TET-dependent regulation of retrotransposable elements in mouse embryonic stem cells
© The Author(s). 2016
Received: 25 July 2016
Accepted: 1 November 2016
Published: 18 November 2016
Ten-eleven translocation (TET) enzymes oxidise DNA methylation as part of an active demethylation pathway. Despite extensive research into the role of TETs in genome regulation, little is known about their effect on transposable elements (TEs), which make up nearly half of the mouse and human genomes. Epigenetic mechanisms controlling TEs have the potential to affect their mobility and to drive the co-adoption of TEs for the benefit of the host.
We performed a detailed investigation of the role of TET enzymes in the regulation of TEs in mouse embryonic stem cells (ESCs). We find that TET1 and TET2 bind multiple TE classes that harbour a variety of epigenetic signatures indicative of different functional roles. TETs co-bind with pluripotency factors to enhancer-like TEs that interact with highly expressed genes in ESCs whose expression is partly maintained by TET2-mediated DNA demethylation. TETs and 5-hydroxymethylcytosine (5hmC) are also strongly enriched at the 5′ UTR of full-length, evolutionarily young LINE-1 elements, a pattern that is conserved in human ESCs. TETs drive LINE-1 demethylation, but surprisingly, LINE-1s are kept repressed through additional TET-dependent activities. We find that the SIN3A co-repressive complex binds to LINE-1s, ensuring their repression in a TET1-dependent manner.
Our data implicate TET enzymes in the evolutionary dynamics of TEs, both in the context of exaptation processes and of retrotransposition control. The dual role of TET action on LINE-1s may reflect the evolutionary battle between TEs and the host.
Transposable elements (TEs) are mobile genetic elements that are present in organisms from all kingdoms of life and account for nearly half of the human and murine genomes. Retrotransposons, which act through an RNA intermediate, are recognised as major evolutionary contributors to genome structure and organisation. Namely, retrotransposons bear enormous potential for affecting gene expression at multiple levels, from creating novel transcription factor (TF) binding sites to regulating post-transcriptional processes [1, 2]. The co-adoption, or exaptation, of TEs by the host genome is often associated with the creation of species-specific cis-acting elements that help to shape gene regulatory networks [1, 3]. In embryonic stem cells (ESCs), long-terminal repeat (LTR) TEs (also termed endogenous retroviruses; ERVs) have made a substantial contribution to the distribution of binding sites for pluripotency factors across both human and mouse genomes [4, 5], and the expression of HERVH TEs is required to safeguard the identity of human ESCs [6, 7].
On the other hand, active TEs such as LINE-1 (or L1) elements carry substantial mutagenic potential, as evidenced by numerous examples of disease-causing retrotransposition events . Somatic retrotransposition of L1s has also been linked to disease, especially in the context of cancer [9, 10]. Mobility restriction of these elements is therefore of paramount importance to maintain genome stability, especially during the crucial developmental windows of germline establishment and early embryogenesis. Notably, both human ESCs and induced pluripotent stem cells (iPSCs) support L1 mobility  and de novo retrotransposition events from endogenous TEs have been detected in clonally derived iPSCs , with important implications for regenerative medicine strategies.
During early embryogenesis, TEs are kept under control by multiple epigenetic mechanisms such as DNA methylation, histone modifications and small RNA-mediated mechanisms . The large and rapidly evolving family of Krüppel-associated box domain-containing zinc finger proteins (KRAB-ZFPs) play a major role in ERV silencing by recruiting the cofactor KRAB-associated protein 1 (KAP1), which acts as a docking protein for heterochromatin-forming activities, including the H3K9 methyltransferase SETDB1 [14, 15]. Interestingly, KRAB-ZFPs have also been implicated in the silencing of old L1 elements, acting as major drivers of L1 evolutionary dynamics [16, 17]. In contrast, Piwi-interacting RNAs (piRNAs) are important for silencing of young, active L1s in humans iPSCs . DNA methyltransferases, which can act downstream of piRNA action, also play a crucial role in the repression of young L1 families in both mouse and human ESCs [15, 16, 19]. However, DNA methylation (5-methylcytosine; 5mC) undergoes genome-wide erasure during preimplantation development , creating a potential window of opportunity for TE expression. While the primary mechanism underlying DNA demethylation in this period is replication-coupled dilution of 5mC [21, 22], an active mechanism dependent on ten-eleven translocation (TET) enzymes has also been demonstrated for specific loci [23–25]. TETs catalyse the iterative oxidation of 5mC to 5-hydroxymethylcytosine (5hmC), 5-formylcytosine (5fC) and 5-carboxylcytosine (5caC), with the latter two bases being substrates for TDG-dependent repair mechanisms that reintroduce unmodified cytosines into the genome . TET enzymes can also modulate transcription in a non-catalytic way by recruiting other chromatin regulators, such as OGT (O-linked β-d-N-acetylglucosamine (O-GlcNAc) transferase) , the SIN3 co-repressor complex  and the Polycomb repressive complex 2 (PRC2) . Unlike TET-mediated DNA demethylation, the interaction with these proteins can have repressive outcomes, underlying a dual nature of TET action in ESCs [28, 30].
We have previously shown that L1 elements are enriched in 5hmC in mouse ESCs [31, 32], raising the possibility that TET enzymes regulate their transcription. However, a wider and functional investigation of the role of TET enzymes at TEs is missing. Here we asked whether TET enzymes regulate the action of TEs, both as exapted regulatory elements and as mutagenic mobile elements. We found that TETs bind to multiple TE families, including TE-derived ESC-specific enhancers, which are kept hypomethylated in part by the action of TET2, thus helping to maintain the expression of associated genes. We also show that TETs demethylate young L1 elements but that their expression levels are stably maintained upon TET depletion due to the loss of TET-dependent repressive activities, such as SIN3A. Our findings implicate TET enzymes in the evolution of the TE-host relationship, contributing to exaptation processes and forming a hub for the so-called evolutionary arms race.
TET1 binds to repetitive elements with diverse chromatin signatures
TET2 helps to maintain gene expression driven by TE-derived enhancers
TETs and 5hmC are enriched at the 5′ UTR of young L1 elements
We then asked whether a similar mechanism could be implicated in the regulation of human L1 elements. Using TAB-seq data from human ESCs , we uncovered a remarkably similar pattern of 5hmC at a highly active human L1 (L1.4 ), whereby the 5′ UTR displayed relatively high levels of this mark (Fig. 3g). Moreover, 5hmC enrichment across human L1 families mirrored our observations in the mouse, showing that only young, KAP1-unbound elements were enriched for 5hmC and this was accompanied by a decrease in 5mC levels (Fig. 3h). These observations show that, along independent evolutionary paths, L1 elements recruited TET enzymes and accumulated 5hmC at their 5′ UTR in a manner that correlates with their activity.
TET-mediated L1 demethylation has no overall effect on expression
Given that loss of DNA methylation in DNMT KO ESCs leads to L1 derepression [15, 19], we predicted that TET depletion would cause a downregulation of L1s. Surprisingly, however, quantitative reverse transcription polymerase chain reaction (RT-qPCR) analyses showed that the expression levels of L1 elements were unaffected in TET1-depleted or TET2-depleted ESCs, both at four and ten days after lentiviral infection (Fig. 4c). L1 expression was also stable upon DKD of both TETs (Fig. 4c). Analysis of our RNA-seq data supported these observations and further revealed that no TET1-bound TE classes were deregulated in TET-depleted ESCs (Additional file 2: Figure S4A and B); in contrast, substantial changes in the expression of single-copy genes were readily detectable (Additional file 2: Figure S4C). RNA-seq data from WT and Tet1/Tet3 double knockout blastocysts further supported these findings (Additional file 2: Figure S4D) . In line with the RNA data, we detected no alterations in the levels of ORF1p protein (Fig. 4d). Additionally, using northern blot we found no significant differences in the levels of full-length L1Tf elements (Fig. 4e). We considered the possibility that changes in the expression of individual L1 copies that gained 5mC may be undetectable when analysing the whole L1 RNA pool. To test this hypothesis, we designed specific primers for RT-qPCR and oxBS that target three individual full-length L1 loci in the mouse reference genome (referred to here as HL1-2, HL1-6 and HL1-7; see ‘Methods’ for details). All tested L1 copies showed a significant increase in 5mC levels upon DKD of both TETs or single knockdown of TET2, with two of them also displaying changes upon TET1 depletion (Fig. 4f). Despite these differences, none of these L1 copies had significantly altered expression levels upon depletion of TET1, TET2 or both enzymes (Fig. 4f).
Overall, these results demonstrate that TET-mediated changes in DNA methylation at L1s do not translate into expression differences, indicating that although DNA methylation is an important regulator of L1 repression in ESCs, other layers of regulation can play equally prominent roles.
TET-dependent epigenetic activities keep L1s repressed
We argued that any compensatory repressive mechanism that depends on TET1 is likely to be mediated by known interacting partners, namely OGT, SIN3A and the PRC2 complex [27–29]. Publicly available ChIP-seq data revealed that both OGT and SIN3A are enriched at the 5′ UTR of L1 elements, while members of the PRC2 complex are undetectable (Fig. 5b). We confirmed the enrichment of OGT and SIN3A at the 5′ UTR of young L1s by ChIP-qPCR (Fig. 5c). To test the effect of OGT on L1 expression, we depleted ESCs of OGT by siRNA, which led to a global loss of O-GlcNAc (Fig. 5d). Interestingly, OGT depletion caused an upregulation of L1 elements and this effect was partly dependent on TET1 expression (Fig. 5e). We confirmed that L1 derepression involved full-length elements and that ORF1p protein levels were also raised (Additional file 2: Figure S7A and B). However, we noted that TEs that are not bound by TET1 were also derepressed, suggesting that the effect of OGT depletion on L1s may be indirect (Fig. 5e). We then raised cellular levels of O-GlcNAc by inhibiting O-GlcNAc hydrolase (OGA), which led to a small increase in the expression of L1s, but not of TET1-unbound TEs (Additional file 2: Figure S7C and D). The seemingly conflicting outcomes of OGT depletion and OGA inhibition likely reflect the broad and complex roles that O-GlcNAc plays in chromatin regulation, with both activating and repressive outcomes [48, 49]. Nonetheless, it remains possible that cycling of O-GlcNAc through OGT and OGA plays a direct role in L1 silencing. To test whether recruitment of SIN3A to L1s is involved in their regulation, we used siRNA/shRNA-mediated depletion of SIN3A (Fig. 5f). This caused a 50–70% increase on the expression levels of young L1s, whereas TEs unbound by TET1 remained unaffected (Fig. 5g). Moreover, the effect on L1s was TET1-dependent, as the upregulation observed on SIN3A KD samples was lost when we knocked down both SIN3A and TET1, which is in line with the TET1-dependent recruitment of SIN3A to chromatin . We also confirmed that SIN3A-mediated L1 derepression involved full-length elements and a concomitant increase of ORF1p protein levels (Additional file 2: Figure S7A and B). This result is, to the best of our knowledge, the first time that SIN3A has been implicated in TE regulation. Interestingly, SIN3A binding is also seen at the 5′ UTR of human L1s in ESCs (Additional file 2: Figure S8), raising the possibility that human L1s are also kept repressed by TET-dependent deposition of SIN3A. Our data show that there are TET-dependent repressive activities that likely counteract oxidative DNA demethylation at L1 elements. Indeed, we show that, in a SIN3A-null context, TET1 is a positive regulator of L1 expression. Our work uncovers TET enzymes as epigenetic hubs with dual roles in L1 regulation.
At L1 elements, TET binding displays a striking pattern whereby it targets exclusively the 5′ UTR of young, active L1 clades. Moreover, the same pattern is apparent in human ESCs (as judged by the presence of 5hmC), which is suggestive of an evolutionarily convergent mechanism that allows for L1 activation and expansion. Parallel evolution of L1 silencing mechanisms is also seen for KAP1 action, which targets and silences older L1 elements in both mouse and human through the binding of KRAB-ZFPs [16, 17]. Interestingly, none of the young TET1/5hmC-enriched L1s are KAP1 targets. KAP1 binding leads to recruitment of SETDB1 and deposition of H3K9me3, which has been suggested to impair TET1 binding at TEs . Therefore, the evolutionary escape of L1s from KAP1-mediated repression may have created permissive conditions for TET binding, which in turn led to DNA demethylation (Fig. 6b). Accordingly, we found that TET depletion leads to increased methylation at the 5′ UTR of young L1s. It is possible that the scale of this effect is dependent on cell culture conditions, as these are known to deeply affect DNA methylation levels in ESCs . We note that L1 methylation levels in ESCs grown with standard serum-containing medium, as used for all our experiments, closely match those seen in blastocysts and inner cell mass cells (Additional file 2: Figure S9). In contrast, cells grown with the so-called ‘2i’ formulation (with or without vitamin C) display substantially lower L1 methylation levels (Additional file 2: Figure S9), suggesting that the epigenetic modifiers targeting L1s in 2i cells differ to those acting in vivo.
Given that young L1s are derepressed in DNMT-null ESCs [15, 16], TET-mediated DNA demethylation is expected to activate L1 expression. However, our data clearly show that TET depletion has no overall effect on L1 expression levels, potentially arguing against a role of TET recruitment in TE expansion. Mechanistically, the maintenance of L1 expression upon TET-mediated demethylation suggests that DNA methylation may not play as prominent a role in the regulation of these TEs as previously thought. It is possible that the L1 derepression seen in DNMT mutants [15, 16, 19] are independent of the enzymes’ catalytic activity , although ESCs expressing catalytically inactive DNMT1 fail to repress IAPs . Alternatively, TETs may drive other catalytically independent alterations to chromatin that compensate for changes in DNA methylation. In line with this hypothesis, we found that TET2 depletion leads to a reduction of H3K9me3 levels at L1s and that TET1 recruits the SIN3A co-repressor complex, which in turn plays a role in L1 silencing. TET1 also recruits OGT to L1s and our data indicate a possible role for O-GlcNAc turnover in L1 repression. Interestingly, SIN3A is modified by OGT to synergistically repress transcription , raising the possibility that the OGT-mediated and SIN3A-mediated repressive effects on L1s are mechanistically linked. However, OGT may also affect TE expression through RNA Pol II [53, 54] or other effects on chromatin [48, 49].
Could TET-dependent repressive activities constitute a host strategy for ensuring L1 silencing upon TET-mediated DNA demethylation? In one possible evolutionary scenario, as a newly formed L1 clade (not necessarily the most recent ones) escaped from the repressive action of KRAB-ZFP/KAP1, permissive TET-binding conditions would have been created that would lead to DNA demethylation, contributing to L1 expansion (Fig. 6b). Selective pressure to maintain genome stability could then have driven the evolution of TET-dependent host strategies to control L1 expansion, such as sequestration of epigenetic silencing activities (e.g. SIN3A) by TET enzymes (Fig. 6b). In this scenario, TETs would have served as ‘double agents’ in an arms race between L1s and the host genome. As each subsequent emerging L1 family escaped KRAB-ZFP/KAP1 repression, the activating effects of TET binding would have been dampened by the accompanying repressive activities. More effective and permanent silencing of an active L1 family would then require the recognition by a novel KRAB-ZFP . In an alternative scenario, the main selective pressures driving the interactions between TETs and epigenetic silencers could have come from elsewhere. Yet the added need to maintain control of L1 expression during preimplantation could have arguably re-enforced these epigenetic relationships and further contributed to their evolution and fine-tuning.
It remains possible that in other cell types TET-mediated control of DNA methylation regulates L1 transcription due to differences in the underlying epigenome. For example, TET1 and TET2 silencing during early ESC differentiation may be important to ensure full L1 repression through DNA methylation. And in neurons, where 5hmC levels are particularly high , TET-mediated DNA demethylation could help drive the high levels of L1 expression seen therein [56, 57].
We have unveiled several novel roles of TET enzymes in the regulation of TEs, both catalytically dependent and independent. TETs mediate DNA demethylation of several TEs, but also serve as a hub for additional epigenetic activities, both of which impact on the evolution of the TE-host relationship. The intriguing dual nature of TET action on L1 elements appears to reflect the competing interests of host and TEs, raising new questions on TE-driven evolution of epigenetic mechanisms.
ESC culture, gene knockdown and PUGNAc treatment
E14 ESCs were grown in feeder-free conditions using DMEM-based medium with 15% FBS and 1000 U/mL ESGRO LIF (Millipore). For shRNA-mediated gene knockdown, ESCs were infected with viral particles carrying pLKO.1 constructs harbouring gene-specific shRNAs or a non-targeting sequence (Scr: CCTAAGGTTAAGTCGCCCTCGCTC; shTET1: TTTCAACTCCGACGTAAATAT, TRCN0000341848; shTET2: TTCGGAGGAGAAGGGTCATAA, TRCN0000250894). After 24 h, cells were selected with 10 μg/mL puromycin or 50 μg/mL hygromycin for 3–9 days (see ‘Results’). For siRNA-mediated knockdown, cells were transfected twice (second transfection two days after the first one) with gene-specific (see below) or non-targeting siRNAs (Dharmacon siGENOME Non-Targeting siRNA #2 Cat.D-001210-02-20) using Lipofectamine 3000 (Thermo Scientific, Cat. L3000008) and collected four days post transfection. Tet1 (Dharmacon siGENOME Mouse Tet1 Cat. D-062861-01-0020) and Sin3a siRNAs (Dharmacon Custom siRNA ON-TARGET Cat. CTM-220747, sequence: gctgttccgattgtccttaaa) were used at 125 nM, whereas Ogt esiRNAs (Sigma-Aldrich, Cat. EMU006701) were used at 50 nM. Knockdown was confirmed by RT-qPCR and western blot. PUGNAc (Sigma-Aldrich, Cat. A7229) treatment was performed either with 100 μM of DMSO-dissolved PUGNAc or matched amount of DMSO in control conditions and its efficiency was tested by western blot. A list of antibodies used can be found in Additional file 3.
RNA isolation, RT-qPCR and northern blot
RNA was extracted using Quick-RNA™ MiniPrep (Zymo Research D7003) and DNAse treated in-column either with the DNAse provided on the kit or with the TURBO DNA-free™ Kit (Ambion, AM1907). RNA (1 μg) was retrotranscribed using SuperScript® III Reverse Transcriptase (Invitrogen, Cat. 18080044) and the cDNA was diluted 1/10 for qPCRs using MESA BLUE MasterMix (Eurogenentec, 10-SY2X-03 + NRWOUB) on a LightCycler® 480 Instrument II (Roche). A list of primers used can be found in Additional file 3: Table S2. For northern blotting, 0.5 μg of glyoxylated total RNA was separated on a 1.2% BPTE gel, then blotted and probed as described in . RNA probes were amplified from mouse genomic DNA using the primers detailed in Additional file 3.
ChIP was performed as described in , with modifications. For the detection of TFs (TET1, TET2, SIN3A, OGT, OCT4), cells were fixed with an initial cross-linking step of 45 min with 2 mM Di(N-succinimidyl) glutarate (Sigma-Aldrich Cat. 80424) in PBS at room temperature, followed by a PBS wash and a second fixation step of 12 min with 1% formaldehyde in DMEM. For histone ChIPs (H3K4me3, H3K9me3, H3) a single cross-linking step with 1% formaldehyde for 15 min was used. After quenching with glycine, washes and lysis, chromatin was sonicated using a Bioruptor Pico from Diagenode, on a 30 s on/off cycle for 12 (TFs) or eight (histones) cycles. Immunoprecipitation was performed using 150 μg of chromatin and 7.5 μg of antibody (TFs) or 30 μg of chromatin and 5 μg of antibody (histones). Final DNA purification was performed using the GeneJET PCR Purification Kit (Thermo Scientific. Cat. K0701) and elution in 80 μL of elution buffer. This was diluted 1/10 and analysed by qPCR, using the KAPA SYBR® FAST Roche LightCycler® 480 2X qPCR Master Mix (Kapa Biosistems, Cat. KK4611). A list of antibodies and primers used can be found in Additional file 3.
RNA-seq and small RNA-seq
Ribosomal RNA-depleted RNA-seq libraries were prepared from 400–600 ng of total RNA using the low input ScriptSeq Complete Gold Kit (Epicentre). For small RNA-seq, RNA was isolated with QIAzol (QIAGEN) and libraries prepared from 400–600 ng of RNA using the NEBNext small RNA library prep kit (NEB), followed by size selection (~120–150 bp, including adaptors) using gel extraction. Libraries were sequenced on an Illumina NextSeq 500 with single-end 75 bp reads.
Oxidative bisulphite sequencing
Deep sequencing of PCR products from BS-converted and oxBS-converted DNA was performed as previously described . Briefly, precipitated DNA (without glycogen) was resuspended in water and further purified using Micro Bio-Spin columns (Bio-Rad), after which half of the DNA was oxidised with 15 mM KRuO4 (Alpha Aesar) in 0.5 M NaOH for 1 h. Following bisulphite conversion of both DNA fractions with the EpiTect Bisulfite kit (QIAGEN), a two-step PCR amplification was used: a first PCR amplifies the region of interest and adds part of the sequencing adaptors; a second PCR on pooled amplicons then completes the adaptors and adds sample barcodes, allowing for multiplexing (see primers in Additional file 3). Paired-end sequencing of pooled samples was done using an Illumina MiSeq.
Individual L1 elements
For RT-qPCR and oxBS analysis of individual L1s in the mouse reference genome, assays were first designed for several full-length elements with uniquely mapped reads and high 5hmC levels in TAB-seq data. We then selected elements whose RT and oxBS amplicons were validated by Sanger sequencing and for which the abundance in genomic DNA was close to that of single copy genes, as judged by qPCR. Elements have the following mm9 coordinates: chr2:125230942–125237372 (HL1-2; contains L1Tf monomers), chr5:70814798–70821450 (HL1-6; L1Tf monomers), chr6:107895070–107901677 (HL1-7; L1A monomers).
High-throughput sequencing data processing
External datasets were downloaded from GEO (Additional file 4). Reads were trimmed using Trim_galore! v0.3.3 with default parameters, except for small RNA-seq, where the options ‘-q 0 --length 15’ were used. Alignments were performed either to the whole mouse genome (mm9), to the L1Orl sequence (accession number D84391) or the L1.4 sequence (accession number L19092). ChIP-seq and small RNA-seq data were aligned using Bowtie2 v2.1.0  with default parameters, which assigns reads with multiple hits of equal mapping quality to one of those locations at random (inclusive mapping). To obtain uniquely mapped data, the output SAM file was filtered using a custom script. ChIP-seq peak detection was performed using the MACS algorithm implemented within Seqmonk. RNA-seq data were aligned using Tophat v2.0.9  with -g 1 option, which yields inclusive mapping. BS-seq and TAB-seq data were aligned to representative L1 elements (as above) using Bismark ; for genome-wide analysis, processed CpG calls from uniquely mapped data were obtained directly from GEO (see Additional file 4). OxBS data were aligned with Bismark to a custom genome containing the amplicon sequences; only CpGs covered by at least 100 reads were used to calculate 5mC/5hmC levels.
TET1 ChIP-seq analysis
To detect TE classes that were enriched for TET1 binding events, the number of TET1 peaks (from unique or inclusive mapping) overlapping each TE class (using the Repeatmasker annotation) were summed. This was compared with the overlaps obtained using a matched list of regions (equal number and lengths than the TET1 peaks) randomly distributed anywhere in the genome (for inclusive mapping) or within mappable regions of the genome, as defined by the uniqueome  (for unique mapping).
Raw read counts for each gene or repeat (from RepeatMasker annotation) were generated in Seqmonk with the RNA-seq quantitation pipeline. Reads from repeats belonging to the same class were pooled. DESeq2 was used for differential expression analysis and for generating normalised gene and repeat expression values. To couple TE-derived enhancers to gene expression values, promoter capture Hi-C data from ESCs , as processed by CHiCAGO  (kindly provided by Paula Freire-Pritchett, Babraham Institute), was used.
We thank the Barts Genome Centre for high-throughput sequencing; Claire Senner and Myriam Hemberger for their help with chromatin immunoprecipitation; Jesper Christensen and Kristian Helin for the TET2 antibody; Alex Bortvin for the ORF1p antibody; and Matthew Lorincz and Vardhman Rakyan for their critical reading of the manuscript.
MRB is a Sir Henry Dale Fellow (101225/Z/13/Z), jointly funded by the Wellcome Trust and the Royal Society. LR and OD have received funding from the People Programme (Marie Curie Actions) of the European Union’s Seventh Framework Programme (FP7/2007-2013) under REA grant agreement no. 608765. JH is funded by the Wellcome Trust (110216) and CC by the BBSRC.
Availability of data and materials
The datasets generated during the current study (RNA-seq and small RNA-seq) are available in the GEO repository under the accession number GSE84460. Accession numbers for previously published data used herein can be found in Additional file 4.
LR and MB designed the study and experiments. LR performed siRNA/shRNA knockdowns, RT-qPCR, western blotting, oxBS and ChIP experiments. OD performed OGT/SIN3A knockdowns and ChIP experiments. KC performed western blotting and qPCR analyses. CT designed bioinformatic analyses. CC and JH performed northern blotting. MB performed oxBS, RNA-seq and small RNA-seq experiments, and bioinfomatic analyses. LR and MB wrote the manuscript with all other authors. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
Ethics approval and consent to participate
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