Structure of a P element transposase–DNA complex reveals unusual DNA structures and GTP-DNA contacts

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Abstract

P element transposase catalyzes the mobility of P element DNA transposons within the Drosophila genome. P element transposase exhibits several unique properties, including the requirement for a guanosine triphosphate cofactor and the generation of long staggered DNA breaks during transposition. To gain insights into these features, we determined the atomic structure of the Drosophila P element transposase strand transfer complex using cryo-EM. The structure of this post-transposition nucleoprotein complex reveals that the terminal single-stranded transposon DNA adopts unusual A-form and distorted B-form helical geometries that are stabilized by extensive protein-DNA interactions. Additionally, we infer that the bound guanosine triphosphate cofactor interacts with the terminal base of the transposon DNA, apparently to position the P element DNA for catalysis. Our structure provides the first view of the P element transposase superfamily, offers new insights into P element transposition and implies a transposition pathway fundamentally distinct from other cut-and-paste DNA transposases.

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Fig. 1: Reconstituted strand transfer complex represents the active form of TNP.
Fig. 2: Structure of the Drosophila P element STC.
Fig. 3: dDNA adopts a non-canonical geometry within the STC.
Fig. 4: Each subunit makes extensive contacts with a single dDNA.
Fig. 5: The tDNA is severely bent at AT-rich sites.
Fig. 6: The unsymmetrized reconstruction suggests a mechanism for 5′ and 3′ P element end pairing.

Data availability

Atomic models are available through the Protein Data Bank with accessions codes 6P5A (C2) and 6PE2 (C1); cryo-EM reconstructions are available through the EMDB with accession codes EMD-20254 (C2) and EMD-20321 (C1).

References

  1. 1.

    Kidwell, M. G. Horizontal transfer of P-elements and other short inverted repeat transposons. Genetica 86, 275–286 (1992).

  2. 2.

    Engels, W. R. P elements in Drosophila. Curr. Top. Microbiol. Immunol. 204, 103–123 (1996).

  3. 3.

    MajumdarS. & RioD. C. P transposable elements in Drosophila and other eukaryotic organisms. Microbiol. Spectr. 3, MDNA3-0004-2014 (2015).

  4. 4.

    Sekelsky, J. DNA repair in Drosophila: mutagens, models and missing genes. Genetics 205, 471–490 (2017).

  5. 5.

    Khurana, J. S. et al. Adaptation to P element transposon invasion in Drosophila melanogaster. Cell 147, 1551–1563 (2011).

  6. 6.

    Teixeira, F. K. et al. piRNA-mediated regulation of transposon alternative splicing in the soma and germ line. Nature 552, 268–272 (2017).

  7. 7.

    Laski, F. A., Rio, D. C. & Rubin, G. M. Tissue specificity of Drosophila P element transposition is regulated at the level of mRNA splicing. Cell 44, 7–19 (1986).

  8. 8.

    Siebel, C. W., Fresco, L. D. & Rio, D. C. The mechanism of somatic inhibition of Drosophila P-element pre-mRNA splicing: multiprotein complexes at an exon pseudo-5′ splice site control U1 snRNP binding. Genes Dev. 6, 1386–1401 (1992).

  9. 9.

    Roussigne, M. et al. The THAP domain: a novel protein motif with similarity to the DNA-binding domain of P element transposase. Trends Biochem. Sci. 28, 66–69 (2003).

  10. 10.

    Majumdar, S., Singh, A. & Rio, D. C. The human THAP9 gene encodes an active P-element DNA transposase. Science 339, 446–448 (2013).

  11. 11.

    Quesneville, H., Nouaud, D. & Anxolabehere, D. Recurrent recruitment of the THAP DNA-binding domain and molecular domestication of the P-transposable element. Mol. Biol. Evol. 22, 741–746 (2005).

  12. 12.

    Hammer, S. E. Homologs of Drosophila P transposons were mobile in zebrafish but have been domesticated in a common ancestor of chicken and human. Mol. Biol. Evol. 22, 833–844 (2005).

  13. 13.

    O’Hare, K. & Rubin, G. M. Structures of P transposable elements and their sites of insertion and excision in the Drosophila melanogaster genome. Cell 34, 25–35 (1983).

  14. 14.

    Mullins, M. C., Rio, D. C. & Rubin, G. M. cis-acting DNA sequence requirements for P-element transposition. Genes Dev. 3, 729–738 (1989).

  15. 15.

    Kaufman, P. D., Doll, R. F. & Rio, D. C. Drosophila P element transposase recognizes internal P element DNA sequences. Cell 59, 359–371 (1989).

  16. 16.

    Rio, D. C., Laski, F. A. & Rubin, G. M. Identification and immunochemical analysis of biologically active Drosophila P element transposase. Cell 44, 21–32 (1986).

  17. 17.

    Tang, M., Cecconi, C., Kim, H., Bustamante, C. & Rio, D. C. Guanosine triphosphate acts as a cofactor to promote assembly of initial P-element transposase–DNA synaptic complexes. Genes Dev. 19, 1422–1425 (2005).

  18. 18.

    Tang, M., Cecconi, C., Bustamante, C. & Rio, D. C. Analysis of P element transposase protein-DNA interactions during the early stages of transposition. J. Biol. Chem. 282, 29002–29012 (2007).

  19. 19.

    Beall, E. L. & Rio, D. C. Drosophila P-element transposase is a novel site-specific endonuclease. Genes Dev. 11, 2137–2151 (1997).

  20. 20.

    Linheiro, R. S. & Bergman, C. M. Testing the palindromic target site model for DNA transposon insertion using the Drosophila melanogaster P-element. Nucleic Acids Res. 36, 6199–6208 (2008).

  21. 21.

    Kaufman, P. D. & Rio, D. C. P element transposition in vitro proceeds by a cut-and-paste mechanism and uses GTP as a cofactor. Cell 69, 27–39 (1992).

  22. 22.

    Roiha, H., Rubin, G. M. & O’Hare, K. P element insertions and rearrangements at the singed locus of Drosophila melanogaster. Genetics 119, 75–83 (1988).

  23. 23.

    Hawley, R. S. et al. Molecular analysis of an unstable P element insertion at the singed locus of Drosophila melanogaster: evidence for intracistronic transposition of a P element. Genetics 119, 85–94 (1988).

  24. 24.

    Yin, Z., Lapkouski, M., Yang, W. & Craigie, R. Assembly of prototype foamy virus strand transfer complexes on product DNA bypassing catalysis of integration. Protein Sci. 21, 1849–1857 (2012).

  25. 25.

    Yin, Z. et al. Crystal structure of the Rous sarcoma virus intasome. Nature 530, 362–366 (2016).

  26. 26.

    Ballandras-Colas, A. et al. A supramolecular assembly mediates lentiviral DNA integration. Science 355, 93–95 (2017).

  27. 27.

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

  28. 28.

    Chow, S. A., Vincent, K. A., Ellison, V. & Brown, P. O. Reversal of integration and DNA splicing mediated by integrase of human-immunodeficiency-virus. Science 255, 723–726 (1992).

  29. 29.

    Melek, M. & Gellert, M. RAG1/2-mediated resolution of transposition intermediates: two pathways and possible consequences. Cell 101, 625–633 (2000).

  30. 30.

    Au, T. K., Pathania, S. & Harshey, R. M. True reversal of Mu integration. EMBO J. 23, 3408–3420 (2004).

  31. 31.

    Polard, P. et al. IS911-mediated transpositional recombination in vitro. J. Mol. Biol. 264, 68–81 (1996).

  32. 32.

    Jonsson, C. B., Donzella, G. A. & Roth, M. J. Characterization of the forward and reverse integration reactions of the Moloney murine leukemia virus integrase protein purified from Escherichia coli. J. Biol. Chem. 268, 1462–1469 (1993).

  33. 33.

    Beall, E. L. & Rio, D. C. Transposase makes critical contacts with, and is stimulated by, single‐stranded DNA at the P element termini in vitro. EMBO J. 17, 2122–2136 (1998).

  34. 34.

    Donzella, G. A., Jonsson, C. B. & Roth, M. J. Coordinated disintegration reactions mediated by Moloney murine leukemia virus integrase. J. Virol. 70, 3909–3921 (1996).

  35. 35.

    Roussigne, M., Cayrol, C., Clouaire, T., Amalric, F. & Girard, J.-P. THAP1 is a nuclear proapoptotic factor that links prostate-apoptosis-response-4 (Par-4) to PML nuclear bodies. Oncogene 22, 2432–2442 (2003).

  36. 36.

    Sabogal, A., Lyubimov, A. Y., Corn, J. E., Berger, J. M. & Rio, D. C. THAP proteins target specific DNA sites through bipartite recognition of adjacent major and minor grooves. Nat. Struct. Mol. Biol. 17, 117–U145 (2010).

  37. 37.

    Lee, C. C., Mul, Y. M. & Rio, D. C. The Drosophila P-element KP repressor protein dimerizes and interacts with multiple sites on P-element DNA. Mol. Cell. Biol. 16, 5616–5622 (1996).

  38. 38.

    Lee, C. C., Beall, E. L. & Rio, D. C. DNA binding by the KP repressor protein inhibits P-element transposase activity in vitro. EMBO J. 17, 4166–4174 (1998).

  39. 39.

    Dunker, A. K. et al. Intrinsically disordered protein. J. Mol. Graph. Model. 19, 26–59 (2001).

  40. 40.

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

  41. 41.

    MorrisE. R., GreyH., McKenzieG., JonesA. C. & RichardsonJ. M. A bend, flip and trap mechanism for transposon integration. eLife 5, e15537 (2016).

  42. 42.

    Maertens, G. N., Hare, S. & Cherepanov, P. The mechanism of retroviral integration from X-ray structures of its key intermediates. Nature 468, 326–329 (2010).

  43. 43.

    Hickman, A. B., Chandler, M. & Dyda, F. Integrating prokaryotes and eukaryotes: DNA transposases in light of structure. Crit. Rev. Biochem. Mol. Biol. 45, 50–69 (2010).

  44. 44.

    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).

  45. 45.

    Beall, E. L. & Rio, D. C. Drosophila IRBP/Ku p70 corresponds to the mutagen-sensitive mus309 gene and is involved in P-element excision in vivo. Genes Dev. 10, 921–933 (1996).

  46. 46.

    Fuller, J. R. & Rice, P. A. Target DNA bending by the Mu transpososome promotes careful transposition and prevents its reversal. eLife 6, 257 (2017).

  47. 47.

    Wright, A. V. et al. Structures of the CRISPR genome integration complex. Science 357, 1113–1118 (2017).

  48. 48.

    Rodgers, K. K. Riches in RAGs: revealing the V(D)J recombinase through high-resolution structures. Trends Biochem. Sci. 42, 72–84 (2017).

  49. 49.

    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).

  50. 50.

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

  51. 51.

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

  52. 52.

    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).

  53. 53.

    Chuong, E. B., Elde, N. C. & Feschotte, C. Regulatory activities of transposable elements: from conflicts to benefits. Nat. Rev. Genet. 18, 71–86 (2017).

  54. 54.

    Sano, K.-I., Maeda, K., Oki, M. & Maéda, Y. Enhancement of protein expression in insect cells by a lobster tropomyosin cDNA leader sequence. FEBS Lett. 532, 143–146 (2002).

  55. 55.

    Trowitzsch, S., Bieniossek, C., Nie, Y., Garzoni, F. & Berger, I. New baculovirus expression tools for recombinant protein complex production. J. Struct. Biol. 172, 45–54 (2010).

  56. 56.

    Ballandras-Colas, A. et al. Cryo-EM reveals a novel octameric integrase structure for betaretroviral intasome function. Nature 530, 358–361 (2016).

  57. 57.

    Mastronarde, D. N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol. 152, 36–51 (2005).

  58. 58.

    Tan, Y. Z. et al. Addressing preferred specimen orientation in single-particle cryo-EM through tilting. Nat. Methods 14, 793–796 (2017).

  59. 59.

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

  60. 60.

    Zhang, K. Gctf: real-time CTF determination and correction. J. Struct. Biol. 193, 1–12 (2016).

  61. 61.

    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).

  62. 62.

    Zivanov, J. et al. New tools for automated high-resolution cryo-EM structure determination in RELION-3. eLife 7, 163 (2018).

  63. 63.

    Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D 60, 2126–2132 (2004).

  64. 64.

    Frenz, B., Walls, A. C., Egelman, E. H., Veesler, D. & DiMaio, F. RosettaES: a sampling strategy enabling automated interpretation of difficult cryo-EM maps. Nat. Methods 14, 797–800 (2017).

  65. 65.

    Adams, P. D. et al. The Phenix software for automated determination of macromolecular structures. Methods 55, 94–106 (2011).

  66. 66.

    Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).

  67. 67.

    Goddard, T. D. et al. UCSF ChimeraX: meeting modern challenges in visualization and analysis. Protein Sci. 27, 14–25 (2018).

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Acknowledgements

We thank the Rio Lab members for help and advice. We are grateful to P. Grob, E. Montabana and D. Toso for help with cryo-EM data acquisition and for general microscope maintenance. We thank A. Chintangal for computational support. We are grateful to A. Ban and A. Zanghellini (Arzeda Corporation) for the gift of the codon-optimized P element gene. We thank F. Dimaio and O. Sobolev for advice on modeling with RosettaES and PHENIX, respectively. We thank J. Berger (JHUMS) for examining our DNA and protein modeling and for advice. We thank K. Collins, J. Berger, T.H.G. Nguyen and Y. Lee for critical reading of the manuscript. Work in the Rio Lab was supported by NIH grant R35GM118121. E.H.K. was supported by NIH grant no. K99GM124463. E.N. is an Investigator of the Howard Hughes Medical Institute.

Author information

D.C.R. and E.N. supervised the study. G.G. developed the transposase expression conditions and purification procedure, performed in vitro biochemical assays and in vivo assays and prepared complexes for imaging. E.H.K. performed negative-stain and cryo-EM specimen preparation, data collection and data processing. E.H.K. interpreted the protein density with feedback from E.N., G.G. and D.C.R. G.G. and E.H.K. built the DNA model into the map. E.H.K. built the protein model into the map and refined the structure. G.G. and D.C.R. wrote the initial manuscript. All authors contributed to the final manuscript.

Correspondence to Elizabeth H. Kellogg or Donald C. Rio.

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

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Integrated supplementary information

Supplementary Figure 1 Assembly of the CDC, authentic STC, and STC bound to stDNA.

a, Diagram of the cleaved donor complex (CDC) assembly pathway. (TIR, terminal inverted repeat). b, Diagram of the authentic strand transfer complex (STC) assembly pathway. CDCs were assembled as in Supplementary Fig. 1a, then provided with an idealized hotspot target DNA from the Drosophila singed locus (tDNA). (GTP, guanosine triphosphate; TSM, target sequence motif). c, Diagram of the strand transfer complexes assembled on strand transfer product DNA (stDNA). d, Representative Coomassie-stained SDS-PAGE of the overnight dialysis and gel filtration fractions. ‘o/n dialysis’ and ‘spin’ lanes were diluted 1:10 before loading. (HMS*, 6xHis-Maltose binding protein-SUMO* tandem solubility tag). e, Negative stain electron micrographs of authentic STC and stDNA bound STC.

Supplementary Figure 2 Image processing of tilted dataset leading to a 3.6 Å resolution cryo-EM reconstruction.

a, Representative cryo-EM image collected with a 40° tilt showing well-defined, monodispersed particles (scale bar represents 100 nm). b, Angular distribution of particles from the tilted dataset is cone-like, corresponding to a majority of top-views. c, A single, well-defined reconstruction was produced using cryoSPARC and subsequently refined to high resolution using RELION-3.0 (see Methods for details). d, Reference-free 2D classes of the tilted data reveal secondary structure features. e, The secondary structure features are consistent with the estimated resolution of the map, with well-defined secondary structure and distinctive densities for large side-chain. f, The overall resolution (based on the Fourier shell correlation (FSC) 0.143 criterion) for the symmetrized reconstruction is 3.5 Å (3.6 Å if using randomized phases), and 3.9 Å for the unsymmetrized reconstruction. The map versus model resolution is 3.7 and 4 Å, respectively, for the symmetrized and unsymmetrized maps. g, 3D map for the C2 (symmetrized, left) and C1 (unsymmetrized, right) reconstructions colored by local resolution showing the core of the structure to be around 3.5 Å. To show some of the most disordered regions, the C1 map is shown low-pass filtered to both 4 Å and 6 Å.

Supplementary Figure 3 The STC is dimeric and contains disordered regions.

a, PONDR scores of predicted disordered regions. Disordered regions predicted with high confidence (indicated by black bars), are within the leucine zipper dimerization domain and the RNase H – CTD linker region. Contrary to the prediction results, we observe the dimerization domain to be largely ordered in the C1 reconstruction, in spite of the prediction suggesting that this region may undergo a disorder-to-order transition upon dimerization (Dunker, A. K. et al., J. Mol. Graph. Model. 19, 26–59, 2001). b, The disordered linker spanning the RNase H and CTD domains is represented by a red dashed line, with density for ordered regions colored by domain as in Fig. 2c. c, STC subunit organization. Densities are shown as in Fig. 2c but are colored by subunit (blue and green). Donor DNAs are colored in red, and target DNA in purple. The density corresponding to GTP is indicated in yellow.

Supplementary Figure 4 A comparison of RNase H insertion domains among different transposases and RNase H catalytic mutants are inactive.

a, Architectures of insertion domains found in other DNA transposases. The RNase H domains (grey) of other structurally characterized DNA transposases (or the transposase-related RAG1 protein) were aligned by their respective catalytic residues (indicated in red) and ordered by increasing insertion domain size (blue). Insertion domain sizes (indicated below) were determined by approximate start and end insertion positions. The PDB numbers from which these structures were derived are in parentheses. b, Structural alignment of the P element transposase insertion domain, the Hermes insertion domain (1DWY) and the RAG1–RAG2 insertion domain (6CIK) reveals structural similarities at the fold level. c, Bar graph of relative in vivo P element excision activity of alanine-substituted catalytic mutants (D230, D303 and E531). Cell-based excision assays were performed as previously described (Rio, D. C. et al., Cell. 44, 21–32, 1986)(Beall, E. L. et al., Genes Dev. 10, 921–933, 1996). Single alanine mutants were generated by site-directed mutagenesis of pPBSKS (+) pAc-TNP and verified by sequencing over the entire coding sequence. The assay was conducted in triplicate (n = 3). Error bars indicate standard deviations. (WT, wild type). d, Representative immunoblot of wild type transposase and catalytic mutant protein expression levels. Cells were harvested 24hr after transfection and lysates were normalized to cell number. Membrane was cut and then immunoblotted with anti-transposase antibodies (α-TNP) or a loading control (α-HRP48).

Supplementary Figure 5 Characteristics of A-form DNA are well resolved and base pairing between distant donor DNA regions is required for strand transfer activity.

a, Ideal A-form DNA fitted into donor DNA reconstruction, depicting widened minor groove, base pair tilt, axial rise and helical axis dislocation relative to base pairs. A single donor DNA is depicted for clarity. The reconstruction is colored green and yellow, for transferred strand and non-transferred strand, respectively. Relevant regions of DNA are indicated. b, Atomic model of donor DNA depicting A-form DNA characteristics. Views are as in a, except only relevant regions the donor DNA atomic model are depicted. c, Schematic of the secondary structure of a donor DNA terminal inverted repeat (left). Watson-Crick base pairing is indicated by solid lines. Non-canonical base pairing is indicated by dots, or dotted lines. Nucleotides of the transferred strand are numbered -1 to -31, starting at the 3′ terminal guanosine. Distant noncanonical A-form helical base pairing between the transferred and non-transferred strand is highlighted (dashed red box). Agarose gel of a strand transfer assay with 5′-radiolabeled mutant and rescue donor DNAs (right). Assays were largely performed as previously described (Beall, E. L., et al., EMBO J. 17, 2122–2136, 1998). The base pairs are shown above each lane, with the substituted bases highlighted in red (mutant, lanes 2 - 8). Compensatory substitutions in the non-transferred strand are shown above each lane, with substitutions to restore base pairing highlighted in blue (rescue, lanes 10 - 16). The expected positions of single-ended integration (SET) and double-ended integration (DET), as well as free donor DNA (Probe), are indicated. wt, wild type donor DNA.

Supplementary Figure 6 A single transposase subunit engages both P element donor DNAs.

Left, the RNase H domain of one subunit of transposase is catalytically engaged with one P element donor DNA (red DNA). The domains are colored as in Fig. 2c. For clarity, the dimerization domain, the GBD, and the other transposase subunit are not shown. Catalytic residues are depicted in red. Right, a 90° rotated view shows the same subunit contacting the other P element donor DNA (blue DNA), through the HTH domain, a long loop in the RNase H domain, and through the CTD. This mode of engagement likely acts as a regulatory step to ensure proper assembly with both P element ends before proceeding to catalysis.

Supplementary Figure 7 TNP nucleotide density is consistent with GTP and distinguishable from GDP within the reported resolution regime.

The GTP density in our cryo-EM map (top-left) is consistent with the GTP density observed the cryo-EM reconstruction of the U4–U6.U5 tri-snRNP Snu114 (3.7Å, top-right) and the non-hydrolyzable GTP analog (GMPPCP) of dynamin (3.7Å, bottom-left) and inconsistent with GDP in the β-tubulin subunit (3.5 Å, bottom-right), for which GTP is hydrolyzed during microtubule assembly.

Supplementary Figure 8 Target DNA binds in a positively charged groove.

a, Plot of target DNA minor groove width. The minor groove width was calculated from the target DNA model using the 3DNA webserver (Li, S., et al., Nucleic Acids Research. 47, W26–W34, 2019), with a 2 bp sliding window, accounting for phosphate van der Waals radii. The target DNA sequence is depicted on the x-axis and colored as in Fig. 5a. Red dots indicate transposition sites, on either the top or bottom strand of the target DNA. b, Electrostatic surface potential of the STC as viewed from below the target DNA binding site. Calculations were performed in UCSF Chimera (Pettersen, E. F., et al., J Comput. Chem. 25, 1605–1612, 2004). Blue denotes a positive charge and red denotes a negative charge. Target DNA is shown as in Fig. 5a. c, Schematic representation of observed base-specific and backbone contacts between transposase and the target DNA. Target DNA (purple border) is numbered as in Fig. 5e (target site duplication, pink fill; AT-rich flanks, green fill). Residue numbers are indicated and outlined in a solid or dashed border to indicate transposase subunit A, or transposase subunit B, respectively. Residues are colored according to domain (RNase H, orange; GBD, blue). Direct contacts are shown as solid lines; aromatic base stacking interactions are shown as dashed lines; major groove, minor groove and main chain contacts are indicated; interacting phosphates are highlighted yellow. d, Unmodeled density at the C-terminus is oriented towards the target DNA. The map is low-pass filtered to 4 Å to more clearly show the presence of additional, poorly-ordered density. While we could not confidently build into the density beyond position 734, the highly basic nature and positioning of the weak density near DNA suggests that this region likely plays a role in target DNA binding. Consistent with this, C-terminal tags on transposase decrease the overall excision and strand transfer activity (unpublished results, D. Rio).

Supplementary information

Supplementary Information

Supplementary Figs. 1–8 and Supplementary Tables 1 and 2.

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Supplementary Data Set 1

Source images for all data obtained by gel electrophoresis indicated in the figures.

Supplementary Video 1

Rotating view of the P element transposase strand transfer complex. Rotating view of the cryo-EM reconstruction and atomic model for the Drosophila P element transposase strand transfer complex.

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Ghanim, G.E., Kellogg, E.H., Nogales, E. et al. Structure of a P element transposase–DNA complex reveals unusual DNA structures and GTP-DNA contacts. Nat Struct Mol Biol 26, 1013–1022 (2019) doi:10.1038/s41594-019-0319-6

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