Nuclear factor (erythroid-derived 2)-like 2-fl/fl mice were a kind gift from Prof. Pi, China Medical University, Shenyang, China. CD11c-Cre mice were obtained from Prof. Steinkasserer, University of Erlangen-Nürnberg, Erlangen, Germany. Mice harboring a transgenic MOG-specific T cell receptor [2D2 mice (35)]; were a kind gift from Prof. V. Kuchroo, Boston, MA, USA. Wild-type (wt), 2D2, Nrf2KO, and CD11c-Cre/Nrf2-fl/fl mice were backcrossed on the C57BL/6J background and all mice were obtained from an in-house breed at the local animal care facility of the University of Erlangen-Nürnberg or the Ruhr-University Bochum under a 12-h day–night cycle and standardized environment. All experiments were performed in accordance with the German laws for animal protection and were approved by local ethic committees (Government of Unterfranken, Bavaria, Germany, ref. # 55.2-2532-2-451).

Mice received 15 mg/kg body weight DMF (Sigma-Aldrich, St. Louis, MO, USA) in 0.8 mg/ml hydroxypropyl methylcellulose (Sigma-Aldrich) twice daily by oral gavage, while control mice obtained 0.8 mg/ml hydroxypropyl methylcellulose. Mice were adapted to DMF treatment 2 weeks before EAE induction. Since the majority of DMF is rapidly metabolized to MMF by esterases in the intestine and only MMF can be detected in the systemic circulation (36, 37), we employed DMF to examine effects of Nrf2 pathway activation in our in vivo studies but MMF for in vitro stimulation experiments.

Induction of active EAE was performed as previously described (38, 39). For DC transfer experiments, 1 × 10⁵ DC, cultured in vitro for 10 days with or without 200 µM MMF (Sigma-Aldrich) and stimulated with 1 µg/ml lipopolysaccharide (LPS) (Sigma-Aldrich) for 24 h, were injected intraperitoneally at d2 and d7 after EAE induction together with 200 ng pertussis toxin (List Biological Laboratories).

Bone marrow cells were obtained from tibias and femurs and maintained in R10 medium (RPMI-1640 + 10% FCS + 1% penicillin/streptomycin + 1% l-glutamine) containing granulocyte macrophage colony-stimulating factor (GM-CSF) (5 ng/ml, Biolegend, San Diego, CA, USA) and interleukin (IL)-4 (5 ng/ml, Biolegend) for 10 days with/without 200 µM MMF (Sigma-Aldrich). Cells were matured on day 8 with 1 µg/ml LPS (Sigma-Aldrich). For flow cytometry, cells were stained with αCD11c (N418, Biolegend), αCD80 (16-10A1, Biolegend), αCD86 (GL-1, Biolegend), and αMHCII (AF6-120.1, BD Biosciences, San Diego, CA, USA).

Phagocytosis was assessed by stimulating DC with 1 µg/ml LPS (Sigma-Aldrich) for 48 h before adding opsonized Fluoresbrite^® YG Carboxylate Microspheres 0.75 µm (Polysciences Inc., Warrington, PA, USA). After 20 h of incubation, cells were subjected to flow cytometry and phagocytic cells were detected in the FITC channel.

The migratory capacity of DC was investigated using a modified Boyden chamber assay and transwell inserts with a 5 µm porous bottom (Corning Life Sciences, Corning, NY, USA). Mature DC were seeded into the migration chamber in medium containing 0.5% FCS. The lower chamber was supplemented with 10% FCS medium. After 20 h of migration, cells residing in the lower chamber (transmigrated fraction) were stained for CD11c and quantified by flow cytometry (FACSCantoII, BD Biosciences).

Mature DC were loaded with MOG_35–55 peptide (20 µg/ml) for 1 h and seeded with naive transgenic T cells harboring a MOG specific TCR at a ratio of 1:6. Naive MOG-specific transgenic T cells were isolated form the spleen of 2D2 mice by magnetic activated cell sorting using the “Naive CD4⁺ T Cell Isolation Kit, mouse” according to the manufacturer’s instructions (Miltenyi Biotech, Bergisch Gladbach, Germany). For Th17 differentiations, cocultured cells were additionally stimulated with IL-6 (40 ng/ml, R&D Systems, Minneapolis, MN, USA) and transforming growth factor beta (TGFβ1) (2 ng/ml, Biolegend). For Th1 differentiations, cells were cultured in the presence of IL-12p70 (20 ng/ml, R&D Systems) and anti-IL-4 (10 mg/ml, 11B11, Biolegend). For Treg differentiations, TGFβ1 (10 ng/ml, Biolegend) was added to culture media. T cell frequencies were analyzed by flow cytometry (FACSCantoII, BD Biosciences) using the antibodies listed in the section “Murine FACS analysis.” To monitor proliferation, cells were pulsed with [3H]-thymidine 24 h before analysis.

Differentiation of murine T cells was performed as previously described (40). Splenic T cells were isolated by magnetic activated cell sorting using the “pan T cell isolation kit II” according to the manufacturer’s instructions (Miltenyi Biotech). Cells were fluorescently stained with an antibody cocktail containing αCD4-FITC (RM4-5, eBioscience), αCD44-PE (IM7, Biolegend), αCD62L-APC (MEL-14, eBioscience), and αCD25-PE-Cy5 (PC61.5, eBioscience) and were subsequently isolated by fluorescence activated cell sorting on MoFlo (Beckman-Coulter, Brea, CA, USA) in the FACS-core unit of the University Hospital Erlangen. Sorted naive T cells (CD4⁺CD62L⁺CD44^lowCD25⁻) were stimulated by plate-bound anti-CD3 (2 µg/ml, 145-2C11, BD Pharmingen, San Diego, CA, USA) and soluble anti-CD28 (2 µg/ml, 37.51, BD Pharmingen) and cultured for 4 days in the presence of IL-6 (40 ng/ml, R&D Systems, Minneapolis, MN, USA) and TGFβ1 (2 ng/ml, Biolegend) for Th17, IL-12p70 (20 ng/ml,R&D Systems) and anti-IL-4 (10 µg/ml, Biolegend) for Th1 or TGFβ1 (10 ng/ml, Biolegend) alone for Treg differentiation. T cell frequencies were analyzed by flow cytometry (FACSCantoII, BD Biosciences) using the antibodies listed in the section “Murine FACS analysis.”

Spinal cords were disrupted with a 5 ml glass homogenizer followed by Percoll (GE Healthcare) density gradient centrifugation. Cell frequencies were analyzed by flow cytometry (FACSCantoII, BD Biosciences) using the antibodies listed in the section “Murine FACS analysis.”

As previously described (40), dead cells were excluded by the fixable viability dye eFluor^®780 (0.2 μl/test, eBioscience). Nonspecific Fc-mediated interactions were blocked by addition of 0.5 µl αCD16/32 (93, eBioscience). For surface staining, cells were treated with the respective fluorochrome-conjugated antibodies: αCD4-BV510 (RM4-5, Biolegend), αCD11c-FITC (HL3, BD Biosciences), αCD25-APC (PC61.5, eBioscience), αCD80-APC (16-10A1, Biolegend), αCD86-PerCP/Cy5.5 (GL-1, Biolegend), and αMHCII-BV510 (M5/114.15.2, Biole-gend).

For intracellular cytokine staining, cells were stimulated for 4 h with ionomycin (1 µM, Sigma-Aldrich) and PMA (50 ng/ml, Sigma-Aldrich) in the presence of monensin (2 µM, eBioscience), fixed with 1% paraformaldehyde and made permeable by saponin buffer treatment. Intracellular cytokines were stained with the respective fluorochrome-conjugated antibodies: αFoxp3-PE (FJK-16s, eBioscience), αIFNγ-APC (XMG1.2, eBioscience), and αIL-17A-PE (eBio17B7, eBioscience). Samples were measured with a flow cytometer (FACSCantoII, BD Biosciences).

For the generation of monocyte-derived DCs from leukoreduction system chambers of healthy donors (HDs) the positive vote from the local ethics committee has been obtained (Re.-Nr.: 4556). Generation of human monocyte-derived DC was performed as previously described (41). Peripheral blood mononuclear cells were prepared from leukoreduction system chambers of HDs by density centrifugation, followed by plastic adherence on tissue culture dishes (BD Falcon, San Diego, CA, USA). The non-adherent fraction was cryopreserved at −80°C for isolation of allogeneic T cells as described before (42), while the adherent cell fraction was cultured for 4 days with or without 50 or 200 µM MMF in DC-medium consisting of RPMI 1640 (Lonza, Basel, Switzerland) supplemented with 1% of heat-inactivated human serum type AB (Sigma-Aldrich), 1% Penicillin/Streptomycin/l-glutamine (Sigma-Aldrich), and 10 mM Hepes (Lonza) as well as 800 IU/ml (day 0) or 400 IU/ml (day 3) recombinant human GM-CSF and 250 IU/ml (days 0 and 3) recombinant IL-4 (both Miltenyi). Maturation of DC was induced by adding a maturation cocktail consisting of 200 U/ml IL-1β, 1,000 U/ml IL-6 (both CellGenix, Freiburg, Germany), 10 ng/ml tumor necrosis factor alpha (TNF-α) (Beromun, Boehringer Ingelheim, Germany), and 1 µg/ml PGE2 (Prostin E2, Pfizer, NJ, NY, USA) for 48 h. DC were analyzed by flow cytometry for activation marker expression: αCD80-PE (2D10.4, eBioscience), αMHCII-PE/Cy5 (LN3, eBioscience), αCD25-FITC (M-A251, BD Biosciences), αCD86-PE (IT2.2, eBioscience), αMHCI-FITC (W6/32, eBioscience), αCCR7-APC (3D12, eBioscience), αILT-4-FITC (27D6, eBioscience), and αCD83-PE (HB15e, eBioscience).

Mature human DC were generated as described above (41) and were either left untreated or were stimulated with maturation cocktail for 24 h. Allogeneic T cells were generated from cryopreserved non-adherent fraction. For this purpose, frozen cells were thawed, filtrated using a cell-strainer (Falcon) to remove dead cells and finally adjusted to 2 × 10⁶ cells/ml in MLR medium consisting of RPMI 1640 (Lonza) supplemented with 5% heat-inactivated human serum type AB (Sigma-Aldrich), 1% Penicillin/Streptomycin/l-glutamine (Sigma-Aldrich), and 10 mM HEPES (Lonza). Afterward, triplicates of 2 × 10⁵ allogeneic T cells were cocultured in 96-well flat cell culture plates (Falcon) with allogeneic DCs at different ratios for 72 h in MLR medium, and were finally pulsed with [3H]-thymidine (1 μC/well) for 8–16 h. Cultures were harvested onto glassfiber filtermates using an ICH-110 harvester (Inotech, Dottikon, Switzerland) and filters were counted in a 1450 microplate counter (Wallac, Turku, Finland).

Determination of cytokines (IL-6, TNF-α) secreted by human DC and/or T cells into the supernatant was assessed by Human Inflammation LEGENDplex™ (13-plex, Biolegend) according to the manufacturer’s protocol.

Cytokine concentrations in murine cell culture supernatants were analyzed by ELISA for the secretion of IL-6, TNF-α, IL-23, and IL-12 (R&D Systems) according to the manufacturer’s instructions.

RNA isolation, reverse transcription and PCR reactions were performed as previously described (40). Relative quantification was performed by the ΔΔCT method, normalizing target gene expression on actb/β-Actin or gapdh as housekeeping gene. The following TaqMan^® real-time PCR assays from Thermo Fisher Scientific were used: actb (β-Actin) Mm00607939_s1, nfe2l2 (nrf2) Mm00477784_m1, il6 Mm99999064_m1, nqo1 Mm01253561_m1, hmox1 Mm00516005_m1, il10 Mm01288386_m1, il12a Mm00434169_m1, arg1 Mm00475988_m1, tgfb Mm01178819_m1, nqo1 Hs02512143_s1, akr1c1 Hs04230636_sH, hcar2 Hs02341584_s1, and gapdh Hs02758991_g1.

Cultured cells were harvested in 1× Ripa lysis buffer [15 mM NaCl, 3.85 mM SDS, 5 mM Tris, 13.4 mM SDOX, 0.05 mM EDTA, 0.1% NP40, complete protease inhibitor cocktail Complete Mini and phosphatase inhibitor cocktail PhosStop (Roche Diagnostics, Basel, Switzerland)]. Protein concentration was determined using the BC Assay Protein Quantitation Kit (Interchim, Montluçon, France). Protein was detected using a rabbit polyclonal Nqo1 antibody (1:1,000, ab34173, Abcam, Cambridge, United Kingdom). Mouse anti-β-Actin (1:1,000) was obtained from Abcam (ab8226). Donkey antirabbit Alexa Fluor 488-conjugated and goat antimouse Alexa Fluor 647-conjugated secondary antibodies (1:1,000, Invitrogen, Carlsbad, CA, USA) were used. Detection was performed using a Fusion Capt Advance FX7 (peqlab, Erlangen, Germany).

Statistical testing was performed using GraphPad Prism (GraphPad Software Inc., San Diego, CA, USA). All in vitro and ex vivo data were analyzed by either one-/two-way ANOVA followed by Tukey’s posttest, unpaired t-test or Mann–Whitney test after checking for normal distribution (unless otherwise indicated). EAE data were analyzed by Mann–Whitney U-test. Data are presented as mean ± SEM; *p < 0.05, **p < 0.01, or ***p < 0.001 were considered to be statistically significant.

To analyze Nrf2 signaling in DC, we employed Nrf2-deficient cells. In a complementary approach, MMF treatment was used to induce Nrf2 signaling in DC. BMDC were cultured with 200 µM MMF either from the beginning (day 0) or starting at the day of LPS stimulation (i.e., day 8) to analyze if the time point of Nrf2 activation is crucial for its effect on DC maturation and surface marker expression. First, we examined whether activation of Nrf2-dependent signaling pathways influences BMDC generation in vitro. Yet CD11c⁺ cell frequencies were not altered in MMF-treated cell cultures (Figure 1A). However, the expression of MHCII, CD80 and CD86 was reduced when BMDC were generated under Nrf2 activating conditions (day 0) whereas addition of MMF on day 8 together with LPS, had almost no effect on activation marker expression (Figure 1B; Figure S1A in Supplementary Material).

To investigate whether the effect of MMF on BMDC surface marker expression is indeed dependent on Nrf2 induced signaling pathways, we treated Nrf2-deficient BMDC generated from either complete Nrf2 knockout mice (Nrf2KO) or conditional DC specific knockout mice (CD11c-Cre/Nrf2-fl/fl) with MMF. Again, the yield of CD11c⁺ cells did not differ between bone marrow obtained from wt, Nrf2KO, or CD11c-Cre/Nrf2-fl/fl mice and treatment with 200 µM MMF did not influence the generation of BMDC in the cell culture (Figure 1C). However, flow cytometric analyses revealed enhanced expression of CD80, CD86 (Figure 1D), and CD40 (Figure S1B in Supplementary Material) on Nrf2-deficient BMDC generated from either Nrf2KO or CD11c-Cre/Nrf2-fl/fl mice compared to wt BMDC. Additionally, MMF treatment had no influence on the frequency of CD80⁺/CD86⁺ and CD80⁺/CD40⁺ Nrf2-deficient BMDC but reduced the expression of CD80, CD86, and CD40 on wt BMDC (Figure 1D; Figure S1B in Supplementary Material).

In conclusion, modulation of the Nrf2 signaling pathway critically affects the activation status of DC in vitro.

Next, we assessed the influence of Nrf2 on different essential BMDC functions in vitro. As shown in Figure 2A, migratory capacities of BMDC isolated from Nrf2KO mice was significantly increased compared to BMDC from wt mice. Furthermore, MMF-treated wt BMDC showed a strongly reduced migratory capacity compared to untreated wt mice (Figure 2A). The phagocytosis rate of MMF-treated wt BMDC was not altered compared to untreated wt BMDC but decreased in Nrf2-deficient BMDC which correlates well with their enhanced maturation status (Figure 2B).

Expression analysis revealed that MMF treatment enhanced Nrf2 mRNA levels (Figure 2C) in wt BMDC in vitro. In turn, Nrf2 activation not only induced its antioxidative target genes nqo1 and hmox1 but also the anti-inflammatory cytokine il10 (Figures 2D–F). As expected, nfe2l2 (Nrf2), nqo1, and hmox1 expression was almost undetectable in Nrf2-deficient BMDC and tgfb expression levels were also significantly lower compared to wt BMDC (Figures 2C–E,G; Figure S2 in Supplementary Material). Furthermore, Nrf2 activation enhanced, while Nrf2 deficiency reduced, the expression of arginase 1 (arg1), a marker for tolerogenic DC (Figure 2H). Nrf2KO BMDC displayed a higher expression level of il12a compared to wt BMDC, while Nrf2 activation decreased proinflammatory cytokine expression on the mRNA level (Figures 2I,J). Nrf2 induction significantly reduced the production of IL-6, IL-12, IL-23, and TNF-α, proinflammatory cytokines that mediate T helper (Th)1 and Th17 differentiation, in wt but not Nrf2KO (data not shown) or CD11c-Cre/Nrf2-fl/fl BMDC (Figures 2K–N). Furthermore, Nrf2-deficient BMDC tended to produce higher baseline proinflammatory but lower tolerogenic cytokine levels compared to wt cells (Figure 2).

Immunoblot analysis of Nqo1 protein in wt and Nrf2-deficient BMDC cultures confirmed that the induction of Nqo1 expression in BMDC by MMF treatment is Nrf2 dependent (Figure 2O; Figure S3 in Supplementary Material).

Taken together, these data indicate that Nrf2 signaling alters DC functionality by modulating maturation, migration, expression of costimulatory molecules, and cytokine/chemokine production, which are required for effective antigen presentation to T cells.

In coculture experiments, untreated or MMF pretreated wt and Nrf2KO BMDC were cultured together with naive CD4⁺ T cells under Th1, Th17, or Treg cell polarizing conditions to demonstrate effects of Nrf2 signaling on DC-mediated T cell activation. Nrf2-deficient BMDC increased Th1 and Th17 but reduced Treg differentiation in coculture (Figures 3A–C). MMF pretreatment of wt BMDC had no effect on Th1 and Th17 frequencies (Figures 3A,B) but increased Treg differentiation (Figure 3C). Thymidine incorporation showed an enhanced proliferation rate of T cells that were cultured together with Nrf2KO BMDC and a reduced proliferation in the coculture with MMF pretreated wt BMDC (Figure 3D).

T cell monocultures demonstrated that Nrf2 had no direct effect on Th1 and Th17 cell differentiation (Figures 3E,F). However, Nrf2 deficiency induced a distinct reduction in Treg differentiation (Figure 3G).

These results show that Nrf2 signaling in DC may modulate their potential to regulate T cell differentiation and proliferation.

To demonstrate the relevance of the Nrf2 pathway in DC function for T cell-mediated autoimmunity in vivo, we employed the EAE animal model.

Spinal cords of DMF or control-treated wt and Nrf2KO mice were analyzed for DC frequencies and surface marker expression on day 14 after MOG-EAE induction (Figures 4A–C). Neither Nrf2 activation nor deficiency had an effect on the frequency of CD11c⁺ cells in the spinal cord of EAE mice (Figure 4A). In good accordance with the previous data, Nrf2 deficiency caused a significant increase in the expression of the activation markers CD80 and CD86 and enhanced MHCII expression on CD11c⁺ DC in the spinal cord (Figures 4B,C).

To further elucidate the effect of Nrf2 modulation on DC function during neuroinflammation, we employed MMF treatment as a potent Nrf2 inducer. In vitro cultured untreated or MMF-treated BMDC were injected into untreated wt mice at d2 and d7 after EAE induction. Mice which received MMF-treated BMDC showed an ameliorated clinical disease course compared to the control group (Figure 4D) and also a significantly reduced amount of total CD4⁺ as well as Th1 and Th17 cells in the spinal cord (Figure 4E).

Therefore, Nrf2 induction in DC in vitro may promote a tolerogenic phenotype enabling these cells to mitigate active EAE in vivo.

In a next step, we translated our findings from murine to human DC. Treatment of in vitro cultured human monocyte-derived DC with increasing amounts of the potent Nrf2 inducer MMF caused a concentration-dependent reduction of DC costimulatory, activation and inflammatory molecules including CD80, CD86, MHCI, MHCII, CD25, CD83, and C-C chemokine receptor type 7 (CCR7) as well as an increased expression of the tolerogenic surface receptor Ig-like transcript (ILT)-4 (Figures 5A–H).

Expression analysis showed that Nrf2 activation enhanced the expression of nqo1 and aldo-keto reductase family 1 member C1 (akr1c1) in human monocyte-derived DC analogously to murine BMDC (Figures 6A,B). Furthermore, MMF induced the expression of hydroxycarboxylic acid receptor 2 (hcar2), a niacin receptor involved in the regulation of inflammation and oxidative stress independently from Nrf2 signaling (Figure 6C). Human DC-treated with MMF also exhibited reduced tnfa and il6 but increased il10 and tgfb expression compared to the control group (Figures 6D–G).

Thymidine incorporation revealed a robust reduction in the T cell proliferation rate when T cells were cultured together with MMF pretreated human monocyte-derived DC compared to untreated DC (Figure 6H). To verify our PCR results on the protein level, cytokine production was measured in the supernatant of the aforementioned coculture assays. Here, IL-6 (Figure 6I) and TNF-α (Figure 6J) secretion was significantly reduced when DC were pretreated with MMF.

Complementary to the murine model, induction of Nrf2 signaling in human monocyte-derived DC modulates maturation marker expression, cytokine production and DC induced T cell proliferation.