Menin, a tumor suppressor protein, is encoded by the MEN1 gene in humans. Certain germinal mutations of MEN1 induce an autosomal-dominant syndrome that is characterized by concurrent parathyroid adenomas and several other tumor types. Although menin is also expressed in hematopoietic lineages, its role in CD8+ T cells remains unclear. We generated Meninflox/flox CD4-Cre (Menin-KO) mice by crossing Meninflox/flox mice with CD4-Cre transgenic (Tg) mice to determine the role of menin in CD8+ T cells. Wild-type (WT) and Menin-KO mice were infected with Listeria monocytogenes expressing OVA to analyze the immune response of Ag-specific CD8+ T cells. Menin deficiency resulted in an impaired primary immune response by CD8+ T cells. On day 7, there were fewer Menin-KO OVA-specific CD8+ T cells compared with WT cells. Next, we adoptively transferred WT and Menin-KO OT-1 Tg CD8+ T cells into congenic recipient mice and infected them with L. monocytogenes expressing OVA to determine the CD8+ T cell–intrinsic effect. Menin-KO OT-1 Tg CD8+ T cells were outcompeted by the WT cells upon infection. Increased expression of Blimp-1 and T-bet, cell cycle inhibitors, and proapoptotic genes was observed in the Menin-KO OT-1 Tg CD8+ T cells upon infection. These data suggest that menin inhibits differentiation into terminal effectors and positively controls proliferation and survival of Ag-specific CD8+ T cells that are activated upon infection. Collectively, our study uncovered an important role for menin in the immune response of CD8+ T cells to infection.
The main function of Ag-experienced CD8+ T cells is to eliminate tumors and cells infected with intracellular pathogens (1, 2). The goal of our vaccine and immunotherapeutic strategies is to gain a better understanding of the immune response of CD8+ T cells. In response to infection, naive CD8+ T cells are activated by specific Ag stimulation and then Ag-experienced CD8+ T cells undergo robust proliferation, giving rise to effector and immunological memory subsets (3, 4). The effector CD8+ T cells exhibit antiviral and antibacterial activities, including cytotoxic activity against infected cells; therefore, they are defined as the cells that produce functional molecules, such as IFN-γ and granzyme B (GzmB), which are important for protection against infection (5). To acquire these functions, CD8+ T cells need to differentiate from naive T cells into effectors. The fate of activated CD8+ T cells is determined by TCR signal strength, costimulation, transcription factors, inflammatory cytokines, and metabolic regulators (6, 7). In particular, a number of transcriptional activators and repressors were reported to be involved in the proliferation and differentiation of CD8+ T cells upon infection (4, 6, 8–12). There appears to be a delicate balance in the proliferation, differentiation, exhaustion, and survival regulated by transcription factors, indicating that a lack of balance could be harmful for the homeostasis of activated T cells. However, precisely which molecules contribute to the adequate immune response of Ag-experienced CD8+ T cells for the maximal function of effectors remains to be clearly elucidated.
Ag-specific effector CD8+ T cells highly express the lectin-like NKR KLRG1 in response to infection, whereas memory CD8+ T cells highly express cytokine IL-7Rα for long-term protection (4). CD8+ T cells with a high expression of KLRG1, referred to as short-lived effector cells, represent terminally differentiated effector cells with a lower survival than memory CD8+ T cells (13). It is known that the differentiation into short-lived effector cells is regulated by inflammatory conditions, followed by an increased expression of transcription factors, such as Blimp-1 and T-bet (14). In addition to the well-known regulatory function of these transcription factors, recent data indicate that transcriptional repressors, including Bcl-6, are involved in the regulation of differentiation in CD8+ T cells, suggesting that there is a network of transcriptional regulation rather than a single master regulator for cell fate determination (15). The identification of the molecules that play a role in the differentiation and maintenance of effector CD8+ T cells is important to improve immunological protection against infections; however, the detailed mechanism behind this process remains unclear.
We focused on a transcriptional repressor, menin, which is also known as a tumor suppressor (16–19). We recently reported that menin is a critical regulator of CD4+ T cell senescence and cytokine homeostasis (20). Despite the importance of menin for the maintenance of CD4+ T cells, its role in other lymphocytes remains to be elucidated. Thus, we hypothesized that menin is also involved in the regulation of differentiation and homeostasis in Ag-stimulated CD8+ T cells. The goal of this study was to investigate how menin is involved in the immune response of CD8+ T cells. In this article, we demonstrate that tumor-suppressor menin is critical for the primary immune response to infections using T cell–specific Menin-deleted mice. Menin-deficient CD8+ T cells showed a severe defect in proliferation and survival during expansion upon Listeria infection in a cell-intrinsic manner. Because Menin deficiency enhanced the expression of effector lineage-specific transcription factors, Blimp-1 and T-bet, menin could play a role in the differentiation checkpoint for cell fate determination and homeostasis of activated CD8+ T cells by controlling the expression of transcription factors. Taken together, our findings could provide important targets within this novel control pathway of the immune response to enhance the response to vaccination and immunotherapy.
Materials and Methods
Mice and cells
Meninflox/flox mice, CD4-Cre transgenic (Tg) mice, Rosa26-Cre-ERT2 Tg mice, C57BL/6 Thy1.1+ mice, and OT-1 Tg mice were purchased from The Jackson Laboratory. Meninflox/flox mice were crossed with CD4-Cre Tg mice to generate Meninflox/flox CD4-Cre (Menin-KO) mice. Next, we crossed wild-type (WT) or Menin-KO mice with OT-1 Tg mice to generate WT OT-1 Tg or Menin-KO OT-1 Tg mice. For tamoxifen-inducible gene deletion, Meninflox/flox OT-1 Tg mice were crossed with Rosa-Cre Tg mice to generate Meninflox/flox Rosa26-Cre-ERT2 OT-1 Tg (Rosa-Cre) mice. The mice were genotyped by PCR using genomic DNA isolated from tails. All mice were used at 6–12 wk of age, and both sexes were included in the experiments. All experiments using mice were performed with the approval of the Ehime University Administrative Panel for Animal Care. All animal care was conducted in accordance with the guidelines of Ehime University.
Cell suspensions were prepared by manual disruption of spleens and lymph nodes with frosted glass slides, followed by lysis of erythrocytes with an ammonium chloride/potassium solution. The livers and lungs were perfused with ice-cold PBS, as previously described (12). Both tissues were homogenized and incubated in PBS containing collagenase III (400 U/ml; Funakoshi, Tokyo, Japan) at 37°C for 30 min. Digested tissues were applied to a Percoll gradient (GE Healthcare Life Sciences) to collect the lymphocytes, according to the manufacturer’s protocol. The OVA-specific CD8+ T cells were detected using MHC class I pentamer H-2Kb SIINFEKL (#F093-0A-G) and FLUOROTAG R-PE label (#K2A; both from PROIMMUNE), according to the manufacturer’s protocol. The following Abs were used for cell surface staining and intracellular staining: anti-Thy1.1 FITC (HIS51; eBioscience), anti-Thy1.1 Alexa Fluor 647 (OX-7; BioLegend), anti-Thy1.2 allophycocyanin-Cy7 (30-H12; BioLegend), anti-Thy1.2 PE (53-2.1; BD Biosciences), anti-CD3 violetFluor 450 (17A2; TONBO Biosciences), anti-CD8 violetFluor 450 (2.43; TONBO Biosciences), anti-CD8 Alexa Fluor 488 (53-6.7; BioLegend), anti-CD4 eFluor 780 (RM4-5; eBioscience), anti-CD62L FITC (MEL-14; BD Biosciences), anti-CD62L allophycocyanin (MEL-14; TONBO Biosciences), anti-CD44 PE (IM7; TONBO Biosciences), anti-CD27 PE (LG.3A10; BD Biosciences), anti-CD127 Alexa Fluor 488 (SB/199; BioLegend), anti-CD25 PE (3C7; BD Biosciences), anti-CD69 PE (H1.2F3; BD Biosciences), anti-KLRG1 PE (2FI/KLRG1; BD Biosciences), anti–granzyme B Alexa Fluor 647 (GB11; BD Biosciences), anti–IFN-γ PE (XMG11.2; BD Biosciences), anti–PD-1 PE (J43; BD Biosciences), anti-2B4 PE (m2B4.B6.458.1; BioLegend), anti–LAG-3 PE (C9B7W; BioLegend), anti–CTLA-4 PE (UC10-4B9; BioLegend), anti-CD160 PE (7H1; BioLegend), anti–Tim-3 PE (8B.2C12; eBioscience), anti–Blimp-1 PE (C-21; Santa Cruz Biotechnology), anti–T-bet PE (4B10; BioLegend), anti-Eomes PE (Dan11mag; eBioscience), anti–p-Stat1 Alexa Fluor 647 (58D6; Cell Signaling Technology), anti–p-Stat4 Alexa Fluor 647 (38/p-Stat4; BD Biosciences), and anti–p-Stat5 Alexa Fluor 647 (47/Stat5-pY694; BD Biosciences). For IFN-γ and GzmB staining, intracellular staining was performed as previously described (21). Briefly, the splenocytes were isolated and stimulated with 1 μg/ml of H-2Kb OVA peptide SIINFEKL (MBL, Nagoya, Japan) in the presence of monensin (2 μM) in a 96-well culture plate for 6 h. Next, they were stained with anti-Thy1.1, anti-Thy1.1, and anti-CD8α and then fixed and permeabilized, followed by intracellular staining of anti–IFN-γ or GzmB. For intracellular staining of Blimp-1, T-bet, Eomes, p-Stat1, p-Stat4, and p-Stat5, the cell surface was stained as described above and then fixed and permeabilized using a Transcription Factor Staining Buffer Kit (TONBO Biosciences), followed by intracellular staining. For analysis of apoptosis, cells were stained with anti-CD8α, anti-Thy1.1, and anti-Thy1.2 and then incubated with 7-aminoactinomycin D and Annexin V–PE using an Annexin V Apoptosis Detection Kit (#556422; BD Biosciences), according to the manufacturer’s protocol. Flow cytometry was performed using a Gallios instrument (Beckman Coulter), and data were analyzed with FlowJo software (TreeStar).
Adoptive transfer of CD8+ T cells and Listeria infection
Naive CD44loCD8+ T cells were purified from the spleens of WT OT-1 Tg (Thy1.1+ or Thy1.2+) and Menin-KO OT-1 Tg (Thy1.2+) mice using a Naive CD8a+ T Cell Isolation Kit and an autoMACS Pro Separator (Miltenyi Biotec), and their purities were checked by flow cytometry (>95% purity). The purified cells were mixed at a 1:1 ratio (WT/Menin KO) and adoptively transferred into double-congenic (Thy1.1+Thy1.2+) mice (1 × 104 cells per mouse, i.v.). Next, the mice were infected with Listeria monocytogenes expressing OVA (Lm-OVA) at 5 × 103 CFU i.v. 18–24 h later. For the analysis or purification of donor cells, the splenocytes isolated from recipient mice were stained with anti-CD3, anti-CD8, anti-CD4, anti-Thy1.1, and anti-Thy1.2 Abs. The donor cells were then analyzed or purified using a Gallios instrument or a BD FACSAria II cell sorter (BD Biosciences) by gating on CD3+CD4−CD8+Thy1.1+Thy1.2− (WT) cells or CD3+CD4−CD8+Thy1.1−Thy1.2+ (Menin-KO) cells. All experiments using Lm-OVA were performed according to the protocols approved by the Ehime University Institutional Biosafety Committee.
Measurement of bacterial burden
WT and Menin-KO mice were infected with Lm-OVA (4 × 105 CFU, i.v.). The bacterial burden in the spleen, liver, and lung was determined on day 7 after Lm-OVA infection in culture with BHI bacterial plates, as previously described (22).
Induction of gene deletion with tamoxifen
Naive CD8+ T cells from Rosa-Cre OT-1 Tg (Thy1.2+) mice were purified and mixed with naive OT-1 CD8+ T cells at a 1:2 ratio (WT/Rosa-Cre) and then adoptively transferred into double-congenic (Thy1.1+Thy1.2+) mice (1 × 104 cells/mouse, i.v.). Mice were infected with Lm-OVA (5 × 103 CFU, i.v.) 18–24 h later. For the induction of gene deletion, we used tamoxifen (#T5648-1G) dissolved in corn oil (#23-0320-5; both from Sigma-Aldrich). Tamoxifen was administered to the mice (3 mg per mouse, i.p.) on day 4 after Lm-OVA infection.
In vivo proliferation assay by eFluor 670 staining and BrdU incorporation
For eFluor 670 staining, OT-1 CD8+ T cells were incubated with 5 μM Cell Proliferation Dye eFluor 670 (#65-0840; eBioscience), according to the manufacturer’s protocol. eFluor 670–labeled cells were adoptively transferred into recipient mice, which were infected with Lm-OVA (5 × 103 CFU, i.v.). At different time points postinfection, splenocytes from the recipients were stained with anti-CD8, anti-Thy1.1, and anti-Thy1.2, and the dilution of eFluor 670 was measured by flow cytometry to evaluate cell proliferation. The proliferation index (PI) was calculated using FlowJo software to compare WT and Menin-KO CD8+ T cells. For BrdU incorporation, BrdU (Sigma-Aldrich) was injected into mice (2 mg per mouse, i.p.) on day 5 after Lm-OVA infection, and the spleens were harvested 14 h later. Splenocytes were stained with anti-CD8α violetFluor 450, anti-Thy1.1 Alexa Fluor 647, and anti-Thy1.2 allophycocyanin-Cy7, and BrdU incorporation was measured by flow cytometry with an FITC BrdU flow kit (#559619; BD Biosciences), according to the manufacturer’s protocol.
In vitro proliferation assay by stimulation with OVA-altered peptide ligands
OT-1 CD8+ T cells were labeled with eFluor 670, as described above. Splenocytes were pulsed with OVA-altered peptide Q4 (SIIQFEKL) (#AS-64402; AnaSpec) at 37°C for 60 min. Then, OT-1 CD8+ T cells were cocultured with peptide-pulsed splenocytes. After 3 d of stimulation, the dilution of eFluor 670 in OT-1 CD8+ T cells was analyzed by flow cytometry.
RNA isolation and quantitative RT-PCR
Total RNA was isolated using TRIzol reagent (Life Technologies) or NucleoSpin RNA XS (Takara Bio), according to the manufacturers’ protocols. Then, cDNA was synthesized using a SuperScript VILO cDNA Synthesis Kit (Life Technologies). Quantitative PCR was performed using the StepOnePlus Real-Time PCR System (Life Technologies). The levels of gene expression were normalized to that of CD3e. The following specific primers and Roche Universal Probes were used: Hprt: 5′-TCCTCCTCAGACCGCTTTT-3′ (forward), 5′-CCTGGTTCATCATCGCTAATC-3′ (reverse), probe #95; Menin: 5′-ACCCACTCACCCTTTATCACA-3′ (forward), 5′-ACATTTCGGTTGCGACAGT-3′ (reverse), probe #20; Bach2: 5′-CAGTGAGTCGTGTCCTGTGC-3′ (forward), 5′-TTCCTGGGAAGGTCTGTGAT-3′ (reverse), probe #79; Pmaip1 (Noxa): 5′-CAGATGCCTGGGAAGTCG-3′ (forward), 5′-TGAGCACACTCGTCCTTCAA-3′ (reverse), probe #15; Perp: 5′-GACCCCAGATGCTTGTTTTC-3′ (forward), 5′-ACCAGGGAGATGATCTGGAA-3′ (reverse), probe #71; Cdkn1a (p21): 5′-TCCACAGCGATATCCAGACA-3′ (forward), 5′-GGACATCACCAGGATTGGAC-3′ (reverse), probe #21; Cdkn2d (p19): 5′-GGGTTTTCTTGGTGAAGTTCG-3′ (forward), 5′-TTGCCCATCATCATCACCT-3′ (reverse), probe #106; Cdkn2a (p16): 5′-AATCTCCGCGAGGAAAGC-3′ (forward), 5′-GTCTGCAGCGGACTCCAT-3′ (reverse), probe #91; Cdkn2b (p15): 5′-GGCTGGATGTGTGTGACG-3′ (forward), 5′-GCAGATACCTCGCAATGTCA-3′ (reverse), probe #41; Cdkn2a (p16): 5′-AATCTCCGCGAGGAAAGC-3′ (forward), 5′-GTCTGCAGCGGACTCCAT-3′ (reverse), probe #91; Bim: 5′-GGAGACGAGTTCAACGAAACTT-3′ (forward), 5′-AACAGTTGTAAGATAACCATTTGAGG-3′ (reverse), probe #41; Puma: 5′-TTCTCCGGAGTGTTCATGC-3′ (forward), 5′-TACAGCGGAGGGCATCAG-3′ (reverse), probe #79; Bcl-2: 5′-GTACCTGAACCGGCATCTG-3′ (forward), 5′-GGGGCCATATAGTTCCACAA-3′ (reverse), probe #75; Bcl-xL: 5′-TGACCACCTAGAGCCTTGGA-3′ (forward), 5′-GCTGCATTGTTCCCGTAGA-3′ (reverse), probe #2; Mcl-1: 5′-GGTATTTAAGCTAGGGTCATTTGAA-3′ (forward), 5′-TGCAGCCCTGACTAAAGGTC-3′ (reverse), probe #41; Prdm1: 5′-TGCGGAGAGAGGCTCCACTA-3′ (forward), 5′-TGGGTTGCTTTCCGTTTG-3′ (reverse), probe #80; Tbx21: 5′-AAACATCCTGTAATGGCTTGTG-3′ (forward), 5′-TCAACCAGCACCAGACAGAG-3′ (reverse), probe #19; Bcl6: 5′-CTGCAGATGGAGCATGTTGT-3′ (forward), 5′-GCCATTTCTGCTTCACTCG-3′ (reverse), probe #4; Id3: 5′-GAGGAGCTTTTGCCACTGAC-3′ (forward), 5′-GCTCATCCATGCCCTCAG-3′ (reverse), probe #19; Bax: 5′-GTGAGCGGCTGCTTGTCT-3′ (forward), 5′-GGTCCCGAAGTAGGAGAGGA-3′ (reverse), probe #83; Cdkn6 (p18): 5′-AAATGGATTTGGGAGAACTGC-3′ (forward), 5′-AAATTGGGATTAGCACCTCTGA-3′ (reverse), probe #79; Cdkn1b (p27): 5′-GAGCAGTGTCCAGGGATGAG-3′ (forward), 5′-TCTGTTCTGTTGGCCCTTTT-3′ (reverse), probe #62; CD3ε: 5′-CCAGCCTCAAATAAAAACACG-3′ (forward), 5′-GATGATTATGGCTACTGCTGTCA-3′ (reverse), probe #10; and β-actin: 5′-CTAAGGCCAACCGTGAAAAG-3′ (forward), 5′-ACCAGAGGCATACAGGGACA-3′ (reverse), probe #64. Specific reagents for Tcf7 were purchased from Applied Biosystems (catalog no. Mm00493445_m1). Gene expressions were calculated as the relative expression, normalized to the expression of HPRT or CD3ε.
Menin deficiency impairs the immune response of Ag-specific CD8+ T cells
T cell–specific Menin-deleted mice were generated by crossing Menin-floxed mice with CD4-promoter derived Cre-recombinase Tg mice. A markedly reduced expression of menin was observed in peripheral CD8+ T cells from Meninflox/flox CD4-Cre Tg (Menin-KO) mice by quantitative RT-PCR (qRT-PCR) (Supplemental Fig. 1A). Before the infection study, an immunophenotypic analysis of CD8+ T cells was performed in the steady-state. There was no significant difference in the CD44hiCD62Lhi population among CD8+ T cells in the thymus between WT and Menin-KO mice (Supplemental Fig. 1B). However, Menin-KO mice had a larger CD44hiCD62Lhi population among CD8+ T cells in the spleen, blood, inguinal lymph node (iLN), mesenteric LN (mLN), liver, and lung (Supplemental Fig. 1C) but a lower total number of CD8+ T cells than WT mice (Supplemental Fig. 1D). Naive CD8+ T cells from WT and Menin-KO mice did not express the activation markers CD69 and CD25, nor did they express GzmB in the steady-state without stimulation (Supplemental Fig. 2A). Intriguingly, naive Menin-KO OT-1 CD8+ cells showed enhanced proliferation upon stimulation with low-affinity OVA peptides and a more rapid increase in activation markers CD69 and CD25 than WT cells, suggesting a reduced activation threshold in naive Menin-KO cells (Supplemental Fig. 2B–F). Next, we infected the mice with the intracellular pathogen Lm-OVA to determine the impact of Menin deficiency on the immune response of CD8+ T cells, as previously described (23). We investigated the primary immune response of endogenous CD8+ T cells to Lm-OVA infection by flow cytometry using a pentamer for the detection of OVA-specific CD8+ T cells. Although there were no significant differences between WT and Menin-KO mice on days 3 and 5 after Lm-OVA infection, Menin-KO CD8+ T cells showed a much lower frequency of OVA-specific pentamer-positive (Pent+) cells in the spleen on day 7, at the peak of the immune response of WT cells (Fig. 1A). The kinetics showed a significantly lower frequency and absolute number of Pent+ cells in Menin-KO cells after day 7 of infection (Fig. 1B). Other tissues, including the liver, lung, iLN, and mLN, also exhibited a significantly lower percentage and absolute number of Pent+ cells among Menin-KO CD8+ T cells compared with WT, indicating that the lower number of Ag-specific CD8+ T cells in the spleen was not due to a difference in tissue distribution (Fig. 1C). Correlating with these results, Menin-KO mice showed a significantly lower clearance of Lm-OVA in the spleen, liver, and lung than did WT mice (Fig. 1D). In contrast, heterozygous Menin-deleted (Meninflox/+ CD4-Cre Tg) mice showed an expansion in the spleen that was comparable to WT mice (Supplemental Fig. 3A).
Immune response of Menin-KO CD8+ T cells is impaired in a cell-intrinsic manner
In the T cell–specific gene-deletion system using CD4-Cre Tg mice, menin is absent in CD4+ T cells, as well as CD8+ T cells. This raises the possibility that the immune response of Menin-deleted CD8+ T cells was impaired as a result of functional defects in Menin-deleted CD4+ T cells (i.e., in a cell-extrinsic manner). To exclude this possibility, we used an adoptive-transfer approach by transferring donor cells from OT-1 Tg mice into congenic recipient mice. We confirmed the presence of >90% Pent+ cells in naive CD8+ T cells from WT OT-1 Tg and Menin-KO OT-1 Tg mice by flow cytometry (data not shown). We purified naive CD44loCD8+ cells from the spleens of WT OT-1 Tg (Thy1.1+) and Menin-KO OT-1 Tg (Thy1.2+) mice and then adoptively transferred a 1:1 mixture of WT/Menin-KO OT-1 Tg cells into double-congenic (Thy1.1+Thy1.2+) recipient mice to distinguish cells by flow cytometry using cell surface staining [Fig. 2A (1)]. Consistent with the results of the endogenous immune response, donor Menin-KO cells were outcompeted by WT cells in the spleen, liver, and lung on day 7 after Lm-OVA infection (Fig. 2B). It seems clear that menin plays a critical role in the immune response of Ag-specific CD8+ T cells to infection.
Next, we induced gene deletion after Lm-OVA infection using a drug, tamoxifen. For tamoxifen-induced Menin deletion, we crossed Meninflox/flox OT-1 Tg mice with Rosa26-promoter-derived Cre-ERT2 recombinase Tg mice to generate Rosa-Cre mice to analyze the role of menin during expansion following infection. Naive CD44loCD8+ T cells were purified from WT OT-1 Tg (Thy1.1+) mice and Rosa-Cre OT-1 Tg (Thy1.2+) mice and mixed at a 1:2 ratio (WT/Rosa-Cre) to compare the expansion of Ag-specific activated CD8+ T cells in the same recipient host. Recipient mice were infected with Lm-OVA (5 × 103 CFU, i.v.) the day after adoptive transfer, and they were administered tamoxifen (3 mg per mouse) or vehicle control (corn oil) by i.p. injection on day 4 postinfection [Fig. 2A (2)]. A high rate of gene deletion was observed in donor CD8+ T cells from Rosa-Cre OT-1 Tg mice by genomic PCR (Supplemental Fig. 3B). The results showed a much lower percentage and absolute number of tamoxifen-treated Rosa-Cre OT-1 CD8+ T cells compared with WT or vehicle-treated Rosa-Cre OT-1 CD8+ T cells in the spleen, liver, and lung (Fig. 2C). However, donor Menin-KO OT-1 CD8+ T cells showed a comparable proliferation to WT cells at the early time points (days 3 and 4) postinfection, with a slightly higher PI in Menin-KO cells on day 3 (Fig. 2D). These observations suggest that a key change related to an impaired expansion occurs in Menin-KO cells after day 4 of infection.
Menin enhances proliferation and inhibits apoptosis in activated CD8+ T cells
We compared proliferation in Ag-specific CD8+ T cells on day 5 postinfection by BrdU incorporation. BrdU was injected i.p. 14 h before spleen harvest and was detected by flow cytometry with anti-BrdU. Menin-KO CD8+ T cells exhibited a significantly lower percentage of BrdU+ cells than did WT cells (Fig. 3A). Next, we isolated donor CD8+ T cells from recipient mice on day 5 postinfection by cell sorting and isolated total RNA to compare the gene expression of cell cycle inhibitors in WT and Menin-KO cells. The results revealed that Menin-KO cells proliferated less, with a greater expression of cell cycle inhibitor genes (p15, p16, p19, and p27) than in WT cells on day 5 postinfection (Fig. 3B).
Next, we compared apoptosis in Ag-specific CD8+ T cells from WT and Menin-KO mice by flow cytometry, with Annexin V staining on day 5 after Lm-OVA infection. Menin-KO CD8+ T cells exhibited a significantly higher percentage of AnnexinV+ cells than did the WT cells (Fig. 3C). We then measured the expression of proapoptotic genes (Fig. 3D, upper panels) and antiapoptotic genes (Fig. 3D, lower panels) in donor CD8+ T cells on day 5 postinfection by qRT-PCR (Fig. 3B). qRT-PCR analysis showed higher expression of proapoptotic genes (Bim and Puma) and lower expression of an antiapoptotic gene (Bcl-xL) in Menin-KO cells than in WT cells. To identify the factors involved in these defects in Menin-KO cells, we analyzed cytokine responsiveness by measuring phosphorylated Stats upon cytokine stimulation. The response to cytokines IL-2, IL-12, and IFN-α was normal in Menin-KO cells (Supplemental Fig. 4).
Menin deficiency enhances differentiation into terminal effectors in activated CD8+ T cells during expansion
Next, we focused on the analysis of differentiation from naive T cells into effectors and memory precursor cells by comparing immunophenotypes using flow cytometry. We examined phenotypic markers of Ag-specific activated CD8+ T cells in the spleen on days 3–30 after Lm-OVA infection to compare differentiation during primary expansion and memory development. Activated CD8+ T cells can be divided into short-lived effector and memory precursor populations based on the expression of KLRG1 and CD127 (IL-7Rα) (13). The time-course study revealed enhanced differentiation of Menin-KO CD8+ T cells into short-lived effectors during expansion compared with WT cells before the peak of expansion on day 7 (Fig. 4A). Menin-KO CD8+ T cells exhibited a significantly higher percentage of CD127loKLRG1hi cells than did WT cells on day 5 (Fig. 4C, left panel). WT CD8+ T cells exhibited a gradual increase in the percentage of CD127hiKLRG1lo memory precursors after day 7. In contrast, Menin-KO CD8+ T cells did not show an increase in memory precursors and maintained a high percentage of CD127loKLRG1hi terminal effectors. Surprisingly, we detected Menin-KO CD8+ T cells only rarely as a result of the large decrease in cell number after 14 d (Supplemental Fig. 3C).
We also examined the immunophenotype of activated CD8+ T cells by flow cytometry using anti-CD62L and anti-CD27 Abs. In agreement with the findings from CD127/KLRG1 analysis, Menin-KO CD8+ T cells showed enhanced differentiation into CD62LloCD27lo terminal effectors and maintained a high percentage of terminal effectors (Fig. 4B). Menin-KO CD8+ T cells exhibited a significantly higher percentage of terminal effectors compared with WT cells on day 5 (Fig. 4C, right panel). Finally, we did not detect any memory Menin-KO CD8+ T cells on day 60 in a cotransfer setting, suggesting that there was a cell-intrinsic defect in memory development (Fig. 4D). Next, we used flow cytometry to analyze effector cells from WT and Menin-KO mice to assess their production of functional molecules GzmB and IFN-γ by comparing mean fluorescent intensity (MFI) on day 5 postinfection. Menin-KO CD8+ T cells showed greater production of GzmB and IFN-γ compared with WT cells (Fig. 4E). Correlating with an enhanced terminal differentiation, Menin-KO CD8+ T cells also showed greater expression of inhibitory receptors, including PD-1 and 2B4, in a cotransfer setting on day 5 postinfection (Fig. 4F).
Menin negatively regulates the expression of effector-promoting transcription factors and positively regulates the expression of memory-promoting transcription factors
We used intracellular staining to compare the expression of the transcription factors related to differentiation into terminal effectors and memory precursors in activated CD8+ T cells after Lm-OVA infection. The time-course study revealed rapid enhancement of Blimp-1 expression related to terminal effector differentiation in Menin-KO CD8+ T cells (Fig. 5A). Menin-KO CD8+ T cells also showed significantly higher expression of Blimp-1 and T-bet compared with WT cells on day 5 postinfection, whereas there was no significant difference in Eomes expression in the Ag-specific activated CD8+ T cells between Menin-KO and WT cells, as determined by MFI and gene expression (Fig. 5B). Intriguingly, higher expression of Blimp-1 was sustained in Menin-KO cells beyond day 7, in contrast to WT cells. However, the expression of Bcl6 and Id3 was significantly lower in Menin-KO cells at time points later than day 7, and the expression of Tcf7 was reduced significantly in Menin-KO cells on day 12 compared with WT cells, although there was no significant difference on day 5 postinfection (Fig. 5C). These results indicate that menin inhibits terminal effector differentiation and enhances memory development by controlling the expression of transcription factors important for differentiation into effector cells and memory cells.
The understanding of differentiation and homeostasis in activated CD8+ T cells upon infection is required for the development of vaccination and immunotherapeutic protocols. Transcription factors are involved in cell fate determination of Ag-experienced CD8+ T cells (4). In this article, we demonstrated that a tumor suppressor, menin, is also involved in the regulation of differentiation, proliferation, and survival in activated CD8+ T cells upon infection. In the steady-state without infection, Menin-KO CD8+ T cells exhibited a larger percentage of CD44hiCD62Lhi cells in peripheral tissues, but not in the thymus, compared with WT cells. Interestingly, we observed the altered peptide ligands induced more proliferation in Menin-KO CD8+ T cells than in WT cells, which suggests that Menin deficiency leads to a reduced activation threshold upon TCR stimulation. Akt inhibition by menin could be involved in this reduced activation threshold in Menin-KO cells because menin interacts with Akt and suppresses its kinase activity, which, in turn, is important for T cell activation (24). Thus, Menin-KO naive CD8+ T cells could be more easily activated by encountering unknown low-affinity Ags in the steady-state and may increase the CD44hi population. For the initial investigation into the impact of menin in the immune response, we infected mice and compared the response of CD8+ T cells from WT and Menin-KO mice. Notably, T cell–specific Menin deficiency resulted in the dramatic decrease in Ag-specific CD8+ T cells on day 7 after Lm-OVA infection compared with the WT cells. In contrast, heterozygous Menin-deleted mice showed expansion similar to the WT mice after Lm-OVA infection, indicating that Menin is haplosufficient in the immune response of CD8+ T cells. In agreement with the reduced clonal expansion in Menin-KO CD8+ T cells, Menin-KO mice showed less effective clearance of intracellular pathogens compared with WT mice, which could be due to the reduced number of Ag-specific CD8+ T cells. Moreover, adoptive-transfer experiments using OT-1 Tg mice showed a lower number of Ag-specific Menin-KO CD8+ T cells than WT cells on day 7 after Lm-OVA infection, and Menin-KO CD8+ T cells were rarely detected later than 14 d postinfection. Intriguingly, tamoxifen treatment for Menin deletion postinfection reduced the cell number on day 7, suggesting that menin is required in the expansion phase postinfection to obtain a proper immune response.
With regard to proliferation, Menin-KO CD8+ T cells had a slightly higher PI compared with WT cells on day 3 postinfection, which is likely due to a reduced activation threshold. However, the BrdU-incorporation results revealed less proliferation in Menin-KO CD8+ T cells on day 5 compared with WT cells, which might be related to the increased apoptosis of Menin-KO CD8+ T cells. These proliferation differences in Menin-KO cells cannot be attributed to cytokine IL-2, IL-12, or type-I IFN responsiveness because there were no significant differences in cytokine signaling between WT and Menin-KO cells. There could be an abnormal epigenetic change in cell cycle–related genes in Menin-KO CD8+ T cells between days 3 and 5 because menin plays a role in histone modification to epigenetically control the expression of genes involved in cell proliferation (25). Thus, Menin deficiency may contribute to cell cycle arrest during expansion by enhancing the expression of cell cycle inhibitor genes (p15, p16, p19, and p27). Among these cell cycle inhibitors, the gene expression of p15 (Ink4b) and p16 (Ink4a) is known to be regulated by the Polycomb group (PcG) (26), which indicates that cell cycle arrest could be regulated by epigenetic modifications in CD8+ T cells. Trithorax group gene products can antagonize the effect of PcG gene products (27). Thus, menin could regulate the gene expression of cell cycle inhibitors in an epigenetic fashion because it forms a Trithorax complex with mixed-lineage leukemia 1, a histone H3K4 methyltransferase (28, 29). Moreover, p16 is associated with cellular senescence in CD8+ T cells from elderly persons (30). This may be related to senescence as observed in activated Menin-KO CD4+ T cells from in vitro culture by β-galactosidase staining (20), although the detailed mechanism of cell cycle arrest regulated by menin remains to be explored. p16 also increases apoptosis in T lymphocytes (31), which suggests that there is a positive correlation between cell cycle arrest and apoptosis. In addition to proliferation, the survival of activated Ag-specific CD8+ T cells has to be maintained during expansion to eliminate the pathogen. Analysis using Annexin V demonstrated enhanced apoptosis in Menin-KO CD8+ T cells, indicating a reduced survival in the absence of menin that could be due to the increased expression of proapoptotic genes (Bim and Puma) and the decreased expression of an antiapoptotic gene (Bcl-xL). Proapoptotic genes are known to be regulated by the PcG gene Bmi1 in CD4+ T cells (32, 33), which suggests that apoptosis could be induced by epigenetic modifications in CD8+ T cells by menin as well. Thus, apoptosis could be connected with reduced proliferation via epigenetic changes regulated by menin. Finally, the combination of lower proliferation and impaired survival dramatically decreased the number of Menin-KO cells on day 7 postinfection.
Based on immunophenotypic analysis, Menin-KO CD8+ T cells showed enhanced differentiation into CD127loKLRG1hi and CD62LloCD27lo terminal effectors with higher expression of GzmB and IFN-γ and diminished the CD127hiKLRG1lo long-lived memory precursor subset in the expansion phase. Therefore, our activated Menin-KO CD8+ T cells do not appear to retain sufficient plasticity to give rise to other subsets, such as effector memory T cells, and may be primarily destined for death or senescence as with exhausted CD8+ T cells during chronic infection (34, 35). Terminal effector differentiation is known to be enhanced by inflammatory signals produced upon infection (14, 36, 37). However, with regard to the reduced cell number in Menin-KO cells, extrinsic environmental influences can be excluded because adoptive-transfer experiments also resulted in an impaired immune response of Menin-KO CD8+ T cells. Therefore, an important finding was the fact that menin plays a role in the differentiation and homeostasis of activated CD8+ T cells during expansion, which is due to its cell-intrinsic effect. In the current study, we linked the role of transcription factors to the differentiation of terminal effectors. Menin deficiency increased the expression of Blimp-1 and T-bet, which are important transcription factors for differentiation into terminal effector CD8+ T cells (38–41). Menin could negatively regulate the expression of those factors as a transcriptional repressor of AP-1 by interacting with JunD (16, 28, 29, 42–44). In contrast, there was no significant difference in the expression of the transcription factor Eomes, which is important for memory differentiation (45, 46), suggesting that the disrupted balance in Menin-KO CD8+ T cells predominantly leads to terminal effectors. Concomitantly with enhanced terminal effector differentiation, Menin-KO CD8+ T cells exhibited higher expression of inhibitory receptors (PD-1 and 2B4) with reduced proliferation compared with WT cells, as observed in exhausted CD8+ T cells during chronic infection (47). The high expression of Blimp-1 is known to play a role in the exhaustion of activated CD8+ T cells during chronic infection (48, 49). In addition, increased T-bet expression was reported to be associated with senescence of virus-specific CD8+ T cells (50). Thus, our results may imply that the higher expression of Blimp-1 and T-bet is likely to increase the expression of inhibitory receptors and p19 (Arf), a senescence-associated gene (51), which could contribute to the reduced number of Menin-KO cells compared with WT cells during expansion upon infection. As is the case with Menin-KO CD4+ T cells, menin might be targeting Bach2, which is known to regulate immune homeostasis (20). Bach2 is also known to promote memory CD8+ T cell development (52). Importantly, Bach2 binds to the Prdm1 gene locus and represses its transcription. Indeed, Menin-KO CD8+ T cells showed reduced expression of Bach2 (data not shown), suggesting that it could be involved in the regulation of the immune system downstream of menin. Thus, overexpression of Bach2 potentially rescues the impaired immune response in Menin-KO CD8+ T cells.
A key finding of our study is that menin is required to obtain a proper immune response during clonal expansion. We identified that menin plays an important role in the immune response of CD8+ T cells to infection, although it may also have many unexplored effects on immune responses beyond the regulation of differentiation and homeostasis. As a result, these findings could be used for the optimization of immunotherapies for cancer, as well as for vaccination against infectious pathogens.
We thank Hao Shen and Shiki Takamura for providing Lm-OVA, Aya Tamai for valuable technical support, Kenji Kameda for cell sorting assistance, and Nicholas P. Casey for critically reading the manuscript.
This work was supported by JSPS KAKENHI Grant 26460579, the Takeda Science Foundation, BioLegend/TOMY Digital Biology, the Naito Foundation, and the Waksman Foundation.
The online version of this article contains supplemental material.
Abbreviations used in this article:
The authors have no financial conflicts of interest.