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RNA-binding proteins in bacteria

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.

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Fig. 1: Overview of bacterial RBPs.
Fig. 2: RBP-mediated regulation of transcription termination.
Fig. 3: RBP-based regulation of translation.
Fig. 4: RBP-dependent regulation of RNA decay.

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References

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

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

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

    Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    Google Scholar 

  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.

    Google Scholar 

  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.

    Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    Google Scholar 

  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).

    CAS  Google Scholar 

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

    PubMed  PubMed Central  Google Scholar 

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

    Google Scholar 

  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.

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

  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.

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

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

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

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

    Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

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

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

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

    PubMed  PubMed Central  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  Google Scholar 

  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.

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    PubMed  PubMed Central  Google Scholar 

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

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  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).

    CAS  Google Scholar 

  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).

    CAS  PubMed  PubMed Central  Google Scholar 

  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.

    CAS  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

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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).

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

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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.

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Supplementary information

Glossary

Ribosomal proteins

(r-proteins). All proteins that, together with the ribosomal RNA, form the two subunits of the ribosome, the molecular machine that synthesizes polypeptides from an mRNA template.

Small non-coding RNAs

(sRNAs). Bacterial, in most cases non-coding, RNAs in the size range of 50–300 nucleotides that carry out regulatory functions, either by base pairing to complementary sequences in other RNAs or by functioning as binding partners for proteins.

RNA sequencing

(RNA-seq). A method for the parallel determination of sequence and abundance of RNA molecules in a biological sample.

CRISPR–Cas systems

Bacterial adaptive immune systems consisting of short RNAs that guide Cas proteins to target invading nucleic acids for destruction.

Ribonucleoprotein particles

(RNPs). Macromolecular complexes that consist of RNA and RNA-binding proteins.

Ribonucleases

Enzymes that catalyse the cleavage of RNA. Endoribonucleases cleave RNA internally, whereas exoribonucleases degrade RNA from either the 5′ end or the 3′ end.

RNA-modification proteins

Proteins that introduce chemical modifications in RNA.

Intrinsic termination

A mechanism used by bacteria to stop transcription elongation and release the newly synthesized RNA. RNA polymerase is released from the nascent transcript as it encounters a palindromic GC-rich DNA sequence followed by a stretch of T residues. Also called Rho-independent termination.

Rho-dependent termination

A bacterial transcription termination mechanism whereby the release of the transcribing RNA polymerase from the DNA template is mediated by the Rho protein.

Rho

A homohexameric protein that promotes termination of transcription elongation by means of ATP hydrolysis. Rho is the target of the antibiotic bicyclomycin.

Ribosome binding site

(RBS). An mRNA sequence that recruits the 30 S ribosomal subunit to initiate translation. The Shine–Dalgarno sequence of the RBS is complementary to the 16 S ribosomal RNA (rRNA) and enables 30S–mRNA interaction.

trans-encoded

A concept to describe the relation between two genetic elements. A small non-coding RNA (sRNA) is trans-encoded with respect to its mRNA target if the two are encoded by different genetic loci. By contrast, a cis-encoded sRNA is encoded by the same locus as its mRNA target, for instance, when the respective genes overlap.

Degradosomes

Multi-protein complexes that carry out RNA degradation in bacteria.

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Holmqvist, E., Vogel, J. RNA-binding proteins in bacteria. Nat Rev Microbiol 16, 601–615 (2018). https://doi.org/10.1038/s41579-018-0049-5

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