FTO suppresses DNA repair by inhibiting PARP1

Zhu, Tianyi; Tan, Jing Zhi Anson; Zhang, Lingrui; Huang, He; Das, Sooraj S.; Cheng, Flora; Padmanabhan, Pranesh; Jones, Mathew J. K.; Lee, Mihwa; Lee, Albert; Widagdo, Jocelyn; Anggono, Victor

doi:10.1038/s41467-025-58309-0

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Published: 25 March 2025

FTO suppresses DNA repair by inhibiting PARP1

Nature Communications volume 16, Article number: 2925 (2025) Cite this article

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Abstract

Maintaining genomic integrity and faithful transmission of genetic information is essential for the survival and proliferation of cells and organisms. DNA damage, which threatens the integrity of the genome, is rapidly sensed and repaired by mechanisms collectively known as the DNA damage response. The RNA demethylase FTO has been implicated in this process; however, the underlying mechanism by which FTO regulates DNA repair remains unclear. Here, we use an unbiased quantitative proteomic approach to identify the proximal interactome of endogenous FTO protein. Our results demonstrate a direct interaction with the DNA damage sensor protein PARP1, which dissociates upon ultraviolet stimulation. FTO inhibits PARP1 catalytic activity and controls its clustering in the nucleolus. Loss of FTO enhances PARP1 enzymatic activity and the rate of PARP1 recruitment to DNA damage sites, accelerating DNA repair and promoting cell survival. Interestingly, FTO regulates PARP1 function and DNA damage response independent of its catalytic activity. We conclude that FTO is an endogenous negative regulator of PARP1 and the DNA damage response in cells beyond its role as an RNA demethylase.

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Introduction

Human cells experience a high frequency of DNA lesions induced by endogenous and exogenous insults that must be rapidly repaired to avoid the accumulation of catastrophic mutations and genomic instability, which can lead to cell death or the development of diseases such as cancer^1,2. To ensure genomic integrity, cells have evolved versatile mechanisms, collectively known as the DNA damage response (DDR), that sense and repair DNA lesions. The recruitment of poly(ADP-ribose) polymerase 1 (PARP1) to DNA damage sites represents one of the earliest events in multiple DNA repair pathways, including single-strand break repair, base excision repair, nucleotide excision repair and double-strand break repair through homologous recombination or non-homologous end-joining pathways (NHEJ)^3,4. It is well established that active PARP1 catalyzes the post-translational attachment of poly(ADP-ribose) (PAR) onto itself and essential DNA repair proteins, recruiting them to the damaged sites. However, the mechanisms and factors that restrict PARP1 activation in undamaged conditions remain poorly understood.

Fat mass and obesity-associated (FTO) protein is an Fe²⁺/α-ketoglutarate-dependent dioxygenase primarily known to demethylate multiple RNA species at the N⁶ (m⁶A) or N¹ (m¹A) position of adenosine and the N⁶, 2’-O-dimethyladenosine (m⁶A_m)^5,6,7. FTO expression is misregulated in many types of cancers⁸. Upregulation of FTO, which leads to downregulation of m⁶A marks in its target mRNAs such as RARA, MYC, ASB2, PARP1 and CEBPA, contributes to tumorigenesis by promoting cancer stem-cell renewal and differentiation, cell growth, migration, and invasion in most cancer types^8,9,10,11,12. Small molecule inhibitors targeting FTO are therefore of clinical importance for the treatment of acute myeloid leukemia, glioblastoma, breast and pancreatic cancers^13,14,15. However, FTO can also exhibit a tumor suppressor function in lung, breast, liver and cervical cancers by modulating gene expression in an m⁶A-dependent manner^8,16,17. Thus, a better understanding of the cellular function of FTO in DDR may provide mechanistic insight into the contribution of FTO in maintaining genomic stability in cells.

N⁶-methyladenosine (m⁶A) signaling has recently emerged as a critical regulator of the DDR^18,19,20,21. It is the most prevalent post-transcriptional RNA modification in eukaryotes, which is deposited co-transcriptionally by the m⁶A writer complex containing the catalytically active methyltransferase-like 3 (METTL3) and reversed by the m⁶A RNA demethylases, FTO or AlkB homolog 5 (ALKBH5)^{6,22,23,24,25}. Upon ultraviolet (UV) light exposure, METTL3 is recruited to the DNA damage site and increases the m⁶A level in a PARP1-dependent manner, which through an unknown mechanism then facilitates the recruitment of polymerase κ (Polκ) for efficient DNA repair¹⁸. Although the role of FTO in the DDR pathway has recently been indicated in a few studies^26,27,28, the exact mechanisms and functions of FTO in these processes are unclear.

Results

FTO loss promotes UV-induced DNA repair

To determine the role of FTO in the UV-induced DDR, we quantified the levels of cyclobutene pyrimidine dimers (CPDs), the dominant DNA lesions caused by UV exposure²⁹, in two independent lines of FTO knockout (KO) HeLa cells (Supplementary Fig. 1a)³⁰. Our results revealed that FTO KO cells exhibited significant reductions in CPD levels (Fig. 1a, b). Notably, there were no significant differences in the level of CPDs formed shortly after UV exposure between wild-type (WT) and FTO KO cells (Supplementary Fig. 1b–e), indicating that loss of FTO promotes CPD removal in HeLa cells. Next, we examined the phosphorylation of the histone variant H2A.X at Ser-139 (γH2A.X), a marker of DNA double-strand breaks³¹. Consistent with our CPD data, we observed lower UV-induced γH2A.X in FTO KO cells than in WT HeLa cells by immunostaining (Fig. 1c, d) and western blot assays (Fig. 1e, f). The reductions in DNA damage markers in FTO KO HeLa cells are associated with a more robust cell survival after 5–25 J UV irradiation than WT cells (Fig. 1g, h). To validate our findings, we generated FTO KO in two distinct cell lines, namely U2OS (one clone) and HCT116 (two clones), and determined the impact of FTO depletion on DDR. Corroborating the findings in HeLa cells, our results showed that FTO KO U2OS and HCT116 cells had reduced CPD and γH2A.X levels following UV treatment, resulting in higher percentages of cell survival after UV irradiation (Supplementary Fig. 2). Collectively, these findings demonstrate that FTO suppresses the repair of UV-induced DNA lesions and cellular resistance to UV.

**Fig. 1: Loss of FTO expression promotes DNA repairs and cell survival following UV exposure.**

Identification of endogenous FTO proximal proteome

To determine the underlying mechanisms of FTO function in DDR, we performed an unbiased screen to identify FTO proximal interactome in HeLa cells by combining the biotinylation by antibody recognition (BAR) labeling technique with stable isotope labeling with amino acids in cell culture (SILAC)-based mass spectrometry (MS) analysis (Supplementary Fig. 3a, b)³². This method does not require overexpression of FTO protein tagged with a biotin ligase or peroxidase, which enabled us to capture the endogenous FTO proximal interactome recognized only by specific antibodies against FTO. In WT cells, FTO BAR generated significantly stronger biotin labeling as revealed by immunostaining with Alexa-488-conjugated streptavidin than that observed in FTO KO HeLa cells (Supplementary Fig. 3c). Moreover, only low to negligible levels of biotin labeling could be detected in conditions where no primary antibodies, biotin phenol or hydrogen peroxide (H₂O₂) were added to the reactions (Supplementary Fig. 3c), validating the specificity of the assay. The normalized MS data revealed 243 proteins in the FTO proximal proteome (adjusted P-value < 0.05), which had significantly higher levels of biotinylation in WT than FTO KO cells (Fig. 2a, b and Supplementary Data 1). Many of these proteins function in mRNA splicing (DDX17, SFPQ, SRSF6, PUF60 and WTAP) and the spliceosome complex (snRNP70, PRPFs and HNRNPs) (Fig. 2c and Supplementary Data 2), consistent with the established role of FTO in these processes^33,34,35. In addition, we also identified proteins involved in the ribonucleoprotein biogenesis complex in the FTO proximal proteome (multiple members of the DDX, RPL and RPS family of proteins), indicating a cellular function of FTO in ribosome biogenesis. We also identified several FTO interactors that had been reported in previous proteomic studies^27,36, including DDX17, KIFC1, HNRNPH1 and SFPQ (Supplementary Fig. 4a), confirming the ability of our assay to isolate known interacting proteins.

**Fig. 2: The proximal proteome of endogenous FTO protein.**

Despite being a critical component in RNA processing, we found many FTO proximal interacting proteins are involved in DNA replication, telomere maintenance, DNA repair, chromosomal organization and nuclear division, suggesting a prominent role of FTO in regulating the stability and integrity of the nuclear genome in cells (Fig. 2d, e). Many of these FTO proximity interactors can function in multiple cellular pathways that maintain genomic integrity, such as members of the MCM replicative helicases, XRCC5 (Ku80), PRKDC (DNA-PK), ERCC4 and PARP1. Although FTO itself was not identified in the list of 585 PARP1 proximal-interacting proteins³⁷, we found approximately 37% of FTO proximity interactors (89 out of 243 proteins) in the PARP1-proximal proteome (Supplementary Fig. 4b). Many of them are nuclear proteins with nucleic acid binding and helicase activities, indicating the functional interaction between these two proteins.

FTO interacts with PARP1 and regulates its clustering in cells

To validate the interaction between FTO and PARP1, we performed BAR assays with anti-FTO antibodies followed by western blotting. We could pull down biotinylated PARP1 and its interacting proteins that form the core component of the NHEJ machinery, DNA-PK and Ku80 in WT but not in FTO KO cells (Fig. 3a, b)^38,39. Proximity-based ligation assays (PLA) confirmed the interaction between endogenous FTO and PARP1. FTO-PARP1 PLA puncta were observed in the nuclei of WT cells but were significantly reduced in two independent lines of FTO KO cells (Fig. 3c, d). Furthermore, in vitro binding assays revealed that recombinant Flag-PARP1 protein only co-precipitated on a nickel column in the presence of His-FTO (Fig. 3e, f), demonstrating that PARP1 can directly interact with FTO.

**Fig. 3: FTO controls PARP1 clustering in the nucleus.**

UV exposure in cells could modulate the strength of the interaction between FTO and PARP1. Interestingly, the FTO-BAR assays revealed a significant reduction in biotinylated PARP1, Ku80 and DNA-PK 4 min post-UV irradiation (Fig. 3a, b). We also observed a similar decrease in FTO-PARP1 PLA puncta in cells (Fig. 3g, h), suggesting that UV exposure weakens the interaction between FTO and PARP1. The reduced FTO-PARP1 interaction was not due to changes in PARP1 expression because FTO knockout or overexpression did not affect the abundance of PARP1 protein (Supplementary Fig. 5a, b). Furthermore, there were no significant changes in the levels of PARP1 mRNA in FTO KO compared to wild-type HeLa cells (Supplementary Fig. 5c). Next, we determined the localization of FTO and PARP1 using super-resolved structured illumination microscopy. As expected, high-intensity clusters of PARP1 molecules were localized within the nucleoli of HeLa cells (Fig. 3i). Conversely, FTO had a more even distribution across the nucleus although a higher level of labeling could be observed close to PARP1 within the nucleoli (Fig. 3i). Consistent with the results obtained with the BAR and PLA assays, we found that the nearest distance between FTO and PARP1 molecules increased post-UV treatment, further confirming the weakening of the FTO-PARP1 interaction in cells (Fig. 3i, j and Supplementary Fig. 6a). Based on the high-resolution images, we also observed a dispersal of the high-intensity PARP1 clusters that localized to the nucleoli following UV treatment (Fig. 3i, k). This was evidenced by the reduced number of high-intensity PARP1 clusters as well as the wider area occupied by the dispersed PARP1 clusters in response to UV treatment (Fig. 3k–m). This phenomenon was accompanied by a significant decrease in the volume of individual PARP1 puncta and an alteration in their three-dimensional shape (Fig. 3n and Supplementary Fig. 6b). Strikingly, the loss of FTO expression led to PARP1 dispersal into smaller clusters that occupied larger surface areas (Fig. 3k–n). UV exposure did not induce further dispersal of PARP1 molecules in FTO KO cells (Fig. 3k–n). Moreover, UV treatment only had minor effects on the volume and three-dimensional shape of FTO puncta (Supplementary Fig. 6c, d). Collectively, these data indicate that FTO plays a structural role in maintaining PARP1 clusters in cells.

FTO negatively regulates PARP1 function

Our results predicted that UV-induced dissociation of the FTO-PARP1 interaction allows PARP1 to disperse and be efficiently recruited to DNA damage sites. To test this hypothesis, we directly visualized the recruitment of GFP-PARP1 to the sites of DNA lesions elicited by UVA micro-irradiation in live cells. UV micro-irradiation caused rapid association of GFP-PARP1 to the DNA damage sites with a time constant (τ) of ~8.1 s (Fig. 4a–d). By contrast, HeLa cells lacking FTO exhibited a significantly faster rate of GFP-PARP1 recruitment (τ ~ 5.8 s) in two independent KO lines without affecting the amount of GFP-PARP1 bound to damaged DNA (Fig. 4a–e). Although overexpression of myc-FTO did not slow the recruitment rate, the abundance of GFP-PARP1 at sites of DNA damage was dramatically reduced (Supplementary Fig. 7a–d).

**Fig. 4: Loss of FTO enhances PARP1 function.**

The binding of PARP1 to DNA strand breaks results in a conformational change that dramatically enhances its enzymatic activity and catalyzes rapid poly(ADP)ribosylation (PARylation) of itself (auto-PARylation) and of many other nuclear proteins^40,41. Consistent with the faster recruitment of PARP1 to the UV damage sites, we found that UV-induced global protein PARylation was markedly upregulated in FTO KO cells (Fig. 4f, g). On the contrary, overexpression of myc-FTO significantly suppressed PARP1 enzymatic activity following UV treatment (Supplementary Fig. 7e, f). Changes in the kinetics of PARP1 relocation to DNA strand breaks, which are associated with its polymerase activity, also prompted us to test the possibility that FTO directly affects the enzymatic activity of PARP1. Indeed, we observed that His-FTO inhibited PARP1 activity by limiting the length of the ADP-ribose polymer formed in vitro (Fig. 4h, i). Moreover, significant increases in PARP1 auto-PARylation could be observed in FTO KO cells (Fig. 4j, k), underscoring the role of FTO as an endogenous negative regulator of PARP1 function in cells.

To establish a direct involvement of FTO in DDR through PARP1 interaction, we utilized AlphaFold to predict the binding interface between FTO and PARP1⁴². AlphaFold generated several models with different interfaces; all showed relatively low confidence scores. However, this may be expected as there is currently no experimentally determined structure of full-length PARP1. Despite this, several models predicted the N-terminal domain of FTO to be involved in the interaction with PARP1 (Fig. 5a and Supplementary Fig. 8a). Based on this prediction, we produced recombinant His-FTO protein with a deletion in the first 28 amino acids at the N-terminal region (FTO-Δ28). In vitro binding assays revealed a significant reduction in the binding of recombinant Flag-PARP1 protein to His-FTO-Δ28 compared to full-length His-FTO protein (Fig. 5b, c), demonstrating that FTO directly interacts with PARP1 in part through its N-terminal helix. Next, we transfected full-length myc-FTO or myc-FTO-Δ28 into FTO KO HeLa cells and performed PLA assays to determine their proximal interactions with endogenous PARP1. We found that myc-FTO-Δ28 formed significantly less PLA puncta than full-length myc-FTO, which could be further reduced when the cells were exposed to UV irradiation (Fig. 5d, e). Consequently, re-expression of myc-FTO-Δ28 could only partially rescue dispersed PARP1 clusters, while full-length myc-FTO WT completely restored high-intensity PARP1 clusters in the FTO KO cells (Fig. 5f, g).

**Fig. 5: FTO-PARP1 interaction, but not FTO demethylase activity, is required for PARP1 clustering in cells.**

Using a molecular replacement strategy, we re-introduced myc-FTO (WT or Δ28) in FTO KO HeLa cells and determined their effects on PARP1 functions in DDR. Firstly, we found that the expression of myc-FTO WT not only normalized the rate of GFP-PARP1 association to lesioned DNA but also reduced the amount of GFP-PARP1 recruitment (Fig. 6a–e), mimicking the effect of FTO overexpression in cells (Supplementary Fig. 7a–d). In contrast, the effects of myc-FTO-Δ28 expression on the rate and extent of GFP-PARP1 recruitment to the DNA damage sites were indistinguishable from those in FTO KO cells (Fig. 6a–e). Secondly, UV-induced global protein PARylation was markedly downregulated in FTO KO cells that expressed myc-FTO WT, but was only partially rescued by myc-FTO-Δ28 (Fig. 6f, g). Consistent with these findings, re-expression of myc-FTO WT significantly reduced the rate of DNA repair (measured by the levels of CPD, Fig. 6h, i) and resulted in poor cell survival compared to FTO KO cells (Fig. 6j). On the other hand, the expression of myc-FTO-Δ28, which significantly reduced the interaction with PARP1, only exhibited partial effects (Fig. 6h–j). These data demonstrate the requirement of the FTO N-terminal helix in interacting with and regulating PARP1 functions in cells.

**Fig. 6: FTO-PARP1 interaction, but not FTO demethylase activity, is required for the regulation of PARP1 function and DDR.**

However, it is important to note that FTO contains two nuclear localization signals (NLS), one at the N-terminus (residues 1–31) and another at the C-terminus⁴³. Due to the deletion of the N-terminal NLS, myc-FTO-Δ28 became partially mislocalized in the cytoplasm (Fig. 5d, f and Supplementary Fig. 8b, c), which may also explain why this FTO deletion mutant failed to rescue the phenotypes associated with FTO KO cells. Regardless of the exact underlying mechanisms, FTO-Δ28 represents a loss-of-function mutant that allows us to demonstrate the functional importance of direct FTO-PARP1 interaction and/or FTO nuclear localization in DDR.

FTO regulates PARP1 function independent of its RNA demethylase activity

Given the emerging role of m⁶A signaling in DDR^18,19,20,21, we next investigated the involvement of FTO RNA demethylase activity in regulating PARP1 function and the DNA repair pathway using two complementary approaches. First, we used a small molecule FTO inhibitor FB23-2 (ref. ¹⁴) and confirmed that 72 h treatment with this compound resulted in a significant accumulation of m⁶A levels in HeLa cells (Supplementary Fig. 9a, b). Interestingly, we found that pharmacological inhibition of FTO demethylase activity with 10 μM FB23-2 did not significantly affect the UV-induced GFP-PARP1 recruitment in cells (Supplementary Fig. 9c–f), PARP1 catalytic activity (Supplementary Fig. 9g, h) or the interaction between FTO and PARP1 in control or UV-treated cells (Supplementary Fig. 9i, j). Furthermore, we also found that FB23-2 treatment did not change the levels of CPD formation or cell survival following UV exposure (Supplementary Fig. 9k–m).

To corroborate these findings, we performed molecular replacement analyses of FTO KO cells overexpressing a catalytically inactive FTO H231A/D233A (DD) mutant⁴⁴, which abrogates its ability to bind Fe²⁺ (Fig. 5a). Myc-FTO-DD mutant displayed the expected nuclear localization and interacted with PARP1 to a level comparable to wild-type FTO (Fig. 5d, e). The re-introduction of myc-FTO-DD also effectively restored high-intensity PARP1 clusters in the nucleoli of FTO KO cells (Fig. 5f, g). Like my-FTO-WT, overexpression of myc-FTO-DD suppressed the recruitment of GFP-PARP1 to the DNA damage sites (Fig. 6a–e), reduced the levels of UV-induced global protein PARylation (Fig. 6f, g) and the rate of DNA repair (Fig. 6h, i), resulting in poorer cell survival compared to FTO KO cells (Fig. 6j). Overall, these results provide strong evidence supporting the notion that the role of FTO as a negative regulator of PARP1 function in UV-induced DDR is independent of its m⁶A-demethylase activity.

FTO is a substrate of PARP1

Considering the close interaction between FTO and PARP1, we also examined whether FTO is a direct target of PARP1, revealing that FTO could indeed be PARylated by PARP1 in vitro (Fig. 7a). Due to its proximity to PARP1, a low FTO PARylation could be detected in cells under basal conditions (Fig. 7b, c). Consistent with the activation of PARP1 catalytic activity, FTO became hyper-PARylated 1 min post-UV exposure. However, as the activity of PARP1 waned, the FTO PARylation was significantly down-regulated at 4 min post-UV exposure and returned to baseline at 10 min. Interestingly, the level of PARP1 auto-PARylation showed an inverse correlation with the extent of FTO-PARP1 interaction in cells measured by two independent assays, namely PLA (Fig. 7d, e) and BAR followed by western blotting analysis (Fig. 7f, g). Moreover, the time course of FTO PARylation also inversely correlated with the dispersal of PARP1 clusters in cells following UV treatment (Supplementary Fig. 10a–c). These data suggest a role for protein PARylation in regulating the interaction between these two proteins. To address this possibility, we performed an in vitro binding assay and found that the addition of the PARP1 substrate, nicotinamide adenine nucleotide (NAD + ), dramatically enhanced the interaction between FTO and PARP1 (Fig. 7h). In contrast, when auto-PARylation of PARP1 occurred before His-FTO was added, no augmentation of FTO-PARP1 binding was observed. The dynamic interaction between FTO and PARP1 was not affected by inhibition of FTO demethylase activity (Fig. 5d, e). Based on these results, we propose a model in which a low level of FTO PARylation supports the interaction between FTO and PARP1 under basal conditions. As the activity of PARP1 increases following UV exposure, its auto-PARylation creates steric hindrance and charge repulsion, which subsequently promotes its dissociation from FTO (Fig. 7i).

**Fig. 7: FTO is a direct target of PARP1.**

Discussion

In this study, we used quantitative MS to map the proximal interactome of endogenous FTO and identified a functional interaction with the DNA damage sensor PARP1. Interestingly, this interaction was not observed in previous studies that rely on overexpressing the FTO-BioID2 construct or chemical cross-linking approaches^27,36. Furthermore, there are minimal overlaps with those reported in those studies^27,36 or the BioGRID database⁴⁵. Strikingly, approximately one-third of FTO interacting proteins were also reported in a recent PARP1 proximal interactome study³⁷, suggesting that they share common cellular functions, which include DNA repair, ribosome biogenesis, RNA splicing and chromatin remodeling.

Recent studies have implicated FTO in the DDR pathway induced by various stressors^12,26,27,28. Consistent with our findings that the loss of FTO promotes DNA repair in 3 different cell lines (HeLa, U2OS and HCT116), epidermis-specific deletion of FTO also prevents arsenic- and UVB-induced skin tumorigenesis in mice²⁸. In contrast, some studies have reported that FTO is required to protect cells from DNA-damaging agents such as 2-hydroxyurea, H₂O₂, cisplatin or UVC irradiation^12,26,27. Generally, these studies indicate that FTO functions as an RNA demethylase to control the expression of genes involved in the DNA repair pathway, including PARP1 (ref. ¹²). However, we did not observe any significant changes in the expression of PARP1 protein or PARP1 mRNA in FTO KO HeLa cells. More importantly, pharmacological inhibition of FTO demethylase activity with FB23-2 or overexpression of the catalytic-inactive FTO-DD mutant did not have any significant effect on UV-induced DNA repair and cell survival, suggesting that FTO has an RNA demethylase-independent function in the DDR pathway.

Here we demonstrate that FTO regulates the DDR by physically interacting with PARP1 through the N-terminal helix and regulating its function. Firstly, FTO acts as a scaffolding molecule to cluster inactive PARP1 inside the nucleolus. Upon UV exposure, PARP1 rapidly dissociates from FTO and is recruited to the sites of DNA lesions to promote DNA repair. In FTO KO HeLa cells, PARP1 molecules appear more dispersed and are recruited at a faster rate to DNA damage sites. Secondly, FTO inhibits PARP1 enzymatic activity both in vitro and in vivo. The apparent increases in the levels of global protein PARylation and PARP1 auto-PARylation coincide with more efficient DNA repair in FTO KO cells, likely by promoting the recruitment of DNA repair proteins and facilitating the release of PARP1 from damaged DNA. Moreover, FTO overexpression suppresses the recruitment of PARP1 and decreases the level of UV-induced protein PARylation. Furthermore, we showed that the N-terminal deletion of FTO significantly reduces PARP1 binding and can only partially regulate PARP1 function in cells. These lines of evidence directly support the role of FTO as an endogenous suppressor of the DDR by negatively regulating PARP1 function.

FTO is a direct target of PARP1 and is PARylated in cells. Mechanistically, the interaction between FTO and PARP1 relies on the basal level of FTO PARylation. The activation of PARP1 triggers auto-PARylation, which may create a steric hindrance and force its dissociation from FTO. The high level of FTO PARylation observed shortly after UV exposure could also contribute to its dissociation not only with PARP1 but also with other repair proteins, including DNA-PK and Ku80. Interestingly, the interaction of FTO and PARP1 is largely independent of the former’s RNA demethylase function. Therefore, our study highlights an important mechanism of FTO protein in the DDR beyond its role as an RNA demethylase. Given that METTL3 recruitment to DNA lesions is dependent on PARP1 function, our findings are consistent with the apparent increase in the level of m⁶A observed in FTO KO cells following UV irradiation¹⁸. However, based on our working model, we propose that the increase in m⁶A level is due to enhanced PARP1 function, thus facilitating the more efficient recruitment of METTL3 to sites of DNA damage instead of the loss of RNA demethylase function in FTO KO cells per se.

The enzymatic activity of PARP1 can be positively modulated by several interacting proteins, including the mammalian sirtuin SIRT6 (ref. ⁴⁶), the DNA repair protein XPC⁴⁷, the microtubule-associated protein TPX2 (ref. ³⁷), and the histone PARylation factor HPF1 (ref. ⁴⁸). HPF1 robustly interacts with PARP1 and completes the enzymatic active site, which in turn determines the specificity of PARP1 towards serine-linked PARylation^48,49,50. Interestingly, none of these PARP1 positive modulators was identified in the FTO proximal proteome, further supporting the role of FTO as a unique suppressor of PARP1 activity in cells. One possible mechanism of action may involve the ability of FTO to bind NADP+ and NAD+ (ref. ⁵¹), which limits the availability of PARP1 substrate. FTO could therefore be responsible for preventing hyperactivation of PARP1 which could be detrimental to cells due to excessive consumption of NAD+ substrate, leading to energy collapse and cell death⁵².

In conclusion, our study uncovers an unexpected role of FTO in regulating multiple aspects of PARP1 function, which are not affected by the inhibition of its RNA demethylase activity. We suggest that FTO acts as a scaffolding molecule that is required for the clustering of PARP1 in the nucleolus. The dissociation of FTO regulates the dynamic dispersal of nucleolar PARP1 upon UV-induced DNA damage, suggesting a regulatory mechanism that controls the recruitment of PARP1 to the sites of DNA lesions. Our work also highlights that FTO negatively modulates PARP1 catalytic activity, suggesting the potential clinical use of FTO antisense oligonucleotide to enhance PARP1 sensitivity and increase the efficacy of PARP1 inhibitors for the treatment of many cancer types in human.

Methods

Generation of FTO KO cell lines

HeLa (CCL-2), U2OS (HTB-96) and HCT116 (CCL-247) cells were obtained from ATCC. FTO KO cell lines were generated using the CRISPR/Cas-9 system as previously described³⁰. Three single-guide RNAs (sgRNAs) were designed to target exon 1 of the FTO gene (sgRNA#1 – CACCGAAGCGCACCCCGACTGCCG, sgRNA#2 – CACCGAGCTTCGCGCTCTCGTTCCT and sgRNA#3 – CACCGCTCTCGTTCCTCGGCAGTCG) and cloned into pSpCas9(BB)-2A-GFP vector (a gift from Feng Zhang, Addgene plasmid #48138)⁵³. After transfection, GFP-positive cells were isolated by fluorescence-activated cell sorting and grown as single clones. FTO KO cells were screened and confirmed by genomic DNA sequencing and western blot analysis. The resulting cell lines were HeLa FTO KO.1 (sgRNA#1), FTO KO.2 (sgRNA#2), U2OS FTO KO (sgRNA#1), HCT116 FTO KO.1 (sgRNA#1) and HCT116 KO.3 (sgRNA#3). Most experiments were performed with FTO KO.2 HeLa cells, unless otherwise stated.

Antibodies

The following antibodies were obtained commercially: mouse anti-β-actin (clone C4, Santa Cruz Biotechnology, sc-47778, WB 1:5000), mouse anti-CPD (clone TDM-2, Cosmo Bio USA, CAC-NM-DND-001, WB 1:1000), rabbit anti-DNA PK (Cell Signaling Technology, #4602, WB 1:1000), rabbit anti-FLAG (clone D6W5B, Cell Signaling Technology, #14793, WB 1:1000), rabbit anti-FTO (clone EPR6894, Abcam, ab126605, WB 1:1000, IF 1:250, BAR 1:2000, PLA 1:250), rabbit anti-phospho-Ser-139 Histone H2A.X (clone 20E3, Cell Signaling Technology, #9718, WB 1:1000, IF 1:500), rabbit anti-Ku80 (clone C48E7, Cell Signaling Technology, #2180, WB 1:1000), rabbit anti-m6A (Abcam, ab151230, WB 1:1000), rabbit anti-myc (clone A-14, Santa Cruz Biotechnology, sc-789, IF 1:500, PLA 1:500), mouse anti-Myc (clone 9E10, Santa Cruz Biotechnology, sc-40, WB 1:500), mouse anti-PARP1 (clone A6.4.12, Bio-Rad, MCA1522G, WB 1:1000, IF 1:250, PLA 1:250), mouse anti-PAR (clone 10HA, Trevigen, 4335-MC-100, WB 1:1000), mouse anti-PAR (10H, Merck, MABC547, WB 1:1000) and mouse anti-α-tubulin (clone DM1A, Cell Signaling Technology, #3873, WB 1:10,000). Alexa-conjugated secondary antibodies or streptavidin were purchased from Thermo Scientific. Horseradish peroxidase (HRP)-conjugated secondary antibodies were obtained from GE Healthcare.

DNA plasmids

FTO cDNA was isolated from the human cDNA library by standard PCR with the following primers: 5′- TTGTCGACCAGCATGAAGCGCACCCCGACT-3′ (sense) and 5′- TTGCGGCCGCCTAGGGTTTTGCTTCCAGAA-3′ (anti-sense), and cloned into the pRK5-myc vector or FUW-myc lentiviral vector using the SalI and NotI restriction sites. FTO-Δ28 and FTO-DD (H231A/D233A) were generated using a standard and an overlapping PCR protocol, respectively.

Cell culture

All cell lines were grown in Dulbecco’s modified Eagle’s medium (DMEM) with 4.5 g/L glucose supplemented with 10% fetal bovine serum (FBS), 50 U/mL penicillin and 50 mg/mL streptomycin in a humidified 5% CO₂ incubator at 37 °C. Cells were transfected using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol and processed the next day or transduced with lentiviral particles. For SILAC labeling, HeLa cells were cultured in DMEM (Silantes) containing unlabeled or labeled ²H₄-lysine (Lys-4) and ¹³C₆-arginine (Arg-6) for 8 passages. Labeling efficiency reached ~96% after five passages. To induce DNA damage, cells were pre-treated with 10 μM bromodeoxyuridine (BrdU, Sigma) overnight before UV irradiation exposure. UVC irradiation (254 nm) was performed in a Stratalinker 2400 (Stratagene) or the CL-508 UV cross-linker (UVITec) at the indicated dose (0–100 J/m²). UVA micro-irradiation (405 nm) was performed on an LSM710 confocal microscope (Zeiss). The PARP1 inhibitor Olaparib and FTO blocker FB23-2 were purchased from Selleck. They were dissolved in DMSO and used at the indicated concentrations.

Lentiviral packaging

Lentiviral particles encoding myc-FTO (WT, Δ28 or DD) were generated in HEK293T cells (ATCC, CRL-3216) that had been transfected with 10 μg of FUW-myc-FTO plasmid, 5 μg each of pMD2.G envelope plasmid, pRSV-Rev encoding plasmid and pMDLg/pRRE packaging constructs via the calcium-phosphate precipitation method. Lentivirus-containing supernatant was collected 48 h after transfection and passed through a 0.45 μm cellulose acetate low protein binding membrane. Lentiviral particles were pelleted by ultracentrifugation at 106,559 g at 4 °C for 2 h, resuspended in OptiMEM medium (Invitrogen), snap-frozen in liquid nitrogen and stored at -80 °C.

Cyclobutene pyrimidine dimer (CPD) dot blot assay

Genomic DNAs were extracted from HeLa cells using the DNAzol reagent (Invitrogen) according to the manufacturer’s instructions. DNAs (500 ng) were spotted onto Hybond-N+ nylon membranes (GE Healthcare) and air-dried for 10 min. The membrane was stained with 0.04% methylene blue solution (in sodium acetate pH 5.2) for 5 min to visualize the loading of total RNA on a ChemiDoc XRS+ gel imaging system (Bio-Rad). After extensive washing with methanol, membranes were blocked in 5% skim milk (in PBS containing 0.1% Tween-20) for 1 h and incubated with anti-CPD antibodies (Cosmo Bio USA) overnight at 4 °C. Membranes were washed the next day and incubated with HRP-secondary antibody for 1 h at room temperature. They were then washed extensively and developed using the enhanced chemiluminescence (ECL) method (PerkinElmer). Images were acquired with an Odyssey Fc imaging system (LiCOR) and quantified using ImageStudio software.

γH2A.X immunostaining

Cells were fixed with Parafix solution (4% paraformaldehyde, 4% sucrose in PBS) for 10 min at room temperature, followed by a 2 min methanol fixation at -20 °C. They were then permeabilized with 0.025% Triton X-100 in PBS for 10 min, washed and blocked with 10% normal goat serum (NGS) in PBS for 1 h. After incubation with anti-phospho-Ser-139 H2A.X antibodies (Cell Signaling Technologies) overnight at 4 °C, cells were washed five times in PBS and incubated with Alexa-conjugated secondary antibodies, washed and stained with DAPI in PBS. Images were taken with Zeiss Axio Imager fluorescence microscope and analyzed using ImageJ software (National Institutes of Health).

Colony formation assay

Cells were seeded at low density (500 cells per 6 cm dish) a day prior to UV treatments at the indicated dose, then returned to culture for 14 days prior to being fixed and stained with crystal violet solution (0.05% w/v crystal violet, 1% v/v formaldehyde, 1% v/v methanol in PBS) for 20 min. Tissue culture plates were washed extensively and air-dried.

Biotinylation by antibody recognition (BAR) assay

BAR assays were performed according to a previously published protocol with minor changes³². Briefly, cells were fixed, permeabilized and oxidized with 0.5% H₂O₂. They were then blocked in 3% bovine serum albumin (BSA) in PBS containing 0.1% Tween-20 (PBS-T) for 1 h at room temperature and incubated with anti-FTO antibodies (Abcam; diluted in 3% BSA containing PBS-T) overnight at 4 °C. After extensive washes with PBS-T, samples were incubated with HRP-conjugated anti-rabbit antibodies (in 3% BSA containing PBS-T) for 2 h at room temperature. Samples were washed three times with PBS and incubated with 500 mM of biotin tyramide (Iris Biotech) in PBS for 30 min. The biotin labeling reaction was induced by adding 1 mM H₂O₂ for 1 min and stopped by adding 500 mM of sodium ascorbate to the reaction. After washing with PBS-T, cells were processed for immunostaining or affinity-purification for biochemical analyses.

For immunostaining, biotinylated proteins and FTO immunostaining were detected with Alexa-488-conjugated streptavidin and Alexa-568-conjugated anti-rabbit antibodies, respectively, followed by analysis using a Zeiss Axio Imager epifluorescence microscope.

For biochemical analysis, cells were lyzed with ice-cold RIPA buffer (50 mM Tris, 100 mM NaCl, 2 mM EDTA, 2 mM EGTA, 10 mM Na-pyrophosphate, 50 mM NaF, 1% Triton X-100, 0.5% Na-deoxycholate and 0.1% SDS) supplemented with Complete-EDTA free protease inhibitor cocktails (Roche). Cell lysates were sonicated, boiled for 30 min and centrifuged at 20,627 g for 20 min at 4 °C. Cleared lysates were then incubated with Dynabeads M-280 streptavidin magnetic beads (Invitrogen) or neutravidin agarose resin (Thermo Scientific), followed by mass spectrometry or western blot analyses, respectively.

Mass spectrometry

Immunoprecipitated samples were separated by SDS-PAGE stained with Coomassie Brilliant Blue R-250. Each sample was excised into 3 equal fractions and cut into 0.5-1 mm gel cubes. Proteins were equilibrated in 100 µL of 100 mM ammonium bicarbonate (pH 8.0), destained in 50 mM ammonium bicarbonate/50% acetonitrile (pH 8.0), and dehydrated in 100% acetonitrile. The gel pieces were dried and incubated with 10 mM 1,4-dithiothreitol (Roche) for 30 min at 55 °C and then alkylated in 20 mM iodoacetamide (Sigma) for 30 min at room temperature. After washing, trypsin (12.5 ng/µL, Promega) was added to the gel pieces and they were incubated overnight at 37 °C. Following overnight digestion, tryptic peptides were extracted with 50% acetonitrile/2% formic acid and dried under vacuum centrifugation. They were resuspended in 0.1% formic acid for desalting using pre-equilibrated C18 OMIX tips (Agilent). Bound peptides were eluted with 50% acetonitrile/0.1% formic acid, dried under vacuum centrifugation, and stored at -80 °C until analysis. The peptides were resuspended in 0.1% formic acid, bath sonicated for 15 min and centrifuged at 14,000 g for 15 min to remove any insoluble debris. The clarified peptides were transferred into glass vials for liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis.

The tryptic peptides were separated on an Ultimate 3000 nano-LC (Thermo Scientific) fitted with the Acclaim PepMap C₁₈ reversed-phase column (15 cm length, 75 μm diameter and 2 μm particle size, Thermo Fisher Scientific) and eluted with a linear gradient from 2 to 80% acetonitrile containing 0.1% formic acid) for 60 min. Eluted peptides were subsequently ionized into a Q Exactive Plus mass spectrometer (Thermo Scientific). Precursor ions were selected for MS/MS fragmentation using a data-dependent “Top 10” method operating in Fourier transform (FT) acquisition mode with higher-energy C-trap dissociation (HCD) fragmentation. FT-MS analysis on the Q Exactive Plus was carried out at 70,000 resolution with an automated gain control (AGC) target of 5 × 10⁵ions in full MS. MS/MS scans were carried out at 17,500 resolution with an AGC target of 1 × 10⁵ions. Maximum injection times were set to 50 and 110 ms, respectively. The charge exclusion was set to unassigned and 1+ charged state with a dynamic exclusion of 20 s. The ion selection threshold for triggering MS/MS fragmentation was set to 44,000 counts, and an isolation width of 1.5 m/z was used to perform HCD fragmentation with a normalized collision energy of 27.

Peptide identification and bioinformatic analyses

Raw spectra files were processed using the Proteome Discoverer software 2.2 (Thermo Scientific) incorporating the Sequest search algorithm. Peptide identifications were determined using a 20-ppm precursor ion tolerance and a 0.1-Da MS/MS fragment ion tolerance for FT-MS and HCD fragmentation. Carbamidomethylation modification ( + 57.021 Da) of cysteines was considered a static modification while oxidation ( + 15.995 Da) of methionine, ¹³C₆¹⁵N₂ ( + 8.014 Da) and ²H₄ ( + 4.025 Da) of lysine, ¹³C₆¹⁵N₄ ( + 10.008 Da) and ¹³C₆ ( + 6.020 Da) of arginine, biotin ( + 226.078 Da) of lysine and acetyl ( + 42.011 Da) modification on N-terminal residues were set as variable modifications allowing for maximum two missed cleavages. The data were processed through Percolator to estimate false discovery rates (FDRs). Protein identifications were validated using a q-value of 0.01. PSMs and peptides were validated using a strict FDR for PSMs of 0.01 and 0.05 for a relaxed FDR, respectively. Peptides shorter than 6 amino acids were filtered out. Peptide-spectrum matches (PSMs) were chromatographically aligned for each input file in a sample set with a mass tolerance of 10 ppm and a maximum retention time (RT) shift of 10 min. Peptide groups used for protein quantification were analyzed using the default parameters which set a peptide as unique if it was included in only one protein group. The quantification was processed using unique and razor peptides (peptides shared among multiple proteins group or proteins) with the precursor abundance based on the intensity. Protein abundance was calculated as a sum of the individual peptide group abundances and the ratio was based on a pairwise ratio using a geometric median of the peptide group ratios. A one-way ANOVA was used for the hypothesis test, using the background population of ratios for all peptides and proteins to determine whether any given single peptide or protein was significantly changing relative to that background.

Differential enrichment protein analysis was performed with the R-package DEP⁵⁴. Only proteins detected in at least 2 out of 3 replicates with an adjusted P-value < 0.05 were considered significantly enriched. Gene ontology (GO) enrichment analysis was performed using the R-package clusterProfiler⁵⁵. The protein-protein interaction (PPI) network of selected GO terms was generated with STRING (minimum required interaction score > 0.7) and visualized with Cytoscape^56,57.

Proximity ligation assay (PLA)

Monolayers of WT and FTO KO HeLa cells were fixed with Parafix solution for 10 min at room temperature, followed by a further 2 min fixation with 100% methanol at -20 °C. Cells were then permeabilized with 0.25% Triton X-100/PBS for 10 min and blocked in 10% normal goat serum for 1 h, after which they were incubated with rabbit anti-FTO (Abcam), rabbit anti-myc (Santa Cruz Biotechnology) and mouse anti-PARP1 (Bio-Rad) antibodies at 4 °C overnight. PLA was carried out with the Duolink® kit (Sigma) according to the manufacturer’s protocol. Briefly, cells were incubated with mouse PLUS and rabbit MINUS Duolink PLA probes at 37 °C for 1 h, ligated at 37 °C for 30 min and amplified (Texas Red) at 37 °C for 100 min. Nuclei were visualized with DAPI staining. Images were acquired on a Zeiss LSM510 confocal microscope with a 63× oil-immersion objective (1.4 N.A.). PLA puncta number and fluorescence intensities within cell nuclei were quantified using the BioVoxxel Toolbox (Speckle Inspector) plugin in ImageJ software (NIH). Nuclei were set as big objects with a minimum object size of 3000 pixels and the PLA fluorescence was set as small speckles with a minimum speckle size of 7 pixels.

Structured illumination and confocal microscopy

Cells were fixed, blocked, permeabilized and stained with rabbit anti-FTO and mouse anti-PARP1 antibodies. FTO and PARP1 were visualized by staining with Alexa-488-conjugated anti-rabbit and Alexa-568-conjugated anti-mouse secondary antibodies, respectively. Nuclei were visualized with DAPI staining. Coverslips were mounted on a glass slide with Vectashield (Vector Laboratories). Images were acquired on a Zeiss ELYRA inverted structured illumination microscope equipped with a 63x Plan-Apochromat oil-immersion objective (1.4 N.A.). Raw SIM files were manually processed with the Zeiss Zen microscopy software using the structured illumination function (noise filter at -5.0 to -6.0 and SR frequency weighting at 1.0), followed by spatial misalignment adjustment (channel alignment method) with calibrated polystyrene beads coated with multiple fluorophores. In some experiments, images were acquired on a Zeiss LSM 980 NLO Airyscan 2 confocal microscope built around an Axio Observer 7 body and equipped with a 63x Plan-Apochromat oil-immersion objective (1.4 N.A.), Airyscan 2 super-resolution detector, a 34-channel spectral PMT (photomultiplier tube) array, two internal GaAsP PMTs, a transmission PMT, and controlled by Zeiss Zen Blue software. Raw images were processed on the Zeiss Zen Blue software using the Airyscan joint deconvolution, and the number of iterations was set at 8.

The rendering of FTO and PARP1 surfaces in the nucleus was created using the surface tool (smooth surface detail set at 0.02 mm) in the Imaris image analysis software (Oxford Instruments), which were subsequently used to calculate the volume (μm³), sphericity and nearest distance (μm) of FTO and PARP1 particles. Nuclear PARP1 clusters and area analyses were performed using the ImageJ software. The minimum cluster radius of PARP1 was thresholded at 3.0 pixels. The number of PARP1 clusters per nucleus was manually counted. The area of the nucleus and each PARP1 cluster was determined by tracing the region of interest (ROI).

Laser micro-irradiation and GFP-PARP1 recruitment assay

A plasmid expressing GFP-PARP1 was generated by subcloning PARP1 cDNA from the pCMV-PARP1-3x-Flag construct (a gift from Thomas Muir, Addgene plasmid #111575)⁵⁸ into the pEGFP-C3 vector (Clontech). Cells were transfected with the pEGFP-PARP1 construct and treated with 10 μM BrdU in DMEM without phenol red but containing 10% FBS and 1% penicillin and streptomycin overnight. The recruitment of GFP-PARP1 was imaged at 5 Hz on a Zeiss LSM710 confocal microscope. DNA damage was induced by irradiating a 10 pixel-width of the nucleus with a 405 nm laser (line scan mode). The laser was set with a line attenuator transmission of 92%, 1 iteration and a pixel dwell time of 177.32 µs. ROIs were drawn to the DNA damage site to measure the GFP fluorescence over time using the ImageJ software. DNA damage-induced changes in fluorescence were calculated by normalizing the background subtracted values to the baseline fluorescence prior to UVA micro-irradiation (ΔF/Fo). The recruitment time constant (τ) was calculated by fitting the fluorescence recovery after micro-irradiation to a single exponential function.

Cellular PARylation assay

Cells were lyzed in 1% SDS (in PBS) and diluted in 10 volumes with ice-cold cell lysis buffer (1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 50 mM NaF, 5 mM Na-pyrophosphate in PBS) supplemented with Complete EDTA-free protease inhibitor cocktails (Roche). Cell lysates were sonicated and centrifuged at 20,627 g for 20 min at 4 °C. Global PARylation of cellular proteins were detected by SDS-PAGE and western blotting with anti-PAR antibodies (Trevigen or Merck). α-tubulin or β-actin blots were used as loading controls. To detect PARP1 auto-PARylation, total cell lysates were probed with anti-PARP1 antibodies (BioRad) to reveal the slower migrating bands, which were not present in Olaparib-treated cells (negative control). To detect PARylation of endogenous FTO protein, cell lysates were subjected to immunoprecipitation with anti-FTO antibodies (Abcam) that were coupled to protein A Sepharose beads at 4 °C overnight. Bound proteins were washed and eluted with 2X SDS sample buffer for 10 min at 95 °C. Eluates were resolved on SDS-PAGE and blotted with anti-PAR and anti-FTO antibodies.

In vitro binding assay

The full-length FTO open-reading frame was subcloned into the pRSFDuet1 bacterial expression vector. FTO construct lacking the first 28 amino acids (Δ28) was also subcloned into the same bacterial expression vector. The expression and purification of recombinant His-FTO proteins (full-length or Δ28) in Escherichia coli BL21(DE3) cells were performed according to a previously published method⁵⁹. For the in vitro binding assay, 5 μg of purified His-FTO recombinant proteins were pre-coupled to 30 μL of Ni Sepharose beads (Cytiva, pre-blocked with 1.5% BSA in PBS-T) in binding buffer (0.5% Triton X-100, 0.5 mM EDTA, 0.5 mM EGTA, 25 mM NaF, 2.5 mM Na-pyrophosphate, 10 mM Na phosphate, 0.25 M NaCl, 10 mM imidazole in 0.5× PBS, PH 7.4) for 1 h at room temperature. The beads were washed twice in the binding buffer before incubation with 1 μg of recombinant Flag-PARP1 proteins (Abcam) for 1 h at room temperature. The beads were then washed four times and eluted by boiling them in 2× SDS sample buffer for 10 min at 95 °C. Eluates were resolved on SDS-PAGE, and either blotted with anti-Flag and anti-FTO antibodies or stained with Coomassie Brilliant Blue R-250.

In vitro PARylation assay

The in vitro PARylation assay was performed by adding 1 μg of recombinant Flag-PARP1 protein in 50 μL of reaction buffer (20 mM Tris-Cl, 100 mM NaCl, 10 mM MgCl₂, 10 mM ZnCl₂, 10% glycerol, 300 mM NAD + , pH 8.0). The addition of 10 μM of PARP1 inhibitor Olaparib or the omission of NAD+ in a reaction served as negative controls. Recombinant His-FTO recombinant proteins (5 μg) were added to the reaction mix as specified. The reactions were performed at 30 °C for 30 min. PARP1 PARylation was analyzed by western blotting using anti-Flag antibodies (Cell Signalling Technology). To detect FTO PARylation, the reaction containing His-FTO was incubated with Ni Sepharose beads, washed and eluted in 2X SDS sample buffer for 10 min at 95 °C. FTO PARylation was analyzed by western blotting using anti-PAR antibodies.

Western blotting

Samples were loaded in 7.5 or 10% SDS-PAGE gels and separated at 110 V for 1-2 h. Proteins were then transferred to a PVDF membrane at 100 V for 2 h. Membranes were blocked in 5% skim milk (in TBS containing 0.1% Tween-20, TBS-T) for 1 h and washed in TBS-T three times at 5 min intervals prior to overnight incubation with primary antibodies at 4 °C. Membranes were washed in 1% milk/TBS-T five times and incubated with HRP-conjugated secondary antibodies (GE Healthcare, 1:10,000) for 1 h at room temperature. They were then washed extensively and developed using the enhanced chemiluminescence method (PerkinElmer). In some experiments, the total protein load was visualized by Revert Protein Stains (LiCOR) according to the manufacturer’s instructions. Images were acquired with a LiCOR imaging system and quantified with the ImageStudio software. Uncropped blots are provided in the source data file.

m⁶A dot blot

Total RNA was extracted using Trizol (Invitrogen) and normalized to the same concentration. RNA was denatured at 95 °C for 3 min and then chilled on ice. An equal volume of RNA was spotted on a Hybond-N+ nylon membrane (GE Healthcare), air-dried and UV crosslinked. The membrane was stained with 0.04% methylene blue solution (in sodium acetate pH 5.2) for 5 min to visualize the loading of total RNA on a ChemiDoc XRS+ gel imaging system (Bio-Rad). After extensive washing with methanol, the membrane was blocked with 5% skim milk in PBS containing 0.1% Tween, incubated with anti-m⁶A antibody (Abcam) overnight at 4 °C, followed by incubation with HRP-conjugated secondary antibodies (GE Healthcare, 1:10,000) and detection with the enhanced chemiluminescence method (PerkinElmer).

Quantitative RT-PCR

Reverse transcription of total RNA from wild-type or FTO KO HeLa cells was performed using SuperScript IV (Invitrogen) according to the manufacturer’s instructions. Quantitative real-time PCR with Rotor-Gene Q real-time cycler (Qiagen) was performed using SYBR-green amplification (Takara Bio). PARP1 and ACTB transcript levels were measured using the following primers: PARP1 FP 5′-GGGCAAGCACAGTGTCAAAG-3’ and PARP1 RP 5′-CCAGATGAAATCCCGGTCCC-3′, ACTB FP 5′-GCAGGAGTATGACGAGTCCGGC-3′ and ACTB RP 5′-GTAACAACGCATCTCATATTTG-3′, respectively. PARP1 expression was calculated using the standard comparative Ct method after normalizing to the housekeeping gene ACTB.

Statistical analysis

The sample size (N) reported in figure legends represents individual cells or wells generated from at least three independent experiments, unless otherwise stated. Statistical analysis was performed in Graph Pad Prism 9.0 using one-way or two-way analysis of variance (ANOVA). A two-tailed student’s t-test was employed to compare between two groups. All data are reported as mean ± standard error of the mean (SEM), unless stated otherwise.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE⁶⁰ partner repository with the dataset identifier PXD041638. Source data are provided with this paper.

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Acknowledgements

We thank R. Amor and A. Gaudin for their support with microscopy; V. Nink for assisting with the flow cytometry; and R. Tweedale for comments on the manuscript. This work was partly supported by Australian Medical Research Future Fund (Clem Jones Centre for Ageing Dementia Research Flagship Project Grant to V.A.). V.A. holds an Australian Research Council (ARC) Future Fellowship (FT220100485). J.W. was supported by an ARC Discovery Early Career Researcher Award (DE170100112), ARC Discovery Project Grant (DP230102041), a University of Queensland (UQ) Amplify Award and the Peter Hilton Alzheimer’s Fund. J.Z.A.T. was supported by a UQ Researcher Retention (RSA2) Fellowship. T.Z., L.R., S.S.D. and H.H. were supported by UQ Research Training Scholarships. Imaging was performed at the Queensland Brain Institute’s Advanced Microscopy Facility, supported by the Australian Government through ARC LIEF grant LE130100078.

Author information

These authors jointly supervised this work: Jocelyn Widagdo, Victor Anggono.

Authors and Affiliations

Clem Jones Centre for Ageing Dementia Research, Queensland Brain Institute, Faculty of Health, Medicine and Behavioural Sciences, The University of Queensland, Brisbane, Australia
Tianyi Zhu, Jing Zhi Anson Tan, Lingrui Zhang, He Huang, Sooraj S. Das, Pranesh Padmanabhan, Jocelyn Widagdo & Victor Anggono
The Solomon H. Snyder Department of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, USA
He Huang
Motor Neuron Disease Research Centre, Macquarie Medical School, Faculty of Medicine Health and Human Sciences, Macquarie University, Sydney, Australia
Flora Cheng & Albert Lee
School of Biomedical Sciences, Faculty of Health, Medicine and Behavioural Sciences, The University of Queensland, Brisbane, Australia
Pranesh Padmanabhan
NHMRC Centre for Research Excellence in Mechanisms in NeuroDegeneration – Alzheimer’s Disease (MIND-AD CRE), Brisbane, Australia
Pranesh Padmanabhan & Victor Anggono
Frazer Institute, Faculty of Health, Medicine and Behavioural Sciences, The University of Queensland, Brisbane, Australia
Mathew J. K. Jones
School of Chemistry & Molecular Biosciences, Faculty of Science, The University of Queensland, Brisbane, Australia
Mathew J. K. Jones
Department of Biochemistry and Chemistry, La Trobe Institute for Molecular Science, La Trobe University, Melbourne, Australia
Mihwa Lee
School of Chemistry, Bio21 Molecular Science and Biotechnology Institute, University of Melbourne, Parkville, Australia
Mihwa Lee

Authors

Tianyi Zhu
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Jing Zhi Anson Tan
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Lingrui Zhang
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He Huang
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Sooraj S. Das
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Flora Cheng
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Albert Lee
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Jocelyn Widagdo
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Victor Anggono
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Contributions

Conceptualization: T.Z., J.W., and V.A. Methodology: T.Z., J.Z.A.T., L.Z., H.H., S.S.D., M.J.K.J., M.L., A.L., J.W., and V.A. Investigation: T.Z., J.Z.A.T., L.Z., F.C., P.P., M.L., A.L., J.W. and V.A. Visualization: T.Z., J.Z.A.T., L.Z., H.H., M.L., J.W., and V.A. Funding acquisition: J.W. and V.A. Project administration: J.W. and V.A. Supervision: J.W. and V.A. Writing—original draft: T.Z., J.W. and V.A. Writing—review & editing: T.Z., J.Z.A.T., L.Z., P.P, M.J.K.J., M.L., A.L., J.W., and V.A.

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Correspondence to Jocelyn Widagdo or Victor Anggono.

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Zhu, T., Tan, J.Z.A., Zhang, L. et al. FTO suppresses DNA repair by inhibiting PARP1. Nat Commun 16, 2925 (2025). https://doi.org/10.1038/s41467-025-58309-0

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Received: 06 July 2023
Accepted: 17 March 2025
Published: 25 March 2025
DOI: https://doi.org/10.1038/s41467-025-58309-0