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.

  • Letter
  • Published:

Architecture of an HIV-1 reverse transcriptase initiation complex

Abstract

Reverse transcription of the HIV-1 RNA genome into double-stranded DNA is a central step in viral infection1 and a common target of antiretroviral drugs2. The reaction is catalysed by viral reverse transcriptase (RT)3,4 that is packaged in an infectious virion with two copies of viral genomic RNA5 each bound to host lysine 3 transfer RNA (tRNALys3), which acts as a primer for initiation of reverse transcription6,7. Upon viral entry into cells, initiation is slow and non-processive compared to elongation8,9. Despite extensive efforts, the structural basis of RT function during initiation has remained a mystery. Here we use cryo-electron microscopy to determine a three-dimensional structure of an HIV-1 RT initiation complex. In our structure, RT is in an inactive polymerase conformation with open fingers and thumb and with the nucleic acid primer–template complex shifted away from the active site. The primer binding site (PBS) helix formed between tRNALys3 and HIV-1 RNA lies in the cleft of RT and is extended by additional pairing interactions. The 5′ end of the tRNA refolds and stacks on the PBS to create a long helical structure, while the remaining viral RNA forms two helical stems positioned above the RT active site, with a linker that connects these helices to the RNase H region of the PBS. Our results illustrate how RNA structure in the initiation complex alters RT conformation to decrease activity, highlighting a potential target for drug action.

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

Access options

Rent or buy this article

Prices vary by article type

from$1.95

to$39.95

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

Fig. 1: RTIC constructs and purification.
Fig. 2: Global architecture of the RTIC.
Fig. 3: Structure of the RTIC core.
Fig. 4: The +1 RTIC adopts an inactive conformation.

Similar content being viewed by others

References

  1. Gilboa, E., Mitra, S. W., Goff, S. & Baltimore, D. A detailed model of reverse transcription and tests of crucial aspects. Cell 18, 93–100 (1979).

    Article  CAS  Google Scholar 

  2. Sarafianos, S. G. et al. Structure and function of HIV-1 reverse transcriptase: molecular mechanisms of polymerization and inhibition. J. Mol. Biol. 385, 693–713 (2009).

    Article  CAS  Google Scholar 

  3. Baltimore, D. RNA-dependent DNA polymerase in virions of RNA tumour viruses. Nature 226, 1209–1211 (1970).

    Article  ADS  CAS  Google Scholar 

  4. Temin, H. M. & Mizutani, S. RNA-dependent DNA polymerase in virions of Rous sarcoma virus. Nature 226, 1211–1213 (1970).

    Article  ADS  CAS  Google Scholar 

  5. Paillart, J. C., Shehu-Xhilaga, M., Marquet, R. & Mak, J. Dimerization of retroviral RNA genomes: an inseparable pair. Nat. Rev. Microbiol. 2, 461–472 (2004).

    Article  CAS  Google Scholar 

  6. Huang, Y. et al. Incorporation of excess wild-type and mutant tRNA(3Lys) into human immunodeficiency virus type 1. J. Virol. 68, 7676–7683 (1994).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Ratner, L. et al. Complete nucleotide sequence of the AIDS virus, HTLV-III. Nature 313, 277–284 (1985).

    Article  ADS  CAS  Google Scholar 

  8. Lanchy, J. M., Ehresmann, C., Le Grice, S. F., Ehresmann, B. & Marquet, R. Binding and kinetic properties of HIV-1 reverse transcriptase markedly differ during initiation and elongation of reverse transcription. EMBO J. 15, 7178–7187 (1996).

    Article  CAS  Google Scholar 

  9. Lanchy, J. M. et al. Contacts between reverse transcriptase and the primer strand govern the transition from initiation to elongation of HIV-1 reverse transcription. J. Biol. Chem. 273, 24425–24432 (1998).

    Article  CAS  Google Scholar 

  10. Isel, C., Ehresmann, C. & Marquet, R. Initiation of HIV reverse transcription. Viruses 2, 213–243 (2010).

    Article  CAS  Google Scholar 

  11. Beerens, N. & Berkhout, B. The tRNA primer activation signal in the human immunodeficiency virus type 1 genome is important for initiation and processive elongation of reverse transcription. J. Virol. 76, 2329–2339 (2002).

    Article  CAS  Google Scholar 

  12. Beerens, N., Groot, F. & Berkhout, B. Initiation of HIV-1 reverse transcription is regulated by a primer activation signal. J. Biol. Chem. 276, 31247–31256 (2001).

    Article  CAS  Google Scholar 

  13. Goldschmidt, V., Ehresmann, C., Ehresmann, B. & Marquet, R. Does the HIV-1 primer activation signal interact with tRNA3 Lys during the initiation of reverse transcription? Nucleic Acids Res. 31, 850–859 (2003).

    Article  CAS  Google Scholar 

  14. Goldschmidt, V. et al. Structural variability of the initiation complex of HIV-1 reverse transcription. J. Biol. Chem. 279, 35923–35931 (2004).

    Article  CAS  Google Scholar 

  15. Goldschmidt, V. et al. Direct and indirect contributions of RNA secondary structure elements to the initiation of HIV-1 reverse transcription. J. Biol. Chem. 277, 43233–43242 (2002).

    Article  CAS  Google Scholar 

  16. Isel, C. et al. Structural basis for the specificity of the initiation of HIV-1 reverse transcription. EMBO J. 18, 1038–1048 (1999).

    Article  CAS  Google Scholar 

  17. Iwatani, Y., Rosen, A. E., Guo, J., Musier-Forsyth, K. & Levin, J. G. Efficient initiation of HIV-1 reverse transcription in vitro. Requirement for RNA sequences downstream of the primer binding site abrogated by nucleocapsid protein-dependent primer-template interactions. J. Biol. Chem. 278, 14185–14195 (2003).

    Article  CAS  Google Scholar 

  18. Liang, C. et al. The importance of the A-rich loop in human immunodeficiency virus type 1 reverse transcription and infectivity. J. Virol. 71, 5750–5757 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Liu, S., Harada, B. T., Miller, J. T., Le Grice, S. F. & Zhuang, X. Initiation complex dynamics direct the transitions between distinct phases of early HIV reverse transcription. Nat. Struct. Mol. Biol. 17, 1453–1460 (2010).

    Article  CAS  Google Scholar 

  20. Huang, H., Chopra, R., Verdine, G. L. & Harrison, S. C. Structure of a covalently trapped catalytic complex of HIV-1 reverse transcriptase: implications for drug resistance. Science 282, 1669–1675 (1998).

    Article  ADS  CAS  Google Scholar 

  21. Jacobo-Molina, A. et al. Crystal structure of human immunodeficiency virus type 1 reverse transcriptase complexed with double-stranded DNA at 3.0 Å resolution shows bent DNA. Proc. Natl Acad. Sci. USA 90, 6320–6324 (1993).

    Article  ADS  CAS  Google Scholar 

  22. Das, K., Martinez, S. E., Bauman, J. D. & Arnold, E. HIV-1 reverse transcriptase complex with DNA and nevirapine reveals non-nucleoside inhibition mechanism. Nat. Struct. Mol. Biol. 19, 253–259 (2012).

    Article  CAS  Google Scholar 

  23. Peisley, A. & Skiniotis, G. 2D projection analysis of GPCR complexes by negative stain electron microscopy. Methods Mol. Biol. 1335, 29–38 (2015).

    Article  Google Scholar 

  24. Das, R., Karanicolas, J. & Baker, D. Atomic accuracy in predicting and designing noncanonical RNA structure. Nat. Methods 7, 291–294 (2010).

    Article  CAS  Google Scholar 

  25. Watts, J. M. et al. Architecture and secondary structure of an entire HIV-1 RNA genome. Nature 460, 711–716 (2009).

    Article  ADS  CAS  Google Scholar 

  26. Foley, B. et al. HIV Sequence Compendium 2013 (Los Alamos National Laboratory, Los Alamos, 2013).

    Google Scholar 

  27. Coey, A., Larsen, K., Puglisi, J. D. & Viani Puglisi, E. Heterogeneous structures formed by conserved RNA sequences within the HIV reverse transcription initiation site. RNA 22, 1689–1698 (2016).

    Article  CAS  Google Scholar 

  28. Puglisi, E. V. & Puglisi, J. D. Secondary structure of the HIV reverse transcription initiation complex by NMR. J. Mol. Biol. 410, 863–874 (2011).

    Article  CAS  Google Scholar 

  29. Li, A., Gong, S. & Johnson, K. A. Rate-limiting pyrophosphate release by HIV reverse transcriptase improves fidelity. J. Biol. Chem. 291, 26554–26565 (2016).

    Article  CAS  Google Scholar 

  30. Beerens, N. et al. Role of the primer activation signal in tRNA annealing onto the HIV-1 genome studied by single-molecule FRET microscopy. RNA 19, 517–526 (2013).

    Article  CAS  Google Scholar 

  31. Zuker, M. Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res. 31, 3406–3415 (2003).

    Article  CAS  Google Scholar 

  32. Götte, M. et al. HIV-1 reverse transcriptase-associated RNase H cleaves RNA/RNA in arrested complexes: implications for the mechanism by which RNase H discriminates between RNA/RNA and RNA/DNA. EMBO J. 14, 833–841 (1995).

    Article  Google Scholar 

  33. Marshall, R. A., Dorywalska, M. & Puglisi, J. D. Irreversible chemical steps control intersubunit dynamics during translation. Proc. Natl Acad. Sci. USA 105, 15364–15369 (2008).

    Article  ADS  CAS  Google Scholar 

  34. Aitken, C. E., Marshall, R. A. & Puglisi, J. D. An oxygen scavenging system for improvement of dye stability in single-molecule fluorescence experiments. Biophys. J. 94, 1826–1835 (2008).

    Article  CAS  Google Scholar 

  35. Johansson, M., Chen, J., Tsai, A., Kornberg, G. & Puglisi, J. D. Sequence-dependent elongation dynamics on macrolide-bound ribosomes. Cell Rep. 7, 1534–1546 (2014).

    Article  CAS  Google Scholar 

  36. O’Leary, S. E., Petrov, A., Chen, J. & Puglisi, J. D. Dynamic recognition of the mRNA cap by Saccharomyces cerevisiae eIF4E. Structure 21, 2197–2207 (2013).

    Article  Google Scholar 

  37. Aitken, C. E. & Puglisi, J. D. Following the intersubunit conformation of the ribosome during translation in real time. Nat. Struct. Mol. Biol. 17, 793–800 (2010).

    Article  CAS  Google Scholar 

  38. Chen, J., Tsai, A., Petrov, A. & Puglisi, J. D. Nonfluorescent quenchers to correlate single-molecule conformational and compositional dynamics. J. Am. Chem. Soc. 134, 5734–5737 (2012).

    Article  CAS  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).

    Article  CAS  Google Scholar 

  40. Rohou, A. & Grigorieff, N. CTFFIND4: Fast and accurate defocus estimation from electron micrographs. J. Struct. Biol. 192, 216–221 (2015).

    Article  Google Scholar 

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

    Article  ADS  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  43. Scheres, S. H. Semi-automated selection of cryo-EM particles in RELION-1.3. J. Struct. Biol. 189, 114–122 (2015).

    Article  ADS  CAS  Google Scholar 

  44. Scheres, S. H. Processing of structurally heterogeneous cryo-EM data in RELION. Methods Enzymol. 579, 125–157 (2016).

    Article  CAS  Google Scholar 

  45. Penczek, P. A., Grassucci, R. A. & Frank, J. The ribosome at improved resolution: new techniques for merging and orientation refinement in 3D cryo-electron microscopy of biological particles. Ultramicroscopy 53, 251–270 (1994).

    Article  CAS  Google Scholar 

  46. Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).

    Article  CAS  Google Scholar 

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

    Article  Google Scholar 

  48. Wriggers, W. Conventions and workflows for using Situs. Acta Crystallogr. D 68, 344–351 (2012).

    Article  CAS  Google Scholar 

  49. Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D 66, 213–221 (2010).

    Google Scholar 

  50. Davis, I. W. et al. MolProbity: all-atom contacts and structure validation for proteins and nucleic acids. Nucleic Acids Res. 35, W375–W383 (2007).

    Article  ADS  Google Scholar 

  51. Lavender, C. A., Gorelick, R. J. & Weeks, K. M. Structure-based alignment and consensus secondary structures for three HIV-related RNA genomes. PLoS Comput. Biol. 11, e1004230 (2015).

    Article  ADS  Google Scholar 

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

    Article  CAS  Google Scholar 

  53. Isel, C., Ehresmann, C., Keith, G., Ehresmann, B. & Marquet, R. Initiation of reverse transcription of HIV-1: secondary structure of the HIV-1 RNA/tRNA(3Lys) (template/primer). J. Mol. Biol. 247, 236–250 (1995).

    Article  CAS  Google Scholar 

  54. Isel, C. et al. Specific initiation and switch to elongation of human immunodeficiency virus type 1 reverse transcription require the post-transcriptional modifications of primer tRNA3Lys. EMBO J. 15, 917–924 (1996).

    Article  CAS  Google Scholar 

Download references

Acknowledgements

We thank A. Frost and L. Stryer for suggesting beta-octyl glucoside as an additive for cryo-EM, R. Kornberg, M. Levitt, P. Geiduschek and W. Sundquist for reading the manuscript, M. Levitt for discussion of alternative tRNA folds and general support, D. Herschlag for discussions, and N. R. Latorraca for discussions and assistance with the Sherlock cluster. Supported by National Institutes of Health grant GM082545 to E.V.P., T32-GM008294 (Molecular Biophysics Training Program) to K.P.L., A.T.C. and K.K., National Science Foundation Graduate Research Fellowship Program (DGE-114747) to A.T.C and K.K., and Gabilan Stanford Graduate Fellowship to K.K. We thank Stanford University and the Stanford Research Computing Center for providing the Sherlock cluster resources. Additional calculations were performed on the Stanford BioX3 cluster, supported by NIH Shared Instrumentation Grant 1S10RR02664701.

Reviewer information

Nature thanks N. Sluis-Cremer and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Author information

Authors and Affiliations

Authors

Contributions

K.P.L., Y.K.M. and D.-H.C. acquired preliminary cryo-EM data and performed initial cryo-EM map calculations. Y.K.M. acquired cryo-EM data and obtained the 3D reconstructions shown in the main manuscript. K.P.L. acquired Mg2+ cryo-EM data and performed corresponding cryo-EM map calculations. A.T.C. purified the vRNA used for single-molecule experimentation and performed the single-molecule experiments. K.P.L., D.B. and L.M. performed all vRNA and RT sample preparations. K.P.L. performed all α-32P-dTTP incorporation assays. D.B. performed the RT activity assays. K.P.L. designed the purification scheme and purified the RTIC used in all experimentation. K.K. performed the vRNA–tRNA model building with input from K.P.L. K.P.L. and Y.K.M. performed final RTIC model building and refinement. K.P.L., Y.K.M, G.S., J.D.P. and E.V.P. interpreted the data. K.P.L and E.V.P wrote the manuscript with input from J.D.P., K.K., Y.K.M. and G.S.

Corresponding author

Correspondence to Elisabetta Viani Puglisi.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

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

Extended data figures and tables

Extended Data Fig. 1 Purification and activity of RTIC.

a, Initial anion-exchange purification of the RTIC away from free RT and vRNA–tRNA. This purification was repeated for each sample (>10) used in the manuscript, with only slight variations in the chromatogram. b, Polishing step using size-exclusion chromatography purification of the RTIC after anion exchange. This purification was repeated for each sample used in the manuscript (>10), with only slight variations in the chromatogram. c, A final 10% native TBE gel on the purified components. RT barely enters the gel under these running conditions. The RTIC runs as a single band, but trace amounts of free vRNA and/or vRNA–tRNA complex are sometimes present. This native gel is a representative result that was repeated independently for all purified RTIC samples used in the paper (>10). d, Autoradiograph image illustrating that the RTIC is capable of incorporating an incoming α-32P-dTTP nucleotide when extended and purified using dCTP instead of ddCTP. This gel is a representative result that was repeated independently for crosslinked and uncrosslinked samples (>6 independently prepared samples) used in dTTP incorporation assays. e, The RTIC incorporates α-32P-dTTP at roughly 89% efficiency compared to the free components after reaching a plateau. Values are mean ± s.d. (n = 3 independent experiments) with normalization to total incorporation of free RT + vRNA–tRNA reactions. f, Autoradiograph image showing that the incorporation of dTTP is inhibited in the presence of nevirapine (NNRTI). Images have been adjusted to allow identification of the NNRTI-inhibited band. This gel is a representative result that was repeated independently for crosslinked and uncrosslinked samples (3 samples each). g, Relative activities, judged by primer usage, of wild-type, Q258C, and Q258C/E478Q reverse transcriptase mutants used in this study. Values are mean ± s.d. (n = 3 independent experiments) with normalization to the primer usage of wild-type RT. h, RTIC (triangles), RTIC with NNRTI (circles) or vRNA–tRNA + excess RT (squares) reactions were initiated by addition of α-32P-dTTP and quenched at different time points. Data were fit using the relationship for the free vRNA–tRNA + RT reaction: \({\rm{Intensity}}=A\left(1-{e}^{-{k}_{{\rm{pol}}}t}\right)+B\left(1-{e}^{-{k}_{{\rm{slow}}}t}\right)\). Data were fit using the relationship for the RTIC (with or without NNRTI) reaction: \({\rm{Intensity}}=B\left(1-{e}^{-{k}_{{\rm{slow}}}t}\right)\) where A and B represent the amplitude of the fast and slow processes, respectively, kpol is the apparent extension rate constant, and kslow is the rate of the slow process. The second relationship was used for the RTIC data, as the slow process appears to dominate incorporation when the vRNA–tRNA substrate is crosslinked to RT. The best fits were obtained with: A = 0.7166 AU, kpol = 0.1078 s−1, B = 0.2754, kslow = 0.01002 s−1 for the vRNA–tRNA + excess RT; B = 0.9808, kslow = 0.003140 s−1 for the RTIC; and B = 1.095, kslow = 0.0001714 s−1 for the RTIC with NNRTI. kslow is about 3.19 times slower for crosslinked RTIC than for un-crosslinked components. Assays were independently repeated three times to ensure reproducibility.

Extended Data Fig. 2 Representative negative-stain EM images, cryo-EM images, and 2D averages of the RTIC.

a, Representative negative-stain EM image of HIV RTIC reveals a mono-disperse sample that is free of aggregates. Approximately a dozen images were taken of each sample before cryo-EM grid preparation to ensure sample quality. b, Cryo-EM image of RTIC without β-OG. The long chains correspond to RNA from the complex with very few particles resembling the protein. Results are reproducible in the absence of β-OG (>10 samples tested). c, Cryo-EM image of RTIC with β-OG. Single particles corresponding to the complex appear similar to the negative-stain visualization. All 5,107 images used in both cryo-EM datasets have a similar appearance with slight differences in particle density. d, Representative 2D averages of RTIC complex from the cryo-EM data collected with β-OG. Both datasets exhibit very similar 2D classes.

Extended Data Fig. 3 Data processing workflow for RTIC complex.

a, Data processing workflow for the 8.0 Å global and 4.5 Å core maps. b, Gold standard FSC curve of RTIC core and global maps. c, The final 4.5 Å map is coloured according to local resolution estimated by Relion. d, Angular distribution of particle projections. The length of each projection direction is proportional to the number of assigned particles. e, Data processing workflow for the 8.2 Å global Mg2+ map. f, Gold standard FSC curve of RTIC Mg2+ global map.

Extended Data Fig. 4 Quality of the cryo-EM density for the core RTIC map.

a, View of HIV-1 RT from the front. The subdomains of RT are coloured. Underneath the main RTIC view, each subdomain of RT, plus the p51 subunit, is shown fit into the 4.5 Å map. b, View of HIV-1 RT from the polymerase active site side. The subdomains of RT are coloured. Underneath the main RTIC view, each subdomain of RT, plus the p51 subunit, is shown fit into the 4.5 Å map. In a, b, regions of protein, namely loops and linkers, that lacked sufficient density were removed after comparison with previously published structures of RT. These regions are indicated by dotted lines and are most commonly found in the finger and palm subdomains. c, Representative regions of 4.5 Å map fitted with protein secondary structure that display densities for side chains. A view of the PBS helix fit into the 4.5 Å map is also shown; phosphates of the RNA backbone are partially resolved. Regions are coloured with respect to the main text models.

Extended Data Fig. 5 Mg2+ global map views and structure comparison.

a, Side and top views of the 8.2 Å global map at different density thresholds. The orientation of the peripheral vRNA and tRNA elements is within the variability seen among the different RTIC conformers. b, A model of the RTIC built into the Mg2+ density using the main text global RTIC model. vRNA and tRNA helices were treated as rigid bodies derived from main text model (see Extended Data Fig. 6 and Methods). c, Comparison of the global RTIC model RNA (grey) with the Mg2+ model RNA (coloured). All three regions of RNA structure (H1, H2, and tRNA) differ in the Mg2+ model, but are adequately described by rigid body movements of the RNA helical elements taken from the global RTIC model. Both H1 and H2 represent a substantial structural barrier to initiation. d, Partial accommodation of H1 into high monovalent salt classes 3, 4 and 7.

Extended Data Fig. 6 Low-resolution tRNA density and fold comparison.

a, Top and side views of the elongated helical tRNA density observed in the low-resolution global map of the RTIC. b, Top and side views of the vRNA–tRNA model generated using the hypothesized elongated tRNA helical fold. The tRNA model fits the long helical density well. Corresponding secondary structure is in d. c, Top and side views of the vRNA–tRNA model generated using previously hypothesized tRNA secondary structures that have the anticodon and D-stem loops independently folded. Corresponding secondary structure is in e. d, Secondary structure depiction of the new vRNA–tRNA and canonical clover-leaf fold of the tRNA. The different domains are coloured and correspond with the models in panels b and c. e, Secondary-structure depiction of the old vRNA–tRNA fold with independent anticodon and D-stem loops. The domains are coloured and correspond with the model in c and clover-leaf fold of the tRNA in d.

Extended Data Fig. 7 Peripheral RNA heterogeneity of the RTIC conformers.

a, Tiled views of eight conformations emerging from 3D classification of RTIC. Each class is numbered and class 7 was used for the global RTIC reconstruction. b, Superposition of the eight classes from a. The main areas of RNA heterogeneity are focused on the orientations of vRNA H2, H1 and the connection loop, and the tRNA. With no stabilizing protein contacts, vRNA H2, H1, and the tRNA sample a wide range of conformations, limiting the resolution of the global map. c, Additional RTIC models built into classes 3 (tan) and 4 (blue). The models for the tRNA, vRNA H1, and vRNA H2 were all derived from the global RTIC model and treated as rigid bodies for model building. The connecting loop was not built in these models as the density for this region was not clear in these maps, though there is reasonable density to model a loop near H1. Junctions between the helices serve as hinges that allow movement of the independent domains. The main text global RTIC model (grey) is included as a comparison. d, The vRNA and tRNA helices treated as rigid bodies for modelling are shown in bold. Hinge points for each helix are highlighted with grey circles and serve as points of flexibility for the RTIC.

Extended Data Fig. 8 Single-molecule experimentation and analysis.

a, Secondary structure depiction of the vRNA–tRNA construct used for single-molecule experiments. The labelling scheme is shown, with the Cy3 dye located on the 5′ end of the vRNA helix 1 and Cy5 dye located on an oligonucleotide positioned near the 5′ end of helix 1. The vRNA–tRNA complex was crosslinked to RT for the experiments. b, Ninety-five per cent of the RTIC complexes are in the high FRET, helix 1 formation, state (480 traces analysed, see Methods). c, Example trace of the ones used for final FRET analysis. The high FRET state of the RTIC complex, which is attributed to helix 1 formation. Photobleaching events for both Cy5 and Cy3 are indicated. d, Examples of traces removed from final FRET analysis. Traces exhibit the presence of multiple molecules (multiple single-dye photobleaching events) or poor dye behaviour (blinking and quenching).

Extended Data Fig. 9 Comparison with NNRTI bound and active RT–nucleic acid complexes in the cryo-EM map.

All alignments between structures and the RTIC were done using the p51 subunit. a, Comparison of an active conformation RT–nucleic acid structure (pink, 1RTD) with the RTIC core (RT, purple; tRNA primer, red; vRNA template, yellow). The EM map overlay shows the poor fit of the 1RTD model in the fingers, thumb, and primer grip of RT. Deviations of the nucleic acid primer and template of 1RTD away from the RTIC density are also apparent. b, Comparison of an NNRTI-bound RT–nucleic acid structure (dark grey, 3V81) with the RTIC core. The EM map overlay shows the closer fit of the fingers and primer grip regions of RT in the 3V81 model. The thumb region also overlays well, but with slight deviations. Most noticeably, the nucleic acid primer/template in the 3V81 model deviates, although not as dramatically as in 1RTD, from the RTIC core EM density.

Extended Data Table 1 Cryo-EM data collection, refinement, and validation statistics

Supplementary information

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Larsen, K.P., Mathiharan, Y.K., Kappel, K. et al. Architecture of an HIV-1 reverse transcriptase initiation complex. Nature 557, 118–122 (2018). https://doi.org/10.1038/s41586-018-0055-9

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41586-018-0055-9

This article is cited by

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.

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