effect for selective detection of RNA and DNA m5C methyltransferase ...pharmacy.nus.edu.sg/wp-content/uploads/2019/11/gkz1047-2.pdf · effect for selective detection of RNA and DNA

Nucleic Acids Research, 2019 1doi: 10.1093/nar/gkz1047

A general strategy exploiting m5C duplex-remodellingeffect for selective detection of RNA and DNA m5Cmethyltransferase activity in cellsTianming Yang, Joanne J.A. Low and Esther C.Y. Woon *

Department of Pharmacy, National University of Singapore, 18 Science Drive 4, 117543 Singapore

Received July 20, 2019; Revised October 07, 2019; Editorial Decision October 21, 2019; Accepted October 30, 2019

ABSTRACT

RNA:5-methylcytosine (m5C) methyltransferases arecurrently the focus of intense research followinga series of high-profile reports documenting theirphysiological links to several diseases. However, nomethods exist which permit the specific analysisof RNA:m5C methyltransferases in cells. Herein, wedescribed how a combination of biophysical stud-ies led us to identify distinct duplex-remodelling ef-fects of m5C on RNA and DNA duplexes. Specifi-cally, m5C induces a C3′-endo to C2′-endo sugar-pucker switch in CpG RNA duplex but triggers aB-to-Z transformation in CpG DNA duplex. Inspiredby these different ‘structural signatures’, we devel-oped a m5C-sensitive probe which fluoresces spon-taneously in response to m5C-induced sugar-puckerswitch, hence useful for sensing RNA:m5C methyl-transferase activity. Through the use of this probe,we achieved real-time imaging and flow cytome-try analysis of NOP2/Sun RNA methyltransferase 2(NSUN2) activity in HeLa cells. We further appliedthe probe to the cell-based screening of NSUN2inhibitors. The developed strategy could also beadapted for the detection of DNA:m5C methyltrans-ferases. This was demonstrated by the developmentof DNA m5C-probe which permits the screening ofDNA methyltransferase 3A inhibitors. To our knowl-edge, this study represents not only the first exam-ples of m5C-responsive probes, but also a new strat-egy for discriminating RNA and DNA m5C methyl-transferase activity in cells.

INTRODUCTION

The DNA and RNA of all living organisms, as well as thatof viruses, mitochondria and chloroplasts, undergo a widerange of modifications (1–2). These modifications not onlyexpand the structural diversity of nucleic acids, but also pro-

vide an epigenetic mechanism to fine-tune their biologicalfunctions (3–4). To date, at least 160 naturally-occurringchemical modifications have been identified, amongst which5-methylcytosine (m5C) is currently one of the most inten-sively studied epigenetic modifications. The m5C modifica-tion is prevalent in DNA and multiple RNA classes (5–6).Its biological functions are best understood in DNA, whereit is involved in the regulation of gene expression, genomereprogramming, organismal development and cellular dif-ferentiation (5,7,8). In contrast, there has been compara-tively less studies on the biological roles of m5C in RNA.Indeed the significance of m5C modification in mRNA wasnot fully appreciated until recently, following the landmarkdiscovery of widespread m5C sites in the transcriptomes ofdiverse organisms (9–16), suggesting that the m5C modifica-tion is far more pervasive in human mRNA than previouslyrealised. This has reignited intense interest in the study ofthis epitranscriptomic mark. At present, the exact biolog-ical function of m5C modification in mRNA remains elu-sive although it has been linked to many cellular processes,such as nuclear export regulation (14), modulation of pro-tein translation (17–19), and stress response (20).

The m5C methylation landscape is regulated by a com-plex array of m5C methyltransferases (MTases) and m5Cdemethylases, which specifically add and remove a 5-methylmark from cytosine base, respectively (6,21–23). In hu-mans, C-5 cytosine methylation of DNA is catalysed byat least three DNA:m5C MTases (i.e. DNMT1, 3A and3B), whereas m5C methylation of RNA is catalysed by theNOL1/NOP2/Sun domain (NSUN) RNA methyltrans-ferase family, which includes NSUN1-7, as well as the DNAMTase homologue TRDMT1 (formerly DNMT2). No-tably, both the DNA and RNA m5C MTases employ a com-mon S-adenosyl-L-methionine (SAM) dependent mecha-nism for methyl group transfer, although different active-site cysteine residues are likely involved in the catalysis (24).

In recent years, it has become increasingly clear that aber-rant expression of m5C MTases may underlie the pathogen-esis of several diseases. For instance, DNMT3A, NSUN1,NSUN2 and NSUN4 were found to be overexpressed in anumber of human cancers, including breast, prostate, cervi-

*To whom correspondence should be addressed. Tel: +65 6516 2932; Fax: +65 6779 1554; Email: [email protected]

C© The Author(s) 2019. Published by Oxford University Press on behalf of Nucleic Acids Research.This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/), whichpermits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.

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2 Nucleic Acids Research, 2019

cal, and colorectal cancers (25–29). NSUN3 is linked to mi-tochondrial diseases (30,31), whilst NSUN5 and NSUN7are likely implicated in Williams-Beuren syndrome (a neu-rodevelopmental disorder) (32) and male infertility (33), re-spectively. Given the strong physiological links, there is cur-rently immense interest in studying the biological roles ofthese enzymes and their potentials as therapeutic targets.However, at present, no methods exist which permit the di-rect analysis of specific m5C MTase in living cells. There arealso no reports of cell-based assays capable of discriminat-ing between DNA and RNA MTase activities. This severelyimpedes our understanding of these medically-importantenzymes.

It is known that several nucleic acid base methyla-tions can directly impact the secondary structure of DNAand RNA. For instance, we (34–36), and others (37–40), have recently shown that the presence of a singleN6-methyladenosine (m6A) or N1-methyladenosine (m1A)modification can trigger a duplex-hairpin transformationin certain RNA sequence contexts. It is also clear from anumber of NMR and X-ray crystallographic studies thatm5C modification is able to induce a major B–Z structuralchange in some DNA sequences. This phenomenon was firstreported by Behe and Felsenfeld (41), and subsequently byothers (42–44), for a number of DNA duplexes with alter-nating CpG sequences, such as d(m5CGCGm5CG), polyd(m5CG)n and poly d(Gm5C)n. The structural influenceof m5C on RNA duplexes, however, is significantly less-studied. One early study on a poly(CG) RNA sequence sug-gests that m5C methylation likely elicit a conformationalchange from an A-type duplex to an atypical duplex struc-ture (45), whereas a separate study using a different se-quence context indicates no significant structural alteration(46). To the best of our knowledge, these two reports pro-vide the only experimental data available in this regard and,to date, it remains unclear whether m5C modification hasany major impact on the secondary structures of RNA du-plexes.

Herein we demonstrated, through a combination ofNMR, circular dichroism (CD), and thermodynamic stud-ies, that m5C has different conformational effects on DNAand RNA duplexes, even for duplexes with identical CpGsequence. In the case of CpG RNA duplex, it induces aconsiderable distortion of the phosphate backbone and aC3′-endo to C2′-endo sugar pucker switch in the terminalresidues whereas, in the case of CpG DNA duplex, m5Ctriggered a remarkable B-to-Z structural transformation.m5C therefore produces distinctly different ‘structural sig-natures’ on CpG RNA and DNA duplexes under the samephysiologically-relevant conditions.

Inspired by these interesting findings, we envisioned thatthe different duplex-remodelling effects of m5C could pro-vide a basis for the selective detection of RNA and DNAm5C MTase activity in cells. This was demonstrated bythe development of a novel m5C-sensitive nucleic acidprobe which, by design, is capable of switching its termi-nal sugar pucker conformation spontaneously in responseto m5C methylation (Figure 1). When coupled with anenvironment-sensitive fluorophore, it provides a powerfulvisual tool for sensing RNA:m5C methylation changes incells. Prior to this work, we are not aware of any assay

methods which are based on m5C-induced conformationalchange.

As we shall demonstrate, the m5C-probe is highly-selective and could specifically target NSUN2 over otherRNA:m5C MTases (including structurally-related subfam-ily members NSUN3, NSUN5A, NSUN6) and DNA:m5CMTases (including DNMT1 and DNMT3A). Through them5C-probe approach, we achieved live cell imaging and flowcytometry analyses of NSUN2 activity in HeLa cells. Wealso successfully applied the probe to the high-throughputcell-based screening of NSUN2 inhibitors. The discovery ofsuch highly selective probes is rarely achieved and may pro-vide insights on the functions of NSUN2 in m5C-regulatedprocesses. Although this study focus on the sensing ofRNA:m5C MTase activity, the m5C-switchable probe strat-egy outlined here could also be adapted for the study ofDNA:m5C MTase. This was demonstrated by the success-ful development of DNA m5C-probe which is useful for thein vitro screening of DNMT3A inhibitors.

To the best of our knowledge, this study representsnot only the first examples of m5C-responsive fluorescentprobes, but also a new strategy for the discrimination ofRNA and DNA m5C MTases activity in cells, which hope-fully will inspire the development of epigenetic biosensorsand diagnostics tools.

MATERIALS AND METHODS

Preparation of m5C-probes

All oligonucleotide probes investigated in this studywere synthesized using standard �-cyanoethyl phospho-ramidite chemistry on an automated DNA/RNA synthe-siser (Applied Biosystems 394). All synthesiser reagentswere purchased from Glen Research; 2′-O-methyl 6-phenylpyrrolocytidine phosphoramidite (47,48) and pyrenedeoxynucleoside phosphoramidite (49) were synthesised ac-cording to literature procedure. For details of synthesis, seeSupplementary Data. All oligonucleotides have a purity ofat least 95% (Supplementary Tables S1-S3).

Studies on the duplex-remodelling effects of m5C usingNMR, CD and UV-based thermodynamic analyses

All 1H NMR experiments were recorded on a Bruker DRX-500 spectrometer operating at 500.23 MHz and the dataprocessed using the standard Bruker processing softwareTopSpin. Estimated accuracy for protons is within 0.02ppm. The oligos were dissolved in either D2O (99.98%) orbuffer/D2O mix; the buffer used was 10 mM sodium phos-phate buffer (pH 7.4) containing 150 mM NaCl and 20 mMMgCl2. Final concentrations were 3 mM for the unmethy-lated duplexes (1a, 7a) and 2 mM for the m5C-methylatedduplexes (1b, 7b). The proton chemical shifts were refer-enced relative to residual H2O signal (4.75 ppm at 25◦C).The 1H imino spectra of duplexes in 9:1 buffer/D2O solventmix (pH 7.4) were acquired at 4◦C and 10◦C with suppres-sion of water signal by gradient-tailored excitation (WA-TERGATE) sequence, as previously described (34,50); atthese temperatures, the oligos exist in the duplex state, asverified by UV-melting and CD analyses. Two-dimensional

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Nucleic Acids Research, 2019 3

Figure 1. The m5C-switchable probe strategy. (A) The m5C-probe 8a contains a 5′-terminal fluorescent nucleotide PC (2′-O-methyl 6-phenylpyrrolocytidine), which lights up spontaneously in response to m5C-induced terminal sugar pucker switch. (B) When the probe is unmethylated,pC is able to base-pair with guanine in the complementary strand and stack strongly with its adjacent base. This results in efficient quenching of pC flu-orescence through photoinduced electron transfer. m5C methylation of the probe by RNA:m5C MTase (e.g. NSUN2), however, is expected to trigger aC2′-endo to C3′-endo sugar pucker switch in PC and, since the sugar ring pucker defines the glycosidic bond angle, such a change in sugar puckering willalso convert the orientation of PC base from axial to equatorial. This, in turn, disrupts its base-pairing and base-stacking interactions, leading to fluores-cence activation. (C) Schematic representation of 2′-OMe RNA probes and their methylated counterparts. The composition of the probes was confirmedthrough MALDI-TOF mass spectrometric analysis (see Supplementary Table S2). The structure of ‘locked 6-phenylpyrrolocytidine’ (LC) is shown. TheC2′−C4′ covalent link is geometrically incompatible with C2′-endo pucker mode, it therefore locks the nucleotide into the C3′-endo conformation.

1H NMR spectra of duplexes in D2O and buffer/D2O wereacquired in the phase-sensitive mode using previously de-scribed method (51). Two-dimensional NOESY spectra ofduplexes in D2O (52–54) were recorded at 25◦C with a spec-tral width of 4000 Hz using 1000 t2 complex points, 1000t1 (real) points, pulse repetition delay of 2.5 s and mixingtimes (� m) of 80, 150 and 300 ms. Two-dimensional NOESYspectra of duplexes in 9:1 buffer/D2O were acquired at 10◦Cwith water suppression by WATERGATE sequence (34,50).Spectra were recorded using 2048 t2 complex points, 256t1 (real) points, pulse repetition delay of 1.7 s and mixingtime of 200 ms. Proton-decoupled 31P NMR spectra of du-plexes in 9:1 buffer/D2O were acquired at 10◦C on a BrukerAV300 spectrometer equipped with an autotune 5 mm QNPprobe. Spectra were recorded at 121.4 MHz, with a spectral

width of 10 kHz. The 31P chemical shifts were externallyreferenced to 85% phosphoric acid (H3PO4). Estimated ac-curacy for phosphorus is within 0.03 ppm. For CD spec-troscopy and UV-based thermal denaturation experiments,see Supplementary Data.

Fluorescence analyses of m5C-probes

Fluorescence analysis was performed using a Cary Eclipsefluorescence spectrophotometer, as previously reportedwith modifications (36). The fluorescence emission spectra(�em = 400–600 nm) of the probes were recorded in a quartzcuvette at 37◦C, at a total stand concentration 5 �M in abuffer of 10 mM sodium phosphate buffer (pH 7.4) contain-ing 150 mM NaCl and 20 mM MgCl2. An average of five

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scans was recorded. Background fluorescence spectra wereacquired after the probe has been incubated for 30 min at37◦C. For determination of fluorescence quantum yields ofthe m5C-probes, see Supplementary Data.

In vitro m5C-probe methyltransferase assay

RNA methyltransferase assay: the assay was performed at37◦C in triplicate in a Corning Costar 96-well flat bot-tom black plate. Reaction consisted of enzyme (0.5 �M),methyl donor S-adenosyl-L-methionine (SAM; 200 �M),m5C-probe 9a (substrate; 5 �M) or locked-probe 10a (con-trol; 5 �M) in a 50 mM HEPES buffer (pH 7.4) contain-ing 150 mM NaCl and 20 mM MgCl2. The final reactionvolume is 25 �l. The time course of fluorescence activationwas recorded immediately after the addition of enzyme witha Tecan ultra microplate reader (�ex 360 nm; �em 465 nm).An average of five scans was recorded. DNA methyltrans-ferase assay: the assay was performed as described above.Reaction consisted of enzyme (NSUN2 or DNMT3A; 0.5�M), methyl donor S-adenosyl-L-methionine (SAM; 200�M), DNA m5C-probe 11a (substrate; 5 �M) in a 50 mMHEPES buffer (pH 7.4) containing 150 mM NaCl and 20mM MgCl2. The fluorescence of the reaction was moni-tored at �em 400 nm (�ex 340 nm).

Steady-state kinetic analyses of the methylation of m5C-probe, tRNA and ssRNA by NSUN2

The substrates investigated include m5C-probe 9a, hu-man tRNALeu(CAA) (5′-CCAGACUCAAGUUCUGG-3′), and CpG-rich ssRNA (5′-CGCGCGCGCGCG-3′).The kinetic constants (Vmax, Km, kcat) of the substrate weredetermined using an NSUN2 concentration of 0.1 �M andsubstrate concentrations of 1, 2, 3, 5, 8 and 10 �M. Theamount of m5C-methylated product formed at differentsubstrate concentrations was determined based on the rela-tive intensities of the substrate and the product peaks ob-served in MALDI-TOF mass spectra. All reactions wereperformed at 37◦C in triplicate.

Fluorescence microscopy

The HeLa cells were grown in 35 mm glass bottom cul-ture dishes (Thermo Scientific). The cells were transfectedwith either m5C-probe 9a (10 �M) or locked-probe 10a (10�M; control) using Lipofectamine 2000 (Invitrogen) follow-ing the manufacturer’s protocols. Fluorescence images werethen acquired at 37◦C in the presence of 5% CO2 using aNikon BioStation IM-Q live cell imaging system equippedwith 20× and 63× oil immersion objective lens. The fluo-rescence images were captured using the DAPI filter setting(i.e. �ex 340–380 nm; �em 435–485 nm) and the fluorescenceintensity quantified using Biostation IM multichannel soft-ware by collecting mean gray values for regions exhibitingfluorescence. All experiments were performed in triplicatesand the mean fluorescence intensities were normalised tothat of the control. The corresponding transmitted light im-age for each fluorescence image was also acquired.

Single-cell flow cytometry analysis

The HeLa cells were transfected with m5C-probe 9a (10�M) or control probe 10a (10 �M) using Lipofectamine2000. They were then collected by trypsinization at 37◦Cwith 5% CO2, washed with phosphate buffered saline (PBS)media, pH 7.4, resuspended in PBS media, and analysed us-ing a BD LSRFortessa™ flow cytometer (BD Bioscience).Fluorescence was measured using an excitation laser of 355nm, and a 450/50 bandpass emission filter. Acquisition wasstopped when 20, 000 events per sample were acquired. Thefluorescence data were then analysed using BD FACS soft-ware. Violin plot was produced using the Origin software.

Cell-based m5C-probe methyltransferase assay

The HeLa cells were pre-incubation with various concen-trations of non-specific MTase inhibitors, including sine-fungin, S-adenosyl-L-homocysteine (SAH), adenosine, andhomocysteine at 37◦C for 30 min. Nine different concentra-tions of inhibitors were used (0.1, 0.3, 1, 3, 10, 50, 100, 250,1000 �M) and the final DMSO concentration was kept atless than 1% (v/v) of the assay mix. These concentrations ofinhibitors were shown to be non-toxic to HeLa cells in MTTcytotoxicity assays. This was followed by the delivery of ei-ther the m5C-probe 9a (10 �M) or control probe 10a (10�M) into the cells via Lipofectamine 2000. Aliquots werethen collected for flow cytometry measurements (�ex 355nm; �em 425–475 nm) at the indicated time points. The meanfluorescence intensity of at least 20 000 live cells was deter-mined. IC50 determination: the IC50 values were then calcu-lated from the variation in fluorescence at different inhibitorconcentrations using nonlinear regression, with normaliseddose-response fit on GraphPad Prism 6.0™. The assay wasperformed in triplicate for each inhibitor concentration. Z’factor determination: the Z’ factor for our m5C-probe assaywas calculated using previously described method (55). Fordetails, see Supplementary Data.

Cell culture, gene silencing and gene overexpression

HeLa cells (ATCC) were cultured in Dulbecco’s modifiedEagle’s medium (DMEM; Life Technologies) supplementedwith 10% fetal bovine serum (FBS) and antibiotics (100units/ml penicillin, 100 �g/ml streptomycin). Cells weregrown at 37◦C in a 5% CO2 humidified incubator. To silencethe expression of specific gene transiently, the cells were firstplated in a 24-well plate (0.5 × 105 cells/well). The next day,the cells were transfected with either siNSUN2, siNSUN6,siTRDMT1, siDNMT3A or siControl using LipofectamineRNAiMAX (Invitrogen), according to manufacturer’s pro-tocols. The sequences used for siRNAs and siControlsare provided in the Supplementary Data. The protein lev-els were analysed by western blot using specific antibod-ies. Overexpression of specific gene. The construction ofpcDNA3-Flag vectors expressing NSUN2, TRDMT1 andDNMT3A was previously described (56–58). Full lengthNSUN2, TRDMT1 and DNMT3A genes were amplifiedby PCR using human HeLa cells cDNA, then subclonedinto the EcoRI-BamHI sites of mammalian pcDNA3-Flag vector (Invitrogen) to obtain pcDNA3-Flag-NSUN2,pcDNA3-Flag-TRDMT1, and pcDNA3-Flag-DNMT3A

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plasmids, respectively. The plasmids were then transfectedinto HeLa cells using the calcium chloride transfectionmethod. For construction of pcDNA3-Flag-NSUN6 plas-mid (59), the cDNA encoding NSUN6 was PCR-amplifiedusing Phusion DNA polymerase and subcloned into theKpnI-XhoI sites of pcDNA3.1-Flag with CloneExpress IIOne step Cloning Kit (Vazyme). The primer sequences usedfor vector construction are given in Supplementary Data.

Western blot analysis

The HeLa cells were first lysed using radioimmunopre-cipitation assay (RIPA) buffer (i.e. 25 mM Tris–HCl (pH7.6), 150 mM NaCl, 0.1% SDS, 1% sodium deoxycholate,1% NP-40 and 1× protease inhibitors), followed by im-munoblotting using standard protocol. In brief, the extractwas fractionated by sodium dodecyl sulfate-polyacrylamidegel electrophoresis (SDS-PAGE), transferred onto nitrocel-lulose filter membranes (Whatman), then incubation withthe respective antibodies. This was followed by incubationwith either horseradish peroxidase (HRP)-conjugated anti-rabbit IgG antibody (Bio-Rad) or HRP-conjugated anti-mouse IgG antibody (Bio-Rad) for 2 h at room temper-ature. Enhanced chemiluminescence substrates (LuminataCrescendo, EMD Millipore) were then applied and the sig-nals exposed to autoradiography film. The immunoblotswere then quantified by densitometric analyses using Im-ageJ software. All antibodies used were purchased fromcommercial sources (for details, see Supplementary Data).

RESULTS AND DISCUSSION

m5C induces a local distortion of the phosphate backbone anda C3′-endo to C2′-endo sugar pucker switch in CpG RNA du-plex

To date, there has not been a systematic study directly com-paring the structural impact of m5C methylation on CpGRNA duplexes and CpG DNA duplexes. Such an analysismight provide insights into the respective conformationaland/or mechanistic roles of m5C in DNA and RNA, henceis of special interest. To investigate this question in multi-ple sequence contexts, we prepared a set of 6-mer and 12-mer RNA duplexes containing either alternating CpG se-quences (1a–2a), out-of-alternation CpG sequence 3a orrandom sequences (4a–6a), as well as their correspondingm5C-methylated counterparts (1b–6b; Table 1). To facilitatecomparison, the CpG DNA duplexes 7a and 7b were alsogenerated. For details of their chemical synthesis, see Sup-plementary Data.

We began by analysing the secondary structure of CpGRNA hexamer r(CG-C-GCG)2 1a and its methylated equiv-alent r(CG-m5C-GCG)2 1b using 1H and 31P NMR. As-signment of the proton resonances was accomplished by2D NOESY experiments and by comparison with reportedNMR data for 1a (60). The results are summarised in Figure2 and Table 2. Consistent with previous NMR studies (60),the imino proton spectrum of 1a exhibited three NH signalsat 10◦C, which is expected for a palindromic duplex with sixbase-pairs (Figure 2A). Upon m5C methylation (1b), we ob-served a downfield shift in G2 resonance by ∼0.35 ppm anda disappearance of the terminal G6 resonance. The latter is

likely due to ‘fraying’ of the terminal base-pairs as the G6signal could again be detected at a lower temperature of 4◦C(Supplementary Figure S1). This result suggests that m5Cmodification likely promotes the disruption of base-pairinginteractions at duplex terminal ends. As anticipated, thereis little or no change in the imino resonance of G4 whichis involved in base-pairing with m5C, confirming that C5-cytosine methylation does not hinder C:G Watson–Crickbase-pairing interactions. Nevertheless, we did observe con-siderable variations in 31P resonances between the methy-lated and unmethylated duplexes, indicating a significantdistortion of the phosphate backbone (Figure 2B).

Prompted by this finding, we proceeded to examinewhether changes in phosphate backbone also alter the sugarring geometry. To this end, we determined the coupling con-stant between sugar proton H1′ and H2′ (3JH1′−H2′), whichis highly sensitive to the dihedral angle and, therefore, pro-vides a direct indication of the sugar pucker conformation.In general, a 3JH1′−H2′ value <2 Hz is suggestive of a C3′-endo sugar pucker (North type), whereas 3JH1′−H2′ value >8Hz is characteristic of a C2′-endo pucker (South type) (61).The 3JH1′−H2′ values for 1a and 1b are summarized in Table2 and the representative spectra are shown in Supplemen-tary Figure S2. In the absence of cytosine methylation, all ri-bose rings in 1a exhibit a C3′-endo conformation (3JH1′−H2′

∼1–2 Hz), which is the preferred sugar puckering mode forA-form RNA duplex. Upon m5C methylation (1b), how-ever, the 5′-terminal cytosine (C1) undergoes a sugar puckerswitch from C3′-endo to C2′-endo, as evidenced by a largeincrease in 3JH1′−H2′ from 1.1 Hz to 8.2 Hz. This is accom-panied with a switch in sugar puckering of the 3′-terminalguanosine (G6) to an intermediate conformation betweenC3′-endo and C2′-endo (3JH1′−H2′ = 6.0 Hz). No changesin sugar pucker mode are observed for the four internalresidues G2 to C5, which continue to adopt a C3′-endo ori-entation. Our data support the findings of an earlier NMRstudy (45) and, together, they demonstrate that m5C can al-ter the sugar pucker conformation of the 5′- and 3′-terminalresidues in CpG RNA duplexes.

To confirm our results, we further analysed the intensityof intranucleotide NOEs between the base protons (H6 incytosine; H8 in guanine) and the sugar protons (H2′ andH3′), which correlates with the distance between the pro-tons (Figures 2C, D and Table 2) (62,63). Consistent withour 3JH1′−H2′ data, the C1 residue of 1b displays a strongNOE between its base proton H6 and sugar proton H2′(∼2.2 A) and a weak NOE between H6 and H3′ (∼3.9 A),which is indicative of a C2′-endo conformation, and the G6residue of 1b shows approximately similar NOE intensitiesfor H8-H2′ (∼2.7 A) and H8-H3′ (∼3.3 A), suggesting anintermediate C2′-endo/C3′-endo conformation. In compar-ison, both the C1 and G6 residues of unmethylated 1a ex-hibit a C3′-endo sugar pucker, thus m5C methylation indeedproduces a switch in the terminal sugar pucker orientation.

Notably, however, these structural perturbations appearto have little or no effect on the overall secondary struc-ture of the RNA duplex. This is apparent from CD anal-ysis where the spectra of 1a and 1b exhibit characteristicswhich are typical of a right-handed A-form duplex, andthey superimpose almost perfectly with each other (Figure3A). Hence, m5C modification likely only affects the local

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Table 1. Sequences of RNA and DNA duplexes investigated in this study. The 3JH1′−H2′ values for the 5′- and 3′-terminal residues of each sequence areindicated in red and green, respectively. m5C-induced terminal sugar pucker switch was observed in 6-mer and 12-mer CpG duplexes (1b and 2b) but notin out-of-alternation CpG duplex 3b and random duplexes 4b–6b, suggesting that the structural effect of m5C may be unique to alternating CpG RNAsequences. m5C methylation of 7a i.e. the DNA equivalent of 1a caused a C2′-endo to C3′-endo sugar pucker switch in all guanosine residues, whilst allcytosine residues preserved their C2′-endo orientation, consistent with a B-Z structural conversion

aThe composition of the oligos was confirmed through MALDI-TOF mass spectrometric analysis (see Supplementary Table S1).bObserved at 25◦C in D2O.

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Figure 2. m5C methylation directly alters the phosphate backbone in CpG RNA duplex, leading to a C3′-endo to C2′-endo sugar pucker switch of theterminal residues under physiologically-relevant conditions. (A) Representative 1H imino NMR spectra of r(CG-C-GCG)2 1a (top) and its methylatedcounterpart r(CG-m5C-GCG)2 1b (bottom) in 9:1 buffer/D2O solvent mix acquired at 10◦C. The buffer used was 10 mM sodium phosphate buffer(pH 7.4) containing 150 mM NaCl and 20 mM MgCl2. The G6 imino resonance disappeared upon m5C methylation, presumably due to ‘fraying’ ofthe terminal base-pairs. (B) Proton-decoupled 31P NMR spectra of 1a (top) and 1b (bottom) in 9:1 buffer/D2O solvent mix acquired at 10◦C. There isconsiderable variations in 31P resonances between the methylated and unmethylated duplexes, suggesting a significant distortion of the phosphate backbone.Representative 2D NOESY spectra of the H2′/H3′-aromatic region of (C) 1a and (D) 1b in D2O at 25◦C (pH 7.4; �m = 150 ms) showing the H6/8-H2′NOE connectivity pathways (blue dashed-line) and the H6/8-H3′ NOE connectivity pathways (black solid line). The schematic illustrations (left) showedthe likely sugar pucker mode for terminal residues C1 and G6, and their calculated intranucleotide base-sugar distances (labelled in red; determined basedon the intensity of NOEs between the base and sugar protons).

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Table 2. 1H NMR chemical shifts �H (ppm) for CpG RNA duplex 1a (top) and its m5C-methylated counterpart 1b (bottom) observed at 25◦C in D2Owith WATERGATE suppression (�m 150 ms). Assignment of the proton resonances was accomplished by 2D NOESY experiments, and by comparisonwith reported NMR data for 1a (60). The representative NMR spectra for the determination of 3JH1′−H2′ coupling constants are given in SupplementaryFigure S2. For calculation of sugar-base proton distances, see Supplementary Data

aObserved at 10◦C in 9:1 buffer/D2O solvent mix. The buffer used was 10 mM sodium phosphate buffer (pH 7.4) containing 150 mM NaCl and 20 mMMgCl2.bChemical shift for 5-methyl group on m5C.cNot observed even at 4◦C.

Figure 3. m5C methylation induces distinctly different conformational ef-fects on RNA and DNA duplexes. Superimposition of the CD spectra of(A) CpG RNA duplex 1a with that of its m5C-methylated counterpart 1b,and (B) CpG DNA duplex 7a with that of its m5C-methylated equivalent7b. Under physiologically-relevant salt and pH conditions, m5C methyla-tion has negligible effect on the overall conformation of 1a. In sharp con-trast, methylation of 7a triggered a transformation from right-handed B-DNA to left-handed Z-DNA, as apparent from the characteristic inversionof its CD spectrum.

nucleotide geometry without altering the overall conforma-tion of CpG RNA duplexes.

m5C-induced terminal sugar pucker switch is likely unique toalternating CpG RNA sequences

We next examined whether m5C could also elicit terminalsugar pucker switch in other RNA sequence contexts. Tothis end, we determined the 3JH1′−H2′ coupling constant val-ues for the 5′- and 3′-terminal residues of each sequence(Table 1). Our results revealed a similar C2′-endo to C3′-endo switch in the longer 12-mer CpG duplex 2b, but notin the out-of-alternation CpG sequence 3a and random se-quences 4a–6a. On the basis of these results, we performedfurther NMR experiments to investigate how the position of

m5C methylation affects terminal sugar pucker conforma-tion in alternating CpG duplexes 1a and 2a. The 3JH1′−H2′

values are summarised in Supplementary Table S4. For 6-mer CpG duplexes, terminal sugar switch occurred onlywhen the third cytosine residue (C3) was methylated (i.e.1b), and not when the first (1c) or second (1d) cytosineresidue was methylated. Similarly for 12-mer CpG duplexes,m5C-induced sugar switch was observed only when methy-lation was on C3 (2b), and not on C1 (2c) or C5 (2d), thusthe structural effect of m5C is highly-dependent on the po-sition of m5C methylation.

The sequence requirements and mechanisms by whichm5C induces terminal sugar pucker switch are unclear atpresent and require further investigations. Nevertheless, thepresent study, the first to investigate the structural influenceof m5C on multiple RNA sequence contexts, suggests thatthis conformational effect might be unique to alternatingCpG RNA sequences.

m5C triggers a B-to-Z DNA transformation in CpG DNAduplex

We next investigated the structural influence of m5C onCpG DNA duplex 7a, i.e. the DNA equivalent of 1a un-der the same physiologically-relevant conditions. A prelim-inary analysis of the 3JH1′−H2′ values and sugar-base protondistances of 7a and 7b revealed a switch in sugar puckeringof all guanosine residues from C2′-endo to C3′-endo mode,whilst all cytosine residues preserve their C2′-endo orienta-tion, which is highly suggestive of a B-to-Z structural trans-formation (Table 3; the representative 2D NOESY spec-trum of 7b is shown in Supplementary Figure S3). As fur-ther evidence, CD analysis showed a characteristic inversionof the CD spectrum upon m5C methylation, confirming that

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m5C does indeed triggers a major B-Z structural transfor-mation in CpG DNA duplex 7a (Figure 3B). This result isnotable because it demonstrates that m5C produces differ-ent structural outcomes on RNA and DNA duplexes.

The observed B-Z conversion is likely attributed to theability of m5C to stabilize the Z-form DNA over its B-formstructure (42,64). Indeed, UV-based thermodynamic analy-sis of 7a showed that m5C methylation increases the stabil-ity of its Z-duplex by ∼2.5 kcal/mol relative to its B-duplex(compare �G◦

310 of 7a with 7b; Supplementary Table S3and Supplementary Figure S4). Moreover, m5C-induced B-Z helicity change is primarily an entropy-driven process,as judged by the more favourable entropic contributions(��S◦

7a→7b = 31.6 cal/mol/K) which counteracts the en-thalpy cost (��H◦

7a→7b = 7.3 kcal/mol). Further thermo-dynamic analysis of 7b in the absence of MgCl2 (a knownZ-conformation inducer) revealed a free energy of B-to-Ztransition (��G◦

T) of ∼1.5 kcal/mol (compare �G◦310 of

7b with 7b*); since 7b does not undergo B-Z conversion inthe absence of MgCl2, any free energy differences observedat these two salt conditions, i.e. between 7b and 7b* is largelyascribed to conformational transition (Supplementary Ta-ble S3 and Supplementary Figure S5).

m5C does not promote A–Z transition in CpG RNA duplex

To date, the effects of m5C methylation on the stability ofZ-form RNA is unclear. We are also not aware of any priorstudies investigating whether m5C could promote an anal-ogous A–Z transition in RNA duplex. Therefore, we per-formed further CD experiments to monitor the conforma-tion of CpG RNA duplex 1a and its methylated equiva-lent 1b at a range of MgCl2 concentrations (1–300 mM).The results showed that both 1a and 1b maintain an A-type double helical structure at all MgCl2 concentrationsinvestigated (Supplementary Figure S6). In fact, the methy-lated RNA 1b could not be induced to form Z-RNA even atMgCl2 concentration as high as 300 mM. Consistent withthis observation, the presence of m5C does not significantlyimprove the stability of 1a (��G◦

310 ∼0.8 kcal/mol; Sup-plementary Table S3). Thus m5C methylation favours B–Zconversion in CpG DNA duplex, but not A–Z transitionin CpG RNA duplex under the same physiological salt andpH conditions.

Overall, our results demonstrated that m5C methylationcan directly influence the conformation of RNA and DNAduplexes. In the case of CpG RNA hexamer, m5C modifica-tion induces a local distortion of the phosphate backboneand a C3′-endo to C2′-endo terminal sugar pucker switchwhereas, in CpG DNA hexamer, m5C triggers a transforma-tion from a right-handed B-DNA to a left-handed Z-DNA.We appreciate that detailed crystallographic studies are re-quired to fully characterise these structural changes, nev-ertheless, our available data clearly demonstrated that m5Cmodification produces distinctly different ‘structural signa-tures’ on RNA and DNA duplexes, even for duplexes withidentical CpG sequence. The significance of this finding isunclear at present, however it might infer a possible involve-ment of m5C-induced conformational change in biologicalregulatory processes.

Design principle of m5C-switchable probes

Inspired by these interesting findings, we envisaged thatthe different structure-remodelling effects of m5C on RNAand DNA duplexes could be exploited for the design of‘methylation-switchable probe’ useful for studying m5Cmethylation in DNA and RNA. Notably, at present, nomethods exist which permit the direct analysis of m5CMTase activity in living cells. There are also no reports ofassay methods that are able to distinguish between the ac-tivities of DNA m5C MTases and RNA m5C MTases.

As a proof-of-principle study, we developed a novel m5C-probe which is able to fluoresce spontaneously in responseto m5C-induced terminal sugar pucker switch, hence use-ful for sensing RNA:m5C MTase activity. The probe (8a;Figure 1) is modelled after the CpG RNA duplex 1a butcontains two important modifications. First, in order to vi-sualise m5C-induced sugar pucker switch, the 5′-cytosineresidues on the forward strand was replaced with the flu-orescent cytosine analogue 6-phenylpyrrolocytosine (pC)(47,48). pC is well-suited as reporter for our probe be-cause it is highly fluorescence (quantum yield (ΦF) 0.31)and its emission is extremely sensitive to microenviron-ment changes (65–67). In particular, previous photophys-ical studies showed that pC emits brightly when in a single-stranded environment, but becomes significantly quenchedwhen hybridised with its complementary strand (65–67).Second, to render the m5C-probe sufficiently stable for live-cell applications, the RNA backbone was replaced with a2′-O-methyl backbone, which is reasonably resistant to cel-lular nuclease degradation.

The analytical principle of the m5C-probe is illustrated inFigures 1A and B. By design, when the probe is unmethy-lated, the pC fluorophore is able to base-pair with guanineand stack strongly with its adjacent base. This results in ef-ficient quenching of pC fluorescence through photoinducedelectron transfer (PET). m5C methylation of the probe byRNA:m5C MTases, however, is expected to trigger a spon-taneous C3′-endo to C2′-endo sugar pucker switch in pC.Since the sugar ring pucker defines the glycosidic bond an-gle, a change in sugar puckering mode will also convertthe orientation of PC base from axial to equatorial, therebydisrupting its base-pairing and base-stacking interactions,leading to fluorescence activation.

The m5C-probe has the sensitivity to detect a single m5Cmark change

To verify our probe design, we first compared the fluo-rescence emission of the probes at different m5C methyla-tion levels (�ex 360 nm; �em 465 nm; Supplementary Fig-ure S7). Consistent with our detection strategy, the start-ing unmethylated probe 8a displayed weak fluorescence,with a relatively low fluorescence quantum yield (ΦF) of∼0.03, whilst its methylated counterpart 8b yielded a 2-fold increase in fluorescence intensity (ΦF ∼0.06; Supple-mental Figure S7). Similarly the unmethylated probe 8c,which was designed to simultaneously exploit sugar puckerswitch at both 5′-ends, also gave negligible fluorescence re-sponse (ΦF ∼0.04; Supplementary Figure S7). This resultis notable because most DNA/RNA duplexes are known

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Table 3. 1H NMR chemical shifts �H (ppm) for CpG DNA duplex 7a (top) and its m5C-methylated counterpart 7b (bottom) observed at 25◦C in D2Owith WATERGATE suppression. The representative 2D NOESY spectrum is given in Supplementary Figure S3. For calculation of sugar-base protondistances, see Supplementary Data

aObserved at 10◦C in 9:1 buffer/D2O solvent mix. The buffer used was 10 mM sodium phosphate buffer (pH 7.4) containing 150 mM NaCl and 20 mMMgCl2.bChemical shift for 5-methyl group on m5C.cNot observed even at 4◦C.

to undergo transient opening of the terminal base-pairs insolution. Conceivable, this might cause auto-activation ofpC fluorescence, which will severely limit the applicationsof these probes. Nevertheless, our fluorescence data fromstarting probes 8a and 8c suggests that emission arisingfrom such transient end-fraying event is relatively insignif-icant. Indeed, with the improved probe 8c, we are able toclearly detect a 2.8-fold increase in fluorescence intensityupon m5C methylation (8d; ΦF ∼ 0.11; Supplemental Fig-ure S7), which is remarkable in light of only a single m5Cmark change. Thus our m5C-probe has the sensitivity to de-tect subtle methylation changes. Moreover, the introductionof a second m5C modification, i.e. 8e led to approximately adoubling of fluorescence emission (5.3-fold), hence the flu-orescence intensity of the probe increases proportionatelywith m5C methylation level.

Interestingly, the fluorescence output of the probe couldbe further increased by incorporating additional pC fluo-rophores, as demonstrated by probe 9a (Figure 4A) wherethe concurrent replacement of cytosine 1 and 5 in bothstrands with pC led to significant improvements in fluores-cence light-up response (�ΦF (9b) 3.6-fold; (9c) 7.2-fold).Despite the introduction of 2′-O-Me backbone and four rel-atively bulky pC residues, NMR data indicate that both the5′- and 3′-terminal residues of probe 9a continue to adopta C3′-endo pucker conformation (3JH1′−H2′ ∼1–2 Hz; Sup-plementary Table S4), and they spontaneously assume aC2′-endo pucker orientation (3JH1′−H2′ ∼8 Hz) upon m5Cmethylation (9c), thus these modifications do not interferewith fluorescence activation of the probe. There is also min-imal disturbance to the overall duplex structure of probe9a relative to parent probes 8a and 1a, as verified by CDanalysis (Supplementary Figure S8). Amongst the probesinvestigated, 9a gives the greatest fluorescence response, itwas therefore selected as representative m5C-probe for sub-sequent evaluations.

The m5C-probe is highly-selective for NSUN2 over otherRNA/DNA:m5C MTases

We next examined whether m5C-probe 9a could functionas fluorogenic substrate for RNA:m5C MTase by testing itagainst NSUN2, which is of particular interest in light ofemerging evidence linking NSUN2 with m5C methylationof human mRNA (14) and its association with human can-cers (25–29). In a typical assay, 5 �M of 9a was incubatedwith NSUN2 (0.5 �M) and methyl donor S-adenosyl-L-methionine (SAM; 200 �M) under physiologically-relevantconditions (at 37◦C in 50 mM HEPES buffer containing 150mM NaCl and 20 mM MgCl2, pH 7.4). The formation ofmethylated probe was measured by an increase in fluores-cence signal at 465 nm (�ex 360 nm). Our result showed thatthe probe could be readily methylated by NSUN2, yielding∼8-fold increase in fluorescence signal after 3 h (Figure 4B).

To rule out fluorescence activation due to other non-specific mechanisms, we repeated the assay in the absence ofNSUN2 or methyl donor SAM; in both cases, there was nodetectable fluorescence light-up response (SupplementaryFigure S9). Furthermore, pre-incubation of NSUN2 with10 �M of sinefungin (a non-specific DNA/RNA MTase in-hibitor) (25,68,69) resulted in a significant reduction in flu-orescence intensity, confirming that fluorescence activationwas specifically mediated by NSUN2 MTase activity (Fig-ure 4B). Notably, the fluorescence intensity increases lin-early with the concentration of methylated probe, thus pro-viding a direct read-out of NSUN2 MTase activity (Figure4C).

We next verified whether m5C-induced sugar puckerswitch in PC was indeed responsible for fluorescence turn-on. For this purpose, we synthesised ‘locked-probe’ 10a(Figure 1C) wherein the pC fluorophore was conformation-ally restricted in a C3′-endo sugar pucker orientation viaa methylene bridge connecting O2′ with C4′ (LC; Figure1C). As a result of this modification, the locked-probe will

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Figure 4. Verification of the m5C-probe design. (A) The fluorescence emission spectra of probe 9a (�ex 360 nm; �em 465 nm) were recorded at 5 �M strandconcentration in 10 mM sodium phosphate buffer (pH 7.4) containing 150 mM NaCl and 20 mM MgCl2, 37◦C. The probe is highly responsive to its m5Cmethylation level; introduction of one and two m5C modifications led to a considerable 3.6-fold (9b) and 7.2-fold (9c) increase in fluorescence intensity,respectively. (B) Time-course fluorescence analysis of 9a (5 �M). Significant fluorescence could be detected after a short 30-minute incubation with NSUN2(0.5 �M; blue line); maximum emission was reached after ∼3 h, giving ∼8 fold increase in fluorescence intensity. Probe fluorescence reduces significantlyin the presence of generic MTase inhibitor sinefungin (10 �M; grey line), thus the probe is specifically activated by NSUN2 MTase activity. Incubationof NSUN2 with locked-probe 10a (5 �M; black line) gave negligible fluorescence response, confirming that probe activation is dependent on a changein sugar puckering of PC. (C) The fluorescence intensity increases linearly with the concentration of methylated probe, this enables a direct read-out ofNSUN2 MTase activity. Data are expressed as mean ± SD of three replicates. (D, E) Steady-state kinetics analyses of the methylation of m5C-probe, humantRNALeu(CAA) (5′-CCAGACUCAAGUUCUGG-3′) and CpG-rich ssRNA (5′-CGCGCGCGCGCG-3′) by NSUN2. Data are expressed as mean ± SDof three replicates.

no longer be able to undergo sugar pucker switch uponm5C methylation. Consistent with our hypothesis, 10a gavenegligible fluorescence light-up response when exposed toNSUN2 (Figure 4B), even though MALDI-TOF MS assayclearly showed that this probe can be methylated by NSUN2(Supplementary Figure S10). Thus probe activation mecha-nism is strictly dependent on a change in sugar pucker modeof the 5′-pC residues.

We then evaluated the specificity of probe 9a by testingit against a panel of human RNA:m5C MTases, includingthree structurally-related homologues NSUN3, NSUN5A,and NSUN6, as well as DNMT homologue TRDMT1 (for-merly DNMT2) (6,22). In all cases, there was no significantfluorescence activation (Supplementary Figure S9). Fur-thermore, MALDI-TOF MS assay showed no methylatedproduct formation even after a prolonged 8 h-incubationwith these enzymes, suggesting that 9a is highly selectivefor NSUN2 over other RNA:m5C MTases (SupplementaryFigure S10). Remarkably, 9a also gave no appreciable flu-orescence response and methylated product when exposedto key human DNA:m5C MTases, namely DNMT1 andDNMT3A (21,22), hence the probe is also able to discrim-inate against DNA:m5C MTases (Supplementary FiguresS9 and S10). Steady-state kinetic analysis using MALDI-TOF MS assay indicated that the m5C-probe is a relativelygood substrate for NSUN2 (kcat/Km = 0.3 min-1�M-1), andis only ∼2-fold less efficiently methylated compared with itsnative tRNA substrate, tRNALeu(CAA), and CpG-rich ss-RNA (Figures 4D and E).

The m5C-probe is applicable for live-cell imaging, single-cellflow cytometry analysis and cell-based inhibitor screening

On the basis of these promising data, we proceeded to in-vestigate whether the m5C-probe strategy could provide di-rect visualisation of NSUN2 activity in living cells. Ac-cordingly, HeLa cells, which constitutively express NSUN2,were transfected with either 9a (10 �M) or the locked-probe 10a (control) via Lipofectamine 2000, and then im-aged using fluorescence microscopy (�ex 340–380 nm, �em435–485 nm). Cells treated with probe 9a gave off brightblue fluorescence after 1 h; this is ∼7-fold higher intensitycompared with control (Figures 5A–C).

Further characterisation of the probe by time course flu-orescence analysis showed that the probe remained stronglyemissive in HeLa cell lysate for at least 3 h, suggesting goodphotostability (Supplementary Figure S11A). Moreover,there was no obvious increase in probe fluorescence in celllysate which had been spiked with 1 mM sinefungin, sug-gesting no significant fluorescence arising from NSUN2-independent mechanisms (Supplementary Figure S11A).The lack of fluorescence increase further suggests that theprobe is reasonably resistant to cellular degradation, whichwould otherwise release unstacked fluorophore. Consistentwith this result, a repeat of the cell lysate experiment us-ing locked-probe 10a gave very little or no increase in flu-orescence in HeLa cell lysate, both in the presence and ab-sence of 1 mM sinefungin (Supplementary Figure S11B).These data suggest that probe degradation by cellular nu-

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Figure 5. The m5C-probe provides real-time visualisation of cellular NSUN2 MTase activity. Live cell imaging of HeLa cells following treatment withm5C-probe 9a (10 �M) using (A) bright field and (B) fluorescence microscopy. (C) Fluorescence image of HeLa cells treated with locked-probe 10a (10�M) obtained from a separate experiment than the correlated images in panels A and B. It is displayed using the same brightness and contrast settingsas that in panel B. Scale bar, 20 �m. (D) Violin plots showing the flow cytometry analysis of HeLa cells transfected with probe 9a (green), and withNSUN2 knockdown (orange) or NSUN2 overexpression (red) (�ex 355 nm; �em 425–475 nm). The boxes in violin plots show the median (white dot),25–75 percentile (black box) and 5–95 percentile values (black bar). (E) The fluorescence intensity of m5C-probe changes with the expression levels ofNSUN2 in HeLa cells, but is unresponsive towards NSUN6, TRDMT1 and DNMT3A. A minimum of 20 000 live cells were analysed for determinationof mean fluorescence intensity (MFI). Data are expressed as mean ± SD of three biological replicates. **P < 0.01; n.s. = not significant.

cleases and other nucleic acid modifying enzymes do notinterfere with the performance of the m5C-probe 9a sig-nificantly, at least not within the time frame of our cell-based assay (typically 1 h). Furthermore, the probe is rel-atively non-cytotoxic, as evidenced by MTT toxicity assayand cell morphological assessment, where >80% of the cellsremained viable after being exposed to 40 �M probe for 24h (Supplementary Figure S11C).

Besides live-cell imaging, the m5C-probe is also suitablefor use in single-cell flow cytometry analysis (�ex 355 nm;�em 425–475 nm). In particular, NSUN2 knockdown inHeLa cells caused a marked reduction in probe fluores-cence, whereas overexpression of NSUN2 led to signifi-cant increase in fluorescence signal (Figures 5D and E). Re-markably, the fluorescence intensity was unchanged withNSUN6, TRDMT1, and DNMT3A knockdown and over-expression, thus, consistent with our in vitro data, the probeis highly selective for NSUN2 over other m5C MTases. Priorto this work, we are not aware of any assay method that is

able to discriminate between DNA and RNA m5C MTaseactivities in cells.

Finally, the developed m5C-probe could also be usedto directly measure NSUN2 inhibition in cells. Treat-ment of HeLa cells with increasing concentrations ofknown MTase inhibitors, sinefungin or S-adenosyl-L-homocysteine (SAH), for 30 min prior to the addition ofprobe 9a (10 �M) led to a concentration-dependent de-crease in fluorescence signal (Figures 6A–C). The deter-mined IC50 values of sinefungin (8.9 �M) and SAH (3.1�M) against NSUN2 are comparable with their inhibitoryactivities against other RNA MTases (IC50 values typicallyrange between 0.1 �M to 20 �M (25,68); Figure 6D). Im-portantly, the negative controls, adenosine and homocys-teine, showed poor inhibition towards NSUN2, as antici-pated (IC50s >200 �M and >400 �M, respectively). Thusour assay is able to identify known NSUN2 inhibitorsfrom amongst structurally-related analogues. We furtherperformed Z’ factor analysis (55) to evaluate the reliabil-

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Figure 6. Application of the m5C-probe assay in cell-based screening of NSUN2 inhibitors. (A) IC50 values of non-specific MTase inhibitors againstNSUN2. Results are from m5C-probe assay. (B) Analysis of NSUN2 inhibition in live HeLa cells by real-time flow cytometry (�ex 355 nm; �em 425–475nm). Cells were pre-incubated with sinefungin (0, 10 and 1000 �M; colour coded) for 30 min prior to treatment with m5C-probe 9a (10 �M) or locked-probe 10a (10 �M; control). (C) The flow cytometry profile at 1 h showed a concentration-dependent decrease in mean fluorescence intensity (MFI) in cellstreated with 9a. (D) Inhibition of NSUN2 by selected MTase inhibitors. The m5C-probe assay could distinguish known inhibitors from structurally-relatednegative controls. Data are expressed as mean ± SD of three replicates. (E) Screening validation of the m5C-probe assay in a 96-well format. The assay hasa Z’ factor of 0.73, which demonstrates excellent reproducibility.

ity of our assay for high-throughput screening of inhibitor(for experimental details, see Supplementary Data). Ourassay produces an average Z’ factor of 0.73 in a 96-wellplate format, indicating that it has the required statistical re-producibility for cell-based screening of NSUN2 inhibitors(Figure 6E).

The m5C-switchable probe strategy could be adapted for thestudy of DNA:m5C MTase activity

In light of present finding that m5C methylation can inducea spontaneous B–Z conformational change in CpG DNAduplex 7a (Figure 3B and Supplementary Figure S6), ourm5C-probe strategy may, in principle, also be adapted forthe study of DNA:m5C MTase activity. To examine this pos-sibility, we prepared DNA probe 11a, which is an analogueof 7a containing a 5′-overhang pyrene deoxyriboside (Py)in the forward strand (Figure 7). The design of this probeexploits differences in the ability of Py to participate in end-stacking interactions in the B-DNA and Z-DNA forms. Itis clear, from a number of studies, that whereas the B-DNA

(diameter of helix ∼20 A) is able to adopt a continuousbase-stacking arrangement, the more compact Z-DNA (di-ameter of helix ∼18 A) only permits discontinuous series offour-base stack (70). Because of these inherent differencesin base-stacking pattern, the unpaired Py will only be ableto end-stack on adjacent G:C base pair when it is in a B-DNA environment, and not in the Z-DNA environment.Accordingly, conversion of probe 11a from B-DNA to Z-DNA form upon m5C methylation is expected to cause adestacking of Py fluorophores and, consequently, fluores-cence activation.

Consistent with our probe detection strategy, m5C methy-lation readily triggered a B-Z transformation in probe 11a,with concomitant increase in fluorescence emission (�ΦF∼5.5-fold; �ex 340 nm; �em 380 nm and 400 nm; Figures8A and B). Notably, there was no fluorescence activationin the absence of MgCl2 where 11a* did not undergo B-to-Z conversion, thus the observed fluorescence response wasprimarily mediated by a B–Z conversion of the probe (Fig-ure 8B). Moreover, our thermodynamic analysis supportsthe notion that B-Z transition triggers a destacking of the

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Figure 7. The m5C-probe approach could be adapted for the detection of DNA:m5C MTases activity. (A) The DNA m5C-probe 11a is conformationally-responsive to m5C methylation. It exists as a right handed B-DNA structure when unmethylated, but undergoes spontaneous and rapid transformation tothe more compact, left-handed Z-DNA structure upon m5C methylation by DNA:m5C MTases (e.g. DNMT3A). Such a major B-Z conversion severelydisrupts end-stacking interaction of the fluorophore, pyrene deoxyriboside (Py), leading to fluorescence activation. (B) Schematic representation of DNAm5C-probe 11a and its methylated counterpart 11b. The structure of Py monomer is shown.

dangling Py (Supplementary Table S3). In particular, in theabsence of m5C methylation, Py is able to stack readily atthe 5′-end of probe 11a, giving an end-stacking stabilisationof ∼2.8 kcal/mol (compare the difference in duplex stabil-ity between 11a and 7a; Supplementary Table S3 and Sup-plementary Figures S12 and S13). Upon m5C methylation,however, end-stacking interaction is completely abolished(negligible difference in stability between 11b and 7b). Thissuggests that the end-stacking property of Py and, hence, itsfluorescence output is sensitive to m5C methylation status.

Preliminary biochemical analysis further demonstratedthat probe fluorescence is only activated by DNA:m5CMTases DNMT3A, and not by RNA:m5C MTases NSUN2(Figure 8C). Importantly, m5C-probe 11a may also be ap-plied to the inhibition study of DNMT3A. As shown in Fig-ure 8D, incubation of probe 11a (5 �M) and DNMT3A (0.5�M) with increasing concentrations of sinefungin (knownDNMT3A inhibitor) or adenosine (negative control) for 30min led to clear concentration-dependent decrease in fluo-rescence intensity. The determined IC50 values of sinefun-gin (0.7 �M) and adenosine (106.3 �M) are comparablewith previously reported values (71), thus the developedm5C-probe strategy could be used for in vitro screening ofDNMT3A inhibitors. Work is currently underway to ex-plore the possibility of employing both probes concurrentlyfor the simultaneous detection of RNA and DNA m5CMTase activity.

CONCLUSION

Overall, we use a combination of NMR, CD and ther-modynamic analyses to investigate the structural impactof m5C methylation on RNA and DNA duplexes. Our re-sults revealed that, although m5C does not hinder canoni-cal Watson–Crick base-pairing interactions, this modifica-tion can, in fact, elicits significant conformational changein certain RNA and DNA sequence contexts. We furtherdemonstrated that m5C produces distinctly different ‘struc-tural signatures’ on RNA and DNA duplexes, including du-plexes with identical CpG sequences. In the case of CpGRNA duplex (as exemplified by 1a), m5C methylation in-duces a local distortion of the phosphate backbone and aC3′-endo to C2′-endo sugar pucker switch in the terminalresidues. However, in the case of CpG DNA duplex (as ex-emplified by 7a), m5C triggers a remarkable B-Z structuraltransformation. The significance of this result is unclear atpresent nevertheless, given that majority of the m5C mod-ifications in DNA and mRNA are distributed within theCpG consensus motif, sequence information alone is in-sufficient to discriminate m5C marks on RNA and DNACpG sites. Therefore, one question which arises from thisfinding is whether the duplex-remodelling property of m5Cmight provide a mechanism for its specific recognition byRNA/DNA:m5C binding proteins.

On the basis of this interesting finding, we further pro-vided proof-of-principle that m5C-induced conformational

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Figure 8. DNA probe 11a provides highly sensitive detection of DNA:m5C methylation. (A) Fluorescence measurement of probe 11a (5 �M) showedtwo distinct pyrene emission peaks at 380 nm and 400 nm (�ex 340 nm). The spectra were recorded at 37◦C under physiologically-relevant conditions (10mM sodium phosphate buffer (pH 7.4) containing 150 mM NaCl and 20 mM MgCl2). The probe exhibited a significant 5.5-fold increase in fluorescenceintensity upon m5C methylation (11b), but gave no appreciable light-up response in the absence of MgCl2 (11b*), implying that fluorescence switch-on wasprimarily mediated by B–Z structural conversion of the probe. (B) m5C methylation triggered a major inversion of the CD spectrum of probe 11a underphysiologically-relevant conditions. There was little or no conformational change in the absence of MgCl2 (11b*). (C) Time-course fluorescence analysis(�ex 340 nm; �em 400 nm) of probe 11a (5 �M) in the presence of DNMT3A (0.5 �M; red line) or NSUN2 (0.5 �M; black line). The results demonstrate thatprobe 11a is selectively activated by DNMT3A, and not by NSUN2. (D) Probe 11a could be used for in vitro inhibition assay of DNMT3A. Both sinefungin(known DNMT3A inhibitor) and adenosine (negative control) displayed clear concentration-dependent decrease in fluorescence, with determined IC50values of 0.7 �M and 106.3 �M, respectively. Data are expressed as mean ± SD of three replicates.

change can be exploited for direct detection of m5C MTaseactivity in cells. This was demonstrated by the develop-ment of the first m5C-responsive probe 9a, which switchessugar pucker conformation spontaneously according to itsm5C methylation status, hence useful for sensing RNA:m5CMTase activity.

The m5C-probe is simple, inexpensive and highly se-lective for NSUN2 over other RNA:m5C MTases (in-cluding NSUN3, NSUN5A, NSUN6 and TRDMT1) andDNA:m5C MTases (including DNMT1 and DNMT3A).Through the use of this probe, we achieved fluorescenceimaging and real-time flow cytometry analysis of NSUN2activity in live HeLa cells. We further demonstrated theutility of the probe in cell-based screening of NSUN2 in-hibitors. Prior to this study, we are not aware of any meth-ods that permit direct sensing of RNA:m5C MTase activityin cells. There are also no reports of assays that selectivelytarget NSUN2 over other RNA/DNA:m5C MTases. Thediscovery of such highly selective probes is rarely achievedand may prove valuable in advancing our knowledge ofNSUN2 in m5C-regulated processes. Importantly, our m5C-probe approach could also be adapted for the analysis ofDNA:m5C MTase activity. This was demonstrated by thedevelopment of DNA m5C-probe 11a, which is useful for invitro screening of DNMT3A inhibitors.

We appreciate that m5C-induced terminal sugar puckerswitch is likely an interesting anomaly that is unique to alter-nating CpG RNA duplexes, since this phenomenon was notobserved in majority of other sequences investigated in this

study. Therefore, one limitation of the proposed approachis that it is not applicable to RNA:m5C methyltransferasesthat do not methylate CpG substrates. Nevertheless, giventhe importance of NSUN2, both as a key regulator of m5Cmarks in mRNA and a potential therapeutic target, we en-visaged that the proposed NSUN2 assay strategy would beof broad scientific interest.

SUPPLEMENTARY DATA

Supplementary Data are available at NAR Online.

ACKNOWLEDGEMENTS

We would like to thank the Chemical, Molecular and Ma-terials Analysis Centre at the National University of Singa-pore for their support in NMR measurements and helpfuldiscussions.

FUNDING

Singapore Ministry of Health’s National Medical ResearchCouncil [NMRC/BNIG/2008/2013]; Singapore Ministryof Education [AcRF Tier 1 Grant R148-000-231-114 andR148-000-238-114]. Funding for open access charge: Sin-gapore Ministry of Education [AcRF Tier 1 Grant R148-000-238-114].Conflict of interest statement. None declared.

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https://academic.oup.com/nar/article-lookup/doi/10.1093/nar/gkz1047#supplementary-data


REFERENCES1. Boccaletto,P., Machnicka,M.A., Purta,E., Piatkowski,P., Baginski,B.,

Wirecki,T.K., de Crecy-Lagard,V., Ross,R., Limbach,P.A., Kotter,A.et al. (2018) MODOMICS: a database of RNA modificationpathways. 2017 update. Nucleic. Acids. Res., 46, D303–D307.

2. Sood,A.J., Viner,C. and Hoffman,M.M. (2019) DNAmod: the DNAmodification database. J. Cheminform., 11, 30.

3. Grosjean,H. (2005) Fine-Tuning of RNA Functions by Modificationand Editing. Springer, Berlin, Heidelberg.

4. Motorin,Y. and Helm,M. (2011). RNA nucleotide methylation. WileyInterdiscip. Rev. RNA, 2, 611–631.

5. Breiling,A. and Lyko,F. (2015) Epigenetic regulatory functions ofDNA modifications: 5-methylcytosine and beyond. Epigenet.Chromatin, 8, 24.

6. Motorin,Y., Lyko,F. and Helm,M. (2010) 5-methylcytosine in RNA:detection, enzymatic formation and biological functions. NucleicAcids Res., 38, 1415–1430.

7. Smith,Z.D. and Meissner,A. (2013) DNA methylation: roles inmammalian development. Nat. Rev. Genet., 14, 204–220.

8. Jones,P.A. (2012) Functions of DNA methylation: islands, start sites,gene bodies and beyond. Nat. Rev. Genet. 13, 484–492.

9. Squires,J.E., Patel,H.R., Nousch,M., Sibbritt,T., Humphreys,D.T.,Parker,B.J., Suter,C.M. and Preiss,T. (2012) Widespread occurrenceof 5-methylcytosine in human coding and non-coding RNA. NucleicAcids Res., 40, 5023–5033.

10. Edelheit,S., Schwartz,S., Mumbach,M.R., Wurtzel,O. and Sorek,R.(2013) Transcriptome-wide mapping of 5-methylcytidine RNAmodifications in bacteria, archaea, and yeast reveals m5C withinarchaeal mRNAs. PLoS Genet., 9:e1003602.

11. Khoddami,V. and Cairns,B.R. (2013) Identification of direct targetsand modified bases of RNA cytosine methyltransferases. Nat.Biotechnol., 31, 458–464.

12. Legrand,C., Tuorto,F., Hartmann,M., Liebers,R., Jacob,D.,Helm,M. and Lyko,F. (2017). Statistically robust methylation callingfor whole-transcriptome bisulfite sequencing reveals distinctmethylation patterns for mouse RNAs. Genome Res., 27, 1589–1596.

13. Amort,T., Rieder,D., Wille,A., Khokhlova-Cubberley,D., Riml,C.,Trixl,L., Jia,X.-Y., Micura,R. and Lusser,A. (2017) Distinct5-methylcytosine profiles in poly(A) RNA from mouse embryonicstem cells and brain. Genome Biol., 18, 1.

14. Yang,X., Yang,Y., Sun,B.-F., Chen,Y.-S., Xu,J.-W., Lai,W.-Y., Li,A.,Wang,X., Bhattarai,D.P., Xiao,W. et al. (2017). 5-methylcytosinepromotes mRNA export––NSUN2 as the methyltransferase andALYREF as an m5C reader. Cell Res., 27, 606–625.

15. David,R., Burgess,A., Parker,B., Li,J., Pulsford,K., Sibbritt,T.,Preiss,T. and Searle,I.R. (2017). Transcriptome-wide mapping ofRNA 5-Methylcytosine in Arabidopsis mRNAs and non-codingRNAs. Plant Cell, 29, 445–460.

16. Cui,X., Liang,Z., Shen,L., Zhang,Q., Bao,S., Geng,Y., Zhang,B.,Leo,V., Vardy,L.A., Lu,T. et al. (2017). 5-Methylcytosine RNAmethylation in Arabidopsis thaliana. Mol. Plant, 10, 1387–1399.

17. Roundtree,I.A., Evans,M.E., Pan,T. and He,C. (2017) Dynamic RNAmodifications in gene expression regulation. Cell, 169, 1187–1200.

18. Tang,H., Fan,X., Xing,J., Liu,Z., Jiang,B., Dou,Y., Gorospe,M. andWang,W. (2015). NSun2 delays replicative senescence by repressingp27 (KIP1) translation and elevating CDK1 translation. Aging, 7,1143–1158.

19. Xing,J., Yi,J., Cai,X., Tang,H., Liu,Z., Zhang,X., Martindale,J.L.,Yang,X., Jiang,B., Gorospe,M. et al. (2015). NSun2 promotes cellgrowth via elevating cyclin-dependent kinase 1 translation. Mol. Cell.Biol., 35, 4043–4052.

20. Aguilo,F., Li,S., Balasubramaniyan,N., Sancho,A., Benko,S.,Zhang,F., Vashisht,A., Rengasamy,M., Andino,B., Chen,C.H. et al.(2016). Deposition of 5-Methylcytosine on enhancer RNAs enablesthe coactivator function of PGC-1�. Cell Rep., 14, 479–492.

21. Lyko,F. (2018) The DNA methyltransferase family: a versatile toolkitfor epigenetic regulation. Nat. Rev. Genet., 19, 81–92.

22. Grosjean,H. (2009) DNA and RNA Modification Enzymes: Structure,Mechanism, Function and Evolution. Landes Bioscience, Austin.

23. Kohli,R.M. and Zhang,Y. (2013) TET enzymes, TDG and thedynamics of DNA demethylation. Nature, 502, 472–479.

24. Liu,Y. and Santi,D.V. (2000) m5C RNA and m5C DNA methyltransferases use different cysteine residues as catalysts. Proc. Natl.Acad. Sci. U.S.A., 97, 8263–8265.

25. Boriack-Sjodin,P.A., Ribich,S. and Copeland,R.A. (2018)RNA-modifying proteins as anticancer drug targets. Nat. Rev. DrugDiscov., 17, 435–453.

26. Freeman,J.W., Hazlewood,J.E., Auerbach,P. and Busch,H. (1988)Optimal loading of scraped HeLa cells with monoclonal antibodiesto the proliferation associated Mr 120, 000 Nucleolar Antigen.Cancer Res., 48, 5246–5250.

27. Frye,M. and Watt,F.M. (2006) The RNA methyltransferase Misu(NSun2) mediates Myc-induced proliferation and is upregulated intumors. Curr Biol., 16, 971–981.

28. Hussain,S., Benavente,S.B., Nascimento,E., Dragoni,I., Kurowski,A.,Gillich,A., Humphreys,P. and Frye,M. (2009) The nucleolar RNAmethyltransferase Misu (NSun2) is required for mitotic spindlestability. J. Cell Biol., 186, 27–40.

29. Kar,S.P., Beesley,J., Amin,A., Olama,A., Michailidou,K., Tyrer,J.,Kote-Jarai,Z., Lawrenson,K., Lindstrom,S., Ramus,S.J. et al. (2016)Genome-wide meta-analyses of breast, ovarian, and prostate cancerassociation studies identify multiple new susceptibility loci shared byat least two cancer types. Cancer Discov., 6, 1052–1067.

30. Van Haute,L., Dietmann,S., Kremer,L., Hussain,S., Pearce,S.F.,Powell,C.A., Rorbach,J., Lantaff,R., Blanco,S., Sauer,S. et al. (2016)Deficient methylation and formylation of mt tRNA Met wobblecytosine in a patient carrying mutations in NSUN3. Nat. Commun.,7, 2039.

31. Haag,S., Sloan,K.E., Ranjan,N., Warda,A.S., Kretschmer,J.,Blessing,C., Hubner,B., Seikowski,J., Dennerlein,S., Rehling,P. et al.(2016) NSUN3 and ABH1 modify the wobble position ofmt-tRNAMet to expand codon recognition in mitochondrialtranslation. EMBO J., 35, 2104–2119.

32. Aguilo,F., Li,S., Balasubramaniyan,N., Sancho,A., Benko,S.,Zhang,F., Vashisht,A., Rengasamy,M., Andino,B., Chen,C.H. et al.(2016) Deposition of 5-methylcytosine on enhancer RNAs enablesthe coactivator function of PGC-1�. Cell Rep., 14, 479–492.

33. Harris,T., Marquez,B., Suarez,S. and Schimenti,J. (2007) Spermmotility defects and infertility in male mice with a mutation in Nsun7,a member of the Sun domain-containing family of putative RNAmethyltransferases. Biol. Reprod., 77, 376–382.

34. Yang,T., Cheong,A., Mai,X., Zou,S. and Woon,E.C.Y. (2016) Amethylation-switchable conformational probe for the sensitive andselective detection of RNA demethylase activity. Chem. Commun., 52,6181–6184.

35. Zou,S., Toh,J.D.W., Wong,K.H.Q., Gao,Y.-G., Hong,W. andWoon,E.C.Y. (2016) N6-Methyladenosine: a conformational markerthat regulates the substrate specificity of human demethylases FTOand ALKBH5. Sci. Rep., 6, 25677.

36. Cheong,A., Low,J.J.A., Lim,A., Yen,P.M. and Woon,E.C.Y. (2018) Afluorescent methylation-switchable probe for highly sensitive analysisof FTO N6-methyladenosine demethylase activity in cells. Chem. Sci.,9, 7174–7185.

37. Engel,J.D. and von Hippel,P.H.J. (1978) Effects of methylation on thestability of nucleic acid conformations. Studies at the polymer level.Biol. Chem., 253, 927–934.

38. Micura,R., Pils,W., Hobartner,C., Grubmayr,K., Ebert,M.O. andJaun,B. (2001) Methylation of the nucleobases in RNAoligonucleotides mediates duplex–hairpin conversion. Nucleic. Acids.Res., 29, 3997–4005.

39. Kierzek,E. and Kierzek,R. (2003) The thermodynamic stability ofRNA duplexes and hairpins containing N6-alkyladenosines and2-methylthio-N6-alkyladenosines. Nucleic Acids Res., 31, 4472–4480.

40. Roost,C., Lynch,S.R., Batista,P.J., Qu,K., Chang,H.Y. and Kool,E.T.(2015) Structure and thermodynamics of N6-methyladenosine inRNA: a spring-loaded base modification. J. Am. Chem. Soc., 137,2107–2115.

41. Behe,M. and Felsenfeld,G. (1981) Effects of methylation on asynthetic polynucleotide: the B-Z transition inpoly(dG-m5dC).poly(dG-m5dC). Proc. Natl. Acad. Sci. U. S. A., 78,1619–1623.

42. Tran-Dinh,S., Taboury,J., Neumann,J.-M., Huynh-Dinh,T.,Genissel,B., Langlois d’Estaintot,B. and Igolen,J. (1984) ProtonNMR and circular dichroism studies of the B and Z conformations ofthe self-complementary deoxyhexanucleotide

Dow






ber 2019


d(m5C-G-C-G-m5C-G): mechanism of the Z-B-coil transitions.Biochemistry, 23, 1362–1371.

43. Butkus,V., Klirnaauskas,S., Petrauskiene,L., Maneliene,Z.,Janulaitis,A., Minchenkoval,L.E. and Schyolkinal,A.K. (1987)Synthesis and physical characterization of DNA fragmentscontaining N4-methylcytosine and 5-methycytosine. Nucleic AcidsRes., 15, 8467–8478.

44. Orbons,L.P. and Altona,C. (1986) Conformational analysis of the Band Z forms of the d(m5C-G)3 and d(br5C-G)3 hexamers in solution.A 300-MHz and 500-MHz NMR study. Eur. J. Biochem., 160,141–148.

45. Ceolin,F., Babin,F., Huynh-Dinh,T. and Igolen,J. (1987) RNAfragment r(CGm5CGCG) that exhibits two conformations in slowexchange on the NMR time scale in low salt solution. J. Am. Chem.Soc., 109, 2539–2541.

46. Bloch,G., Neumann,J.M., Babin,F. and Huynh-Dinh,T. (1987)Sequence-dependence of the conformational changes induced by the5-methyl cytosine in synthetic RNA oligomers. FEBS Lett., 219,464–468.

47. Hudson,R.H.E. and Ghorbani-Choghamarani,A. (2007) Selectivefluorometric detection of guanosine-containing sequences by6-phenylpyrrolocytidine in DNA. Synlett., 6, 870–873.

48. Cho,S.J., Ghorbani-Choghamarani,A., Saito,Y. and Hudson,R.H.E.(2019) 6-Phenylpyrrolocytidine: an intrinsically fluorescent,environmentally responsive nucleoside analogue. Curr. Protoc.Nucleic Acid Chem., 76, e75.

49. Ren,R.X.-F., Chaudhuri,N.C., Paris,P.L., Rumney,S. andKool,E.T.(1996) Naphthalene, phenanthrene, and pyrene as DNAbase analogues: synthesis, structure, and fluorescence in DNA. J. Am.Chem. Soc., 118, 7671–7678.

50. Sklenar,V., Piotto,M., Leppik,R. and Saudek,V. (1993)Gradient-tailored water suppression for 1H-15N HSQC experimentsoptimized to retain full sensitivity. J. Magn. Reson. A, 102, 241–250.

51. States,D.J., Haberkorn,R.A. and Ruben,D.J. (1982) Atwo-dimensional nuclear overhauser experiment with pure absorptionphase in four quadrants. J. Magn. Reson., 48, 286–292.

52. Jeener,J., Meier,B.H., Bachmann,P. and Ernst,R.R. (1979)Investigation of exchange processes by two-dimensional NMRspectroscopy. J. Chem. Phys., 71, 4546–4593.

53. Barkhuisen,H., de Beer,R., Bovee,W.M.M.J. and van Ormondt,K.(1985) Retrieval of frequencies, amplitudes, damping factors, andphases from time-domain signals using a linear least-squaresprocedure. J. Magn. Reson., 61, 465–481.

54. Otting,G., Widmer,H., Wagner,G. and Wuthrich,K. (1986) Origin of�2 and �2 ridges in 2D NMR spectra and procedures for suppression.J. Magn. Reson., 66, 187–193.

55. Zhang,J.H., Chung,T.D.Y. and Oldenburg,K.R. (1999) A simplestatistical parameter for use in evaluation and validation of highthroughput screening assays. J. Biomol. Screen, 4, 67–73.

56. Zhang,X., Liu,Z., Yi,J., Tang,H., Xing,J., Yu,M., Tong,T., Shang,Y.,Gorospe,M. and Wang,W. (2012) NSun2 stabilizes p16INK4 mRNAby methylating the 3′-untranslated region of p16. Nat. Commun., 3,712.

57. Dev,R.R., Ganji,R., Singh,S.P., Mahalingam,S., Banerjee,S. andKhosla,S. (2017). Cytosine methylation by DNMT2 facilitates

stability and survival of HIV-1 RNA in the host cell during infection.Biochem. J., 474, 2009–2026.

58. Chen,Z.X., Mann,J.R., Hsieh,C.L., Riggs,A.D. and Chedin,F. (2005)Physical and functional interactions between the human DNMT3Lprotein and members of the de novo methyltransferase family. J. CellBiochem., 95, 902–917.

59. Wang,Z.-L., Li,B., Luo,Y.-X., Lin,Q., Liu,S.-R., Zhang,X.-Q.,Zhou,H., Yang,J.-H. and Qu,L.-H. (2018) Comprehensive genomiccharacterization of RNA-binding proteins across human cancers.Cell Rep., 22, 286–298.

60. Popenda,M., Biala,E., Milecki,J. and Adamiak,R.W. (1997) Solutionstructure of RNA duplexes containing alternating CG base pairs:NMR study of r(CGCGCG)2 and 2′-O-Me(CGCGCG)2 under lowsalt conditions. Nucleic. Acids. Res., 25, 4589–4598.

61. Altona,C. and Sundaralingam,M. (1973) Conformational analysis ofthe sugar ring in nucleosides and nucleotides. Improved method forthe interpretation of proton magnetic resonance coupling constants.J. Am. Chem. Soc., 95, 2333–2344.

62. Neuhaus,D. and Williamson,M.P. (2000) The Nuclear OverhauserEffect in Structural and Conformational Analysis. John Wiley & Sons,Inc., NY.

63. Butts,C.P., Jones,C.R., Towers,E.C., Flynn,J.L., Appleby,L. andBarron,N.J. (2011) Interproton distance determinations by NOE –surprising accuracy and precision in a rigid organic molecule. Org.Biomol. Chem., 9, 177–184.

64. Fujii,S., Wang,A.H.-J., van der Marel,G., van Boom,J.H. and Rich,A.(1982) Molecular structure of (m5dC-dG)3: the role of the methylgroup on 5-methyl cytosine in stabilizing Z-DNA. Nucleic Acids Res.,10, 7879–7892.

65. Wojciechowski,F. and Hudson,R.H.E. (2008) Fluorescence andhybridization properties of peptide nucleic acid containing asubstituted phenylpyrrolocytosine designed to engage guanine withan additional H-bond. J. Am. Chem. Soc., 130, 12574–12575.

66. Wahba,A.S., Esmaeili,A., Damha,M.J. and Hudson,R.H.E. (2010) Asingle-label phenylpyrrolocytidine provides a molecular beacon-likeresponse reporting HIV-1 RT RNase H activity. Nucleic Acids Res.,38, 1048–1056.

67. Wahba,A.S., Azizi,F., Deleavey,G.F., Brown,C., Robert,F.,Carrier,M., Kalota,A., Gewirtz,A.M., Pelletier,J., Hudson,R.H. et al.(2011) Phenylpyrrolocytosine as an unobtrusive base modification formonitoring activity and cellular trafficking of siRNA. ACS Chem.Biol., 6, 912–919.

68. Yebra,M.J., Sanchez,J., Martin,C.G., Hardisson,C. and Barbes,C.(1991), The effect of sinefungin and synthetic analogues on RNA andDNA methyltransferases from Streptomyces. J. Antibiot., 44,1141–1147.

69. Woon,E.C.Y., Arcieri,M., Wilderspin,A.F., Malkinson,J.P. andSearcey,M. (2007) Solid-phase synthesis of chlorofusin analogues. J.Org. Chem., 72, 5146–5151.

70. Neidle,S. (1999) Oxford Handbook of Nucleic Acid Structure. OxfordUniversity Press, NY.

71. Kilgore,J.A., Du,X., Melito,L., Wei,S., Wang,C., Chin,H.G.,Posner,B., Pradhan,S., Ready,J.M. and Williams,N.S. (2013)Identification of DNMT1 selective antagonists using a novelscintillation proximity assay. J. Biol. Chem., 288, 19673–19684.

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