YTHDC1 helps build telomerase by supporting the connection between its protein and RNA parts

Human alveolar epithelial cells (AECs) were purchased from ATCC and cultured at 37°C in 5% CO₂ with Dulbecco's modified Eagle's medium (DMEM, Gibco, USA) supplemented with 10% fetal bovine serum and 100 units/mL penicillin and streptomycin. Cells were transfected with 100 pmol METTL3 siRNA (siMETTL3), YTHDC1 siRNA (siYTHDC1), or negative control siRNA (siNC) using Lipofectamine RNAiMAX transfection reagent (Invitrogen, USA), according to the manufacturer's instructions. The corresponding siRNA sequences are listed in Table S1. Plasmid transfections were performed using Lipofectamine 3000 (Invitrogen) according to standard protocols. To construct stable knockdown cell lines, viruses expressing shCtrl, shMETTL3, or shYTHDC1 shRNA were packaged using a triple plasmid system (psPAX2 and pMD2.G) and transfected into AECs. Positive cells were screened using puromycin (2 ug/mL) to establish stable polyclonal populations.

To construct PHBLV‐shMETTL3‐GFP or PHBLV‐ shYTHDC1‐GFP plasmids, oligos of shRNA listed in Table were purchased and annealed to double‐stranded shRNA, subsequently inserted between BamH I and EcoR I sites of the PHBLV‐U6‐GFP plasmid. S1

For targeted m6A demethylation, AECs were co‐transfected with TERC targeting sgRNAs (TERC‐sgRNA‐1 and sgRNA‐2, listed in Table S1) and the dCas13‐ALKBH5 plasmid using Lipofectamine3000. LacZ‐sgRNA was used as a negative control. Transfected cells were harvested 48 h post‐transfection for downstream analyses.

Western blotting was performed according to standard procedures. Antibodies listed in Table were used to detect the respective bands at 4°C overnight. S1

Total RNA was extracted from cells using the Trizol reagent (Invitrogen) following the manufacturer's protocol. After reverse transcription, cDNA was subjected to real‐time quantitative PCR (Q‐PCR) to detect the expression of the target gene. The primer pairs used in this study are provided in Table . S1

The RNAscope Multiplex Fluorescent Assay (ACD Bio, USA) was performed according to the manufacturer's protocol. Briefly, AECs were initially seeded into confocal Petri dishes and fixed with 10% neutral formalin. Subsequently, the cells were subjected to digestion with RNAscope Protease III for 10 min, and the digestion was terminated by washing with PBS. This was followed by probe hybridization, signal cascade amplification with a Z probe and multi‐channel fluorescent staining to label the endogenous TERC RNA. The cells were then blocked with 5% BSA for 1 h at room temperature. Subsequently, the cells were incubated with a primary antibody against YTHDC1 overnight at 4°C. After washing three times with PBST (containing 0.1% Tween 20), a fluorescent secondary antibody (1:200) with the corresponding properties was added and incubated for 2 h at room temperature. The cells were washed again with PBST, followed by counterstaining with a blocking agent containing 4′,6‐diamidino‐2‐phenylindole (DAPI) to visualize nuclei. The co‐localization of YTHDC1 and TERC RNA within the cells was then visualized and imaged using confocal fluorescence microscopy.

TRAP and IP‐TRAP assays were performed as previously described (Tang et al. 2018). Briefly, the cells were collected and lysed using NP‐40 lysis buffer (150 mM NaCl, 10 mM Tris–HCl pH 8.0, 1 mM MgCl₂, 1 mM EDTA, 1% NP‐40, 0.25 mM Sodium deoxycholate, 5 mM β‐ME, and protein inhibitor) at a concentration of 1000 cells/μL. Two microliters of lysate were subjected to the extension reaction at 37°C incubation and followed by PCR amplification with TS, ACX, NT, and TSNT primers. The sequences are listed in Table S1. The products were separated on non‐denatured polyacrylamide gels (12%) and stained with SYBR Safe (Invitrogen). The bands were visualized using an Odyssey system. The intensity of the bands was quantified using image J and the intensity of the telomere bands was calculated against that of the internal control. For IP‐TRAP, the cells were lysed in lysis buffer (0.01% NP‐40,10 mM Tris pH 7.5, 50 mM KCl, 5 mM MgCl₂, 2 mM dithiothreitol, 20% glycerol, and protease inhibitors) at a concentration of 100,000 cells/mL and sonicated at 50 J/W‐s. After centrifugation at 16,000 × g for 30 min, the supernatant was collected and used for immunoprecipitation with the corresponding antibody at 4°C for 4–6 h. The mixture was then added to protein A/G agarose beads (Invitrogen) and incubated at 4°C for 1 h. After washing with lysis buffer four times, the beads were resuspended in lysis buffer and subjected to a TRAP assay to detect telomerase activity.

A telomere‐restricted fragment length assay was performed using a commercial kit (cat. no. 12 209136 001; Roche, Switzerland) according to the manufacturer's instructions. Briefly, genomic DNA was digested using HinfI and RsaI and then separated using a 0.8% agarose gel. After fixing, denaturing, and neutralizing, the gel was transferred to a nylon membrane. Subsequently, the membrane was UV‐crosslinked, hybridized with a Digoxin‐labeled TTAGGG probe, and incubated with an alkaline phosphatase‐coupled anti‐digoxin antibody. After addition of the reaction substrate, the membrane was exposed to a chemiluminescence apparatus (Thermo Fisher, USA). Mean TRF length has been defined according to the following formula:TRF¯=∑ODi∑ODi/Liwhere OD_i is the chemiluminescent signal and L_i is the length of the TRF at position i. The calculation considered the higher signal intensity from larger TRFs due to multiple hybridizations of the telomere‐specific hybridization probe.

RNA pull‐down experiments were performed as previously described (Liu et al. 2022) with the following modifications. Biotinylated TERC RNA probes (wild‐type and mutant variants) were synthesized in vitro using the TranscriptAid T7 High‐Yield Transcription Kit (Thermo Fisher). AECs lysates were freshly prepared using RIPA lysis buffer with proteinase inhibitor. The freshly prepared cell lysates (500 μg total protein) were incubated with the indicated biotinylated probes (500 ng) and 100 μM S‐adenosylmethionine (SAM, NEB, USA) in TENT buffer (20 mM Tris–HCl pH = 7.5, 250 mM NaCl, 1 mM EDTA, 0.5% TritonX‐100) supplemented with RNase inhibitor (0.5 U/μL, NEB) and protease inhibitor cocktail (Roche) at 37°C for 60 min with gentle rotation. Subsequently, the reaction mixture was isolated using Dynabeads M‐280 Streptavidin (Thermo Fisher), and the proteins were subjected to western blotting. For dot‐blot analysis, the beads‐RNA‐Protein complexes were subjected to RNA isolation using Trizol, and the RNA was denatured at 65°C for 10 min and spotted onto positively charged nylon membranes. M6A methylation level of TERC probes was assessed using m6A antibody and visualized by chemiluminescence. Subsequently, the membrane was stripped and subjected to streptavidin‐HRP antibody to detect the loading control.

For UV‐crosslinking RNA immunoprecipitation (RIP), cells were exposed to UVC (400 mJ/cm²) for crosslinking, followed by lysis using RIP Lysis buffer (50 mM HEPES pH 7.5, 150 mM KCl, 2 mM EDTA, and 1% NP‐40) supplemented with RNase inhibitor (0.5 U/μL, NEB) and protease inhibitor cocktail (Roche). Lysates were clarified by centrifugation (16,000 × g, 15 min at 4°C) and pre‐cleared with Protein A/G beads (Thermo fisher). Immunoprecipitations were performed overnight at 4°C with 3–5 μg of target‐specific antibodies or species‐matched IgG controls, followed by incubation with Protein A/G magnetic beads for 4 h. The IP materials were washed with IP wash buffer (100 mM Tris–HCl pH 7.4, 500 mM LiCl, 0.1% TritonX‐100, and 1 mM DTT) for three times, and RNA was extracted using Trizol (Vazyme, China) and assessed by RT‐qPCR. Enrichment was calculated as fold‐change relative to IgG controls after normalization to input RNA.

The meRIP‐qPCR assay was performed according to the standard protocol. Briefly, 10–20 μg RNA was fragmented into 200–300 nt fragments via a 15‐min incubation at 70°C in fragmentation buffer (10 mM ZnCl₂, and 10 mM Tris–HCl pH 7.0). For exogenous TERC methylation analysis, RNA fragmentation was omitted to preserve full‐length transcripts. Thereafter, the fragmented RNA was incubated with 2–3 μg m6A antibody (Synaptic Systems) at 4°C overnight, and then the complex was incubated with DynaGreen Protein A/G magnetic beads (Thermo Fisher) at 4°C for 1 h. After washing with meRIP lysis buffer (150 mM NaCl, 0.1% NP‐40, and 10 mM Tris–HCl), the bound RNAs were recovered by proteinase K digestion, phenol–chloroform extraction, and ethanol preparation. One‐tenth of the fragmented RNA was used as an input control. The input or bound RNA was subjected to reverse transcription and subsequent qPCR. Relative enrichment was calculated as fold‐change relative to IgG controls after normalization to input RNA.

Cytoplasmic and nuclear proteins were extracted using a commercial subcellular protein fractionation kit (Beyotime Biotechnology). Briefly, cells were harvested, washed with ice‐cold PBS, and lysed in Cytoplasmic Lysis Buffer on ice for 15 min. The lysates were centrifuged at 12,000 × g for 5 min at 4°C. The supernatant (cytoplasmic fraction) was collected. The resulting pellet (nuclei) was further extracted with Nuclear Extraction Buffer containing on ice for 30 min. The nuclear extract was clarified by centrifugation at 12,000 × g for 10 min. Protein concentrations were determined using the BCA assay. The purity of the fractions was verified by Western blotting using antibodies against GAPDH (cytoplasmic marker) and Histone 3 (nuclear marker). RNA and proteins were extracted by Trizol (Invitrogen), according to the manufacturer's instructions. Quantification of the TERC in cytoplasmic and nuclear fractions was performed with specific primers.

Cells were lysed in IP lysis buffer (150 mM NaCl, 25 mM Tris–HCl pH 7.5, 1 mM EDTA, 0.5% NP‐40, 5% glycerol, and protein inhibitor). After centrifugation (16,000 × g, 30 min, 4°C), 500 μg lysate was immunoprecipitated overnight at 4°C with 3 μg antibody and Protein A/G beads. For RNase treatment, lysates were incubated with RNase A/T1 (100 μg/mL and 1000 U/mL respectively) at 37°C for 30 min prior to IP. The mixture was washed four times with the IP lysis buffer. The beads were resuspended in protein‐loading buffer and subjected to western blotting analysis.

Cells were fixed with 4% paraformaldehyde for 15 min and permeabilized in PBST (0.25% Triton X‐100) for 10 min at room temperature. After washing with PBS, the cells were blocked with 5% BSA for 30 min and incubated with the following antibodies: Flag (1:200; Sigma‐Aldrich, Germany), YTHDF1 (1:200; Proteintech, USA), YTHDF2 (1:200; Proteintech), YTHDF3 (1:200; Santa Cruz Biotechnology, USA), YTHDC1 (1:200; Cell Signaling Technology, USA), YTHDC2 (1:200; Proteintech), or coilin (1:200; Proteintech) overnight at 4°C. After washing three times with PBS, the cells were incubated with secondary antibodies (FITC/TRITC, 1:200, 1 h), followed by DAPI counterstaining. Imaging was performed on a Leica DMi8 (Germany) using standardized parameters.

Cell proliferation and cell cycle assays were conducted as previously described (Cheng et al. 2021). EdU incorporation was performed using a kit (RiboBio, Guangzhou, China) following the manufacturer's instructions. SA‐β‐gal activity was assessed using a senescence β‐Galactosidase Staining Kit (GENMED Scientifics Inc., USA).

To establish the presence of m6A modifications on TERC, we preliminarily performed the m6A‐RNA immunoprecipitation (MeRIP) assays. TERC showed significant enrichment with m6A antibody compared to IgG control (Figure 1A), similar to the positive control ADAM19 mRNA. Although previous transcriptome‐wide studies suggested A435 as the primary m6A site, public nanopore sequencing data (DirectRMDB) indicated A111 and A241 as potential sites. To resolve this discrepancy, we designed experiments to screen and identify the actual methylation sites. Since N6‐Methyladenosine preferentially occurs at the consensus motif DRACH (R is A or G; H is U, A, or C) (Nombela et al. 2021), we used the deepSRAMP tool (A sequence‐based m6A site predictor by a combined deep learning framework) to screen and speculate on the potential m6A methylation site (Figure 1B). Based on the computational screening, we generated wild‐type (WT) and mutant TERC constructs (Figure 1C) by subcloning the corresponding sequences into the pcDNA3.1(+) plasmid. These plasmids were transfected into AECs, and after 48 h, total RNA was extracted and subjected to MeRIP‐qPCR to assess the m6A methylation levels in exogenous TERC. MeRIP analysis revealed a significant reduction in m6A enrichment upon mutation of either A111 or A435 (Figure 1D), confirming these as major methylation sites. Strikingly, the double mutant (A111G/A435G, M2/M6) exhibited an even more pronounced decrease in m6A methylation (Figure S1A,B). Meanwhile, METTL3 deficiency dramatically decreased the enrichment of TERC (Figure 1E), confirming METTL3‐dependent methylation. Notably, mutations near the methylation sites (C108T, r.110_113delGACU, r.378_451delGACU) are closely related to pulmonary fibrosis, although their mechanistic contribution remains unclear. We engineered TERC transcripts harboring the disease‐associated variants (Figure S1A). Strikingly, these mutants exhibited a significant reduction in m6A methylation levels (Figure 1F), implicating m6A dysregulation as a potential contributor to disease pathogenesis. Interestingly, the methylation sites are highly conserved across mammals (Figure S1C), suggesting that m6A methylation at these positions may play a critical and evolutionarily conserved role in TERC function. Collectively, these findings clearly state that METTL3‐mediated m6A modification occurs in TERC RNA with A111 and A435 serving as the primary methylation sites.

While a prior study reported that ALKBH5 overexpression reduces telomerase activity by 22% (Han et al. 2020), the mechanistic basis for this observation remained unclear. To rigorously investigate the role of m6A methylation in telomerase function, we generated AECs with stable METTL3 knockdown (Figure S1D). Strikingly, telomerase activity decreased by 57.5% in METTL3‐depleted cells compared to controls, as measured by TRAP assay (Figure 2A,B). To further dissect the role of TERC m6A methylation, we utilized U2OS cells—a well‐established reconstitution system lacking endogenous TERT and TERC (Tang et al. 2018). When co‐transfected with Flag‐TERT and either wild‐type or methylation‐deficient mutant TERC (M2 [A111], M6 [A435], or double mutant), we observed a moderate reduction in telomerase activity with single‐site mutants (A111 or A435) and a dramatic impairment in the double mutant condition (Figure 2C,D). This dose‐dependent effect aligns with the progressive reduction in m6A methylation observed in these mutants (Figure 1D, Figure S1B). Expression levels of TERC and Flag‐TERT were confirmed by real‐time qPCR and Western blotting (Figure S2A,B). To rule out the potential impact of mutagenesis on TERC structure, we employed a dCasRx‐alkbh5 RNA editing system to selectively erase m6A methylation on TERC. The results suggested that the TERC‐targeting sgRNA‐guided dCasRx‐ALKBH5 system efficiently demethylated the TERC, but not ADAM19 (Figure S2C,D). Strikingly, TERC demethylation recapitulated the telomerase activity defect observed in methylation‐site mutants (Figure 2E,F), confirming that the phenotype stems from m6A loss rather than structural perturbations. To ascertain the requirement for METTL3's catalytic activity, we used its specific inhibitor, STM2457, which is currently in clinical trials. STM2457 treatment potently suppressed global RNA m6A methylation, as measured by RNA dot blot, while METTL3 protein levels were largely unaffected (Figure S2E). Consequently, telomerase activity was severely impaired (Figure 2G,H). To assess long‐term functional consequences, we performed Southern blotting in METTL3‐depleted cells, which exhibited a 30% reduction in mean telomere length (Figure 2I,J). Meanwhile, we performed longitudinal telomere length analysis in AECs transfected with control (pcDNA), wild‐type TERC, or mutant TERC plasmids every 3 days, with samples collected on the indicated days for Southern blot analysis. Remarkably, cells expressing wild‐type TERC exhibited progressive telomere elongation, while those with mutant TERC showed minimal length changes (Figure S2F,G). Collectively, these results provide compelling evidence that m6A modification of TERC is critical for maintaining telomerase activity and telomere length homeostasis.

To elucidate the mechanism by which TERC m6A modifications regulate telomerase activity, we first examined TERC and TERT expression levels, finding only minor alterations upon METTL3 knockdown (Figure 3A). Furthermore, to determine whether METTL3 knockdown affects TERC localization, we used RNAscope and cellular fractionation assays. RNAscope analysis revealed that TERC was primarily localized in the nucleus, and this distribution pattern remained largely unaltered upon METTL3 silencing (Figure 3B). This observation was corroborated by detecting TERC in separated cytoplasmic and nuclear fractions (Figure S3A,B). However, immunofluorescence revealed slightly reduced co‐localization of TERT with Cajal bodies (marked by Coilin) following METTL3 depletion (Figure S3C), hinting that defective holoenzyme assembly. To further confirm our hypothesis, we conducted immunoprecipitation assays and modified RNA pulldown. The TERC level in Flag‐TERT‐enriched materials decreased upon METTL3 knockdown (Figure 3C). Meanwhile, RNA pulldown assays further demonstrated weakened TERT binding to m6A‐deficient TERC upon METTL3 knockdown, corroborated by reduced m6A signals in biotin‐labeled TERC via RNA dot blot (Figure 3D). Consistent with these findings, RNA pulldown with biotin‐labeled wild‐type or mutant TERC (M2/M6), the association of TERT with mutant TERC was dramatically decreased compared with the wild‐type TERC (Figure S3B). Additionally, IP‐TRAP assays were performed as previously described (Tang et al. 2018; Ford et al. 2000). METTL3 knockdown resulted in decreased enrichment of the telomerase holoenzyme immunoprecipitated with Flag‐TERT (Figure 3E), indicating that m6A methylation facilitates the TERT‐TERC interaction. To compare the catalytic activity of m6A‐modified versus unmethylated telomerase, we quantified its specific activity by normalizing the TRAP signal from the immunoprecipitated complexes to the levels of co‐precipitated TERC. The normalized activity of telomerase was also decreased upon METTL3 depletion, implying that the holoenzyme containing m6A‐modified TERC possesses higher intrinsic catalytic activity (Figure 3F,G, Figure S3E,F). Furthermore, the TERC‐targeting sgRNA‐guided dCasRx‐ALKBH5 system significantly impaired the interaction between Flag‐TERT and TERC (Figure 3H). In summary, our findings imply that m6A modification of TERC maintains telomerase activity by facilitating the assembly of TERT and m6A‐methylated TERC.

Previous studies have established that the assembly and activity of the telomerase holoenzyme depend on dyskerin, GAR1, NOP10, and NHP2, which bind directly to TERC (Schmidt and Cech 2015). However, we observed that neither the expression levels of these proteins nor their association with TERC was affected by METTL3 knockdown (Figure S3G,H). We therefore turned our attention to YTH domain‐containing reader proteins, which are known to mediate m6A‐dependent RNA–protein interactions (Boulias and Greer 2023; Zhao et al. 2017), as potential alternative regulators. Bioinformatics analysis using POSTAR3 (Zhao et al. 2022) predicted YTHDC1 as a putative TERC‐binding protein (Table S2). RNA pulldown and immunoprecipitation (RIP) assays confirmed YTHDC1 as the predominant m6A reader protein interacting with TERC (Figure 4A and Figure S4A). Furthermore, IP‐TRAP assays showed that, similar to Flag‐TERT, YTHDC1 co‐precipitated with the telomerase holoenzyme (Figure S4B,C), corroborating that YTHDC1 may bind to telomerase RNA TERC. Additionally, immunofluorescence and RNAscope demonstrated YTHDC1 co‐localization with both coilin (a Cajal body marker) and TERC itself (Figure 4B and Figure S4D). Unsurprisingly, RIP assays revealed that METTL3 knockdown reduced YTHDC1 binding to TERC (Figure 4C). RNA pulldown experiments using biotin‐labeled TERC transcripts (wild‐type, M2 [A111], M6 [A435], or double mutant) further showed impaired YTHDC1 binding with TERC upon mutation of the methylation sites (Figure 4D). Notably, single‐site mutants exhibited partial reductions in binding, while the double mutant showed near‐complete loss of YTHDC1 interaction. Therefore, these data indicate that YTHDC1 participates in telomerase holoenzyme assembly by binding to TERC, a process that is influenced by the m6A methylation status of TERC.

Based on the finding that TERC m6A modification facilitates TERT‐TERC association, we hypothesized that YTHDC1 mediates this interaction. To test this, we performed co‐immunoprecipitation (co‐IP) assays using antibodies against YTHDC1 or TERT. As shown in Figure 4E,F, western blotting revealed that TERT associated with YTHDC1. Importantly, this interaction persisted after RNase A/T1 treatment (Figure 4G), demonstrating it occurs in an RNA‐independent manner. Consistent with this finding, YTHDC1 also co‐immunoprecipitated with Flag‐tagged TERT in TERC‐deficient U2OS cells (Figure S5A), further demonstrating that the YTHDC1‐TERT interaction occurs independently of TERC. As reported previously, YTHDC1 comprises six regions and one classic YTH domain responsible for binding m6A‐modified RNA (Dattilo et al. 2023). To identify the TERT‐binding domain of YTHDC1, we generated a series of Flag‐tagged truncation mutants encompassing the six structural regions of YTHDC1 (Figure S5B). Co‐IP experiments showed that wild‐type YTHDC1, but not deletion mutants Δ4, Δ5, or Δ6, maintained the ability to bind TERT (Figure S5C). This identified amino acids 345‐648 (encompassing the YTH and R/P domains) as the critical interaction domain. Intriguingly, a mutant YTHDC1 lacking the R/P domain also exhibited weakened TERT binding while maintaining its association with TERC (Figure 4H and Figure S5D). Additionally, deletion of the YTH domain severely compromised the interaction with TERC (Figure S5E), confirming that the YTH domain is essential for recognizing m6A‐modified TERC. Furthermore, the TERT‐YTHDC1 interaction was unaffected by METTL14 knockdown (Figure S5F). Collectively, these data demonstrate that YTHDC1 directly binds TERT through its C‐terminal domain (amino acids 345‐648) in a TERC‐independent manner.

To dissect YTHDC1's functional role, we first assessed its effect on telomerase activity. TRAP assays revealed a striking 65.6% reduction in telomerase activity upon YTHDC1 knockdown (Figure 5A,B), accompanied by progressive telomere shortening in YTHDC1‐deficient cells (Figure 5C and Figure S6A). Crucially, similar to METTL3 depletion, YTHDC1 knockdown did not alter TERC/TERT transcript or TERT protein levels, hinting that YTHDC1 might also affect telomerase assembly (Figure 5D,G). Supporting this, IP‐TRAP assays showed significantly diminished telomerase enrichment by Flag‐TERT in YTHDC1‐deficient cells (Figure S6B,C). Furthermore, RNA immunoprecipitation and RNA pulldown experiments demonstrated that YTHDC1 knockdown weakened the TERT‐TERC association (Figure 5E,F), phenocopying the effect of METTL3 depletion. Notably, although YTHDC1 knockdown did not alter the expression of the core telomerase‐associated proteins dyskerin, GAR1, NOP10, or NHP2, it significantly impaired the association between TERT and these TERC‐binding factors (Figure S6D). These results further support the conclusion that YTHDC1 influences telomerase holoenzyme assembly. To determine whether YTHDC1's effect on telomerase activity depends on m6A methylation, we silenced the expression of YTHDC1, METTL3,or both. Western blotting confirmed that TERT protein levels remained unchanged under all conditions (Figure 5G). Strikingly, while YTHDC1 depletion alone reduced telomerase activity, its knockdown in METTL3‐deficient cells failed to cause a further suppression (lane 2 vs. lane 4, Figure 5H,I). This indicates that YTHDC1's regulation of telomerase is strictly m6A‐dependent. Taken together, these data support the hypothesis that YTHDC1 promotes telomerase holoenzyme assembly by strengthening the combination of m6A‐methylation TERC with TERT.

Given the established role of telomerase activity in maintaining cellular proliferation and preventing senescence (Blackburn et al. 2015; Shawi and Autexier 2008; Schmidt and Cech 2015), we investigated whether YTHDC1‐mediated telomerase assembly impacts these processes in AECs. To this end, we performed rescue experiments in YTHDC1‐deficient AECs by continuously transfecting plasmids expressing either wild‐type YTHDC1 or a truncated mutant (Flag‐YTHDC1‐ΔR/P, lacking the critical TERT‐binding domain) every 3 days until population doubling 60 (PD60). Since telomere dysfunction or telomerase deficiency is known to activate DNA damage signaling, we first assessed the levels of γ‐H2AX and p‐CHK2, both markers of DNA damage. YTHDC1 knockdown increased the expression of these markers. However, this effect was reversed by expression of wild‐type YTHDC1, but not the truncated mutant YTHDC1 (Figure 6A and Figure S7A). Consistently, expression of wild‐type YTHDC1, but not the mutant, rescued the loss of telomerase activity and telomere length erosion (Figure S7B,C and Figure 6B,C). We next examined cellular proliferation and senescence using EdU incorporation assays (Figure 6D,E), ki67 immunofluorescence (Figure S7D,E), senescence‐associated β‐galactosidase assays (Figure 6F,G), senescence‐associated secretory phenotype (SASP) evaluation (Figure 6H), and cell cycle analysis (Figure S7F). As expected, YTHDC1 deficiency induced S phase arrest, decreased EdU incorporation, lower ki67‐positive rates, and accelerated cellular senescence (increased SA‐β‐gal positive staining and elevated SASP level). Crucially, these defects were rescued by wild‐type YTHDC1 expression, whereas the R/P domain‐truncated mutant failed to restore the proliferation or delay senescence. Altogether, these findings demonstrate that YTHDC1 promotes telomerase assembly by strengthening the TERT–TERC interaction, which in turn is critical for sustaining proliferation and suppressing senescence in AECs.