of Th response in vivo challenging the idea that exhausted DCs induce preferentially Th2 response. Altogether, these observations should be taken into account in all experiments based on the transfer of ex vivo purified DCs.
Keywords: activation, antigen-presenting cell, T cell response
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Introduction |
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It is generally accepted that dendritic cells (DCs) display opposite functions in the immune system as they control both tolerance and immunity according to their maturation state (1). In the steady-state conditions, immature DCs reside within most tissues in which they participate in the maintenance of tolerance by continuously sampling their environment and presenting self-antigens to T cells. Activation of immature DCs directly by a pathogen or indirectly by inflammatory cytokines results in DC maturation characterized by the up-regulation of their stimulatory capacity and their migration to lymphoid organs. After activation, mature DCs are the most potent antigen-presenting cells to prime naive T cells against microbial antigens encountered earlier in periphery. Altogether, these observations strongly suggest that immature DCs play a role of sentinels in the periphery, whereas mature DCs switch on the immune response in the lymphoid organs (2).
To sense the modifications in their environment, DCs use a large panel of receptors such as Toll-like receptors (TLRs) or cytokine receptors and DC maturation results from the specific stimulation through these receptors (3). However, spontaneous maturation of ex vivo purified DCs undergo when cultured in vitro for a few hours in the absence of any specific activator indicating that DCs are exquisitely sensitive to any perturbation (4). While this spontaneous activation can occur during culture through the spontaneous release of type I IFNs, the sole DC isolation procedure could also deliver strong enough activation signals to induce DC maturation, even when they are further transferred into a physiological environment. This hypothesis is consistent with several studies reporting that transfer of peptide-loaded splenic DCs induces specific T cell responses and not tolerance, in the absence of deliberate DC activation and culture (5–7). As only mature DCs are able to prime naive T cells, this observation raises the question of DC activation state after their purification and transfer into host mice.
Furthermore, it was clearly shown that DCs have the ability to polarize CD4 T cell responses according to the signals they receive from the pathogens, the cytokine microenvironment or the antigen dose (8, 9). Kinetics studies have shown that recently activated DCs preferentially induced Th1 responses, whereas ‘exhausted’ DCs primed Th2 and non-polarized T cells indicating that kinetics of maturation have an important impact on polarization of primed T cells (10).
In this study, we investigated the induction of CD4+ T cell responses following injection of ex vivo unstimulated DCs after their purification from mouse spleen and the impact of the kinetics of DC maturation on in vivo priming of antigen-specific CD4+ T cells. We show that the sole isolation of DCs induces their maturation after their intravenous (i.v.) transfer into host mice and leads to the induction of Th1 response. These ‘non-activated’ DCs primed CD4+ T cells similarly to early and late CpG-activated DCs indicating that kinetics of maturation do not impact on the activation of T helper responses in vivo.
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Methods |
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Mice
Seven- to ten-weeks-old female or male C57BL/6 mice (H-2b) were purchased from CER Janvier (Le Gesnet St-Isle, France). OT-II mice transgenic for the TCR recognizing the I-Ab-restricted peptide corresponding to the residues 323–339 of ovalbumin (OVA) peptide (11) were bred on C57BL/6 background at the animal facilities of the Pasteur Institute. Ly5.1 C57BL/6 mice were obtained from CDTA (Orléans, France). TLR4-KO mice were kindly given by M. Chignard (Pasteur Institute). All mice were maintained under specific pathogen-free conditions. All animal work was approved by institutional animal experimentation committees.
Culture medium
Culture medium (CM) consisted of RPMI-1640 containing L-alanyl-L-glutamine dipeptide supplemented with 10% FCS, 5 x 10–5 M of 2-mercaptoethanol and antibiotics (penicillin 100 U ml–1 and streptomycin 100 µg ml–1). SFEM Stem Span (Stem Cell Technology, Meylan, France) containing antibiotics was used for DC loading with the OVA peptide.
Peptides and oligonucleotides
The synthetic OVA323–339 (ISQAVHAAHAEINEAGR) peptide corresponding to the immunodominant I-Ab-restricted CD4+ T cell epitope from OVA was purchased from Neosystem (Strasbourg, France). CpG oligodeoxynucleotide (ODN 1826; TCCATGACGTTCCTGACGTT, CpG ODN) and control (ODN 1982; TCCAGGACTTCTCTCAGGTT) were synthesized by GENSET (Paris, France).
LPS quantification
All media, buffers and reagents were tested for the presence of LPS using the Limulus, amebocyte lysate assay (Cambrex, Walkersville, MD, USA). The LPS content of the collagenase/DNase preparation and of the SFEM Stem Span medium were, respectively, 25 and 8 EU ml–1. For all other products, LPS content was <1 EU ml–1.
DC activation
For in vivo DC activation, 100 µg of CpG or control ODN were i.v. injected to C57BL/6 mice. DCs were purified from injected mice 4 or 18 h after ODN injection.
Cell purification
DCs were isolated from spleens of untreated, CpG ODN-injected or control ODN-injected mice. Spleens were harvested and treated for 45 min with 400 U ml–1 collagenase type IV and 50 µg ml–1 DNase I (Boehringer, Mannheim, Germany). For some experiments presented in Table 1, spleens were not treated by collagenase/DNase before purification. Spleens were then dissociated and single-cell suspensions were incubated with anti-CD11c-coated magnetic beads (N418 clone; Miltenyi Biotec, Bergisch-Gladbach, Germany) according to the manufacturer's procedure in PBS containing 2.5 mM of EDTA and 0.5% of BSA (Sigma–Aldrich, Saint Quentin Fallavier, France). After incubation, cells were washed and CD11c-positive cells were selected on an automated magnetic cell sorter (AutoMACS, Miltenyi Biotec) using posseld2 program. Harvested cell suspensions contained 90–95% of CD11c+ cells. For some experiments shown in Table 1, DCs were enriched by negative selection. RBCs were lysed using ACK lysing buffer (Cambrex) and then, cells were incubated with biotinylated anti-CD3 (clone 17A2, BD PharMingen) and anti-CD19 (clone 1D3, BD PharMingen) antibodies followed by incubation with streptavidin magnetic microbeads (Miltenyi Biotec) according to the manufacturer's procedure. Unlabeled cells were selected on AutoMACS (Miltenyi Biotec) using deplete program. Following this procedure, CD11c+ cells represented 30% of the cell suspension. OT-II transgenic T cells were isolated from lymph nodes (LNs) and labeled with 5 µM of 5,6-carboxyfluorescein diacetate succinimidyl ester (CFSE, Molecular Probes, Leiden, The Netherlands) for 15 min at room temperature, in the dark. Cells were then washed with CM.
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DC loading with the OVA peptide
Purified DCs were incubated with 1 µg ml–1 of OVA323–339 peptide in serum-free SFEM Stem Span medium for 30 min at 37°C and extensively washed to eliminate free peptide.
Adoptive transfer of T cells and DC immunization
A total of 106 CFSE-labeled OT-II T cells were i.v. injected to Ly5.1 C57BL/6 mice. After 24 h, the mice received 2 x 105 OVA323–339 peptide-pulsed or -unpulsed DCs by i.v. route.
Flow cytometry
All mAbs and isotype control antibodies were purchased from BD PharMingen (Le Pont de Claix, France). Isolated cells were suspended in PBS supplemented with 1% BSA and 0.1% NaN3. Cells were incubated with a rat anti-CD16/32 mAb (2.4G2 clone) in order to avoid non-specific binding and then labeled for 30 min in dark with the following mAbs: anti-CD11c (HL-3 clone), anti-CD40 (HM40-3 clone), anti-CD86 (GL1 clone), anti-Kb (AF6-88.5 clone), anti-CD45.1 (A20 clone), anti-CD45.2 (104 clone), anti-CD4 (L3T4 clone), anti-CD44 (IM7 clone) or anti-CD62L (MEL-14 clone).
For intracellular cytokine staining, unstimulated or CpG-activated DCs were incubated for 5–6 h in the presence of Golgi Plug (BD PharMingen). Then, cells were fixed and permeabilized before adding anti-IL-12p40 (C15.6 clone) or anti-tumor necrosis factor (TNF)- (MP6-XT22 clone) mAb.
Events were acquired on a FACSCan or FACSCalibur flow cytometer (BD Biosciences, San Diego, CA, USA) and analyzed using CellQuest Software (BD Biosciences).
Cytokine ELISA assay
Splenocytes from immunized mice were re-stimulated in vitro in the presence or absence of 10 µg ml–1 of OVA peptide in CM and culture supernatants were harvested 72 h later. IL-5, IL-10 and IFN-
concentrations were then measured by a standard sandwich ELISA. Maxisorp plates (Nunc, Roskilde, Denmark) were coated with unconjugated anti-IL-5, anti-IL-10 or anti-IFN- antibody (TRFK5, JES5-2A5 and R4-6A2 clones, respectively; BD PharMingen) and detection was done using the corresponding biotinylated mAb (TRFK4, SXC-1, XMG1.2 and C17.8 clones; BD PharMingen). The plates were developed using streptavidin–HRP (BD PharMingen) and o-phenylenediamine (Sigma–Aldrich) as substrate. All dosages were performed in duplicate. Assays were standardized with murine recombinant cytokines (BD PharMingen). Results shown are obtained by subtracting non-specific cytokine secretion in the absence of OVA323–339 peptide from specific secretion obtained in the presence of this peptide.
ELISPOT assay
The frequency of specific IFN--producing cells in the spleen of immunized mice was determined by an enzyme-linked immunospot (ELISPOT) assay as previously described. Ninety-six-well Multiscreen-HA sterile plates (Millipore, Molsheim, France) were coated overnight at 4°C with purified anti-IFN- mAb at 2 µg ml–1 in sterile PBS. Plates were washed five times with PBS and blocked with CM. Various numbers of splenocytes from immunized and control mice were then added to syngeneic-irradiated splenocytes (5 x 105 per well) in the presence or absence of 10 µg ml–1 of the OVA323–339 peptide, in CM. Plates were cultured for 36 h at 37°C and were then washed twice with sterile H20 Tween 0.5% and five times with PBS Tween 0.5%. Biotinylated antibody was added at 2 µg ml–1 in PBS Tween 0.5% and BSA 1%. Two hours later, the plates were washed five times with PBS Tween 0.5% and streptavidin–AKP was added to the wells in PBS Tween 0.5% FCS 1%. Two hours later, the plates were washed five times with PBS Tween 0.5% and twice with PBS and 100 µl of 5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium (BCIP/NBT) substrate (Sigma) were added in each well. After 10–15 min, the revelation was stopped with water. The spots were counted using the automated Bioreader-3000 pro counter (Bioreader, Karben, Germany). For each mouse, the number of peptide-specific IFN--producing cells was determined by calculating the difference between the numbers of spot generated in the absence and in the presence of OVA323–339 peptide. Results are expressed as spot-forming cells per number of splenocytes in the wells.
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Results |
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In vivo DC activation upon TLR9 triggering
Culturing DCs induces their maturation by a mechanism partially dependent upon autocrine type I IFN (4). In order to limit the effect of culture-induced signaling, and to analyze physiological DC activation as far as possible, DCs were stimulated in vivo by injection of CpG ODNs known to activate them through TLR9 stimulation (12). C57BL/6 mice were injected with 100 µg of CpG by i.v. route. Four and 18 h after CpG injection, splenic CD11c+ cell were enriched by magnetic cell sorting and expression of maturation markers and MHC molecules was compared with DCs purified from untreated mice by FACS analysis. Figure 1 shows that DCs freshly purified from untreated mice expressed low levels of co-stimulatory molecules and can be considered as immature DCs. By contrast, DCs purified from mice that received CpG 4 h before, expressed high levels of co-stimulatory and MHC class I and II molecules. Eighteen hours after CpG injection, DCs still expressed increased levels of these molecules. However, the expression of CD40, CD80 and MHC class II molecules was slightly lower whereas CD86 and class I expression was higher than on early-activated DC.
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We then analyzed cytokines produced by DCs following CpG stimulation. Freshly purified DCs were cultured for 6 h in the presence of brefeldine A and were then stained with specific mAb. As shown in Fig. 2, 30% of the 4 h CpG-activated DCs (CD11c+ cells) were positive for IL-12 and 24% for TNF-
. After 18 h, a dramatic decrease of the percentage of positive cells was observed for both cytokines indicating that most DCs reached final maturation. It is noteworthy that IL-12 and TNF--positive CD11c+ cells were also observed for DCs purified from control mice (mice injected with PBS or control ODN) indicating that these cells produced spontaneously these cytokines after purification and in vitro incubation in the presence of brefeldine. No positive cells were detected using control isotype antibody indicating the specificity of the labeling. This result indicates that DCs were activated to produce IL-12 and TNF- either during the purification protocol or during the incubation.
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Spontaneous phenotypic maturation of DCs upon i.v. transfer into host mice
To determine whether the purification procedure directly stimulates DCs, we purified splenic DCs from C57BL/6 mice based on their CD11c expression and transferred them directly into Ly5.1 congeneic mice without any culture step. Eighteen hours later, spleens of recipient mice were harvested and DCs were isolated and labeled. For comparison, we also analyzed DCs purified from spleens of mice previously injected with CpG, 4 or 18 h prior to purification. Owing to Ly5.1 and Ly5.2 markers, host and transferred DCs were distinguished and the level of CD86 expression by these DCs was compared. Figure 3(A) shows the phenotype of DCs purified from Ly5.1 C57BL/6 mice just before their transfer into Ly5.2 C57BL/6 mice. DCs purified from untreated mice were immature since these cells expressed low level of CD86, while DCs purified from CpG-treated mice were fully mature. Eighteen hours after their transfer, CD11c+ cells from spleen of the recipient mice were purified and analyzed. Figure 3(B) shows that 1–2% of splenic DCs were transferred DCs (characterized as CD11c+, CD45.2+ and CD45.1–). These cells showed a mature phenotype since they expressed high levels of CD86 molecules at their surface as compared with endogenous host DCs, whatever the treatment they received before purification and transfer. Thus, purification of DCs by positive selection of CD11c+ cells and further transfer of these cells lead to signaling that induced phenotypic maturation. Furthermore, these DCs entered the spleens of recipient mice after i.v. injection, and their migration was not affected by in vivo CpG activation prior to their purification. In order to determine if DC maturation was due to LPS-contaminating medium and/or reagents used during DC purification, their LPS content was tested. Low LPS concentration was found in the collagenase/DNase preparation (25 EU ml–1) and in the SFEM Stem Span medium (8 EU ml–1) that was used for peptide loading. Thus, the same type of experiment was repeated without collagenase/DNase treatment of the spleen and without any incubation into SFEM Stem Span medium. Under these conditions, a similar up-regulation of CD86 on DCs was observed after their transfer into recipient mice (Table 1). Furthermore, DC maturation was also observed following transfer of spleen DCs purified from TLR4-KO mice (Table 1) which are not able to respond to LPS. These results strongly suggest that the DC maturation observed was not due to LPS contamination. In addition, these data exclude that DC maturation was induced by cross-linking of CD11c during purification since DCs enriched by negative selection from splenocytes also matured after their transfer.
DCs purified from normal or CpG-treated mice induce similar T cell proliferation and activation in vivo
To determine the capacity of DCs to induce clonal expansion and activation of antigen-specific T cells in vivo, 1 x 106 CFSE-labeled LN cells from Ly5.2 OT-II transgenic mice were i.v. transferred into Ly5.1 C57BL/6 host mice. One day after, mice received 2 x 105 Ly5.2 DCs pulsed with the I-Ab OVA323–339 peptide specifically recognized by the OT-II T cells. Spleens of immunized mice were recovered 7 days later and spleen cells were labeled with anti-CD45.1 and anti-CD45.2 mAbs. CD45.2+ OT-II cells were analyzed by FACS for CFSE profiles and for the expression of the CD44 activation marker (Fig. 4). Mice immunized with unpulsed DCs showed neither T cell proliferation nor up-regulation of CD44 indicating that transferred OT-II cells did not proliferate spontaneously into recipient mice. By contrast, immunization of mice with OVA323–339 peptide-pulsed DCs lead to similar CD4+ T cell proliferation whether DCs were purified from untreated or CpG-injected mice. Furthermore, all cells that entered into division up-regulated CD44. These results indicate that untreated DCs or CpG-activated DCs induced similar proliferation and activation of CD4+ T cells.
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Untreated and CpG-activated DCs elicit Th1 response
To characterize the immune response of OT-II cells activated by DCs according to their maturation state, C57BL/6 mice received 1 x 106 LN cells from OT-II transgenic mice and then, 2 x 105 OVA323–339 peptide-pulsed DCs from untreated or CpG-injected mice. Spleen cells from immunized mice were harvested 7 days after DC injection, stimulated in vitro with the OVA333–341 peptide and cytokine production was measured by determining the concentration of cytokine released in culture supernatants. As shown in Fig. 5(A), splenocytes of mice immunized with OVA323–339 peptide-pulsed DCs produce a high rate of IFN-
. This production was associated with IFN--producing cells as detected by ELISPOT (Fig. 5B). Although the difference was not statistically different, higher response was observed following immunization with 4 h CpG-activated DCs, both by ELISA and ELISPOT assays. This result could indicate that early CpG-activated DCs are more efficient to induce IFN- production by CD4+ T cells. In some experiments and for some mice, low antigen-specific production of IL-4 and IL-5 was detected regardless the maturation status of injected DCs (data not shown). However, these responses were very heterogeneous among the different groups of mice. Finally, IL-10 was never detected in these experiments (data not shown).
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Discussion |
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DC maturation is a prerequisite for DCs to acquire T cell-priming ability (13). However, several studies have shown that DCs freshly purified from spleen and loaded with an epitopic peptide induce both CD4+ and CD8+ T cell responses in vivo, in the absence of deliberate DC activation (5–7). These observations suggest that DCs receive an activation signal either during purification procedure or during antigen loading that leads to their maturation. Here we show that the sole DC purification using the most common procedure of splenic DC isolation, i.e. immunomagnetic sorting with anti-CD11c microbeads after collagenase/DNAse treatment, induces up-regulation of co-stimulatory and MHC molecules after transfer into a syngeneic host by i.v. route. Furthermore, analysis of cytokine secretion shows that these purified DCs spontaneously produce IL-12 and TNF-
, two cytokines known to be strongly linked to DC activation (14). However, it is well established that in vitro culture induces DC maturation (4). Since cytokine intracellular staining requires a 6-h in vitro culture step, we cannot exclude that cytokine production could be induced by signal given during this in vitro step. DCs secreting these cytokines cannot correspond to DCs pre-activated in vivo since all ex vivo purified DCs display an immature phenotype. Altogether, these results demonstrate that splenic DCs receive one or several signals during their isolation, leading to full maturation.
DC activation could result either from direct stimulation or from the secretion of inflammatory cytokines (2) spontaneously released by DCs themselves or produced at their injection site. However, it was recently shown that indirect activation by inflammatory mediators alone induces DCs that support T cell expansion but fail to promote Th effector differentiation and to produce IL-12 (15). In contrast, the present study shows that freshly purified DCs produce IL-12 and induce proliferation and activation of antigen-specific IFN--producing CD4+ T cell effectors in vivo. These responses are similar to those induced by DCs activated by CpG, a strong adjuvant promoting Th1 response. Furthermore, this signal is strong enough for conferring to DCs the capacity to license CD8+ T cells to kill since we and others have shown that injection of freshly purified DCs loaded with antigenic peptide induce in vivo strong anti-viral-protective cytotoxic T cell responses and IFN--producing CD8+ T cells (5, 6). These results suggest that DC activation by purification is not only due to inflammatory mediators but rather to a strong and direct activation signal. Noteworthy, we can exclude that DC maturation was due to collagenase treatment or to CD11c cross-linking since DCs purified in the absence of collagenase/DNase digestion or following negative enrichment of spleen DCs after depletion of T and B cells by magnetic sorting still matured following in vivo transfer. However, this stimulation could be due to several non-physiological and non-specific factors such as the isolation of DCs from their physiological context.
These results are in contrast with the study of de Heusch et al. (16), showing that injection of freshly isolated splenic immature DCs induced the clonal expansion of CD4+ T cell that did not differentiate into effector cells. However, when IL-10 was neutralized, these DCs primed a Th1 response with IFN- production, indicating that they were able to generate effector CD4+ T cells, but also to induce the production of IL-10 that blocked activation and differentiation of T cells. The difference observed between this and our study could be due to the route of injection (injection of DCs into footpads and analysis of LN T cell responses in comparison to i.v. route and study of spleen cell responses). Indeed, after i.v. injection, DCs go directly into the spleen and are rapidly in contact with T cell. In contrast, following DC injection into footpads, DCs home to LNs by an active process and could encounter and interact with several cell types leading to the production of IL-10.
The induction of Th1 response could be related to the capacity of freshly isolated DCs to produce IL-12. This cytokine is known as a major Th1-promoting factor and is essential in generation of Th1-biaised cells from naive precursors (17). Low but significant numbers (5%) of freshly purified DCs spontaneously produced IL-12 during the 6 h that followed their isolation. Moreover, we can expect that higher percentage of the transferred cells produced IL-12 after their transfer since all DCs were mature 18 h after injection into recipient mice (18, 19). This would explain the quite similar efficacy of CpG-activated and untreated DCs to stimulate CD4+ T cell responses.
The present data challenge the idea that late-activated DCs prime Th2 and non-polarized T cells (10). Indeed, as expected, a higher IFN- response was obtained following immunization by early CpG-activated DCs as with to non-activated DCs, which could be correlated to the level of IL-12 production by these DCs. However, the kinetics of DC activation did not affect the polarization of T cell response since non-stimulated, early and late CpG-activated DCs promoted a Th1 T cell response although a lower response was obtained with late CpG-activated DCs. Several elements may explain this discrepancy between the study of Langenkamp et al. (10) and our results, as they worked in vitro with human DCs activated by LPS or poly-IC, while we used murine splenic DCs activated in vivo and injected back to their physiological environment. In our study, late DCs (18 h-activated DCs) were characterized phenotypically by a down-regulation of MHC class II molecule expression and a dramatic decrease of IL-12 production as compared with early CpG-activated DCs (4 h). Thus, they could be considered as exhausted DCs (10). However, since we did not check whether 18 h CpG-activated DCs were refractory to further activation by CD40-L, we cannot formally exclude that these DCs were still be capable to produce IL-12 in vivo after their transfer, upon interaction with T cells (18, 19). However, our results clearly establish that the injection of late-activated DCs neither induces Th2-biased T cell responses nor favors Th0 phenotype, showing that kinetics of DC activation do not significantly impact the polarization of T cell responses in vivo.
In conclusion, the present study highlights the bias that DC purification introduces in the analysis of the impact of maturation signals on DC ability to prime and polarize T cell responses. In line with other studies (20–22), our results suggest that immature DCs have to be studied directly in vivo, as their sole isolation may definitively alter their functional properties.
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Acknowledgements |
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This work was supported by a fellowship of the French Government to G.S. and by the Association pour la Recherche sur le Cancer. We thank M. Chignard for the kind gift of TLR4-KO mice and the European Community (grant NMP4-CT-2004-500039).
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Abbreviations |
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| CFSE | 5,6-carboxyfluorescein diacetate succinimidyl ester |
| CM | culture medium |
| DC | dendritic cell |
| ELISPOT | enzyme-linked immunospot |
| i.v. | intravenous |
| LN | lymph node |
| ODN | oligodeoxynucleotide |
| OVA | ovalbumin |
| TLR | Toll-like receptor |
| TNF | tumor necrosis factor |
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Notes |
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Transmitting editor: E. Vivier
Received 28 June 2005, accepted 13 December 2005.
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