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m1A and m1G disrupt A-RNA structure through the intrinsic instability of Hoogsteen base pairs

Nature Structural & Molecular Biology volume 23, pages 803–810 (2016)Cite this article

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Abstract

The B-DNA double helix can dynamically accommodate G-C and A-T base pairs in either Watson–Crick or Hoogsteen configurations. Here, we show that G-C+ (in which + indicates protonation) and A-U Hoogsteen base pairs are strongly disfavored in A-RNA. As a result,N1-methyladenosine and N1-methylguanosine, which occur in DNA as a form of alkylation damage and in RNA as post-transcriptional modifications, have dramatically different consequences. Whereas they create G-C+ and A-T Hoogsteen base pairs in duplex DNA, thereby maintaining the structural integrity of the double helix, they block base-pairing and induce local duplex melting in RNA. These observations provide a mechanism for disrupting RNA structure through post-transcriptional modifications. The different propensities to form Hoogsteen base pairs in B-DNA and A-RNA may help cells meet the opposing requirements of maintaining genome stability, on the one hand, and of dynamically modulating the structure of the epitranscriptome, on the other.

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Figure 1: Absence of detectable WC ⇌ HG exchange in A-RNA by NMR relaxation dispersion.
Figure 2: Absence of detectable WC ⇌ HG exchange across diverse sites and conditions in the E-gc RNA duplex.
Figure 3: m1A and m1G disrupt A-RNA structure and do not form HG base pairs.
Figure 4: Source of HG instability in A-RNA.
Figure 5: Different propensities for HG base-pair formation in B-DNA and A-RNA enable contrasting roles at the genome and transcriptome level.
Figure 6: Modulation of RNA structure by m1rA and m1rG, owing to the instability of HG base pairs.

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Acknowledgements

We thank M. Juen (University of Innsbruck, Austria), N. Orlovsky, Y. Xue, A. Shakya, M. Clay, A. Rangadurai, E. Szymanski, members of the Al-Hashimi laboratory, and T. Mustoe (UNC–Chapel Hill) for assistance and critical input. We acknowledge technical support and resources from the Duke Magnetic Resonance Spectroscopy Center, the Duke Compute Cluster and the Shared Materials Instrumentation Facility at Duke University. This work was supported by NIH grants (R01GM089846 to I.A. and H.M.A.; 5P50GM103297 to H.M.A.) and Austrian Science Fund (FWF) grants (P26550 and P28725 to C.K.). H.Z. acknowledges support from the China Scholarship Council.

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Author notes

  1. Bharathwaj Sathyamoorthy

    Present address: Present address: Department of Chemistry, Indian Institute of Science Education and Research Bhopal, Bhopal, India.,

Authors and Affiliations

  1. Department of Biochemistry, Duke University School of Medicine, Durham, North Carolina, USA

    Huiqing Zhou, Isaac J Kimsey, Bharathwaj Sathyamoorthy, Tianyu Bai & Hashim M Al-Hashimi

  2. Department of Molecular Biology, The Scripps Research Institute, La Jolla, California, USA

    Evgenia N Nikolova

  3. Department of Chemistry, University of California Irvine, Irvine, California, USA

    Gianmarc Grazioli, James McSally & Ioan Andricioaei

  4. Bachem Americas, Torrance, California, USA

    Christoph H Wunderlich

  5. Institute of Organic Chemistry, University of Innsbruck, Innsbruck, Austria

    Christoph Kreutz

  6. Center for Molecular Biosciences Innsbruck, University of Innsbruck, Innsbruck, Austria

    Christoph Kreutz

  7. Department of Chemistry, Duke University, Durham, North Carolina, USA

    Hashim M Al-Hashimi

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Contributions

H.Z., E.N.N., and H.M.A. conceived the project and experimental design. H.Z. prepared NMR samples, with assistance from I.J.K. and E.N.N., and performed NMR experiments and analyzed NMR data, with assistance from I.J.K. and B.S. H.Z. performed DFT calculations and modeling of steric analysis. H.Z., I.J.K., and E.N.N. performed the structure-based survey of RNA Hoogsteen base pairs. I.A., G.G., and J.M. performed and analyzed the MD simulations. C.H.W. and C.K. prepared the 13C-C8-adenosine phosphoramidite. H.Z. and T.B. carried out the UV melting experiments and performed the data analysis. H.M.A., H.Z., and I.A. wrote the manuscript with critical input from I.J.K., B.S., E.N.N., G.G., J.M., T.B., C.H.W., and C.K.

Corresponding authors

Correspondence to Ioan Andricioaei or Hashim M Al-Hashimi.

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Integrated supplementary information

Supplementary Figure 1 Resonance assignment and NMR spectra for hairpin and duplex A6 constructs.

(a) 2D HSQC spectra of hairpin construct (hp-A6-RNA) with labeled assignments. (b) 2D HSQC or SOFAST-HMQC spectra of m1A or m1G modified RNA (in violet) duplexes with labeled assignments overlaid on their unmodified counterparts (in grey). Arrows indicate significant chemical shift perturbations induced by m1A or m1G. Resonances that are exchanged broadened out of detection due to m1A or m1G are highlighted on the corresponding resonance of the unmodified duplexes with a dashed circle. In A6-RNAm1A, G13C-C1′ has a ≍6.0 p.p.m. upfield shift suggesting the ribose adopts C2′-endo conformation rather than the C3′-endo conformation typical in A-RNA. (c) Overlay of 1D 1H spectra showing the imino G-H1 and T/U-H3 resonances at high temperature (35°C). Modified and unmodified duplexes are shown in color and grey, respectively.

Supplementary Figure 2 Lack of detectable exchange across diverse RNA sequence and structural contexts.

(a) Secondary structures with bps showing no detectable RD highlighted in red. (b) Off-resonance RD profiles for the highlighted bps with error bars representing experimental uncertainty (one s.d.) estimated from mono-exponential fitting of n = 6 independently measured peak intensities using a Monte-Carlo based method (Methods). RD profiles not shown for wtTAR have been reported previously (Lee, J. et al., Proc Natl Acad Sci USA. 111, 9485–9490, 2014).

Supplementary Figure 3 1H assignment of duplex A6 constructs with m1A and m1G.

Chemical structures of m1dA–dT and m1dG–dC+ HG bps and color-coded duplexes as in Fig. 3a and Fig. 3b, respectively. Representative 2D NOESY spectra showing NOE connectivity (labeled in orange) indicating HG hydrogen-bonding. Residues in grey are broadened out of detection. Non-exchangeable and exchangeable 1H–1H sequential connectivity used to generate assignments and intra-nucleotide H1′–H8 NOEs used to assess syn conformation are labeled for each residue on the spectra. Resonance assignments for corresponding DNA duplexes A6-DNAm1A and A6-DNAm1G were reported in other studies (Nikolova, E.N. et al., Nature. 470, 498–502, 2011). The NOE walk is indicated using grey arrows on the duplex structure. Interruption of the sequential walk due to exchange broadening of resonances or syn conformation is shown using dashed lines on the spectra as well as the duplex structure.

Supplementary Figure 4 Resonance assignment and NMR spectra for duplex gc and A2 constructs.

2D HSQC or SOFAST-HMQC spectra of m1A or m1G modified DNA (in blue) and RNA (in violet) duplexes with labeled assignments overlaid on their unmodified counterparts (in grey). Folded resonances are indicted using an asterisk. Arrows indicate significant chemical shift perturbations induced by m1A or m1G. Resonances that are exchanged broadened out of detection due to m1A or m1G are highlighted on the corresponding resonance of the unmodified duplexes with a dashed circle. Note that in gc-RNAm1A, there is an additional resonance showing characteristic chemical shifts of a flipped out uridine based on C6-H6 (in red) and C5-H5 (data not shown) chemical shifts. This resonance is likely U5 that is complementary to m1A.

Supplementary Figure 5 1H assignment of duplex gc and A2 constructs with m1A.

Representative 2D NOESY spectra showing NOE connectivity (labeled in orange) indicating HG hydrogen-bonding or syn purine conformation. Residues in grey are broadened out of detection. Non-exchangeable and exchangeable 1H–1H sequential connectivity used to generate assignments and intra-nucleotide H1′–H8 NOEs used to assess syn conformation are labeled for each residue on the spectra. Resonance assignments for A2-DNAm1A will be reported in other study (B.S., H.Z., Y.X., H.M.A., unpublished). Color-coded duplexes are shown as in Fig. 3b. The NOE walk is indicated using grey arrows on the duplex structure. Interruption of the sequential walk due to exchange broadening of resonances or syn conformation is shown using dashed lines on the spectra as well as the duplex structure.

Supplementary Figure 6 NMR analysis of ribonucleotide-substituted A6-DNA.

(a) Shown are overlays of 2D HSQC spectra of A6-DNArA (in red), A6-DNArG (in green), and unmodified A6-DNA (in black) with arrows indicating significant chemical shift perturbations induced by the rA16 or rG10. (b) Shown are 2D HSQC spectra of A6-DNAm1rA (in red), A6-DNAm1rG (in green), overlaid with unmodified (in black) A6-DNArA and A6-DNArG respectively, with arrows indicating significant chemical shift perturbations induced by the m1rA16 or m1rG10. A6-DNArA and A6-DNArG duplexes with13C/15N labeled residues (Methods) colored in red. rA16 and rG10 are shown in bold. Sites used as NMR RD probes are highlighted with a circle. (c) Comparison of 2-state and 3-state BM fitting of RD profiles measured on dC15-C6 in A6-DNArG. Error bars refer to one s.d. estimated from mono-exponential fitting of n = 6 independently measured peak intensities using the Monte-Carlo method (Methods). The data is statistically better satisfied using a 3-state rather than 2-state fit though this minimally impacts the exchange parameters obtained for the HG state (Supplementary Table 4). (d) van’t Hoff analysis (Nikolova, E.N. et al., Nature. 470, 498–502, 2011), using combined data from the current study (dA16-C8 RD at 10ºC) with previously published varying temperature A16-C8 RD data (Nikolova, E.N. et al., Nature. 470, 498–502, 2011), shows that RD of dA16-C8 at 10ºC is in better agreement with the 2-state exchange model with AVG than GS initial alignment of magnetization during the BM fitting. (e) Shown are chemical shift perturbations (Δω = ωHoogsteen – ωWatson-Crick) for syn purine-C8 and purine-C1′ in A-RNA, apical loop or mispairs from experimental data (“EXPT”) and DFT calculations (“DFT”) (Supplementary Note). Error bars shown for fitted parameters from RD measurements represent experimental uncertainty (one s.d.) estimated from mono-exponential fitting using the Monte-Carlo method (Methods).

Supplementary information

Supplementary Text and Figures

Supplementary Figures 1–6, Supplementary Tables 1–5 and Supplementary Note (PDF 2783 kb)

dA–dT WC→HG transition in B-DNA

Shown is the transition of dA16-dT9 bp (in green and purple) from WC to HG bp in a B-DNA duplex (A6-DNA) from the biased MD simulation. dA16 is being flipped around the glycosidic bond under the biasing force. The purple sphere keeps track of dA16-H2, which has a close contact with the 5′-neighbouring bp (dC15-dG10). The steric contact is effectively accommodated by minimal structural adjustment of dC15-dG10 without disrupting hydrogen-bonding. (MOV 1198 kb)

rA–rU WC→HG transition in A-RNA

Shown is a representative transition of rA16-rU9 bp (rA16 in green) from WC to HG bp in an A-RNA hairpin construct (hp-A6-RNA) from the biased MD simulation. In this case, although the biasing force is eventually able to force the rA16 residue shown in bright green to flip 180° around the X angle, it does so at the cost of disrupting the hydrogen bonding of the 5′- neighboring base pair shown in orange and cyan. (MOV 502 kb)

Stable Hoogsteen base pair during unbiased MD simulations for dA-dT HG bp in A6-DNA duplex (MOV 4205 kb)

Stable Hoogsteen base pair during unbiased MD simulations for rA-dT HG bp in A6-DNA duplex (MOV 2547 kb)

Stable Hoogsteen base pair during unbiased MD simulations for m1dA-dT HG bp in A6-DNA duplex (MOV 2926 kb)

m1rA–rU induced melting in hp-A6-RNA

In stark contrast to the stably accommodated m1dA-dT bp in B-DNA (Supplementary Movie 5), m1rA (in grey) leads to the melting of most bps in the unbiased MD simulation. The melting starts near the m1rA -rU bp and then propagates through both sides of the strand, thus leading to melting of almost the entire helix. (MOV 9027 kb)

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Zhou, H., Kimsey, I., Nikolova, E. et al. m1A and m1G disrupt A-RNA structure through the intrinsic instability of Hoogsteen base pairs. Nat Struct Mol Biol 23, 803–810 (2016). https://doi.org/10.1038/nsmb.3270

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