Structural insights into influenza A virus ribonucleoproteins reveal a processive helical track as transcription mechanism

Abstract

The influenza virus genome consists of eight viral ribonucleoproteins (vRNPs), each consisting of a copy of the polymerase, one of the genomic RNA segments and multiple copies of the nucleoprotein arranged in a double helical conformation. vRNPs are macromolecular machines responsible for messenger RNA synthesis and genome replication, that is, the formation of progeny vRNPs. Here, we describe the structural basis of the transcription process. The mechanism, which we call the ‘processive helical track’, is based on the extreme flexibility of the helical part of the vRNP that permits a sliding movement between both antiparallel nucleoprotein-RNA strands, thereby allowing the polymerase to move over the genome while bound to both RNA ends. Accordingly, we demonstrate that blocking this movement leads to inhibition of vRNP transcriptional activity. This mechanism also reveals a critical role of the nucleoprotein in maintaining the double helical structure throughout the copying process to make the RNA template accessible to the polymerase.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Fig. 1: 3D reconstructions of different conformations of the helical part of vRNPs.
Fig. 2: Local resolution map of vRNPs.
Fig. 3: Effect of nucleozin on vRNPs structure.
Fig. 4: Decreased in vitro transcriptional activity of vRNPs pre-incubated with nucleozin at different concentrations.
Fig. 5: Images of the transcription process.
Fig. 6: The transcription process mechanism steps.

Data availability

The EM maps have been deposited with the Electron Microscopy Data Bank (https://www.ebi.ac.uk/pdbe/emdb) under accession numbers EMD-0175, EMD-4412, EMD-4423, EMD-4426 and EMD-4430. The atomic coordinates of the docking have been deposited with the Protein Data Bank (https://www.rcsb.org/) under accession numbers 6H9G, 6I54, 6I7B, 6I7M and 6I85.

References

  1. 1.

    Ortega, J. et al. Ultrastructural and functional analyses of recombinant influenza virus ribonucleoproteins suggest dimerization of nucleoprotein during virus amplification. J. Virol. 74, 156–163 (2000).

  2. 2.

    Ye, Q., Krug, R. M. & Tao, Y. J. The mechanism by which influenza A virus nucleoprotein forms oligomers and binds RNA. Nature 444, 1078–1082 (2006).

  3. 3.

    Ng, A. K. et al. Structure of the influenza virus A H5N1 nucleoprotein: implications for RNA binding, oligomerization, and vaccine design. FASEB J. 22, 3638–3647 (2008).

  4. 4.

    Arranz, R. et al. The structure of native influenza virion ribonucleoproteins. Science 338, 1634–1637 (2012).

  5. 5.

    Eisfeld, A. J., Neumann, G. & Kawaoka, Y. At the centre: influenza A virus ribonucleoproteins. Nat. Rev. Microbiol. 13, 28–41 (2015).

  6. 6.

    Engelhardt, O. G., Smith, M. & Fodor, E. Association of the influenza A virus RNA-dependent RNA polymerase with cellular RNA polymerase II. J. Virol. 79, 5812–5818 (2005).

  7. 7.

    Lukarska, M. et al. Structural basis of an essential interaction between influenza polymerase and Pol II CTD. Nature 541, 117–121 (2017).

  8. 8.

    Rodriguez, A., Pérez-González, A. & Nieto, A. Influenza virus infection causes specific degradation of the largest subunit of cellular RNA polymerase II. J. Virol. 81, 5315–5324 (2007).

  9. 9.

    Serna Martin, I. et al. A mechanism for the activation of the influenza virus transcriptase. Mol. Cell 70, 1101–1110 (2018).

  10. 10.

    Bouloy, M., Plotch, S. J. & Krug, R. M. Globin mRNAs are primers for the transcription of influenza viral RNA in vitro. Proc. Natl Acad. Sci. USA 75, 4886–4890 (1978).

  11. 11.

    Kouba, T., Drncová, P. & Cusack, S. Structural snapshots of actively transcribing influenza polymerase. Nat. Struct. Mol. Biol. 26, 460–470 (2019).

  12. 12.

    Li, X. & Palese, P. Characterization of the polyadenylation signal of influenza virus RNA. J. Virol. 68, 1245–1249 (1994).

  13. 13.

    Robertson, J. S., Schubert, M. & Lazzarini, R. A. Polyadenylation sites for influenza virus mRNA. J. Virol. 38, 157–163 (1981).

  14. 14.

    Hay, A. J., Skehel, J. J. & McCauley, J. Structure and synthesis of influenza virus complementary RNAs. Phil. Trans. R. Soc. Lond. B 288, 341–348 (1980).

  15. 15.

    Vreede, F. T. & Brownlee, G. G. Influenza virion-derived viral ribonucleoproteins synthesize both mRNA and cRNA in vitro. J. Virol. 81, 2196–2204 (2007).

  16. 16.

    Jorba, N., Coloma, R. & Ortín, J. Genetic trans-complementation establishes a new model for influenza virus RNA transcription and replication. PLoS Pathog. 5, e1000462 (2009).

  17. 17.

    York, A., Hengrung, N., Vreede, F. T., Huiskonen, J. T. & Fodor, E. Isolation and characterization of the positive-sense replicative intermediate of a negative-strand RNA virus. Proc. Natl Acad. Sci. USA 110, E4238–E4245 (2013).

  18. 18.

    Huang, T. S., Palese, P. & Krystal, M. Determination of influenza virus proteins required for genome replication. J. Virol. 64, 5669–5673 (1990).

  19. 19.

    Moeller, A., Kirchdoerfer, R. N., Potter, C. S., Carragher, B. & Wilson, I. A. Organization of the influenza virus replication machinery. Science 338, 1631–1634 (2012).

  20. 20.

    Gallagher, J. R., Torian, U., McCraw, D. M. & Harris, A. K. Structural studies of influenza virus RNPs by electron microscopy indicate molecular contortions within NP supra-structures. J. Struct. Biol. 197, 294–307 (2017).

  21. 21.

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

  22. 22.

    Egelman, E. H. A robust algorithm for the reconstruction of helical filaments using single-particle methods. Ultramicroscopy 85, 225–234 (2000).

  23. 23.

    Vilas, J. L. et al. MonoRes: automatic and accurate estimation of local resolution for electron microscopy maps. Structure 26, 337–344 (2018).

  24. 24.

    Gerritz, S. W. et al. Inhibition of influenza virus replication via small molecules that induce the formation of higher-order nucleoprotein oligomers. Proc. Natl Acad. Sci. USA 108, 15366–15371 (2011).

  25. 25.

    Kao, R. Y. et al. Identification of influenza A nucleoprotein as an antiviral target. Nat. Biotechnol. 28, 600–605 (2010).

  26. 26.

    Amorim, M. J., Kao, R. Y. & Digard, P. Nucleozin targets cytoplasmic trafficking of viral ribonucleoprotein-Rab11 complexes in influenza A virus infection. J. Virol. 87, 4694–4703 (2013).

  27. 27.

    Te Velthuis, A. J. & Fodor, E. Influenza virus RNA polymerase: insights into the mechanisms of viral RNA synthesis. Nat. Rev. Microbiol. 14, 479–493 (2016).

  28. 28.

    Gerlach, P., Malet, H., Cusack, S. & Reguera, J. Structural insights into bunyavirus replication and its regulation by the vRNA promoter. Cell 161, 1267–1279 (2015).

  29. 29.

    Luo, G. X., Luytjes, W., Enami, M. & Palese, P. The polyadenylation signal of influenza virus RNA involves a stretch of uridines followed by the RNA duplex of the panhandle structure. J. Virol. 65, 2861–2867 (1991).

  30. 30.

    Pflug, A., Guilligay, D., Reich, S. & Cusack, S. Structure of influenza A polymerase bound to the viral RNA promoter. Nature 516, 355–360 (2014).

  31. 31.

    Reich, S. et al. Structural insight into cap-snatching and RNA synthesis by influenza polymerase. Nature 516, 361–366 (2014).

  32. 32.

    Resa-Infante, P., Recuero-Checa, M. A., Zamarreño, N., Llorca, O. & Ortín, J. Structural and functional characterization of an influenza virus RNA polymerase-genomic RNA complex. J. Virol. 84, 10477–10487 (2010).

  33. 33.

    Turrell, L., Lyall, J. W., Tiley, L. S., Fodor, E. & Vreede, F. T. The role and assembly mechanism of nucleoprotein in influenza A virus ribonucleoprotein complexes. Nat. Commun. 4, 1591 (2013).

  34. 34.

    Honda, A., Uéda, K., Nagata, K. & Ishihama, A. RNA polymerase of influenza virus: role of NP in RNA chain elongation. J. Biochem. 104, 1021–1026 (1988).

  35. 35.

    Coloma, R. et al. The structure of a biologically active influenza virus ribonucleoprotein complex. PLoS Pathog. 5, e1000491 (2009).

  36. 36.

    Ortín, J., Nájera, R., López, C., Dávila, M. & Domingo, E. Genetic variability of Hong Kong (H3N2) influenza viruses: spontaneous mutations and their location in the viral genome. Gene 11, 319–331 (1980).

  37. 37.

    Munier, S., Rolland, T., Diot, C., Jacob, Y. & Naffakh, N. Exploration of binary virus–host interactions using an infectious protein complementation assay. Mol. Cell. Proteomics 12, 2845–2855 (2013).

  38. 38.

    Fodor, E. et al. Rescue of influenza A virus from recombinant DNA. J. Virol. 73, 9679–9682 (1999).

  39. 39.

    Matrosovich, M., Matrosovich, T., Garten, W. & Klenk, H-D. New low-viscosity overlay medium for viral plaque assays. Virol. J. 3, 63 (2006).

  40. 40.

    Parvin, J. D., Palese, P., Honda, A., Ishihama, A. & Krystal, M. Promoter analysis of influenza virus RNA polymerase. J. Virol. 63, 5142–5152 (1989).

  41. 41.

    Compans, R. W., Content, J. & Duesberg, P. H. Structure of the ribonucleoprotein of influenza virus. J. Virol. 10, 795–800 (1972).

  42. 42.

    de la Rosa-Trevín, J. M. et al. Scipion: a software framework toward integration, reproducibility and validation in 3D electron microscopy. J. Struct. Biol. 195, 93–99 (2016).

  43. 43.

    Abrishami, V. et al. Alignment of direct detection device micrographs using a robust Optical Flow approach. J. Struct. Biol. 189, 163–176 (2015).

  44. 44.

    Li, X. et al. Electron counting and beam-induced motion correction enable near-atomic-resolution single-particle cryo-EM. Nat. Methods 10, 584–590 (2013).

  45. 45.

    Mindell, J. A. & Grigorieff, N. Accurate determination of local defocus and specimen tilt in electron microscopy. J. Struct. Biol. 142, 334–347 (2003).

  46. 46.

    Sorzano, C. O. et al. A clustering approach to multireference alignment of single-particle projections in electron microscopy. J. Struct. Biol. 171, 197–206 (2010).

  47. 47.

    Shaikh, T. R. et al. SPIDER image processing for single-particle reconstruction of biological macromolecules from electron micrographs. Nat. Protoc. 3, 1941–1974 (2008).

  48. 48.

    de la Rosa-Trevín, J. M. et al. Xmipp 3.0: an improved software suite for image processing in electron microscopy. J. Struct. Biol. 184, 321–328 (2013).

  49. 49.

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

  50. 50.

    Schneider, C. A., Rasband, W. S. & Eliceiri, K. W. NIH Image to ImageJ: 25 years of image analysis. Nat. Methods 9, 671–675 (2012).

  51. 51.

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

Download references

Acknowledgements

We thank S.H.W. Scheres, C. Savva and the Laboratory of Molecular Biology (Cambridge) for access to the Titan Krios microscope and technical assistance. We thank E. Sahagún for the generation of the Supplementary video (https://scixel.es/). The professional editing service NB Revisions (https://www.nbrevisions.com/) was used for technical editing of the manuscript before submission. We acknowledge the cryo-EM facility of the CNB–CIB (CSIC) for its technical advice and support throughout this work. This work was supported by the Spanish Ministry of Science, Innovation and Universities (Ministerio de Ciencia, Innovación y Universidades) grant nos. BFU2017-90018-R and BFU2011-25090/BMC (J.M.-B.) and Integrative Biology of Emerging Infectious Diseases LabEx grant no. 10-LABX-0062 (N.N.).

Author information

J.M.-B. and J.O. designed the experiments. R.C., R.A., D.C. and J.M.-B. carried out the experiments. J.M.R-T. and C.O.S.S. contributed the analysis tools. S.M. and N.N. contributed materials. R.C., R.A., J.O. and J.M.-B. analysed the data. J.M.-B. wrote the paper with contributions from all the other authors.

Correspondence to Juan Ortín or Jaime Martín-Benito.

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

Extended Data Fig. 1 Cryo-EM of wild type vRNPs.

a, Cryo-EM image of isolated vRNPs showing the extreme flexibility of the particles. b, Gallery of 2D averages with some examples of the different structures of the helical part of influenza vRNPs. Scale bars represent 100Å.

Extended Data Fig. 2 Variation of the relative position of two nucleoprotein monomers extracted from opposite strands of the helixes shown in Fig. 1 a-d of the main text.

For clarity, the equatorial region between the head (yellow) and body (green) domains has been marked with a red line, with the arrows indicating the direction of displacement. In all cases the position of the helical axis of the vRNP is represented with a vertical black line and the position of the dihedral axis is perpendicular to the plane of the figure and depicted by the () symbol.

Extended Data Fig. 3 Docking reliability of the nucleoprotein atomic structure (pdb 2IQH) into the cryo-EM density maps.

From top to bottom, six views representing two rotated positions (top and bottom raw for each case) of the cryo-EM density maps shown at three different thresholds (2.2, 2.8 and 3.2 σ) corresponding to the volumes shown in Fig. 1a–c, respectively. It is important to notice that cryo-EM maps contain the genomic ARN not present in the atomic structure. In all cases, the quality of the fit of the nucleoprotein atomic structure into the map is evident.

Extended Data Fig. 4 Negative staining electron microscopy characterization of nucleozin-treated vRNPs.

a, After nucleozin treatment, vRNPs appear as straight structures with a characteristic dark centerline along the helix. b, Longer vRNPs appear as broken helixes showing sharp corners. These structures probably formed from the initial binding of nucleozin at different points of one long helix followed by a cooperative extension of the conformational change induced by the drug; the growth of the straight segments generates these striking bends in the joint zones (see Extended Data Fig. 5 for more information). c, Gallery of averaged kinks selected from the nucleozin-treated vRNPs. The treated particles always show a characteristic central line and an increase in the diameter of the helix is clearly visible (compare with Extended Data Fig. 1b), indicating that a conformational change has occurred. Representative individual images of each class are shown on the right. Scale bar represents 200 Å in a and b and 150 Å in c.

Extended Data Fig. 5 Effect of nucleozin on vRNP structure.

a, Another structure obtained by cryo-EM after incubation of native vRNPs in the presence of nucleozin; the nucleozin binding site is marked with magenta circles. The structural changes are manifested in a decrease of z-rise, from 28–35 Å in untreated vRNPs to approximately 20 Å in the treated ones. Additionally, the phi angle drops from about 55–60 degrees to around 20 degrees. In consequence, the diameter of the helix increases (compare Fig. 3c and Extended Data Fig. 5a with Fig. 1a–d) and the vRNPs become shorter. Scale bar represents 50 Å. b, Scheme of the collapse of the major groove upon nucleozin treatment. On the left side, a draft of the wild type vRNP structure is shown, the upper (red) and lower (blue) strands and two pairs of nucleoproteins have been outlined, indicating the position of the nucleozin binding sites (magenta circles). In most cases, nucleozin probably binds to one nucleoprotein and the flexibility of the helix allows contact with the neighboring nucleoprotein of the contiguous turn, which produces the cross-linkage of both nucleoproteins, collapsing the major groove of the structure and opening the minor one. The right side of this panel shows two schemes for the helix after nucleozin binding corresponding to the structures shown in Fig. 3c and Extended Data Fig. 5a, respectively. c, Despite the cross-linking produced by nucleozin, vRNPs retain their flexibility due to the possibility of small movements among nucleoprotein dimers along the z-axis (left) or residual ability of the monomers to rotate (right). In this scheme, the nucleozin binding sites have been removed for clarity.

Extended Data Fig. 6 Decreased in vitro transcriptional activity of vRNPs pre-incubated with nucleozin.

a, Gel and quantification of mRNA synthesized in 1 h of transcription after 10 min of incubation with different concentrations of nucleozin. b, Gel and quantification of the shorter mRNAs synthesized in 1 h of transcription after 10 min of incubation with different concentrations of nucleozin. c, Effect of duration of nucleozin pre-incubation on mRNA synthesis by vRNPs. For treatments with small amounts of nucleozin (left panel, 1μM), the pre-treatment time had little influence on activity, but again affected longer mRNAs to a greater extent. One result of three independent experiments is shown.

Extended Data Fig. 7 Electron microscopy of vRNPs during transcription.

a, Image gallery of negatively stained vRNPs during the transcription process. In all cases a nucleoprotein loop is clearly visible at each end of the vRNP and the polymerase is located at some point along the helical part (blue arrows). The first two images show vRNPs where the His-tagged PB2 polymerase has been labeled with 5-nm Ni-NTA-Nanogold nanoprobe (black arrows). In some cases, an mRNA thread emerging from the polymerase was visible (green arrows). b, Gallery of non-activated control vRNPs showing the polymerase Ni-NTA-Nanogold labeled at one end. c, Average images obtained after alignment and classification of the region where the polymerase was located on the vRNPs during transcription. Each average was obtained from around 200 images; representative single images of the classes are shown on the right. Scale bars represent 150 Å.

Supplementary information

41564_2020_675_MOESM3_ESM.mp4

Schematic diagram of the movement of the vRNP for the processive helical track mechanism during transcription.

Supplementary Information

Supplementary Tables 1 and 2.

Reporting Summary

Supplementary Video

Schematic diagram of the movement of the vRNP for the processive helical track mechanism during transcription.

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Coloma, R., Arranz, R., de la Rosa-Trevín, J.M. et al. Structural insights into influenza A virus ribonucleoproteins reveal a processive helical track as transcription mechanism. Nat Microbiol (2020). https://doi.org/10.1038/s41564-020-0675-3

Download citation