The Loss of H3K27 Histone Demethylase Utx in T Cells Aggravates Allergic Contact Dermatitis

It has been well established in research that allergic contact dermatitis (ACD) is induced by exposure to environmental allergens. The prevalence of ACD in general individuals is ∼20.1% in adults and 16.5% in nonadults, with a female predominance (1). Because contact allergens exist in very common products used in daily life, as well as in industrial and occupational settings (2, 3), it is difficult to remove potential allergens from daily life and prevent the onset of ACD in association with allergic inflammation.

Allergic inflammation after exposure to a certain contact allergen is caused by delayed-type hypersensitivity reactions through Ag-specific T cells (3). Ag-specific effector T cells contribute to the development of the pathogenesis of ACD by producing proinflammatory cytokines and chemokines. In addition, Ag-specific regulatory T (T_reg) cells are involved in the regulation of inflammation with their anti-inflammatory functions (3, 4).

Epigenetic modifications such as histone modifications have effects on the functions of immune cells (e.g., T cells and macrophages) and other participants (e.g., epidermal keratinocytes and dermal fibroblasts) that are associated with the development of allergic inflammation (5, 6). In addition, the responses of Ag-specific T cells, including the functional differentiation and the phenotypic plasticity of each subset of T cells, are tightly controlled by the gene expression through epigenetic modifications (7–9). Among the modifications of histones, the methylation mark of H3K27 represents a repressive transcription, whereas the methylation mark of H3K4 is a hallmark of an active transcription (10, 11). Thus, the modifications of the methylation of H3K4 and H3K27 are involved in the determination of the cell fate and function of CD4⁺ T cells (12).

Ezh2, Utx, and Jmjd3 regulate the methylation status of histone H3K27 (13, 14). Ezh2 methylates the residue of histone H3K27, whereas Utx and Jmjd3 demethylate the di- or trimethylated H3K27 (15). Utx has a role in promoting the differentiation of CD4⁺ T cells into Th2 cells (M. Kuwahara, M. Izumoto, H. Honda, K. Inoue, Y. Imai, J. Suzuki, S. Maruyama, M. Yasukawa, and M. Yamashita, manuscript posted on bioRxiv, DOI: 10.1101/184648). However, Jmjd3 has roles in promoting the differentiation of CD4⁺ T cells into Th1 cells and in suppressing differentiation into Th17 cells (16). Two studies reporting the function of Jmjd3 in the differentiation into Th2 cells have reported conflicting results (M. Kuwahara, et al., manuscript posted on bioRxiv, DOI: 10.1101/184648) (16). Furthermore, Jmjd3 is involved in the peripheral differentiation of CD4⁺ T cells into T_reg cells. Although functional roles of Utx and Jmjd3 in the differentiation of T cells into effector cells remain partially unknown, both the studies consistently showed that Utx and Jmjd3 could be involved in the differentiation of T cells into effector cells. Most important, the role of Utx and Jmjd3 in the pathogenesis of T cell–mediated immune disorders remains to be elucidated.

In this study, we examined the effects of either Utx deficiency or Jmjd3 deficiency in T cells on the pathogenesis of ACD. As a result, unlike Jmjd3 deficiency in T cells, the Utx deficiency aggravated the symptoms of ACD. The cellular and histological analyses of the skin lesions and the draining lymph nodes (LNs) showed that Utx deficiency in T cells increased the production of proinflammatory cytokines in the skin lesions or the draining LNs but decreased the ratio of T_reg cells to conventional CD4⁺ T (T_conv) cells in skin lesions and the draining LNs. In addition, Utx deficiency in CD4⁺ T cells modified their differentiation into effector cells and T_reg cells. We thus concluded that in T cells, Utx but not Jmjd3 negatively regulates the pathogenesis of ACD.

T cell–specific, Utx-deficient mice and Jmjd3-deficient mice have been established by crossing Cre recombinase transgenic mice under the control of the Cd4 promoter (Cd4-Cre Tg mice) with either Utx^flox/flox mice or Jmjd3^flox/flox mice. The Cd4-Cre Tg mice were purchased from the BioResource Research Center at the RIKEN Institute, Japan. Utx^flox/flox mice (17) and Jmjd3^flox/flox mice (18) were established by Drs. Kazuki Inoue and Yuki Imai (Ehime University) and Dr. Hiroaki Honda (Tokyo Women’s Medical University), respectively. C57BL/6 mice that were purchased from Clea Japan (Tokyo, Japan) were used as control mice after the mice became acclimatized to the environment of the animal facility at least for 1 wk. All mice at 8–13 wk of age with a C57BL/6 background were maintained under specific pathogen-free conditions in the animal facility at Ehime University Graduate School of Medicine. All experiments were performed on either female or male mice, except for Utx^flox/flox mice crossed with Cd4-Cre Tg mice, for which experiments were performed on female mice. All the animal procedures were approved by the Ehime University Animal Care and Use Committee and the Gene Recombination Experiment Safety Committee of Ehime University Graduate School of Medicine.

The method of inducing ACD in our present study was established on the basis of previously described methods (19). Mice were anesthetized, and then 100 μl of 0.15% 2,4-dinitro-1-fluorobenzene (DNFB; Sigma-Aldrich, Tokyo, Japan) dissolved in acetone/olive oil (3:1) was applied on the surface of their shaved abdomen. Five days after the first application of 0.15% DNFB solution, either 25 μl of 0.15% DNFB solution was applied to each side of the auricles (the auricle skin model) or 50 μl of 0.15% DNFB solution was applied onto the dorsal skin of the lumbar area that was shaved with hair clippers (the back skin model) three times with 5-d intervals. In some experiments, acetone/olive oil was applied instead of 0.15% DNFB solution as a control. The ear thickness was measured at the indicated durations using a thickness gauge (Dial thickness gauge; Mitsutoyo, Tokyo, Japan). The increase in ear thickness versus baseline was calculated by subtraction.

Auricle skin samples were taken 24 h after the final application of DNFB, fixed in 4% paraformaldehyde (Thermo Fisher Scientific, Tokyo, Japan), embedded in paraffin, sectioned at 5 μm, and stained with H&E. Images were photographed using a BZ-9000 digital microscope (Keyence Co., Osaka, Japan).

For the preparation of cells from the draining LNs, one pair of axillary, brachial, and inguinal LNs were pooled. For the preparation of cells from the auricle skin, pieces of the auricle skin were pooled, minced, and digested with 0.13 U/ml Liberase Thermolysin Medium (Sigma-Aldrich) and 50 U/ml DNase I (Nippon Gene, Tokyo, Japan) for 45 min at 37°C. Six pieces of auricle from wild-type (WT) mice and four pieces of auricle from Utx KO mice were used for the analysis of the granulocytes or other innate cells. Six pieces of auricle from WT or Utx KO mice were used for flow cytometry of T cells. Total CD4⁺ T cells from LNs for ex vivo analysis were positively selected using anti–CD4-FITC with anti-FITC Microbeads Ultrapure (Miltenyi Biotech, Tokyo, Japan) on an autoMACS separator at the purity of >95%. Total CD4⁺ T cells from spleen for in vitro culture were negatively selected using a MojoSort Mouse CD4 T Cell Isolation Kit (BioLegend, Tokyo, Japan) at >95% purity. CD4⁺CD44^loCD62L^hi naive CD4⁺ T cells from spleen for in vitro culture were negatively selected using a MojoSort Mouse CD4 T Cell Isolation Kit in combination with biotinylated antimouse/antihuman CD44 mAb (clone IM7; BioLegend) and biotinylated antimouse CD25 mAb (clone PC61; BioLegend) on an autoMACS separator at the purity of >95%.

Whole or enriched single cells were stained with Abs. The following mAbs were used: anti-CD3 (clone 145-2C11), anti-CD4 (clone RM4-5), anti-CD8 (clone 53-6.7), anti-CD11b (clone M1/70), anti-CD11c (clone N418), anti-CD19 (clone eBio 1D3), anti-CD44 (clone IM7), anti-CD45 (clone 30-F11), anti-CD45R (clone RA3-6B2), anti-CD49b (clone DX5), anti-CD62L (clone MEL-14), anti-CD64 (clone X54-5/7.1), anti-CD69 (clone [¹H].2F3), anti-CD103 (clone 2E7), anti-Foxp3 (clone 150D), anti-IA/IE (clone M5/114.15.2), anti–IFN-γ (clone HMG1.2), anti–IL-2 (clone JES6-5H4), anti–IL-4 (clone 11B11), anti–IL-17A (clone TC11-18H10.1), anti–IL-17F (clone 9D3.1C8), anti-Ly6G (clone 1A8), anti–Siglec-F (clone E50-2440), anti–TCR-β (clone H57-597), and anti-TER119 (clone TER-119) (purchased from BioLegend, BD Biosciences, eBioscience, Thermo Fisher Scientific, or TONBO Biosciences, all located in Tokyo, Japan). The following fluorescent conjugations of Abs were used: FITC, PE, PE—cyanine 7, allophycocyanin, Alexa Fluor 647, allophycocyanin–cyanine 7, Brilliant Violet 421, Violet 450, or Brilliant Violet 510. For the staining of lineage markers, biotinylated Abs were used in combination with Brilliant Violet 605 streptavidin. For the staining of cells in the auricle skin, the cells were treated with anti-CD16/32 (clone 93; BioLegend) before the staining with fluorescent conjugated Abs. Stained cells were analyzed on a Gallios cytometer with the Kaluza software program (Beckman Coulter, Tokyo, Japan), the CytoFLEX cytometer with the CytExpert 1.0 software program (Beckman Coulter), or a FACSAria II cell sorter with the FACSDiva software program (Beckton Dickinson, Tokyo, Japan). Cell sorting was performed using a BD FACSAria II cell sorter with the FACSDiva software program at the purity of >98%. All data were analyzed using the FlowJo software program, version 10.4.2 (BD Biosciences, Ashland, OR). The gating strategy was designed on the basis of a previous study (20).

CD4⁺ T cells were cultured in RPMI 1640 medium supplemented with 2 mM l-glutamine, 1 mM sodium pyruvate, 10 mmol/l nonessential amino acids, 10 mM HEPES, 100 U/ml penicillin, 100 μg/ml streptomycin, 55 μM 2-ME, and 10% FBS in a humidified atmosphere of 5% CO₂ at 37°C. The cells were stimulated with immobilized anti–TCR-β mAb (10 μg/ml, clone H57-597; BioLegend) and soluble anti-CD28 mAb (1 μg/ml, clone 37.5; BioLegend) for the first 2 d and expanded for 5 d (ThN, Th1, and Th2 conditions), 3 d (Th17 and inducible Treg [iTreg] conditions), or the indicated periods. The following culture conditions were used: ThN condition, 10 ng/ml recombinant murine (rm)IL-2 (TONBO Biosciences); Th1 culture condition, 10 ng/ml rmIL-2, 0.1 ng/ml rmIL-12 (PeproTech, Rocky Hill, NJ), and 2.5 μg/ml anti-IL-4 mAb (clone 11B11; BioLegend); Th2 culture condition, 10 ng/ml rmIL-2, 1 ng/ml rmIL-4 (PeproTech), and 2.5 μg/ml anti–IFN-γ mAb (clone R4-6A2; BioLegend); Th17 culture condition, 10 ng/ml rmIL-1β (FUJIFILM Wako, Osaka, Japan), 5 ng/ml rmIL-6 (Fujifilm Wako), 1 ng/ml recombinant human TGF-β (R&D Systems, Minneapolis, MN), 2.5 μg/ml anti–IFN-γ mAb, 2.5 μg/ml anti-IL-4 mAb, and 2.5 μg/ml anti–IL-2 mAb (clone JES6-1A12; BioLegend); iTreg condition, 10 ng/ml rmIL-2, 1 ng/ml recombinant human TGF-β, 2.5 μg/ml anti–IFN-γ mAb, and 2.5 μg/ml anti–IL-4 mAb.

For intracellular cytokine staining, the cultured cells were restimulated with immobilized 10 μg/ml anti–TCR-β mAb for 6 h in the presence of 2 μM monensin. For intracellular cytokine staining, cells were fixed in 4% paraformaldehyde, permeabilized, blocked, and stained with the indicated mAbs. Cytokine production was analyzed on a flow cytometer. For ELISA, the cells were restimulated with immobilized 10 μg/ml anti–TCR-β mAb for 24 h. The cytokines contained in the restimulated culture supernatant were measured by a sandwich ELISA using an optimal pair of Abs as follows: IFN-γ, clone R4-6A2 and clone XMG1.2; IL-2, clone JES6-1A12 and clone JES6-5H4; IL-4, clone BVD4-1D11 and clone BVD6-24G2; and IL-5, clone TRFK5 and clone TRFK4 (BD Biosciences). ELISAs for IL-10 and GM-CSF were performed with Ready-SET-Go! ELISA (Thermo Fisher Scientific) according to the manufacturer’s protocols. The ELISAs for IL-17, IL-17F, and IL-13 were performed with the DuoSet ELISA (R&D Systems) according to the manufacturer’s protocols.

For ex vivo analysis, cells were stimulated with or without 10 μg/ml anti–TCR-β for 4 h, then lysed in TRI reagent (Molecular Research Center, Cincinnati, OH). For in vitro analysis, cultured cells were harvested and lysed in TRI reagent. Total RNA was extracted. cDNA was synthesized using SuperScript VILO reverse transcriptase (Invitrogen/Thermo Fisher Scientific). cDNA of stimulated cells was used as a template for quantitative PCRs for the detection of cytokine genes, whereas cDNA of unstimulated cells was used for the detection of other genes. The expression of genes was detected with specific primers and Roche Universal Probes (Nippon Gene). The primers and probes used were as follows: 18s rRNA: 5′-GCAATTATTCCCCATGAACG-3′ (forward), 5′-GGGACTTAATCAACGCAAGC-3′ (reverse), probe 48; Ccr4: 5′-CTCAGGATCACTTTCAGAAGAGC-3′ (forward), 5′-GGCATTCATCTTTGGAATCG-3′ (reverse), probe 18; Ccr8: 5′-AGAAGAAAGGCTCGCTCAGA-3′ (forward), 5′-GGCTCCATCGTGTAATCCAT-3′ (reverse), probe 4; Ccr10: 5′-TGGCAATGGCCTAGTCTTG-3′ (forward), 5′-AGGTGGGAGATCGGGTAGTT-3′ (reverse), probe 71; Csf2: 5′-GCATGTAGAGGCCATCAAAGA-3′ (forward), 5′-CGGGTCTGCACACATGTTA-3′ (reverse), probe 79; Foxp3: 5′-TCAGGAGCCCACCAGTACA-3′ (forward), 5′-TCTGAAGGCAGAGTCAGGAGA-3′ (reverse), probe 78; Gata3: 5′-TTATCAAGCCCAAGCGAAG-3′ (forward), 5′-TGGTGGTGGTCTGACAGTT-3′ (reverse), probe 108; Ifng: 5′-ATCTGGAGGAACTGGCAAAA-3′ (forward), 5′-TTCAAGACTTCAAAGAGTCTGAGGTA-3′ (reverse), probe 21; Il4: 5′-GATCGGCATTTTCAACGAG-3′ (forward), 5′-CGAGCTCACTCTCTGTGGTG-3′ (reverse), probe 92; Il10: 5′-CAGAGCCACATGCTCCTAGA-3′ (forward), 5′-TGTCCAGCTGGTCCTTTGTT-3′ (reverse), probe 41; Il13: 5′-ACCCAGAGGATATTGC-3′ (forward), 5′-TGGGCTACTTCGATTT-3′ (reverse), probe 75; Il17f: 5′-CCCAGGAAGACATACT-3′ (forward), 5′-CAACAGTAGCAAAGAC-3′(reverse), probe 46; Il22: 5′-TTTCCTGACCAAACTCAGCA-3′ (forward), 5′-TCTGGATGTTCTGGTCGTCA-3′ (reverse), probe 17; Itgae: 5′-GCAGAGAACCACAGGACGA-3′ (forward), 5′-CAATGATGAGAGGCAAAGAGC-3′ (reverse), probe 63; Kdm6a: 5′-CGGGTTCGTGAGGTTTCAT-3′ (forward), 5′-GAGATTCGTAGCAGCGAACA-3′ (reverse), probe 3; Kdm6b: 5′-GTACAGACCCCCGGAACC-3′ (forward), 5′-TGGTGGAGAAAAGGCCTAAG-3′ (reverse), probe 20; Rorc: 5′-CAGCGCACCAACCTCTTT-3′ (forward), 5′-CCCACATCTCCCACATTGA-3′ (reverse), probe 52; Tbx21: 5′-TCAACCAGCACCAGACAGAG-3′ (forward), 5′-AAACATCCTGTAATGGCTTGTG-3′ (reverse), probe 19; Tnfa: 5′-TCTTCTCATTCCTGCTTGTGG-3′ (forward), 5′-GGTCTGGGCCATAGAACTGA-3′ (reverse), probe 49; Kdm6a (exon 2, for deleted region in Utx KO): 5′-CGGGTTCGTGAGGTTTCAT-3′ (forward), 5′-GAGATTCGTAGCAGCGAACA-3′ (reverse), probe 3; Kdm6b: 5′-GTACAGACCCCCGGAACC-3′ (forward), 5′-TGGTGGAGAAAAGGCCTAAG-3′ (reverse), probe 20; Kdm6b (exon 17, for deleted region in Jmjd3 KO): 5′-GGGTAACATCCACAGGCAAC-3′ (forward), 5′-TGTATAGCTGCACGGTGTTCAT-3′ (reverse), probe 102. For the detection of Il5 (Mm00439646_m1) and Il17a (Mmm004396180_m1) genes, predeveloped assay reagent kits of primers and probes (TaqMan; Thermo Fisher Scientific) were used.

Cells were washed with ice-cold PBS, and nuclear and cytoplasmic lysates were prepared using NE-PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific). The lysates were denatured, separated with SDS-PAGE on a 10% acrylamide gel, and transferred onto a polyvinylidene difluoride membrane. The blots were probed with rabbit anti-UTX mAb (clone D3Q1l), rabbit anti–β-actin mAb (clone 13E5), rabbit anti-H3K27me2 polyclonal Ab (PoAb) (39245), rabbit anti-H3K27me3 PoAb (39155), and mouse anti-histone H3 mAb (39763) (from either Cell Signaling Technology, Tokyo, Japan, or Active Motif, Carlsbad, CA). The immunoreactive proteins were visualized with peroxidase-conjugated antirabbit IgG (H+L) or antigoat IgG (H+L) (Jackson ImmunoResearch Laboratory, West Grove, PA) using an ECL detection system (GE Healthcare, Tokyo, Japan). The images were captured using a ChemiDoc XRS Plus luminescent image analyzer (Bio-Rad Laboratories, Hercules, CA).

The ChIP assay was performed using the Magna ChIP G kit (17-611; Merck Millipore, Burlington, MA) according to the manufacturer’s protocol. Briefly, 1 million cells per one ChIP reaction of freshly isolated CD4⁺ T cells or CD4⁺ T cells that were cultured for 5 d under one ThN condition were cross-linked with 1% paraformaldehyde in PBS and sonicated. The sheared chromatin lysates were precipitated with 2 μg of rabbit PoAb anti–histone H3K27me2 (39245; Active Motif) or 5 μg of rabbit PoAb anti–histone H3K27me3 (39155; Active Motif) Ab. The precipitated genome was amplified with gene-specific primers and Roche Universal Probes (Nippon Gene). The primers and probes used were as follows: Ifng promoter region (Ifng-p2): 5′-GAGGAGCCTTCGATCAGGTA-3′ (forward), 5′-TTCTCTAGGTCAGCCGATGG-3′ (reverse), probe 106; Csf2 promoter region: 5′-GCAGTGAGCCCAGTACTCAGA-3′ (forward), 5′-CACAATGCCCAGGAAAAGTAA-3′ (reverse), probe 5; Foxp3 exon 1 region: 5′- CATCTCCTCCATCCAAGCTC-3′ (forward), 5′-CTTCCAAGTCTCGTCTGAAGG-3′ (reverse), probe 2.

All statistical analyses were performed using the Prism software program (version 6 or version 9; GraphPad Software, La Jolla, CA). Differences between the two groups were tested using two-tailed unpaired t tests. Differences among multiple groups were tested using either one-way or two-way ANOVA tests. The statistical significance is shown in the figures (*p < 0.05, **p < 0.01, ***p < 0.005, and ****p < 0.001).

Ag-specific CD4⁺ and CD8⁺ T cells (21) are involved in the development of ACD. Both Utx and Jmjd3 play an important role in the differentiation and function of CD4⁺ and CD8⁺ T cells (M. Kuwahara, et al., manuscript posted on bioRxiv, DOI: 10.1101/184648) (16, 22). Thus, we first assessed the impact of the loss of Utx or Jmjd3 in T cells upon the induction of ACD using T cell–specific Utx KO mice and Jmjd3 KO mice. Of note, the mRNA expression of Kdm6a coding Utx and Kdm6b coding Jmjd3 was definitely deleted in CD4⁺ T cells of Utx KO mice and Jmjd3 KO mice, respectively (Supplemental Fig. 1A). In addition, we confirmed the depletion of Utx protein in cultured Utx KO CD4⁺ T cells by immunoblotting (Supplemental Fig. 1B). However, the level of histone H3K27me2 and H3K27me3 was almost identical among WT, Utx KO, and Jmjd3 KO CD4⁺ T cells (Supplemental Fig. 1C). Repeated applications of a hapten DNFB resulted in scaly and swollen skin in Utx KO mice in comparison with WT mice (Fig. 1A). In contrast, visual observation and the measurement of the ear thickness revealed no difference in the skin of Jmjd3 KO mice and WT mice (Fig. 1B).

In order to evaluate the morphological differences in the skin, we analyzed histological sections of skin specimens, as shown in (Fig. 1C and 1D. In comparison with auricle skin that was treated with vehicle (a mixture of acetone and olive oil), the skin treated with DNFB showed epidermal hyperplasia accompanied by mitotic figures in the epidermal keratinocytes, hyperkeratosis, hypertrophy of keratinocytes in the spinous and granular layers, and an increase in the intracellular space in the epidermis. In addition, cellular infiltration, including granulocyte infiltration and an increase in the number of fibroblasts, was observed in the dermis of WT mice that were treated with DNFB. DNFB-treated Utx KO mice had epidermal hyperplasia and increased mitotic figures, a decrease in granular layer keratinocytes, cellular hypertrophy in spinous layer keratinocytes, and microabscesses and massive infiltration of cells into the epidermis, dermis, and s.c. tissue. On the one hand, the junction between the dermis and s.c. tissue was not clear in the skin of Utx KO mice that received DNFB (Fig. 1C). On the other hand, Jmjd3 KO mice that were treated with DNFB had epidermal hyperplasia along with hypertrophy of keratinocytes and larger intracellular spaces in the spinous layer of keratinocytes, whereas the intensity of symptoms of Jmjd3 KO mice was milder than the symptoms of Utx KO mice (Fig. 1D). No inflammation was observed in the auricle of either Jmjd3 KO mice or WT mice that were treated with vehicle (Fig. 1C and 1D). As described below, this morphological analysis led us to focus on the role of the Utx expression in T cells in the development of DNFB-derived ACD.

Upon repeated exposure to a hapten, inflammatory cells, including granulocytes, macrophages, and Ag-specific effector T cells, infiltrated the lesional skin (4). In order to compare the composition of immune cells that accumulated in ACD skin between WT mice and Utx KO mice, we established gating strategies for granulocytes (Supplemental Fig. 2A) and for monocytes, macrophages, and dendritic cells (DCs) (Supplemental Fig. 2B) and analyzed immune cells in the skin at 24 h after a third challenge with DNFB (Fig. 2A–2C). Consistent with the histological observations (Fig. 1C and 1D), the frequencies and cell numbers of granulocytes, neutrophils, and eosinophils, per auricle, were all increased in Utx KO mice: neutrophils expressing Ly6G (a neutrophil marker) (23) as a CD45⁺ lineage marker (Lin)⁻ TCR-β⁻ CD11b⁺ Ly6G⁺, WT versus Utx KO, 53.2 ± 4.0% versus 50.2 ± 9.8% [52.7 ± 3.6 × 10³ cells versus 152.7 ± 30.2 × 10³ cells], a 2.9-fold increase in the mean cell number; eosinophils expressing Siglec-F (an eosinophil marker) (24) as CD45⁺ Lin⁻ TCR-β⁻ Ly6G⁻ IA/IE⁻ CD11b⁺ Siglec-F⁺, WT versus Utx KO, 23.5 ± 15.5% versus 60.4 ± 13.7% [9.7 ± 4.4 × 10³ cells versus 96.5 ± 57.7 × 10³ cells], a 9.9-fold increase in the mean cell number (Fig. 2A and 2D).

In contrast, the cell numbers of monocytes in DNFB-treated Utx KO mice were slightly increased compared with those in the DNFB-treated WT mice: monocytes expressing CD64 (a monocyte/macrophage marker) as CD45⁺ Lin⁻ Ly6G⁻ IA/IE⁻ SSC-A^lo CD11b⁺ CD64⁺, WT versus Utx KO, 46.9 ± 24.9% versus 52.3 ± 27.1% [8.3 ± 5.8 × 10³ cells versus 11.2 ± 13.3 × 10³ cells], a 1.3-fold increase in the mean cell number. Only the cell numbers of macrophages and DCs were increased in Utx KO mice: macrophages as CD45⁺ Lin⁻ Ly6G⁻ IA/IE^+/hi CD11b^+/hi CD64⁺, WT versus Utx KO, 50.3 ± 24.4% versus 27.2 ± 12.7% [14.3 ± 10.2 × 10³ cells versus 42.3 ± 42.0 × 10³ cells], a 3.0-fold increase in the mean cell number; DCs as CD45⁺ Lin⁻ Ly6G⁻ IA/IE^hi CD11b^−/+ CD11c⁺ CD64^-, WT versus Utx KO, 16.1 ± 3.1% versus 4.2 ± 1.8% [1.9 ± 0.7 × 10³ cells versus 3.4 ± 0.4 × 10³ cells], 1.8-fold increase as the mean cell number (Fig. 2B and 2E). Of note, the changes in the numbers of cells in Utx KO mice showed identical trends between experiments. Only the changes in neutrophils and DCs showed large statistically significant variations in the analysis (Fig. 2D–2F). The cell numbers of total αβT cells as CD45⁺ TCR-β⁺ SSC-A^lo, CD4⁺ T cells as CD45⁺ TCRβ⁺ SSC-A^lo CD4⁺, and CD8⁺ T cells as CD45⁺ TCR-β⁺ SSC-A^lo CD8⁺ were almost identical between Utx KO mice and WT mice, although the frequency varied: CD4⁺ T cells, WT versus Utx KO, 30.5 ± 7.6% versus 21.7 ± 4.2% [5.5 ± 2.5 × 10³ cells versus 5.3 ± 2.0 × 10³ cells]; CD8⁺ T cells, WT versus Utx KO, 51.4 ± 12.8% versus 47.6 ± 6.9% [11.3 ± 9.1 × 10³ cells versus 12.1 ± 5.3 × 10³ cells] (Fig. 2C and 2F).

In response to repeated Ag exposures, effector memory T (T_EM) cells and tissue-resident memory T (T_RM) cells accumulate in local sites of nonlymphoid tissues, such as the skin exposed to Ag rechallenge (25). In order to classify the type of memory T cells in the skin, we analyzed the expression profiles of surface markers. The frequency of CD4⁺ T_EM cells (CD45⁺ Lin⁻ TCR-β⁺ CD4⁺ CD62L^lo CD44^hi) in the skin of DNFB-treated WT and Utx KO mice was identical: WT versus Utx KO, 64.7 ± 4.6% versus 57.4 ± 1.4% [4.8 ± 3.0 × 10³ cells versus 5.0 ± 2.4 × 10³ cells] (Supplemental Fig. 3A and data not shown). The frequency and cell number of CD4⁺ T_RM cells (CD45⁺ Lin⁻ TCR-β⁺ CD4⁺ CD69⁺ CD103^+/−) in the skin of Utx KO mice was slightly higher than that in the skin of WT mice, although the trend was not statistically significant: WT versus Utx KO, 19.3 ± 17.9% versus 31.4 ± 18.0% [1.1 ± 1.1 × 10³ cells versus 1.9 ± 1.8 × 10³ cells] (Supplemental Fig. 3B and data not shown). Similarly to CD4⁺ T cells, the numbers of CD8⁺ T_EM cells (CD45⁺ TCR-β⁺ CD8⁺ CD62L^lo CD44^hi) in the DNFB-treated skin of WT mice and Utx KO mice were statistically identical: WT versus Utx KO, 71.8 ± 13.0% versus 70.9 ± 14.8% [10.1 ± 7.6 × 10³ cells versus 5.3 ± 2.3 × 10³ cells] (Supplemental Fig. 3C and data not shown). In addition, the numbers of CD8⁺ T_RM cells (CD45⁺ TCR-β⁺ CD8⁺ CD69⁺ CD103⁺) in the skin of Utx KO mice were also statistically identical to those of WT mice (Supplemental Fig. 3D, WT versus Utx KO, 19.0 ± 4.5% versus 13.8 ± 0.5% [1.9 ± 1.4 × 10³ cells versus 1.7 ± 0.7 × 10³ cells]).

Although the numbers of CD4⁺ T cells and CD8⁺ T cells in the lesional skin were identical between WT and Utx KO mice, the numbers of CD25⁺ Foxp3⁻ T_conv cells were significantly increased in the skin of Utx KO mice, whereas the numbers of Foxp3⁺ Treg cells were decreased in Utx KO mice (Supplemental Fig. 3E and 3F). The result suggests that a larger number of CD4⁺ T cells that accumulated in the lesional skin of Utx KO mice could become activated. The mRNA expression of inflammatory cytokine genes was analyzed at the same time point as (Fig. 2A–2F to clarify the functionality of CD4⁺ T cells and CD8⁺ T cells in the skin (Fig. 2G and 2H). Total CD4⁺ T cells in the DNFB-treated skin of Utx KO mice expressed proinflammatory cytokine mRNAs, including Ifng, Il13, and Csf2, at levels almost identical to those of the cells of WT mice (Fig. 2G). Similarly, total CD8⁺ T cells in the skin of Utx KO mice expressed proinflammatory cytokines such as Ifng and Tnfa at levels identical to those of the cells of WT mice. However, the Il13 expression of Utx KO CD8⁺ T cells in the skin was significantly higher than that in WT cells (Fig. 2H). It is noteworthy that the relative quantities of Ifng and Il13 in CD4⁺ T cells and CD8⁺ T cells were calculated using the same standardized relative quantity (Fig. 2G and 2H). The Ifng expression of WT CD8⁺ T cells was ∼10 times higher than the expression of WT CD4⁺ T cells, whereas the Il13 expression in WT CD8⁺ T cells and WT CD4⁺ T cells was identical at the time point that we tested. Taken together, repeated applications of DNFB to Utx KO mice caused the accumulation of larger numbers of neutrophils and DCs as well as possibly larger numbers of eosinophils and macrophages in the skin lesion than in WT mice. Furthermore, the activation of Utx KO CD4⁺ T cells and CD8⁺ T cell–derived proinflammatory cytokines such as IL-13 might be involved in the aggravation of skin symptoms.

The sensitization of skin by having contact with haptens primes the naive T cells in the draining LNs into effector T cells through migratory DCs that present peptides from haptenated proteins (3, 4). To compare the kinetics of surface markers on T cells in the draining LNs along with Ag applications between WT mice and Utx KO mice, we analyzed naive, central memory (CM), and EM phenotypes on CD4⁺ or CD8⁺ T cells according to the expression of CD62L and CD44. As shown in (Fig. 3A and 3B, the frequencies and cell numbers of total CD4⁺ and CD8⁺ T cells were both lower in Utx KO mice at 24 h after the second challenge with DNFB. At this point, a significant difference in ear thickness was observed between WT and Utx KO mice (Fig. 1A). Furthermore, the numbers of CD62L^hi CD44^lo naive CD4⁺ T cells, CD62L^lo CD44^hi CD4⁺ T_EM cells, and CD62L^hi CD44^hi CD4⁺ T_CM cells were all decreased in Utx KO mice until 24 h after the second challenge, whereas the frequencies were dramatically decreased in CD4⁺ T_EM cells at 24 h after the third challenge (Fig. 3A). In contrast, significant differences in the frequencies and cell numbers in CD8⁺ T cells were only observed in the CM populations, with the exception of the slight difference in cell numbers of naive CD8⁺ T cells between WT and Utx KO mice (Fig. 3B). Because at least two challenges with DNFB caused swelling responses in the auricle skin that were more severe in Utx KO mice than in WT mice (Fig. 1A), the cell numbers of memory and/or naive phenotypes of CD4⁺ T cells and CD8⁺ T cells in the draining LNs might not be linked to the severity of the skin lesion.

To assess the capability of cytokine production in CD4⁺ T cells of skin-draining LNs after challenge with DNFB, we analyzed the proinflammatory phenotypes of CD4⁺ T cells and compared the cells between WT or Utx KO mice at 24 h after the first, second, and third challenges with DNFB. Among the cytokines that we tested, the Ifng and Csf2 mRNA expression levels at 24 h after the second challenge with DNFB were higher, whereas the Il17a, Il17f, and IL22 mRNA expression levels were lower, in Utx KO CD4⁺ T cells of the draining LNs than in WT CD4⁺ T cells (Fig. 4A). Consistent with the lower Il17a and IL17f mRNA expression in Utx KO CD4⁺ T cells, the master regulator for Th17 cell differentiation, Rorc (26), at 24 h after the second challenge with DNFB was lower in Utx KO CD4⁺ T cells than in WT CD4⁺ T cells. In CD4⁺ T cells, the expression levels of Tbx21 coding T-bet, the master regulator for Th1 cells (27), and Gata3, the master regulator for Th2 cells (28), were identical in WT and Utx KO mice, regardless of the number of challenges with DNFB (Fig. 4B).

It is noteworthy that the expression of Il13 was higher in Utx KO CD4⁺ T cells at 24 h after the first and second challenges with DNFB than that in WT CD4⁺ T cells. However, the difference was not statistically significant. The expression of Il4 and Il5 did not differ between Utx KO CD4⁺ T cells and WT CD4⁺ T cells, regardless of the number of challenges (Fig. 4A). The expression of Gata3 dropped after the third challenge in CD4⁺ T cells, regardless of the presence of Utx (Fig. 4B). These findings suggested that Utx KO CD4⁺ T cells showed greater capacity to express Ifng and Csf2 (and probably Il13) than did WT CD4⁺ T cells in the draining LNs after challenges with the hapten, but that Utx KO CD4⁺ T cells had a smaller capacity to express Il17a, Il17f, and Il22 than did WT CD4⁺ T cells in the draining LNs after challenges with the hapten.

To address the role of Utx in the functional differentiation of CD4⁺ T cells, we cultured splenic naive CD4⁺ T cells from either WT or Utx KO mice under various conditions. First, we analyzed the kinetic expression of Kdm6a and Kdm6b in naive CD4⁺ T cells after stimulation with anti–TCR-β plus anti-CD28 for up to 72 h (Fig. 5A). Interestingly, the kinetics of the expression of Kdm6a and Kdm6b in activated CD4⁺ T cells were different: The expression of Kdm6a was almost stable within the time frame that we analyzed, whereas the expression of Kdm6b was transiently increased at just after 2 h of stimulation and then gradually decreased (Fig. 5A). Second, the expression levels of Kdm6a in differentiating CD4⁺ T cells that were cultured for 2 d under neutral (ThN), Th1, Th2, Th17 or iTreg conditions were all lower than those in naive CD4⁺ T cells. Similarly, the expression levels of Kdm6b in differentiating CD4⁺ T cells were all lower, regardless of the culture conditions (Fig. 5B).

To assess the effects of Utx deficiency in effector CD4⁺ T cell differentiation in vitro, we cultured naive CD4⁺ T cells from WT or Utx KO mice under various cytokine conditions. Activated Utx KO CD4⁺ T cells cultured with one ThN condition gave rise to a greater frequency of IFN-γ–producing cells and greater IFN-γ production than did activated WT CD4⁺ T cells. The Utx KO cells produced slightly a larger amount of IL-2 and a significantly larger amount of GM-CSF than did WT cells (Fig. 5C and Supplemental Fig. 4A). The increase in the production of IFN-γ and GM-CSF in Utx KO CD4⁺ T cells under one ThN condition was also supported by the status of H3K27me2 and H3K27me3 (Supplemental Fig. 4B). The basal level of H3K27me3 at the Ifng promoter region (Ifng-p2) in Utx KO unstimulated CD4⁺ T cells was lower than in WT unstimulated CD4⁺ T cells. After cultivation for 5 d, the level of H3K27me3 at the Ifng-p2 in Utx KO cells showed no reduction, whereas the level in WT cells was decreased. The level of H3K27me2 at the Csf2 promoter region (Csf2-p) of unstimulated Utx KO CD4⁺ T cells was almost the same as in unstimulated WT cells. However, the level of H3K27me2 at the Csf2-p gene locus of differentiated Utx KO cells was lower than that in differentiated WT cells (Supplemental Fig. 4B).

Similarly, Utx KO CD4⁺ T cells under one Th1 condition were capable of inducing slightly higher IL-2 production than were WT cells. To the contrary, the amount of IFN-γ in differentiated CD4⁺ T cells under one Th1 condition was slightly smaller than that in WT cells (Fig. 5D and Supplemental Fig. 4A). It is noteworthy that the amount of IFN-γ produced by Utx KO CD4⁺ T cells under one ThN condition (Fig. 5C) was almost identical to the amount produced under one Th1 condition (Fig. 5D). In addition, on the one hand, Utx KO CD4⁺ T cells that were cultured under one Th1 condition were capable of producing a significantly larger amount of GM-CSF than were WT CD4⁺ T cells (Fig. 5D). On the other hand, the capability to produce proinflammatory cytokines (e.g., IL-4, IL-5, and IL-13) and the regulatory cytokine IL-10 was decreased in Utx KO CD4⁺ T cells cultured under one Th2 condition as compared with WT CD4⁺ T cells, whereas the production of GM-CSF in WT and Utx KO CD4⁺ T cells was identical (Fig. 5E and Supplemental Fig. 4A). The capability to produce IL-17A in Utx KO CD4⁺ T cells cultured under one Th17 condition was also inhibited in comparison with WT CD4⁺ T cells, whereas the IL-17F secretion was identical (Fig. 5F and Supplemental Fig. 4A). In addition, Utx KO CD4⁺ T cells cultured under one Th17 condition were able to secrete larger amounts of GM-CSF than the WT CD4⁺ T cells. However, the amount of GM-CSF produced by the Utx KO CD4⁺ T cells under one Th17 condition was relatively much smaller than the amount of GM-CSF produced by the Utx KO cells under one Th2 condition (Fig. 5E and 5F).

The expression of chemokine receptors is required for the migration of CD4⁺ T cells from the draining LNs to the skin (29). We therefore analyzed the expression of skin-tropic chemokine receptors, Ccr4, Ccr8, and Ccr10, on CD4⁺ T cells cultured under various conditions in vitro. Ccr4 was expressed at the highest level in differentiating Th2 and Th17 cells, whereas Ccr8 and Ccr10 were expressed in Th2 and ThN at the highest level in differentiating WT CD4⁺ T cells, respectively. The loss of Utx slightly enhanced the Ccr4, Ccr8, and Ccr10 expression in differentiating Th2 cells. In contrast, Utx KO Th1 cells showed lower Ccr4 and Ccr8 expression levels with the higher expression of Ccr10. Utx KO CD4⁺ T cells cultured under one ThN condition also expressed a lower level of Ccr10. However, the expression levels of Ccr4 and Ccr8 in Utx KO ThN cells were identical (Supplemental Fig. 4C). These analyses of Utx KO CD4⁺ T cells cultured under various conditions in vitro suggested that activated Utx KO CD4⁺ T cells might be able to produce a large amount of IFN-γ and GM-CSF and that Utx KO effector CD4⁺ T cells could be prone to traffic to the skin.

Foxp3⁺ CD4⁺ T_reg cells play suppressive roles in hapten-specific effector responses (30). In order to address the effect of Utx deficiency in T_reg cells upon aggravation of DNFB-induced Utx KO ACD in mice and the role of Utx in the generation of T_reg cells in vitro, we first analyzed the frequencies and cell numbers of T_reg cells in our models of ACD. As shown in (Fig. 6A and 6B, the frequency and cell numbers of Foxp3⁺ CD4⁺ T_reg cells in the skin of Utx KO mice at 24 h after the third challenge with DNFB were lower than those in WT mice. However, the changes of Foxp3⁺ CD4⁺ T_reg cells were not statistically significant: WT versus Utx KO, 7.1 ± 1.2% versus 2.1 ± 0.4% [0.5 ± 0.2 × 10³ cells versus 0.2 ± 0.1 × 10³ cells]. Furthermore, the frequency and cell numbers of Foxp3⁻ CD4⁺ T_conv cells in the skin of Utx KO mice were higher. However, the changes of Foxp3⁻ CD4⁺ T_conv cells were not statistically significant: WT versus Utx KO, 40.8 ± 5.1% versus 61.9 ± 3.7% [2.8 ± 0.4 × 10³ cells versus 5.2 ± 1.8 × 10³ cells]. The ratio of Foxp3⁺ CD4⁺ T_reg cells to Foxp3⁻ CD4⁺ T_conv cells in the skin was also lower in Utx KO mice than in WT mice: WT versus Utx KO, 0.18 ± 0.04 versus 0.04 ± 0.01. Consistently, the expression of Foxp3 in the CD4⁺ T cells in the skin of Utx KO mice was slightly lower than that in the cells in the skin of WT mice: WT versus Utx KO, 1.20 versus 0.97 (Fig. 6C). In addition to the phenotype in the skin, the frequencies and cell numbers of Foxp3⁺ CD4⁺ T_reg cells in the draining LNs of Utx KO mice and the ratio of T_reg cells to T_conv cells were all lower than those in WT mice, regardless of the number of DNFB challenges (Fig. 6D–6F). Furthermore, the expression of another T cell–mediated regulating factor, Il10, was also lower in CD4⁺ T cells of Utx KO mice than in those of WT mice at 24 h after the third challenge with DNFB, whereas the expression after the first and second challenges was identical (Fig. 6G). These data suggested that the regulatory functions of CD4⁺ T cells could become weaker without the expression of Utx.

In addition to the above analyses, we investigated the effect of the Utx deficiency in naive CD4⁺ T cells on the differentiation of T_reg cells in vitro. The dimethylation or trimethylation status of histone H3K27 at the Foxp3 exon 1 region in Utx KO freshly isolated CD4⁺ and activated CD4⁺ T cells was almost comparable to that in WT cells (Supplemental Fig. 4B). However, the induction of Foxp3 in activated CD4⁺ T cells cultured under iTreg conditions was decreased in Utx KO CD4⁺ T cells as compared with that in WT CD4⁺ T cells (Fig. 6H). It is noteworthy that Utx KO Foxp3⁺ T_reg cells under one culture condition in the presence of TGF-β showed lower expression of CCR4 at both the mRNA (Fig. 6I) and protein (Fig. 6J) levels than did WT Foxp3⁺ T_reg cells. The efficiency of CD4⁺ T cell differentiation into T_reg cells and of CCR4 induction that is required for infiltration of the skin (31) was lower in the absence of Utx. All of the results suggested that lower numbers of Foxp3⁺ T_reg cells and the weaker expression of CCR4 in Utx KO CD4⁺ T cells could be linked to the aggravation of skin symptoms induced by the application of DNFB.

Because CD4⁺ T cells are required to have an early inflammatory response for the induction of contact hypersensitivity (3), we focused on CD4⁺ T cells in the present study. We demonstrated that T cell–specific Utx KO mice showed severer symptoms of ACD than did WT mice, whereas Jmjd3 KO mice showed conditions of inflammation identical to those in WT mice. We therefore concluded that the following factors in Utx KO mice could potentially exacerbate the pathogenesis of ACD: (1) the massive accumulation of myeloid cells and the activation of Ag-specific T cells that express proinflammatory cytokines, (2) the decreased capability of CD4⁺ T cells to differentiate into Foxp3⁺ T_reg cells, and (3) the decreased expression of CCR4 (a skin-tropic chemokine receptor) in T_reg cells.

The analysis of the affected skin by flow cytometry showed a greater accumulation of myeloid cells, including neutrophils, DCs, eosinophils, and macrophages, in Utx KO mice than in WT mice. Furthermore, CD8⁺ T cells that were capable of producing larger amounts of proinflammatory cytokines (e.g., Il13) infiltrating the skin of Utx KO mice than were seen in the skin of WT mice after a third challenge with DNFB. Although the mRNA expression of cytokine genes in CD4⁺ T cells of the skin at one time point during the process of ACD development did not differ between Utx KO and WT mice to a statistically significant extent (Fig. 2G and 2H), the higher levels of cytokine expression, such as Ifng and Csf2, in Utx KO CD4⁺ T cells in the skin at a different time point, after a second challenge with DNFB, could be interpreted as follows. First, Utx KO CD4⁺ T cells expressed Ifng and Csf2 mRNAs at higher levels than did WT CD4⁺ T cells during the development of ACD (Fig. 4A). Second, Utx KO CD4⁺ T cells cultured under one ThN condition gave rise to the expression of IFN-γ and GM-CSF at higher levels than did WT CD4⁺ T cells (Fig. 5C). The production of IL-13 could attract T cells via the induction of CCL22 in keratinocytes (32) and could downregulate the expression of structural molecules in the epidermis (33, 34). In addition, the IFN-γ expression in the skin could result in the accumulation of T cells and inflammatory DCs (35), the production of chemoattracting and activating factors for monocytes in keratinocytes (36), and the induction of the expression of transglutaminase, all processes that are involved in the cornification of keratinocytes (37). Furthermore, the presence of GM-CSF could become a mediator of tissue inflammation through DCs, macrophages, and granulocytes (38). Thus, the increase in the production of all proinflammatory cytokines in Utx KO T cells could be associated with the accumulation of the other immune cells in the skin, as we observed histologically in Utx KO mice. Further analysis of the kinetic expression of cytokines and comprehensive analysis of cytokine production in Utx KO T cells in the skin could further elucidate the mechanisms of skin symptoms of ACD.

As reported in one previous study (39), the expression of cell surface molecules, including CD44, may be affected by the deletion of Utx in CD4⁺ T cells. Throughout our analysis, we have observed the downregulated expression of CD44 in Utx KO T cells in the skin-draining LNs and spleen and a slightly downregulated expression level in the skin (Supplemental Fig. 3 and data not shown). Although CD44 may play a role in the development of ACD and the expression of CD44 in Utx KO T cells in the skin was relatively low (Fig. 2), we hypothesize that the lower expression of CD44 on Utx KO CD4⁺ T cells and CD8⁺ T cells could not affect their capability to express cytokines in the skin or the lesional skin.

Because Li et al. demonstrated that the generation of Foxp3⁺ T_reg cells was reduced in Jmjd3-deficient CD4⁺ T cells cultured under one iTreg condition in vitro (16), we examined whether the regulatory functions could become weakened in Utx KO mice. Strikingly, the numbers of T_reg cells in the skin of Utx KO mice were decreased, as were those in the draining LNs of Utx KO mice. The expression of the regulatory cytokine Il10 was also decreased in both CD4⁺ T cells of Utx KO mice with ACD and Utx KO CD4⁺ T cells cultured under one Th2 condition in vitro. The decreased differentiation of Utx KO CD4⁺ T cells into Foxp3⁺ T_reg cells in in vitro culture under one iTreg condition with the lower expression of CCR4 could be linked to the lower numbers of T_reg cells in both the skin and the draining LNs. All of the aspects that we have observed in Utx KO mice with ACD and in Utx KO CD4⁺ T cells in vitro could be explained as the aggravation of the pathogenesis of ACD, partially due to the deterioration of the regulatory functions.

Based on our data on Jmjd3 KO CD4⁺ T cells in vitro (M. Omori-Miyake, unpublished observations), as well as the data described by Li et al. (16), the increase in IFN-γ production in in vitro cultures of Utx KO CD4⁺ T cells under one ThN condition was not observed in Jmjd3 KO CD4⁺ T cells. Furthermore, Utx and Jmjd3 deficiency had different influences on histone H3K27 methylation at the Ifng and Csf2 promoter regions (Supplemental Fig. 4B). Therefore, Utx and Jmjd3 might have a distinct function for the differentiation of CD4⁺ T cells. Because the frequency of Ag-specific T cells in the draining LNs in response to an Ag could be quite low, we believe that the ThN conditions of culture, with only IL-2, would mimic the events in our experimental model of ACD. We therefore hypothesize that Utx could be involved in the Ag-driven expression of IFN-γ and GM-CSF in CD4⁺ T cells for the following reasons: (1) Utx KO CD4⁺ T cells expressed Ifng and Csf2 mRNAs at higher levels than WT CD4⁺ T cells in response to the application of DNFB (Fig. 4); (2) the trimethylation status of the promoter region of the Ifng gene locus in Utx KO CD4⁺ T cells was originally lower than that in WT CD4⁺ T cells (Supplemental Fig. 4B); and (3) the TCR stimulation–driven downregulation of H3K27me2 status at the promoter region of the Csf2 gene locus in Utx KO CD4⁺ T cells was greater in than that in WT CD4⁺ T cells (Supplemental Fig. 4B).

To envision our further studies, we also determined the cytokine-driven differentiation of Utx KO CD4⁺ T cells under Th1, Th2, Th17, or iTreg conditions in vitro (Fig. 5). The ablation of Utx in CD4⁺ T cells resulted in the decreased expression of each characteristic cytokine or transcription factor: IFN-γ for one Th1 condition; IL-4, IL-13, and IL-5 for one Th2 condition; IL-17A for one Th17 condition; and Foxp3 for one iTreg condition. In addition, the trends of IL-4, IL-5, and IL-13 production under one Th2 culture condition, the IL-17 production under one Th17 culture condition, and the Foxp3 expression under one iTreg culture condition in Utx KO CD4⁺ T cells and Jmjd3 KO CD4⁺ T cells were all identical, as other groups have shown (Ref. 16 and M. Kuwahara, et al., manuscript posted on bioRxiv, DOI: 10.1101/184648). What these results suggest to us is that Utx and Jmjd3 might have distinct function in a given condition but redundant functions in another condition for differentiation of CD4⁺ T cells. Although the molecular mechanisms of these changes in Utx KO CD4⁺ T cells remain to be investigated, the results have become a touchstone for our future studies.

In our present study, we assessed the effects of the loss of Utx and Jmjd3 expression in T cells on the pathogenesis of ACD. The exacerbated symptoms of ACD in Utx KO mice could occur via the following potential mechanisms: the accumulation of T cells that secrete proinflammatory cytokines, the infiltration of proinflammatory myeloid cells that could be attracted and activated by the T cells, and the reduced number of T_reg cells in the skin. Further studies will be needed to elucidate each of these potential mechanisms and their interplay and thus to facilitate our further understanding of how and why the symptoms of ACD in Utx KO mice would be exacerbated.

The authors express their sincere thanks to Aya Tamai and Mei Imamura-Miyazaki (Department of Immunology, Ehime University Graduate School of Medicine) and the staff of the Division of Laboratory Animal Research at the Advanced Research Support Center (Ehime University) for the maintenance of the experimental mice. The authors also thank Dr. Kenji Kameda for his technical assistance with the multicolor cell-sorter operation and the staff for their technical assistance in the histological analysis (all at Division of Analytical Bio-Medicine, Advanced Research Support Center, Ehime University), Drs. Akira Matsumoto (Department of Infections and Host Defenses, Ehime University), and Kazuki Inoue and Yuuki Imai (Ehime University) for their technical support.

The authors have no financial conflicts of interest.