Female C57BL/6 (H-2^b) and BALB/C (H-2^d) mice were purchased from Shanghai Laboratory Animal Center (Shanghai, China). All mice were maintained in a specific pathogen-free room at Animal Facilities of Soochow University. Experiments were carried out and approved according to the guidelines of the animal care and use committee at Soochow University. A20 lymphoma cells were purchased from American Type Culture Collection (Rockville, MD, USA). Cells were cultured at 37°C in a 5% CO₂ incubator in RPMI 1640 culture media supplemented with 10% fetal bovine serum (Biological Industries, Co., Haemek, Israel). For bioluminescent imaging, stable luciferase-expressing A20 cells (A20-luc) were generated in our laboratory.

Murine aGVHD and GVL models were induced as described previously (20, 21). Briefly, BALB/c mice were given lethally 650 cGy (one dose) total body irradiation from X-ray, irradiated BALB/c mice was transplanted with 1 × 10⁷ C57BL/6 bone marrow cells and 5 × 10⁶ C57BL/6 spleen cells via the tail vein. DMF (30 mg/kg body weight, Item No. 50744, Sigma-Aldrich, USA) was administrated to the recipient mice by gavage once daily starting from day −3 to day 3 after bone marrow transplantation (BMT). 0.8% methocel (Sigma-Aldrich Fluka, USA) at the same volume was used as vehicle control. For GVL model, 1 × 10⁶ A20-luc cells were added to bone marrow graft as mentioned above, and injected into lethally irradiated BALB/c mice. In vivo bioluminescence imaging was performed as described previously (20). Briefly, mice were given an intraperitoneal injection of 200 µg firefly luciferin and then anesthetized and imaged using Xenogen, IVIS 100 Bioluminescent Imaging System (Caliper Life Sciences, Hopkinton, MA, USA). Treg depletion was performed as described previously (22, 23). Briefly, lethally irradiated BALB/c mice were transplanted with 5 × 10⁶ TCD-BM plus 1 × 10⁶ total spleen T cells or CD25-depleted T cells from B6 mice, DMF was administered to these recipients, with vehicle treatment as control. T cell depletion was performed by anti-Thy1.2 mAb (30H12, Biolegend, USA) and rabbit complement (24). T cell purification was performed by using mouse T cell isolation kit (catalog #19851, Stemcell technologies, Vancouver, BC, USA), CD25 depletion was performed by using mouse CD25 regulatory T cell positive selection kit (catalog #18782, Stemcell technologies, USA) according to the manufacturer’s protocols. Unlabeled CD25 negative cells were collected. CD25 depletion efficiency was confirmed by FACS. The recipients were monitored daily for survival and every three days for body weight changes and clinical signs of GVHD. The severity of GVHD was assessed using a GVHD scoring system as described previously (20, 21).

Fourteen days after transplantation, liver, lung, small intestine and skin were obtained from the transplanted recipients and fixed in 10% formalin. Samples were then embedded in paraffin, sectioned were stained with hematoxylin and eosin. Tissue damage was assessed based on a semiquantitative scoring system as described previously (25, 26).

Mixed lymphocyte reaction assay was performed as described previously (27). Briefly, responder T cells were isolated from spleen of C57BL/6 mice by mouse T cell enrichment kit (StemCell Technologies, Vancouver, BC, Canada). Stimulators were DCs from BALB/c cells. BM-derived DCs were generated and expanded from BALB/c mice with GM-CSF (10 ng/ml) and IL-4 (10 ng/ml) for 7 days. DCs were pretreated with DMF or DMSO for 24 h, then washed twice with PBS. 1 × 10⁴ DCs treated as above were irradiated (30 Gy) and cocultured with 1 × 10⁵ allogeneic T cells in U-bottom microwell plates. 5 days later, tritiated thymidine (³H-TdR, 1 mCi/well) (Shanghai Institute of Physics, Chinese Academy of Sciences) were added to the culture for 16–18 h prior to harvesting and were counted on a β-plate reader (PerkinElmer Instruments, Meriden, CT, USA). Cytokines in the supernatants were collected and measured by ELISA. In some experiments, T cells were labeled with CellTrace CFSE (5 µmol/L, Invitrogen) according to the manufacturer’s protocol. The CFSE dilution was examined by flow cytometry. For ex vivo MLR, splenocytes from transplanted recipients 14 days after BMT were as responders, irradiated splenocytes from BALB/c mice were as stimulators. Cytotoxicity assays were performed as described previously (20). Splenocytes from transplanted recipients 14 days after BMT were used as killing cells, and their killing ability of A20 targets was measured using CytoTox 96 nonradioactive cytotoxicity assay kit (Promega, Fitchburg, WI, USA).

The procedure for isolating single-cell suspensions from spleens has been described previously (21). Antibodies against CD3, CD4, CD8, CD11c, CD25, CD40, CD44, CD69, CD86, PD-1, PD-L1, IFN-γ, H-2k^b, H-2k^d used in this study were all purchased from BioLegend (San Diego, CA, USA). For cell surface staining, cell samples were stained with fluorescent dye-conjugated mAb for 20 min at 4°C in the presence of FcR-Block. For intracellular cytokine staining, cells were stimulated for 5 h with PMA (50 ng/ml) and ionomycin (500 ng/ml) in the presence of brefeldin A (10 µg/ml). Cells were harvested, washed, and stained with surface molecule antibodies in the presence of FcR-Block (eBioscience, San Diego, CA, USA). After the wash, cells were then fixed using CytoFix/CytoPerm buffer (BD Biosciences, USA) and stained with antibodies against intracellular cytokines or isotype control on ice for 30 min. Intracellular staining for FoxP3 was performed by using a Foxp3 staining kit (eBioscience, San Diego, CA, USA). Data were acquired on a NovoCyte Flow cytometer (ACEA Biosciences, San Diego, CA, USA) and analyzed using Flowjo software (FlowJo, Ashland, OR, USA).

Blood samples were obtained from recipients 14 days after BMT, serum was separated by centrifugation and was stored at −80°C. Culture supernatants were collected at indicated time by centrifugation. The levels of IL-2, IL-6, IFN-γ, TNF-α, TGF-β were examined by ELISA kit according to the manufacturer’s instructions (R&D system, Minneapolis, MN, USA).

Total RNA were extracted with TRIzol reagent (Takara, Japan) from aGVHD targets organs according to the manufacturer’s instructions. Transcription levels of Nrf2, Keap1, HO-1, GST-α1, IL-1β, IL-2, IL-6, IFN-γ genes were analyzed by real-time PCR using SYBR Green Master Mix (Applied Biosystems, Warrington, UK). The primers used were: Nrf2, Forward 5′-TAGATGACCATGAGTCGCTTGC-3′, reverse 5′-GCCAAACTTGCTCCATGTCC-3′; keap1, forward 5′-TGCCCCTGTGGTCAAAGTG-3′, reverse 5′-GGTTCGGTTACCGTCCTGC-3′; HO-1, forward 5′-AAGCCGAGAATGCTGAGTTCA-3′, reverse 5′-GCCGTGTAGATATGGTACAAGGA-3′; GST-α1, forward 5′-AAGCCCGTGCTTCACTACTTC-3′, reverse 5′-GGGCACTTGGTCAAACATCAAA-3′; β-actin, forward 5′-ATCTGGCACCACACCTTC-3′, reverse 5′-AGCCAGGTCCAGACGCA-3′. The relative expression of the gene was quantified using the comparative 2^−ΔΔCt method relative to the housekeeping gene β-actin.

CD3⁺ T cells were isolated from spleen of C57BL/6 mice and activated by plate bound anti-CD3 (5 µg/ml) and anti-CD28 (1 µg/ml) in the presence of DMF or DMSO for 12 h. ROS levels were examined by Reactive Oxygen Species Assay Kit according to the manufacturer’s instructions (Beyotime, Shanghai, China). Briefly, cells were harvested and washed three times using RPMI-1640 media, 2′-7′-dichlorofluoresci1n diacetate was added at a final concentration of 10 µM to the cells. After a 20-min incubation at 37°C, cells were washed and resuspended in PBS. Data were acquired on a NovoCyte Flow cytometer and analyzed using Flowjo software.

For cell proliferation, 2 × 10⁴ A20 cells were seeded into 96-well plates and treated with DMF or DMSO for 48 h. Then 10 µl of CCK-8 solution was added to each well and incubated for additional 2 h, the absorbance was measured at 450 nm. Cell apoptosis was evaluated using a Annexin V Apoptosis Detection Kit (eBioscience, San Diego, CA, USA) according to the manufacturer’s instructions. Briefly, 48 h after DMF treatment, the cells were washed with PBS and resuspended in 500 µl binding buffer with 5 µl annexin V and 10 µl propidium iodide. Data were acquired by flow cytometry and analyzed using Flowjo software.

Survival data were analyzed by log-rank test and Kaplan-Meier survival curves were generated using GraphPad Prism version 5 (GraphPad 6.0, San Diego, CA, USA). The data were expressed as the mean ± SD. Two-tailed Student’s t-test was used for statistical comparison between two groups. One-way ANOVA with Dunnet’s test was used for multiple comparisons. The significance levels are marked *P < 0.05; **P < 0.01; ***P < 0.001.

Dimethyl fumarate has been demonstrated to potently inhibit NF-κB activity while promote Nrf2 activation. Therefore, we investigated the impact of DMF on alloreactive T cell responses in MLR assays. BMDCs from BALB/c mice were cultured with allogeneic splenic CD3⁺T cells purified from C57BL/6 mice, in the presence of the Nrf2 activator DMF (Figure 1A). The results showed that DMF significantly inhibited the proliferation of alloreactive T cells in a dose-dependent manner on day 5 determined by ³H-TdR and CFSE dye dilution (Figures 1B,C). Consistent with the T cell proliferation results, cytokine analysis showed that proinflammatory cytokines IL-2, IL-6, IFN-γ, and TNF-α production were significantly decreased in a dose-dependent manner upon DMF treatment (Figure 1D). In addition, cell apoptosis assay showed that the apoptosis of alloreactive CD4⁺ T cells were significantly increased in the presence of DMF (Figures 1E,F). More interesting, the apoptotic cells were mainly CFSE^low populations. To address whether DMF has a direct effect on T cells, we performed a T cell activation assay with anti-CD3/CD28 stimulation, the result showed that DMF could significantly inhibit T cells proliferation directly (Figure S1A in Supplementary Material). Additionally, we examined the effect of DMF on antigen presenting cells (APC). We cultured bone marrow DCs and then treated with DMF. The data showed that DMF had no effect on DC maturation, however, when DCs were preactivated by LPS, DMF could reduce CD80, CD86, and CD40 expression in a dose-dependent manner (Figure S1B–D in Supplementary Material). Similar results were observed in IL-6 and TNF-α expression (Figure S1E,F in Supplementary Material). Taken together, the results demonstrated that DMF could inhibit alloreactive T responses by suppressing cell proliferation and inducing cell apoptosis in vitro, through both direct effects on T cells and indirect effects on DCs.

To determine the possible role of Nrf2 activation in aGVHD, we first examined the Nrf2 expression after allo-HSCT. We observed that Nrf2 mRNA levels were significantly decreased in the spleen, liver, lung and intestine of allogeneic BMT mice compared to those of syngeneic control animals (Figure 2A), suggesting that Nrf2 suppression could be involved in the pathogenesis of aGVHD. We then evaluated the potential impact of DMF on the aGVHD development in vivo by using a murine aGVHD model. As shown in Figure 2B, DMF treatment significantly prolonged survival (log rank, P = 0.005) of the hosts following allo-HSCT (Figure 2B). In addition, aGVHD scores were reduced in DMF-treated mice compared with vehicle control recipients (Figure 2C). Histological analysis revealed that there was decreased pathological damage in the liver, lung, intestine and skin of recipients receiving DMF 14 days after allo-HSCT (Figure 2D). Similar protective effect was observed when the dose of DMF was changed to 60 mg/kg from day −3 to day +3 post-allo-HSCT (data not shown). Moreover, DMF administration did not affect donor chimerism 14 days posttransplant (data not shown). Therefore, these data suggested that Nrf2 activation by DMF treatment in vivo significantly improved aGVHD outcomes as evidenced by prolonged survival and reduced aGVHD scores following allo-HSCT.

To investigate the potential mechanisms responsible for reduced aGVHD severity by DMF treatment, we analyzed the alloreactive T cell responses in vivo. On day 14 after allo-HSCT, decreased infiltration of donor CD3⁺ T and CD4⁺ T cells was observed in spleen of DMF-treated recipients compared with vehicle control recipients (Figure 3A). The ex vivo MLR assay further demonstrated that DMF treatment significantly reduced donor T cell alloreactivity (Figure 3B). In addition, both donor CD4⁺ T and CD8⁺ T cells had a reduced activation phenotype indicated by CD69 levels in the hosts treated with DMF (Figure 3C). Furthermore, purified CD4⁺ T cells were activated in vitro by anti-CD3/CD28, and DMF treatment significantly inhibited T cell activation and downregulated CD69, CD44, as well as PD-1 expressions. The CD25 levels, however, were slightly upregulated by DMF treatment (Figure 3D). Intracellular staining revealed that the frequencies of IFN-γ-producing donor CD4⁺ T and CD8⁺ T cells in spleen were significantly decreased in recipients given DMF (Figure 3E). Analysis of serum samples on day 14 following allo-HSCT showed that the levels of proinflammatory cytokines IL-6, IFN-γ as well as TNF-α were significantly downregulated in DMF-treated recipients compared with vehicle control recipients (Figure 3F).

Since DMF is a potent activator of Nrf2 and can induce the Nrf2-dependent antioxidant response, we then examined the ROS production and antioxidant genes expression. We found that DMF could significantly inhibit ROS production by activated T cells in a dose-dependent manner in vitro (Figure 3G). Moreover, the Nrf2 levels and antioxidant defense enzymes HO-1 and GST-α1 expressions were upregulated in recipients treated with DMF (Figure 3H). We then explored the effect of DMF on nuclear translocation of Nrf2, which has been demonstrated as a major mechanism of function for DMF. We observed a dose-dependent effect of DMF on nuclear translocation of Nrf2 in the CD3⁺ T cells by immunofluorescent staining (Figure 3I). Taken together, these results suggested that DMF could inhibit donor T cell alloreactivity, and production of proinflammatory cytokines, as well as upregulate antioxidant enzymes, which led to the alleviation of aGVHD severity.

Dimethyl fumarate upregulated CD25 expression on CD4 T cells, suggesting that DMF may promote the generation of Treg cells. We examined the frequencies and absolute numbers of CD4⁺Foxp3⁺ Treg cells in spleen on day 14 after allo-HSCT. As shown in Figures 4A,B, the frequencies, as well as the numbers of Treg cells in spleen were significantly increased in DMF-treated compared with vehicle control recipients, suggesting that DMF could increase Treg cells in vivo. In vitro MLR assay showed that DMF promoted Treg cell differentiation in a dose-dependent manner (Figure 4C). To further confirm that the effect of DMF on aGVHD was partially dependent on the promotion of Treg cells, mice were injected with anti-CD25 antibody to deplete Treg cells in vivo. As shown in Figure 4D, depletion of CD25⁺ cells in DMF recipients aggravated aGVHD mortality compared with IgG control recipients, while it had no effect on vehicle treatment recipients. TGF-β has been known to promote Treg generation, we therefore examined the TGF-β levels in serum of GVHD mice and in MLR culture supernatants. Date shown that TGF-β was increased in DMF recipients (Figure 4E) and upregulated upon DMF treatment in vitro (Figure 4F). Therefore, these results suggested that the protective effect of DMF treatment on aGVHD was at least partially dependent on the promotion of Treg cells.

To evaluate the impact of DMF administration on GVL effects, aGVHD mice were challenged with A20-luc leukemia cells post-allo-HSCT. As shown in Figure 5A, mice transplanted with allo-BM alone plus A20-luc leukemia cells all died from leukemia within 40 days after transplant, regardless of whether DMF was administered. It has been reported that DMF could inhibit cell proliferation and induce apoptosis in a number of malignant cell lines including myeloid and lymphoid leukemia cell lines. However, the results in Figure 5A suggested that DMF may not directly affect A20 cell growth in vivo. DMF administration to mice receiving allo-BM and splenocytes, plus A20-luc showed prolonged survival compared with mice receiving vehicle control (Figure 5B), and low tumor burden was observed in DMF recipients as shown in bioluminescence imaging (Figure 5C), indicating the presence of GVL effect in DMF treated mice. The cell proliferation and apoptosis assay showed that low concentrations of DMF could not affect tumor growth and apoptosis in vitro, suggesting that DMF treatment inhibits A20 cell growth only at high concentrations, which may not be reached in vivo (Figures 5D,E). We also observed that donor T cells from DMF-treated recipients showed comparable, or even increased CTL killing activity against A20 leukemia cells compared to vehicle controls (Figure 5F). These findings further supported the observations that DMF treatment could preserve GVL effect after allo-HSCT.