The transcription factor promyelocytic leukemia zinc finger (PLZF) is encoded by the BTB domain–containing 16 (Zbtb16) gene. Its repressor function regulates specific transcriptional programs. During the development of invariant NKT cells, PLZF is expressed and directs their effector program, but the detailed mechanisms underlying PLZF regulation of multistage NKT cell developmental program are not well understood. This study investigated the role of acetylation-induced PLZF activation on NKT cell development by analyzing mice expressing a mutant form of PLZF mimicking constitutive acetylation (PLZFON) mice. NKT populations in PLZFON mice were reduced in proportion and numbers of cells, and the cells present were blocked at the transition from developmental stage 1 to stage 2. NKT cell subset differentiation was also altered, with T-bet+ NKT1 and RORγt+ NKT17 subsets dramatically reduced and the emergence of a T-betRORγt NKT cell subset with features of cells in early developmental stages rather than mature NKT2 cells. Preliminary analysis of DNA methylation patterns suggested that activated PLZF acts on the DNA methylation signature to regulate NKT cells’ entry into the early stages of development while repressing maturation. In wild-type NKT cells, deacetylation of PLZF is possible, allowing subsequent NKT cell differentiation. Interestingly, development of other innate lymphoid and myeloid cells that are dependent on PLZF for their generation is not altered in PLZFON mice, highlighting lineage-specific regulation. Overall, we propose that specific epigenetic control of PLZF through acetylation levels is required to regulate normal NKT cell differentiation.

This article is featured in Top Reads, p.751

Invariant NKT (iNKT) cell subsets are a unique regulatory αβ T cell population selected by self-lipid Ags presented by the nonclassical MHC molecule CD1d (1). Stimulation of NKT cells by α-galactosylceramide (α-GalCer), a synthetic CD1d ligand recognized by all NKT cell subsets, triggers the release of multiple cytokines; the specific mixture depends on the NKT cell subset (2). NKT1 cells mainly produce Th1 cytokines, and these cells have been reported to be essential in a range of immune responses, such as tumor rejection (3). NKT17 cells produce Th17 cytokines and were shown to be involved in autoimmune diseases and antibacterial responses (4, 5). A third subset of NKT cells, NKT2 cells, has features of Th2 cells; these cells are involved in airway hyperactivity (6).

In contrast to mainstream αβ T cell progenitors, NKT cell progenitors express a semi-invariant TCR. In mice, this receptor is composed of a Vα14-Jα18 chain paired with one of three Vβ segments (Vβ8, -2, or -7) (7). Although conventional αβ TCRs are positively selected by peptides presented in the context of MHC molecules expressed on thymic epithelial cells, NKT Vα14i TCRs are positively selected by glycolipid-presenting nonclassical class I CD1d molecules expressed on CD4+CD8+ cortical thymocytes (1). Although they derive from a common CD4+CD8+ cortical thymocyte progenitor (8), surface phenotype, developmental program, and function start to diverge shortly after the positive selection of conventional and NKT cells (9). During their development, mature NKT cells pass through three developmental stages characterized by differential expression of CD44 and NK1.1. In C57BL/6 mice, the stages are defined as follows: stage 1 (CD44lowNK1.1+), stage 2 (CD44highNK1.1), and stage 3 (CD44highNK1.1+). Stage 3 NKT cells produce mainly Th1 type cytokines, whereas stage 1 NKT cells produce Th2 type cytokines (9). Before stage 1, NKT cells are said to be stage 0 and correspond to early heat stable Ag (HSA)high nonproliferating precursors.

Recently, a classification was published categorizing NKT cell development into the subtypes NKT1, NKT2, and NKT17 based on their cytokine profiles and expression of specific transcription factors (TFs) (10).

The molecular mechanisms by which NKT cell progenitors commit to the NKT lineage and differentiate have been extensively studied. CD1d/TCR and homotypic interactions through SLAM family proteins expressed by double-positive (DP) cells play important roles in guiding developing T cells into the NKT cell pool (11). The TF promyelocytic leukemia zinc finger (PLZF) is a significant developmental regulator of the NKT lineage. Indeed, PLZF is specifically expressed in these cells, but not in conventional T cells (12, 13). Expression of PLZF in developing NKT cells was shown to be epigenetically regulated and linked to TCR-induced Egr2 expression, which in turn triggers PLZF gene expression (14). PLZF directs the acquisition of several key components of the NKT cell effector program during development, including functional and migratory properties (12, 13). Mutation or deletion of the PLZF gene Zbtb16 causes NKT cells to regress from effector memory to a naive phenotype and their redistribution to the lymph nodes. On the contrary, ectopic CD4 promoter–driven expression of PLZF converts conventional naive CD4+ thymocytes into CD44highCD62Llow effector cells, producing both type 1 and type 2 cytokines (12, 13). These observations suggest that PLZF is not only required but also sufficient for acquisition of the innate T cell effector program. PLZF is also involved in the development of other unconventional T cells, namely MAIT cells, γδ T cells expressing the Vγ1.1-Vδ6.3 TCR, and Vγ6 IL-17+ T cells (12, 15, 16). In addition, this TF was recently found to influence the development and function of innate lymphoid cells (ILCs) (17).

The PLZF protein is subdivided into two main domains: a BTB/POZ evolutionarily conserved N-terminal domain that promotes recruitment and dimerization with other poxviruses and zinc-finger (POZ) and Krüppel members (18) and a Kruppel-like zinc finger C-terminal domain that mainly mediates sequence-specific binding to DNA through a consensus genomic motif (19). In initial studies, PLZF was shown to be an important regulator of cell growth, self-renewal, and differentiation, influencing these events through its sequence-specific negative transcriptional activity (20, 21). Following recruitment of histone deacetylases (HDACs), DNA methyltransferases, and nuclear corepressors to sequence-specific binding sites, PLZF exerts its epigenic function and maintains propagation of a repressive chromatin environment and further chromatin remodeling activity to cause gene silencing (21, 22). Through its ninth zinc finger, PLZF’s DNA binding activity may be modulated by coactivators such as histone acetyltransferase (HAT) protein p300, which targets a specific acetylation site (aa 632–652) (22). Mutation of the target lysines to residues that cannot be acetylated abolishes the ability of PLZF to bind DNA, repress transcription, and suppress cell growth. Replacing wild-type (WT) PLZF protein by a mutant form (constitutive acetylation of PLZF [PLZFON]) in which target lysines K647/650/653 in zinc finger 9 were mutated to glutamine (Q), mimicking acetylated lysine residues, and revealed that acetylation stimulates PLZF DNA binding and promotes its transcription repression activity (22, 23).

It is currently unknown whether acetylation-based PLZF regulation plays a role in NKT cell development. Therefore, we addressed this question by analyzing the development of NKT cells in mice expressing PLZFON (23). Our findings indicated that active PLZF promotes the early steps of NKT cell development but inhibits the progression of NKT cell precursors to subsequent differentiation stages. In this article, we investigate this model further and show potential links between PLZF activation and specific DNA methylation patterns resulting in differential expression of genes related to commitment to the NKT lineage rather than genes controlling progress to later differentiation stages. Thus, our data suggest that specific epigenetic control of PLZF mediated by acetylation levels is crucial to regulating NKT cell differentiation.

PLZFON mice have been previously described (22). Jα18−/− and CD1d−/− mice are described elsewhere (3, 24). WT C57BL/6J mice were purchased from Janvier Laboratories; C57BL/6 Ly-5.1 mice were purchased from Charles River Laboratories. All mice were bred and maintained under specific pathogen–free conditions, and experiments were performed in accordance with the Institutional Animal Care and Use Guidelines. The study was approved by the local ethics committee Comité d’Ethique Paris-Nord (C2EA-121) affiliated with the Comité National de réflexion Ethique en Expérimentation Animale and to the French ministry for higher education and research.

Thymus, pooled peripheral lymph nodes (PLNs; comprising axillary, subaxillary, maxillary, inguinal, and popliteal lymph nodes), and spleen were isolated, mechanically disrupted, and filtered through a 40-μm stainless steel mesh to obtain single-cell suspensions. Bone marrow (BM) cell suspensions were obtained by flushing femurs and tibias with sterile medium, followed by gentle crushing of the suspension before filtration.

To measure intracellular cytokines, cells were stimulated with 50 ng/ml PMA (Sigma-Aldrich) and 1 µM ionomycin (Cell Signaling Technologies) in the presence of 5 µg/ml brefeldin A (Sigma-Aldrich) for 4 h.

Mice received a single intranasal administration of 2 µg α-GalCer (Kirin Brewery) or control PBS. Twenty-four hours later, mice were anesthetized by i.p. injection of ketamine and xylazine, and their airways were flushed twice with NaCl 0.9% solutions. The number of cells recovered in bronchoalveolar lavage fluid (BALF) was determined by counting.

Single-cell suspensions were incubated with anti-CD16/32 (2.4G2; BD) to block Fc receptors before staining with PE, allophycocyanin, or BV421 CD1d–α-GalCer–loaded tetramer, as previously described (25). Binding was revealed with fluorochrome-conjugated Abs diluted in FACS buffer (PBS containing 5% FCS and 0.02% sodium azide). Fluorochrome-conjugated Abs (purchased from BD, eBioscience, or BioLegend) were directed against the following molecules: FITC CD4 (RM4-5), PE-Cy5 CD8 (53-6.7), Alexa Fluor 700 TCRαβ (H57-597), BV510 HSA (M1/69), BV510 B220 (RA3-6B2), Alexa Fluor 700 CD44 (IM7), PE-Cy5 NK1.1 (PK136), PE-Cy7 NK1.1 (PK136), Alexa Fluor 647 IL-17RB (9B10), FITC CD45.1 (A20), PE CD45.2 (104), PE-Cy7 GR-1 (RB6-8C5), BV421 c-KIT (2B8), APC TCRγδ (GL3), PE PLZF (9E12), PerCP-EF710 RORγt (B2D), PE-Cy7 T-bet (eBIO4B10), APC T-bet (4B10), FITC Ki-67 (SoLA15), PE IL-17A (TC11-18H10), APC IL-17A (TC11-18H10), PerCP-EF710 IL-17A (eBIOGL3), PE-Cy7 IL-4 (11B11), BV421 IL-4 (11B11), and PerCP-EF710 IL-13 (eBIO13A). For intranuclear staining, cells were fixed and permeabilized after staining, then the Foxp3 staining kit was used according to the manufacturer’s instructions (eBioscience). For intracellular cytokine staining, cells were fixed with 2% paraformaldehyde (Sigma-Aldrich) and permeabilized with saponin. The annexin V/7-aminoactinomycin D (7-AAD) staining kit was used according to the protocol provided by BD Biosciences. Flow cytometry was performed on a four-laser LSR Fortessa cytometer (BD Biosciences) and analyzed using FACSDiva software v8 (BD Biosciences).

Thymi were pooled and depleted of CD8+ cells using the Dynabeads untouched mouse CD4 cells kit. Cells were then stained with anti-HSA Abs and CD1d tetramers; HSAlow CD1d-positive NKT cells were sorted on an FACSAria III (BD Biosciences) for microarray or methylated DNA immunoprecipitation sequencing (MeDIP-Seq) analysis.

Recipients (PLZFON or Jα18−/−) for chimera generation were lethally irradiated (9 Gy) before injecting 1 × 107 to 2 × 107 donor BM cells (C57BL/6J Ly-5.2, C57BL/6 Ly-5.1, or PLZFON) into the lateral tail vein within 24 h. All chimeric mice were allowed to reconstitute for at least 2 mo before analysis.

RNA was extracted with TRIzol reagent (Invitrogen), followed by a cleanup procedure using QIAGEN RNeasy Mini columns according to the manufacturer’s protocol. RNA concentrations were measured with a Nanodrop 2000/2000c (Thermo Fisher Scientific).

RNA samples were processed for hybridization on MoGene2.0.st chips (Affymetrix) by the Core Genomics Facility at Cochin Institute. For data preprocessing and normalization, raw chip data were extracted and summarized to the gene level on a log2 scale, quantile normalization was performed using the Affymetrix RMA-sketch routine, and annotation was based on Affymetrix MoGene2.0.st library files. Metabolic pathways were analyzed by applying gene set enrichment analysis (GSEA) (26). Data were obtained from whole-genome microarray datasets under Gene Expression Omnibus accession number GSE128558: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE128558. The same microarray data are used in all of the figures.

MeDIP-Seq was performed as previously described (27). A single assay was performed with pooled samples. Briefly, 1 × 105 freshly sorted thymic CD1d tetramer–positive NKT cells from PLZFON, and WT mice were digested with GenDNA Digestion Buffer and Proteinase K for 12–18 h at 50°C with shaking. Genomic DNA extracted from each sample was fragmented by sonication with a Diagenode Bioruptor to produce a median fragment length of 180–230 bp. The size range of genomic DNA fragments was assessed using a 2100 Bioanalyzer with DNA1000 chips (Agilent Technologies). Sonicated genomic DNA was then used for end repair and adaptor ligation according to the protocol provided with the iDeal Library Preparation Kit (Diagenode).

The reaction mix was purified using AMPure XP beads (Beckman Coulter). After adapter ligation, MeDIP-Seq was performed. Adaptor-ligated DNA was incubated with the anti–5-methylcytosine Ab (Diagenode), and methylated DNA immunoprecipitation (MeDIP) was performed using the MagMeDIP kit (Diagenode), including unmethylated and methylated spike-in controls. Incubation with Abs was performed at 4°C overnight. Immunoprecipitated samples were magnetically purified using the Auto iPure v2 kit (Diagenode) according to the manufacturer’s instructions. Recovery and enrichment were evaluated by quantitative PCR using primer sets specific for the spike-in controls. MeDIP experiments were considered successful when methylated fragments were specifically recovered (≥95%) compared with unmethylated fragments. After successful validation of the MeDIP reaction, immunoprecipitation and input DNA were amplified using the iDeal Library Preparation Kit (Diagenode) according to the manufacturer’s instructions. Samples were then purified and size selected with AMPure XP beads. The prepared library was sequenced using Illumina HiSeq 2500. The quality of sequencing reads was verified using FastQC. Adapters were removed using Trim Galore. Reads were then aligned to the mouse reference genome (mm10) using bwa, before identifying enriched regions and comparing them between the two conditions. The R packages MEDIPS was used for comparative analysis to identify regions that are differentially enriched between WT and PLZFON samples. To identify these regions, the difference in coverage, the CpG density, and the background pattern based on the input samples were all considered, and duplicate reads were removed. The threshold for significance of differences was set to q < 0.01, and a minimum 10 reads per 100-pb window was also applied. The Integrative Genomics viewer (version mm10) was then used to present the data by converting the output from the analysis pipeline into Browser Extensible Data files. MeDIP data are deposited under Gene Expression Omnibus accession number GSE171984: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE171984

FACS data are summarized as mean + SD. The statistical significance of differences between populations was assessed based on a nonparametric Mann–Whitney U test performed using GraphPad Prism software; p values ≤0.05 were considered significant.

PLZF is a significant factor in NKT cell development. To determine the impact of PLZF acetylation on this process, we analyzed NKT cell subsets in PLZFON mice. These mice express a mutant form of PLZF in which lysine residues K647/650/653 in zinc finger 9 were mutated to glutamine. The glutamine side chains mimic the structure of acetylated lysine residues, neutralizing of the positive charge (22). The mutated protein thus behaves like acetylated PLZF without the need for acetyltransferase activity. In vitro and in vivo systems indicated that these K-to-Q mutations appear to stimulate PLZF DNA binding and to promote its transcriptional repression (23). In PLZFON mice, the frequency and number of double-negative, DP, CD4+, and CD8+ T cells in the thymus and of T and B lymphocytes in the spleen and PLNs were normal (data not shown). The frequency and absolute numbers of thymic NKT cells, as assessed based on CD1d tetramer staining, were decreased in PLZFON mice compared with WT mice (Fig. 1A). We next assessed NKT cell subset distribution based on expression of the TFs T-bet (NKT1) and RORγt (NKT17). NKT2 cells are defined as lacking expression of both factors. In the thymus of PLFZON animals, we observed a drastic reduction in the frequency and absolute numbers of NKT1 and NKT17 cells. However, the frequency and absolute numbers of NKT2 increased (Fig. 1A). Although the frequency and absolute numbers of NKT cells in the spleen and PLNs of PLZFON mice were not significantly affected, analysis of NKT cell subsets showed, like in the thymus, an almost complete absence of NKT17 and a drastic reduction in NKT1 cells. These effects were compensated for by a corresponding increase in NKT2 cells (Fig. 1B, 1C). We went on to assess the status of other hematopoietic subsets dependent on PLZF for their development. PLZF is important for the development of IL-17–producing, fetal-derived Vγ6+ γδ T cells expressing RORγt (16) and IFN-γ–producing Vγ1 γδ T cells expressing T-bet (15). The frequency and absolute numbers of γδ T cells in the thymus of PLZFON mice was similar to that recorded for WT mice (Supplemental Fig. 1A). The distribution of RORγt+ and T-bet+ cells among γδ T cells was also normal (Supplemental Fig. 1A). The cytokine-production capacity of these γδ T cell subsets was unaltered, as assessed by intracellular IFN-γ and IL-17 staining after PMA/ionomycin stimulation (Supplemental Fig. 1A). Normal frequencies, numbers, subset distribution, and function of γδ T cells were also observed in PLNs and spleen (Supplemental Fig. 1B, 1C). We also assessed the status of myeloid subsets, which depend on PLZF for their development, and found similar frequencies of c-kit+ and Gr-1+ cells in BM from PLZFON and WT mice (Supplemental Fig. 1D).

FIGURE 1.

Constitutive activation of PLZF inhibits NKT cell differentiation. Representative staining of NKT1 (T-bet+), NKT2 (T-betRORγt), and NKT17 (RORγt+) subsets among thymic (A), PLN (B), and spleen (C) NKT cells from WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute numbers of these distinct subsets are represented (right). Data are pooled from four independent experiments in which at least three 7–8-wk-old mice were used in each experiment and presented as mean + SD. Significant results are indicated by an asterisk and were assessed by applying a nonparametric Mann–Whitney U test. *p < 0.05.

FIGURE 1.

Constitutive activation of PLZF inhibits NKT cell differentiation. Representative staining of NKT1 (T-bet+), NKT2 (T-betRORγt), and NKT17 (RORγt+) subsets among thymic (A), PLN (B), and spleen (C) NKT cells from WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute numbers of these distinct subsets are represented (right). Data are pooled from four independent experiments in which at least three 7–8-wk-old mice were used in each experiment and presented as mean + SD. Significant results are indicated by an asterisk and were assessed by applying a nonparametric Mann–Whitney U test. *p < 0.05.

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Overall, these results indicate that constitutive expression of an active form of PLZF in PLZFON mice affects the development of NKT cells but has no impact on the other PLZF-dependent myeloid or lymphoid cells tested in this study.

We next investigated whether constitutive expression of an active form of PLZF affected NKT cell function acquisition. Production of IFN-γ, IL-17, IL-4, and IL-13 was determined following PMA/ionomycin stimulation. The frequency and absolute numbers of thymic NKT cells producing IFN-γ and IL-17 were severely reduced in PLZFON mice, with almost no cells producing IL-17 (Fig. 2A). In contrast, the proportion of cells producing IL-4 and IL-13 increased drastically (Fig. 2A). These results concur with the findings presented above (Fig. 1A), showing a severe reduction in IFN-γ–producing NKT1 cells, an almost complete absence of NKT17 cells producing IL-17, and a dramatic increase in NKT2 cells producing IL-13 and IL-4. The same cytokine-production pattern was observed in PLNs and spleen (Fig. 2B, 2C) and correlates with the observed frequencies of NKT subsets in PLZFON mice (Fig. 1B, 1C). Previous reports indicated that lung NKT cells, through their IL-17 production, play a determinant role in recruiting neutrophils to airways upon LPS or α-GalCer instillation (28). To assess the consequences of the altered NKT cell function in PLZFON mice, we delivered α-GalCer intranasally to specifically target NKT cells and measured neutrophil recruitment in BALF. In the conditions used, neutrophil infiltration was only observed in the lung from WT, but not PLZFON or NKT cell–deficient CD1d−/−, animals (Fig. 2D). These results are consistent with the almost complete absence of IL-17–producing NKT cells in PLZFON mice.

FIGURE 2.

Constitutive activation of PLZF inhibits NKT cell functionality. Representative staining of IFN-γ, IL-17, IL-4, and IL-13 secretion among total NKT thymic (A), PLN (B), and spleen (C) NKT cells from WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute counts of secreted cytokines are represented (right). (D) CD1d−/−, WT, and PLZFON mice received a single intranasal (i.n.) dose of 2 μg α-GalCer and were sacrificed 24 h later. The absolute number of neutrophils recruited in BALF is indicated. Data from four independent experiments with at least three 7–8-wk-old mice were pooled for each experiment, and data are presented as mean + SD. Significant results are indicated by an asterisk; significance was determined by a nonparametric Mann–Whitney U test. *p < 0.05.

FIGURE 2.

Constitutive activation of PLZF inhibits NKT cell functionality. Representative staining of IFN-γ, IL-17, IL-4, and IL-13 secretion among total NKT thymic (A), PLN (B), and spleen (C) NKT cells from WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute counts of secreted cytokines are represented (right). (D) CD1d−/−, WT, and PLZFON mice received a single intranasal (i.n.) dose of 2 μg α-GalCer and were sacrificed 24 h later. The absolute number of neutrophils recruited in BALF is indicated. Data from four independent experiments with at least three 7–8-wk-old mice were pooled for each experiment, and data are presented as mean + SD. Significant results are indicated by an asterisk; significance was determined by a nonparametric Mann–Whitney U test. *p < 0.05.

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Overall, these results indicate that the expression of a constitutively active form of PLZF shuts off the development of NKT1 and NKT17 cells without affecting the development of NKT2 cells producing IL-4 and IL-13.

To determine at which stage NKT cell development was blocked in the thymus in PLZFON mice, we assessed CD44 and NK1.1 expression. These markers allow us to distinguish stages 1 to 3 of NKT cell development (9). In 3-wk-old mice, the frequency and absolute numbers of stage 1 NKT cells were increased in PLZFON mice, whereas the proportion of stage 3 NKT cells was markedly decreased (Fig. 3A, upper panel). The same pattern was observed in 12-wk-old PLZFON mice, indicating that the developmental defect is not corrected with age (Fig. 3A, lower panel). To identify which NKT cell subset is blocked at the CD44low stage, we analyzed CD44 expression in NKT cell subsets in 3-wk-old mice. The NKT1 cells that developed in PLZFON mice were almost all CD44high, like those observed in their WT littermates (Fig. 3B). In these young mice, T-betRORγt NKT cells represent both NKT2 and precursor cells, whereas in older mice, T-betRORγt cells almost exclusively represent NKT2 cells (29). In PLZFON mice, a higher proportion of T-betRORγt NKT cells were CD44low than in WT mice (Fig. 3B). The frequency of CD44low cells among T-betRORγt NKT cells remained high in 12-wk-old PLZFON mice, whereas their frequency decreased in WT littermate controls, in which they had progressed to the CD44high stage (Fig. 3B). Analysis of PLNs and spleen also showed an unusually high frequency of CD44low cells among T-betRORγt NKT cells from PLZFON animals compared with WT controls (data not shown).

FIGURE 3.

NKT cell maturation is blocked at the transition between stage 1 and stage 2 in PLZFON mice. (A) Representative staining of CD44 and NK1.1 among thymic NKT cells from 3-wk-old and 3-mo-old WT controls and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute numbers of these stages are represented (right). (B) Representative staining of CD44 among NKT1 (T-bet+), NKT17 (RORγt+), and T-betRORγt subsets in thymic NKT cells from WT control and PLZFON littermates. Frequency of CD44low among T-betRORγt cells in 3-wk-old and 3-mo-old WT controls compared with PLZFON littermates is represented (right). Data from four independent experiments using at least three mice in each experiment were pooled and presented as mean + SD. Asterisks indicate significant results, as determined by a nonparametric Mann–Whitney U test. *p < 0.05.

FIGURE 3.

NKT cell maturation is blocked at the transition between stage 1 and stage 2 in PLZFON mice. (A) Representative staining of CD44 and NK1.1 among thymic NKT cells from 3-wk-old and 3-mo-old WT controls and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies and absolute numbers of these stages are represented (right). (B) Representative staining of CD44 among NKT1 (T-bet+), NKT17 (RORγt+), and T-betRORγt subsets in thymic NKT cells from WT control and PLZFON littermates. Frequency of CD44low among T-betRORγt cells in 3-wk-old and 3-mo-old WT controls compared with PLZFON littermates is represented (right). Data from four independent experiments using at least three mice in each experiment were pooled and presented as mean + SD. Asterisks indicate significant results, as determined by a nonparametric Mann–Whitney U test. *p < 0.05.

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These results indicate that NKT cells in PLZFON mice have a defect preventing their progression from CD44low developmental stage 1 to CD44high stage 2 in the thymus and that the defect specifically affects T-betRORγt NKT cells.

The aberrant expression of CD44 in T-betRORγt cells from PLZFON mice suggests that these cells could represent immature NKT cell developmental intermediates rather than mature NKT2 cells. To distinguish between these two possibilities, we analyzed IL-17RB expression. Indeed, published results indicate that expression of this receptor is shared by early NKT cell precursors at CD44low stage 1 and is upregulated in mature NKT2 cells at CD44high stage 2 (10, 30). The frequency of IL-17RB+ NKT cells increased from stage 1 to stage 2 as expected in WT mice (62 versus 90%, respectively; see representative histogram plot in (Fig. 4A, upper panel) in line with the emergence of mature CD44high NKT2 cells at stage 2. In PLZFON mice in contrast, the frequency remained unchanged (66 versus 67%, respectively; see representative histogram plot in (Fig. 4A, lower panel), indicating that these cells probably represent cells at early stages of development rather than mature NKT2 cells.

FIGURE 4.

T-betRORγt NKT cells in PLZFON mice represent NKT cell precursors rather than NKT2 cells. (A) Representative staining of IL-17RB among RORγt/T-bet subsets in thymic stage 1 (CD44low) and stage 2 (CD44high) NKT cells in WT control and PLZFON littermates. Values indicated on dot and histogram plots represent frequencies. Frequency of IL-17RB expression is represented (right). (B) Representative staining of IL-17RB and the TF T-bet or RORγt [named TFs and defined based on IL-17RB substages (stages A, B, C, and D)–] among stage 1 (CD44lowNK1.1) and stage 2 (CD44highNK1.1) thymic NKT cells in WT control (left) and PLZFON (right) littermate mice. (C) Schematic representation of developmental stages of NKT cells defined based on their acquisition of IL-17RB and the TFs T-bet or RORγt in WT mice compared with PLZFON mice. The red cross in the diagram for PLZFON mice indicates a block at the (stage B)-to-(stage C) transition due to the lack of expression of TFs. In WT cells, these genes are mainly expressed at the CD44high stage 2 NKT cells; their lack indicates a block in NKT cell subset differentiation. Data shown in (A) and (B) were pooled from four independent experiments, using at least three 3–4-wk-old mice in each experiment, and results are presented as mean + SD. Significant results are indicated by an asterisk; significance was determined using a nonparametric Mann–Whitney U test. *p < 0.05.

FIGURE 4.

T-betRORγt NKT cells in PLZFON mice represent NKT cell precursors rather than NKT2 cells. (A) Representative staining of IL-17RB among RORγt/T-bet subsets in thymic stage 1 (CD44low) and stage 2 (CD44high) NKT cells in WT control and PLZFON littermates. Values indicated on dot and histogram plots represent frequencies. Frequency of IL-17RB expression is represented (right). (B) Representative staining of IL-17RB and the TF T-bet or RORγt [named TFs and defined based on IL-17RB substages (stages A, B, C, and D)–] among stage 1 (CD44lowNK1.1) and stage 2 (CD44highNK1.1) thymic NKT cells in WT control (left) and PLZFON (right) littermate mice. (C) Schematic representation of developmental stages of NKT cells defined based on their acquisition of IL-17RB and the TFs T-bet or RORγt in WT mice compared with PLZFON mice. The red cross in the diagram for PLZFON mice indicates a block at the (stage B)-to-(stage C) transition due to the lack of expression of TFs. In WT cells, these genes are mainly expressed at the CD44high stage 2 NKT cells; their lack indicates a block in NKT cell subset differentiation. Data shown in (A) and (B) were pooled from four independent experiments, using at least three 3–4-wk-old mice in each experiment, and results are presented as mean + SD. Significant results are indicated by an asterisk; significance was determined using a nonparametric Mann–Whitney U test. *p < 0.05.

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To further characterize the early developmental block of NKT cells in PLZFON mice, we examined the expression of IL-17RB and the TFs T-bet and RORγt in developing NKT cells. In WT mice, this analysis showed that IL-17RB is acquired by stage 1 CD44low cells (see red gates in (Fig. 4B, left panel) before the TFs, which are mainly acquired at stage 2, CD44high (see blue gates in (Fig. 4B, left panel). This observation suggests a four-step developmental sequence from stage 1 to stage 2 (Fig. 4B, middle panel). Analysis of thymic NKT cells from PLZFON mice revealed that NKT cells acquire IL-17RB expression at stage 1 (see red gates in (Fig. 4B, left panel) but that the few cells that progress to stage 2, CD44high, fail to appropriately express the TFs (see blue gates in (Fig. 4B, right panel).

Overall, our results indicate that CD44low immature stage 1 NKT cells in WT mice pass through an IL-17RB+ stage before expressing specific TFs (i.e., RORγt+ and T-bet+), mainly from stage 2 of development, in confirmation of our previous results (31). In PLZFON mice, these cells acquire IL-17RB, but not the TFs, and thus remain blocked at the transition between CD44low stage 1 and CD44high stage 2 (see model in (Fig. 4C).

We next wanted to determine whether the reduced numbers of thymic NKT cells in PLZFON mice was due to altered proliferation. We therefore measured the capacity of thymic NKT cells to proliferate in vivo by determining the frequency of Ki-67–positive cells in 4-wk-old mice. As shown in (Fig. 5A, upper panel, Ki-67+ cells among total NKT cells were more frequent in PLZFON mice than in WT mice (59 versus 28% Ki-67+ cells, respectively, as shown in representative histograms). Gene expression profiles for WT versus PLZFON thymic NKT cells support enhanced proliferation in PLZFON mice based on GSEA (Fig. 5B, left panel) (see Materials and Methods for microarray data). When we examined Ki-67 expression at different developmental stages, we found that stage 1 NKT cells from PLZFON mice express Ki-67 at a higher frequency (∼81%) compared with the equivalent population in WT mice (∼63%) (Fig. 5A, upper panel). The contrary was observed at stage 2, in which NKT cells from PLZFON mice expressed Ki-67 to a lesser extent (40%) than the equivalent population in WT mice (60%, as shown in representative histograms highlighted with a red gate) (Fig. 5A, upper panel). No difference was observed at stage 3, during which cells are in a nonproliferating state in WT mice (Fig. 5A, upper panel). These results indicate that the proliferation status of NKT cells is altered in PLZFON mice, starting with a high proliferation rate at stage 1. Unlike in WT mice, this proliferation is not sustained at stage 2, and this finding likely contributes to the reduced numbers of cells at this stage in PLZFON animals.

FIGURE 5.

NKT cell proliferation and mortality are affected in PLZFON mice. (A) Representative staining of Ki-67 (upper panel) and 7-AAD/annexin V–positive cells (lower panel) among stage 1, 2, and 3 or total thymic NKT cells in WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies are represented on histograms. Data were pooled from four independent experiments, using at least four 4-wk-old mice in each experiment, and results are presented as mean + SD. Significant results are indicated by an asterisk and were determined using a nonparametric Mann–Whitney U test. *p < 0.05. (B) GSEA of proliferation, apoptosis, and p53 signaling in thymic NKT cells from WT control and PLZFON mice. Microarray data are the same as those used in (Fig. 7.

FIGURE 5.

NKT cell proliferation and mortality are affected in PLZFON mice. (A) Representative staining of Ki-67 (upper panel) and 7-AAD/annexin V–positive cells (lower panel) among stage 1, 2, and 3 or total thymic NKT cells in WT control and PLZFON littermates. Values indicated on dot plots represent frequencies. Frequencies are represented on histograms. Data were pooled from four independent experiments, using at least four 4-wk-old mice in each experiment, and results are presented as mean + SD. Significant results are indicated by an asterisk and were determined using a nonparametric Mann–Whitney U test. *p < 0.05. (B) GSEA of proliferation, apoptosis, and p53 signaling in thymic NKT cells from WT control and PLZFON mice. Microarray data are the same as those used in (Fig. 7.

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We next examined whether reduced NKT cell numbers in PLZFON mice could also be caused by increased cell death. To do so, thymic NKT cells were stained with 7-AAD and annexin V. The proportion of apoptotic and dead cells was significantly increased in total NKT cells from PLZFON mice compared with WT mice (60 versus 22% annexin V+ cells, respectively, as shown in representative dot plots) (Fig. 5A, lower panel). GSEA analysis also indicated higher apoptosis in PLZFON mice (Fig. 5B, middle and right panel) (see Materials and Methods for microarray data). In particular, cell death and apoptosis were specifically increased in stage 2 NKT cells in PLZFON mice (67% annexin V+ cells versus 40% in WT controls, as shown in representative dot plots highlighted with a red gate) (Fig. 5A, lower panel, see red gate). These results indicate that increased apoptosis of thymic NKT cells in PLZFON mice may also contribute to their reduced numbers.

Taken together, our results indicate that the reduced number of thymic NKT cells in PLZFON mice is caused by their reduced proliferation and increased apoptosis when they reach stage 2 of their thymic development.

To exclude the possibility that impaired development of thymic NKT cells was due to a failure of Ag presentation, we examined expression of CD1d on DP thymocytes in PLZFON and WT control. No difference was noted (data not shown). To investigate whether the NKT cell developmental defect was cell intrinsic, we generated mixed BM chimeras by transferring a 1:1 ratio of WT and PLZFON BM donor cells into sublethally irradiated Jα18−/− hosts, which lack NKT cells (Fig. 6, upper panel). After 2 mo of engraftment, we isolated cells from the thymus and determined the ability of each donor compartment to reconstitute NKT cells. Whereas WT donor cells fully reconstituted NKT cell developmental subsets and stages, NKT cells in the PLZFON donor compartment displayed the same NKT cell developmental defect as observed in the PLZFON mice or in single PLZFON ≥ WT chimeras (Fig. 6; data not shown). These results indicate that the presence of a WT stromal or BM compartment could not rescue the NKT developmental defect in the PLZFON compartment, confirming that the defect is inherent to the NKT cells.

FIGURE 6.

PLZF constitutive activation is intrinsic. Lethally irradiated CD45.2 Jα18KO mice were reconstituted with BM from CD45.1 C57BL/6 (WT) and CD45.2 PLZFON mice (scheme in upper panel). NKT-enriched thymic cells were analyzed 8 wk after BM reconstitution. Representative staining of CD44/NK1.1 and RORγt/T-bet expression among iNKT cells is shown (upper panel). Values in dot plots represent frequencies. Frequency of total iNKT cells, the iNKT cell subsets indicated, and developmental stages are represented (lower panel). Data were pooled from at least four independent experiments in which four to five mice in each experiment were used and presented as mean + SD. Significant results are indicated by an asterisk; significance was determined by a nonparametric Mann–Whitney U test. *p <0.05.

FIGURE 6.

PLZF constitutive activation is intrinsic. Lethally irradiated CD45.2 Jα18KO mice were reconstituted with BM from CD45.1 C57BL/6 (WT) and CD45.2 PLZFON mice (scheme in upper panel). NKT-enriched thymic cells were analyzed 8 wk after BM reconstitution. Representative staining of CD44/NK1.1 and RORγt/T-bet expression among iNKT cells is shown (upper panel). Values in dot plots represent frequencies. Frequency of total iNKT cells, the iNKT cell subsets indicated, and developmental stages are represented (lower panel). Data were pooled from at least four independent experiments in which four to five mice in each experiment were used and presented as mean + SD. Significant results are indicated by an asterisk; significance was determined by a nonparametric Mann–Whitney U test. *p <0.05.

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To investigate the impact of the PLZFON mutation on general gene expression in NKT cells and attempt to elucidate the molecular mechanisms by which constitutive expression of active PLZF affected NKT cell development, we generated gene expression profiles for sorted thymic NKT cells from PLZFON and WT mice. RNA was isolated from CD1d tetramer–positive NKT cells, shown in the postsort gates in (Fig. 7A. Unsupervised analysis conducted on the 30,920-probe sets obtained for duplicate samples of WT and PLZFON yielded 197 genes for which the coefficient of variation was >0.2 (Supplemental Fig. 2). This unsupervised analysis can be used to discriminate between WT and PLZFON samples using hierarchical clustering with Euclidean distance between samples (Supplemental Fig. 2). In accordance with FACS analysis, genes related to NKT cell maturation programs (e.g., the NK lineage–related klr gene family) are downregulated in PLZFON cells.

FIGURE 7.

PLZF is required for the lineage-specific expression of signature genes in NKT cell development. (A) Total thymic cells from control WT and PLZFON 4-wk-old littermates were CD8 depleted. Representative dot plots show pre- and postsorted NKT cells stained with CD1d tetramer (CD1d-tet), HSA, CD44, and NK1.1. Values indicated on dot plots represent frequencies. CD1d-tet–positive NKT cells shown in the postsort gates correspond to the population used for RNA isolation (B) GSEA of pathways upregulated in WT (n = 2) (left) and PLZFON (n = 2) (right) mice. (C) Heatmap of selected immune genes classified as TFs, cytokines, chemokines, and receptors, and cell-surface molecules in thymic NKT cells from WT control compared with PLZFON littermates.

FIGURE 7.

PLZF is required for the lineage-specific expression of signature genes in NKT cell development. (A) Total thymic cells from control WT and PLZFON 4-wk-old littermates were CD8 depleted. Representative dot plots show pre- and postsorted NKT cells stained with CD1d tetramer (CD1d-tet), HSA, CD44, and NK1.1. Values indicated on dot plots represent frequencies. CD1d-tet–positive NKT cells shown in the postsort gates correspond to the population used for RNA isolation (B) GSEA of pathways upregulated in WT (n = 2) (left) and PLZFON (n = 2) (right) mice. (C) Heatmap of selected immune genes classified as TFs, cytokines, chemokines, and receptors, and cell-surface molecules in thymic NKT cells from WT control compared with PLZFON littermates.

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Supervised analysis of data for WT and PLZFON samples, with a false discovery rate of 20%, yielded 312 regulated genes (168 upregulated and 144 downregulated genes). GSEA of the genes downregulated in PLZFON NKT cells revealed enrichment of genes encoding products related to NKT cells and involved in NKT cell differentiation, as well as genes involved in signaling pathways, including IFNs, IL-12, and IL-17 (Fig. 7B, left panel). These signaling molecules have been reported to play essential roles in NKT cells (32). In contrast, the genes upregulated in response to constitutive expression of active PLZF-encoded products involved in the cell cycle, cytoskeleton organization, mitochondrial activity, and stemness (Fig. 7B, right panel).

(Fig. 7C shows a heatmap of selected immune genes that are directly or indirectly regulated by PLZF, as reported by Mao et al. (33). These genes are classified as cell TFs, cytokines/chemokines and receptors, and other cell-surface molecules. In WT mice, downregulation of the Sell gene, which encodes CD62L, characterizes the NKT cell effector program, and the gene product is required for recirculation to lymph nodes (in (Fig. 7D, upregulated genes in PLZFON that will be mentioned hereafter are indicated by a red rectangle, downregulated genes by a green rectangle). PLZF was shown to directly bind this gene in NKT cells (33). Furthermore, PLZF binds and downregulates the TF Klf2, a major positive regulator of Sell (34, 35). Expression of Sell and Klf2 in NKT cells from PLZFON mice was upregulated compared with WT controls (Fig. 7C). Cd44 expression was downregulated in NKT cells from PLZFON in line with its cell-surface expression profile (Figs. 3A, 7C). Thus, although PLZF directly regulates the CD44high CD62Llow nonrecirculating effector phenotype of NKT cells in WT mice, the active form of PLZF promotes the generation of a reverse CD44low CD62Lhigh naive-like phenotype.

PLZF was previously shown to directly bind the promoters and activate the expression of Il12rb1 and Il18r1, as well as Ifngr1, in NKT cells (33). It is also known to activate expression of IL-1R1 and IL-23R. Expression of all these cytokine receptors was downregulated in PLZFON mice relative to WT mice (Fig. 7C).

Not all cytokine or chemokine receptor genes controlled by PLZF reacted in the same way. For example, native PLZF binds and activates Ccr4, which encodes a chemokine receptor attracting cells toward CCL17-producing DCs and enhances Ag cross-presentation to CD8 T cells (36). In NKT cells from PLZFON mice, CCR4 expression was not repressed. Interestingly, like IL-17RB, CCR4 is expressed in the early immature stages of NKT cell development and on NKT2 cells (37). The results for these two genes suggest that genes expressed early during NKT cell development are not repressed by the active form of PLZF.

Data from the literature indicate that although PLZF does not bind cytokine loci encoding IL-4, IL-13, IFN-γ, or IL-17, it directly binds and/or activates multiple Th-specific TFs (33). These factors include Runx3 and Rorc, and their gene expression was repressed in NKT cells from PLZFON mice. Tbx21, which is indirectly activated by PLZF, was also repressed in NKT cells from PLZFON mice. In contrast, Gata3, which is expressed early during NKT cell development, was not repressed. Thus, the acetylated form of PLZF might be needed to initiate the early stages of NKT cell development, characterized in particular by Gata3, CCR4, and IL17RB induction while limiting the expression of other genes promoting the differentiation of NKT cell subsets.

The TF Bach2 is normally highly expressed in naive T cells and downregulated in effector T cells (38, 39). According to our data, Bach2 was profoundly directly repressed by PLZF in WT NKT but upregulated in NKT cells from PLZFON mice. Other genes such as themis and CCR9 are expressed early during T cell development, and their expression is downregulated as cells mature. These two genes are expressed by CD4+CD8+ thymocytes and are important for thymocyte selection and localization, respectively (40). Interestingly, we found that expression of both genes was upregulated in NKT cells from PLZFON mice, suggesting that these cells resemble early developing NKT cells rather than mature cells (Fig. 7C). LEF1 has been reported to be required for the transition of NKT0 cells to other stages by inducing the expression of Cd127 (which encodes the cytokine receptor IL-7Ra [CD127]) and Myc (coding for the TF c-Myc). LEF1 expression is reported to be increased in the rare NKT0 cells (41). In PLZFON NKT cells, we found that expression of LEF1 was upregulated, supporting the notion that these cells are blocked at the precursor stage of NKT cell development. A recent study by the Hogquist group (42) indicated that immature iNKT cells expressing CCR7 represent a multipotent progenitor pool. We found upregulated expression of CCR7 in iNKT cells from PLZFON mice (Fig. 7C). In addition, we found that the frequency of CCR7+ iNKT cells at the immature CD44low stage 1 was higher in PLZFON mice than in WT mice (data not shown).

Overall, these findings, although not providing mechanistic insights, support the notion that active PLZF promotes the early steps of NKT cell development, but not the efficient progression of iNKT cells to subsequent differentiation stages.

PLZF is known to shape the cellular genome by inducing a specific pattern of epigenetic marks, including DNA methylation (23). In an attempt to understand the specific gene expression signature induced by PLZF, we characterized the DNA methylation status in whole-thymic NKT cells from PLZFON and WT mice. To do so, we isolated thymic NKT cells from these mice by cell sorting and analyzed the genome-wide distribution of DNA methylation, at CpG sites, using the anti–5-methylcytidine Ab to immunoprecipitate methylated DNA (MeDIP), followed by next-generation sequencing. MeDIP-Seq reads were quantified to obtain a DNA methylation score in overlapping 1-kb bins across the genome. This score provides a high-resolution assessment of the distribution of DNA methylation patterns throughout the genome. In total, 143,801 common methylated CpG regions were validated in PLZFON and WT samples (Supplemental Fig. 3A). We also found 103,931 methylated CpG regions specific to WT mice and 34,742 CpG methylated regions specific to PLZFON mice (Supplemental Fig. 3A). The DNA methylation pattern was differentially distributed between WT and PLZFON NKT cells, highlighting regions of high DNA methylation preferentially located in intergenic regions (87.87% of CpG methylated regions) in PLZFON samples (Supplemental Fig. 3B). We identified a total of 66 specific differentially methylated regions (DMRs) induced by activated PLZF (Supplemental Fig. 3C). Of these DMRs, 58 were distributed across and enriched in intergenic regions on chromosomes 2, 9, 12, 14, and Y. Chromosome 9 was particularly extensively targeted, bearing 48 DMRs (Supplemental Fig. 3C). An expanded view of the methylation tracks in all regions of all chromosomes, comparing PLZFON to WT NKT cells, is shown in Supplemental Fig. 3D. These results reveal nonidentical DNA methylation signatures for PLZFON and WT samples, with an overall hyper methylation of the genome in the intergenic regions observed in several chromosomes from PLZFON cells. The differences in methylation may reflect differences in the type of regulation that DNA methylation exerts on several genes in the NKT cells’ developmental network. For instance, in PLZFON mice, greater methylation variations were noted in actively transcribed genes compared with genes with reduced activity (Supplemental Fig. 3E, for representative genes, and Supplemental Fig. 3F for total genes showing significance above baseline of average methylation variation of 0.054 versus 0.014 for downregulated and upregulated genes, respectively; see Supplemental Fig. 4 for total gene list). These results suggest that the repressive action of PLZFON in mice might be less reflected in terms of variations in methylation at the gene level. Interestingly, this distinction in methylation levels was more obvious for the group of TF genes (Supplemental Fig. 3F, for TFs showing significance above baseline of 0.438 versus 0.016 for downregulated and upregulated genes, respectively, see Supplemental Fig. 4 for total gene list). These results highlight differential methylation between NKT-related active and repressed genes in PLZFON mice, suggesting differences in methylation-related control of their expression, particularly for TFs.

To assess a possible generalized effect on gene expression induced by PLZF, and knowing that PLZF can specifically target repeat elements [such as transposable elements (23)], we analyzed genomic distribution of NKT-related gene aforementioned. We detected a high frequency of downregulated TF genes located in telomeric regions (82%), compared with an almost complete absence from the opposing centromeric region (6%) (Supplemental Fig. 3G, left panel, see Supplemental Fig. 4 for gene list). The mirror image was observed for actively transcribed TFs, with only 28% of genes coding for these factors located in the telomeric region of chromosomes (Supplemental Fig. 3G, right panel). These results suggest that specific clustering of NKT cell-related genes to the telomeric region might make it possible for PLZF to maintain overall control over their expression.

Taken together, our results suggest that acetylated PLZF induces a specific methylation signature that may provide the necessary activation signals related to early stages of NKT cell development while repressing specification signals. This process probably exists to avoid uncontrolled premature differentiation.

In this study, we analyzed the consequences of the presence of a constitutively acetylated form of PLZF on NKT cell development. We found that although activated PLZF promoted the development of early NKT cell precursors, it repressed NKT cell subset differentiation. Thus, in WT mice, regulation of PLZF acetylation/deacetylation controls NKT cell differentiation.

In PLZFON mice, we observed decreased thymic NKT cell numbers that could be explained by reduced expansion of NKT cells at stage 2 associated with increased apoptosis of cells entering this stage. Interestingly, NKT cell proliferation in these mice increased at stage 1 before being reduced at stage 2. PLZF was previously shown to inhibit myeloid cell growth and differentiation (43). In this lineage, PLZF expression dramatically suppressed growth and was associated with accumulation of cells at the G0/G1 stage of the cell cycle and an increased incidence of apoptosis. Because PLZF expression protected myeloid lineages from apoptotic death after IL-3 withdrawal, it was hypothesized that the increased apoptosis observed might be due to a clash in cellular growth signals (43). By analogy, developing NKT cells in PLZFON mice may receive a signal to proliferate from IL-7 at the same time as an antiproliferative signal from PLZF, which is known to repress genes involved in cell cycle progression. In line with this hypothesis, expression of LEF1 was increased in NKT cells from PLZFON mice; this TF controls IL-7R and c-Myc expression (41). Along the same lines, our results recall data showing that serum deprivation, which is a growth-inhibitory signal, clashes with constitutive c-Myc or E2F expression to induce apoptosis (44, 45). Significantly, the G2 checkpoint, E2F targets, and mitotic spindle were the most upregulated pathways in thymic NKT cells from PLZFON mice.

In addition to their reduced numbers, NKT cell differentiation was altered in PLZFON mice, with a dramatic reduction in NKT1 and NKT17 cells and a relative accumulation of T-betRORγt NKT cells. We believe that these cells represent NKT cell precursors rather than the mature NKT2 cell subset. The first argument in favor of this hypothesis is the fact that we found a block at the CD44low stage of development (stage 1). This stage corresponds to immature NKT cell subsets; mature stages of development are characterized by the upregulation of CD44. The block at the CD44low stage was not the result of delayed maturation because, in 12-wk-old mice, the CD44low NKT cells had not progressed to the CD44high stage. The second argument relates to the expression of IL-17RB on these cells. Mature NKT2 cells are reported to express higher level of IL-17RB than mature NKT17 and NKT1 cells (10, 46), suggesting that this surface marker could be used to define this subpopulation. In WT mice, we observed an increased frequency of IL-17RB+ cells among CD44high cells compared with CD44low cells, reflecting the emergence of mature NKT2 cells at this stage. However, in PLZFON animals, although a considerable proportion of CD44high cells was present among the T-betRORγt NKT cells, the frequency of IL-17RB+ cells in the CD44high population was not increased compared with among CD44low NKT cells. This result indicates that the total population of T-betRORγt NKT cells, regardless of CD44 expression, represents NKT cells at early developmental stages rather than mature NKT2 cells. Specific genes, such as LEF1 and CCR9, are reported to be upregulated at the NKT0 stage of development (37). Expression of LEF1 and CCR9 were effectively upregulated in NKT cells from PLZFON mice, further supporting the hypothesis that these cells are immature NKT cells rather than mature NKT2 cells. As further evidence, we also observed an increase in the proportion of cells expressing CCR7, the specific marker of NKT cell precursors (42).

Based on the expression of IL-17RB and the TFs T-bet and RORγt, our results indicated that the developmental block of NKT cells in PLZFON mice is linked to their incapacity to express TFs in response to IL-17RB signaling. In WT mice, NKT cell precursors passed through an IL-17RB+ stage at the CD44low stage 1 before specific TFs were expressed, mainly during the CD44high stage 2 (see model (Fig. 4C). According to this sequence, all maturing NKT cell precursors pass through an IL-17RB+ stage during development, a scenario that supports our earlier results (31) and completes previous studies relating to the developmental steps during NKT cell differentiation (47, 48).

Transcriptomic analysis of purified NKT cells from WT and PLZFON mice indicated that PLZF activation by acetylation promotes the entry of NKT cells into a specific program impeding the signal to proceed with differentiation. This appears to occur through the repressive action of PLZF by inducing DNA methylation of specific genomic regions to restrict the expression of differentiating genes and allow the activation of genes expressed during early NKT cell development.

Several questions arise from this analysis: the first is how PLZF controls the repression of multiple genes simultaneously? A previous study showed that in addition to the ability of PLZF to remodel chromatin by recruiting HDACs to regulatory PLZF binding sites, its homodimerization status could mediate long-range interactions between regulatory regions containing several PLZF binding sites. This capacity may play an important role in coordinating the spatial expression of distant genes within a locus (49). In addition, PLZF was shown to interact with polycomb group members such as Bmi-1, allowing it to act as a trans-repressive factor controlling target gene expression. Acetylated PLZF may use similar stratagems to suppress gene activity during early NKT cell developmental stages. In support of this suggestion, previous reports indicated that PLZF epigenetic functions could be driven by regions located at a distance from promoter regions, such as intronic (21), 3′ gene regulatory (23), and enhancer regions (50). Thus, PLZF could exert its repressive activity over long distances by inducing chromatin spreading or looping mechanisms (21, 49). Chromatin immunoprecipitation sequencing experiments may help to determine which genes are directly associated with active PLZF during the early stages of NKT cell development. In addition, three-dimensional DNA experiments could be used to investigate spatial control of active PLZF on gene repression. Our attempts to perform such experiments have been unsuccessful so far because of a limited number of NKT cell subsets.

The second question is how PLZF, while promoting repression, simultaneously allows activation of genes, some of which were described to interact directly with PLZF. The repressive action of PLZF may require the recruitment of repressive factors to regulatory regions containing multiple PLZF binding sites, leading to accumulation of these factors and, thus, impeding repression at other geographically opposed PLZF binding sites. This situation would establish a gradient of positive and negative transcriptional influence governing the spatial expression of genes expressed at different stages of NKT cell development. This scenario is supported by our evidence of a close distribution of late-stage differentiating TFs in telomere-rich PLZF regulating regions, leading to general repression of these genes while concomitantly allowing expression of geographically distant TFs to promote early NKT cell development. Thus, in addition to allowing CD1d-selected cells to enter into the NKT cell program, PLZF, through its dual action, avoids uncontrolled premature differentiation.

The third question is how do physiological PLZF acetylation levels promote NKT cell development? PLZF acetylation was shown to be associated with the acetylase p300 (22); it is deacetylated by both HDAC3 and the NAD+-dependent deacetylase silent mating type information regulation 2 homolog 1 (SIRT1) (51). Interestingly, our analysis of a previously published dataset (52) indicated that CBP/p300 is expressed at stage 1 of NKT cell development (Fig. 8A). In addition, gene expression analysis indicated that although CBP/p300 expression decreased between stages 1 and 2, the expression of HDAC3 and SIRT1 increased (Fig. 8A); thus, these genes follow reverse regulation during NKT cell development. Consequently, we propose that CBP/p300 and HDAC/SIRT1-mediated PLZF acetylation/deacetylation could potentially allow rapid control and fine-tuning of PLZF activity through posttranscriptional modification of regulatory gene expression and normal NKT cell development (see model in (Fig. 8B). This hypothetical model does not exclude the possibility that PLZF undergoes passive deacetylation during NKT cell development.

FIGURE 8.

Ep300 and Hdac3/Sirt1 expression are differently regulated during NKT cell development. (A) Heatmap of selected genes in the various subsets of steady-state thymic differentiation from a published dataset (52). In the heatmaps, rows are mean centered and normalized, and local scaling was applied. Averaged data were combined from at least three independent experiments for each population, with cells pooled from three or more mice in each experiment. (B) A hypothetical model showing the potential role of PLZF acetylation/deacetylation in the emergence of NKT cells in WT mice (upper panel). At stage 1, PLZF acetylation allows PLZF to bind to target genes on which it exerts its repressive action. This acetylation could be mediated by the acetylase EP-300, as reported in other cell types (22). Loss of PLZF-associated repression occurs at stage 2 after passive or active PLZF deacetylation, allowing gene expression. Active deacetylation could potentially be mediated by HDAC3/SIRT1 deacetylase, as reported for other cell types (51). When PLZF is continuously acetylated in PLZFON mice, lineage differentiation is blocked because of the continuous repressive action of PLZF due to its fixed acetylation state and thus absence of its acetylation/deacetylation regulation (lower panel).

FIGURE 8.

Ep300 and Hdac3/Sirt1 expression are differently regulated during NKT cell development. (A) Heatmap of selected genes in the various subsets of steady-state thymic differentiation from a published dataset (52). In the heatmaps, rows are mean centered and normalized, and local scaling was applied. Averaged data were combined from at least three independent experiments for each population, with cells pooled from three or more mice in each experiment. (B) A hypothetical model showing the potential role of PLZF acetylation/deacetylation in the emergence of NKT cells in WT mice (upper panel). At stage 1, PLZF acetylation allows PLZF to bind to target genes on which it exerts its repressive action. This acetylation could be mediated by the acetylase EP-300, as reported in other cell types (22). Loss of PLZF-associated repression occurs at stage 2 after passive or active PLZF deacetylation, allowing gene expression. Active deacetylation could potentially be mediated by HDAC3/SIRT1 deacetylase, as reported for other cell types (51). When PLZF is continuously acetylated in PLZFON mice, lineage differentiation is blocked because of the continuous repressive action of PLZF due to its fixed acetylation state and thus absence of its acetylation/deacetylation regulation (lower panel).

Close modal

Our study is reminiscent of that of Wang et al. (53), showing that the histone acetyltransferase GCN5 catalyzes EGR2 lysine acetylation and regulates NKT cell development. GCN5 does not appear to impair NKT cell development through effects on the expression of multiple acetyltransferases such as p300; rather, it controls the expression of Runx1, T-bet, PLZF, and IL-2Rβ. Because Egr2 is expressed at stage 0 and promotes PLZF expression, which is observed mainly at stage 1 (12), it appears that distinct acetyltransferases control the acetylation of NKT-related master TFs at successive developmental stages. Thus, GCN5 and p300 control the acetylation of EGR2 and PLZF at stages 0 and 1, respectively. Importantly, our model also suggests that regulation of acetylation levels of these master TFs during NKT cell development is required to allow appropriate development and specialization of NKT cells. This hypothesis is supported by the results presented by Kasler et al. (54), showing that HDAC7 could regulate NKT cell development by directly interacting with PLZF to silence its transcriptional activity.

Overall, the findings presented in this paper indicate that active PLZF promotes the early steps of NKT cell development but inhibits the progression of NKT cell precursors to subsequent differentiation stages. This is reminiscent of the previously described function of PLZF in myeloid cell development, in which PLZF was shown to be involved in maintaining the progenitor pool (55, 56). Our study suggests that specific epigenetic control of PLZF through modulation of acetylation levels regulates normal NKT cell differentiation. Further studies will be required to precisely define upstream signals, allowing the control of PLZF acetylation levels.

We thank staff at Institut de Recherche Saint-Louis genomic and FACS facility, particularly Christelle Doliger and Sophie Duchez, and at the institute’s animal facility, particularly Veronique Parietti and Marika Pla. We are indebted to the National Institutes of Health Tetramer Facility for providing CD1d tetramers, and all members of the Benlagha group for helpful discussion, particularly Ines and Elies Benlagha.

This work was supported by INSERM, idex-SLI (Grant DXCAIHUSLI/SPC-KB14); idex-MelaTNK (Grant DXCAR1MTNK/SPC-23S1L) (to K.B. and J.K.); Université Paris Diderot, Ministère de l’Enseignement supérieur, de la Recherche et de l’Innovation (to C.J.); and the Fondation pour la Recherche Médicale (Grant FDT20160434991) (to C.J.).

The microarray datasets presented in this article have been submitted to Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/) under accession number GSE128558.

The online version of this article contains supplemental material.

Abbreviations used in this article

7-AAD

7-aminoactinomycin D

BALF

bronchoalveolar lavage fluid

BM

bone marrow

DMR

differentially methylated region

DP

double-positive

α-GalCer

α-galactosylceramide

GSEA

gene set enrichment analysis

HDAC

histone deacetylase

HSA

heat stable Ag

iNKT

invariant NKT

MeDIP

methylated DNA immunoprecipitation

MeDIP-Seq

methylated DNA immunoprecipitation sequencing

PLN

peripheral lymph node

PLZF

promyelocytic leukemia zinc finger

PLZFON

constitutive acetylation of PLZF

SIRT1

silent mating type information regulation 2 homolog 1

TF

transcription factor

WT

wild-type

1.
Bendelac
A.
,
M. N.
Rivera
,
S. H.
Park
,
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The authors have no financial conflicts of interest.

Supplementary data