Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Article
  • Published:

Cutting antiparallel DNA strands in a single active site

Nature Structural & Molecular Biology volume 27, pages 119–126 (2020)Cite this article

Subjects

Abstract

A single enzyme active site that catalyzes multiple reactions is a well-established biochemical theme, but how one nuclease site cleaves both DNA strands of a double helix has not been well understood. In analyzing site-specific DNA cleavage by the mammalian RAG1–RAG2 recombinase, which initiates V(D)J recombination, we find that the active site is reconfigured for the two consecutive reactions and the DNA double helix adopts drastically different structures. For initial nicking of the DNA, a locally unwound and unpaired DNA duplex forms a zipper via alternating interstrand base stacking, rather than melting as generally thought. The second strand cleavage and formation of a hairpin–DNA product requires a global scissor-like movement of protein and DNA, delivering the scissile phosphate into the rearranged active site.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: Reactions catalyzed by RAG recombinase.
Fig. 2: Structure of the DNA zipper in NFC.
Fig. 3: A DNA zipper is associated with untwisted DNA.
Fig. 4: Protein-DNA interactions in NFC.
Fig. 5: Different configurations of L12 loops in PRC, NFC and HFC.
Fig. 6: Cutting two DNA strands in a single active site.

Similar content being viewed by others

Data availability

The accession numbers for the cryo-EM structures and associated density maps of the mouse PRC and NFC complexes reported in this paper have been deposited to the PDB and EMDB under accession codes PDB 6OEM to 6OER and 6V0V and EMD-20030 to EMD-20035, EMD-20038, EMD-20039 and EMD-21003, as specified in Table 1.

References

  1. Mizuuchi, K. Transpositional recombination: mechanistic insights from studies of Mu and other elements. Annu. Rev. Biochem. 61, 1011–1051 (1992).

    CAS  PubMed  Google Scholar 

  2. Gellert, M. V(D)J recombination: RAG proteins, repair factors, and regulation. Annu. Rev. Biochem. 71, 101–132 (2002).

    CAS  PubMed  Google Scholar 

  3. Schatz, D. G. & Swanson, P. C. V(D)J recombination: mechanisms of initiation. Annu. Rev. Genet. 45, 167–202 (2011).

    CAS  PubMed  Google Scholar 

  4. Kim, M. S., Lapkouski, M., Yang, W. & Gellert, M. Crystal structure of the V(D)J recombinase RAG1–RAG2. Nature 518, 507–511 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  5. Sakano, H., Huppi, K., Heinrich, G. & Tonegawa, S. Sequences at the somatic recombination sites of immunoglobulin light-chain genes. Nature 280, 288–294 (1979).

    CAS  PubMed  Google Scholar 

  6. Lewis, S. M. The mechanism of V(D)J joining: lessons from molecular, immunological, and comparative analyses. Adv. Immunol. 56, 27–150 (1994).

    CAS  PubMed  Google Scholar 

  7. Lapkouski, M., Chuenchor, W., Kim, M. S., Gellert, M. & Yang, W. Assembly pathway and characterization of the RAG1/2–DNA paired and signal-end complexes. J. Biol. Chem. 290, 14618–14625 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. Ru, H. et al. Molecular mechanism of V(D)J recombination from synaptic RAG1–RAG2 complex structures. Cell 163, 1138–1152 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. Kim, M. S. et al. Cracking the DNA code for V(D)J recombination. Mol. Cell 70, 358–370 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  10. Ru, H. et al. DNA melting initiates the RAG catalytic pathway. Nat. Struct. Mol. Biol. 25, 732–742 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  11. Davies, D. R., Goryshin, I. Y., Reznikoff, W. S. & Rayment, I. Three-dimensional structure of the Tn5 synaptic complex transposition intermediate. Science 289, 77–85 (2000).

    CAS  PubMed  Google Scholar 

  12. Richardson, J. M., Colloms, S. D., Finnegan, D. J. & Walkinshaw, M. D. Molecular architecture of the Mos1 paired-end complex: the structural basis of DNA transposition in a eukaryote. Cell 138, 1096–1108 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Hare, S., Gupta, S. S., Valkov, E., Engelman, A. & Cherepanov, P. Retroviral intasome assembly and inhibition of DNA strand transfer. Nature 464, 232–236 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Montano, S. P., Pigli, Y. Z. & Rice, P. A. The Mu transpososome structure sheds light on DDE recombinase evolution. Nature 491, 413–417 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Hickman, A. B. et al. Structural basis of hAT transposon end recognition by Hermes, an octameric DNA transposase from Musca domestica. Cell 158, 353–367 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  16. Passos, D. O. et al. Cryo-EM structures and atomic model of the HIV-1 strand transfer complex intasome. Science 355, 89–92 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  17. Yusa, K. piggyBac transposon. Microbiol. Spectr. 3, MDNA3-0028-2014 (2015).

  18. Lesbats, P., Engelman, A. N. & Cherepanov, P. Retroviral DNA integration. Chem. Rev. 116, 12730–12757 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Nowotny, M., Gaidamakov, S. A., Crouch, R. J. & Yang, W. Crystal structures of RNase H bound to an RNA/DNA hybrid: substrate specificity and metal-dependent catalysis. Cell 121, 1005–1016 (2005).

    CAS  PubMed  Google Scholar 

  20. Grundy, G. J., Yang, W. & Gellert, M. Autoinhibition of DNA cleavage mediated by RAG1 and RAG2 is overcome by an epigenetic signal in V(D)J recombination. Proc. Natl Acad. Sci. USA 107, 22487–22492 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. Grundy, G. J., Hesse, J. E. & Gellert, M. Requirements for DNA hairpin formation by RAG1/2. Proc. Natl Acad. Sci. USA 104, 3078–3083 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Mills, J. B. & Hagerman, P. J. Origin of the intrinsic rigidity of DNA. Nucleic Acids Res. 32, 4055–4059 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. Yakovchuk, P., Protozanova, E. & Frank-Kamenetskii, M. D. Base-stacking and base-pairing contributions into thermal stability of the DNA double helix. Nucleic Acids Res. 34, 564–574 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Zgarbova, M. et al. Refinement of the Cornell et al. nucleic acids force field based on reference quantum chemical calculations of glycosidic torsion profiles. J. Chem. Theory Comput. 7, 2886–2902 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Hart, K. et al. Optimization of the CHARMM additive force field for DNA: improved treatment of the BI/BII conformational equilibrium. J. Chem. Theory Comput. 8, 348–362 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  26. Rubio-Cosials, A. et al. Transposase–DNA complex structures reveal mechanisms for conjugative transposition of antibiotic resistance. Cell 173, 208–220 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Atkinson, P. W. hAT transposable elements. Microbiol. Spectr. 3, MDNA3-0054-2014 (2015).

  28. Ru, H., Zhang, P. & Wu, H. Structural gymnastics of RAG-mediated DNA cleavage in V(D)J recombination. Curr. Opin. Struct. Biol. 53, 178–186 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Hickman, A. B. et al. Structural insights into the mechanism of double strand break formation by Hermes, a hAT family eukaryotic DNA transposase. Nucleic Acids Res. 46, 10286–10301 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Ramsden, D. A., McBlane, J. F., van Gent, D. C. & Gellert, M. Distinct DNA sequence and structure requirements for the two steps of V(D)J recombination signal cleavage. EMBO J. 15, 3197–3206 (1996).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Hesse, J. E., Lieber, M. R., Mizuuchi, K. & Gellert, M. V(D)J recombination: a functional definition of the joining signals. Genes Dev. 3, 1053–1061 (1989).

    CAS  PubMed  Google Scholar 

  32. Hu, J. et al. Chromosomal loop domains direct the recombination of antigen receptor genes. Cell 163, 947–959 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Nowotny, M. Retroviral integrase superfamily: the structural perspective. EMBO Rep. 10, 144–151 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Yuan, Y. W. & Wessler, S. R. The catalytic domain of all eukaryotic cut-and-paste transposase superfamilies. Proc. Natl Acad. Sci. USA 108, 7884–7889 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. Boboila, C., Alt, F. W. & Schwer, B. in Adv. Immunol Vol 116 (ed. Alt, F. A.) Ch. 1 (Elsevier, 2012).

  36. Deriano, L. & Roth, D. B. Modernizing the nonhomologous end-joining repertoire: alternative and classical NHEJ share the stage. Annu. Rev. Genet. 47, 433–455 (2013).

    CAS  PubMed  Google Scholar 

  37. Grundy, G. J. et al. Initial stages of V(D)J recombination: the organization of RAG1/2 and RSS DNA in the postcleavage complex. Mol. Cell 35, 217–227 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. Suloway, C. et al. Automated molecular microscopy: the new Leginon system. J. Struct. Biol. 151, 41–60 (2005).

    CAS  PubMed  Google Scholar 

  39. Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Fernandez-Leiro, R. & Scheres, S. H. W. A pipeline approach to single-particle processing in RELION. Acta Crystallogr. D Struct. Biol. 73, 496–502 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Punjani, A., Rubinstein, J. L., Fleet, D. J. & Brubaker, M. A. cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat. Methods 14, 290–296 (2017).

    CAS  PubMed  Google Scholar 

  42. Scheres, S. H. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  43. Bai, X. C., Rajendra, E., Yang, G., Shi, Y. & Scheres, S. H. Sampling the conformational space of the catalytic subunit of human γ-secretase. Elife 4, e11182 (2015).

    PubMed  PubMed Central  Google Scholar 

  44. Swint-Kruse, L. & Brown, C. S. Resmap: automated representation of macromolecular interfaces as two-dimensional networks. Bioinformatics 21, 3327–3328 (2005).

    CAS  PubMed  Google Scholar 

  45. Kucukelbir, A., Sigworth, F. J. & Tagare, H. D. Quantifying the local resolution of cryo-EM density maps. Nat. Methods 11, 63–65 (2014).

    CAS  PubMed  Google Scholar 

  46. Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Barad, B. A. et al. EMRinger: side chain-directed model and map validation for 3D cryo-electron microscopy. Nat. Methods 12, 943–946 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. Páll, S., Abraham, M. J., Kutzner, C., Hess, B. & Lindahl, E. in Solving Software Challenges for Exascale (eds Markidis, S. & Laure, E.) 3–27 (Lecture Notes in Computer Science 8759, Springer, 2015).

  49. Tribello, G. A., Bonomi, M., Branduardi, D., Camilloni, C. & Bussi, G. PLUMED2: new feathers for an old bird. Comput. Phys. Commun. 185, 604–613 (2014).

    CAS  Google Scholar 

  50. Bussi, G., Donadio, D. & Parrinello, M. Canonical sampling through velocity rescaling. J. Chem. Phys. 126, 014101 (2007).

    PubMed  Google Scholar 

  51. Parrinello, M. & Rahman, A. Polymorphic transitions in single crystals: a new molecular dynamics method. J. Appl. Phys. 52, 7182–7190 (1981).

    CAS  Google Scholar 

  52. Domanski, J., Sansom, M. S. P., Stansfeld, P. J. & Best, R. B. Balancing force field protein–lipid interactions to capture transmembrane helix–helix association. J. Chem. Theory Comput. 14, 1706–1715 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. Grosse-Kunstleve, R. W. & Adams, P. D. Substructure search procedures for macromolecular structures. Acta Crystallogr. D Biol. Crystallogr. 59, 1966–1973 (2003).

    CAS  PubMed  Google Scholar 

  54. McCoy, A. J., Storoni, L. C. & Read, R. J. Simple algorithm for a maximum-likelihood SAD function. Acta Crystallogr. D Biol. Crystallogr. 60, 1220–1228 (2004).

    PubMed  Google Scholar 

Download references

Acknowledgements

This research was supported by the National Institute of Diabetes and Digestive and Kidney Diseases (M.G., DK036167; W.Y., DK036147 and DK036144; Z.H.Z., GM071940). The authors acknowledge the use of instruments at the Electron Imaging Center for NanoMachines supported by the NIH (1S10RR23057, 1S10OD018111 and U24GM116792), NSF (DBI-1338135 and DMR-1548924) and CNSI at UCLA.

Author information

Authors and Affiliations

  1. Laboratory of Molecular Biology, NIDDK, National Institutes of Health, Bethesda, MD, USA

    Xuemin Chen, Huaibin Wang, Wei Yang & Martin Gellert

  2. California NanoSystems Institute, University of California, Los Angeles, Los Angeles, CA, USA

    Yanxiang Cui & Z. Hong Zhou

  3. Laboratory of Chemical Physics, NIDDK, National Institutes of Health, Bethesda, MD, USA

    Robert B. Best

  4. Department of Microbiology, Immunology and Molecular Genetics, University of California, Los Angeles, Los Angeles, CA, USA

    Z. Hong Zhou

Authors
  1. Xuemin Chen
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Yanxiang Cui
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Robert B. Best
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Huaibin Wang
    View author publications

    You can also search for this author in PubMed Google Scholar

  5. Z. Hong Zhou
    View author publications

    You can also search for this author in PubMed Google Scholar

  6. Wei Yang
    View author publications

    You can also search for this author in PubMed Google Scholar

  7. Martin Gellert
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

X.C. carried out all experiments and structure determination. Y.C. collected cryo-EM micrographs on the Krios microscope at UCLA and helped with structure determination and refinement. H.W. helped with cryo-EM data collection on the TF20 and Krios at NIH. R.B.B. carried out molecular dynamics simulations. Z.H.Z., W.Y. and M.G. supervised the research project. X.C., R.B.B., W.Y. and M.G. prepared the manuscript.

Corresponding authors

Correspondence to Wei Yang or Martin Gellert.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information Anke Sparmann was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data

Extended Data Fig. 1 Cleavage and cryo-EM analysis of DNA substrates in NFC.

a, b, Cleavage efficiencies (nicking and hairpinning) of the three DNA variants by WT mRAG at 22 and 37 °C (mean and s.d., n= 3 independent samples). c, Percentage of NFC and PRC (NFC/PRC) in cryo-EM 3D classification from samples made of DNA0, DNA1 or DNA2 substrate with E962Q mutant mRAG at 22 and 37 °C. Asterisk (*) indicates that the dataset was collected on a Tecnai F20 electron microscope instead of Titan Krios.

Extended Data Fig. 2 Structure determination of mouse NFC with DNA1 by cryo-EM.

a, Flow chart for cryo-EM data processing of mRAG complexed with DNA1. The maps with red bold letter are used for final model building. b, A surface presentation of the 3.7 Å NFC (DNA1) map (C1 symmetry). Colors are according to the local resolution estimated by ResMap, and the color scale bar is shown on its right. c, Angular distributions of all particles used for the final three-dimensional reconstruction shown in b. d, The FSC curves of the NFC (DNA1) map (C1). The “gold standard” FSC between two independent halves of the map (black line) indicates a resolution of 3.7 Å, and the blue line is the FSC between the final refined model and the final map. e to i, Representative regions of the C1 map (transparent grey surface). The maps are shown with the final structural models (cartoon or stick) superimposed.

Extended Data Fig. 3 Structure determination of mouse NFC with DNA2 by cryo-EM.

a, Flow chart for cryo-EM data processing of mRAG complexed with DNA2. The maps with red bold letter are used for final model building. b, A surface presentation of the 3.3 Å NFC (DNA2) map (C1 symmetry). Colors are according to the local resolution estimated by ResMap, and the color scale bar is shown on its right. c, Angular distributions of all particles used for the final three-dimensional reconstruction shown in b. d, The FSC curves of NFC (DNA2) map (C1). The “gold standard” FSC between two independent halves of the map (black line) indicates a resolution of 3.3 Å, and the blue line is the FSC between the final refined model and the final map. e to i, Representative regions of the C1 map (transparent grey surface). The maps are shown with the final structural models (cartoon or stick) superimposed.

Extended Data Fig. 4 Structural comparisons of mouse PRC and NFC.

a, Cryo-EM structures of NFC with DNA1 (green) and DNA2 (blue) are superimposable. b, Comparison of mouse PRC and NFC structures. Superposition of cryo-EM PRC (red) and NFC (DNA1) (green) structures reveals limited NBD and nonamer movement, which is marked with blue dashed circle (right panel). c, Superposition of crystal (grey) and cryo-EM (red) PRC structures reveals the different NBD and nonamer region (circled in red dashes) due to crystal-lattice contacts. d, e, The zoom-in views of the active center and DNA distortions in the superimposed structures shown in a-b. The catalytic DDE motif and two metal ions (a and b) are labeled; the heptamer of RSS DNA is shown in detailed cartoon presentation; the scissile phosphate of top strand is marked by a large ball. In panel d, bases forming the DNA zipper are labeled. f, A zoom-in view of boxed area in panel b. RAG2 interacts with the minor groove in PRC or the major groove in NFC.

Extended Data Fig. 5 Structure determination of mouse WT NFC with DNA0 by cryo-EM.

a, Flow chart for cryo-EM data processing of WT mRAG complexed with DNA0. Data processing was done using RELION. The 3.6Å map labeled in red was used for final model building. b, A surface presentation of the 3.6 Å map of WT NFC (DNA0). Colors are according to the local resolution estimated by ResMap, and the color scale bar is shown on its right. c, Angular distributions of all particles used for the final three-dimensional reconstruction. d, The FSC curves of WT NFC (DNA0) map. The “gold standard” FSC between two independent halves of the map indicates an overall resolution of 3.6 Å. e–h, Representative regions of the map (transparent grey surface). The refined zipper DNA fits the map better (e) than the melted DNA (PDB: 6DBR) (f).

Extended Data Fig. 6 Molecular simulations of the untwisted region of DNA.

a, b, Unbiased simulations were run starting from the base-flipped out structure (PDB 6DBR) (a), and the zipper structure (b) using the Amber 14 force field. Plotted in each case are the all-atom RMSD to the base-flipped out structure (black) and the zippered structure (red). The structures at the start and end of each run are shown above the RMSD plots. c, d, The analogous results are given for simulations with the CHARMM 36 force field. e, Biased simulations were run from a canonical B-DNA form, in which the terminal residues were driven to mimic the stretched and untwisted DNA observed in the mouse and zebrafish NFC structures. The RMSD and initial and final structures are shown as before.

Extended Data Fig. 7 Structure determination of mouse PRC with DNA0 by cryo-EM.

a, Flow chart for the cryo-EM data processing of mRAG complexed with DNA0. The maps with red bold letter are used for final model building. b, A surface presentation of the 3.6 Å NFC map (C1 symmetry). Colors are according to the local resolution estimated by ResMap, and the color scale bar is shown on its right. c, Angular distributions of all particles used for the final three-dimensional reconstruction. d, The FSC curves of PRC (DNA0) map (C1). The “gold standard” FSC between two independent halves of the map (black line) indicates a resolution of 3.6 Å, and the blue line is the FSC between the final refined model and the final map. e to i, Representative regions of the C1 map (transparent grey surface). The maps are shown with the final structural models (cartoon or stick) superimposed. j, The maps of 23RSS in PRC (DNA0), NFC (DNA1) and NFC (DNA2).

Extended Data Fig. 8 Remodeling of the active site in NFC and HFC.

a, Repositioning of αX in the RNH domain during PRC to NFC transition (both are cryo-EM structures). E962 is far from the active site in PRC (light blue) but is positioned for catalysis in NFC (green). b, Anomalous X-ray scattering of the PRC crystals confirms that one Mn2+ and one Zn2+ are bound to each RAG1 subunit. The anomalous map is contoured at 3σ in red. The blue 2Fo-Fc map (contoured at 1σ) highlights R848, which is buried in the minor groove. c, The reconfigured E962 in HFC (pink) after the first DNA cleavage by nicking (green).

Supplementary information

Supplementary Information

Supplementary Fig. 1 and Supplementary Table 1.

Reporting Summary

Supplementary Video 1

DNA zipper formation. The two base pairs (AC/(TG)) undergoing zipper formation are shown as sticks, and the rest of the DNA is simplified as a tube and ladder cartoon. The active site DDE are shown in red sticks, and divalent cations are represented as green spheres. The second Mg2+ ion appears only at the end because it binds in a fully formed active site with the substrate in place, but not in PRC. The red sphere on the DNA strand marks the scissile phosphate for nicking, and the DNA becomes nicked when two Mg2+ ions are properly bound.

Supplementary Video 2

Morphing of structural transition from PRC to NFC. The mRAG protein is shown as a semitransparent molecular surface. ZnH2 domains (green and light blue cartoon) of RAG1 move significantly and two L12 loops (green and light blue coils) become extended. The second metal ion (green sphere) is captured in the active site (marked by D600, D708 and E962 in red sticks) only when E962 and the scissile phosphate are in the active configuration. The 12/23RSS DNAs are shown as yellow and orange tube-and-ladders.

Supplementary Video 3

Morphing of the structural transition from NFC to HFC. One RAG1–RAG2 heterodimer (bound to 12RSS, yellow) is superimposed between the two structures and shown as a semitransparent molecular surface. The second RAG1–RAG2 heterodimer (bound to 23RSS, orange), shown as a colored cartoon, moves towards RAG-12RSS, delivering the bottom strand on each RSS DNA into the active site. Two L12 loops (green and light blue coil) of RAG1 subunits move away from the RAG1 interface. The configuration of the active center (red stick-and-balls for catalytic residues), except for E962, remains unchanged, and the closing motion positions the scissile phosphates on the bottom strands into the active centers. The green spheres represent Mg2+ ions. The second Mg2+ ion appears only when the active site is fully formed, but not when DNA undergoes conformational changes. When both Mg2+ ions are properly bound in the active site, the DNA cleavage/transesterification reaction takes place as shown.

Supplementary Video 4

The reaction cycle of DNA nicking and hairpin formation catalyzed by RAG recombinase. The scissile phosphates for nicking and hairpinning of DNA1 are highlighted by red and pink spheres. The active site DDE are shown as red sticks, and two metal ions in the NFC and HFC states are shown as green spheres. When the catalytic carboxylates DDE are not fully aligned or the scissile phosphate is not captured in the active site, such as during conformational changes of RAG and DNA, only one metal ion may be retained.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Chen, X., Cui, Y., Best, R.B. et al. Cutting antiparallel DNA strands in a single active site. Nat Struct Mol Biol 27, 119–126 (2020). https://doi.org/10.1038/s41594-019-0363-2

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41594-019-0363-2

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing