FTO controls CD8+ T cell survival and effector response by modulating m6A methylation of Fas

Sun, Lina; Zhang, Tianzhe; Ge, Yao; Yao, Zhihong; Su, Yanhong; Wang, Qianhao; Chen, Yang; He, Boxiao; Ding, Renyi; Zhang, Cangang; Lan, Linbo; Liu, Ruonan; Ping, Huanxin; Zhang, Dan; Shi, Lin; Zhou, Xiaobo; Jia, Xiaoxuan; Sun, Chenming; Liang, Lingli; Zhang, Lianjun; Zhang, Baojun

doi:10.1038/s41419-025-07606-z

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Published: 15 April 2025

FTO controls CD8⁺ T cell survival and effector response by modulating m⁶A methylation of Fas

Lina Sun ORCID: orcid.org/0009-0008-5650-7480^1,2,3,4^na1,
Tianzhe Zhang^1,2,3,4^na1,
Yao Ge⁵^na1,
Zhihong Yao^1,6^na1,
Yanhong Su^1,2,3,4^na1,
Qianhao Wang^1,2,3,4,
Yang Chen⁵,
Boxiao He^1,2,
Renyi Ding^1,2,
Cangang Zhang ORCID: orcid.org/0000-0001-6443-7023^1,2,
Linbo Lan⁷,
Ruonan Liu^1,2,
Huanxin Ping^1,2,
Dan Zhang^1,2,
Lin Shi^1,2,4,
Xiaobo Zhou^1,2,4,
Xiaoxuan Jia²,
Chenming Sun ORCID: orcid.org/0000-0002-6199-7225^1,2,3,4,
Lingli Liang ORCID: orcid.org/0000-0001-7594-387X⁸,
Lianjun Zhang ORCID: orcid.org/0000-0001-9448-7718^9,10 &
…
Baojun Zhang ORCID: orcid.org/0000-0001-5972-1011^1,2,3,4

Cell Death & Disease volume 16, Article number: 301 (2025) Cite this article

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Abstract

Functional CD8⁺ T cell immunity is essential for immune surveillance and host defense against infection and tumors. Epigenetic mechanisms, particularly RNA modification, in controlling CD8⁺ T cell immune response is not fully elucidated. Here, by T cell-specific deletion of fat mass and obesity-associated protein (FTO), a critical N6-methyladenosine (m⁶A) demethylase, we revealed that FTO was indispensable for adequate CD8⁺ T cell immune response and protective function. FTO ablation led to considerable cell death in activated CD8⁺ T cells, which was attributed to cell apoptosis. MeRIP-seq analysis revealed an increase in m⁶A methylation on Fas mRNA in FTO-deficient CD8⁺ T cells. The loss of FTO promoted Fas expression via enhancing the Fas mRNA stability, which depended on the m⁶A reader insulin-like growth factor-2 mRNA-biding proteins 3 (IGF2BP3). Mutation of the Fas m⁶A sites or knockdown IGF2BP3 could normalize the upregulated Fas expression and apoptosis levels caused by FTO ablation in CD8⁺ T cells. Our findings delineate a novel epigenetic regulatory mechanism of FTO-mediated m⁶A modification in supporting CD8⁺ T cell survival and effector responses, providing new insights into understanding the post-transcriptional regulation in CD8⁺ T cell immunological functions and the potential therapeutic intervention.

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Introduction

CD8⁺ T cells are pivotal cell populations in adaptive immunity for protecting the host against pathogenic infections and eliminating malignancy [1]. Upon recognizing the antigen, naïve CD8⁺ T cells undergo rapid activation, expansion, and differentiation into effector cells, which carry out cytotoxic function and memory cells afterwards, which provide long-term protection [2]. In response to acute infections, antigen-specific CD8⁺ T cells experience distinct stages: expansion, contraction (with massive apoptosis), and memory differentiation [3]. The CD8⁺ T cell immune responses are orchestrated by complex signal cascades, including T cell receptor (TCR), co-stimulation, cytokines, and transcription factors (TFs) [2]. In addition, epigenetic regulations, such as DNA methylation and histone modifications, have gained much attention in modulating CD8⁺ T cell differentiation and function by altering the chromatin accessibility of the regulatory regions within key genes in CD8⁺ T cells [4, 5]. Besides DNA and protein, RNA-based epigenetic regulations, including small RNAs, long non-coding RNAs (lncRNAs), and RNA modifications, have recently emerged as crucial regulators of gene expression, silencing, and chromatin structure, involving various biological processes [6,7,8]. The regulatory mechanisms mediated by small RNAs and lncRNAs have been reported to govern CD8⁺ T cell effector functions [9]. However, the contributions of post-transcriptional RNA modifications to CD8⁺ T cell immune response remain largely unknown. m⁶A is the most abundant and prevalent RNA (particularly mRNA) modification in eukaryotic cells. m⁶A modification is dynamically and reversibly modulated by methyltransferases (writers) consisting of methyltransferase 3 (METTL3), METTL14, Wilms Tumor Associated Protein (WTAP), vir-like m⁶A methyltransferase associated (VIRMA) etc., and demethylases (erasers) including two members, alpha-ketoglutarate-dependent dioxygenase ABH5 (ALKBH5) and FTO [10]. m⁶A modifications affect mRNAs' fate and downstream functions, such as decay, splicing, subcellular translocation, and translation, which are determined by m⁶A binding proteins (readers) [11]. To date, three distinct groups of proteins have been identified as m⁶A readers: YT521-B homology (YTH)-RNA binding ___domain family, including YTH ___domain family 1–3 (YTHDF1-3) and YTH ___domain containing 1–2 (YTHDC1-2); the heterogeneous nuclear ribonucleoprotein (HNRNP) family members (e.g. HNRNPC, HNRNPG, and HNRNPA2B1); and insulin-like growth factor-2 mRNA-biding proteins 1–3 (IGF2BP1-3) [12]. Among those readers, YTHDF2 has been shown to mediate mRNA decay, and YTHDF1, YTHDF3, and YTHDC2 promote translation, while IGF2BPs mainly enhance mRNA stability [13]. Previous studies have demonstrated that m⁶A modifications exert pivotal regulatory effects on T cell homeostasis and functionality [14]. For instance, m⁶A writer METTL3 was required for T cells homeostasis by regulating the IL-7/STAT5/SOCS pathway [15]. In addition, m⁶A modifications directly modulated T cell differentiation as METTL3 ablation in T cells exhibited disrupted T helper (Th)1 [15], Th17 [15], regulatory T (Treg) [16] and T follicular helper T (Tfh) [17] cell differentiation, as well as impaired CD8⁺ T cell effector differentiation and memory formation [18]. Moreover, m⁶A modifications can influence T cell immune response indirectly by affecting antigen-presenting cells (APCs), such as dendritic cells (DCs) [19] and macrophages [20, 21].

FTO is the first identified m⁶A demethylase and primarily functions to remove m⁶A methylation through its oxidative demethylase activity, playing critical roles in a variety of pathological conditions, including obesity, diabetes, heart failure, neurological disorders, and tumorigenesis [22,23,24]. However, the roles of FTO in modulating immune cell functions have not been well understood, with limited studies focusing on innate immune cells. FTO promotes polarization of both M1 and M2 macrophages by sustaining NF-κB signaling pathway and the mRNA stability of STAT1 and PPAR-γ [25]. However, under diabetic conditions, FTO inhibits M1 macrophages activation and inflammation via m⁶A-regulated distinct signaling pathways [26, 27]. Additionally, FTO negatively regulates the cytotoxic activity of natural killer (NK) cells by increasing the mRNA stability of suppressor of cytokine signaling protein (SOCS) family genes [28]. While the direct regulatory roles of FTO in T cells remain unclear, indirect effects have been reported in the context of tumors, where FTO influences T cell function by modulating tumor cells [29, 30]. For example, FTO supports cancer (stem) cell growth and maintenance and the expression of immune checkpoint molecule PD-L1, thereby enhancing immune evasion [31,32,33]. Recently, FTO-mediated m⁶A demethylation promotes glycolytic activity in tumor cells, which impedes CD8⁺ T cell infiltration and effector function [24, 34]. Here, by T cell-specific deletion of FTO, we unveiled that FTO was crucial for supporting CD8⁺ T cell immune response and protective function during pathogenic infection. Ablation of FTO resulted in a pronounced increase in CD8⁺ T cell apoptosis upon activation, which was attributed to the upregulated expression of Fas receptor. Mechanistically, FTO deficiency caused an elevated level of m⁶A methylation on Fas mRNA, which enhanced the mRNA stability and, thereby, Fas expression in a manner dependent on the m⁶A reader protein IGF2BP3. Our findings define a crucial role of RNA epigenetic regulator via FTO-mediated m⁶A modification in CD8⁺ T cell survival and effector response.

Results

FTO is required for effector CD8⁺ T cell responses during acute infection

To explore the effects of FTO-mediated m⁶A modification on CD8⁺ T cell function, we first generated T cell-specific Fto knockout (KO) mice (Fto^fl/flCD4-Cre⁺) and (Fto^fl/flCD4-Cre⁺OT-1⁺) mice by crossing Fto^fl/fl mice with CD4-Cre transgenic strain, and further crossing with OT-1 transgenic mice in which the T cells express TCR specific for ovalbumin peptide (OVA_257-264) SIINFEKL (Supplementary Fig. 1A, B). The deletion of FTO was confirmed by quantitative PCR (qPCR) at the mRNA level in CD8⁺ T cells from FTO KO (Fto^fl/flCD4-Cre⁺) mice compared to wild-type (WT) (Fto^fl/flCD4-Cre⁻) mice (Supplementary Fig. 1C). FTO ablation in T cells did not affect T cell development under the homeostatic conditions (Supplementary Fig. 2), which was consistent with the previous study [35]. However, FTO is essential for the protective function of CD8⁺ T cells against pathogenic infections. In an in vivo acute infection model (Fig. 1A), naïve CD8⁺ T cells from WT (Fto^fl/flCD4-Cre⁻OT-1⁺, CD45.1⁺) and FTO KO (Fto^fl/flCD4-Cre⁺OT-1⁺, CD45.2⁺) mice were transferred separately into the WT recipient mice (CD45.1⁺CD45.2⁺) followed by infection with Listeria monocytogenes (LM) expressing OVA (LM-OVA). Bacterial load was determined in the spleen and liver tissues 5 days post-infection. The results showed that mice transferred with FTO KO CD8⁺ T cells, compared to those with WT CD8⁺ T cells, had significantly increased bacterial colonies on agar plates in both spleen and liver (Fig. 1B), suggesting that FTO-deficient CD8⁺ T cells had impaired effector function against infection.

To further determine the intrinsic role of FTO in CD8⁺ T cells, we next co-transferred naïve CD8⁺ T cells from WT (Fto^fl/flCD4-Cre⁻OT-1⁺, CD45.1⁺) and FTO KO (Fto^fl/flCD4-Cre⁺OT-1⁺, CD45.2⁺) mice at a ratio of 1:1 into the same WT recipient mice (CD45.1⁺CD45.2⁺). After LM-OVA infection, the CD8⁺ T cell immune response in the secondary lymphoid tissue was monitored at different time periods (Fig. 1C). Compared to an equal number transferred on day 0, the ratio of FTO KO to WT OT-1⁺CD8⁺ T cells was significantly reduced in peripheral blood lymphocytes (PBL) following infection (Fig. 1D, E), strongly suggesting that FTO-deficient CD8⁺ T cells had a diminished immune response. Given that CD8⁺ T cell expansion reaches its peak around day 7 [2], we further examined the CD8⁺ T cell response at earlier timepoints before the expansion peak in the acute infection model. Consistently, FTO KO OT-1⁺CD8⁺ T cells exhibited a remarkable reduction in cell expansion on day 4, 5, and 6 in the spleen compared to WT counterparts (Fig. 1F and Supplementary Fig. 3A, B). Based on the expression of KLRG1 and CD127, effector CD8⁺ T cells at the peak of expansion can be divided into two subpopulations: KLRG1⁺CD127⁻ terminally differentiated short-lived effector cells (SLECs) and KLRG1⁻CD127⁺ long-lived memory precursor effector cells (MPECs) [36]. The evaluation of CD8⁺ T cell differentiation revealed that the frequencies of SLECs were comparable between WT and FTO KO groups, while the frequencies of MPECs were increased on days 4 and 5 but unchanged on day 6 (Fig. 1G, H and Supplementary Fig. 3C–F). However, owing to the massive reduction of total FTO KO CD8⁺ T cells, the absolute cell numbers of both SLECs and MPECs were decreased during the immune response (Fig. 1G, H and Supplementary Fig. 3C–F). Consistently, there was no difference in the expression of effector and memory CD8⁺ T cell-specific TFs T-bet and Eomes between WT and FTO KO CD8⁺ T cells (Fig. 1I, J). In addition, FTO-deficient CD8⁺ T cells displayed comparable cytokine production of IL-2 and IFN-γ and very slightly changed TNF-α, granzyme B, and perforin compared to their WT counterparts (Fig. 1K, L). Similarly, no differences in cytokine production were also observed in WT (Fto^fl/flCD4-Cre⁻OT-1⁺) and FTO KO (Fto^fl/flCD4-Cre⁺OT-1⁺) CD8⁺ T cells stimulated in vitro with OVA_257-264 peptide (Supplementary Fig. 3G). Collectively, these data elucidate a pivotal role of FTO in facilitating the CD8⁺ T cell population expansion during acute response, yet with limited effects on effector/memory cell differentiation and cytokine production.

FTO deficiency affects CD8⁺ T cell survival during immune responses

Upon antigen stimulation, CD8⁺ T cells undergo rapid cell activation, proliferation, and apoptosis before developing into effector and memory T cells [2]. To further explore the underlying mechanisms responsible for the diminished CD8⁺ T cell population after FTO ablation, we assessed the cellular activation, proliferation, and cell death in WT and FTO-deficient CD8⁺ T cells in both the in vitro stimulation model and in vivo co-transfer model. When naïve CD8⁺ T cells from WT (Fto^fl/flCD4-Cre⁻OT-1⁺) and FTO KO (Fto^fl/flCD4-Cre⁺OT-1⁺) mice were stimulated in vitro with cognate peptide (OVA_257-264), FTO-deficient OT-1⁺CD8⁺ T cells, compared to WT controls, showed comparable T cell activation after 24 h assessed by the expression of activation markers CD69 and CD25 (Supplementary Fig. 4A, B). Similarly, when naïve CD8⁺ T cells from WT (Fto^fl/flCD4-Cre⁻) and FTO KO (Fto^fl/flCD4-Cre⁺) mice were stimulated in vitro with anti-CD3/CD28 antibodies, T cell activation after 24 h was also unaltered between the two groups (Supplementary Fig. 4C, D). In terms of cell proliferation, we first measured the expression of cell-cycle marker Ki67 on CD8⁺ T cells obtained from the in vivo co-transfer model (Fig. 1C) at the expansion phase [2]. The results showed that FTO ablation did not affect CD8⁺ T cell proliferation on days 4 and 5, as evidenced by comparable expression of Ki67 (Fig. 2A, B). Similar results were observed with BrdU incorporation in WT and KO CD8⁺ T cells in the in vivo co-transfer model (Supplementary Fig. 4E). Moreover, when stimulated in vitro with OVA_257-264 peptide, FTO-deficient OT-1⁺CD8⁺ T cells also exhibited unchanged proliferation determined by both Ki67 staining and BrdU incorporation (Fig. 2C, D).

**Fig. 2: FTO deficiency affects effector CD8⁺ T cell survival.**

Nevertheless, FTO deficiency in CD8⁺ T cells caused massive cell death upon antigen stimulation. In the in vivo co-transfer model, FTO KO CD8⁺ T cells had significantly increased expression levels of Annexin V and 7AAD on both day 4 and 5 post-infection (Fig. 2E, F). Specifically, the proportions of Annexin V⁺7AAD⁻ early apoptotic cells, but not Annexin V⁺7AAD⁺ late apoptotic cells and Annexin V⁻7AAD⁺ necroptotic cells, were markedly elevated in FTO KO CD8⁺ T cells (Fig. 2G). Consistently, the frequencies of total apoptotic and early apoptotic cells were significantly increased in FTO-deficient CD8⁺ T cells stimulated in vitro with OVA_257-264 peptide (OT-1⁺CD8⁺ T cells) (Fig. 2H) and anti-CD3/CD28 antibodies (WT CD8⁺ T cells) (Fig. 2I) for both 24 h and 48 h. These data indicate that FTO deletion has a limited impact on CD8⁺ T cell activation and proliferation but causes significant cell death during immune responses.

FTO deficiency results in increased CD8⁺ T cell apoptosis

To reveal the mechanisms underlying the T cell death in FTO KO CD8⁺ T cells, we employed small molecule inhibitors of different cell death pathways. WT (Fto^fl/flCD4-Cre⁻) and FTO KO (Fto^fl/flCD4-Cre⁺) CD8⁺ T cells were stimulated in vitro with anti-CD3/CD28 antibodies in the presence of the necroptosis inhibitor necrostatin-1 (NEC-1) [37], the pyroptosis inhibitor VX-765 [38], the ferroptosis inhibitor ferrostatin-1 (Fer-1) [39] and the apoptosis inhibitor Z-VAD-FMK (Z-VAD) [40]. Interestingly, NEC-1, VX-765, and Fer-1 could not rescue the impaired CD8⁺ T cell survival caused by FTO ablation (Supplementary Fig. 5A–C). Notably, the apoptosis inhibitor Z-VAD significantly restored the upregulated apoptosis in FTO KO CD8⁺ T cells (Fig. 3A). Genome-wide transcriptomic analysis (RNA-seq) was performed to compare the transcriptomic signatures between WT and FTO KO OT-1⁺CD8⁺ T cells isolated from recipient mice 5 days after LM-OVA infection in the in vivo co-transfer model (Fig. 1C). We found that 522 and 286 genes were upregulated and downregulated (fold-change greater than 2, P-value < 0.05), respectively in FTO KO CD8⁺ T cells (Supplementary Data 1). Gene Ontology (GO) analysis of the differentially expressed genes (DEGs) from RNA-seq data revealed that in FTO-deficient CD8⁺ T cells, genes associated with programmed cell death and apoptotic process were significantly enriched while genes associated with activation of immune response and negative regulation of apoptotic process were significantly downregulated compared to WT controls (Fig. 3B). Gene set enrichment analysis (GSEA) of the hallmark pathways identified enriched gene sets involved in cell survival pathways, such as TNF signaling, p53 pathway, MYC targets and apoptosis in KO CD8⁺ T cells (Fig. 3C).

**Fig. 3: FTO-deficient effector CD8⁺ T cells have increased cell apoptosis.**

Extrinsic apoptotic pathways triggered by death receptors (e.g., Fas) can activate the intrinsic apoptotic pathway involving mitochondrial and DNA damages [41]. In line with this notion, the expression of the anti-apoptotic protein Bcl-2 was remarkably decreased in FTO KO (Fto^fl/flCD4-Cre⁺OT-1⁺) CD8⁺ T cells obtained from the in vivo co-transfer model on days 4 and 5, compared to WT (Fto^fl/flCD4-Cre⁻OT-1⁺) controls (Fig. 3D). And the expression of cleaved caspase-3, a hallmark of apoptosis, was also remarkably elevated in FTO-deficient CD8⁺ T cells stimulated in vitro with OVA_257-264 peptide (OT-1⁺CD8⁺ T cells) (Fig. 3E) and anti-CD3/CD28 antibodies (WT CD8⁺ T cells) (Fig. 3F). Meanwhile, the downregulation of Bcl-2 and upregulation of cleaved caspase-3 were further verified by Western blot in WT and FTO-deficient CD8⁺ T cells stimulated with anti-CD3/CD28 antibodies (Fig. 3G). Together, these data indicate that the significantly reduced FTO-deficient CD8⁺ T cells during immune response are attributed to cell apoptosis.

FTO regulates CD8⁺ T cell survival via m⁶A modification on Fas

FTO, as one critical m⁶A demethylase, plays vital roles in a variety of biological processes through regulating the mRNA m⁶A levels and expression of target genes [22]. To identify the underlying mechanisms by which FTO controls CD8⁺ T cell survival during immune response, we performed m⁶A-methylated RNA immunoprecipitation sequencing (MeRIP-seq) to map the m⁶A landscape in WT and FTO KO CD8⁺ T cells obtained from the recipient mice 5 days after LM-OVA infection in the in vivo co-transfer model. The m⁶A sites distribution was displayed across the transcript regions 5′ UTR, coding sequence (CDS) and 3′ UTR. The density of m⁶A peaks was significantly enriched in the 3′ UTR region with slightly decreased in abundance at the 3′ UTR and increased abundance at the 5′ UTR and CDS regions in FTO KO groups (Fig. 4A, B). Comparing the differentially expressed methylation peaks, there were 581 and 3241 hyper-methylated and hypo-methylated (fold-change greater than 1.5, P-value < 0.05) transcripts, respectively, in FTO KO CD8⁺ T cells (Supplementary Data 2). Given that loss of FTO causes an upregulation of m⁶A level, we focused specifically on those hyper-methylated transcripts. GO term analysis of those transcripts uncovered a significant enrichment in pathways associated with stress response, DNA damage, programmed cell death and apoptotic signaling (Fig. 4C). In addition, pathway enrichment analysis of RNA-seq data based on C7 immunologic gene sets revealed that the deletional tolerance-associated apoptotic pathway was enriched in FTO-deficient CD8⁺ T cells (Fig. 4D). It is well-appreciated that T cell apoptosis mediated by death receptors—the most well-studied one is Fas (also known as CD95)—represents a key mechanism for periphery immunological tolerance, a process known as activation-induced cell death (AICD) [42]. Notably, among those genes in deletional tolerance pathway with m⁶A hyper-methylation, the Fas mRNA was highly marked by m⁶A modification at CDS regions visualized by Integrative Genomics Viewer (IGV) (Fig. 4E). Further, the m⁶A-RIP-qPCR results also confirmed that the m⁶A level on Fas mRNA was substantially increased in FTO-deficient CD8⁺ T cells (Fig. 4F), indicating that FTO modulates the m⁶A methylation on Fas mRNA in CD8⁺ T cells upon antigen activation.

Accordingly, the Fas expression was significantly elevated in FTO-deficient OT-1⁺CD8⁺ T cells, compared to WT counterparts, in the in vivo co-transfer model on days 4 and 5 post-infection (Fig. 4G). Similar results were also observed in FTO-deficient CD8⁺ T cells stimulated in vitro with both OVA_257-264 peptide (OT-1⁺CD8⁺ T cells) (Fig. 4H) and anti-CD3/CD28 antibodies (WT CD8⁺ T cells) (Fig. 4I) for 8 h and 24 h. It is well-appreciated that the extrinsic apoptosis pathways, induced by death receptor (i.e., Fas), lead to the recruitment and activation of caspase-8, which in turn directly cleaves caspase-3 and thereby induces cell apoptosis [41, 43]. Hence, we next measured the caspase-8 activity in WT and FTO CD8⁺ T cells upon antigen stimulation. In both OT-1⁺ and WT CD8⁺ T cells, stimulated in vitro with OVA_257-264 peptide and anti-CD3/CD28 antibodies respectively, FTO deficiency led to markedly elevated caspase-8 activity compared to WT groups (Fig. 4J). To provide further evidence of upregulated Fas impairing CD8⁺ T cell survival, we examined the cell apoptosis in WT and FTO KO CD8⁺ T cells stimulated in vitro with anti-CD3/CD28 antibodies in the presence or absence of anti-FasL blocking antibodies. As expected, Fas signaling blockade significantly mitigated the upregulated cell death and early apoptosis caused by FTO deficiency in activated CD8⁺ T cells (Fig. 4K). Furthermore, to evaluate the impact of FTO on endogenous CD8⁺ T cell responses, we conducted a direct infection model on WT and FTO conditional KO mice. WT (Fto^fl/flCD4-Cre⁻) and FTO KO (Fto^fl/flCD4-Cre⁺) mice were infected by intravenous injection of LM that expresses the GP33 peptide (LM-GP33) [44], a well-characterized CD8⁺ T cell epitope derived from the glycoprotein of lymphocytic choriomeningitis virus (LCMV) (Supplementary Fig. 6A). 7 days post-infection, the frequency and cell number of both total CD8⁺ T cells (Supplementary Fig. 6B) as well as antigen-specific tetramer-GP33⁺ CD8⁺ T cells (Supplementary Fig. 6C) were remarkably reduced in FTO KO mice compared to WT counterparts. In addition, the cell apoptosis (Supplementary Fig. 6D) and Fas expression (Supplementary Fig. 6E) were also upregulated in FTO KO CD8⁺ T cells. Together, these data suggest that FTO directly demethylates m⁶A on Fas mRNA; thus, FTO ablation leads to an upregulation of m⁶A modification and subsequent Fas expression, which promotes cell apoptosis in FTO KO CD8⁺ T cells upon antigen stimulation.

Increased m⁶A methylation on Fas mRNA enhances its stability

Given that m⁶A methylation on mRNA modulates gene expression through influencing mRNA stability and translation efficiency [10], we next sought to explore whether the upregulated m⁶A modification on Fas mRNA affects its mRNA stability and translation. We first detected the mRNA levels of Fas in WT and FTO KO CD8⁺ T cells stimulated with anti-CD3/CD28 antibodies in vitro for different periods of time. The qPCR data showed that FTO-deficient CD8⁺ T cells had augmented mRNA expression of Fas along with the stimulation compared to WT cells (Fig. 5A). RNA decay assay was further performed by administration of actinomycin D (ActD), a transcription inhibitor, during CD8⁺ T cell stimulation to evaluate the stability of Fas mRNA. We found that the Fas mRNA had decelerated decay kinetics in FTO-deficient CD8⁺ T cells (Fig. 5B, C), suggesting an enhancement of Fas mRNA stability upon FTO ablation. To explore the effects of protein translation and degradation, we next applied protein-synthesis inhibitor cycloheximide (CHX) and proteasome inhibitor MG132 during CD8⁺ T cell activation. WT and FTO KO naïve CD8⁺ T cells were stimulated in vitro with anti-CD3/CD28 antibodies for 12 h before adding CHX to block protein synthesis. In agreement with the above results, Fas expression was rapidly elevated upon TCR stimulation, and FTO-deficient CD8⁺ T cells exhibited markedly increased Fas levels (Fig. 5D, E). CHX treatment led to a downregulation of Fas expression in both WT and KO groups (Fig. 5D, E). However, the decline of Fas expression after CHX treatment was comparable between WT and KO cells when comparing to its original expression level at 0 h (before CHX treatment) (Fig. 5F). Additionally, WT and FTO KO naïve CD8⁺ T cells were stimulated with anti-CD3/CD28 antibodies for 4 h before adding in MG132 to block protein degradation. The results showed that FTO-deficient CD8⁺ T cells displayed significantly enhanced Fas expression upon stimulation irrelative with MG132 treatment (Fig. 5G, H). Similarly, the dynamic expression of Fas after MG132 was comparable between WT and KO cells (Fig. 5I). All these data suggest that FTO ablation promotes Fas expression via enhancing the Fas mRNA m⁶A methylation and its stability.

**Fig. 5: FTO deficiency enhances *Fas* mRNA stability in CD8⁺ T cells.**

FTO modulates Fas expression dependent on m⁶A reader IGF2BP3

To validate the FTO regulation of Fas expression dependent on m⁶A modification, we analyzed the m⁶A methylation sites on Fas mRNA based on the conserved m⁶A motif RRACH (R = G or A; H = A, C, or U) [45], and yielded four upregulated m⁶A sites within the CDS region in FTO-deficient CD8⁺ T cells (Fig. 6A). We then generated constructs containing both WT and mutant Fas with m⁶A methylation sites mutation. Owing to a close proximity (within 10 bp) between the first two sites (shown in IGV), we constructed three Fas mutants with mutations at both site 1 and 2 (mutant 1), site 3 (mutant 2) and site 4 (mutant 3) (Fig. 6A). As anticipated, overexpression of WT Fas in both WT and FTO KO CD8⁺ T cells could significantly elevate the Fas expression and cell apoptosis compared to that overexpressed with the empty vector (Mock) (Fig. 6B, C, first four panels). More noticeably, overexpression of WT Fas in FTO KO CD8⁺ T cells, compared to WT CD8⁺ T cells, resulted in even higher levels of Fas expression and cell apoptosis (Fig. 6B, C, first four panels). Notably, transfection with both Fas WT and mutants resulted in comparable Fas expression levels (Supplementary Fig. 7A). However, overexpression of Fas mutant 1, but not mutant 2 and 3, into FTO KO CD8⁺ T cells significantly decreased the Fas expression, total cell apoptosis as well as early apoptosis levels (Fig. 6B, C, last three panels), suggesting that FTO likely regulates CD8⁺ T cell apoptosis through modifying the m⁶A methylation on Fas mRNA at site 1 and 2.

**Fig. 6: Loss of FTO enhances the IGF2BP3-mediated transcription of Fas.**

Furthermore, m⁶A modification is a dynamic process in which the methyltransferases and demethylases mainly control the m⁶A levels on target transcripts, whereas the RNA binding proteins (RBPs), namely m⁶A readers, determine the biological function of these modifications [10]. Given that IGF2BPs (IGF2BP1/2/3), among distinct types of m⁶A reader proteins, play a dominant role in modulating the stability of m⁶A-targeted mRNAs [46], we mainly focused on this family of proteins. We first measured the expression of IGF2BPs in WT and Fas KO CD8⁺ T cells isolated from recipient mice 5 days after infection in the in vivo co-transfer model by qPCR, and found that IGF2BP3, but not IGF2BP1 and IGF2BP2, had enhanced expression in KO cells, suggesting it might play a regulatory role in FTO ablated CD8⁺ T cells (Supplementary Fig. 7B). In addition, according to the m⁶A2Target website (http://rm2target.canceromics.org), a comprehensive database containing m⁶A writer, eraser, and reader target genes, IGF2BP3 has been shown to bind to Fas with m⁶A modification (Supplementary Table 1) [47]. To validate its role in mediating the upregulation of Fas in FTO-deficient CD8⁺ T cells, the knockdown of Igf2bp3 using shRNA was performed. Transduction of Igf2bp3 shRNA into WT CD8⁺ T cells could efficiently downregulate IGF2BP3 expression assessed by qPCR (Supplementary Fig. 7C). When Igf2bp3 shRNA was introduced into FTO KO CD8⁺ T cells, compared to WT and FTO KO CD8⁺ T cells transduced with Mock, the knockdown of Igf2bp3 significantly reduced the enhanced Fas expression and cell apoptosis caused by FTO ablation (Fig. 6D, E). Collectively, these data suggest that FTO-mediated m⁶A methylation regulates Fas expression in activated CD8⁺ T cells in a manner dependent on m⁶A reader protein IGF2BP3.

Discussion

Understanding the underlying regulatory mechanisms involved in CD8⁺ T cell immune responses is profoundly essential in protecting against infection and tumors, as well as offering potential opportunities in the clinical development of T cell based immunotherapy [48]. Mechanisms regulating CD8⁺ T cell differentiation and functionality during infections at the transcriptional, epigenetic (DNA and histone modification), and metabolic levels have been extensively characterized [2, 4]. Epigenetic regulations at the RNA level, such as RNA m⁶A methylation, have gained emerging attention for their essential roles in a variety of biological and pathological conditions [10]. In this study, we investigated the effects of m⁶A modification in CD8⁺ T cell immune responses by targeting the m⁶A demethylase FTO. We unveiled that FTO is indispensable for maintaining CD8⁺ T cell survival, expansion, and immunological function during acute infection. Our data revealed that T cell-specific deficiency in FTO led to an elevation of apoptosis in CD8⁺ T cells upon antigen stimulation. Loss of FTO resulted in an upregulation of m⁶A modification on Fas mRNA, which promoted the mRNA stability dependent on m⁶A reader IGF2BP3. The increased mRNA and consequent protein expression of Fas triggered the apoptotic process in activated CD8⁺ T cells (Fig. 7).

**Fig. 7: Mechanistic scheme showing the FTO-mediated m⁶A methylation of Fas in regulating CD8⁺ T cell survival and effector response.**

Emerging evidence has demonstrated that m⁶A modification plays a key role in regulating the immunological functions of T cells, yet the majority of early studies focused on CD4⁺ T cell homeostasis and differentiation [49]. Recently, Guo et al. investigated the intrinsic role of m⁶A methyltransferase METTL3 in CD8⁺ T cell responses during acute viral infection [18]. They reported that METTL3-deficient CD8⁺ T cells exhibited impaired effector cell differentiation, defective cell survival, cytokine production, and subsequent memory formation, which relied on the regulatory mechanism of METTL3-m⁶A-T-bet axis [18]. Interestingly, our study, by targeting m⁶A demethylase FTO in T cells, revealed significantly different phenotypes in CD8⁺ T cells during acute infection. Our results elucidated that FTO ablation led to a dramatic CD8⁺ T cell apoptosis attributed to the upregulated Fas expression upon antigen stimulation. However, FTO deficiency had limited effects on CD8⁺ T cell proliferation, cytokine production, and effector cell differentiation. These divergent data explicitly suggest that m⁶A methyltransferases and demethylases regulate distinct biological events or functions, likely through modifying distinct genes, even in the same type of cells. Similar results were observed in m⁶A regulation of Th17 cell differentiation. METTL3-deficient naïve CD4⁺ T cells showed less differentiation into Th17 cells, whereas ALKBH5 deficiency in CD4⁺ T cells also resulted in impaired IL-17 signaling [35]. METTL3 or METTL14 knockdown in CD4⁺ T cells could promote Tfh cell development upon LCMV infection [50]. However, other studies reported that METTL3 ablation in CD4⁺ T cells had impaired Tfh cell differentiation upon LCMV infection [17]. Moreover, METTL3 deficiency induced cell apoptosis in CD8⁺ T cells [18] but not in CD4⁺ T cells upon TCR stimulation [15]. Distinct phenotypes could be observed in CD8⁺ T cells with METTL3 ablation driven by different Cre promoter (Cd4-Cre versus Gzmb-Cre), which was related to different T cell activation status [18]. All these findings strongly indicate that m⁶A modification is a biological context-dependent process, highly associated with enzymatic activity, cell types, and immunological environment.

Since FTO was first discovered to be associated with fat mass and obesity, a large number of studies have demonstrated that FTO plays critical roles in metabolic diseases, such as obesity and diabetes, and cardiovascular diseases [51, 52]. FTO modulates adipocyte biology by promoting differentiation, adipogenesis and lipid metabolism [52, 53]. FTO inhibits insulin secretion in pancreatic β cells through NF-κB activation [54, 55]. Emerging recent evidence has revealed the essential function of FTO in promoting tumorigenesis, and FTO can serve as a potential cancer therapeutic target [24, 56]. FTO exerts oncogenic effects on various types of cancer cells, promoting tumor cell growth by activating survival and cell-cycle signaling, such as PI3K/Akt/mTOR pathway, MYC, β-catenin, and CEBPA [10, 57]. However, the regulatory roles of FTO in immune cells are much less studied. A few evidence has revealed that FTO, through meditating m⁶A methylation, regulates differentiation and function in macrophages and NK cells [25,26,27,28]. These findings indicate that FTO-mediated m⁶A regulates gene expression in cell type- and biological context-dependent manners. In this study, we evaluate the function of FTO in CD8⁺ T cells in the adaptive immune system. FTO plays a pivotal role in maintaining cell survival in CD8⁺ T cells during immune responses through controlling the m⁶A levels and mRNA stability of the death receptor Fas. This data underscores the special mechanism of FTO-mediated m⁶A modification on apoptosis is uniquely pronounced in CD8⁺ T cells.

As a member of the TNFR superfamily of death receptors, Fas interacts with FasL, initiating a complex cascade of cell apoptosis and playing important roles in T cell immune responses [58]. Apoptosis is primarily mediated by two classical pathways: the extrinsic pathway triggered by death receptors (e.g., Fas) and the intrinsic pathway involving mitochondrial and DNA damages [41, 59]. Extrinsic apoptotic pathways can activate the intrinsic apoptotic pathway, linking death receptor signaling to mitochondrial apoptosis. Activation of Fas/FasL signaling recruits FADD, leading to caspase-8 activation. Active caspase-8 cleaves Bid into tBid, which translocates to mitochondria and inhibits anti-apoptotic Bcl-2 proteins, thereby initiating the intrinsic pathway, caspase-3 cleavage, and final cell apoptosis [41, 60]. In this study, we observed upregulated Fas expression, increased caspase-8 activity, which is the primary downstream effector of Fas/FasL signaling, elevated caspase-3 activity, and enhanced cell apoptosis, accompanied by reduced Bcl-2 levels in FTO KO CD8⁺ T cells. These findings indicate that the interplay between extrinsic and intrinsic apoptotic pathways cooperatively contributes to the cell apoptosis of FTO-deficient CD8⁺ T cells. Furthermore, Fas-triggered T cell apoptosis depends on TCR ligation, which increases Fas expression and provides additional “competency signal” to ensure antigen-specific T cell apoptosis [61]. However, TCR stimulation does not elicit immediate T cell death [62]. The underlying mechanism was suggested to be associated with a few days-delay of surface expression of Fas or FasL after TCR ligation [63]. Besides regulation by gene transcription and translation, we here found that Fas expression was governed by a novel epigenetic regulation of RNA m⁶A modification. Under the physiological condition, TCR stimulation induces Fas gene transcription with a balanced m⁶A modification on Fas mRNA, which results in a gradual increase of Fas expression during the first few days of CD8⁺ T cell activation. The upregulated Fas expression subsequently induces apoptosis to avoid T cell overactivation [43]. However, with the ablation of m⁶A eraser FTO, the m⁶A levels on Fas mRNA were significantly increased, which promoted its mRNA stability. Therefore, the elevated m⁶A modification on Fas mRNA induces an accelerated elevation of Fas expression and sensitivity to Fas-induced apoptosis during CD8⁺ T cell immune response.

The m⁶A methylation is dynamically controlled by methyltransferases and demethylases, whereas the eventual biological functions of m⁶A modification are determined by m⁶A reader proteins. The m⁶A reader proteins recognize and bind to m⁶A-marked transcripts to mediate a variety of processes, such as mRNA stability, splicing, translocation, and translation [10]. Distinct m⁶A readers exist in different cell types and conduct biological context-dependent functions. For instance, previous work has shown that METTL3-mediated m⁶A modification exhibited discrete functions in naïve CD4⁺ T cells and Treg cells [15, 64]. m⁶A reader proteins can be classified into three categories: YTH family proteins, HNRNP family proteins and IGF2BP family proteins [11]. IGF2BPs are recently identified an RBP family as m⁶A readers, mainly promoting the stability and storage of their target mRNAs [65]. Our data demonstrate that FTO ablation leads to increased m⁶A methylation on Fas mRNA, which promotes Fas expression via enhancing the Fas mRNA stability. Therefore, in the investigation of m⁶A readers that mediate FTO’s regulation of Fas mRNA stability, we mainly focused on IGF2BP family members. IGF2BPs have been shown to play critical roles in promoting cancer cell growth and creating immunosuppressive tumor microenvironment in various types of cancer [66]. However, the direct regulatory roles of IGF2BPs in T cells are currently undefined. Here, we found that IGF2BP3 mediated the enhanced Fas mRNA stability in FTO-deficient CD8⁺ T cells, thus, knockdown IGF2BP3 significantly decreased the Fas expression and cell apoptosis caused by FTO ablation. These findings suggest that the m⁶A-mediated regulation in CD8⁺ T cells is a sophisticated process orchestrated by the full m⁶A machinery of readers, erasers, and readers.

The current findings of FTO regulating m⁶A modification on Fas mRNA through IGF2BP3 are supported by the following pieces of evidence: [1] MeRIP-seq and MeRIP-PCR data identifying upregulated m⁶A peaks on Fas mRNA in FTO-deficient CD8⁺ T cells, which is consistent with the demethylase activity of FTO [2]. Site-directed mutagenesis confirms the functional relevance of FTO-mediated m⁶A methylation in Fas expression and apoptosis regulation [3]. The public database (m⁶A2Target website) demonstrates a direct IGF2BP3 binding to Fas mRNA depending on m⁶A. However, more direct experimental evidence of FTO/IGF2BP3 binding to m⁶A sites of Fas mRNA, such as CLIP-PCR, biotin-labeled RNA pull-down, and RNA binding protein immunoprecipitation (RIP), would further strengthen this study. In addition, as a critical m⁶A demethylase, FTO is broadly expressed and exerts diverse regulatory roles in diverse biological processes depending on different cellular contexts [24, 67]. Here, we found that FTO ablation in CD8⁺ T cells mostly led to increased apoptosis. Thus, to identify FTO targets, we primarily focused on cell death/apoptosis-related genes in the MeRIP-seq data. However, the m⁶A levels of majority of those genes, such as Bcl-2, Myc, Faslg, Fadd, Bcl2l1 (Bcl-xl), Cflar (cFLIP), Bax, Bak, Casp3, Casp7, and Casp8, were not upregulated, while Fas was emerged as a key candidate in FTO KO CD8⁺ T cells. While we acknowledge that FTO may also influence CD8⁺ T cell immune responses indirectly through other signaling pathways, further systematic exploration of FTO’s broader regulatory network, for example, employing genome-wide CRISPR screens and multi-omics approaches, is required in future research.

In conclusion, our study defines a novel mechanism of FTO-mediated m⁶A modification in maintaining CD8⁺ T cell survival upon antigen stimulation, providing new insights into understanding the RNA-based epigenetic regulation in CD8⁺ T cell immune responses. Notably, the discrepant findings observed in FTO and METTL3-deficient CD8⁺ T cells evidently suggest that m⁶A readers and erasers have markedly distinct downstream genes. It will be intriguing to explore the exact regulatory mechanisms that m⁶A readers and erasers employ in CD8⁺ T cells in the future.

Materials and methods

Mice

Fto^fl/fl, CD4-Cre ⁺, CD45.1⁺, and OT-1 strains were purchased from the Jackson Laboratory (Bar Harbor, ME, USA). Fto^fl/fl mice were crossed with CD4-Cre and OT-1 mice to generate Fto^fl/flCD4-Cre⁺(OT-1⁺) (KO) and Fto^fl/flCD4-Cre⁻(OT-1⁺) (WT) mice. CD45.1⁺ mice were crossed with OT-1⁺ or CD45.2⁺ mice to generate OT-1⁺CD45.1⁺ or CD45.1⁺CD45.2⁺ mice. 8-12-week-old female mice were preferentially used in this study. All mice were housed in specific-pathogen free conditions by the Xi’an Jiaotong University Division of Laboratory Animal Research.

Antibodies and flow cytometry

Single-cell preparations were stained with the monoclonal antibodies purchased from Biolegend (San Diego, CA, USA): anti-mouse CD45.1 (#110716, clone A20), anti-mouse CD45.2 (#109806, clone 104), anti-mouse CD3E (#100334, clone 145-2C11), anti-mouse CD8A (#100725, clone 53-6.7), anti-mouse KLRG1 (#138412, clone 2F1/KLRG1), anti-mouse IL-7R (#135014, clone A7R34), anti-mouse CD44 (#103028, clone IM7), anti-mouse CD62L (#104412, clone MEL-14), anti-mouse CD25 (#102030, clone PC61), anti-mouse CD69 (#104508, clone H1.2F3), anti-mouse IL-2 (#503820, clone JES6-5H4), anti-mouse IFN-γ (#505826, clone XMG1.2), anti-mouse TNF-α (#506308, clone Mp6-XT22), anti-mouse granzyme B (#372208, clone QA16A02), anti-mouse perforin (#154306, clone S16009a), anti-mouse T-bet (#644824, clone 4B10), anti-mouse Eomes (#157706, clone W17001A), anti-mouse Bcl-2 (#658706, clone 100), anti-mouse CD95 (#152608, clone SA367H8), anti-mouse BrdU (#364116, clone 3D4) and anti-mouse Ki67 monoclonal antibody (#652422, clone 16A8). Anti-mouse active Caspase-3 monoclonal antibody (#570334, clone C92-605.rMAb) was obtained from BD Biosciences. Phorbol 12-myristate 13-acetate (PMA, # HY-18739), Ionomycin (#HY-13434), brefeldin A (#HY-16592), and monensin (#HY-N0150) were purchased from MedChemExpress.

For analysis of surface markers, cells were stained in PBS containing 1% FBS on ice for 30 min. For analysis of intracellular cytokine staining, cells were stimulated for 4 h in vitro with PMA/Ionomycin in the presence of brefeldin A and monensin. The stimulated cells were fixed and permeabilized using a Fixation/Permeabilization Solution Kit (#420801 and #421002, Biolegend). For nuclear protein staining, such as Ki67, T-bet, Eomes, and Bcl-2, cells were fixed and permeabilized using a Transcription Factor Fixation/Permeabilization Concentrate and Diluent purchased from eBioscience (#00-5523-00, San Diego, CA, USA).

For BrdU staining, cells were first stained with indicated surface markers followed by BrdU staining according to the manufacturer’s instruction (#554714, BrdU Flow Kit, BD Biosciences). For apoptosis, cells were stained with 7-Aminoactinomycin D (7AAD, #420404)/Annexin V (#640947) (Biolegend). Cells were analyzed using a CytoFLEX flow cytometer (BECKMAN COULTER). FlowJo software and CytExpert software were used for data analysis. For CD8⁺ T cell sorting, FACSAria II sorter was applied (BD Biosciences).

Quantitative PCR (qPCR)

To examine the mRNA levels, Quick-RNA Microprep Kit (#R1051, Zymo Research) was used to extract total RNA from purified T cells according to the manufacturer’s instruction. cDNA was reverse-transcribed using the cDNA synthesis kit (#FSQ-301, TOYOBO) and amplified with SYBR Green RT-qPCR Mastermix (#A311-05, GenStar) at StepOnePlusTM Real-Time PCR System (ThermoFisher). Primers were synthesized from Tsingke Biotechnology. Primer sequences used in this study were: mouse FTO forward 5′-AAAGTTTGAAGGAGGGGAGAAGTG-3′, reverse 5′-ACCAAAGAGGGGGAGACAGTTACG-3′; mouse Fas forward 5′- AAGTCCCAGAAATCGCCTATG-3′, reverse 5′- GGTATGGTTTCACGACTGGAG-3′; mouse IGF2BP1 forward 5′- CCTGGCTCATAACAACTTCGTCG-3′, reverse 5′- CCTTCACAGTGATGGTCCTCTC-3′; mouse IGF2BP2 forward 5′- TGAAGCCTGTGCCAATGCTGAG-3′, reverse 5′- CCAGTCGAAAAGATGCCAAGTGC-3′; IGF2BP3 forward 5′- CCACCCAGTTTGTTGGAGCCAT-3′, reverse 5′- GGATAGTAATGGACTTCTCCGCG-3′; and mouse HPRT forward 5′- AGTACAGCCCCAAAATGGTTAAG-3′, reverse 5′- CTTAGGCTTTGTATTTGGCTTTTC-3′.

Western blotting

Total proteins were extracted from CD8⁺ T cells at indicated experiments by cell lysis in RIPA buffer (#P0013B, Beyotime Biotechnology). Protein samples were separated on SDS-polyacrylamide gels and electro-transferred onto Polyvinylidene fluoride (PVDF) membranes (#IPVH00010, Millipore). Membranes were then blocked with 5% skimmed milk before incubation with indicated primary antibodies at 4 °C overnight, followed by incubation with secondary antibodies at room temperature for one hour. The protein expression was measured by Fusion-Solo.6s (VILBER). Primary antibodies used in Western blot assay were: Bcl-2 (#sc-7382, clone C-2, Santa Cruz), cleaved caspase-3 (#9664, clone 5A1E, CST) and Actin (#60008-1-Ig, clone 7D2C10, Proteintech). The secondary antibodies purchased from Cwbio were HRP goat anti-rabbit IgG (#CW0103S) and HRP goat anti-mouse IgG (#CW0102S). Uncropped western blots were included in Supplementary Data 3.

T cell adoptive transfer and LM Infection

In the in vivo infection model, naïve OT-1⁺CD8⁺ T cells were sorted using FACSAria II (BD Bioscience) cell sorter from WT (CD45.1⁺) and KO (CD45.2⁺; Fto^fl/flCD4-Cre⁺) mice and either separately (each 1 × 10⁵) transferred or co-transferred (each 5 × 10⁴) at a ratio of 1:1 into WT recipient mice (CD45.1⁺CD45.2⁺). Twenty-four hours after cell transfer, mice were infected with 2 × 10⁴ colony-forming units (CFU) of LM-OVA intravenously (i.v.). The lymphocytes in peripheral blood and spleen tissues were analyzed on the indicated days post-infection. To examine in vivo proliferation, mice were injected intraperitoneally (i.p.) with 1 mg BrdU (#B5002, Sigma-Aldrich) for 4 h before cell analysis. For quantification of bacterial burden, 1 × 10⁶ CFU LM-OVA was applied, and spleens and livers from infected mice were homogenized in PBS. Serial dilutions of the tissue homogenate were prepared, and 50 μl of each dilution was cultured on brain-heart infusion (BHI) agar plates. After incubation overnight at 37 °C, colonies were counted, and the LM CFUs recovered from each tissue were calculated. In the direct infection model, WT and FTO KO mice were infected with 1 × 10⁵ CFU of LM-GP33 intravenously (i.v.). The lymphocytes in the spleen were analyzed 7 days post-infection. Antigen-specific CD8⁺ T cells were stained by GP33 tetramer (NIH Tetramer Core Facility, Atlanta, GA, USA).

T cell in vitro stimulation

To examine the T cell activation, proliferation, and apoptosis in vitro, splenocytes were isolated and stimulated with 2.5 µg/ml coated anti-CD3 (#100302, clone 145-2C11, Biolegend) and 1 µg/ml soluble anti-CD28 (#102102, clone 37.51, Biolegend) antibodies. Splenocytes isolated from OT-1⁺ mice were stimulated with 1 µg/ml OVA_257-264 peptide. The following stimulation timepoints were applied: 24 h (activation), 48 h (proliferation), or indicated timepoints (apoptosis). For BrdU incorporation, 10 µM BrdU (#B5002, Sigma-Aldrich) was added 40 min before cell analysis. To examine the cell death pathway, CD8⁺ T cells were stimulated with anti-CD3/CD28 antibodies in the presence of 10 µM NEC-1 (#HY-15760, MedChemExpress), 20 µM VX-765 (#S2228, Selleck), 10 µM Fer-1 (#SML0583, Sigma-Aldrich), 20 µM Z-VAD (#S7023, Selleck) and 10 µg/ml anti-FasL blocking antibodies (#106611, clone MFL3, Biolegend) for 48 h. To evaluate the effects of protein synthesis and degradation, CD8⁺ T cells were stimulated with anti-CD3/CD28 antibodies for 12 h and 4 h before adding in 30 µg/ml CHX (#HY-12320, MedChemExpress) and 10 µM MG132 (#HY-13259, MedChemExpress), respectively, for another indicated time periods.

RNA decay assay

WT and FTO CD8⁺ T cells were stimulated with anti-CD3/CD28 antibodies for 4 h before treatment with the transcription inhibitor actinomycin D (5 µg/ml, # HY-17559, MedChemExpress) for another 1, 2 and 4 h before cell collection. Total RNA was isolated by using a Quick-RNA Microprep Kit (#R1051, Zymo Research). qPCR was performed as described above to quantify the relative levels of target mRNAs.

Analysis of caspase-8 activity

WT and FTO CD8⁺ T cells were stimulated with anti-CD3/CD28 antibodies for 24 h. The caspase-8 activity was assessed by the Caspase-8 Activity Colorimetric Assay Kit (#C1151, Beyotime Biotechnology) according to the manufacturer’s protocol.

cDNA and shRNA constructs, retroviral production, and transfection

Expression plasmids of mouse Fas WT and mutants, and Igf2bp3 shRNA were constructed into MSCV-based expressing vectors, which was engineered to include a EGFP reporter gene. Mutations were introduced via site-directed mutagenesis PCR. BOSC 23 cells (from Dr. Garry Nolan Lab) [68] were used for retrovirus generation. Mycoplasma contamination of cell lines was regularly checked. Retrovirus was produced by transfecting 5 µg of each retroviral vector in combination with 1 μg pCL-Eco retrovirus packaging vector into 6-well plates of cultured BOSC 23 cells using jetPRIME transfection reagent (#101000001, Polyplus). Following 48 h and 72 h, the culture supernatant containing retrovirus was collected through 0.45 µm filters. Purified CD8⁺ T cells were stimulated overnight using plate-bound anti-CD3 (5 µg/ml) and anti-CD28 antibodies (2 µg/mL). Spin infection of stimulated T cells using retrovirus supernatant was performed at 2500 rpm for 90 min at 37 °C. EGFP⁺ cells were identified as successfully transfected cells and examined using flow cytometry analysis or sorted for further experiments. The Igf2bp3 shRNA sequence was GAATCATCTGAAGTTCCTCAA.

RNA-Seq library preparation and sequencing

For RNA-seq analysis, WT and FTO KO CD8⁺ T cells were isolated from recipient mice 5 days after LM-OVA infection in the in vivo co-transfer model. Total RNA was extracted using the RNeasy Mini Kit (#74104, Qiagen) according to the manufacturer’s protocol. RNA-seq analysis was performed by Genergy Biotechnology according to the manufacturer’s standard protocol. RNA concentration was detected by the Qubit RNA broad range assay in the Qubit Fluorometer (Invitrogen). After quality control using an RNase-free agarose gel and Agilent 2100 (Agilent Technologies, Palo Alto, CA, USA), RNA-Seq libraries were prepared using 200 ng total RNA with TruSeq RNA sample prep kit (Illumina). Oligo(dT)-enriched mRNAs were fragmented randomly with fragmentation buffer, followed by the first- and second- strand cDNA synthesis. After a series of terminal repairs, the double-stranded cDNA library was obtained through PCR enrichment and size selection. The prepared cDNA library was sequenced with the Illumina Hiseq 2000 sequencer (Illumina HiSeq 2000 v4 Single-Read 50 bp) after pooling according to its expected data volume and effective concentration. The raw reads were trimmed by using Trim Galore with default parameters. After removing low-quality bases RNAs, the clean data were mapped to mouse genome (GRCm39) using Hisat2 RNA-Seq alignment software, and unique reads were retained to quantify gene expression counts from bam files by using featurecounts. The raw counts were calculated into TPM (Transcripts per million). Data were analyzed and preprocessed in the R environment. Differential expression analysis was performed using R package DESeq2 (P-value < 0.05 and abs| fold-change| greater than 2). Heatmaps and bar graphs were visualized using the R package.

MeRIP-seq

MeRIP-seq was performed at the LC Sciences, LLC, according to the manufacturer’s standard protocol. Total RNA was isolated and purified using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) from WT and FTO KO CD8⁺ T cells isolated from the recipient mice 5 days after LM-OVA infection in the in vivo co-transfer model. m⁶A immunoprecipitation was performed according to standard procedures as described previously [69]. Total RNA integrity was assessed using a Bioanalyzer 2100 (RIN > 7.0) and denaturing agarose gel electrophoresis. Ribosomal RNA was depleted from ~25 μg of total RNA using the Epicentre Ribo-Zero Gold Kit (Illumina). The rRNA-depleted RNA was fragmented using the Magnesium RNA Fragmentation Module (NEB) at 86 °C for 7 min. Fragmented RNA was immunoprecipitated with an m6A-specific antibody (Synaptic Systems) in IP buffer at 4 °C for 2 h. The precipitated RNA was reverse-transcribed into cDNA using SuperScript™ II Reverse Transcriptase (Invitrogen). Second-strand DNA synthesis was performed with E. coli DNA polymerase I, RNase H (NEB), and dUTP. A-tailing and adapter ligation were conducted, followed by size selection using AMPureXP beads. U-labeled second strands were digested with UDG enzyme (NEB), and libraries were PCR-amplified (8 cycles). Final libraries had an average insert size of 300 ± 50 bp. m⁶A-seq was performed at the LC Sciences, LLC on an Illumina Novaseq™ 6000 platform with 2 × 150 bp paired-end sequencing (PE150).

Bioinformatic analysis

For MeRIP-seq data analysis, sequence quality of IP and Input samples were verified using FastQC (https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) and RseQC (http://rseqc.sourceforge.net/). Sequencing reads were mapped to the mouse reference genome (mm39) using HISAT2 (http://daehwankimlab.github.io/hisat2). m⁶A-enriched peaks and diff peaks were identified by exomePeak2 with the corresponding input sample as control, and peaks were annotated by intersection with gene architecture using R package ANNOVAR. HOMER (http://homer.ucsd.edu/homer/motif) was used for de novo followed by localization of the motif with respect to peak summit. StringTie was used to perform expression level for all transcripts and genes from input libraries by calculating FPKM. The differentially expressed transcripts and genes (greater than 1.5-fold, P < 0.05) were selected by the R package. m⁶A peaks were visualized using Integrative Genomics Viewer software.

Differentially expressed genes (DEGs) from RNA-seq data and m⁶A hyper-methylated genes from MeRIP-seq data were subjected to functional annotations. Gene Ontology (GO) analysis was performed using PANTHER (https://geneontology.org/) based on three categories: biological process (BP), cellular component (CC), and molecular function (MF). Hallmark gene sets of mice were extracted from R package msigdbr (version 7.5.1), and GSEA (Gene Set Enrichment Analysis) analysis was performed by enricher function from ClusterProfiler package. The geneset is available at Molecular Signatures Database (MSigDB, https://www.gseamsigdb.org/gsea/downloads.jsp) with either the immunologic signature gene sets (C7) or Hallmark Gene Sets. Enrichment results were visualized using the barplot function for the ggplot2 package (version 3.4.4).

Statistical analysis

Data were presented as means ± SD with n per group and a number of experimental replicates indicated in the respective figure legends. The sample size was not calculated using a statistical approach but instead was determined based on previous experimental observation. Animals were randomly assigned to different experimental groups, and no blinding procedures were applied during the animal studies. The variability between the groups under statistical comparisons is comparable. Two-tailed Student’s t-test was used for all statistical calculations using GraphPad Prism 9 software. For comparison among different groups, one-way or two-way ANOVA was used. All bar graphs include means with error bars to show the distribution of the data. The level of significance is indicated as *P < 0.05, **P < 0.01, and ***P < 0.001.

Data availability

The RNA-seq data and MeRIP-seq generated in this study have been deposited in the Gene Expression Omnibus (GEO) archive under accession GSE270717.

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Acknowledgements

We thank the Mouse Facility of Xi’an Jiaotong University. We also thank Dr. Guohua Zhang from Beckman Coulter, Prof. Baochang Lai and Dr. Xiaofei Wang for flow cytometric technical supports. This work was supported by the National Key Research and Development Program of China grants 2021YFA1100702 (to BZ), National Natural Science Foundation of China grants 82271792 (to LS), 32200727 (to LS), 32170892 (to BZ) and 82071828 (to CS), Innovation Capability Support Program of Shaanxi Province 2024CX-GXPT-45 (to CS), the CAMS Innovation Fund for Medical Sciences (CIFMS) 2023-I2M-2-010 (to LZ), Natural Science Foundation of Shaanxi Province, China 2020JM-065 (to XZ).

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These authors contributed equally: Lina Sun, Tianzhe Zhang, Yao Ge, Zhihong Yao, Yanhong Su.

Authors and Affiliations

Department of Pathogenic Microbiology and Immunology, School of Basic Medical Sciences, Xi’an Jiaotong University, Xi’an, Shaanxi, China
Lina Sun, Tianzhe Zhang, Zhihong Yao, Yanhong Su, Qianhao Wang, Boxiao He, Renyi Ding, Cangang Zhang, Ruonan Liu, Huanxin Ping, Dan Zhang, Lin Shi, Xiaobo Zhou, Chenming Sun & Baojun Zhang
Institute of Infection and Immunity, Translational Medicine Institute, Xi’an Jiaotong University Health Science Center, Xi’an, Shaanxi, China
Lina Sun, Tianzhe Zhang, Yanhong Su, Qianhao Wang, Boxiao He, Renyi Ding, Cangang Zhang, Ruonan Liu, Huanxin Ping, Dan Zhang, Lin Shi, Xiaobo Zhou, Xiaoxuan Jia, Chenming Sun & Baojun Zhang
Key Laboratory of Environment and Genes Related to Diseases, Ministry of Education, Xi’an, Shaanxi, China
Lina Sun, Tianzhe Zhang, Yanhong Su, Qianhao Wang, Chenming Sun & Baojun Zhang
Xi’an Key Laboratory of Immune Related Diseases, Xi’an, Shannxi, China
Lina Sun, Tianzhe Zhang, Yanhong Su, Qianhao Wang, Lin Shi, Xiaobo Zhou, Chenming Sun & Baojun Zhang
Department of Human Anatomy and Histoembryology, School of Basic Medical Sciences, Xi’an Jiaotong University, Xi’an, Shaanxi, China
Yao Ge & Yang Chen
Faculty of Clinical Medicine, Hanzhong Vocational and Technical College, Hanzhong, China
Zhihong Yao
Clinical Teaching and Research Center, School of Nursing, Weinan Vocational and Technical College, Weinan, Shaanxi, China
Linbo Lan
Department of Physiology and Pathophysiology, School of Basic Medical Sciences, Xi’an Jiaotong University Health Science Center, Xi’an, Shaanxi, 710061, PR China; Institute of Neuroscience, Translational Medicine Institute, Xi’an Jiaotong University Health Science Center, Xi’an, Shaanxi, PR China
Lingli Liang
National Key Laboratory of Immunity and Inflammation, Suzhou Institute of Systems Medicine, Chinese Academy of Medical Sciences & Peking Union Medical College, Suzhou, Jiangsu, China
Lianjun Zhang
Key Laboratory of Synthetic Biology Regulatory Elements, Suzhou Institute of Systems Medicine, Chinese Academy of Medical Sciences & Peking Union Medical College, Suzhou, 215123, Jiangsu, China
Lianjun Zhang

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Lina Sun
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Tianzhe Zhang
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Yao Ge
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Zhihong Yao
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Yanhong Su
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Qianhao Wang
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Yang Chen
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Boxiao He
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Renyi Ding
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Cangang Zhang
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Linbo Lan
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Ruonan Liu
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Dan Zhang
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Lin Shi
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Chenming Sun
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Lingli Liang
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Lianjun Zhang
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Contributions

BZ, LS, and CS conceptualized the study and designed experiments. BZ, LS, CS, and XZ contributed to funding acquisition. LS, YG, ZY, YS, QW, CY, XH, RD, LLan, RL, HP, XJ, and DZ performed the experiments and contributed to methodology and data analysis. TZ and CZ analyzed RNA-seq data, and TZ analyzed MeRIP-seq data. L.Liang. provided the Fto^fl/fl mice. Lin Shi and XZ contributed to the scientific discussion. LS and TZ wrote the original draft, and BZ, CS, and LZ provided crucial scientific input and edited the manuscript. All authors have read and approved the article.

Corresponding authors

Correspondence to Chenming Sun, Lingli Liang, Lianjun Zhang or Baojun Zhang.

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Competing interests

The authors declare no competing interests.

Ethics approval and consent to participate

All methods used in this study were performed in accordance with the relevant guidelines and regulations. This study does not include human participants or human materials; therefore, obtaining informed consent for participation is not applicable. The animals used in this study were treated in accordance with the Institutional Animal Care and Use Committee of Xi’an Jiaotong University, Xi’an Center for Disease Control, and Suzhou Institute of Systems Medicine (XJTUAE2022-1160, XJTUAE2022-1161).

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Sun, L., Zhang, T., Ge, Y. et al. FTO controls CD8⁺ T cell survival and effector response by modulating m⁶A methylation of Fas. Cell Death Dis 16, 301 (2025). https://doi.org/10.1038/s41419-025-07606-z

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Received: 20 December 2024
Revised: 19 March 2025
Accepted: 27 March 2025
Published: 15 April 2025
DOI: https://doi.org/10.1038/s41419-025-07606-z