RNA demethylase FTO uses conserved aromatic residues to recognize the mRNA 5′ cap and promote efficient m⁶A_m demethylation

Abstract

The RNA demethylase FTO acts as a methyl ‘eraser’ to remove either internal N⁶-methyladenosine (m⁶A) or 5′ end N⁶-2′-O-dimethyladenosine (m⁶A_m) modifications on mRNA. FTO has an intrinsic preference and significantly faster demethylation rates in vitro for m⁶A_m modifications located at the 5′ mRNA cap structure, but the structural basis for FTO’s ability to discriminate m⁶A versus m⁶A_m modifications has remained unknown. Here we utilize molecular dynamics simulations of FTO-RNA cap complexes to identify conserved aromatic residues on the surface of FTO involved in 5’ cap recognition. Subsequent mutagenesis and enzymology experiments validate the specificity of these residues in engaging the 5′ cap structure to promote m⁶A_m demethylation. We also identify a nonpolar surface on FTO that interacts with the 2′-O-methyl group of m⁶A_m to impact demethylation kinetics. This work provides the first structural insights into how FTO selectively catalyzes m⁶A_m versus m⁶A demethylation on mRNA, suggests why FTO is sensitive to different 5′ cap modifications, and furthers our understanding of how FTO activity is regulated by diverse mechanisms to help control the epitranscriptome.

Introduction

N⁶-methyladenosine (m⁶A) is the most abundant internal chemical modification of eukaryotic mRNA.^1,2 Installed co-transcriptionally by the METTL3/14 methyltransferase ‘writer’ complex, m⁶A can be found throughout the length of mRNA transcripts and is enriched near stop codons and in the 3′ UTR.^1–3 This widely important modification has impacts on mRNA splicing,^1,4–6 export,^7,8 translation,^9,10 and stability.^11,12 Notably, m⁶A plays a crucial role in the regulation of mammalian stem cell differentiation¹³ and is involved in controlling the expression of human oncogenes and tumor suppresser genes.^14–17 Another highly abundant mRNA methyl modification, N⁶,2′-O-dimethyladenosine (m⁶A_m), is found exclusively at the 5′ end of eukaryotic mRNAs in up to 30% of mammalian 5′ mRNA caps.¹⁸ m⁶A_m is formed by the writer enzyme CAPAM (or PCIF1), which carries out N⁶-methylation of 2′-O-methyladenosine (A_m) modifications.^19,20 While the functions m⁶A_m modifications are yet to be fully defined and may vary by cell type, recent work suggests these 5′ cap modifications likely play roles in regulating mRNA translation and splicing.^19,21–28 Together, the highly abundant methyl modifications m⁶A and m⁶A_m help control mRNA fate and function across the transcriptome.

Fat Mass and Obesity Associated Protein (FTO) is an Fe(II)/α-ketoglutarate-dependent dioxygenase enzyme that acts as an m⁶A demethylase or ′eraser′, converting m⁶A or m⁶A_m modifications to A or A_m, respectively, within cellular mRNA transcripts.^29,30 Along with other AlkB homologs in this family (e.g. AlkBH5), FTO allows for reversible m⁶A and m⁶A_m modification and the dynamic regulation of RNA methylation. Although FTO likely only acts on a subset of m⁶A/m⁶A_m modifications found on mRNA,^31–33 its m⁶A/m⁶A_m demethylation activity is linked to splicing,^34–36 cellular metabolism,^37–41 and mammalian development.^42,43 Furthermore, FTO and its methyl eraser functions have close links to the progression of human cancers, such as acute myeloid leukemia^44,45 and glioblastoma,^46,47 where FTO is highly expressed and promotes oncogenesis.^48–50

Although FTO has the ability to demethylate both internal m⁶A and 5′ cap m⁶A_m RNA modifications, previous studies have shown that FTO has a strong intrinsic preference in vitro for m⁶A_m demethylation.⁵¹ Remarkably, FTO has ~100 times greater catalytic efficiency for demethylating m⁶A_m compared to m⁶A modifications on RNA oligonucleotides.⁵¹ Furthermore, in model RNA substrates where the 5′ N⁷-methylguanosine (m⁷G) and/or triphosphate groups of the eukaryotic cap are removed, FTO-mediated demethylation of m⁶A_m is significantly impaired, suggesting that FTO somehow specifically recognizes the 5′ cap structure to promote m⁶A_m demethylation. An existing structure of 6mA-ssDNA in complex with FTO provides information about how the m⁶A base is accommodated in the FTO active site and how DNA (and likely RNA) binds a basic cleft on the surface of FTO.⁵² However, to date there is no structural or biophysical information about how FTO preferentially demethylates 5′ m⁶A_m modifications, how FTO might specifically recognize the eukaryotic 5′ cap structure and m⁷G or m⁶A_m bases, or the general mechanism(s) by which FTO selectively catalyzes removal of m⁶A_m versus m⁶A modifications on RNA.

Here we combine molecular dynamics (MD) simulations of FTO-m⁶A_m cap complexes with mutagenesis and enzymology experiments to identify conserved aromatic residues on the surface of FTO involved in 5′ mRNA cap recognition and selective m⁶A_m demethylation. We also identify nonpolar residues located just outside of the FTO active site that interact with the m⁶A_m 2′-O-methyl group to further mediate m⁶A_m demethylation. These experiments provide the first atomic-level information about how 5′ cap recognition promotes m⁶A_m demethylation by FTO, suggest why additional cap modifications (e.g. 2,2,7-trimethylguanosine) dramatically impair m⁶A_m demethylation,²³ and give new insights into the mechanisms controlling FTO-mediated mRNA demethylation selectivity.

Results

Molecular dynamics simulations identify conserved aromatic FTO surface residues involved in FTO-m⁷G cap interactions.

Previous biochemical studies have shown that FTO more efficiently demethylates 5′ m⁶A_m modifications as compared to internal m⁶A modifications on RNA.⁵¹ Additionally, removing the m⁷G portion of the 5′ mRNA cap structure on model RNA substrates results in a 50% reduction in m⁶A_m demethylation activity, and further removal of the cap triphosphate group reduces demethylation by another 40%. These data suggest that there must be FTO residues that selectively recognize the m⁷G and triphosphate groups adjacent to m⁶A_m and that these surfaces help to recruit and/or position 5′ m⁶A_m modifications for demethylation.⁵¹ To elucidate which residues and surfaces of FTO are interacting with the 5′ mRNA cap structure, we performed microsecond MD simulations of FTO in complex with the small 5′ cap molecule m⁷Gppp(m⁶A_m)pG (Figure 1A). We then surveyed the MD trajectories for m⁷G-FTO contacts ≤ 3.5 Å over all frames and determined the individual FTO residues that contact m⁷G with these criteria and a frequency of > 5% (Figure 1B, Supplementary Figure 1A).

Figure 1. MD reveals FTO residues with high frequency FTO-m⁷G interactions.

(A) m⁷Gppp(m⁶A_m)pG cap structure and initial FTO-m⁶A_m cap model where m⁶A_m was aligned into the FTO active site based on previously determined 6mA-ssDNA-FTO structure PDB 5ZMD. N⁶-methyl group on adenosine is shown in blue, 2′-O-methyl group on ribose is shown in red. (B) m⁷G-FTO residue contacts ≤ 3.5 Å with > 5% interaction frequency from MD simulations on the microsecond timescale. (C) Representative MD frame showing m⁷G stacking with conserved FTO residue H232 and hydrogen bonding with N302, A303, and D233 sidechain and backbone atoms on FTO surface; hydrogen bonds are shown as dashed yellow lines (top). In this binding mode m⁷G interacts with a highly conserved surface of FTO (bottom). (D) Representative MD frame showing m⁷G stacking on strongly conserved FTO residue W278. (E) Representative MD frame showing simultaneous interaction of H232 and W278, in which the m⁷G base stacks with W278 (as in panel D) and the m⁷G ribose interacts with H232.

Analysis of MD frames showing these high-frequency m⁷G-FTO contacts revealed two interesting putative m⁷G binding sites on the surface of FTO, and three binding modes. In the first, m⁷G of cap makes sustained contacts with FTO surface residue H232 (Figure 1C). Here, the positively charged m⁷G base can stack with H232 forming a cation-π interaction, while making extensive hydrogen bond contacts with backbone atoms of FTO residues N302, A303, and D233 (Figure 1C, top). A conservation analysis of FTO showed that H232, located adjacent to metal binding residues on the exterior surface of FTO, is a highly conserved residue (Figure 1C, bottom). The second high frequency m⁷G-FTO contact was observed with FTO residue W278 (Figure 1D). Here, m⁷G stacks with the highly conserved FTO residue W278, located on a flexible loop on the surface of FTO, forming a cation-π interaction (Figure 1D, top). Both the H232- and W278-mediated contacts are strong candidates for FTO surfaces that could mediate selective cap recognition and m⁶A_m demethylation by FTO. Still, a third binding mode was observed, during which H232 engages the m⁷G ribose while W278 simultaneously stacks with the m⁷G guanosine base (Figure 1E). In this mode, H232 contacts the m⁷G ribose via hydrogen bonds, CH-π, or Van der Waals interactions. Concurrent contact of H232 and W278 (occurring with a frequency of 0.14) was associated with a higher computed binding affinity than either residue alone, and represents the most favorable FTO-m⁷Gppp(m⁶A_m)pG interaction mechanism predicted by MD.

Mutations to conserved aromatic surface residues on FTO specifically impair m⁶A_m demethylation.

To experimentally validate the FTO-m⁷G interactions identified in the MD simulations described above, we mutated key, high-frequency m⁷G-interacting FTO residues to alanine and compared WT versus mutant demethylation activities on both an internal m⁶A-containing RNA substrate and the model cap substrate m⁷Gppp(m⁶A_m)pG (Figure 2A). LC-MS-based demethylation activity assays were conducted on the panel of FTO mutants to quantify relative demethylation activity after 150 minutes compared to WT FTO. These biochemical assays showed that while H232A, N302A, W278A, and H232A/W278A mutations had little to no defect on demethylation of a 9-mer m⁶A RNA substrate (Figure 2A, ‘m⁶A-RNA’), H232A, W278A, and H232A/W278A mutants all significantly impaired demethylation of m⁶A_m in the model cap substrate Figure 2A, ‘m⁶A_m cap’). Surprisingly, N302A had ~2-fold higher activity on m⁶A_m cap compared to WT FTO. Kinetic time courses comparing the production of demethylated A_m product from m⁷Gppp(m⁶A_m)pG substrate over time for WT versus H232 and W278 single and double mutants also shows a clear defect in FTO-mediated demethylation activity for H232A, and nearly abolished activity for W278A and H232A/W278A (Figure 2B).

Figure 2. Enzymatic assays validate FTO residues important for m⁶A_m demethylation.

(A) Mutation of conserved aromatic residues in FTO identified as m⁷G interaction sites show reduced m⁶A_m demethylation activity. Endpoint demethylation activity assays were conducted in triplicate with 2 μM CUGG(m⁶A)CUGG (m⁶A-RNA) or m⁷Gppp(m⁶A_m)pG (m⁶A_m cap) and 1 μM FTO enzyme. At 0 and 150 minutes, reaction samples were quenched with EDTA, digested to single nucleosides with RNAase cocktail, and the relative amounts of m⁶A or m⁶A_m were quantified by UPLC-MS. (B) Mutation of conserved aromatic residues in FTO identified as m⁷G interaction sites show reduced A_m production activity. Demethylation time courses were conducted with 2 μM m⁷Gppp(m⁶A_m)pG and 1 μM FTO enzyme in triplicate. At various time points from 0 to 150 minutes, reaction samples were quenched with EDTA, digested to single nucleosides with RNAase cocktail, and the relative amounts of A_m were quantified by UPLC-MS. (C) Mutation of conserved aromatic residues in FTO identified as m⁷G interaction sites show reduced m⁶A_m demethylation activity with a longer m⁶A_m cap RNA substrate. Endpoint demethylation activity assays were conducted with 2 μM m⁷Gppp(m⁶A_m)pCpUpGpG (m⁶A_m cap-RNA) and 0.1 μM FTO enzyme in triplicate. At 0 and 150 minutes, reaction samples were quenched with EDTA, decapped, digested to single nucleosides with RNAase cocktail, and the relative amounts of m⁶A_m were quantified by UPLC-MS. (D) H232/W278 double mutant of conserved aromatic residues in FTO identified as m⁷G interaction sites show reduced A_m production. Demethylation time courses were conducted with 2 μM m⁷Gppp(m⁶A_m)pCpUpGpG and 0.1 μM FTO enzyme in triplicate. At time points from 0 to 150 minutes, reaction samples were quenched with EDTA, decapped, digested to single nucleosides with RNAase cocktail, and the relative amounts of A_m were quantified by UPLC-MS.

We next tested the relative m⁶A_m demethylation activities for WT versus mutant FTO constructs on the longer capped model RNA substrate m⁷Gppp(m⁶A_m)pCpUpGpG, which more closely mimics a capped RNA. With this longer m⁶A_m-containing substrate we saw a small but measurable defect on m⁶A_m demethylation for mutation of FTO H232 and, very similar to the results above with the shorter cap substrate, large defects in activity with W278A and H232A/W278A mutants (Figure 2C). Consistent with these results, a time course comparing A_m product production for FTO WT versus H232A/W278A double mutant with m⁷Gppp(m⁶A_m)pCpUpGpG substrate, clearly shows a large defect for m⁶A_m demethylation activity on this longer capped substrate (Figure 2D). Together, these kinetic data show that mutation of FTO residues H232 and W278 significantly impair FTO-mediated demethylation activity of capped m⁶A_m-containing RNAs, but not internal m⁶A-containing RNAs, suggesting these two conserved FTO surface residues play a key role in recognizing the 5′ cap to promote efficient m⁶A_m demethylation.

MD simulations and kinetic assays identify FTO residues important for recognition of the 2′-O-methyl group in m⁶A_m.

Similar to m⁷G, the 2′-O-methyl group on adenosine (A_m) in the m⁶A_m cap structure has also been shown to be important for promoting FTO-mediated demethylation of m⁶A_m.⁵¹ In vitro activity studies indicate that absence of the A_m 2′-O-methyl group in model m⁷Gppp(m⁶A)-RNA substrates results in a 50% decrease in FTO’s ability to demethylate the 5′ m⁶A group as compared to m⁶A_m. However, there is no structural information about how FTO recognizes the A_m 2′-O-methyl group to help promote m⁶A_m demethylation. To explore this FTO-m⁶A_m interaction, we surveyed A_m methyl-FTO contacts ≤ 3.5Å over all frames in the FTO-m⁶A_m MD simulations and identified two FTO residues, L109 and I85, that contact the A_m 2′-O-methyl group with very high frequencies (Figure 3A, Supplementary Figure 1B). These strongly conserved nonpolar residues are located just outside the active site pocket and provide a hydrophobic surface on which the A_m 2′-O-methyl group frequently rests throughout the majority of the MD simulations (Figure 3B). Demethylation activity assays comparing WT FTO to L109A or I85A mutants revealed that while both of these mutations resulted in only small defects to m⁶A demethylation on an internal m⁶A-RNA, L109A resulted in a nearly 90% reduction in m⁶A_m cap demethylation activity (Figure 3C). Surprisingly, the FTO I85A mutation resulted in a ~50% increase in m⁶A_m cap demethylation relative to WT, suggesting this mutation may open up additional space on this surface or shallow pocket to accommodate the A_m 2′-O-methyl group. Together these experiments suggest that this nonpolar surface on FTO (L109, I85) plays a key role in interacting with the 2′-O-methyl group of the m⁶A_m base to help mediate demethylation.

Figure 3. Nonpolar FTO residues help recognize the A_m 2′-O-methyl group to promote m⁶A_m demethylation.

(A) Graph showing high frequency A_m 2′-O-methyl-FTO residue contacts from MD simulations on the microsecond timescale. (B) Representative frame from MD simulations showing close interactions of the m⁶A_m 2′-O-methyl group with the nonpolar surface just outside the FTO active site pocket composed of conserved residues L109 and I85. (C) FTO L109A and I185A mutations show altered demethylation activity for m⁶A_m cap compared to internal m⁶A-RNA substrates. Endpoint demethylation activity assays were conducted in triplicate with 2 μM CUGG(m⁶A)CUGG or m⁷Gppp(m⁶A_m)pG and 1 μM FTO enzyme.

Discussion

Previous biochemical studies have shown FTO has a clear intrinsic preference for ‘erasing’ m⁶A_m modifications located at the 5′ end of mRNA, as compared to internal m⁶A modifications.⁵¹ While the 6mA-ssDNA-bound FTO structure determined in 2019 provides the structural basis for m⁶A base recognition in the active site,⁵² it remains entirely unknown how FTO recognizes the eukaryotic 5′ cap structure and m⁶A_m 2′-O-methyl group to promote efficient 5′ m⁶A_m demethylation on mRNA. In this work, using MD simulations we identified conserved, aromatic FTO residues H232 and W278 as key surfaces for high frequency interaction with the m⁷G base of the m⁶A_m cap structure. Likewise, MD simulations show that nonpolar FTO residues L109 and I85 form a key interaction surface for the m⁶A_m 2′-O-methyl group. In vitro activity assays validate the structural predictions from MD, and show that mutating FTO residues to H232, W278, or L109 to alanine selectively impairs 5′ m⁶A_m demethylation, with only minimal effects on internal m⁶A demethylation. Together our MD and biochemical data provide the first atomic-level insights into how FTO recognizes the m⁷G base (H232/W278) and the 2′-O-methyl group (L109/I85) of 5′ m⁶A_m-modified mRNAs.

The MD simulations and enzymology data both support the conclusion that FTO surface residues H232 and W278 contribute to m⁷G binding, cap recognition and selective m⁶A_m demethylation. It appears that m⁷G and the 5′ cap dynamically sample multiple surfaces of FTO, as we observe in the MD simulations, and this leads to the importance of both H232 and W278 for cap recognition. Additionally, W278 is located on a flexible loop of FTO, and MD suggests that W278 may come close enough to H232 for both of these residues to interact simultaneously with the m⁷G base of the cap (Supplementary Figure 2A). Therefore, another possibility consistent with our experimental data is that FTO H232 and W278 could sandwich the m⁷G base, making dual cation-π interactions that specifically recognize the 5′ cap structure. This mode of recognition would be very similar to how canonical cap binding proteins, such as the cap binding complex (CPB20/80),⁵³ eIF4E,⁵⁴ or Dcp2,^55,56 recognize the mRNA cap (Supplementary Figure 2B) and would be most consistent with specific m⁷G recognition by FTO.

Furthermore, a ‘sandwiched’ m⁷G binding mode where m⁷G stacks with H232 (as in Figure 1C) and a loop conformational change then brings W278 nearby to make additional interactions, may explain a puzzling piece of biochemical data from the literature. Recently it was shown that N^2,2,7-trimethylguanosine (m^2,2,7G) capped RNAs, like those utilized during splicing, are entirely resistant to m⁶A_m demethylation by FTO;²³ this is odd because other studies have shown that removal of m⁷G entirely only reduces m⁶A_m demethylation activity by 50%.⁵¹ But when m⁷G stacks with H232 as shown in Figure 1C, the N2 nitrogen of guanosine points directly in towards the key metal binding active site residue D233. Dimethylation of the N2 atom as in m^2,2,7G, would sterically clash with D233, and be predicted to disrupt metal binding and inactivate FTO, consistent with the experimental observation that m^2,2,7G-capped RNAs have no m⁶A_m demethylation activity with FTO.

Our combined MD and biochemical studies reveal key, conserved residues on the surface of FTO that recognize the eukaryotic 5′ cap structure to promote efficient m⁶A_m demethylation on mRNA. These results provide a structural basis for FTO’s ability to distinguish 5′ m⁶A_m modifications from internal m⁶A RNA modifications, and suggest that dynamics of FTO residues and/or the 5′ cap may be important for selective cap recognition. Finally, we propose that engineered FTO mutants that specifically impair m⁶A_m demethylation but minimally affect m⁶A demethylation (e.g. H232A/W278A, H232A/W278A/L109A), could be used as separation-of-function mutants in cell-based studies to dissect and understand the different biological roles of FTO-mediated m⁶A_m versus m⁶A demethylation across the transcriptome.

Methods

Model construction.

An all-atom model of FTO in complex with the small 5′ cap molecule m⁷Gppp(m⁶A_m)pG was developed based on PDB 5ZMD.⁵² Missing loop segments in FTO were modeled using AlphaFold2.⁵⁷ The m⁷Gppp(m⁶A_m)pG cap structure was built with xleap⁵⁸ using residue templates from OL3.⁵⁹ The m⁶A_m base was aligned into the FTO catalytic pocket based on the 6mA orientation in the FTO-6mA-ssDNA structure. Crystallographic water molecules were maintained. PDB2PQR⁶⁰ was used to add hydrogen atoms to FTO appropriate for pH 7.0. Local ions were placed around the complex with cionize,⁶¹ and the complex was immersed in a 125 ų box of OPC⁶² water containing 150 mM NaCl.

Model parameterization.

FTO was treated with the ff19SB⁶³ force field. Manganese(II) coordination was parameterized using MCPB.py.⁶⁴ The N-oxalylglycine cofactor was treated with gaff2⁶⁵ parameters and AM1-BCC charges assigned by antechamber.⁵⁸ A custom residue was constructed for m⁷Gppp(m⁶A_m)pG. Parameters were taken from OL3⁵⁹ based on analogy to a GAG sequence, applying updated phosphate oxygen radii.⁶⁶ Methyl and triphosphate parameters were taken from gaff2.⁶⁷ Missing parameters for the methyl-purine moiety in m⁷G (two angles and an improper dihedral), as well as residue charges, were taken from the modified nucleoside force field,⁶⁸ which covers methylated RNA.

Molecular dynamics simulations.

MD simulations in the isothermal-isobaric (NPT) ensemble were performed with pmemd.cuda (SPFP precision model) in AMBER24.⁶⁹ The system was brought to a local energy minimum using 2,500 steepest descent cycles, then 2,500 conjugate gradient cycles. The temperature was raised from 60 K to 310 K over a period of 5 ns, then production sampling was carried out for 1 μs, saving frames every 1 ps. Dynamics were propagated with a timestep of 2.0 fs, enabled by constraining bonds to hydrogen with the SHAKE algorithm. Non-bonded interactions were partitioned into long-and short-range components based on a cutoff of 8.0 Å. System temperature was maintained at a target of 310 K with the Langevin thermostat using a collision frequency of 1.0 ps⁻¹, reinitializing the random number seed upon each simulation restart to avoid synchronization artifacts.⁷⁰ System pressure was maintained at a target of 1.0 bar via isotropic scaling with the Berendsen barostat using a relaxation time of 1.0 ps.

Trajectory analysis.

MD simulation trajectories were analyzed using VMD 1.9.4.⁶¹ Pairwise contacts between FTO and m⁷Gppp(m⁶A_m)pG were defined as being atoms ≤ 3.5 Å to filter for the most relevant interactions.

FTO WT and mutant expression vector construction.

A codon optimized gblock of a human FTO(32–505) construct was obtained from IDT and cloned into a pET28a bacterial expression vector with N-terminal 6xHis tag. The FTO single point mutations were introduced into the 6xHis-FTO(32–505) construct using whole plasmid PCR site-directed mutagenesis.

Protein expression and purification.

FTO WT and mutants were transformed into E. coli BL21(DE3) cells. 20 mL of an overnight culture was added to 2 L of LB media for growth at 37 °C with shaking at 200 rpm. When OD600 reached ~0.6, cells were induced with 1 mM isopropyl β-D-1-thiogalactopyranoside for 18 hours at 18°C with shaking at 200 rpm. Centrifugation at 4100 × g was used to harvest cells and pellets were flash frozen and stored at −70°C. Pellets were thawed and resuspended in lysis buffer (25 mM Tris-Base pH 8.0, 300 mM NaCl, 5 mM 2-mercaptoethanol), lysed by sonication, and centrifuged at 24000 × g for 45 min. Soluble His-tagged FTO was purified using HisPur Ni-NTA affinity resin (ThermoFisher Scientific) at 4°C and eluted in 25 mM Tris-HCl pH 7.5, 300 mM NaCl, 400 mM imidazole. Protein was buffer exchanged into 25 mM Tris-HCl pH 7.5, 25 mM NaCl and then purified using ion exchange chromatography with a 5 mL HiTrapQ column (Cytiva) and linear gradient into 25 mM Tris-HCl pH 7.5, 1 mM NaCl. Further purification was done with size exclusion chromatography using a Superdex 200pg 16/60 gel filtration column (Cytivia) in 10 mM HEPES pH 7.0, 50 mM KCl. Protein fractions were visualized using SDS-PAGE and purified FTO fractions were combined, concentrated, flash frozen, and stored in size exclusion buffer with 10% glycerol at −70°C.

FTO demethylation assays.

In vitro FTO demethylation activity assays were performed by incubating 2 μM substrate (m⁷Gppp(m⁶A_m)pG, m7Gppp(m⁶A_m)CUGG, or CUGG(m⁶A)CUGG) with either 1 μM FTO or 0.1 μM FTO in demethylation reaction buffer (50 mM HEPES pH 7.0, 150 mM KCl, 75 μM (NH₄)₂Fe(SO₄)₂·6H2O, 300 μM 2-oxoglutarate, and 2 mM ascorbic acid). Reactions were initiated by addition of a 2X substrate mixture in reaction buffer to a 2X enzyme and cofactor mixture in reaction buffer and mixed with pipetting. Reaction time points were quenched by addition of EDTA to a final concentration of 1 mM.

RNA nucleoside analysis and quantification.

Quenched reaction samples with substrate m⁷Gppp(m⁶A_m)CUGG were first decapped using NEB mRNA decapping enzyme (M0608S) for 1 hour. All quenched reaction samples were digested with NEB Nucleoside Digestion Mix (M0649S) for 1 hour (CUGG(m⁶A)CUGG) or 18 hours (m⁷Gppp(m⁶A_m)pG or m⁷Gppp(m⁶A_m)pCpUpGpG). Following RNA digestion into single nucleosides, for reaction samples with 1 μM FTO, protein was precipitated by addition of 20% TCA. Samples were then centrifuged and supernatant was collected for LC-MS analysis. Analysis of demethylation status was performed on an Agilent Bio-Inert 1260 Infinity II UHPLC system with Infinity Lab LC/MSD iQ. Separation of individual RNA nucleosides was conducted using an Agilent Zorbax SB-Aq Rapid Resolution HD (2.1 × 100 mm, 1.8 μm particle size) using mobile phase containing 0.1% formic acid (A) and 100% acetonitrile, 0.1% formic acid (B) and then detected by mass spectrometry in positive ionization mode. The gradient was as follows for detection of m⁶A at 0.480 mL/min (m/z = 282, 3.5 min) and A (m/z = 268, 1.6 min): 0–1.0 min, 100% A; 1.0–4.25 min, to 85% A/15% B; 4.25–5.0min, to 25% A/75% B; 5.0–7.0 min, 25% A/75% B; 7.0–7.1 min, to 100% B; 7.1–11.75 min, 100% B. The gradient was as follows for detection of m⁶A_m at 0.450 mL/min (m/z = 296, 3.65 min) and A_m (m/z = 282, 3.5 min): 0–1.0 min, 100% A; 1.0–1.1 min, to 40% A/60% B; 1.1–5.0 min, to 28% A/72% B; 5.0–5.1 min, to 100% B; 5.1.0–8.0 min, 100% B.

RNA substrate preparation.

9-mer m⁶A (CUGGm⁶ACUGG) was synthesized by IDT. Cap m⁶A_m RNA substrates (m⁷Gppp(m⁶A_m)G and m7Gppp(m⁶A_m)CUGG) were synthesized as previously described for similar RNA cap molecules.⁷¹