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. Author manuscript; available in PMC: 2023 Jun 1.
Published in final edited form as: J Immunol. 2019 May 3;202(12):3447–3457. doi: 10.4049/jimmunol.1800018

In Vivo Generation of Gut-Homing Regulatory T Cells for the Suppression of Colitis

Yi Xu *,†,1, Yanmei Cheng *,‡,1, David J Baylink *, Samiksha Wasnik *, Gati Goel §, Mei Huang *,, Huynh Cao †, Xuezhong Qin *,‖, Kin-Hing William Lau *,‖, Christian Chan *, Adam Koch *, Linh H Pham *, Jintao Zhang *,#, Chih-Huang Li *,**,††, Xiaohua Wang *,‡‡, Edmundo Carreon Berumen *, James Smith §§, Xiaolei Tang *
PMCID: PMC10234421  NIHMSID: NIHMS1898214  PMID: 31053627

Abstract

Current therapies for gut inflammation have not reached the desired specificity and are attended by unintended immune suppression. This study aimed to provide evidence for supporting a hypothesis that direct in vivo augmentation of the induction of gut-homing regulatory T (Treg) cells is a strategy of expected specificity for the treatment of chronic intestinal inflammation (e.g., inflammatory bowel disease). We showed that dendritic cells (DCs), engineered to de novo produce high concentrations of both 1,25-dihydroxyvitamin D, the active vitamin D metabolite, and retinoic acid, an active vitamin A metabolite, augmented the induction of T cells that express both the regulatory molecule Foxp3 and the gut-homing receptor CCR9 in vitro and in vivo. In vivo, the newly generated Ag-specific Foxp3+ T cells homed to intestines. Additionally, transfer of such engineered DCs robustly suppressed ongoing experimental colitis. Moreover, CD4+ T cells from spleens of the mice transferred with the engineered DCs suppressed experimental colitis in syngeneic hosts. The data suggest that the engineered DCs enhance regulatory function in CD4+ T cell population in peripheral lymphoid tissues. Finally, we showed that colitis suppression following in vivo transfer of the engineered DCs was significantly reduced when Foxp3+ Treg cells were depleted. The data indicate that maximal colitis suppression mediated by the engineered DCs requires Treg cells. Collectively, our data support that DCs de novo overproducing both 1,25-dihydroxyvitamin D and retinoic acid are a promising novel therapy for chronic intestinal inflammation.

Inflammatory bowel disease (IBD) is a chronic inflammatory disorder in the gastrointestinal tract and affects~1.4 million Americans. It is believed that this disease is caused by a dysregulated immune response to intestinal bacteria and subsequent disruption of mucosal barrier (1, 2). Accordingly, current therapies aim at inhibiting the functions of inflammatory mediators or blocking the entrance of immune cells into intestines. However, these inflammatory mediators and immune cells are also required for immune defense. Hence, current therapies to a certain degree compromise systemic immune defense. Additionally, indiscriminate blockage of immune cell entrance into the intestines can weaken the intestinal immunity, which can lead to severe consequences (3, 4). Furthermore, such blockage also prevents regulatory T (Treg) cells from entering the intestines, which can potentially worsen the already-compromised gut immune tolerance in IBD patients and may also cause systemic immune suppression because of Treg accumulation in blood circulation (5). Finally, the therapeutic effects via blockage of molecules and cells are transient. Consequently, frequent administrations of these medications are necessary, further increasing the chance of causing undesired deleterious effects.

To tackle these challenges, we propose to use gut-homing Treg cells as a novel therapeutic strategy for IBD. In this regard, several lines of evidence support that this new strategy is particularly advantageous. First, gut-homing Treg cells can specifically home to intestines. Consequently, the suppression of inflammation is specific for intestines and will not cause unintended immune suppression outside intestines. Second, it has been demonstrated that Treg-mediated immune regulation can be potentially long lasting (6, 7). Hence, suppression of intestinal inflammation is stable and not transient. Third, because Treg cells target pathogenic mechanisms that are different from current medications, gut-homing Treg cells can potentially be an effective treatment for those IBD patients who do not respond or lose response to current medications.

Theoretically, generation of gut-homing Treg cells can be achieved through both in vitro and in vivo strategies. Because in vitro–generated Treg cells are unstable and can be potentially converted into pathogenic T cells under proinflammatory conditions (8, 9), we reason that an in vivo strategy, which can directly augment the induction of gut-homing Treg cells in peripheral lymphoid tissues, is preferable. In this regard, tolerogenic dendritic cells (TolDCs) are being actively investigated for in vivo augmentation of Treg cells (10). However, TolDCs are themselves produced in vitro and, hence, can be potentially converted into disease-worsening immunogenic dendritic cells (DCs). Therefore, TolDCs also face in vivo instability concern (11, 12). Additionally, previous data have shown that in vitro–generated TolDCs, unless further modified, do not have the capacity to stimulate gut-homing receptors in T cells (13).

To overcome the in vivo instability concern of in vitro–generated TolDCs, we propose to engineer DCs to de novo produce high concentrations of both 1,25-dihydroxyvitamin D [1,25(OH)2D] and retinoic acid (RA) for in vivo augmentation of the induction of gut-homing Treg cells. The rationale is that 1,25(OH)2D has been shown to induce the expressions of regulatory molecules (e.g., Foxp3 and IL-10) (14, 15) and RA has been shown to stimulate the expression of gut-homing receptors (16) in T cells. Importantly, 1,25(OH)2D as a Treg-inducing agent is particularly attractive because it regulates immune responses but does not cause indiscriminate immune suppression. As some examples of its uniqueness in immune regulation, 1,25(OH)2D can also stimulate the secretion of antimicrobials (17) and suppress carcinogenesis (18), which can prevent infections and cancers, respectively, in patients (19, 20). Based on these previous observations, we hypothesize that DCs engineered to de novo produce high concentrations of both 1,25(OH)2D and RA augment the induction of gut-homing Treg cells in peripheral lymphoid tissues to specifically and stably suppress chronic intestinal inflammation (e.g., IBD). This study provided strong evidence that supports this hypothesis.

Materials and Methods

Mice

BALB/c and C57BL/6 (B6) mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and housed in a specific pathogen–free animal facility at Loma Linda University (LLU). All mice were used at ages of 6–8 wk and allowed for an acclimation of minimum 5 d before any experimentation. All in vivo animal study protocols were reviewed and approved by both LLU Institutional Animal Care and Use Committee as well as the Animal Care and Use Review Office of the US Army Medical Research and Materiel Command of the Department of Defense.

Cell lines

A murine bone marrow–derived DC line (DC2.4) was kindly provided by Dr. K. Rock (Dana Farber Cancer Institute, Boston, MA) (21). DC2.4 cells were cultured at 37°C and 5% CO2 in RPMI 1640 culture medium containing 10% FBS (HyClone), 2 mM glutamine, 0.05 mM 2-ME, and 100 U/ml penicillin/streptomycin.

In vitro induction of Foxp3+CCR9+ T cells by 1,25(OH)2D and RA

Mouse (BALB/c) splenocytes were activated by CD3/CD28 Dynabeads (Thermo Fisher Scientific) (cells: beads = 2:1) in 24-well culture plates in a serum-free medium (X-VIVO 15; Lonza Bioscience) for 3 d in the presence of 50 U/ml recombinant human IL-2 and different concentrations of 1,25(OH)2D (Sigma-Aldrich) and RA (Sigma-Aldrich). The cells were then harvested and examined for the expressions of Foxp3 and CCR9 by FACS.

Preparation of plasmid constructs

1.6-kb mouse cytochrome P450 family 27 subfamily B member 1 (CYP27B1) and aldehyde dehydrogenase 1 family member A2 (ALDH1a2) cDNAs were amplified by PCR using a plasmid containing the CYP27B1 cDNA and a plasmid containing the ALDH1a2 cDNA, respectively (GeneCopoeia). The amplified CYP27B1 cDNA fragment with a 5′ KOZAK ribosome entry sequence was cloned into the pRRL-SIN.cPPt.PGKGFP.WPRE lentiviral vector (Addgene). The resulting construct was designated as lenti-CYP-GFP. The amplified ALDH1a2 cDNA fragment was cloned into the lenti-CYP-GFP to replace the GFP and was designated as lenti-CYP-ALDH. This bicistronic plasmid expresses CYP27B1 controlled by spleen focus–forming virus promoter and ALDH1a2 controlled by phosphoglycerate kinase promoter.

Preparation of lentiviruses

Three lentiviral transfer plasmids were used in this study: lenti-CYP-GFP, lenti-ALDH, and lenti-CYP-ALDH (see more descriptions of these lentiviral vectors in the Results). To generate a lentivirus that carried one of the lentiviral transfer plasmids, 293T cells were cultured in a complete DMEM culture containing 10% FBS, 100 U/ml penicillin/streptomycin, 0.05 mM 2-ME, 1 mM sodium pyruvate, 0.1 mM nonessential amino acid, and 2 mM l-glutamine. When the cells reached 70–80% confluence, culture media were replenished, and a transfection solution containing an envelope, a packaging, and the transfer plasmid was added to the cells dropwise. The cells were then cultured at 37°C and 5% CO2 for 24 h and the transfection solution was replaced with a different DMEM culture that contained 4% FBS, 100 U/ml penicillin/streptomycin, and 20 mM HEPES. After the cells were cultured at 37°C and 5% CO2 for 48 h, supernatants were collected, filtered through a 0.45-μm filter, and centrifuged at 4800 × g at 4°C for 24 h. The virus pellet was reconstituted in PBS containing 5% glycerol and titrated by a GFP-based FACS method. The typical titer of a virus was 108–109 transducing units/ml.

Generation of primary DCs from bone marrow and transduction of DCs with lentivirus

Bone marrow mononuclear cells were isolated from mouse tibias and femurs. The cells (1 × 106 cells/ml) were cultured in a RPMI 1640 culture containing 100 U/ml recombinant murine GM-CSF and 10 U/ml murine IL-4 (PeproTech) in a six-well culture plate at 37°C and 5% CO2. Forty-eight hours later, nonadherent cells were gently removed. The remaining adherent clusters were further cultured for another 48 h, and nonadherent cells were then harvested for lentivirus transduction. Briefly, 1 × 106 DCs per well were cultured in a total volume of 0.5 ml culture medium containing 50 μl virus (multiplicity of infection = 40) and 8 μg/ml protamine in a six-well culture plate. Twenty-four hours later, the virus was removed, and culture media were replenished. The cells were cultured for another 24 h and examined for transduction efficiency under a fluorescence microscope. When necessary, the above transduction procedure was repeated one more time. Twenty-four hours after the final virus removal, the DCs were activated by LPS (100 ng/ml for primary DCs and 1 μg/ml for DC2.4 cells) before use.

In vitro induction of Foxp3+CCR9+ T cells by DCs transduced with lenti-CYP-ALDH

Naive CD4+ T cells (5 × 105 cells per well), isolated from mouse (B6) spleens using a Naive CD4+ T Cell Isolation Kit (Miltenyi Biotec), were cocultured with DC2.4 cells (1 × 105 cells per well) or DC2.4-CYP-ALDH cells (1 × 105 cells per well) in 24-well culture plates in the X-VIVO 15 serum-free medium (Lonza Biosciences) in the presence of an anti-CD3 mAb (5 μg/ml) and recombinant human IL-2 (50 U/ml). In addition, the cocultures were supplied with various concentrations of 25-hydroxyvitamin D [25(OH)D] (Sigma-Aldrich) and retinol (Sigma-Aldrich). After 5 d of in vitro stimulation, the cells were collected and analyzed by FACS for the expressions of Foxp3 and CCR9.

In vitro evaluation of the enzymatic activities of 25(OH)D 1α-hydroxylase and retinaldehyde dehydrogenase 2

DCs were generated and transduced with the indicated virus. To evaluate the 25(OH)D 1α-hydroxylase (1α-hydroxylase) enzymatic activity, cells were seeded at a density of 5.0 × 105 cells/ml in a 12-well culture plate and 25(OH)D (Sigma-Aldrich) was added to the final concentration 2.5 μM. After incubation for 24 h, supernatants were collected and 1,25(OH)2D concentrations were measured using a radioimmunoassay by Heartland Assays (Ames, IA).

To evaluate the retinaldehyde dehydrogenase 2 (RALDH2) enzymatic activity, an ALDEFLUOR Kit (STEMCELL Technologies) was used according to the manufacturer protocol. Briefly, RALDH2 substrate (i.e., the BODIPY-aminoacetaldehyde) was added into the cell cultures in the presence or absence of the RALDH2 inhibitor diethylaminobenzaldehyde (DEAB;15 μM). The fluorescence product (i.e., the BODIPY-aminoacetate) was analyzed by FACS.

FACS analysis

FACS analysis was performed as previously described (22, 23). Briefly, ~0.5–1 × 106 cells in 100 μl FACS buffer (PBS containing 1% FBS and 0.05% sodium azide) were stained with various fluorescence-conjugated Abs specific for the cell surface proteins of interest at 4°C for 30 min. The surface-stained cells were then fixed and permeabilized using BD Pharmingen Cytofix/Cytoperm buffer. The cells were then stained with different fluorescence-conjugated Abs specific for the intracellular proteins of interest at 4°C for 30 min in BD Perm/Wash buffer. Finally, the cells were washed twice in the Perm/Wash buffer and twice in the FACS buffer before being analyzed on a BD FACSAria II. Data were analyzed using FlowJo software (Treestar).

Induction of experimental colitis

Experimental colitis was induced by intracolonic injection of 2,4,6-trinitrobenzenesulfonic acid (TNBS; Sigma-Aldrich) according to a previously reported method (24). Briefly, a 2.5% TNBS working solution in 50% ethanol was prepared by dissolving 5% TNBS (Sigma-Aldrich) in an equal volume of absolute ethanol. Subsequently, BALB/c mice were anesthetized by i.p. injection of 100 μl ketamine (12 mg/ml)/xylazine (1.6 mg/ml) solution per 10 g body weight. A 3.5–French gauge catheter connected with a 1-ml syringe was then carefully inserted into colons such that the catheter tip was 4 cm proximal to the anus. To induce mild colitis, 80–100 μl of the TNBS working solution was slowly administered into the colon lumens via the catheter. To induce severe colitis, 150 μl of the TNBS working solution was administered. Mice were then kept in a vertical position for 60 s and returned to their cages. Mortality and body weight were recorded daily.

DC transfer

DCs (1 × 106 cells in 100 μl PBS) were transferred i.v., i.p., or s.c. into mice as indicated in each experimental design. No substrates [i.e., 25(OH) D and retinol] were fed to the transferred mice.

Histological analysis

After being flushed with cold PBS, spleens or colons were fixed in 4% paraformaldehyde. The fixed tissues were then embedded within optimal cutting temperature compound and mounted on a chuck in a cryostat for sectioning. The tissues were cut into 10-μm-thick frozen sections which were collected by adhering to slides and stored at −20°C. Spleen tissue sections were analyzed by fluorescence microscope for Qtracker 655–labeled DCs. Colon tissue sections were examined by H&E (Sigma-Aldrich) staining, which was also used for the determination of microscopic scores, as previously described (25).

Depletion of Foxp3+ Treg cells

The PC61 5.3 anti-CD25 mAb (Bio X Cell) was used to deplete Foxp3+ Treg cells in vivo, as previously reported (26, 27). The Ab was injected i.p. at 250 μg/mouse every other day for four times beginning 2 d before the induction of TNBS colitis.

Statistical analysis

Statistical analyses were performed using GraphPad Prism software (GraphPad Software). For experiments that contained more than two groups, ANOVA tests were used. For experiments that contained only two groups, Student t tests were used. Data were presented as means ± SEM. Results were considered significant when the p value was <0.05.

Results

A combined action of 1,25(OH)2D and RA augmented the induction of Foxp3+CCR9+ T cells in vitro

A necessary premise of our proposed novel strategy is the ability of a combined action of 1,25(OH)2D and RA to program naive T cells when being activated, into gut-homing Treg cells. Hence, we first determined the 1,25(OH)2D concentration that was necessary for augmenting the induction of Foxp3+ T cells in vitro. We used anti–CD3/CD28 for activating splenocytes in the presence of various concentrations of 1,25(OH)2D. Seventy-two hours later, the cells were examined for the expression of Foxp3 in CD3+ T cells by FACS. Our data showed that 100 nM but not 20 nM of 1,25(OH)2D significantly augmented the expression of Foxp3 in CD3+ T cells (Supplemental Fig. 1A, 1B).

Because CCR9 is a gut-homing receptor (28), we next asked whether RA could synergize with 1,25(OH)2D in augmenting the induction of Foxp3+CCR9+ T cells in vitro. To answer this question, we added various concentrations of RA (e.g., 1, 10, and 100 nM) into purified CD4+ T cells that were activated by anti–CD3/CD28 in the presence of the Foxp3-inducing 1,25(OH)2D concentration (100 nM). Our data demonstrated that RA dose-dependently increased the percentage of Foxp3+CCR9+ cells in the activated CD4+ T cell population (Supplemental Fig. 1C, 1D).

DCs engineered to de novo produce high concentrations of both 1,25(OH)2D and RA augmented the induction of Foxp3+CCR9+ T cells in vitro

The 1,25(OH)2D concentration (100 nM) necessary for significantly augmenting the induction of Foxp3 expression is much higher than the physiological levels which are~16–56 pg/ml (0.038–0.13 nM) (29, 30). Although some patients may tolerate up to 300 pg/ml (0.72 nM) of serum 1,25(OH)2D levels without the dose-limiting toxicity (i.e., hypercalcemia) (30), this dose (0.72 nM) is still far below the Foxp3-inducing dose defined in our in vitro system. Additionally, recent data demonstrate that inflamed tissues are insensitive to 1,25(OH)2D (31). Hence, our data and those of others strongly indicate that systemic administration is extremely difficult to achieve the in vivo 1,25(OH)2D concentrations that are necessary for stable augmentation of the induction of Treg cells, especially under chronic inflammatory conditions such as IBD (32, 33). Although not intensively investigated, RA may also face similar challenges. With these foregoing data, as well as our previous success in the delivery of locally high 1,25(OH)2D concentrations without the risk of hypercalcemia (22, 34), we reason that a thorough evaluation of our proposed strategy is warranted to determine the potential use of 1,25(OH)2D and RA for in vivo augmentation of the induction of gut-homing Treg cells.

To facilitate testing our hypothesis, we generated three lentiviral vectors (Fig. 1A). The first lentiviral vector was the lenti-CYP-GFP that carried the CYP27B1 gene encoding for the 1a-hydroxylase. Because the 1α-hydroxylase is the physiological enzyme for catalyzing the synthesis of 1,25(OH)2D, we reason that DCs transduced with the lenti-CYP-GFP (DC-CYP cells) can de novo produce high 1,25(OH)2D concentrations in the presence of 1α-hydroxylase substrate [i.e., 25-hydroxyvitamin D or 25(OH)D, which is the major vitamin D form in blood circulation]. The second lentiviral vector was the lenti-ALDH that carried ALDH1a2 gene encoding for RALDH2. Because the RALDH2 is the physiological enzyme that catalyzes the synthesis of RA, we reason that DCs transduced with the lenti-ALDH (DC-ALDH cells) can de novo produce high RA concentrations in the presence of RALDH2 substrate (i.e., retinol, the major vitamin A form in blood circulation). The third lentiviral vector was the lenti-CYP-ALDH that carried both the CYP27B1 and the ALDH1a2 genes encoding for the 1α-hydroxylase and the RALDH2, respectively. We reason that DCs transduced with the lenti-CYP-ALDH (DC-CYP-ALDH cells) can de novo produce high concentrations of both 1,25(OH)2D and RA.

FIGURE 1.

FIGURE 1.

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Engineered DC-CYP-ALDH cells augmented the induction of Foxp3+CCR9+ T cells in vitro. (A) Lentiviral vectors used in this study. See description of the three vectors in the text. Spleen focus–forming virus (SFFV) and phosphoglycerate kinase (PGK) are promoters. (B) A stable DC2.4-CYP-ALDH cell line was generated by transducing DC2.4 cells with the lenti-CYP-ALDH. Subsequently, CD4+ naive T cells were isolated from mouse spleens. The CD4+ T cells were then activated in cultures by anti-CD3 mAb (5 μg/ml) in the presence of either parental DC2.4 cells or the DC2.4-CYP-ALDH cells. Additionally, the cultures were added with the indicated concentrations of 25(OH)D and retinol. Five days later, the cells were collected and analyzed by FACS for the expressions of Foxp3 and CCR9 in CD3+CD4+ T cell population. Representative FACS plots show the expressions of Foxp3 and CCR9 in the CD3+CD4+ T cell population under the indicated conditions. (C) Cumulative data (n = 4) of (B). *p < 0.05, ANOVA.

Next, we asked whether the DC-CYP-ALDH cells were able to induce gut-homing Treg cells in vitro. To answer this question, we took advantage of the DC2.4 cells [a DC line that was generated from B6 bone marrow and previously shown to recapitulate the functions of primary bone marrow–derived DCs (21, 22, 35, 36)]. Specifically, we transduced the DC2.4 cells with the lenti-CYP-ALDH to generate a transgenic DC2.4 cell line that stably overexpressed both the 1α-hydroxylase and RALDH2. This engineered cell line is hereafter called DC2.4-CYP-ALDH cells. Subsequently, purified naive B6 CD4+ T cells were activated by an anti-CD3 mAb in the presence of either parental DC2.4 cells or the DC2.4-CYP-ALDH cells. Additionally, substrates for the 1α-hydroxylase [i.e., 25(OH)D] and for the RALDH2 (i.e., retinol) were added into the cultures. In this system, differentiation of the CD4+ T cells was determined by the signals derived from either the DC2.4 cells or the DC2.4-CYP-ALDH cells. Our data showed that, in the presence of the substrates, the DC2.4 cells did not significantly change the abundance of Foxp3+CCR9+ cells in the CD4+ T cell populations (Fig. 1B, 1C). In contrast, the DC2.4-CYP-ALDH cells significantly enhanced the abundance of Foxp3+CCR9+ CD4+ T cells. Additionally, the more 25(OH)D added, the greater ability of the DC2.4-CYP-ALDH cells to increase the abundance of Foxp3+CCR9+ CD4+ T cells. Collectively, we conclude that the DC-CYP-ALDH cells are able to augment the induction of Foxp3+CCR9+ T cells in vitro.

Transfer of the DC-CYP-ALDH cells enhanced the induction of Foxp3+CCR9+ T cells in peripheral lymphoid tissues

Based on our in vitro observations, we next asked whether the DC2.4-CYP-ALDH cells were able to enhance the induction of gut-homing Treg cells in peripheral lymphoid tissues in vivo. To facilitate activation of a sufficient amount of Ag-specific T cells in vivo within a short period of time, we transferred the DC2.4-CYP-ALDH cells, which have an MHC background of B6 mice (i.e., H-2b), into allogeneic BALB/c mice that have an MHC background of H-2d. We predicted that the donor DC2.4-CYP-ALDH cells would stimulate a strong allogeneic CD4+ T response in the BALB/c hosts (37). Accordingly, we i.v. transferred DC2.4 cells, DC2.4-CYP cells (DC2.4 cells that were transduced with the lenti-CYP-GFP for a stable overexpression of the 1α-hydroxylase), or DC2.4-CYP-ALDH cells into BALB/c mice. Additionally, mice that did not receive any DC transfer were included as controls. Ten days after the DC transfer, mononuclear cells in spleens and mesenteric lymph nodes were examined by FACS (Fig. 2A). Our data showed that the DC2.4-CYP-ALDH cells, when compared with the controls, significantly enhanced the expressions of Foxp3 (Fig. 2B, 2C), IL-10 (Fig. 2D, 2E), and CCR9 (Fig. 2F, 2G) in CD3+ T cells.

FIGURE 2.

FIGURE 2.

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DC-CYP-ALDH cell transfer enhanced the induction of Foxp3+CCR9+ T cells in peripheral lymphoid tissues. (A) BALB/c mice received i.v. one of the following DC transfers (Transfer): no DC transfer (No transfer), parental DC2.4 cells, DC2.4-CYP cells, or DC2.4-CYP-ALDH cells. Ten days later, spleens and mesenteric lymph nodes (MLNs) were analyzed by FACS. (B) Representative FACS plots show the expression of Foxp3 in T cells. (C) Cumulative data of (B). (D) Representative FACS plots show the expression of IL-10 in T cells. (E) Cumulative data of (D). (F) Representative FACS plots show the expression of CCR9 in T cells. (G) Cumulative data of (F). (H) BALB/c mice received i.p. the DC transfers as shown in (A). Four days later, MLNs were analyzed by FACS. (I) Representative FACS plots show the expressions of Foxp3 and CCR9 in CD3+ T cell population. (J) Cumulative data of (I) show the percentage of Foxp3+CCR9+ cells in CD3+ T cell population. Cells were gated on CD3+ T cells for all the analyses. Where applicable, the data presented are means ± SEM (n = 4–6). *p < 0.05, **p < 0.01, ANOVA.

In another set of experiments, we i.p. transferred the various DC2.4 cell lines into BALB/c mice. Four days after the DC transfer, mesenteric lymph nodes were examined by FACS (Fig. 2H). Our data showed that the DC2.4-CYP-ALDH cell transfer, when compared with the controls, significantly increased the abundance of Foxp3+CCR9+ cells in CD3+ T cell population (Fig. 2I, 2J). Based on these results, we conclude that the DC-CYP-ALDH cell transfer significantly augments the induction of Foxp3+CCR9+ T cells in peripheral lymphoid tissues.

Bone marrow–derived DCs could be engineered to de novo produce high concentrations of both 1,25(OH)2D and RA

Our in vitro and in vivo data from study of the DC2.4-CYP-ALDH cells strongly suggest that the DC-CYP-ALDH cells are a promising therapy for chronic intestinal inflammation (e.g., IBD). Therefore, we sought to determine whether the DC-CYP-ALDH cells were able to suppress ongoing experimental colitis induced by TNBS through augmentation of the induction of gut-homing Treg cells in peripheral lymphoid tissues. The TNBS colitis was chosen because it is a well-established Th1-mediated disease (38), and a recent study, using adoptive transfer of IFN-γ–deficient Th17 cells, demonstrates that Th1 cells are required for the induction of colitis (39).

However, the DC2.4-CYP-ALDH cells are not suitable for the analysis of colitis suppression because DC2.4 cells have an MHC background of H-2b and TNBS colitis is induced in BALB/c mice that have an MHC background of H-2d. For this reason, we decided to use bone marrow–derived primary syngeneic DCs in the following studies. Accordingly, we first asked whether CD11c+ bone marrow–derived DCs (Fig. 3A) could be engineered to overexpress both 1α-hydroxylase and RALDH2 for de novo production of high concentrations of both 1,25(OH)2D and RA. We showed that the DC-CYP-ALDH cells, when compared with parental DCs, displayed enhanced expression of the 1α-hydroxylase as determined by FACS (Fig. 3B). To determine functional activity of the overexpressed 1α-hydroxylase, we added in the DC cultures 1α-hydroxylase substrate [i.e., 25(OH)D]. Twenty-four hours later, 1,25(OH)2D concentrations in the culture supernatants were measured with a radioimmunoassay by Heartland Assay in Ames, IA. Our data showed that 1,25(OH)2D concentrations in culture supernatants of the DC-CYP and DC-CYP-ALDH cells were each~20-fold higher than those of parental DCs and the DC-ALDH cells (Fig. 3C). Hence, our data demonstrated that the 1α-hydroxylase activity was significantly augmented in the DC-CYP and DC-CYP-ALDH cells. To evaluate functional activity of the overexpressed RALDH2, we added a commercially available, nontoxic RALDH2 substrate (i.e., the BODIPY-aminoacetaldehyde) into the cultures of parental DCs and the DC-CYP-ALDH cells. Fluorescent product (i.e., the BODIPY-aminoacetate) in the DCs and the DC-CYP-ALDH cells was then determined by FACS. Our data showed that mean fluorescence intensities of the BODIPY-aminoacetate in the DC-CYP-ALDH cells were~6-fold higher than parental DCs, suggesting that the DC-CYP-ALDH cells, when compared with parental DCs, expressed significantly increased RALDH2 enzymatic activity (Fig. 3D, 3E). Accordingly, we conclude that primary DCs can be engineered to de novo produce high concentrations of both 1,25(OH)2D and RA.

FIGURE 3.

FIGURE 3.

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Bone marrow–derived DCs could be engineered to de novo synthesize high concentrations of both 1,25(OH)2D and RA. Primary DCs were generated from bone marrow and transduced with the lenti-CYP-GFP to generate the DC-CYP cells, the lenti-ALDH to generate the DC-ALDH cells, and the lenti-CYP-ALDH to generate the DC-CYP-ALDH cells. (A) A representative FACS plot shows the expression of DC marker (i.e., CD11c). (B) A representative FACS plot shows the expression of 1α-hydroxylase in parental DCs and the DC-CYP-ALDH cells. (C) 1α-hydroxylase substrate [i.e., 25(OH)D] was added into the DC cultures. Twenty-four hours later, supernatants were collected and measured for 1,25(OH)2D concentrations. The data show concentrations of 1,25(OH)2D in the cultures of parental DCs, DC-CYP cells, DC-ALDH cells, and DC-CYP-ALDH cells (n = 4). **p < 0.01, ANOVA. (D) RALDH2 activity was measured using the ALDEFLUOR assay. Representative FACS plots show the BODIPY-aminoacetate fluorescence in DCs and DC-CYP-ALDH cells in the presence or absence of RALHD2 inhibitor DEAB. (E) Mean fluorescence intensities (MFIs) of BODIPY-aminoacetate in DCs and DC-CYP-ALDH cells in the absence of DEAB (n = 4). *p < 0.05, t test.

DC-CYP-ALDH cells survived and expressed the two overexpressed enzymes for at least 30 d in vivo

Next, we asked whether the DC-CYP-ALDH cells were able to survive for a period of time similar to parental DCs to program T cells in peripheral lymphoid tissues. To address this question, we traced transferred DC-CYP-ALDH cells in spleens of recipient mice. Accordingly, DCs or DC-CYP-ALDH cells were labeled with Qtracker 655 and i.v. transferred into BALB/c mice. At days 7, 14, and 30, spleens were collected from the mice and analyzed by real-time quantitative PCR for the mRNA expressions of CYP27B1 and ALDH1a2. Additionally, the day 30 spleens were also analyzed for the Qtracker 655 fluorescence. Our data showed that the mRNA expression of CYP27B1 in spleens of the mice transferred with the DC-CYP-ALDH cells, when compared with parental DCs, were significantly increased at days 7 and 14 and remained increased at day 30 (Fig. 4A). Additionally, the mRNA expression of ALDH1a2 in spleens of the mice transferred with the DC-CYP-ALDH cells, when compared with parental DCs, were significantly increased at days 14 and 30. Moreover, analysis of the Qtracker 655 fluorescence demonstrated that the DC-CYP-ALDH cells and parental DCs could still be seen in the spleens with similar abundances at day 30 after the in vivo transfer (Fig. 4B, 4C). In conclusion, the DC-CYP-ALDH cells can survive in vivo for a period of time similar to parental DCs (40), and they stably expressed the two overexpressed enzymes within our observation window.

FIGURE 4.

FIGURE 4.

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DC-CYP-ALDH cells expressed transgenes and survived for at least 30 d in vivo. (A) Qtracker 655–labeled DCs or DC-CYP-ALDH cells were i.v. transferred into BALB/c mice. At days 7, 14, and 30, spleens were collected from the mice and analyzed by real-time quantitative PCR for the mRNA expressions of CYP27B1 and ALDH1a2. (B) The day 30 spleens from (A) were analyzed by fluorescence imaging. Left panel shows representative images of Qtracker 655 fluorescence (original magnification ×20). Right panel shows average Qtracker 655+ cells per scan. Where applicable, the data presented are means ± SEM (n = 3). *p < 0.05, **p < 0.01. #, not significant.

s.c. immunization with Ag-pulsed DC-CYP-ALDH cells increased the abundance of Foxp3+ and IL-10+ T cells specific for the Ag in intestines

Using primary DC-CYP-ALDH cells, we then asked whether s.c. immunization with Ag-pulsed DC-CYP-ALDH cells could lead to an increased abundance of Ag-specific Treg cells in intestines, which would be anticipated if the immunization augmented the induction of gut-homing Treg cells in draining lymph nodes. To monitor in vivo migration of newly generated gut-homing Treg cells, we pulsed the DC-CYP-ALDH cells with a peptide of chicken OVA (i.e., OVA323–339) such that the immunization mainly activated and expanded in draining lymph nodes OVA323–339–specific CD4+ T cells, which could be identified by I-Ad/OVA323–339 tetramer (tet) in BALB/c mice (41). Accordingly, we immunized BALB/c mice at days 0 and 10. Ten days after the last immunization, colon tissues were examined by FACS (Fig. 5A, 5B). Our data showed that mice s.c. immunized with the OVA323–339–pulsed DC-CYP-ALDH cells, when compared with the controls, showed significantly increased I-Ad/OVA323–339 tet+ cells in Foxp3+ T cell population in the colons (Fig. 5C, 5D). Because DC-CYP-ALDH cell transfer also increased the abundance of IL-10+ T cells (Fig. 2D, 2E), we analyzed Foxp3− cells. Our data showed that the abundance of I-Ad/OVA323–339 tet+ cells were also significantly increased in the Foxp3− T cell population (Fig. 5E, 5F). Importantly, in the Foxp32I-Ad/OVA323–339 tet+ cell population, the abundance of IL-10+ cells was significantly increased in colons of the mice immunized with the OVA323–339–pulsed DC-CYP and DC-CYP-ALDH cells when compared with the controls (Fig. 5G, 5H).

FIGURE 5.

FIGURE 5.

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s.c. immunization with Ag-pulsed DC-CYP-ALDH cells augmented the abundance of Foxp3+ and IL-10+ T cells specific for the Ag in intestines. (A) BALB/c mice were s.c. immunized (Immun) with OVA323–339–pulsed one of following DCs at days 0 and 10: parental DCs, DC-CYP cells, or DC-CYP-ALDH cells. A group of mice that did not receive any immunizations (No immun) was also included as a control. At day 20, colon tissues from all the mice were analyzed for the OVA323–339–specific T cells, identified by I-Ad/OVA323–339 tet. (B) Gating strategy. (C) Representative FACS plots show I-Ad/OVA323–339 tet+ cells in CD3+Foxp3+ T cell population. (D) Cumulative data of (C). (E) Representative FACS plots show I-Ad/OVA323–339 tet+ cells in CD3+Foxp3− T cell population. (F) Cumulative data of (E). (G) Representative FACS plots show IL-10+ cells in CD3+Foxp3−I-Ad/OVA323–339 tet+ cell population. (H) Cumulative data of (G). Where applicable, the data presented are means ± SEM (n = 4–6). *p < 0.05, **p < 0.01, ***p < 0.001, ANOVA. #, not significant.

Transfer of DC-CYP-ALDH cells robustly suppressed ongoing experimental colitis

The foregoing observations support the possibility that the DC-CYP-ALDH cells can effectively suppress TNBS colitis. To address this possibility, we determined the effects of DC-CYP-ALDH cell transfer on TNBS colitis. Specifically, BALB/c mice received DC transfer once at day 3 after the induction of TNBS colitis (Fig. 6A). Our data showed that the colitic mice transferred with parental DCs displayed 100% mortality (Fig. 6B, upper panel). In contrast, the colitic mice transferred with the DC-CYP-ALDH cells showed 0% mortality and those transferred with the DC-CYP cells had~50% mortality. In addition, the colitic mice transferred with the DC-CYP-ALDH cells, when compared with those transferred with the DC-CYP cells, were much quicker in regaining body weight (Fig. 6B, lower panel). Furthermore, 14 d after colitis induction, colon lengths were measured. Our data showed that the colitic mice transferred with parental DCs, when compared with the normal healthy mice, displayed significantly shortened colon lengths. In contrast, colon lengths in the colitic mice transferred with the DC-CYP-ALDH or the DC-CYP cells were each significantly longer than those transferred with parental DCs. Additionally, colon lengths in the colitic mice transferred with the DC-CYP-ALDH cells were significantly longer than those transferred with the DC-CYP cells. Indeed, colon lengths in the colitic mice transferred with the DC-CYP-ALDH cells were similar to those in the healthy control mice (Fig. 6C). Moreover, H&E staining showed that microscopic scores of colon tissues from the colitic mice transferred with parental DCs, when compared with healthy mice, were significantly higher (Fig. 6D, 6E). Additionally, microscopic scores of colon tissues from the colitic mice transferred with the DC-CYP-ALDH cells, when compared with those transferred with parental DCs and the DC-CYP cells, were significantly lower.

FIGURE 6.

FIGURE 6.

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DC-CYP-ALDH cell transfer, when compared with controls, more robustly suppressed TNBS colitis. (A) BALB/c mice were induced for TNBS colitis. At day 3, mice received i.v. one of the following DC transfers (Transfer): 1 × 106 per mouse parental DCs, 1 × 106 per mouse DC-CYP cells, or 1 × 106 per mouse DC-CYP-ALDH cells. Additionally, a group of healthy mice was also included as a control. The mice were then monitored daily for survival and body weight. At day 14, distal inflamed colons were harvested and analyzed by H&E staining. (B) Percent of survival (upper panel) and baseline body weight (lower panel) are shown over the observation period. (C) A comparison of the lengths of distal inflamed colons among the experimental groups of mice. (D) Representative images of H&E-stained distal inflamed colons from the experimental mice. Original magnification ×5. (E) Average microscopic scores from (D) are shown. Where applicable, the data presented are means ± SEM (n = 4–6). **p < 0.01, ***p < 0.001, ****p < 0.0001, ANOVA.

Maximal colitis suppression following the DC-CYP-ALDH cell transfer required Treg cells

To determine the role of Treg cells in the DC-CYP-ALDH cell–mediated colitis suppression, healthy control BALB/c mice i.v. received different DC transfers. Seven days after the DC transfer, CD4+ T cells purified from the spleens were i.v. injected into syngeneic hosts that were induced for a mild TNBS colitis 3 d before the CD4+ T cell injection (Fig. 7A). Our data did not show significant difference in body weights among all the experimental groups of mice (Fig. 7B). We reasoned that this was because of the mild colitis that caused recovery of all the experimental mice beginning 1 d after the CD4+ T cell injection (day 4). Hence, we performed a second induction of a severe colitis after the mice totally recovered from the first episode of colitis (i.e., day 13). Our data showed that mice injected with the CD4+ T cells from both the DC-CYP and DC-CYP-ALDH cell–transferred mice, when compared with the controls, experienced significantly less reduction of body weight. In conclusion, DC-CYP-ALDH cell transfer augments regulatory activity in CD4+ T cell population in peripheral lymphoid tissues.

FIGURE 7.

FIGURE 7.

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Mice injected with the CD4+ T cells from DC-CYP and DC-CYP-ALDH cell–transferred mice, when compared with those from the control-treated mice, displayed significantly less reduction in body weight following the induction of TNBS colitis. (A) BALB/c mice received i.v. one of the following DC transfers (Transfer): 1 × 106 per mouse parental DCs, 1 × 106 per mouse DC-CYP cells, or 1 × 106 per mouse DC-CYP-ALDH cells. Seven days later, CD4+ T cells were purified from spleens of the mice and i.v. injected into syngeneic hosts that were induced for a mild TNBS colitis 3 d before the CD4+ T cell injection (CD4+/DC, CD4+/DC-CYP, and CD4+/DC-CYP-ALDH, respectively). Additionally, CD4− cells, which were purified from mice transferred with the DC-CYP-ALDH cells (CD4− cells), as well as no cell injection (No cells) were included as controls. Body weight was monitored daily. After having recovered from the first episode of colitis, the mice were induced for colitis at day 13 for a second time. (B) Percentage of baseline body weight of the mice over the observation period. The data presented are means ± SEM and are representative of two independent experiments (n = 4–6): DC-CYP versus DC, p < 0.01; DC-CYP-ALDH versus DC, p < 0.01; DC-CYP-ALDH versus DC-CYP, p > 0.05.

To further determine the role of CD4+ Treg cells in colitis suppression following the DC-CYP-ALDH cell transfer, we investigated the ability of the DC-CYP-ALDH cells to suppress ongoing TNBS colitis in the presence or absence of the PC61 anti-CD25 mAb because previous data published by us and others had shown that this Ab specifically depleted Foxp3+ Treg cells in vivo (22, 26, 27). Our data demonstrated that depletion of Foxp3+ Treg cells significantly reduced the ability of the DC-CYP-ALDH cells to suppress colitis as measured by survival rate (Fig. 8A and upper panel of Fig. 8B) and body weight (Fig. 8A and lower panel of Fig. 8B). Together, our data support that maximal colitis suppression following the DC-CYP-ALDH cell transfer requires Treg cells (Fig. 9).

FIGURE 8.

FIGURE 8.

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Depletion of Foxp3+ Treg cells significantly reduced therapeutic effect of the DC-CYP-ALDH cells for ongoing experimental colitis. (A) BALB/c mice were separated into four groups. In one group, the mice were i.p. administered with the anti-CD25 mAb (PC61) every other day for four times beginning from 2 d before the induction of colitis. All the mice were induced for colitis on day 0. At day 3, the mice that were injected with the anti-CD25 mAb (PC61) and another group of the mice received 1 × 106 DC-CYP-ALDH cells (DC-CYP-ALDH or DC-CYP-ALDH/mAb, respectively). The remaining two groups of mice received either no transfer (No transfer) or parental DC. (B) Percentage of survival (upper panel) and percentage of baseline bodyweight (lower panel) over the observation period are shown. The data presented are means ± SEM and are representative of two independent experiments (n = 4–6). *p < 0.05, ANOVA.

FIGURE 9.

FIGURE 9.

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A proposed model of DC-CYP-ALDH cell–mediated suppression of colitis. See text for description.

Discussion

In this study, we demonstrated that the DC-CYP-ALDH cells augmented the induction of Foxp3+CCR9+ T cells both in vitro and in vivo. In vivo, the DC-CYP-ALDH cell–induced Foxp3+ T cells homed to intestines. Additionally, transfer of the DC-CYP-ALDH cells robustly suppressed ongoing experimental colitis. Furthermore, CD4+ T cells from DC-CYP-ALDH cell–transferred mice suppressed the induction of experimental colitis in syngeneic hosts and depletion of Foxp3+ Treg cells significantly reduced the DC-CYP-ALDH cell–mediated colitis suppression. These findings support the conclusion that maximal colitis suppression following the DC-CYP-ALDH cell transfer requires Treg cells. Importantly, the DC-CYP-ALDH cells did not cause hypercalcemia in vivo (Supplemental Fig. 2), a finding which further corroborates our previous findings (22, 34). Collectively, our data support that the DC-CYP-ALDH cells are a promising novel therapy for IBD.

In our strategy, DCs are engineered to overexpress both the 1α-hydroxylase and RALDH2 so that, when priming naive T cells, the engineered DCs can deliver two strong signals: one signal is 1,25(OH)2D, which has been shown to augment the induction of regulatory molecules (e.g., Foxp3 and IL-10) (Supplemental Fig. 1) (22, 42), and the other signal is RA, which has been shown to enhance the induction of gut-homing receptors (e.g., CCR9) (Supplemental Fig. 1) (16). In this regard, 1,25(OH)2D as an in vivo Treg-inducing agent has particular advantages. First, 1,25(OH)2D does not widely suppress immune responses. Instead, 1,25(OH)2D suppresses the functions of Th1 and Th17 cells, which are major pathogenic T cells associated with IBD (39) but enhances the function of Th2 cells. Second, 1,25(OH)2D stimulates the secretion of antimicrobials (17), suppresses cancer development (18, 43), and, hence, potentially reduces the risks of infections and cancers, which are serious side effects associated with current IBD therapies as a result of unintended immune suppression (3, 4). Third, 1,25(OH)2D has been shown to tolerize DCs as well (44, 45), suggesting that in vivo administered DC-CYP-ALDH cells can be potentially converted into TolDCs through 1,25(OH)2D intracrine and/or autocrine effects. This potential self tolerization is particularly important in preventing the DC-CYP-ALDH cells from worsening the disease because the DCs used in this study were all immunogenic. Indeed, our data support this potential because transfer of the DC-CYP-ALDH cells robustly suppressed ongoing experimental colitis (Figs. 6, 8). However, more investigations are needed to demonstrate the presence of this self tolerization.

With respect to the mechanisms underlying the necessity for high 1,25(OH)2D concentrations, which otherwise cause hypercalcemia if administered systemically, the 1,25(OH)2D concentrations required for the induction of Treg cells are much higher than the physiological levels (Supplemental Fig. 1). In addition, one study showed that inflamed tissues were insensitive to 1,25(OH)2D (31). In this regard, several mechanisms, such as an increasing expression of 1,25(OH)2D catabolizing enzyme (i.e., 25-hydroxyvitamin D 24-hydroxylase) (46–48) and in situ generation of new immune cell subsets that are less sensitive to the action of 1,25(OH)2D, may contribute to this insensitivity during inflammation (31).

This study mainly focused on investigating the role of Foxp3+ Treg cells in the DC-CYP-ALDH cell–mediated suppression of colitis (Fig. 8). However, the DC-CYP-ALDH cells also augmented the expression of IL-10 in T cells in peripheral lymphoid tissues (Fig. 2D, 2E) and colons (Fig. 5G, 5H). In this regard, data from IL-10–deficient mice have clearly demonstrated that IL-10 plays an important role in the maintenance of intestinal immune tolerance (49, 50). Because a major T cell subset that produces high concentrations of IL-10 is type 1 Treg cells (51), it is possible that type 1 Treg cells may also contribute to the DC-CYP-ALDH cell–mediated suppression of colitis.

We acknowledge that this novel strategy is currently not Ag-specific (52). In this regard, recent data suggest that immunization with TolDCs pulsed with enterobacterial extracts or more defined enterobacterial peptides can ameliorate experimental colitis (53–55). However, whether the enterobacteria-derived Ags are the primary targets of the immune system needs further investigation. It remains a possibility that the enterobacteria-derived Ags trigger the initial inflammation and such inflammation then causes subsequent immune responses against intestinal self-antigens. One potential mechanism of the secondary autoimmune responses can be sequence similarity between the enterobacteria and intestinal self-tissues (molecular mimicry) (56). Such secondary autoimmune responses can perpetuate the inflammation observed in IBD patients. In this regard, the DC-CYP-ALDH cells described in this study can be easily pulsed with a human enterobacterium or an intestinal self-antigen, which is associated with IBD pathogenesis, to further enhance the specific targeting. To support this possibility, our data have demonstrated that the DC-CYP cells, when pulsed with an autoantigen, can provide Ag-specific suppression of experimental autoimmune encephalomyelitis (i.e., an autoimmune chronic inflammation in the CNS) (22).

Aside from the aforementioned Ag specificity, it is not yet known whether gut-homing Treg cells induced by the DC-CYP-ALDH cells have memory property. Generation of memory gut-homing Treg cells is critical for the long-term cure because of the perpetuating gut inflammation in IBD patients. Indeed, previous studies support this potential by showing that 1,25(OH)2D stimulates the expression of Helios, which is critical for maintaining stable regulatory property in Treg cells (8, 33, 57). Interestingly, recent data suggest that RA not only stabilizes natural Treg cells (58) but also enhances in vitro generation of induced Treg cells (59). Future studies will determine the effects of RA on the 1,25(OH)2d-mediated induction of Foxp3 expression.

Based on our findings, we propose a model for the DC-CYP-ALDH cell–mediated suppression of colitis (Fig. 9). In this model, the DC-CYP-ALDH cells, when s.c., i.v., or i.p. administered, home to peripheral lymphoid tissues (e.g., spleen and draining lymph nodes) and de novo synthesize locally high concentrations of 1,25(OH)2D and RA. This de novo synthesis prevents a significant elevation of systemic levels of 1,25(OH)2D and RA during the in vivo lifetime of the administered DC-CYP-ALDH cells, thereby circumventing severe systemic toxicities that can be potentially associated with these two molecules [i.e., hypercalcemia is associated with high systemic 1,25(OH)2D concentrations]. Subsequently, when the DC-CYP-ALDH cells prime naive T cells, the high concentrations of 1,25(OH)2D and RA program the naive T cells into gut-homing Treg cells. Consequently, the newly generated gut-homing Treg cells home to intestines and specifically augment gut immune tolerance, which provides a long-lasting control of chronic intestinal inflammation such as IBD.

Supplementary Material

1

Acknowledgments

We thank Dr. David Lo, a distinguished professor at the University of California at Riverside, for critical reading and Penelope Garcia for technical assistance and preparation of this manuscript.

This work was supported by the U.S. Department of Defense’s Office of the Assistant Secretary of Defense for Health Affairs through the Peer Reviewed Medical Research Program under Award W81XWH-15-1-0240 (to X.T.). This work was also partially supported by Loma Linda University Department of Medicine Research Innovation Grants 681207-2967 (to X.T. and G.G.), 681205-2967 (to X.T.), and 325491 (to D.J.B.). The opinions, interpretations, conclusions, and recommendations in this study are those of the authors and are not necessarily endorsed by the U.S. Department of Defense.

Abbreviations used in this article:

ALDH1a2

aldehyde dehydrogenase 1 family member A2

CYP27B1

cytochrome P450 family 27 subfamily B member 1

DC

dendritic cell

DC2.4

a bone marrow–derived DC line

DC-CYP cell

DC transduced with the lenti-CYP-GFP

DC-CYP-ALDH cell

DC transduced with the lenti-CYP-ALDH

DEAB

diethylaminobenzaldehyde

1α-hydroxylase

25(OH)D 1α-hydroxylase

IBD

inflammatory bowel disease

LLU

Loma Linda University

1,25(OH)2D

1,25-dihydroxyvitamin D

25(OH)D

25-hydroxyvitamin D

RA

retinoic acid

RALDH2

retinaldehyde dehydrogenase 2

tet

tetramer

TNBS

2,4,6-trinitrobenzenesulfonic acid

TolDC

tolerogenic dendritic cell

Treg

regulatory T

Footnotes

Disclosures

X.T., D.J.B., and K.-H.W.L. are inventors of a pending patent related to this study. Materials are readily available and will be provided under the material transfer policies of Loma Linda University.

The online version of this article contains supplemental material.

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