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Discovery and evolution of RNA and XNA reverse transcriptase function and fidelity

Abstract

The ability of reverse transcriptases (RTs) to synthesize a complementary DNA from natural RNA and a range of unnatural xeno nucleic acid (XNA) template chemistries, underpins key methods in molecular and synthetic genetics. However, RTs have proven challenging to discover and engineer, in particular for the more divergent XNA chemistries. Here we describe a general strategy for the directed evolution of RT function for any template chemistry called compartmentalized bead labelling and demonstrate it by the directed evolution of efficient RTs for 2′-O-methyl RNA and hexitol nucleic acids and the discovery of RTs for the orphan XNA chemistries d-altritol nucleic acid and 2′-methoxyethyl RNA, for which previously no RTs existed. Finally, we describe the engineering of XNA RTs with active exonucleolytic proofreading as well as the directed evolution of RNA RTs with very high complementary DNA synthesis fidelities, even in the absence of proofreading.

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Fig. 1: CBL selection scheme.
Fig. 2: XNA reverse transcriptases.
Fig. 3: Characterization of 2′-O-Me-RNA reverse transcriptase C8.
Fig. 4: A reverse transcriptase for PS 2′-MOE RNA.
Fig. 5: Proofreading XNA RTs.
Fig. 6: Selection of high-fidelity RTs.

Data availability

The authors declare that the data supporting the findings of this study are available within the article and its Supplementary Information files.

Code availability

Custom scripts used for RT fidelity analysis in this study can be found at GitHub: https://github.com/holliger-lab/fidelity-analysis

References

  1. 1.

    White, A. K. et al. High-throughput microfluidic single-cell RT-qPCR. Proc. Natl Acad. Sci. USA 108, 13999–14004 (2011).

    CAS  PubMed  Google Scholar 

  2. 2.

    Stark, R., Grzelak, M. & Hadfield, J. RNA sequencing: the teenage years. Nat. Rev. Genet. 20, 631–656 (2019).

    CAS  PubMed  Google Scholar 

  3. 3.

    Schaffitzel, C., Hanes, J., Jermutus, L. & Pluckthun, A. Ribosome display: an in vitro method for selection and evolution of antibodies from libraries. J. Immunol. Methods 231, 119–135 (1999).

    CAS  PubMed  Google Scholar 

  4. 4.

    Taylor, A. I., Houlihan, G. & Holliger, P. Beyond DNA and RNA: The Expanding Toolbox of Synthetic Genetics. Cold Spring Harb. Perspect. Biol. 11, a032490 (2019).

    CAS  PubMed  Google Scholar 

  5. 5.

    Pinheiro, V. B. et al. Synthetic genetic polymers capable of heredity and evolution. Science 336, 341–344 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  6. 6.

    Yu, H., Zhang, S. & Chaput, J. C. Darwinian evolution of an alternative genetic system provides support for TNA as an RNA progenitor. Nat. Chem. 4, 183–187 (2012).

    CAS  PubMed  Google Scholar 

  7. 7.

    Biondi, E. et al. Laboratory evolution of artificially expanded DNA gives redesignable aptamers that target the toxic form of anthrax protective antigen. Nucleic Acids Res. 44, 9565–9577 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. 8.

    Arangundy-Franklin, S. et al. A synthetic genetic polymer with an uncharged backbone chemistry based on alkyl phosphonate nucleic acids. Nat. Chem. 11, 533–542 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. 9.

    Kotewicz, M. L., Dalessio, J. M., Driftmier, K. M., Blodgett, K. P. & Gerard, G. F. Cloning and overexpression of Moloney murine leukemia-virus reverse-transcriptase in Escherichia coli. Gene 35, 249–258 (1985).

    CAS  PubMed  Google Scholar 

  10. 10.

    Ong, J. L., Evans, J. T. & Tanner, N. Compositions and methods relating to variant DNA polymerases and synthetic DNA polymerases. US Patent WO 2013/033528 (2013).

  11. 11.

    Zhao, C. & Pyle, A. M. Crystal structures of a group II intron maturase reveal a missing link in spliceosome evolution. Nat. Struct. Mol. Biol. 23, 558–565 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. 12.

    Dunn, M. R. & Chaput, J. C. Reverse transcription of threose nucleic acid by a naturally occurring DNA polymerase. ChemBioChem 17, 1804–1808 (2016).

    CAS  PubMed  Google Scholar 

  13. 13.

    Wang, Y., Ngor, A. K., Nikoomanzar, A. & Chaput, J. C. Evolution of a general RNA-cleaving FANA enzyme. Nat. Commun. 9, 5067 (2018).

    PubMed  PubMed Central  Google Scholar 

  14. 14.

    Choi, H. M., Beck, V. A. & Pierce, N. A. Next-generation in situ hybridization chain reaction: higher gain, lower cost, greater durability. ACS Nano 8, 4284–4294 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. 15.

    Ayadi, L., Galvanin, A., Pichot, F., Marchand, V. & Motorin, Y. RNA ribose methylation (2′-O-methylation): Occurrence, biosynthesis and biological functions. Biochim. Biophys. Acta Gene Regul. Mech. 1862, 253–269 (2019).

    CAS  PubMed  Google Scholar 

  16. 16.

    Burmeister, P. E. et al. Direct in vitro selection of a 2′-O-methyl aptamer to VEGF. Chem. Biol. 12, 25–33 (2005).

    CAS  PubMed  Google Scholar 

  17. 17.

    Voit, T. et al. Safety and efficacy of drisapersen for the treatment of Duchenne muscular dystrophy (DEMAND II): an exploratory, randomised, placebo-controlled phase 2 study. Lancet Neurol. 13, 987–996 (2014).

    CAS  PubMed  Google Scholar 

  18. 18.

    Chen, T. et al. Evolution of thermophilic DNA polymerases for the recognition and amplification of C2′-modified DNA. Nat. Chem. 8, 556–562 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. 19.

    Cozens, C., Pinheiro, V. B., Vaisman, A., Woodgate, R. & Holliger, P. A short adaptive path from DNA to RNA polymerases. Proc. Natl Acad. Sci. USA 109, 8067–8072 (2012).

    CAS  PubMed  Google Scholar 

  20. 20.

    Kropp, H. M., Betz, K., Wirth, J., Diederichs, K. & Marx, A. Crystal structures of ternary complexes of archaeal B-family DNA polymerases. PLoS One 12, e0188005 (2017).

    PubMed  PubMed Central  Google Scholar 

  21. 21.

    Firbank, S. J., Wardle, J., Heslop, P., Lewis, R. J. & Connolly, B. A. Uracil recognition in archaeal DNA polymerases captured by X-ray crystallography. J. Mol. Biol. 381, 529–539 (2008).

    CAS  PubMed  Google Scholar 

  22. 22.

    Rozners, E. & Moulder, J. Hydration of short DNA, RNA, and 2-OMe oligonucleotides determined by osmotic stressing. Nucleic Acids Res. 32, 6153–6153 (2004).

    CAS  Google Scholar 

  23. 23.

    Allart, B. et al. d‐altritol nucleic acids (ANA): hybridisation properties, stability, and initial structural analysis. Chemistry 5, 2424–2431 (1999).

    CAS  Google Scholar 

  24. 24.

    Freier, S. M. & Altmann, K. H. The ups and downs of nucleic acid duplex stability: structure-stability studies on chemically-modified DNA:RNA duplexes. Nucleic Acids Res. 25, 4429–4443 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. 25.

    Wilds, C. J. & Damha, M. J. Duplex recognition by oligonucleotides containing 2′-deoxy-2′-fluoro-d-arabinose and 2′-deoxy-2′-fluoro-d-ribose. Intermolecular 2′-OH-phosphate contacts versus sugar puckering in the stabilization of triple-helical complexes. Bioconjugate Chem. 10, 299–305 (1999).

    CAS  Google Scholar 

  26. 26.

    Ohuchi, S., Nakano, H. & Yamane, T. In vitro method for the generation of protein libraries using PCR amplification of a single DNA molecule and coupled transcription/translation. Nucleic Acids Res. 26, 4339–4346 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  27. 27.

    Okello, J. B. et al. Quantitative assessment of the sensitivity of various commercial reverse transcriptases based on armored HIV RNA. PLoS One 5, e13931 (2010).

    PubMed  PubMed Central  Google Scholar 

  28. 28.

    Taylor, A. I. et al. Catalysts from synthetic genetic polymers. Nature 518, 427–430 (2015).

    CAS  PubMed  Google Scholar 

  29. 29.

    Mutschler, H. et al. Random-sequence genetic oligomer pools display an innate potential for ligation and recombination. eLife. 7, e43022 (2018).

    PubMed  PubMed Central  Google Scholar 

  30. 30.

    Ottesen, E. W. Iss-N1 makes the first fda-approved drug for spinal muscular atrophy. Transl. Neurosci. 8, 1–6 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. 31.

    Ellefson, J. W. et al. Synthetic evolutionary origin of a proofreading reverse transcriptase. Science 352, 1590–1593 (2016).

    CAS  PubMed  Google Scholar 

  32. 32.

    Herr, A. J., Williams, L. N. & Preston, B. D. Antimutator variants of DNA polymerases. Crit. Rev. Biochem. Mol. 46, 548–570 (2011).

    CAS  Google Scholar 

  33. 33.

    Schmitt, M. W. et al. Detection of ultra-rare mutations by next-generation sequencing. Proc. Natl Acad. Sci. USA 109, 14508–14513 (2012).

    CAS  PubMed  Google Scholar 

  34. 34.

    Potapov, V. et al. Base modifications affecting RNA polymerase and reverse transcriptase fidelity. Nucleic Acids Res. 46, 5753–5763 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. 35.

    Yasukawa, K. et al. Next-generation sequencing-based analysis of reverse transcriptase fidelity. Biochem. Bioph. Res. Commun. 492, 147–153 (2017).

    CAS  Google Scholar 

  36. 36.

    Baranauskas, A. et al. Generation and characterization of new highly thermostable and processive M-MuLV reverse transcriptase variants. Protein Eng. Des. Sel. 25, 657–668 (2012).

    CAS  PubMed  Google Scholar 

  37. 37.

    Arezi, B. & Hogrefe, H. Novel mutations in Moloney Murine Leukemia Virus reverse transcriptase increase thermostability through tighter binding to template-primer. Nucleic. Acids Res. 37, 473–481 (2009).

    CAS  PubMed  Google Scholar 

  38. 38.

    Sauter, K. B. & Marx, A. Evolving thermostable reverse transcriptase activity in a DNA polymerase scaffold. Angew. Chem. Int. Ed. Engl. 45, 7633–7635 (2006).

    CAS  PubMed  Google Scholar 

  39. 39.

    Skirgaila, R., Pudzaitis, V., Paliksa, S., Vaitkevicius, M. & Janulaitis, A. Compartmentalization of destabilized enzyme–mRNA–ribosome complexes generated by ribosome display: a novel tool for the directed evolution of enzymes. Protein Eng. Des. Sel. 26, 453–461 (2013).

    CAS  PubMed  Google Scholar 

  40. 40.

    Ong, J. L., Loakes, D., Jaroslawski, S., Too, K. & Holliger, P. Directed evolution of DNA polymerase, RNA polymerase and reverse transcriptase activity in a single polypeptide. J. Mol. Biol. 361, 537–550 (2006).

    CAS  PubMed  Google Scholar 

  41. 41.

    Kool, E. T. Active site tightness and substrate fit in DNA replication. Annu. Rev. Biochem. 71, 191–219 (2002).

    CAS  PubMed  Google Scholar 

  42. 42.

    Franklin, M. C., Wang, J. M. & Steitz, T. A. Structure of the replicating complex of a pol alpha family DNA polymerase. Cell 105, 657–667 (2001).

    CAS  PubMed  Google Scholar 

  43. 43.

    Reha-Krantz, L. J. DNA polymerase proofreading: Multiple roles maintain genome stability. Biochim. Biophys. Acta Proteins Proteomics 1804, 1049–1063 (2010).

    CAS  Google Scholar 

  44. 44.

    Gouge, J., Ralec, C., Henneke, G. & Delarue, M. Molecular recognition of canonical and deaminated bases by P. abyssi family B DNA polymerase. J. Mol. Biol. 423, 315–336 (2012).

    CAS  PubMed  Google Scholar 

  45. 45.

    Tzelepis, K., Rausch, O. & Kouzarides, T. RNA-modifying enzymes and their function in a chromatin context. Nat. Struct. Mol. Biol. 26, 858–862 (2019).

    CAS  PubMed  Google Scholar 

  46. 46.

    Zhou, H. Q. et al. Evolution of a reverse transcriptase to map N 1-methyladenosine in human messenger RNA. Nat. Methods 16, 1281–1288 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 47.

    Diehl, F. et al. BEAMing: single-molecule PCR on microparticles in water-in-oil emulsions. Nat. Methods 3, 551–559 (2006).

    CAS  PubMed  Google Scholar 

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Acknowledgements

This work was supported by the Medical Research Council (MRC) grant programme no. MC_U105178804 (P.H., G.H. and A.I.T.), the UK Biotechnology and Biological Sciences Research Council grant no. 09-EuroSYNBIO-OP-013) (B.T.P.) and a research collaboration between AstraZeneca UK and the Medical Research Council, MRC-AstraZeneca Blue Sky Grant (S.A.-F. and NS).

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Authors

Contributions

G.H,. S.A.-F. and P.H. conceived and designed experiments. G.H. performed all experiments except structural models (with S.A.-F. and B.T.P.), deep sequencing and data analysis (with B.T.P.) and RT characterization (with A.I.T. and N.S.). All authors analysed data, discussed results and co-wrote the manuscript.

Corresponding author

Correspondence to Philipp Holliger.

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The authors declare no competing interests.

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Extended data

Extended Data Fig. 1 CBL model selections, signal amplification and plasmid capture.

a, Cells expressing RT521K were spiked into an excess of cells expressing inactive RT (RT-ΔDTD) at 1:100 (top) and 1:1000 (bottom) ratios and encapsulated in w/o emulsion with beads coated with DNA primer bbFd and template TnotTest7 (Supplementary Table 7). After in-emulsion cell lysis and primer extension, beads were recovered and sorted for fluorescence using FACS (middle panel, dashed box). Plasmids recovered from sorted beads were amplified by qPCR and the amount of RT521K and RT-ΔDTD plasmids post-CBL selection quantified. Enrichment was calculated by determining the ratio of RT521K: RT-ΔDTD plasmids in the input (before CBL selection) (left panels) and comparing it with the output ratio (after CBL selection)(right panels) yielding enrichment factors of ~500-fold in both cases. b, HCR signal amplification. Cells expressing RT521K were encapsulated with beads coated with primer bbTest7 annealed to RNA template tnog_full (Supplementary Table 7) and RNA reverse transcription was performed in emulsion. After recovering the beads from emulsion, the sample was split in half and beads were treated with either a nucleic acid probe (top bead) or underwent HCR (bottom bead). Flow cytometry analysis of the beads labelled with a direct probe (top plot) or HCR (bottom plot) shows approximately an order of magnitude increase in signal and a greater percentage of fluorescently labelled beads in HCR conditions. c, Plasmid capture on microbeads. Plasmids bound to beads during CBL were quantified by qPCR. Cells expressing RT521K were combined with beads in bulk (in solution) or were encapsulated in w/o emulsions and underwent reverse transcription. Beads in w/o emulsion were extracted and bulk and emulsion treated beads were sorted for fluorescence by HCR. The number of plasmids per bead was quantified before and after FACS. Purified plasmid was bound to beads without capture oligo (untreated beads) and with capture oligo as controls showing approximately 10 plasmids captured per bead in emulsion and stably bound (surviving post-sort).

Extended Data Fig. 2 RT mutation screen.

a, Space filling surface model of the ternary structure of KOD polymerase (PDB ID 5OMF) with primer (nascent) strand (red) and template strand (green) shown. The position of mutations selected for screening are shown in blue. b, ELONA (enzyme-linked oligonucleotide assay)-based RT activity assay scheme: (from left to right) RT reactions are performed with a biotinylated primer bbTest7 (Supplementary Table 7), bound to wells in a streptavidin plate and hybridized to 2′-O-Me RNA template (cyan) TFRst 2′-O-Me (Supplementary Table 7). RT synthesized cDNA (red), which remains bound to the plate after template removal. The presence of the (correct) cDNA can be detected by a specific oligonucleotide probe FitcFd (Supplementary Table 7) labelled with FITC (green), which in turn is detected by an anti-FITC antibody (blue) conjugated to horse-radish peroxidase (yellow star). c, RT mutation activity screen: only mutations S383K, N735K and I114T show an improved signal and when combined (RT-TKK) show more than double the signal of wt (RT521K) (NTC, no template negative control, n = 3).

Extended Data Fig. 3 RT-TKK: mutations and electrostatic surface.

a, Sequence alignment of RT521K and RT-TKK with mutations shown in blue. b, Space filling model of the ternary structure of KOD polymerase (PDB ID 5OMF) with RT-TKK mutations I114T, S383K and N735K (blue), primer strand (red), template (green) and incoming deoxynucleotide triphosphate (orange). c, Zoom in of the uracil binding pocket (UBP) with uracil base bound (PDB ID 2VWJ) and V93Q mutation (orange). (present in both RT-521K and RT-TKK) and I114T mutation (blue spheres) (RT-TKK). Note how V93Q narrows the UBP (compared to wild-type UBP shown in Extended Data Fig. 1a) and sterically excludes uracil from the binding pocket. The mechanistic basis of the I114T mutation improvement of cDNA synthesis on 2′-O-Me RNA is currently unclear. The main chain NH of I114 hydrogen bonds with uracil in the wild-type UBP. Mutation to I114T may alter main chain conformation to disrupt this interaction and this may further improve uracil exclusion. d, Electrostatic potential of the primer/template binding surface of KOD (left) and its change upon N735K and S383K mutations. Note the increase of positively charged surface potential in proximity to the template strand (green), which is likely to enhance template binding.

Extended Data Fig. 4 A reverse transcriptase for 2′-MOE RNA.

a, Chemical structure of 2′-MOE RNA. b, denaturing urea-PAGE of cDNA synthesis on 2′-MOE RNA template (sequence: 10–27 Spinraza (Supplementary Table 7)) and c, RT-PCR of cDNA synthesis. Only RT-C8 can reverse transcribe 2′-MOE RNA (as well as PS 2′-MOE RNA (Fig. 4)).

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Supplementary Methods, Figs. 1–8, Tables 1–11 and References.

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Houlihan, G., Arangundy-Franklin, S., Porebski, B.T. et al. Discovery and evolution of RNA and XNA reverse transcriptase function and fidelity. Nat. Chem. 12, 683–690 (2020). https://doi.org/10.1038/s41557-020-0502-8

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