Structural basis of sequence-specific cytosine deamination by double-stranded DNA deaminase toxin DddA.
Journal
Nature structural & molecular biology
ISSN: 1545-9985
Titre abrégé: Nat Struct Mol Biol
Pays: United States
ID NLM: 101186374
Informations de publication
Date de publication:
08 2023
08 2023
Historique:
received:
04
09
2022
accepted:
12
06
2023
medline:
23
8
2023
pubmed:
18
7
2023
entrez:
17
7
2023
Statut:
ppublish
Résumé
The interbacterial deaminase toxin DddA catalyzes cytosine-to-uracil conversion in double-stranded (ds) DNA and enables CRISPR-free mitochondrial base editing, but the molecular mechanisms underlying its unique substrate selectivity have remained elusive. Here, we report crystal structures of DddA bound to a dsDNA substrate containing the 5'-TC target motif. These structures show that DddA binds to the minor groove of a sharply bent dsDNA and engages the target cytosine extruded from the double helix. DddA Phe1375 intercalates in dsDNA and displaces the 5' (-1) thymine, which in turn replaces the target (0) cytosine and forms a noncanonical T-G base pair with the juxtaposed guanine. This tandem displacement mechanism allows DddA to locate a target cytosine without flipping it into the active site. Biochemical experiments demonstrate that DNA base mismatches enhance the DddA deaminase activity and relax its sequence selectivity. On the basis of the structural information, we further identified DddA mutants that exhibit attenuated activity or altered substrate preference. Our studies may help design new tools useful in genome editing or other applications.
Identifiants
pubmed: 37460895
doi: 10.1038/s41594-023-01034-3
pii: 10.1038/s41594-023-01034-3
pmc: PMC10442228
doi:
Substances chimiques
Cytosine
8J337D1HZY
DNA
9007-49-2
Uracil
56HH86ZVCT
Cytidine Deaminase
EC 3.5.4.5
Types de publication
Journal Article
Research Support, U.S. Gov't, Non-P.H.S.
Research Support, N.I.H., Extramural
Langues
eng
Sous-ensembles de citation
IM
Pagination
1153-1159Subventions
Organisme : NCI NIH HHS
ID : P01 CA234228
Pays : United States
Organisme : NIGMS NIH HHS
ID : R35 GM118047
Pays : United States
Organisme : NCRR NIH HHS
ID : S10 RR029205
Pays : United States
Organisme : NIGMS NIH HHS
ID : P30 GM124165
Pays : United States
Informations de copyright
© 2023. The Author(s).
Références
Feng, Y., Seija, N., Di Noia, J. M. & Martin, A. AID in antibody diversification: there and back again. Trends Immunol. 41, 586–600 (2020).
pubmed: 32434680
pmcid: 7183997
doi: 10.1016/j.it.2020.04.009
Green, A. M. & Weitzman, M. D. The spectrum of APOBEC3 activity: from anti-viral agents to anti-cancer opportunities. DNA Repair 83, 102700 (2019).
pubmed: 31563041
pmcid: 6876854
doi: 10.1016/j.dnarep.2019.102700
Muramatsu, M. et al. Class switch recombination and hypermutation require activation-induced cytidine deaminase (AID), a potential RNA editing enzyme. Cell 102, 553–563 (2000).
pubmed: 11007474
doi: 10.1016/S0092-8674(00)00078-7
Swanton, C., McGranahan, N., Starrett, G. J. & Harris, R. S. APOBEC enzymes: mutagenic fuel for cancer evolution and heterogeneity. Cancer Discov. 5, 704–712 (2015).
pubmed: 26091828
pmcid: 4497973
doi: 10.1158/2159-8290.CD-15-0344
Komor, A. C., Kim, Y. B., Packer, M. S., Zuris, J. A. & Liu, D. R. Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533, 420–424 (2016).
pubmed: 27096365
pmcid: 4873371
doi: 10.1038/nature17946
Alexandrov, L. B. et al. Signatures of mutational processes in human cancer. Nature 500, 415–421 (2013).
pubmed: 23945592
pmcid: 3776390
doi: 10.1038/nature12477
Jarvis, M. C., Ebrahimi, D., Temiz, N. A. & Harris, R. S. Mutation signatures including APOBEC in cancer cell lines. JNCI Cancer Spectr. 2, pky002 (2018).
pubmed: 29888758
pmcid: 5993214
doi: 10.1093/jncics/pky002
Shi, K. et al. Structural basis for targeted DNA cytosine deamination and mutagenesis by APOBEC3A and APOBEC3B. Nat. Struct. Mol. Biol. 24, 131–139 (2017).
pubmed: 27991903
doi: 10.1038/nsmb.3344
Gaudelli, N. M. et al. Programmable base editing of A·T to G·C in genomic DNA without DNA cleavage. Nature 551, 464–471 (2017).
pubmed: 29160308
pmcid: 5726555
doi: 10.1038/nature24644
Losey, H. C., Ruthenburg, A. J. & Verdine, G. L. Crystal structure of Staphylococcus aureus tRNA adenosine deaminase TadA in complex with RNA. Nat. Struct. Mol. Biol. 13, 153–159 (2006).
pubmed: 16415880
doi: 10.1038/nsmb1047
de Moraes, M. H. et al. An interbacterial DNA deaminase toxin directly mutagenizes surviving target populations. eLife 10, e62967 (2021).
pubmed: 33448264
pmcid: 7901873
doi: 10.7554/eLife.62967
Mok, B. Y. et al. A bacterial cytidine deaminase toxin enables CRISPR-free mitochondrial base editing. Nature 583, 631–637 (2020).
pubmed: 32641830
pmcid: 7381381
doi: 10.1038/s41586-020-2477-4
Guo, J. et al. DdCBE mediates efficient and inheritable modifications in mouse mitochondrial genome. Mol. Ther. Nucleic Acids 27, 73–80 (2022).
pubmed: 34938607
doi: 10.1016/j.omtn.2021.11.016
Kang, B. C. et al. Chloroplast and mitochondrial DNA editing in plants. Nat. Plants 7, 899–905 (2021).
pubmed: 34211132
pmcid: 8289734
doi: 10.1038/s41477-021-00943-9
Lee, H. et al. Mitochondrial DNA editing in mice with DddA-TALE fusion deaminases. Nat. Commun. 12, 1190 (2021).
pubmed: 33608520
pmcid: 7895935
doi: 10.1038/s41467-021-21464-1
Lei, Z. et al. Mitochondrial base editor induces substantial nuclear off-target mutations. Nature 606, 804–811 (2022).
pubmed: 35551512
doi: 10.1038/s41586-022-04836-5
Lim, K., Cho, S. I. & Kim, J. S. Nuclear and mitochondrial DNA editing in human cells with zinc finger deaminases. Nat. Commun. 13, 366 (2022).
pubmed: 35042880
pmcid: 8766470
doi: 10.1038/s41467-022-27962-0
Mok, B. Y. et al. CRISPR-free base editors with enhanced activity and expanded targeting scope in mitochondrial and nuclear DNA. Nat. Biotechnol. 40, 1378–1387 (2022).
Mok, Y. G. et al. Base editing in human cells with monomeric DddA-TALE fusion deaminases. Nat. Commun. 13, 4038 (2022).
pubmed: 35821233
pmcid: 9276701
doi: 10.1038/s41467-022-31745-y
Silva-Pinheiro, P. et al. In vivo mitochondrial base editing via adeno-associated viral delivery to mouse post-mitotic tissue. Nat. Commun. 13, 750 (2022).
pubmed: 35136065
pmcid: 8825850
doi: 10.1038/s41467-022-28358-w
Cho, S. I. et al. Targeted A-to-G base editing in human mitochondrial DNA with programmable deaminases. Cell 185, 1764–1776.e12 (2022).
pubmed: 35472302
doi: 10.1016/j.cell.2022.03.039
Gallagher, L. A. et al. Genome-wide protein-DNA interaction site mapping in bacteria using a double-stranded DNA-specific cytosine deaminase. Nat. Microbiol 7, 844–855 (2022).
pubmed: 35650286
pmcid: 9159945
doi: 10.1038/s41564-022-01133-9
Blanchet, C., Pasi, M., Zakrzewska, K. & Lavery, R. CURVES+ web server for analyzing and visualizing the helical, backbone and groove parameters of nucleic acid structures. Nucleic Acids Res. 39, W68–W73 (2011).
pubmed: 21558323
pmcid: 3125750
doi: 10.1093/nar/gkr316
Teh, A. H. et al. The 1.48 Å resolution crystal structure of the homotetrameric cytidine deaminase from mouse. Biochemistry 45, 7825–7833 (2006).
pubmed: 16784234
doi: 10.1021/bi060345f
Hollis, T., Ichikawa, Y. & Ellenberger, T. DNA bending and a flip-out mechanism for base excision by the helix-hairpin-helix DNA glycosylase, Escherichia coli AlkA. EMBO J. 19, 758–766 (2000).
pubmed: 10675345
pmcid: 305614
doi: 10.1093/emboj/19.4.758
Hong, S. & Cheng, X. DNA base flipping: a general mechanism for writing, reading, and erasing DNA modifications. Adv. Exp. Med. Biol. 945, 321–341 (2016).
pubmed: 27826845
pmcid: 5542066
doi: 10.1007/978-3-319-43624-1_14
Klimasauskas, S., Kumar, S., Roberts, R. J. & Cheng, X. HhaI methyltransferase flips its target base out of the DNA helix. Cell 76, 357–369 (1994).
pubmed: 8293469
doi: 10.1016/0092-8674(94)90342-5
Matthews, M. M. et al. Structures of human ADAR2 bound to dsRNA reveal base-flipping mechanism and basis for site selectivity. Nat. Struct. Mol. Biol. 23, 426–433 (2016).
pubmed: 27065196
pmcid: 4918759
doi: 10.1038/nsmb.3203
Shi, K. et al. Structural basis for recognition of distinct deaminated DNA lesions by endonuclease Q. Proc. Natl Acad. Sci. USA 118, e2021120118 (2021).
pubmed: 33658373
pmcid: 7958190
doi: 10.1073/pnas.2021120118
Slupphaug, G. et al. A nucleotide-flipping mechanism from the structure of human uracil-DNA glycosylase bound to DNA. Nature 384, 87–92 (1996).
pubmed: 8900285
doi: 10.1038/384087a0
Vassylyev, D. G. et al. Atomic model of a pyrimidine dimer excision repair enzyme complexed with a DNA substrate: structural basis for damaged DNA recognition. Cell 83, 773–782 (1995).
pubmed: 8521494
doi: 10.1016/0092-8674(95)90190-6
Yang, C. G. et al. Crystal structures of DNA/RNA repair enzymes AlkB and ABH2 bound to dsDNA. Nature 452, 961–965 (2008).
pubmed: 18432238
pmcid: 2587245
doi: 10.1038/nature06889
Hendershot, J. M. & O’Brien, P. J. Critical role of DNA intercalation in enzyme-catalyzed nucleotide flipping. Nucleic Acids Res. 42, 12681–12690 (2014).
pubmed: 25324304
pmcid: 4227769
doi: 10.1093/nar/gku919
Olmon, E. D. & Delaney, S. Differential ability of five DNA glycosylases to recognize and repair damage on nucleosomal DNA. ACS Chem. Biol. 12, 692–701 (2017).
pubmed: 28085251
pmcid: 6557264
doi: 10.1021/acschembio.6b00921
Rohs, R. et al. Origins of specificity in protein-DNA recognition. Annu. Rev. Biochem. 79, 233–269 (2010).
pubmed: 20334529
pmcid: 3285485
doi: 10.1146/annurev-biochem-060408-091030
Jurrus, E. et al. Improvements to the APBS biomolecular solvation software suite. Protein Sci. 27, 112–128 (2018).
pubmed: 28836357
doi: 10.1002/pro.3280
Vonrhein, C. et al. Data processing and analysis with the autoPROC toolbox. Acta Crystallogr. D Biol. Crystallogr. 67, 293–302 (2011).
pubmed: 21460447
pmcid: 3069744
doi: 10.1107/S0907444911007773
Kabsch, W. XDS. Acta Crystallogr/ D Biol. Crystallogr. 66, 125–132 (2010).
Winn, M. D. et al. Overview of the CCP4 suite and current developments. Acta Crystallogr. D Biol. Crystallogr. 67, 235–242 (2011).
pubmed: 21460441
pmcid: 3069738
doi: 10.1107/S0907444910045749
Evans, P. R. An introduction to data reduction: space-group determination, scaling and intensity statistics. Acta Crystallogr. D Biol. Crystallogr. 67, 282–292 (2011).
pubmed: 21460446
pmcid: 3069743
doi: 10.1107/S090744491003982X
Evans, P. R. & Murshudov, G. N. How good are my data and what is the resolution? Acta Crystallogr. D Biol. Crystallogr. 69, 1204–1214 (2013).
pubmed: 23793146
pmcid: 3689523
doi: 10.1107/S0907444913000061
French, S. & Wilson, K. On the treatment of negative intensity observations. Acta Crystallogr. A Cryst. Phys. Diffr. Theor. Gen. Crystallogr. 34, 517–525 (1978).
doi: 10.1107/S0567739478001114
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007).
pubmed: 19461840
pmcid: 2483472
doi: 10.1107/S0021889807021206
Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).
pubmed: 20383002
pmcid: 2852313
doi: 10.1107/S0907444910007493
Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr. 66, 213–221 (2010).
pubmed: 20124702
pmcid: 2815670
doi: 10.1107/S0907444909052925
Karplus, P. A. & Diederichs, K. Linking crystallographic model and data quality. Science 336, 1030–1033 (2012).
pubmed: 22628654
pmcid: 3457925
doi: 10.1126/science.1218231