Review Article | Published:

RNA-binding proteins in bacteria

Nature Reviews Microbiologyvolume 16pages601615 (2018) | Download Citation

Abstract

RNA-binding proteins (RBPs) are central to most if not all cellular processes, dictating the fate of virtually all RNA molecules in the cell. Starting with pioneering work on ribosomal proteins, studies of bacterial RBPs have paved the way for molecular studies of RNA–protein interactions. Work over the years has identified major RBPs that act on cellular transcripts at the various stages of bacterial gene expression and that enable their integration into post-transcriptional networks that also comprise small non-coding RNAs. Bacterial RBP research has now entered a new era in which RNA sequencing-based methods permit mapping of RBP activity in a truly global manner in vivo. Moreover, the soaring interest in understudied members of host-associated microbiota and environmental communities is likely to unveil new RBPs and to greatly expand our knowledge of RNA–protein interactions in bacteria.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Additional information

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

Protein Databank: http://www.rcsb.org/pdb/home/home.do

References

  1. 1.

    Fox, G. E. Origin and evolution of the ribosome. Cold Spring Harb. Perspect. Biol. 2, a003483 (2010).

  2. 2.

    Chaikam, V. & Karlson, D. T. Comparison of structure, function and regulation of plant cold shock domain proteins to bacterial and animal cold shock domain proteins. BMB Rep. 43, 1–8 (2010).

  3. 3.

    Hajnsdorf, E. & Boni, I. V. Multiple activities of RNA-binding proteins S1 and Hfq. Biochimie 94, 1544–1553 (2012).

  4. 4.

    Updegrove, T. B., Zhang, A. & Storz, G. Hfq: the flexible RNA matchmaker. Curr. Opin. Microbiol. 30, 133–138 (2016).

  5. 5.

    Koonin, E. V. & Makarova, K. S. CRISPR-Cas: evolution of an RNA-based adaptive immunity system in prokaryotes. RNA Biol. 10, 679–686 (2013).

  6. 6.

    Nicastro, G., Taylor, I. A. & Ramos, A. KH-RNA interactions: back in the groove. Curr. Opin. Struct. Biol. 30, 63–70 (2015).

  7. 7.

    Masliah, G., Barraud, P. & Allain, F. H. RNA recognition by double-stranded RNA binding domains: a matter of shape and sequence. Cell. Mol. Life Sci. 70, 1875–1895 (2013).

  8. 8.

    Swarts, D. C. et al. The evolutionary journey of Argonaute proteins. Nat. Struct. Mol. Biol. 21, 743–753 (2014).

  9. 9.

    Helder, S., Blythe, A. J., Bond, C. S. & Mackay, J. P. Determinants of affinity and specificity in RNA-binding proteins. Curr. Opin. Struct. Biol. 38, 83–91 (2016).

  10. 10.

    Smirnov, A. et al. Grad-seq guides the discovery of ProQ as a major small RNA-binding protein. Proc. Natl Acad. Sci. USA 113, 11591–11596 (2016). This study describes a high-throughput method for analysing global RNA–protein complexes and establishes ProQ as a global bacterial RBP.

  11. 11.

    Hentze, M. W., Castello, A., Schwarzl, T. & Preiss, T. A brave new world of RNA-binding proteins. Nat. Rev. Mol. Cell Biol. 19, 327–341 (2018). This is an excellent overview of the findings from recent screens for eukaryotic RBPs.

  12. 12.

    Gerstberger, S., Hafner, M. & Tuschl, T. A census of human RNA-binding proteins. Nat. Rev. Genet. 15, 829–845 (2014).

  13. 13.

    Morris, K. V. & Mattick, J. S. The rise of regulatory RNA. Nat. Rev. Genet. 15, 423–437 (2014).

  14. 14.

    Wagner, E. G. H. & Romby, P. Small RNAs in bacteria and archaea: who they are, what they do, and how they do it. Adv. Genet. 90, 133–208 (2015).

  15. 15.

    Van Assche, E., Van Puyvelde, S., Vanderleyden, J. & Steenackers, H. P. RNA-binding proteins involved in post-transcriptional regulation in bacteria. Front. Microbiol. 6, 141 (2015).

  16. 16.

    Hör, J., Gorski, S. A. & Vogel, J. Bacterial RNA biology on a genome scale. Mol. Cell 70, 785–799 (2018).

  17. 17.

    van der Oost, J., Westra, E. R., Jackson, R. N. & Wiedenheft, B. Unravelling the structural and mechanistic basis of CRISPR-Cas systems. Nat. Rev. Microbiol. 12, 479–492 (2014).

  18. 18.

    Hui, M. P., Foley, P. L. & Belasco, J. G. Messenger RNA degradation in bacterial cells. Annu. Rev. Genet. 48, 537–559 (2014).

  19. 19.

    Marbaniang, C. N. & Vogel, J. Emerging roles of RNA modifications in bacteria. Curr. Opin. Microbiol. 30, 50–57 (2016).

  20. 20.

    Mohanty, B. K. & Kushner, S. R. Regulation of mRNA decay in bacteria. Annu. Rev. Microbiol. 70, 25–44 (2016).

  21. 21.

    Ray-Soni, A., Bellecourt, M. J. & Landick, R. Mechanisms of bacterial transcription termination: all good things must end. Annu. Rev. Biochem. 85, 319–347 (2016). This review article summarizes the current knowledge about transcription termination and highlights unresolved key questions.

  22. 22.

    Roberts, J. W. Termination factor for RNA synthesis. Nature 224, 1168–1174 (1969).

  23. 23.

    Cardinale, C. J. et al. Termination factor Rho and its cofactors NusA and NusG silence foreign DNA in E. coli. Science 320, 935–938 (2008).

  24. 24.

    Sedlyarova, N. et al. sRNA-mediated control of transcription termination in E. coli. Cell 167, 111–121 e13 (2016).

  25. 25.

    Leela, J. K., Syeda, A. H., Anupama, K. & Gowrishankar, J. Rho-dependent transcription termination is essential to prevent excessive genome-wide R-loops in Escherichia coli. Proc. Natl Acad. Sci. USA 110, 258–263 (2013).

  26. 26.

    Dutta, D., Shatalin, K., Epshtein, V., Gottesman, M. E. & Nudler, E. Linking RNA polymerase backtracking to genome instability in E. coli. Cell 146, 533–543 (2011).

  27. 27.

    Washburn, R. S. & Gottesman, M. E. Transcription termination maintains chromosome integrity. Proc. Natl Acad. Sci. USA 108, 792–797 (2011).

  28. 28.

    Mitra, P., Ghosh, G., Hafeezunnisa, M. & Sen, R. Rho protein: roles and mechanisms. Annu. Rev. Microbiol. 71, 687–709 (2017).

  29. 29.

    Koslover, D. J., Fazal, F. M., Mooney, R. A., Landick, R. & Block, S. M. Binding and translocation of termination factor rho studied at the single-molecule level. J. Mol. Biol. 423, 664–676 (2012).

  30. 30.

    Epshtein, V., Dutta, D., Wade, J. & Nudler, E. An allosteric mechanism of Rho-dependent transcription termination. Nature 463, 245–249 (2010). This study provides evidence that Rho is associated with RNAP throughout the transcription cycle and that transcription termination involves a rearrangement of the RNAP catalytic centre.

  31. 31.

    Zhang, J. & Landick, R. A. Two-way street: regulatory interplay between RNA polymerase and nascent RNA structure. Trends Biochem. Sci. 41, 293–310 (2016).

  32. 32.

    Nudler, E. & Gottesman, M. E. Transcription termination and anti-termination in E. coli. Genes Cells 7, 755–768 (2002).

  33. 33.

    Mondal, S., Yakhnin, A. V., Sebastian, A., Albert, I. & Babitzke, P. NusA-dependent transcription termination prevents misregulation of global gene expression. Nat. Microbiol. 1, 15007 (2016).

  34. 34.

    Qayyum, M. Z., Dey, D. & Sen, R. Transcription elongation factor NusA is a general antagonist of Rho-dependent termination in Escherichia coli. J. Biol. Chem. 291, 8090–8108 (2016).

  35. 35.

    Guo, X. et al. Structural basis for NusA stabilized transcriptional pausing. Mol. Cell 69, 816–827 (2018). This study presents a cryo-EM structure of NusA in complex with a paused RNAP, providing insights into how NusA stimulates transcriptional pausing and termination.

  36. 36.

    Said, N. et al. Structural basis for lambdaN-dependent processive transcription antitermination. Nat. Microbiol. 2, 17062 (2017). This study describes crystal and cryo-EM structures of the phage λ antitermination complex.

  37. 37.

    Gollnick, P., Babitzke, P., Antson, A. & Yanofsky, C. Complexity in regulation of tryptophan biosynthesis in Bacillus subtilis. Annu. Rev. Genet. 39, 47–68 (2005).

  38. 38.

    Antson, A. A. et al. Structure of the trp RNA-binding attenuation protein, TRAP, bound to RNA. Nature 401, 235–242 (1999). This study describes the crystal structure of TRAP in complex with a GAG repeat-containing RNA target to reveal the structural basis for TRAP-mediated regulation of transcription and translation.

  39. 39.

    Babitzke, P., Stults, J. T., Shire, S. J. & Yanofsky, C. TRAP, the trp RNA-binding attenuation protein of Bacillus subtilis, is a multisubunit complex that appears to recognize G/UAG repeats in the trpEDCFBA and trpG transcripts. J. Biol. Chem. 269, 16597–16604 (1994).

  40. 40.

    Turnbough, C. L. Jr & Switzer, R. L. Regulation of pyrimidine biosynthetic gene expression in bacteria: repression without repressors. Microbiol. Mol. Biol. Rev. 72, 266–300 (2008).

  41. 41.

    Figueroa-Bossi, N. et al. RNA remodeling by bacterial global regulator CsrA promotes Rho-dependent transcription termination. Genes Dev. 28, 1239–1251 (2014).

  42. 42.

    Bae, W., Xia, B., Inouye, M. & Severinov, K. Escherichia coli CspA-family RNA chaperones are transcription antiterminators. Proc. Natl Acad. Sci. USA 97, 7784–7789 (2000).

  43. 43.

    Newkirk, K. et al. Solution NMR structure of the major cold shock protein (CspA) from Escherichia coli: identification of a binding epitope for DNA. Proc. Natl Acad. Sci. USA 91, 5114–5118 (1994).

  44. 44.

    Schindelin, H., Jiang, W., Inouye, M. & Heinemann, U. Crystal structure of CspA, the major cold shock protein of Escherichia coli. Proc. Natl Acad. Sci. USA 91, 5119–5123 (1994).

  45. 45.

    Sachs, R., Max, K. E., Heinemann, U. & Balbach, J. RNA single strands bind to a conserved surface of the major cold shock protein in crystals and solution. RNA 18, 65–76 (2012).

  46. 46.

    Xia, B., Ke, H. & Inouye, M. Acquirement of cold sensitivity by quadruple deletion of the cspA family and its suppression by PNPase S1 domain in Escherichia coli. Mol. Microbiol. 40, 179–188 (2001).

  47. 47.

    Zhang, Y. et al. A stress response that monitors and regulates mRNA structure is central to cold shock adaptation. Mol. Cell 70, 274–286 (2018).

  48. 48.

    Michaux, C. et al. RNA target profiles direct the discovery of virulence functions for the cold-shock proteins CspC and CspE. Proc. Natl Acad. Sci. USA 114, 6824–6829 (2017).

  49. 49.

    Phadtare, S., Tadigotla, V., Shin, W. H., Sengupta, A. & Severinov, K. Analysis of Escherichia coli global gene expression profiles in response to overexpression and deletion of CspC and CspE. J. Bacteriol. 188, 2521–2527 (2006).

  50. 50.

    Caballero, C. J. et al. The regulon of the RNA chaperone CspA and its auto-regulation in Staphylococcus aureus. Nucleic Acids Res. 46, 1345–1361 (2018).

  51. 51.

    Feng, Y., Huang, H., Liao, J. & Cohen, S. N. Escherichia coli poly(A)-binding proteins that interact with components of degradosomes or impede RNA decay mediated by polynucleotide phosphorylase and RNase E. J. Biol. Chem. 276, 31651–31656 (2001).

  52. 52.

    Phadtare, S. & Severinov, K. RNA remodeling and gene regulation by cold shock proteins. RNA Biol. 7, 788–795 (2010).

  53. 53.

    Phadtare, S. & Inouye, M. Sequence-selective interactions with RNA by CspB, CspC and CspE, members of the CspA family of Escherichia coli. Mol. Microbiol. 33, 1004–1014 (1999).

  54. 54.

    Kumarevel, T., Mizuno, H. & Kumar, P. K. Structural basis of HutP-mediated anti-termination and roles of the Mg2+ ion and l-histidine ligand. Nature 434, 183–191 (2005).

  55. 55.

    Gopinath, S. C. et al. Insights into anti-termination regulation of the hut operon in Bacillus subtilis: importance of the dual RNA-binding surfaces of HutP. Nucleic Acids Res. 36, 3463–3473 (2008).

  56. 56.

    Amster-Choder, O. The bgl sensory system: a transmembrane signaling pathway controlling transcriptional antitermination. Curr. Opin. Microbiol. 8, 127–134 (2005).

  57. 57.

    Amster-Choder, O. & Wright, A. Modulation of the dimerization of a transcriptional antiterminator protein by phosphorylation. Science 257, 1395–1398 (1992).

  58. 58.

    Goodson, J. R., Klupt, S., Zhang, C., Straight, P. & Winkler, W. C. LoaP is a broadly conserved antiterminator protein that regulates antibiotic gene clusters in Bacillus amyloliquefaciens. Nat. Microbiol. 2, 17003 (2017).

  59. 59.

    DebRoy, S. et al. Riboswitches. A riboswitch-containing sRNA controls gene expression by sequestration of a response regulator. Science 345, 937–940 (2014).

  60. 60.

    Mellin, J. R. et al. Riboswitches. Sequestration of a two-component response regulator by a riboswitch-regulated noncoding RNA. Science 345, 940–943 (2014).

  61. 61.

    Zere, T. R. et al. Genomic targets and features of BarA-UvrY (-SirA) signal transduction systems. PLOS ONE 10, e0145035 (2015).

  62. 62.

    Romeo, T., Gong, M., Liu, M. Y. & Brun-Zinkernagel, A. M. Identification and molecular characterization of csrA, a pleiotropic gene from Escherichia coli that affects glycogen biosynthesis, gluconeogenesis, cell size, and surface properties. J. Bacteriol. 175, 4744–4755 (1993).

  63. 63.

    Liu, M. Y. & Romeo, T. The global regulator CsrA of Escherichia coli is a specific mRNA-binding protein. J. Bacteriol. 179, 4639–4642 (1997).

  64. 64.

    Baker, C. S., Morozov, I., Suzuki, K., Romeo, T. & Babitzke, P. CsrA regulates glycogen biosynthesis by preventing translation of glgC in Escherichia coli. Mol. Microbiol. 44, 1599–1610 (2002).

  65. 65.

    Holmqvist, E. et al. Global RNA recognition patterns of post-transcriptional regulators Hfq and CsrA revealed by UV crosslinking in vivo. EMBO J. 35, 991–1011 (2016). This study describes a CLIP-seq protocol for bacterial RBPs to globally map binding sites of Hfq and CsrA in Salmonella.

  66. 66.

    Potts, A. H. et al. Global role of the bacterial post-transcriptional regulator CsrA revealed by integrated transcriptomics. Nat. Commun. 8, 1596 (2017). This study integrates several global methods, such as CLIP-seq, RNA-seq, ribosome profiling and global RNA stability assays, to link CsrA binding to changes in RNA translation, abundance and stability.

  67. 67.

    Sahr, T. et al. The Legionella pneumophila genome evolved to accommodate multiple regulatory mechanisms controlled by the CsrA-system. PLOS Genet. 13, e1006629 (2017).

  68. 68.

    Dubey, A. K., Baker, C. S., Romeo, T. & Babitzke, P. RNA sequence and secondary structure participate in high-affinity CsrA-RNA interaction. RNA 11, 1579–1587 (2005).

  69. 69.

    Schubert, M. et al. Molecular basis of messenger RNA recognition by the specific bacterial repressing clamp RsmA/CsrA. Nat. Struct. Mol. Biol. 14, 807–813 (2007).

  70. 70.

    Yakhnin, A. V. et al. CsrA activates flhDC expression by protecting flhDC mRNA from RNase E-mediated cleavage. Mol. Microbiol. 87, 851–866 (2013).

  71. 71.

    Weilbacher, T. et al. A novel sRNA component of the carbon storage regulatory system of Escherichia coli. Mol. Microbiol. 48, 657–670 (2003).

  72. 72.

    Duss, O. et al. Structural basis of the non-coding RNA RsmZ acting as a protein sponge. Nature 509, 588–592 (2014). In this study, NMR reveals the three-dimensional structure of RsmE in complex with its RNA sponge RsmZ.

  73. 73.

    Kusmierek, M. & Dersch, P. Regulation of host-pathogen interactions via the post-transcriptional Csr/Rsm system. Curr. Opin. Microbiol. 41, 58–67 (2017).

  74. 74.

    Katsowich, N. et al. Host cell attachment elicits posttranscriptional regulation in infecting enteropathogenic bacteria. Science 355, 735–739 (2017).

  75. 75.

    Yakhnin, H. et al. CsrA of Bacillus subtilis regulates translation initiation of the gene encoding the flagellin protein (hag) by blocking ribosome binding. Mol. Microbiol. 64, 1605–1620 (2007).

  76. 76.

    Mukherjee, S. et al. CsrA-FliW interaction governs flagellin homeostasis and a checkpoint on flagellar morphogenesis in Bacillus subtilis. Mol. Microbiol. 82, 447–461 (2011).

  77. 77.

    Dugar, G. et al. The CsrA-FliW network controls polar localization of the dual-function flagellin mRNA in Campylobacter jejuni. Nat. Commun. 7, 11667 (2016).

  78. 78.

    Agaras, B., Sobrero, P. & Valverde, C. A. CsrA/RsmA translational regulator gene encoded in the replication region of a Sinorhizobium meliloti cryptic plasmid complements Pseudomonas fluorescens rsmA/E mutants. Microbiology 159, 230–242 (2013).

  79. 79.

    Marden, J. N. et al. An unusual CsrA family member operates in series with RsmA to amplify posttranscriptional responses in Pseudomonas aeruginosa. Proc. Natl Acad. Sci. USA 110, 15055–15060 (2013).

  80. 80.

    Vogel, J. & Luisi, B. F. Hfq and its constellation of RNA. Nat. Rev. Microbiol. 9, 578–589 (2011).

  81. 81.

    Gorski, S. A., Vogel, J. & Doudna, J. A. RNA-based recognition and targeting: sowing the seeds of specificity. Nat. Rev. Mol. Cell Biol. 18, 215–228 (2017).

  82. 82.

    Melamed, S. et al. Global mapping of small RNA-target interactions in bacteria. Mol. Cell 63, 884–897 (2016).

  83. 83.

    Tree, J. J., Granneman, S., McAteer, S. P., Tollervey, D. & Gally, D. L. Identification of bacteriophage-encoded anti-sRNAs in pathogenic Escherichia coli. Mol. Cell 55, 199–213 (2014). This study describes the pioneering application of ultraviolet crosslinking and RNA sequencing in bacteria to globally map Hfq binding sites in vivo.

  84. 84.

    Waters, S. A. et al. Small RNA interactome of pathogenic E. coli revealed through crosslinking of RNase E. EMBO J. 36, 374–387 (2017).

  85. 85.

    Kavita, K., de Mets, F. & Gottesman, S. New aspects of RNA-based regulation by Hfq and its partner sRNAs. Curr. Opin. Microbiol. 42, 53–61 (2017).

  86. 86.

    Papenfort, K. & Vanderpool, C. K. Target activation by regulatory RNAs in bacteria. FEMS Microbiol. Rev. 39, 362–378 (2015).

  87. 87.

    Zhang, A., Wassarman, K. M., Ortega, J., Steven, A. C. & Storz, G. The Sm-like Hfq protein increases OxyS RNA interaction with target mRNAs. Mol. Cell 9, 11–22 (2002).

  88. 88.

    Møller, T. et al. Hfq: a bacterial Sm-like protein that mediates RNA-RNA interaction. Mol. Cell 9, 23–30 (2002). These two studies (references 87 and 88) establish Hfq as a bacterial homologue of Sm/Sm-like proteins and demonstrate that Hfq promotes sRNA–mRNA interactions.

  89. 89.

    Dimastrogiovanni, D. et al. Recognition of the small regulatory RNA RydC by the bacterial Hfq protein. Elife 3, e05375 (2014). This paper describes the first crystal structure of Hfq in complex with a natural sRNA.

  90. 90.

    Sauer, E. & Weichenrieder, O. Structural basis for RNA 3′-end recognition by Hfq. Proc. Natl Acad. Sci. USA 108, 13065–13070 (2011).

  91. 91.

    Link, T. M., Valentin-Hansen, P. & Brennan, R. G. Structure of Escherichia coli Hfq bound to polyriboadenylate RNA. Proc. Natl Acad. Sci. USA 106, 19292–19297 (2009).

  92. 92.

    Schumacher, M. A., Pearson, R. F., Moller, T., Valentin-Hansen, P. & Brennan, R. G. Structures of the pleiotropic translational regulator Hfq and an Hfq-RNA complex: a bacterial Sm-like protein. EMBO J. 21, 3546–3556 (2002).

  93. 93.

    Mikulecky, P. J. et al. Escherichia coli Hfq has distinct interaction surfaces for DsrA, rpoS and poly(A) RNAs. Nat. Struct. Mol. Biol. 11, 1206–1214 (2004).

  94. 94.

    Peng, Y., Curtis, J. E., Fang, X. & Woodson, S. A. Structural model of an mRNA in complex with the bacterial chaperone Hfq. Proc. Natl Acad. Sci. USA 111, 17134–17139 (2014).

  95. 95.

    Panja, S., Schu, D. J. & Woodson, S. A. Conserved arginines on the rim of Hfq catalyze base pair formation and exchange. Nucleic Acids Res. 41, 7536–7546 (2013).

  96. 96.

    Schu, D. J., Zhang, A., Gottesman, S. & Storz, G. Alternative Hfq-sRNA interaction modes dictate alternative mRNA recognition. EMBO J. 34, 2557–2573 (2015).

  97. 97.

    Olsen, A. S., Moller-Jensen, J., Brennan, R. G. & Valentin-Hansen, P. C-Terminally truncated derivatives of Escherichia coli Hfq are proficient in riboregulation. J. Mol. Biol. 404, 173–182 (2010).

  98. 98.

    Vecerek, B., Rajkowitsch, L., Sonnleitner, E., Schroeder, R. & Bläsi, U. The C-terminal domain of Escherichia coli Hfq is required for regulation. Nucleic Acids Res. 36, 133–143 (2008).

  99. 99.

    Santiago-Frangos, A., Kavita, K., Schu, D. J., Gottesman, S. & Woodson, S. A. C-terminal domain of the RNA chaperone Hfq drives sRNA competition and release of target RNA. Proc. Natl Acad. Sci. USA 113, E6089–E6096 (2016). This paper clarifies the role of the C-terminal domain of Hfq.

  100. 100.

    Fender, A., Elf, J., Hampel, K., Zimmermann, B. & Wagner, E. G. H. RNAs actively cycle on the Sm-like protein Hfq. Genes Dev. 24, 2621–2626 (2010).

  101. 101.

    Santiago-Frangos, A. & Woodson, S. A. Hfq chaperone brings speed dating to bacterial sRNA. Wiley Interdiscip. Rev. RNA 9, e1475 (2018).

  102. 102.

    Santiago-Frangos, A., Jeliazkov, J. R., Gray, J. J. & Woodson, S. A. Acidic C-terminal domains autoregulate the RNA chaperone Hfq. Elife 6, e27049 (2017).

  103. 103.

    Attia, A. S. et al. Moraxella catarrhalis expresses an unusual Hfq protein. Infect. Immun. 76, 2520–2530 (2008).

  104. 104.

    Sittka, A., Sharma, C. M., Rolle, K. & Vogel, J. Deep sequencing of Salmonella RNA associated with heterologous Hfq proteins in vivo reveals small RNAs as a major target class and identifies RNA processing phenotypes. RNA Biol. 6, 266–275 (2009).

  105. 105.

    Bouloc, P. & Repoila, F. Fresh layers of RNA-mediated regulation in Gram-positive bacteria. Curr. Opin. Microbiol. 30, 30–35 (2016).

  106. 106.

    Chen, J. & Gottesman, S. Hfq links translation repression to stress-induced mutagenesis in E. coli. Genes Dev. 31, 1382–1395 (2017).

  107. 107.

    Sonnleitner, E. et al. Interplay between the catabolite repression control protein Crc, Hfq and RNA in Hfq-dependent translational regulation in Pseudomonas aeruginosa. Nucleic Acids Res. 46, 1470–1485 (2017). This study establishes that the Crc protein of Pseudomonas is an auxiliary protein in translational repression that directly interacts with Hfq and the RNA target.

  108. 108.

    Moreno, R., Fonseca, P. & Rojo, F. Two small RNAs, CrcY and CrcZ, act in concert to sequester the Crc global regulator in Pseudomonas putida, modulating catabolite repression. Mol. Microbiol. 83, 24–40 (2012).

  109. 109.

    Sonnleitner, E. & Bläsi, U. Regulation of Hfq by the RNA CrcZ in Pseudomonas aeruginosa carbon catabolite repression. PLOS Genet. 10, e1004440 (2014).

  110. 110.

    Glover, J. N. et al. The FinO family of bacterial RNA chaperones. Plasmid 78, 79–87 (2015).

  111. 111.

    Holmqvist, E., Li, L., Bischler, T., Barquist, L. & Vogel, J. Global maps of ProQ binding in vivo reveal target recognition via RNA structure and stability control at mRNA 3′ ends. Mol. Cell 70, 971–982 (2018). In this study, ultraviolet CLIP-seq maps of ProQ binding in Salmonella and E. coli suggest a target recognition mode via RNA structure rather than sequence.

  112. 112.

    Smirnov, A., Wang, C., Drewry, L. L. & Vogel, J. Molecular mechanism of mRNA repression in trans by a ProQ-dependent small RNA. EMBO J. 36, 1029–1045 (2017).

  113. 113.

    Chaulk, S. G. et al. ProQ is an RNA chaperone that controls ProP levels in Escherichia coli. Biochemistry 50, 3095–3106 (2011).

  114. 114.

    Attaiech, L. et al. Silencing of natural transformation by an RNA chaperone and a multitarget small RNA. Proc. Natl Acad. Sci. USA 113, 8813–8818 (2016). This paper discovers an essential role of the FinO/ProQ family protein RocC in sRNA-mediated competence control in Legionella.

  115. 115.

    Attaiech, L., Glover, J. N. & Charpentier, X. RNA chaperones step out of Hfq’s shadow. Trends Microbiol. 25, 247–249 (2017).

  116. 116.

    Gonzalez, G. M. et al. Structure of the Escherichia coli ProQ RNA-binding protein. RNA 23, 696–711 (2017).

  117. 117.

    Olejniczak, M. & Storz, G. ProQ/FinO-domain proteins: another ubiquitous family of RNA matchmakers? Mol. Microbiol. 104, 905–915 (2017).

  118. 118.

    Kunte, H. J., Crane, R. A., Culham, D. E., Richmond, D. & Wood, J. M. Protein ProQ influences osmotic activation of compatible solute transporter ProP in Escherichia coli K-12. J. Bacteriol. 181, 1537–1543 (1999).

  119. 119.

    Davis, J. H. & Williamson, J. R. Structure and dynamics of bacterial ribosome biogenesis. Philos. Trans. R. Soc. Lond. B Biol. Sci. 372, 20160181 (2017).

  120. 120.

    Ramakrishnan, V. The ribosome emerges from a black box. Cell 159, 979–984 (2014).

  121. 121.

    Melnikov, S. et al. One core, two shells: bacterial and eukaryotic ribosomes. Nat. Struct. Mol. Biol. 19, 560–567 (2012).

  122. 122.

    Voorhees, R. M. & Ramakrishnan, V. Structural basis of the translational elongation cycle. Annu. Rev. Biochem. 82, 203–236 (2013).

  123. 123.

    Shajani, Z., Sykes, M. T. & Williamson, J. R. Assembly of bacterial ribosomes. Annu. Rev. Biochem. 80, 501–526 (2011).

  124. 124.

    Meyer, M. M. rRNA mimicry in RNA regulation of gene expression. Microbiol. Spectr. 6, RWR-0006-2017 (2018).

  125. 125.

    Merianos, H. J., Wang, J. & Moore, P. B. The structure of a ribosomal protein S8/spc operon mRNA complex. RNA 10, 954–964 (2004).

  126. 126.

    Marzi, S. et al. Structured mRNAs regulate translation initiation by binding to the platform of the ribosome. Cell 130, 1019–1031 (2007).

  127. 127.

    Babitzke, P., Baker, C. S. & Romeo, T. Regulation of translation initiation by RNA binding proteins. Annu. Rev. Microbiol. 63, 27–44 (2009).

  128. 128.

    Duval, M. et al. Escherichia coli ribosomal protein S1 unfolds structured mRNAs onto the ribosome for active translation initiation. PLOS Biol. 11, e1001731 (2013). In this paper, protein S1 is shown to be required for translation initiation at structured mRNAs by two different mechanisms.

  129. 129.

    Boni, I. V., Artamonova, V. S., Tzareva, N. V. & Dreyfus, M. Non-canonical mechanism for translational control in bacteria: synthesis of ribosomal protein S1. EMBO J. 20, 4222–4232 (2001).

  130. 130.

    Butler, J. S., Springer, M., Dondon, J., Graffe, M. & Grunberg-Manago, M. Escherichia coli protein synthesis initiation factor IF3 controls its own gene expression at the translational level in vivo. J. Mol. Biol. 192, 767–780 (1986).

  131. 131.

    Romby, P. & Springer, M. Bacterial translational control at atomic resolution. Trends Genet. 19, 155–161 (2003).

  132. 132.

    Springer, M. et al. Autogenous control of Escherichia coli threonyl-tRNA synthetase expression in vivo. J. Mol. Biol. 185, 93–104 (1985).

  133. 133.

    Caillet, J. et al. The modular structure of Escherichia coli threonyl-tRNA synthetase as both an enzyme and a regulator of gene expression. Mol. Microbiol. 47, 961–974 (2003).

  134. 134.

    Jain, C. & Belasco, J. G. Autoregulation of RNase E synthesis in Escherichia coli. Nucleic Acids Symp. Ser. issue 3, 85–88 (1995).

  135. 135.

    Jarrige, A. C., Mathy, N. & Portier, C. PNPase autocontrols its expression by degrading a double-stranded structure in the pnp mRNA leader. EMBO J. 20, 6845–6855 (2001).

  136. 136.

    Ait-Bara, S. & Carpousis, A. J. RNA degradosomes in bacteria and chloroplasts: classification, distribution and evolution of RNase E homologs. Mol. Microbiol. 97, 1021–1135 (2015). This is an excellent review describing the biology and evolution of degradosomes.

  137. 137.

    Laalami, S., Zig, L. & Putzer, H. Initiation of mRNA decay in bacteria. Cell. Mol. Life Sci. 71, 1799–1828 (2014).

  138. 138.

    Redder, P., Hausmann, S., Khemici, V., Yasrebi, H. & Linder, P. Bacterial versatility requires DEAD-box RNA helicases. FEMS Microbiol. Rev. 39, 392–412 (2015).

  139. 139.

    Py, B., Higgins, C. F., Krisch, H. M. & Carpousis, A. J. A. DEAD-box RNA helicase in the Escherichia coli RNA degradosome. Nature 381, 169–172 (1996).

  140. 140.

    Coburn, G. A., Miao, X., Briant, D. J. & Mackie, G. A. Reconstitution of a minimal RNA degradosome demonstrates functional coordination between a 3′ exonuclease and a DEAD-box RNA helicase. Genes Dev. 13, 2594–2603 (1999).

  141. 141.

    Oun, S. et al. The CshA DEAD-box RNA helicase is important for quorum sensing control in Staphylococcus aureus. RNA Biol. 10, 157–165 (2013).

  142. 142.

    Morita, T., Maki, K. & Aiba, H. RNase E-based ribonucleoprotein complexes: mechanical basis of mRNA destabilization mediated by bacterial noncoding RNAs. Genes Dev. 19, 2176–2186 (2005).

  143. 143.

    Bruce, H. A. et al. Analysis of the natively unstructured RNA/protein-recognition core in the Escherichia coli RNA degradosome and its interactions with regulatory RNA/Hfq complexes. Nucleic Acids Res. 46, 387–402 (2018).

  144. 144.

    Bandyra, K. J. et al. The seed region of a small RNA drives the controlled destruction of the target mRNA by the endoribonuclease RNase E. Mol. Cell 47, 943–953 (2012).

  145. 145.

    Pfeiffer, V., Papenfort, K., Lucchini, S., Hinton, J. C. & Vogel, J. Coding sequence targeting by MicC RNA reveals bacterial mRNA silencing downstream of translational initiation. Nat. Struct. Mol. Biol. 16, 840–846 (2009).

  146. 146.

    Göpel, Y., Papenfort, K., Reichenbach, B., Vogel, J. & Görke, B. Targeted decay of a regulatory small RNA by an adaptor protein for RNase E and counteraction by an anti-adaptor RNA. Genes Dev. 27, 552–564 (2013).

  147. 147.

    Gonzalez, G. M. et al. Structural insights into RapZ-mediated regulation of bacterial amino-sugar metabolism. Nucleic Acids Res. 45, 10845–10860 (2017). In this study, the three-dimensional structure of the adaptor protein RapZ shows its functional configuration to be a tetramer.

  148. 148.

    Vakulskas, C. A. et al. Antagonistic control of the turnover pathway for the global regulatory sRNA CsrB by the CsrA and CsrD proteins. Nucleic Acids Res. 44, 7896–7910 (2016).

  149. 149.

    Potts, A. H., Leng, Y., Babitzke, P. & Romeo, T. Examination of Csr regulatory circuitry using epistasis analysis with RNA-seq (Epi-seq) confirms that CsrD affects gene expression via CsrA, CsrB and CsrC. Sci. Rep. 8, 5373 (2018).

  150. 150.

    Chen, X. et al. An RNA degradation machine sculpted by Ro autoantigen and noncoding RNA. Cell 153, 166–177 (2013). In this study, the Ro protein is shown to work with non-coding RNAs to channel structured RNAs into PNPase for degradation.

  151. 151.

    Moll, I., Afonyushkin, T., Vytvytska, O., Kaberdin, V. R. & Bläsi, U. Coincident Hfq binding and RNase E cleavage sites on mRNA and small regulatory RNAs. RNA 9, 1308–1314 (2003).

  152. 152.

    Chao, Y. et al. In vivo cleavage map illuminates the central role of RNase E in coding and non-coding RNA pathways. Mol. Cell 65, 39–51 (2017).

  153. 153.

    Chao, Y. & Vogel, J. A. 3′ UTR-derived small RNA provides the regulatory noncoding arm of the inner membrane stress response. Mol. Cell 61, 352–363 (2016).

  154. 154.

    Papenfort, K., Sun, Y., Miyakoshi, M., Vanderpool, C. K. & Vogel, J. Small RNA-mediated activation of sugar phosphatase mRNA regulates glucose homeostasis. Cell 153, 426–437 (2013).

  155. 155.

    Pandey, S. P. et al. Central role for RNase YbeY in Hfq-dependent and Hfq-independent small-RNA regulation in bacteria. BMC Genomics 15, 121 (2014).

  156. 156.

    Regnier, P. & Hajnsdorf, E. The interplay of Hfq, poly(A) polymerase I and exoribonucleases at the 3′ ends of RNAs resulting from Rho-independent termination: a tentative model. RNA Biol. 10, 602–609 (2013).

  157. 157.

    Andrade, J. M., Pobre, V., Matos, A. M. & Arraiano, C. M. The crucial role of PNPase in the degradation of small RNAs that are not associated with Hfq. RNA 18, 844–855 (2012).

  158. 158.

    Liou, G. G., Jane, W. N., Cohen, S. N., Lin, N. S. & Lin-Chao, S. RNA degradosomes exist in vivo in Escherichia coli as multicomponent complexes associated with the cytoplasmic membrane via the N-terminal region of ribonuclease E. Proc. Natl Acad. Sci. USA 98, 63–68 (2001).

  159. 159.

    Sharan, M., Forstner, K. U., Eulalio, A. & Vogel, J. APRICOT: an integrated computational pipeline for the sequence-based identification and characterization of RNA-binding proteins. Nucleic Acids Res. 45, e96 (2017).

  160. 160.

    Greenberg, J. R. Ultraviolet light-induced crosslinking of mRNA to proteins. Nucleic Acids Res. 6, 715–732 (1979).

  161. 161.

    Huang, R., Han, M., Meng, L. & Chen, X. Transcriptome-wide discovery of coding and noncoding RNA-binding proteins. Proc. Natl Acad. Sci. USA 115, E3879–E3887 (2018).

  162. 162.

    Tawk, C., Sharan, M., Eulalio, A. & Vogel, J. A systematic analysis of the RNA-targeting potential of secreted bacterial effector proteins. Sci. Rep. 7, 9328 (2017).

  163. 163.

    Nichols, R. J. et al. Phenotypic landscape of a bacterial cell. Cell 144, 143–156 (2011).

  164. 164.

    Hattman, S. Unusual transcriptional and translational regulation of the bacteriophage Mu mom operon. Pharmacol. Ther. 84, 367–388 (1999).

  165. 165.

    Romaniuk, P. J., Lowary, P., Wu, H. N., Stormo, G. & Uhlenbeck, O. C. RNA binding site of R17 coat protein. Biochemistry 26, 1563–1568 (1987).

  166. 166.

    McPheeters, D. S., Stormo, G. D. & Gold, L. Autogenous regulatory site on the bacteriophage T4 gene 32 messenger RNA. J. Mol. Biol. 201, 517–535 (1988).

  167. 167.

    Said, N. et al. In vivo expression and purification of aptamer-tagged small RNA regulators. Nucleic Acids Res. 37, e133 (2009).

  168. 168.

    Lopez-Alonso, J. P. et al. RsgA couples the maturation state of the 30S ribosomal decoding centre to activation of its GTPase pocket. Nucleic Acids Res. 45, 6945-6959 (2017).

  169. 169.

    Ataide, S. F. et al. The crystal structure of the signal recognition particle in complex with its receptor. Science 331, 881–886 (2011).

  170. 170.

    Huter, P., Muller, C., Arenz, S., Beckert, B. & Wilson, D. N. Structural basis for ribosome rescue in bacteria. Trends Biochem. Sci. 42, 669–680 (2017).

  171. 171.

    Neubauer, C., Gillet, R., Kelley, A. C. & Ramakrishnan, V. Decoding in the absence of a codon by tmRNA and SmpB in the ribosome. Science 335, 1366–1369 (2012).

  172. 172.

    Kohler, R., Mooney, R. A., Mills, D. J., Landick, R. & Cramer, P. Architecture of a transcribing-translating expressome. Science 356, 194–197 (2017). In this paper, the cryo-EM structure of E. coli RNAP together with the ribosome provides atomic resolution of transcription–translation coupling.

  173. 173.

    Alen, C. & Sonenshein, A. L. Bacillus subtilis aconitase is an RNA-binding protein. Proc. Natl Acad. Sci. USA 96, 10412–10417 (1999).

  174. 174.

    Benjamin, J. A. & Massé, E. The iron-sensing aconitase B binds its own mRNA to prevent sRNA-induced mRNA cleavage. Nucleic Acids Res. 42, 10023–10036 (2014). In this paper, the moonlighting TCA cycle enzyme aconitase B of E. coli protects the 3′ UTR of its own mRNA against RNase E-dependent degradation.

  175. 175.

    Mitobe, J. et al. RodZ regulates the post-transcriptional processing of the Shigella sonnei type III secretion system. EMBO Rep. 12, 911–916 (2011).

  176. 176.

    Keffer-Wilkes, L. C., Veerareddygari, G. R. & Kothe, U. RNA modification enzyme TruB is a tRNA chaperone. Proc. Natl Acad. Sci. USA 113, 14306–14311 (2016).

  177. 177.

    Beljantseva, J. et al. Negative allosteric regulation of Enterococcus faecalis small alarmone synthetase RelQ by single-stranded RNA. Proc. Natl Acad. Sci. USA 114, 3726–3731 (2017).

  178. 178.

    Qian, Z., Zhurkin, V. B. & Adhya, S. DNA-RNA interactions are critical for chromosome condensation in Escherichia coli. Proc. Natl Acad. Sci. USA 114, 12225–12230 (2017).

  179. 179.

    Brescia, C. C., Kaw, M. K. & Sledjeski, D. D. The DNA binding protein H-NS binds to and alters the stability of RNA in vitro and in vivo. J. Mol. Biol. 339, 505–514 (2004).

  180. 180.

    Deighan, P., Free, A. & Dorman, C. J. A role for the Escherichia coli H-NS-like protein StpA in OmpF porin expression through modulation of micF RNA stability. Mol. Microbiol. 38, 126–139 (2000).

  181. 181.

    Morrison, J. M., Anderson, K. L., Beenken, K. E., Smeltzer, M. S. & Dunman, P. M. The staphylococcal accessory regulator, SarA, is an RNA-binding protein that modulates the mRNA turnover properties of late-exponential and stationary phase Staphylococcus aureus cells. Front. Cell. Infect. Microbiol. 2, 26 (2012).

  182. 182.

    Buskila, A. A., Kannaiah, S. & Amster-Choder, O. RNA localization in bacteria. RNA Biol. 11, 1051–1060 (2014).

  183. 183.

    Wallace, J. G., Zhou, Z. & Breaker, R. R. OLE RNA protects extremophilic bacteria from alcohol toxicity. Nucleic Acids Res. 40, 6898–6907 (2012).

  184. 184.

    Kuwada, N. J., Traxler, B. & Wiggins, P. A. Genome-scale quantitative characterization of bacterial protein localization dynamics throughout the cell cycle. Mol. Microbiol. 95, 64–79 (2015).

  185. 185.

    Landgraf, D., Okumus, B., Chien, P., Baker, T. A. & Paulsson, J. Segregation of molecules at cell division reveals native protein localization. Nat. Methods 9, 480–482 (2012).

  186. 186.

    Short, F. L. et al. Selectivity and self-assembly in the control of a bacterial toxin by an antitoxic noncoding RNA pseudoknot. Proc. Natl Acad. Sci. USA 110, E241–E249 (2013). This study describes an intriguing co-crystal structure of the toxic endoribonuclease ToxN and the pseudoknot-forming antitoxin RNA ToxI.

  187. 187.

    Rajagopala, S. V. et al. The binary protein-protein interaction landscape of Escherichia coli. Nat. Biotechnol. 32, 285–290 (2014).

Download references

Acknowledgements

The authors thank C. Beisel, Y. Chao, K. Papenfort and G. Wagner for comments on the manuscript. J.V. is supported by a DFG Gottfried Wilhelm Leibniz Award (Vo875/20). E.H. is supported by the Wenner-Gren Foundations, the Swedish Research Council (2016–03656) and the Swedish Foundation for Strategic Research (ICA 16–0021).

Reviewer information

Nature Reviews Microbiology thanks M. Hentze, B. Luisi and E. Nudler for their contribution to the peer review of this work.

Author information

Affiliations

  1. Department of Cell and Molecular Biology, Biomedical Center, Uppsala University, Uppsala, Sweden

    • Erik Holmqvist
  2. Helmholtz Institute for RNA-based Infection Research (HIRI), Würzburg, Germany

    • Jörg Vogel
  3. Institute of Molecular Infection Biology, University of Würzburg, Würzburg, Germany

    • Jörg Vogel

Authors

  1. Search for Erik Holmqvist in:

  2. Search for Jörg Vogel in:

Contributions

E.H. and J.V. researched data for the article, made substantial contributions to discussions of the content, wrote the article and reviewed and/or edited the manuscript before submission.

Competing interests

The authors declare no competing interests.

Corresponding author

Correspondence to Jörg Vogel.

Supplementary information

Glossary

About this article

Publication history

Published

DOI

https://doi.org/10.1038/s41579-018-0049-5

Further reading