DNA double-strand breaks (DSBs) are a highly cytotoxic form of DNA damage and the incorrect repair of DSBs is linked to carcinogenesis1,2. The conserved error-prone non-homologous end joining (NHEJ) pathway has a key role in determining the effects of DSB-inducing agents that are used to treat cancer as well as the generation of the diversity in antibodies and T cell receptors2,3. Here we applied single-particle cryo-electron microscopy to visualize two key DNA–protein complexes that are formed by human NHEJ factors. The Ku70/80 heterodimer (Ku), the catalytic subunit of the DNA-dependent protein kinase (DNA-PKcs), DNA ligase IV (LigIV), XRCC4 and XLF form a long-range synaptic complex, in which the DNA ends are held approximately 115 Å apart. Two DNA end-bound subcomplexes comprising Ku and DNA-PKcs are linked by interactions between the DNA-PKcs subunits and a scaffold comprising LigIV, XRCC4, XLF, XRCC4 and LigIV. The relative orientation of the DNA-PKcs molecules suggests a mechanism for autophosphorylation in trans, which leads to the dissociation of DNA-PKcs and the transition into the short-range synaptic complex. Within this complex, the Ku-bound DNA ends are aligned for processing and ligation by the XLF-anchored scaffold, and a single catalytic domain of LigIV is stably associated with a nick between the two Ku molecules, which suggests that the joining of both strands of a DSB involves both LigIV molecules.
This is a preview of subscription content, access via your institution
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 51 print issues and online access
$199.00 per year
only $3.90 per issue
Rent or buy this article
Prices vary by article type
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
Cryo-EM density maps have been deposited in the Electron Microscopy Data Bank under accession numbers EMD-23510 (overall LR complex), EMD-23511 (DNA-PK–N-BRCT in the LR complex), EMD-23512 (LigIV–XRCC4–XLF–XRCC4–LigIV in the LR complex), EMD-23509 (overall SR complex), EMD-23513 (distal Ku–LigIV catalytic domain in the SR complex), EMD-23514 (proximal Ku in the SR complex) and EMD-23515 (LigIV–XRCC4–XLF–XRCC4–LigIV in the SR complex). Model coordinates have been deposited in the Protein Data Bank under accession numbers 7LT3 (the LR complex) and 7LSY (the SR complex).
Aplan, P. D. Causes of oncogenic chromosomal translocation. Trends Genet. 22, 46–55 (2006).
Zhao, B., Rothenberg, E., Ramsden, D. A. & Lieber, M. R. The molecular basis and disease relevance of non-homologous DNA end joining. Nat. Rev. Mol. Cell Biol. 21, 765–781 (2020).
Malu, S., Malshetty, V., Francis, D. & Cortes, P. Role of non-homologous end joining in V(D)J recombination. Immunol. Res. 54, 233–246 (2012).
Gottlieb, T. M. & Jackson, S. P. The DNA-dependent protein kinase: requirement for DNA ends and association with Ku antigen. Cell 72, 131–142 (1993).
Yin, X., Liu, M., Tian, Y., Wang, J. & Xu, Y. Cryo-EM structure of human DNA-PK holoenzyme. Cell Res. 27, 1341–1350 (2017).
Mari, P.-O. et al. Dynamic assembly of end-joining complexes requires interaction between Ku70/80 and XRCC4. Proc. Natl Acad. Sci. USA 103, 18597–18602 (2006).
Yano, K. & Chen, D. J. Live cell imaging of XLF and XRCC4 reveals a novel view of protein assembly in the non-homologous end-joining pathway. Cell Cycle 7, 1321–1325 (2008).
DeFazio, L. G., Stansel, R. M., Griffith, J. D. & Chu, G. Synapsis of DNA ends by DNA-dependent protein kinase. EMBO J. 21, 3192–3200 (2002).
Chen, L., Trujillo, K., Sung, P. & Tomkinson, A. E. Interactions of the DNA ligase IV-XRCC4 complex with DNA ends and the DNA-dependent protein kinase. J. Biol. Chem. 275, 26196–26205 (2000).
Ramsden, D. A. & Gellert, M. Ku protein stimulates DNA end joining by mammalian DNA ligases: a direct role for Ku in repair of DNA double-strand breaks. EMBO J. 17, 609–614 (1998).
Riballo, E. et al. XLF-Cernunnos promotes DNA ligase IV-XRCC4 re-adenylation following ligation. Nucleic Acids Res. 37, 482–492 (2009).
Andres, S. N. et al. A human XRCC4-XLF complex bridges DNA. Nucleic Acids Res. 40, 1868–1878 (2012).
Lees-Miller, S. P., Chen, Y. R. & Anderson, C. W. Human cells contain a DNA-activated protein kinase that phosphorylates simian virus 40 T antigen, mouse p53, and the human Ku autoantigen. Mol. Cell. Biol. 10, 6472–6481 (1990).
Meek, K., Douglas, P., Cui, X., Ding, Q. & Lees-Miller, S. P. trans Autophosphorylation at DNA-dependent protein kinase’s two major autophosphorylation site clusters facilitates end processing but not end joining. Mol. Cell. Biol. 27, 3881–3890 (2007).
Uematsu, N. et al. Autophosphorylation of DNA-PKCS regulates its dynamics at DNA double-strand breaks. J. Cell Biol. 177, 219–229 (2007).
Mahaney, B. L., Hammel, M., Meek, K., Tainer, J. A. & Lees-Miller, S. P. XRCC4 and XLF form long helical protein filaments suitable for DNA end protection and alignment to facilitate DNA double strand break repair. Biochem. Cell Biol. 91, 31–41 (2013).
Chang, H. H. Y. et al. Different DNA end configurations dictate which NHEJ components are most important for joining efficiency. J. Biol. Chem. 291, 24377–24389 (2016).
Conlin, M. P. et al. DNA ligase IV guides end-processing choice during nonhomologous end joining. Cell Rep. 20, 2810–2819 (2017).
Zhao, B. et al. The essential elements for the noncovalent association of two DNA ends during NHEJ synapsis. Nat. Commun. 10, 3588 (2019).
Sibanda, B. L., Chirgadze, D. Y., Ascher, D. B. & Blundell, T. L. DNA-PKcs structure suggests an allosteric mechanism modulating DNA double-strand break repair. Science 355, 520–524 (2017).
Sharif, H. et al. Cryo-EM structure of the DNA-PK holoenzyme. Proc. Natl Acad. Sci. USA 114, 7367–7372 (2017).
Walker, J. R., Corpina, R. A. & Goldberg, J. Structure of the Ku heterodimer bound to DNA and its implications for double-strand break repair. Nature 412, 607–614 (2001).
Andres, S. N., Modesti, M., Tsai, C. J., Chu, G. & Junop, M. S. Crystal structure of human XLF: a twist in nonhomologous DNA end-joining. Mol. Cell 28, 1093–1101 (2007).
Wu, P.-Y. et al. Structural and functional interaction between the human DNA repair proteins DNA ligase IV and XRCC4. Mol. Cell. Biol. 29, 3163–3172 (2009).
Kaminski, A. M. et al. Structures of DNA-bound human ligase IV catalytic core reveal insights into substrate binding and catalysis. Nat. Commun. 9, 2642 (2018).
Graham, T. G. W., Walter, J. C. & Loparo, J. J. Two-Stage synapsis of DNA ends during non-homologous end joining. Mol. Cell 61, 850–858 (2016).
Graham, T. G. W., Carney, S. M., Walter, J. C. & Loparo, J. J. A single XLF dimer bridges DNA ends during nonhomologous end joining. Nat. Struct. Mol. Biol. 25, 877–884 (2018).
Carney, S. M. et al. XLF acts as a flexible connector during non-homologous end joining. eLife 9, e61920 (2020).
Nemoz, C. et al. XLF and APLF bind Ku80 at two remote sites to ensure DNA repair by non-homologous end joining. Nat. Struct. Mol. Biol. 25, 971–980 (2018).
Chaplin, A. K. et al. Dimers of DNA-PK create a stage for DNA double-strand break repair. Nat. Struct. Mol. Biol. 28, 13–19 (2021).
Andres, S. N. & Junop, M. S. Crystallization and preliminary X-ray diffraction analysis of the human XRCC4-XLF complex. Acta Crystallogr. F 67, 1399–1402 (2011).
Ropars, V. et al. Structural characterization of filaments formed by human XRCC4-Cernunnos/XLF complex involved in nonhomologous DNA end-joining. Proc. Natl Acad. Sci. USA 108, 12663–12668 (2011).
Hammel, M. et al. XRCC4 protein interactions with XRCC4-like factor (XLF) create an extended grooved scaffold for DNA ligation and double strand break repair. J. Biol. Chem. 286, 32638–32650 (2011).
Doré, A. S. et al. Structure of an Xrcc4-DNA ligase IV yeast ortholog complex reveals a novel BRCT interaction mode. DNA Repair 5, 362–368 (2006).
Costantini, S., Woodbine, L., Andreoli, L., Jeggo, P. A. & Vindigni, A. Interaction of the Ku heterodimer with the DNA ligase IV/Xrcc4 complex and its regulation by DNA-PK. DNA Repair 6, 712–722 (2007).
Normanno, D. et al. Mutational phospho-mimicry reveals a regulatory role for the XRCC4 and XLF C-terminal tails in modulating DNA bridging during classical non-homologous end joining. eLife 6, e22900 (2017).
Tate, J. G. et al. COSMIC: the Catalogue Of Somatic Mutations In Cancer. Nucleic Acids Res. 47, D941–D947 (2019).
Lees-Miller, J. P. et al. Uncovering DNA-PKcs ancient phylogeny, unique sequence motifs and insights for human disease. Prog. Biophys. Mol. Biol. https://doi.org/10.1016/j.pbiomolbio.2020.09.010 (2020).
Chen, X. et al. Structure of an activated DNA-PK and its implications for NHEJ. Mol. Cell 81, 801–810 (2021).
Yang, H. et al. mTOR kinase structure, mechanism and regulation. Nature 497, 217–223 (2013).
Bao, Z. Q., Jacobsen, D. M. & Young, M. A. Briefly bound to activate: transient binding of a second catalytic magnesium activates the structure and dynamics of CDK2 kinase for catalysis. Structure 19, 675–690 (2011).
Baretić, D. et al. Structures of closed and open conformations of dimeric human ATM. Sci. Adv. 3, e1700933 (2017).
Rao, Q. et al. Cryo-EM structure of human ATR-ATRIP complex. Cell Res. 28, 143–156 (2018).
Hammel, M. et al. An intrinsically disordered APLF links Ku, DNA-PKcs, and XRCC4-DNA ligase IV in an extended flexible non-homologous end joining complex. J. Biol. Chem. 291, 26987–27006 (2016).
Pryor, J. M. et al. Ribonucleotide incorporation enables repair of chromosome breaks by nonhomologous end joining. Science 361, 1126–1129 (2018).
Riballo, E. et al. Cellular and biochemical impact of a mutation in DNA ligase IV conferring clinical radiosensitivity. J. Biol. Chem. 276, 31124–31132 (2001).
Wang, Y., Lamarche, B. J. & Tsai, M.-D. Human DNA ligase IV and the ligase IV/XRCC4 complex: analysis of nick ligation fidelity. Biochemistry 46, 4962–4976 (2007).
Tsai, C. J., Kim, S. A. & Chu, G. Cernunnos/XLF promotes the ligation of mismatched and noncohesive DNA ends. Proc. Natl Acad. Sci. USA 104, 7851–7856 (2007).
Gerodimos, C. A., Chang, H. H. Y., Watanabe, G. & Lieber, M. R. Effects of DNA end configuration on XRCC4-DNA ligase IV and its stimulation of Artemis activity. J. Biol. Chem. 292, 13914–13924 (2017).
Stinson, B. M., Moreno, A. T., Walter, J. C. & Loparo, J. J. A mechanism to minimize errors during non-homologous end joining. Mol. Cell 77, 1080–1091 (2020).
Kysela, B. et al. Ku stimulation of DNA ligase IV-dependent ligation requires inward movement along the DNA molecule. J. Biol. Chem. 278, 22466–22474 (2003).
Goodarzi, A. A. & Lees-Miller, S. P. Biochemical characterization of the ataxia-telangiectasia mutated (ATM) protein from human cells. DNA Repair 3, 753–767 (2004).
Yu, Y. et al. DNA-PK and ATM phosphorylation sites in XLF/Cernunnos are not required for repair of DNA double strand breaks. DNA Repair 7, 1680–1692 (2008).
Han, Y., Reyes, A. A., Malik, S. & He, Y. Cryo-EM structure of SWI/SNF complex bound to a nucleosome. Nature 579, 452–455 (2020).
Patel, A., Toso, D., Litvak, A. & Nogales, E. Efficient graphene oxide coating improves cryo-EM sample preparation and data collection from tilted grids. Preprint at https://doi.org/10.1101/2021.03.08.434344 (2021).
Suloway, C. et al. Automated molecular microscopy: the new Leginon system. J. Struct. Biol. 151, 41–60 (2005).
Mastronarde, D. N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol. 152, 36–51 (2005).
Lander, G. C. et al. Appion: an integrated, database-driven pipeline to facilitate EM image processing. J. Struct. Biol. 166, 95–102 (2009).
Voss, N. R., Yoshioka, C. K., Radermacher, M., Potter, C. S. & Carragher, B. DoG Picker and TiltPicker: software tools to facilitate particle selection in single particle electron microscopy. J. Struct. Biol. 166, 205–213 (2009).
Mindell, J. A. & Grigorieff, N. Accurate determination of local defocus and specimen tilt in electron microscopy. J. Struct. Biol. 142, 334–347 (2003).
van Heel, M., Harauz, G., Orlova, E. V., Schmidt, R. & Schatz, M. A new generation of the IMAGIC image processing system. J. Struct. Biol. 116, 17–24 (1996).
Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).
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).
Kimanius, D., Forsberg, B. O., Scheres, S. H. W. & Lindahl, E. Accelerated cryo-EM structure determination with parallelisation using GPUs in RELION-2. eLife 5, e18722 (2016).
Zhang, K. Gctf: real-time CTF determination and correction. J. Struct. Biol. 193, 1–12 (2016).
Rohou, A. & Grigorieff, N. CTFFIND4: fast and accurate defocus estimation from electron micrographs. J. Struct. Biol. 192, 216–221 (2015).
Scheres, S. H. W. & Chen, S. Prevention of overfitting in cryo-EM structure determination. Nat. Methods 9, 853–854 (2012).
Zivanov, J. et al. New tools for automated high-resolution cryo-EM structure determination in RELION-3. eLife 7, e42166 (2018).
Kucukelbir, A., Sigworth, F. J. & Tagare, H. D. Quantifying the local resolution of cryo-EM density maps. Nat. Methods 11, 63–65 (2014).
Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
Sánchez-García, R. et al. DeepEMhacer: a deep learning solution for cryo-EM volume post-processing. Preprint at https://doi.org/10.1101/2020.06.12.148296 (2020).
Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D 66, 486–501 (2010).
Croll, T. I. ISOLDE: a physically realistic environment for model building into low-resolution electron-density maps. Acta Crystallogr. D 74, 519–530 (2018).
Saltzberg, D. J. et al. SSEThread: integrative threading of the DNA-PKcs sequence based on data from chemical cross-linking and hydrogen deuterium exchange. Prog. Biophys. Mol. Biol. 147, 92–102 (2019).
Hepburn, M. et al. The active DNA-PK holoenzyme occupies a tensed state in a staggered synaptic complex. Structure https://doi.org/10.1016/j.str.2020.12.006 (2021).
Yang, J. et al. Improved protein structure prediction using predicted interresidue orientations. Proc. Natl Acad. Sci. USA 117, 1496–1503 (2020).
Afonine, P. V. et al. Real-space refinement in PHENIX for cryo-EM and crystallography. Acta Crystallogr. D 74, 531–544 (2018).
Chang, H. H. Y., Watanabe, G. & Lieber, M. R. Unifying the DNA end-processing roles of the Artemis nuclease: Ku-dependent Artemis resection at blunt DNA ends. J. Biol. Chem. 290, 24036–24050 (2015).
We thank J. Pattie for computer support, J. Meyers, R. M. Haynes and H. Scott at the PNCC for data collection support, D. Ramsden for the gift of a rabbit anti-phosphoT2609 reagent, A. Rosenzweig and I. Radhakrishnan for discussion and comments on the manuscript. This work was supported by a Cornew Innovation Award from the Chemistry of Life Processes Institute at Northwestern University (to Y.H.), a Catalyst Award by the Chicago Biomedical Consortium with support from the Searle Funds at The Chicago Community Trust (to Y.H.), an Institutional Research Grant from the American Cancer Society (IRG-15-173-21 to Y.H.), an H Foundation Core Facility Pilot Project Award (to Y.H.), a Pilot Project Award under U54CA193419 (to Y.H.) and NIH grant R01 GM135651 (to Y.H.). S.C. is supported by the Molecular Biophysics Training Program from NIGMS/NIH (5T32 GM008382). A portion of this research was supported by NIH grant U24GM129547 and performed at the PNCC at OHSU and accessed through EMSL (grid.436923.9), a DOE Office of Science User Facility sponsored by the Office of Biological and Environmental Research. This work used the Sapphire imager from the Northwestern University Keck Biophysics Facility funded by NIH grant 1S10OD026963-01, as well as the resources of the Northwestern University Structural Biology Facility, which is generously supported by NCI CCSG P30 CA060553 grant awarded to the Robert H. Lurie Comprehensive Cancer Center. The Gatan K2 direct electron detector was purchased with funds provided by the Chicago Biomedical Consortium with support from the Searle Funds at The Chicago Community Trust. Work in the Lees-Miller laboratory was supported by Canadian Institutes of Health grant 16939 and the Engineered Air Chair in Cancer Research. Work in the Tomkinson laboratory was supported by NIH grant R01GM047251, as well as the University of New Mexico Comprehensive Cancer Center supported by NCI CCSG P30 CA118100 grant. The collaboration between the He, Lees-Miller and Tomkinson laboratories was supported by NCI P01 CA092584.
The authors declare no competing interests.
Peer review information Nature thanks the anonymous reviewers for their contribution to the peer review of this work. Peer reviewer reports are available.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
a, Schematic showing the Y35 blunt-end DNA substrate. Complex assembly was attempted (supplying DNA-PKcs, Ku, XLF and LigIV–XRCC4) before purification by RNase H elution. b, A representative negative-staining raw micrograph of the complex assembled as described in a. The raw micrograph is representative of 24 micrographs. c, Representative two-dimensional class averages of the complex assembled as described in a, showing the appearance of only the DNA-PK complex despite the addition of XLF and LigIV–XRCC4. d–f, Same procedure as a–c, showing the complex assembly with the same Y35 substrate, but adding XLF and LigIV–XRCC4 to the purified DNA-PK complex after RNase H elution. The raw micrograph is representative of 27 micrographs. In f, the two-dimensional class averages representing the characteristic view of scarce but existing LR complex were obtained. g–i, Same procedure as a–c, showing the complex assembly using Y30–T40–c8 DNA substrate with 40 nt flexible poly(T) and 8 bp of complementary ends as the 3′ overhang. Although the single-stranded poly(T) overhang and the 8-bp complementary region contribute to the stability of the complex, they are not observed in any part of the reconstructed density map, presumably because these single-strand DNA tethers are too flexible to be aligned with the rest of the complex. The raw micrograph is representative of 24 micrographs. The complex was assembled before RNase H elution as described in a. In i, the majority of the two-dimensional classes correspond to the LR complex. j, A representative cryo-EM raw micrograph (out of 17,114 micrographs in total) of the LR complex assembled with the Y30–T40–c8 DNA substrate shown in g. k, Representative two-dimensional class averages of particles (329,784 in total) contributing to the final reconstruction of the LR complex. l, Silver-stained SDS–PAGE (4–12% gradient, biologically replicated three times) showing the input purified subunits (Ku, DNA-PKcs, LigIV–XRCC4, and XLF) and the RNase H purified LR and SR complex for cryo-EM data collection. All representative micrographs in b, e, h, j are from at least three biologically replicated experiments. For gel source data, see Supplementary Fig. 1. m, Protein–protein interaction network between the components of the LR complex. Major unmodelled regions are shown in grey. Well-documented hetero- or homo-dimers are grouped by red dashed lines. Alternative protein–protein interactions are depicted by black dashed lines. The globular domain within the Ku80 C-terminal region (CTR) is completely flexible in the LR complex, and we do not see evidence of the Ku80 CTR domain swap that was previously observed30. The putative distance between one Ku80 CTR globular domain and the other copy of the Ku80 C-terminal helix is too far to be reached by the 18-amino acid linker within Ku80. BRCTs, tandem BRCT domains; CC, coiled-coil domain; CTD, C-terminal domain; HD, head domain; KD, kinase domain; M-HEAT, middle HEAT domain; N-HEAT, N-terminal HEAT domain; NTD, N-terminal domain; OBD, OB-fold domain.
a, Flow chart of the cryo-EM data processing procedure. The gold-standard Fourier shell correlation (FSC) curves (0.143 cut-off) show the final resolution of the holo-complex and each body. b, sample maps and fitted models of DNA-PKcs (olive) and dsDNA substrate (cyan) from the LR complex are shown at 4.1 Å resolution. Maps are shown as transparent surfaces and models are shown as sticks.
Extended Data Fig. 3 Comparing the structure of Ku among the LR synaptic complex, the SR synaptic complex and previously published models.
a, Ku structure in the LR complex showing outward rotations of both Ku70 and Ku80 vWA domains. b, Crystal structure of the XLF Ku-binding motif (KBM) bound to Ku showing the outward rotation of only the Ku80 vWA domain29. c, Crystal structure of apo Ku showing no rotation of either Ku70 or Ku80 vWA domains22. d, Conformation of Ku shown in the cryo-EM structure of the apo DNA-PK complex30. The Ku70 vWA domain is rotated outward, triggered by binding of DNA-PKcs. e, f, Two copies of the XLF Ku-binding motif bound to Ku in the SR complex. The conformation of both copies is the same as the one in the LR complex (a), despite the fact that DNA-PKcs is not present. Colour codes for Ku70 and Ku80 are the same as in Fig. 1.
Extended Data Fig. 4 Comparing the structure of the LigIV–XRCC4–XLF scaffold among the LR synaptic complex, the SR synaptic complex and previously published models.
a, Structure of XRCC4–XLF from the LR complex is shown in comparison to the XRCC4–XLF filamentous repeat crystal structures and the ones from the SR complex (both copies). The XLF dimer is used to align all of the models shown here. Solid lines are aligned with the coiled-coils of XLF (vertical) and XRCC4 (tilted), and the angles in between are shown. Dashed lines are aligned with the C-terminal half of the coiled-coil domain of XRCC4 when full helices are present, and the bending angles are also shown. b, XRCC4 in the crystal structure of the human and yeast LigIV–XRCC4 complexes24,34 are shown after aligning with XRCC4 in the LR complex shown in a. The bending of the coiled-coil domain of XRCC4 is more similar to the one in the SR than in the LR complex. c, Structure of the LigIV–XRCC4 complex from the LR complex is shown in comparison to human and yeast LigIV–XRCC4 crystal structures and ones from the SR complex (both copies). Colour codes for XLF, XRCC4 and LigIV BRCT domains are the same as in Fig. 1.
Extended Data Fig. 5 Surface electrostatic potential and conservation of different areas in the LR synaptic complex.
a, Magnified view of the interaction surface between LigIV N-BRCT domain and Ku70 vWA domains coloured by sequence conservation. b, Magnified view of the DNA-PKcs–DNA-PKcs interaction surface coloured by sequence conservation. c, Surface electrostatic potential view of DNA-PKcs near its FAT domain, showing its negatively charged interface with the XRCC4 C-terminal region (ribbon). The approximate path of the XRCC4 C-terminal peptide containing multiple phosphorylation sites is depicted. The sphere depicts the location of a cancer-associated truncation mutation that occurs at the interface. d, Surface electrostatic potential view of DNA-PKcs DEB and DEB-A helix. The DNA-interaction surface is positively charged. When models are not coloured by either surface electrostatic potential or sequence conservation, the colour codes are the same as in Fig. 1. We cannot rule out the unlikely possibility that the stabilization of the DEB helix is due to the presence of a 3′ overhang that existed in our DNA substrate design (Extended Data Fig. 1g).
Extended Data Fig. 6 Comparing the dimerization of DNA-PKcs in the LR synaptic complex with other PIKK family dimers.
a, Structure of the two DNA-PKcs in the LR complex. The kinase domain is aligned with the homologous domains in b and c as an anchor point. b, c, Dimer of ATR–ATRIP (b) and ATM (c) showing the aligned kinase domain and corresponding N-HEAT regions in the aligned copy. The symmetric-look front views are shown at the bottom left corner. Each protomer of ATR–ATRIP and ATM is coloured the same as the corresponding DNA-PKcs protomer, in olive (the aligned copy) and dark khaki (the other copy). d, Domain organization of DNA-PKcs compared with ATR and ATM. Abbreviations are as in Fig. 1. In our model, both the ABCDE (T2609, S2612, T2620, S2624, T2638 and T2647) and the PQR (S2023, S2029, S2041, S2053 and S2056) phosphorylation sites are located within disordered loops of DNA-PKcs 2606–2720 and 1993–2084, respectively (Fig. 2b, d). The kinase active centre from the opposite side cannot reach most of the ABCDE sites unless the YRPD-interaction (YRPD-I) loop (residues 2586–2604) is peeled off from the YRPD motif (Fig. 2d). In turn, this conformational change potentially disrupts the DNA-PKcs–DNA-PKcs dimerization interface through loop 2569–2585 (Fig. 1d). Similarly, some PQR sites are located too far from the trans kinase active centre. PQR-autophosphorylation-induced changes could have a direct effect on the Ku80 CTR–DNA-PKcs interface at the bottom of the cradle (Fig. 2d), potentially inducing the domain swap of Ku80 that was previously observed30.
a, Schematic showing the Y30–c4 DNA substrate with 4-nt 3′ complementary overhang. The complex was assembled before RNase H elution. b, A representative negative-staining raw micrograph of the complex assembled as described in a. The raw micrograph is representative of 23 micrographs. c, Representative two-dimensional class averages of the complex assembled as described in a. d, Cryo-EM map reconstructed from a small dataset using Y30–c4 DNA substrate shown in a. The map is coloured according to the estimation of the local resolution. e–h, Same procedure as a–d, showing the complex assembly with the Y30 blunt-end substrate. Stably assembled SR complexes on DNA substrates with either complementary or blunt ends indicates that these complexes are stable in the absence of any bridging effect from DNA. The raw micrograph is representative of 24 micrographs. i–l, Same procedure as a–d, showing the complex assembly with the Y14–T2–c20–n10–10 substrate, with one central single non-ligatable nick. The raw micrograph is representative of 24 micrographs. Strand e is added last after mixing the two halves together with NHEJ factors. m, A representative cryo-EM raw micrograph (out of 32,723 total images) of the SR complex assembled with the Y14–T2–c20–n10–10 DNA substrate shown in i. n, Representative two-dimensional class averages of particles (175,866 in total) contributing to the final reconstruction of the SR complex. All representative micrographs in b, f, j, m are from at least three biologically replicated experiments. o, Protein–protein interaction network between the components of the SR complex. Major unmodelled regions are shown in grey. Well-documented hetero- or homo-dimers are grouped by red dashed circles. Alternative protein–protein interactions are depicted by black dashed lines.
Flow chart of the cryo-EM data processing procedure. The gold-standard Fourier shell correlation curves (0.143 cut-off) show the final resolution of the holo-complex and each body.
a, Magnified view of the interaction surface between the LigIV DBD and Ku70 vWA domain coloured by sequence conservation. DNA-PKcs clashes with LigIV DBD when Ku is aligned between the LR and SR complex. b, Magnified view of the coiled-coil domain of XLF at its C-terminal tip coloured by sequence conservation. When models are not coloured by sequence conservation, the colour codes are the same as in Fig. 3. c, Superimposition of two asymmetric SR complexes after a 180° flip shown in front (top) and top (bottom) views. The XLF homodimer is used for aligning the two conformers. LigIV catalytic domains are hidden for clarity purposes. The transition from the apo state to the flipped state indicates potential conformational changes during the tandem ligation. Paths of DNA are also highlighted by dashed lines. d, Magnified view showing the relative positions of the two off-centred nicks between the two conformers. The two preferential nick positions are separated by approximately 4 bp. Consistently, dsDNA with 4-nt 3′ overhang, a major end-processing product of the NHEJ nuclease Artemis78, is reported to be a favoured substrate for NHEJ50. Our model suggests that dsDNA with a 4-nt 3′ overhang will experience minimum DNA translocation to accommodate the two ligation steps (Supplementary Video 3).
Extended Data Fig. 10 Both LR and SR synaptic complexes are able to perform double ligation during NHEJ in vitro.
a, Substrate design for the ligation assay. An internal Cy5 label is added to only the right half of the substrate to visualize the ligation products. A 4-nt 3′ complementary overhang has been introduced on both sides of the substrate. b, Denaturing gel analysis of end joining by the LR complex. The reaction comprised 100 nM of DNA, 200 nM of DNA-PKcs and Ku70/80, 500 nM of XLF and 70 nM of X4L4. The asterisk indicates an alternative secondary structure or impurity of the Cy5-labelled oligomer. The size of the DNA substrates and ligation products are labelled on the left (unit: bp). c, Denaturing gel analysis of end joining by the SR complex. The final factor concentrations are the same as in b. For gel source data, see Supplementary Fig. 1. Similar conditions for the gels have been replicated as biological replicates twice.
This file contains Supplementary Tables 1-2 and Supplementary Figures 1-7.
Density map and model fitting of the Long-range synaptic complex.
Density map and model fitting of the Short-range synaptic complex.
Model showing the conformational changes within the Short-range synaptic complex during the tandem ligation of two dsDNA nicks. DNA substrate with 4nt 3’ overhang is used as the example.
Model showing the conformational changes during the transition from the Long-range to the Short-range synaptic state.
About this article
Cite this article
Chen, S., Lee, L., Naila, T. et al. Structural basis of long-range to short-range synaptic transition in NHEJ. Nature 593, 294–298 (2021). https://doi.org/10.1038/s41586-021-03458-7
This article is cited by
Nature Structural & Molecular Biology (2023)
Nature Communications (2023)
Human Genetics (2023)
Nature Communications (2022)