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Journal of Leukocyte Biology logoLink to Journal of Leukocyte Biology
. 2008 Apr 22;84(1):124–133. doi: 10.1189/jlb.1107744

Differential regulation of naïve and memory CD4+ T cells by alternatively activated dendritic cells

Amy E Anderson *,1, Bethan L Sayers *,1, Muzlifah A Haniffa *,†,‡, David J Swan *, Julie Diboll *, Xiao-Nong Wang †, John D Isaacs *, Catharien M U Hilkens *,2
PMCID: PMC2504714  PMID: 18430785

Abstract

Promising immunotherapeutic tools for T cell-mediated pathologies are alternatively activated dendritic cells (aaDC), which exert their effect through the regulation and tolerization of T cells. As naïve and memory T cells have different susceptibilities to tolerogenic signals, it is important to understand the modulatory effects of aaDC on these T cell subsets. We have examined regulation of naïve and memory CD4+ T cells by human aaDC generated with dexamethasone, the active form of vitamin D3, 1α,25-dihydroxyvitamin D3, and LPS. Although aaDC induced low, primary, allogeneic responses by naïve and memory T cells, aaDC regulated the differentiation of these T cell subsets in a distinct manner. Naïve T cells primed by aaDC retained a strong, proliferative capacity upon restimulation but were skewed toward a low IFN-γ/high IL-10 cytokine profile. In contrast, memory T cells primed by aaDC became hyporesponsive in terms of proliferation and cytokine production. Induction of anergy in memory T cells by aaDC was not a result of the presence of CD25hi regulatory T cells and could be partially reversed by IL-2. Both T cell subsets acquired regulatory activity and inhibited primary CD4 and CD8 responses. Addition of exogenous IL-12p70 during T cell priming by aaDC prevented anergy induction in memory T cells and cytokine polarization in naïve T cells, indicating that the lack of IL-12p70 is a key feature of aaDC. Our finding that aaDC differentially regulate naïve and memory T cells is important for understanding and maximizing the therapeutic potential of aaDC.

Keywords: tolerance, cytokine deviation, hyporesponsiveness, immunotherapy

INTRODUCTION

Certain diseases, such as autoimmunity or graft-versus-host disease, are caused by an aberrant immune response to “self.” T cells are one of the main mediators of this response and are not only responsible for disease induction but are also involved in the development of a chronic immunopathogenic disease state. Attenuation of this abnormal T cell response is the main goal in the development of new therapies.

Promising, novel, immunotherapeutic tools for inhibiting T cell-mediated pathology are tolerogenic dendritic cells (tolDC). DC are professional APC that initiate and modulate immune responses. They display inherent plasticity and depending on their functional state, induce or inhibit T cell responses [1, 2]. tolDC are characterized by low levels of T cell costimulatory molecules (e.g., CD80, CD86) and/or production of anti-inflammatory cytokines (e.g., IL-10, IL-4). The mechanisms by which tolDC induce tolerance include T cell deletion, anergy, cytokine deviation, and/or the induction of regulatory T cells (Tregs) [3,4,5,6,7,8,9,10,11].

Immature DC (immDC) are an example of tolDC, but they are not stable and can become immunogenic by undergoing final maturation in response to proinflammatory cytokines and/or pathogen-derived molecules. immDC may therefore be unsafe to use as a therapeutic tool. However, stable tolDC can be generated in vitro by, for instance, genetic engineering or pharmacological modulation of DC [12, 13]. tolDC have great therapeutic potential: They can inhibit destructive immune responses in mouse models of bone marrow or organ transplantation and autoimmunity [14,15,16,17,18,19,20,21,22,23].

A novel approach to tolDC generation is “alternative activation” of DC (aaDC), which are first treated with immunosuppressive agents and then activated with LPS [23,24,25,26]. These aaDC resemble semi-mature DC (matDC), are stable, and have been shown to prolong organ allograft survival for longer than DC treated with immunosuppressive agents alone [26].

In relation to T cell-mediated disease, both naïve and memory T cell subsets cause immunopathology [27,28,29], but they have different activation requirements [30], and some tolerance-inducing strategies preferentially tolerize naïve but not memory T cells [31, 32]. The resistance of memory T cells to immunomodulation has been a serious problem. For example, it has been reported that a high frequency of antigen-experienced memory T cells correlates with an enhanced, post-transplant rejection risk [29]. The inclusion of aaDC in immunomodulatory therapeutic regimes in humans may prove beneficial, but it will be important to first establish that aaDC efficiently regulate not only naïve but also memory T cells.

We have investigated regulation of naïve and memory CD4+ T cells by human aaDC generated with dexamethasone (Dex), the active form of vitamin D3, 1α,25-dihydroxyvitamin D3 (VitD3), and LPS. An important and novel finding is that aaDC differentially regulate naïve and memory T cells: Naïve T cells were sensitized and polarized toward a low IFN-γ/high IL-10 cytokine profile, whereas memory T cells were anergized in terms of proliferation and cytokine production. Lack of IL-12p70 production was an important feature of aaDC that was at least partially responsible for their immunomodulatory effects.

MATERIALS AND METHODS

Isolation of cells from peripheral blood

Human samples were obtained with informed consent and following a favorable ethical opinion from North Tyneside Research Ethics Committee (UK). PBMCs were isolated from fresh blood or buffy coats by density centrifugation on Lymphoprep (Axis-Shield Diagnostics, Dundee, UK). CD14+ monocytes were isolated by positive magnetic selection using anti-CD14 magnetic microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). CD45RA+/RO– naïve and CD45RA–/RO+ memory CD4+ T cells were isolated by negative magnetic selection using RoboSep (StemCell Technologies, Vancouver, Canada). Purities of T cell subsets were routinely >95% for naive T cells and >90% for memory T cells. For some experiments, CD25hi cells were depleted from memory T cells with anti-CD25 beads (Miltenyi Biotec). CD4+CD25neg and CD4+CD25hi T cells were separated by first isolating CD4+ T cells using the RosetteSep CD4+ T cell enrichment kit (StemCell Technologies), followed by anti-CD25 beads (Miltenyi Biotec). CD8+ T cells were isolated by negative magnetic selection using the CD8 T cell isolation kit (Miltenyi Biotec).

Generation of DC populations

Monocytes were cultured at 0.5 × 106 cells/ml in the presence of IL-4 and GM-CSF (50 ng/ml each, ImmunoTools, Friesoythe, Germany) for 7 days with fresh medium and cytokines on Day 3. DC were matured on Day 6 by addition of LPS (1 μg/ml, Sigma Chemical Co., Poole, UK) for 24 h. aaDC were generated by addition of Dex (1×10−6 M, Sigma Chemical Co.) at Day 3 and Dex (1×10−6 M), the active form of VitD3, 1α,25-dihydroxyvitamin D3 (1×10−10 M, Leo-Pharma, Ballerup, Denmark), and LPS (1 μg/ml) at Day 6 for 24 h. All DC populations were washed extensively before using them in functional assays. Cells were cultured in RPMI 1640, supplemented with 10% FCS, 2 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin at 37°C with 5% CO2.

Flow cytometry

The following antibodies were used for cell surface marker analysis: CD3-FITC (HIT3a), CD3-PE (UCHT1), CD3-allophycocyanin (HIT3a), CD4-FITC (RPA-T4), CD4-PerCP (SK3), CD8-allophycocyanin (RPA-T8), CD25-FITC (M-A251), CD25-PE (M-A251), CD40-PE (5C3), CD69-PE/CD3-PerCp (L78/SK7), CD80-PE (L307.4), CD86-FITC (2331; all from BD PharMingen, San Jose, CA, USA); CD4-allophycocyanin (MEM-241; ImmunoTools); HLA-DR-FITC (B8.12.2; Immunotech, Marseille, France); CD83-FITC (HB15a; Immunotech); and CCR7-FITC (150503; R&D Systems, Abingdon, UK). Briefly, cells were centrifuged and resuspended in FACS buffer (PBS supplemented with 3% FCS, 2 mM EDTA, and 0.01% sodium azide). Human IgG (Grifols, Los Angeles, CA, USA) was added with antibodies to prevent FcR binding. Cells were incubated on ice for 30 min, centrifuged, and resuspended in FACS buffer. Intracellular forkhead box P3 (FoxP3) was detected using a FoxP3-allophycocyanin staining kit (PCH101; eBioscience, San Diego, CA, USA). Intracellular IFN-γ staining was performed using IFN-γ-PE (25723.11; BD PharMingen). Cells were stimulated with PMA (10 ng/ml) and ionomycin (1 μg/ml), and after 1 h, Brefeldin A (10 μg/ml) was added for an additional 4 h. Cells were harvested, surface-stained for CD3-allophycocyanin, and then fixed using fixation/permeabilization buffer (eBioscience). Cells were permeabilized using permeabilization buffer (eBioscience). To reduce background staining, the cells were blocked with 200 μg/ml mouse IgG (Sigma Chemical Co.) for 15 min prior to addition of the IFN-γ antibody. Cells were incubated at 4°C for 30 min, centrifuged, and resuspended in FACS buffer. Cell viability was assessed using Via-Probe (BD PharMingen), with or without Annexin-V-allophycocyanin (BD PharMingen). For intracellular staining, Via-probe was added prior to fixation and permeabilization. Data were collected on a BD PharMingen FACScan and analyzed using FlowJo (Tree Star, Ashland, OR, USA).

DC cytokine production and migration

Supernatants were harvested 24 h after activation of DC with LPS (matDC) or with Dex, VitD3, and LPS (aaDC) and stored at –20°C. Furthermore, matDC and aaDC were washed and stimulated at 1.5 × 105 DC/ml with CD40 ligand (CD40L)-transfected J558L mouse cells (kindly provided by Prof. Peter Lane, Birmingham University, UK) at a 1:1 ratio. After 24 h, supernatants were harvested and stored at –20°C. Cytokines (IL-12p70, IL-10, IL-6, and TNF-α) were quantified using sandwich ELISA (BD PharMingen). DC migration was assessed in a transwell system (pore size, 8 μm; Corning Life Sciences, UK). DC (2×105; input DC) were added in the upper chamber, and medium, with or without CCL19 (250 ng/ml, R&D Systems), was added to the lower chamber. Migration of DC was assessed after a 2-h incubation period at 37°C by harvesting the DC in the lower chamber and counting using a hemocytometer. DC migration is expressed as the percentage of input DC that had migrated.

DC-T cell cocultures

DC (1×104) were cultured with 1 × 105 allogeneic-naïve or memory T cells in 200 μl cultures. Supernatants were harvested after 4 or 6 days and assayed for IFN-γ by sandwich ELISA (BD PharMingen). Proliferation was assessed by incorporation of 3H-thymidine for the last 18 h of culture by scintillation counting (Microbeta TriLux, Perkin Elmer, Wellesley, MA, USA). Expression of CD69, CD25, and IFN-γ was assessed by flow cytometry as described above. Where indicated, recombinant (r)IL-12p70 (5 ng/ml, Hoffman-La Roche, Nutley, NJ, USA), rTNF-α (10 ng/ml, PeproTech, Rocky Hill, NJ, USA), rIL-6 (100 ng/ml, ImmunoTools), neutralizing anti-IL-10 (10 μg/ml, 25209, R&D Systems), or neutralizing TGF-β (10 μg/ml, 1D11, R&D Systems) were added in the DC-T cell cocultures.

T cell restimulation

Allogeneic-naïve or memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2 (kind gift from Prof. John Robinson, Newcastle University, UK). Anti-IL-10 and rIL-12p70 were added where indicated during priming. No residual aaDC were observed in the T cell lines. Primed T cells were washed and restimulated with matDC from the original DC donor (1:10) or T cell CD3/CD28 expander beads (1:1, Dynal, Invitrogen, Paisley, UK). Where indicated, rIL-2 (2 ng/ml) was added during restimulation. Supernatants were harvested after 72 h and assayed for IL-10 and IFN-γ by sandwich ELISA (BD PharMingen). Cultures were pulsed with 3H-thymidine for a further 8 h to determine proliferation. T cell proliferation was also measured by the red fluorescent dye PKH-26 (Sigma Chemical Co.) or CFSE (Sigma Chemical Co.). Briefly, primed, rested T cells were incubated at 25°C for 5 min with 2 × 10−6 M PKH-26 or at 37°C for 5 min with 0.5 μM CFSE, followed by serum quenching and washing. These cells were restimulated with matDC as described above, and proliferation was quantified with or without TruCOUNT tubes (BD PharMingen) by flow cytometry after 3–4 days. In some experiments, intracellular IFN-γ staining was also performed on CFSE-labeled cells as described above.

CD4 suppression assay

Allogeneic-naïve or memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2. Primed T cells were harvested, irradiated at 50 Gy, and washed. Irradiated, primed T cells or CD4+CD25hi T cells were cocultured with fresh, autologous CD4+CD25neg responder T cells (1:2), and matDC from the original DC donor were added in graded doses in 96-well round-bottom plates. After 5 days, cultures were pulsed with 3H-thymidine for a further 16 h to determine proliferation.

CD8 suppression assay

Allogeneic-naïve or memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2. Fresh, autologous CD8+ responder T cells were labeled with 0.5 μM CFSE for 5 min at 37°C followed by serum quenching and washing. Primed T cells, CD4+CD25neg T cells, or CD4+CD25hi T cells were cocultured with CFSE-labeled CD8+ responder T cells (1:2 or 1:20), and matDC from the original DC donor were added (one DC:80 responder T cells) in 96-well round-bottom plates. After 6 days, cultures were stained with CD8-allophycocyanin and were analyzed by flow cytometry.

Statistics

Paired t-test was performed using Prism 4.0 (GraphPad Software, San Diego, CA, USA). All P values are two-tailed.

RESULTS

aaDC have a semi-mature phenotype and an anti-inflammatory cytokine profile

We first analyzed cell surface phenotype and cytokine production by aaDC generated with Dex, VitD3, and LPS. aaDC displayed a semi-mature surface marker profile (Fig. 1 A). HLA-DR and CD80 expression was similar to matDC, but CD86 levels were lower. The classical DC maturation marker CD83 was only marginally expressed by aaDC. CD40 and the chemokine receptor CCR7 were expressed by aaDC but to a lower extent than by matDC. The cytokine profile of aaDC was anti-inflammatory with high levels of IL-10 but no detectable IL-12p70 and lower levels of TNF-α and IL-6 than matDC (Fig. 1B). The high IL-10/low IL-12p70 was a stable feature of aaDC, which was resistant to washing and restimulation with CD40L (Fig. 1C). Alternative activation of DC did not affect viability, as assessed by flow cytometry using Via-probe (matDC viability=86%±3%; aaDC viability=84%±3%; n=7).

Fig. 1.

Fig. 1.

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aaDC have a semi-mature phenotype and an anti-inflammatory cytokine profile. (A) immDC, LPS-activated matDC, and aaDC were stained with antibodies against surface markers as indicated. Debris and dead cells were excluded on the basis of forward-scatter, side-scatter, and viability staining with Via-probe. One representative experiment of 10 independent donors is shown. (B) Supernatants of DC were harvested 24 h after activation with LPS alone (to generate matDC) or LPS in the presence of Dex and VitD3 (to generate aaDC). Cytokines were quantified by sandwich ELISA. Data are presented as the mean ± sem of four independent experiments. (C) DC populations were washed, cultured at 1.5 × 105 DC/ml, and activated with CD40L-expressing cells (1.5×105/ml) for 24 h. Cytokines in supernatants were quantified by sandwich ELISA. Error bars represent sem of triplicates. One representative experiment of three independent donors is shown. Detection level of IL-12p70 ELISA was 30 pg/ml.

aaDC migrate in response to CCR7 ligand CCL19

As CCR7 expression was reduced on aaDC, we assessed their migratory capacity to the chemokine CCL19, a CCR7 ligand that directs DC to T cell areas in secondary lymph nodes. aaDC migrated in response to CCL19, albeit with 40% of the efficiency of matDC (Fig. 2).

Fig. 2.

Fig. 2.

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aaDC migrate in response to CCR7 ligand CCL19. Migration of matDC and aaDC was measured over a 2-h period in a transwell system. –CK, No chemokine; +CK, CCL19 (250 ng/ml) in the lower compartment. Migration efficiency was calculated as the percentage of the input DC that had migrated. Data are shown as the mean ± sem of three independent experiments.

aaDC have low immunostimulatory capacity for naïve and memory T cells

The T cell stimulatory capacity of aaDC was determined by coculturing them with allogeneic naïve or memory T cells. Compared with matDC, aaDC showed a reduced ability to induce T cell proliferation, production of IFN-γ, and expression of activation markers CD25 and CD69 (Fig. 3). In these primary DC-T cell cocultures, the immunostimulatory capacity of aaDC was low for naïve and memory T cells and was not caused by the induction of apoptosis (data not shown). The possibility that carry-over of immunosuppressive drugs was responsible for the low T cell stimulatory capacity of aaDC was excluded by extensive washing of all DC subsets prior to their use as APC and by demonstrating that aaDC supernatants harvested after washing did not inhibit T cell proliferation (data not shown).

Fig. 3.

Fig. 3.

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aaDC have low stimulatory capacity for naïve and memory T cells. matDC or aaDC (1×104 cells/well) were cocultured with allogeneic-naïve or memory T cells (1×105 cells/well). (A) 3H-Thymidine uptake and IFN-γ production were assessed at Days 4 and 6. (B) Naïve and memory T cells were stained for CD25/CD4 and CD69/CD4 before coculture with DC (left panels) and after 3 days of coculture with matDC (Tmat; middle panels) or aaDC (Taa; right panels). A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. Results are representative of three independent experiments. CD4-ACP, CD4-allophycocyanin.

aaDC sensitize naïve T cells for high IL-10 production but induce hyporesponsiveness in memory T cells

To further investigate the effects of aaDC on naïve and memory T cell responses, T cells were recovered from primary DC-T cell cocultures and restimulated under optimal conditions with matDC or CD3/CD28 beads. Proliferation and cytokine production of T cells that had been primed by aaDC (naïve or memory Taa) were compared with T cells primed by matDC (naïve or memory Tmat), revealing major differences in aaDC regulation of naïve and memory T cell responses. Upon restimulation, proliferative responses of naive Taa and naïve Tmat were comparable (Fig. 4 A). However, the cytokine profile of naïve Taa was significantly skewed toward higher IL-10 and lower IFN-γ production (Fig. 4B). To confirm that the enhanced IL-10 levels in naïve Taa cultures were as a result of its production by T cells, cytokine production was also measured after restimulation with CD3/CD28 beads in the absence of DC (Fig. 4C).

Fig. 4.

Fig. 4.

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Differential modulation of naïve and memory T cells by aaDC. Allogeneic-naïve or memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2. T cell lines primed by matDC (Tmat) and aaDC (Taa) were restimulated with matDC (A, B, D) or CD3/CD28 beads (C). (A) Proliferation was assessed on Day 3 by 3H-thymidine incorporation (left panel) or on Day 4 by measuring dilution of the red fluorescent dye, PKH-26 (right panel). A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. In the left panel, results of six independent experiments are depicted as the percentage proliferation of Tmat cell lines. (B) Supernatants of triplicate cultures were harvested at Day 3, and cytokine levels were measured by sandwich ELISA. Results are depicted as the percentage of cytokine production by Tmat cell lines: Tmat cell lines derived from naïve precursors produced 13.5 ± 9.1 ng/ml IFN-γ and 441 ± 361 pg/ml IL-10; Tmat cell lines derived from memory T cells produced 3.01 ± 2.8 ng/ml IFN-γ and 512 ± 379 pg/ml IL-10. Results of six independent experiments are shown. (C) IL-10 and IFN-γ in supernatants of naïve Tmat and Taa restimulated with CD3/CD28 beads were measured by specific sandwich ELISA. Results are shown as mean ± sem of six independent experiments. (D) CD25 expression and cell viability were tested after 3 days. CD25 expression is shown for T cells in the “Alive” gate; Via-probe/Annexin-V staining is shown for the “Total” gate, as indicated in the top row. A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. Horizontal lines represent median values; Pvalues represent significant differences between Taa and Tmat cells. SSC-H, Side-scatter-height; FSC-H, forward-scatter-height; Annexin V-APC, Annexin V-allophycocyanin.

In contrast to naïve T cells, memory Taa were hyporesponsive upon restimulation. Proliferation was reduced significantly in terms of percentages (Fig. 4A) and absolute numbers (Fig. 5A), T cells had a lower FSC/SSC (Fig. 4D, top row), indicative of less blast formation, and fewer T cells expressed CD25 (Fig. 4D, middle row). No polarization of the cytokine profile was observed; IL-10 and IFN-γ production was decreased compared with memory Tmat (Fig. 4B). The hyporesponsiveness of memory Taa was not a result of cell death, as there are similar percentages of Annexin-Vneg/Via-probeneg cells (viable cells) in memory Taa and memory Tmat populations (Fig. 4D, bottom row). Also, analysis of IFN-γ production by viable cells demonstrates that the lower cytokine production by memory Taa compared with memory Tmat is caused by hyporesponsiveness and not a decrease in cell viability (Fig. 5B). The percentage of IFN-γ-producing cells and the MFI for the IFN-γ+ cells are lower in the memory Taa compared with the memory Tmat. These data indicate that the number of IFN-γ producers and the amounts of IFN-γ produced are lower in the memory Taa than the memory Tmat population.

Fig. 5.

Fig. 5.

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Memory Taa are hyporesponsive: This is not a result of cell death, can be partially reversed by IL-2, and does not require CD4+CD25hi Tregs. (A–D) Allogeneic, total memory T cells or (E) CD25hi-depleted memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2. In some experiments, T cell lines primed by matDC (Tmat) and aaDC (Taa) were labeled with CFSE (A, C). (A) Primed T cells were restimulated with matDC, and absolute numbers of proliferating cells were determined on Day 3 by CFSE dilution and BD PharMingen TruCOUNT tubes. A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. The percentage of proliferating cells is shown, and the absolute number of proliferating cells is shown in bold. (B) Primed T cells were restimulated with 10 ng/ml PMA and 1 μg/ml ionomycin for 5 h in the presence of 10 μg/ml Brefelden A for the final 4 h. Viable cells were determined from the total population using Via-probe (left panel). Intracellular IFN-γ was assessed in viable cells by flow cytometry (right panel). Plots shown are gated on Via-probeneg cells. CD3-APC, CD3-allophycocyanin. MFI, Median fluorescent intensity. (C) Primed T cells were restimulated with matDC. On Day 3, cells were restimulated with PMA and ionomycin in the presence of Brefeldin A as described. Intracellular IFN-γ and proliferation, determined by CFSE dilution, were assessed using flow cytometry. A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. (D, E). Primed T cells were restimulated with matDC in the absence (D, E) or presence (D) of exogenous rIL-2 (20 ng/ml). Supernatants of triplicate cultures were harvested at Day 3, and cytokine levels were measured by sandwich ELISA and depicted as mean ± sem. (F) Memory T cells were stimulated with allogeneic DC for 6 days and rested for an additional 2 days. Expression of FoxP3 and CD25 was determined by flow cytometry. A CD3+CD4+ gate, identifying T cells, was used to exclude DC from subsequent analysis. All results are representative of two independent experiments. Foxp3-APC, FoxP3-allophycocyanin.

The lower cytokine production by memory Taa may be explained in part by the lower proliferative capacity of these cells (Fig. 5C). However, the percentage of proliferating cells, which produce IFN-γ, is lower in the memory Taa population (61%) as compared with the memory Tmat population (91%). Also, the MFI for IFN-γ in the proliferating cell population is lower for the memory Taa than the memory Tmat. These observations suggest that the proliferating Taa cells produce lower levels of IFN-γ than the proliferating Tmat cells, indicating that the memory Taa has a reduced ability to produce cytokine, which is independent of proliferative capacity. The hyporesponsiveness of memory Taa was partially reversible by restimulation in the presence of exogenous rIL-2 (Fig. 5D).

We considered the possibility that naturally occurring CD4+CD25hi Tregs in the memory population played a role in the induction of hyporesponsiveness by aaDC. However, depletion of CD4+CD25hi Tregs prior to the primary coculture with DC populations did not restore the impaired responses of memory Taa: Cytokine production (Fig. 5E) and proliferation (data not shown) remained low. We also could not show that aaDC preferentially induced FoxP3+ Treg expansion from memory (Fig. 5F) or naïve T cell populations (data not shown). However, care should be taken in interpreting data about FoxP3 expression in human T cells, as this is not an exclusive Treg marker but is also expressed in activated T cells [33, 34].

To summarize, these data demonstrate that aaDC differentially regulate naïve and memory T cell responses. aaDC sensitize naïve T cells for high IL-10 production, whereas memory T cells are rendered hyporesponsive. Induction of hyporesponsiveness does not require CD4+CD25hi Tregs and is not a result of the preferential expansion of Tregs.

aaDC-primed naïve and memory T cells acquire regulatory activity

The suppressive capacity of T cells primed by aaDC was assessed. Naïve and memory Taa inhibited proliferation of CD4+CD25neg and CD8+ T cells (Fig. 6). Suppression by memory Taa and (control) naturally occurring Tregs was comparable (Fig. 6), whereas naïve Taa were more potent. However, it should be noted that as a result of the method of isolation, a small proportion of effector T cells, which may have affected the suppressive ability of the Tregs, contaminated the naturally occurring Treg population. Inhibition by naïve Taa was not a result of their own proliferation, as potent, inhibitory effects were still observed at a CD8 responder/naïve Taa ratio of 20:1 (Fig. 6B) and also after irradiation of Taa (Fig. 6A).

Fig. 6.

Fig. 6.

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T cells primed by aaDC inhibit primary CD4+ and CD8+ T cell responses. Allogeneic-naïve or memory T cells were primed with DC (1:10) for 6 days and rested for 4 days with 0.1 ng/ml rIL-2. (A) Taa cells were irradiated (50 Gy) and added to cultures of autologous CD4+CD25neg T cells (ratio 1:2) and allogeneic matDC (ratios indicated in the figure). As controls, CD4+CD25neg or CD4+CD25hi T cells were used at similar ratios. Proliferation was measured after 6 days by 3H-thymidine incorporation, and results are depicted as percentage of suppression. Results are representative of three independent experiments. (B) Taa cells were added to cultures of autologous CFSE-labeled CD8+ T cells (ratio 1:2 or 1:20) and allogeneic matDC (one DC:80 CD8+ T cells). As controls, CD4+CD25neg or CD4+CD25hi T cells were used at similar ratios. CFSE content of CD8+ cells was analyzed after 6 days by flow cytometry. Results are representative of two independent experiments.

Lack of IL-12p70 production is an important feature of aaDC

To dissect the role of cytokines in aaDC regulatory effects, primary DC-T cell cocultures were performed in the presence of neutralizing antibodies to IL-10 or TGF-β or with the addition of the proinflammatory cytokines IL-12p70, TNF-α, or IL-6. None of these additions reinstated the T cell proliferation induced by aaDC to control levels (Fig. 7 A, upper panels), but the addition of IL-12p70 substantially enhanced production of IFN-γ by aaDC-primed T cells.

Fig. 7.

Fig. 7.

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Lack of IL-12p70 production is an important feature of aaDC. matDC or aaDC (1×104 cells/well) were cocultured with allogeneic-naïve or memory T cells (1×105 cells/well). (A) Neutralizing antibodies to IL-10 or TGF-β or the proinflammatory cytokines rIL-12p70, rTNF-α, or rIL-6 were added to the primary DC/T cell cocultures. Proliferation and IFN-γ production were measured at Day 5. (B) After 6 days of DC-T cell coculture, T cells were rested for an additional 4 days with suboptimal levels of rIL-2 (0.1 ng/ml). T cell lines primed by the different DC populations (Tmat or Taa) were restimulated with matDC. Proliferation was assessed on Day 3 by 3H-thymidine incorporation. Supernatants of triplicate cultures were harvested at Day 3. Cytokine levels were measured by sandwich ELISA and depicted as mean ± sem. Results are representative of three independent experiments.

Next, we determined whether exogenous IL-12p70 could overcome the regulatory effects of aaDC on naïve and/or memory T cells. IL-12p70 partially prevented the skewing of naïve T cells toward an anti-inflammatory cytokine profile: IFN-γ levels were restored, but enhanced IL-10 levels were not reversed completely (Fig. 7B). Interestingly, the presence of exogenous IL-12p70 during priming of memory T cells by aaDC reversed the induction of hyporesponsiveness in these T cells (Fig. 7B). Thus, the lack of IL-12p70 production is an important functional characteristic of aaDC.

DISCUSSION

This study focused on the regulation of naïve and memory T cells by aaDC derived with Dex, VitD3, and LPS. We found interesting differences in the modulation of naïve and memory CD4+ T cell responses by aaDC. Naïve T cells were sensitized by aaDC and polarized toward an anti-inflammatory cytokine profile. In marked contrast, memory T cells were anergized by aaDC in terms of proliferation and cytokine production. Naïve and memory Taa exhibited regulatory activity and suppressed primary CD4 and CD8 T cell responses. We found that deficient IL-12p70 production by aaDC was important for their regulatory actions.

This is the first report to demonstrate a differential effect of aaDC on naïve and memory T cells. These T cell subsets are known to have different susceptibilities to tolerogenic signals. For instance, anti-CD3 mAb or fixed APC readily induce anergy in memory T cells in vitro and in vivo but not in naïve T cells [35,36,37]. In contrast, some costimulatory molecule blockade-based, tolerogenic strategies (e.g., anti-CD40) are more effective in naïve than memory T cells [31]. Thus, depending on the tolerogenic method used, memory T cells have a higher or lower sensitivity to immunoregulation than naïve T cells. In our study, aaDC regulated memory and naïve T cells in qualitatively distinct but protolerogenic ways.

In contrast to our data, it was previously shown that DC generated in the presence of Dex induced anergy in naïve and memory T cell subsets [38]. However, in that study, Dex was added at the start of the monocyte culture, which has been shown to prevent their differentiation into DC [39]. We allowed for the differentiation of monocytes for 3 days before adding Dex, resulting in the generation of aaDC that expressed DC functional markers (e.g., CCR7) and retained the capacity to produce large amounts of IL-10 but no IL-12p70. The optimal type of DC for immunomodulatory therapy cannot be determined on the basis of in vitro data alone, but recent in vivo data demonstrated that aaDC (DC treated by Dex followed by LPS activation) prolonged organ allograft survival for longer than non-LPS-activated, Dex-treated DC [26]. The therapeutic potential of DC is determined by their ability to migrate to T cell areas in secondary lymph nodes in addition to their Treg activity. DC treated with immunosuppressive drugs without LPS do not express CCR7 and do not migrate in response to CCL19 (data not shown). It is reasonable to speculate that the superior in vivo therapeutic activity of aaDC may be, in part, a result of their migratory capacity.

aaDC polarized the cytokine profile of naïve T cells toward higher IL-10/lower IFN-γ production. Immune deviation of allo- or autoreactive T cells in this way may be beneficial for the prevention of graft rejection or the treatment of certain autoimmune diseases [40,41,42]. Bacchetta et al. [43] were one of the first groups to demonstrate that transplantation tolerance was induced by cells producing high levels of IL-10 and low levels of IFN-γ. In this regard, it is interesting to note that in patients undergoing liver transplantation, high IL-10 and low IFN-γ serum levels correlated well with graft acceptance [44,45,46], suggesting that cytokine deviation may be a contributing factor to prevention of graft rejection.

Our study did not find any evidence for the involvement of FoxP3+ Tregs in the regulatory activity of aaDC. Induction of anergy in memory T cells could not be prevented by depleting CD4+CD25hi T cells (Fig. 5E), and aaDC did not preferentially expand FoxP3+ T cells from memory (Fig. 5F) or naïve (data not shown) T cells. These observations are in contrast to a recent study in mice showing that aaDC preferentially expand FoxP3+ T cells [23]. The exact reason for this discrepancy is unclear but may be linked to the recent finding that in humans, FoxP3 is not exclusive to Tregs and is induced in all proliferating T cells [33, 34]. Nevertheless, despite the lack of FoxP3+ Treg expansion, aaDC-modulated T cells acquired regulatory activity and inhibited CD4+ and CD8+ T cell responses.

The contribution of the anti-inflammatory cytokine profile of aaDC to their regulatory effects was also investigated. Surprisingly, it was not the high IL-10 levels but lack of IL-12p70 that was important. Cytokine polarization of naïve T cells and anergy induction in memory T cells were reversed by IL-12p70 addition during T cell priming by aaDC, whereas neutralization of IL-10 had no effect (Fig. 6). In contrast to our data, Levings et al. [47] demonstrated that IL-10 is essential for Treg type 1 induction by immDC, although it is interesting to note that IL-10 alone is not sufficient to induce Treg type 1 differentiation [48]. A possible explanation for the requirement of IL-10 for immDC but not aaDC immunoregulation is the regulation of IL-12p70 production by endogenous IL-10. We have noticed that neutralizing IL-10 antibodies enhance IL-12p70 production by immDC but not aaDC (data not shown). Therefore, enhanced production of IL-12p70 may contribute to the reversal of the regulatory actions of immDC by IL-10 neutralization [47]. In our study, anti-IL-10 did not induce production of IL-12p70 by aaDC and did not reverse their Treg effects.

Although IL-10 production by aaDC did not appear important for their direct regulatory actions on T cells, high IL-10 production by these aaDC is likely to play an indirect role by down-regulating IL-12p70 production by other APC. This may be an important feature of aaDC, whose regulatory action may be overcome by IL-12p70 from other sources (Fig. 7). We therefore propose that the lack of IL-12p70 and high levels of IL-10 are important features of aaDC and should be included in quality control criteria for therapeutic aaDC preparations.

IL-12p70 is a well-known and potent, Th1-polarizing cytokine but has also been described to have a T cell costimulatory function. For instance, IL-12p70 can enhance IL-2-dependent proliferation of T cells by up-regulation of IL-2Rα [49]. Like IL-2, it can also reverse the anergic state of T cells in vitro [50], and the administration of IL-12p70 during tolerance induction in vivo can prevent T cell anergy [51, 52]. Interestingly, memory T cells have been shown to depend more on IL-12p70-mediated costimulation than naïve T cells, which were more dependent on costimulation by B7 [49, 53]. This differential dependence on costimulation by IL-12p70 may explain why memory T cells but not naïve T cells were rendered hyporesponsive upon priming by our aaDC.

In conclusion, aaDC generated with Dex, VitD3, and LPS have potent, regulatory effects on T cells and are a promising immunotherapeutic tool. Our finding that aaDC differentially regulate naïve and memory T cells is important for understanding their mechanism of action and will aid maximizing their therapeutic potential. The fact that both subsets are modulated in a protolerogenic direction is important in the context of their future therapeutic application.

Acknowledgments

This work was funded in part by the Arthritis Research Campaign (17750 to C. M. U. H.) and Action Medical Research (RTF1155 to M. A. H.). We thank Pawel Kalinski for helpful comments.

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