Diverse and unified mechanisms of transcription initiation in bacteria

Transcription of DNA is a fundamental process in all cellular organisms. The enzyme responsible for transcription, RNA polymerase, is conserved in general architecture and catalytic function across the three domains of life. Diverse mechanisms are used among and within the different branches to regulate transcription initiation. Mechanistic studies of transcription initiation in bacteria are especially amenable because the promoter recognition and melting steps are much less complicated than in eukaryotes or archaea. Also, bacteria have critical roles in human health as pathogens and commensals, and the bacterial RNA polymerase is a proven target for antibiotics. Recent biophysical studies of RNA polymerases and their inhibition, as well as transcription initiation and transcription factors, have detailed the mechanisms of transcription initiation in phylogenetically diverse bacteria, inspiring this Review to examine unifying and diverse themes in this process.

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References

  1. Crick, F. Central dogma of molecular biology. Nature227, 561–563 (1970). CASPubMedGoogle Scholar
  2. Werner, F. & Grohmann, D. Evolution of multisubunit RNA polymerases in the three domains of life. Nat. Rev. Microbiol.9, 85–98 (2011). CASPubMedGoogle Scholar
  3. Decker, K. B. & Hinton, D. M. Transcription regulation at the core: similarities among bacterial, archaeal, and eukaryotic RNA polymerases. Annu. Rev. Microbiol.67, 113–139 (2013). CASPubMedGoogle Scholar
  4. Pribnow, D. Nucleotide sequence of an RNA polymerase binding site at an early T7 promoter. Proc. Natl Acad. Sci. USA72, 784–788 (1975). CASPubMedPubMed CentralGoogle Scholar
  5. Blombach, F., Smollett, K. L., Grohmann, D. & Werner, F. Molecular mechanisms of transcription initiation — structure, function, and evolution of TFE/TFIIE-like factors and open complex formation. J. Mol. Biol.428, 2592–2606 (2016). CASPubMedGoogle Scholar
  6. Cramer, P. Structure and function of RNA polymerase II. Adv. Protein Chem.67, 1–42 (2004). CASPubMedGoogle Scholar
  7. Burgess, R. R. Separation and characterization of the subunits of ribonucleic acid polymerase. J. Biol. Chem.244, 6168–6176 (1969). CASPubMedGoogle Scholar
  8. Zhang, G. et al. Crystal structure of Thermus aquaticus core RNA polymerase at 3.3 A resolution. Cell98, 811–824 (1999). First crystal structure of a DNA-dependent RNAP that provided high-resolution details of the core RNAP and defined many of the RNAP modules. CASPubMedGoogle Scholar
  9. Cramer, P. Architecture of RNA polymerase II and implications for the transcription mechanism. Science288, 640–649 (2000). CASPubMedGoogle Scholar
  10. Hirata, A., Klein, B. J. & Murakami, K. S. The X-ray crystal structure of RNA polymerase from Archaea. Nature451, 851–854 (2008). CASPubMedPubMed CentralGoogle Scholar
  11. Burgess, R. R., Travers, A. A., Dunn, J. J. & Bautz, E. K. F. Factor stimulating transcription by RNA polymerase. Nature221, 43–46 (1969). Demonstrated that a core RNAP required a σ-factor for transcription. CASPubMedGoogle Scholar
  12. Gruber, T. M. & Gross, C. A. Multiple sigma subunits and the partitioning of bacterial transcription space. Annu. Rev. Microbiol.57, 441–466 (2003). CASPubMedGoogle Scholar
  13. Feklistov, A., Sharon, B. D., Darst, S. A. & Gross, C. A. Bacterial sigma factors: a historical, structural, and genomic perspective. Annu. Rev. Microbiol.68, 357–376 (2014). CASPubMedGoogle Scholar
  14. Campbell, E. A. et al. Structure of the bacterial RNA polymerase promoter specificity sigma subunit. Mol. Cell9, 527–539 (2002). CASPubMedGoogle Scholar
  15. Callaci, S. & Heyduk, T. Conformation and DNA binding properties of a single-stranded DNA binding region of σ 70 subunit from Escherichia coli RNA polymerase are modulated by an interaction with the core enzyme. Biochemistry37, 3312–3320 (1998). CASPubMedGoogle Scholar
  16. Murakami, K. S., Masuda, S. & Darst, S. A. Structural basis of transcription initiation: RNA polymerase holoenzyme at 4 Å resolution. Science296, 1280–1284 (2002). CASPubMedGoogle Scholar
  17. Vassylyev, D. G. et al. Crystal structure of a bacterial RNA polymerase holoenzyme at 2.6 Å resolution. Nature417, 712–719 (2002). High-resolution structure of RNAP holoenzyme showing the housekeeping σ-factor domain architecture in context with RNAP. CASPubMedGoogle Scholar
  18. Danson, A. E., Jovanovic, M., Buck, M. & Zhang, X. Mechanisms of σ54-dependent transcription initiation and regulation. J. Mol. Biol.431, 3960–3974 (2019). CASPubMedPubMed CentralGoogle Scholar
  19. Hawley, D. K. & McClure, W. R. Compilation and analysis of Escherichia coli promoter DNA sequences. Nucleic Acids Res11, 2237–2255 (1983). CASPubMedPubMed CentralGoogle Scholar
  20. Simpson, R. B. The molecular topography of RNA polymerase-promoter interaction. Cell18, 277–285 (1979). CASPubMedGoogle Scholar
  21. Ross, W. et al. A third recognition element in bacterial promoters: DNA binding by the alpha subunit of RNA polymerase. Science262, 1407–1413 (1993). Describes the discovery of upstream elements. CASPubMedGoogle Scholar
  22. Barne, K. A., Bown, J. A., Busby, S. J. & Minchin, S. D. Region 2.5 of the Escherichia coli RNA polymerase sigma70 subunit is responsible for the recognition of the ‘extended-10’ motif at promoters. EMBO J.16, 4034–4040 (1997). CASPubMedPubMed CentralGoogle Scholar
  23. Haugen, S. P. et al. rRNA promoter regulation by nonoptimal binding of σ region 1.2: an additional recognition element for RNA polymerase. Cell125, 1069–1082 (2006). CASPubMedGoogle Scholar
  24. Zhang, Y. et al. Structural basis of transcription initiation. Science338, 1076–1080 (2012). CASPubMedPubMed CentralGoogle Scholar
  25. Feklistov, A. & Darst, S. A. Structural basis for promoter −10 element recognition by the bacterial RNA polymerase σ subunit. Cell147, 1257–1269 (2011). First high-resolution view describing the interaction of the σ-factor with the −10 element. CASPubMedPubMed CentralGoogle Scholar
  26. Hubin, E. A., Lilic, M., Darst, S. A. & Campbell, E. A. Structural insights into the mycobacteria transcription initiation complex from analysis of X-ray crystal structures. Nat. Commun.8, 16072 (2017). CASPubMedPubMed CentralGoogle Scholar
  27. Murakami, K. S. Structural basis of transcription initiation: an RNA polymerase holoenzyme-DNA complex. Science296, 1285–1290 (2002). CASPubMedGoogle Scholar
  28. Bae, B., Feklistov, A., Lass-Napiorkowska, A., Landick, R. & Darst, S. A. Structure of a bacterial RNA polymerase holoenzyme open promoter complex. eLife4, e08504 (2015). High-resolution crystal structure of a transcription initiation complex with a complete bubble and nascent RNA. PubMed CentralGoogle Scholar
  29. Zuo, Y. & Steitz, T. A. Crystal structures of the E. coli transcription initiation complexes with a complete bubble. Mol. Cell58, 534–540 (2015). First crystal structure at mid-range resolution of a transcription initiation complex with a complete bubble and nascent RNA. CASPubMedPubMed CentralGoogle Scholar
  30. Cortes, T. et al. Genome-wide mapping of transcriptional start sites defines an extensive leaderless transcriptome in mycobacterium tuberculosis. Cell Rep.5, 1121–1131 (2013). CASPubMedPubMed CentralGoogle Scholar
  31. Shell, S. S. et al. Leaderless transcripts and small proteins are common features of the mycobacterial translational landscape. PLoS Genet.11, e1005641 (2015). PubMedPubMed CentralGoogle Scholar
  32. Chakraborty, A. et al. Opening and closing of the bacterial RNA polymerase clamp. Science337, 591–595 (2012). Used single-molecule fluorescence energy transfer experiments to detect the motions of the RNAP clamp. CASPubMedPubMed CentralGoogle Scholar
  33. Duchi, D., Mazumder, A., Malinen, A. M., Ebright, R. H. & Kapanidis, A. N. The RNA polymerase clamp interconverts dynamically among three states and is stabilized in a partly closed state by ppGpp. Nucleic Acids Res.46, 7284–7295 (2018). CASPubMedPubMed CentralGoogle Scholar
  34. Feklistov, A. et al. RNA polymerase motions during promoter melting. Science356, 863–866 (2017). CASPubMedPubMed CentralGoogle Scholar
  35. Ruff, E. F. et al. E. coli RNA polymerase determinants of open complex lifetime and structure. J. Mol. Biol.427, 2435–2450 (2015). CASPubMedPubMed CentralGoogle Scholar
  36. Saecker, R. M., Record, M. T. & deHaseth, P. L. Mechanism of bacterial transcription initiation: RNA polymerase - promoter binding, isomerization to initiation-competent open complexes, and initiation of RNA synthesis. J. Mol. Biol.412, 754–771 (2011). CASPubMedPubMed CentralGoogle Scholar
  37. Buc, H. & McClure, W. R. Kinetics of open complex formation between Escherichia coli RNA polymerase and the lac UV5 promoter. Evidence for a sequential mechanism involving three steps. Biochemistry24, 2712–2723 (1985). Early evidence that transcription initiation occurs in multiple steps. CASPubMedGoogle Scholar
  38. Roe, J.-H., Burgess, R. R. & Record, M. T. Temperature dependence of the rate constants of the Escherichia coli RNA polymerase-λPR promoter interaction: assignment of the kinetic steps corresponding to protein conformational change and DNA opening. J. Mol. Biol.184, 441–453 (1985). Early kinetic probing of the transcription rate constants that set the stage for future kinetic studies of promoter melting by RNAP. CASPubMedGoogle Scholar
  39. Ruff, E. F., Record, M. T. & Artsimovitch, I. Initial events in bacterial transcription initiation. Biomolecules5, 1035–1062 (2015). CASPubMedPubMed CentralGoogle Scholar
  40. Hubin, E. A. et al. Structure and function of the mycobacterial transcription initiation complex with the essential regulator RbpA. eLife6, e22520 (2017). One of the first structures of mycobacterial RNAP with detailed quantitative analysis comparing the kinetics of open promoter formation by mycobacteria andE. coliRNAPs and the effects of transcription factors on this process. PubMedPubMed CentralGoogle Scholar
  41. Rammohan, J. et al. CarD stabilizes mycobacterial open complexes via a two-tiered kinetic mechanism. Nucleic Acids Res.43, 3272–3285 (2015). CASPubMedPubMed CentralGoogle Scholar
  42. Haugen, S. P., Ross, W. & Gourse, R. L. Advances in bacterial promoter recognition and its control by factors that do not bind DNA. Nat. Rev. Microbiol.6, 507–519 (2008). CASPubMedPubMed CentralGoogle Scholar
  43. Whipple, F. W. & Sonenshein, A. L. Mechanism of initiation of transcription by Bacillus subtilis RNA polymerase at several promoters. J. Mol. Biol.223, 399–414 (1992). CASPubMedGoogle Scholar
  44. Schroeder, L. A. & deHaseth, P. L. Mechanistic differences in promoter DNA melting by Thermus aquaticus and Escherichia coli RNA polymerases. J. Biol. Chem.280, 17422–17429 (2005). CASPubMedGoogle Scholar
  45. Davis, E., Chen, J., Leon, K., Darst, S. A. & Campbell, E. A. Mycobacterial RNA polymerase forms unstable open promoter complexes that are stabilized by CarD. Nucleic Acids Res.43, 433–445 (2015). CASPubMedGoogle Scholar
  46. Boyaci, H., Chen, J., Jansen, R., Darst, S. A. & Campbell, E. A. Structures of an RNA polymerase promoter melting intermediate elucidate DNA unwinding. Nature565, 382–385 (2019). First structural snapshot of an RNAP promoter melting intermediate. CASPubMedPubMed CentralGoogle Scholar
  47. Boyaci, H. et al. Fidaxomicin jams Mycobacterium tuberculosis RNA polymerase motions needed for initiation via RbpA contacts. eLife7, e34823 (2018). PubMedPubMed CentralGoogle Scholar
  48. Chen, J. et al. Stepwise promoter melting by bacterial RNA polymerase. Mol. Cell78, 275–288.e6 (2020). Captured structural snapshots of promoter melting intermediates that span the RNAP closed complex to the final transcriptionally competent open promoter complex. CASPubMedPubMed CentralGoogle Scholar
  49. McClure, W. R., Cech, C. L. & Johnston, D. E. A steady state assay for the RNA polymerase initiation reaction. J. Biol. Chem.253, 8941–8948 (1978). CASPubMedGoogle Scholar
  50. Carpousis, A. J. & Gralla, J. D. Cycling of ribonucleic acid polymerase to produce oligonucleotides during initiation in vitro at the lac UV5 promoter. Biochemistry19, 3245–3253 (1980). CASPubMedGoogle Scholar
  51. Goldman, S. R., Ebright, R. H. & Nickels, B. E. Direct detection of abortive RNA transcripts in vivo. Science324, 927 (2009). CASPubMedPubMed CentralGoogle Scholar
  52. Hsu, L. M., Vo, N. V., Kane, C. M. & Chamberlin, M. J. In vitro studies of transcript initiation by Escherichia coli RNA polymerase. 1. RNA chain initiation, abortive initiation, and promoter escape at three bacteriophage promoters. Biochemistry42, 3777–3786 (2003). CASPubMedGoogle Scholar
  53. Hsu, L. M. Monitoring abortive initiation. Methods47, 25–36 (2009). CASPubMedGoogle Scholar
  54. Kapanidis, A. N. et al. Initial transcription by RNA polymerase proceeds through a DNA-scrunching mechanism. Science314, 1144–1147 (2006). PubMedPubMed CentralGoogle Scholar
  55. Revyakin, A., Liu, C., Ebright, R. H. & Strick, T. R. Abortive initiation and productive initiation by RNA polymerase involve DNA scrunching. Science314, 1139–1143 (2006). Single-molecule studies revealing the role of scrunching in abortive initiation and promoter escape. CASPubMedPubMed CentralGoogle Scholar
  56. Winkelman, J. T. et al. Crosslink mapping at amino acid-base resolution reveals the path of scrunched DNA in initial transcribing complexes. Mol. Cell59, 768–780 (2015). CASPubMedPubMed CentralGoogle Scholar
  57. Duchi, D. et al. RNA polymerase pausing during initial transcription. Mol. Cell63, 939–950 (2016). CASPubMedPubMed CentralGoogle Scholar
  58. Kulbachinskiy, A. & Mustaev, A. Region 3.2 of the sigma subunit contributes to the binding of the 3’-initiating nucleotide in the RNA polymerase active center and facilitates promoter clearance during initiation. J. Biol. Chem.281, 18273–18276 (2006). CASPubMedGoogle Scholar
  59. Pupov, D., Kuzin, I., Bass, I. & Kulbachinskiy, A. Distinct functions of the RNA polymerase σ subunit region 3.2 in RNA priming and promoter escape. Nucleic Acids Res.42, 4494–4504 (2014). CASPubMedPubMed CentralGoogle Scholar
  60. Samanta, S. & Martin, C. T. Insights into the mechanism of initial transcription in Escherichia coli RNA polymerase. J. Biol. Chem.288, 31993–32003 (2013). CASPubMedPubMed CentralGoogle Scholar
  61. Li, L., Molodtsov, V., Lin, W., Ebright, R. H. & Zhang, Y. RNA extension drives a stepwise displacement of an initiation-factor structural module in initial transcription. Proc. Natl Acad. Sci. USA117, 5801–5809 (2020). CASPubMedPubMed CentralGoogle Scholar
  62. Dulin, D. et al. Pausing controls branching between productive and non-productive pathways during initial transcription in bacteria. Nat. Commun.9, 1478 (2018). PubMedPubMed CentralGoogle Scholar
  63. Jensen, D., Manzano, A. R., Rammohan, J., Stallings, C. L. & Galburt, E. A. CarD and RbpA modify the kinetics of initial transcription and slow promoter escape of the Mycobacterium tuberculosis RNA polymerase. Nucleic Acids Res.47, 6685–6698 (2019). CASPubMedPubMed CentralGoogle Scholar
  64. Mooney, R. A., Darst, S. A. & Landick, R. Sigma and RNA polymerase: an on-again, off-again relationship? Mol. Cell20, 335–345 (2005). CASPubMedGoogle Scholar
  65. Harden, T. T. et al. Bacterial RNA polymerase can retain σ70 throughout transcription. Proc. Natl Acad. Sci. USA113, 602–607 (2016). CASPubMedPubMed CentralGoogle Scholar
  66. Mukhopadhyay, J. et al. Translocation of σ70 with RNA polymerase during transcription: fluorescence resonance energy transfer assay for movement relative to DNA. Cell106, 453–463 (2001). CASPubMedGoogle Scholar
  67. Bar-Nahum, G. & Nudler, E. Isolation and characterization of sigma(70)-retaining transcription elongation complexes from Escherichia coli. Cell106, 443–451 (2001). CASPubMedGoogle Scholar
  68. Kapanidis, A. N. et al. Retention of transcription initiation factor sigma70 in transcription elongation: single-molecule analysis. Mol. Cell20, 347–356 (2005). CASPubMedGoogle Scholar
  69. Mooney, R. A. & Landick, R. Tethering sigma70 to RNA polymerase reveals high in vivo activity of sigma factors and sigma70-dependent pausing at promoter-distal locations. Genes Dev.17, 2839–2851 (2003). CASPubMedPubMed CentralGoogle Scholar
  70. Ring, B. Z., Yarnell, W. S. & Roberts, J. W. Function of E. coli RNA polymerase sigma factor sigma 70 in promoter-proximal pausing. Cell86, 485–493 (1996). CASPubMedGoogle Scholar
  71. Davis, M. C., Kesthely, C. A., Franklin, E. A. & MacLellan, S. R. The essential activities of the bacterial sigma factor. Can. J. Microbiol.63, 89–99 (2017). CASPubMedGoogle Scholar
  72. Gross, C. A. et al. The functional and regulatory roles of sigma factors in transcription. Cold Spring Harb. Symp. Quant. Biol.63, 141–155 (1998). CASPubMedGoogle Scholar
  73. Losick, R. & Pero, J. Cascades of sigma factors. Cell25, 582–584 (1981). CASPubMedGoogle Scholar
  74. Paget, M. Bacterial sigma factors and anti-sigma factors: structure, function and distribution. Biomolecules5, 1245–1265 (2015). CASPubMedPubMed CentralGoogle Scholar
  75. Staroń, A. et al. The third pillar of bacterial signal transduction: classification of the extracytoplasmic function (ECF) σ factor protein family: ECF σ factor classification. Mol. Microbiol.74, 557–581 (2009). A comparative genomic analysis revealing the range and diversity of the ECF σ-factor in bacteria. PubMedGoogle Scholar
  76. Hughes, K. T. & Mathee, K. The anti-sigma factors. Annu. Rev. Microbiol.52, 231–286 (1998). CASPubMedGoogle Scholar
  77. Campbell, E. A. et al. A conserved structural module regulates transcriptional responses to diverse stress signals in bacteria. Mol. Cell27, 793–805 (2007). CASPubMedPubMed CentralGoogle Scholar
  78. Maillard, A. P. et al. The crystal structure of the anti-σ factor CnrY in complex with the σ factor CnrH shows a new structural class of anti-σ factors targeting extracytoplasmic function σ factors. J. Mol. Biol.426, 2313–2327 (2014). CASPubMedGoogle Scholar
  79. Sineva, E., Savkina, M. & Ades, S. E. Themes and variations in gene regulation by extracytoplasmic function (ECF) sigma factors. Curr. Opin. Microbiol.36, 128–137 (2017). CASPubMedPubMed CentralGoogle Scholar
  80. Campbell, E. A. et al. Crystal structure of the Bacillus stearothermophilus anti-sigma factor SpoIIAB with the sporulation sigma factor sigmaF. Cell108, 795–807 (2002). CASPubMedGoogle Scholar
  81. Campbell, E. A. et al. Crystal structure of Escherichia coli sigmaE with the cytoplasmic domain of its anti-sigma RseA. Mol. Cell11, 1067–1078 (2003). CASPubMedGoogle Scholar
  82. Sorenson, M. K., Ray, S. S. & Darst, S. A. Crystal structure of the flagellar σ/anti-σ complex σ28/FlgM reveals an intact σ factor in an inactive conformation. Mol. Cell14, 127–138 (2004). CASPubMedGoogle Scholar
  83. Schumacher, M. A. et al. The crystal structure of the RsbN–σBldN complex from Streptomyces venezuelae defines a new structural class of anti-σ factor. Nucleic Acids Res.46, 7405–7417 (2018). CASPubMedPubMed CentralGoogle Scholar
  84. Gallagher, K. A. et al. c-di-GMP arms an anti-σ to control progression of multicellular differentiation in streptomyces. Mol. Cell77, 586–599.e6 (2020). CASPubMedPubMed CentralGoogle Scholar
  85. Shukla, J., Gupta, R., Thakur, K. G., Gokhale, R. & Gopal, B. Structural basis for the redox sensitivity of the Mycobacterium tuberculosis SigK-RskA σ-anti-σ complex. Acta Crystallogr. D Biol. Crystallogr.70, 1026–1036 (2014). CASPubMedGoogle Scholar
  86. Devkota, S. R., Kwon, E., Ha, S. C., Chang, H. W. & Kim, D. Y. Structural insights into the regulation of Bacillus subtilis SigW activity by anti-sigma RsiW. PLoS ONE12, e0174284 (2017). PubMedPubMed CentralGoogle Scholar
  87. Herrou, J., Rotskoff, G., Luo, Y., Roux, B. & Crosson, S. Structural basis of a protein partner switch that regulates the general stress response of α-proteobacteria. Proc. Natl Acad. Sci. USA109, 7973–7973 (2012). CASGoogle Scholar
  88. Campagne, S. et al. Structural basis for sigma factor mimicry in the general stress response of Alphaproteobacteria. Proc. Natl Acad. Sci. USA109, E1405–E1414 (2012). CASPubMedPubMed CentralGoogle Scholar
  89. Campbell, E. A., Westblade, L. F. & Darst, S. A. Regulation of bacterial RNA polymerase sigma factor activity: a structural perspective. Curr. Opin. Microbiol.11, 121–127 (2008). CASPubMedPubMed CentralGoogle Scholar
  90. Patikoglou, G. A. et al. Crystal structure of the Escherichia coli regulator of sigma70, Rsd, in complex with sigma70 domain 4. J. Mol. Biol.372, 649–659 (2007). CASPubMedPubMed CentralGoogle Scholar
  91. Shi, J. et al. Structural basis of σ appropriation. Nucleic Acids Res.47, 9423–9432 (2019). CASPubMedPubMed CentralGoogle Scholar
  92. Hu, Y., Morichaud, Z., Chen, S., Leonetti, J.-P. & Brodolin, K. Mycobacterium tuberculosis RbpA protein is a new type of transcriptional activator that stabilizes the σ A -containing RNA polymerase holoenzyme. Nucleic Acids Res.40, 6547–6557 (2012). CASPubMedPubMed CentralGoogle Scholar
  93. Bortoluzzi, A. et al. Mycobacterium tuberculosis RNA polymerase-binding protein A (RbpA) and its interactions with sigma factors. J. Biol. Chem.288, 14438–14450 (2013). CASPubMedPubMed CentralGoogle Scholar
  94. Tabib-Salazar, A. et al. The actinobacterial transcription factor RbpA binds to the principal sigma subunit of RNA polymerase. Nucleic Acids Res.41, 5679–5691 (2013). CASPubMedPubMed CentralGoogle Scholar
  95. Vishwakarma, R. K. et al. Single-molecule analysis reveals the mechanism of transcription activation in M. tuberculosis. Sci. Adv.4, eaao5498 (2018). PubMedPubMed CentralGoogle Scholar
  96. Hubin, E. A. et al. Structural, functional, and genetic analyses of the actinobacterial transcription factor RbpA. Proc. Natl Acad. Sci. USA112, 7171–7176 (2015). CASPubMedPubMed CentralGoogle Scholar
  97. Haakonsen, D. L., Yuan, A. H. & Laub, M. T. The bacterial cell cycle regulator GcrA is a σ70 cofactor that drives gene expression from a subset of methylated promoters. Genes Dev.29, 2272–2286 (2015). CASPubMedPubMed CentralGoogle Scholar
  98. Holtzendorff, J. et al. Oscillating global regulators control the genetic circuit driving a bacterial cell cycle. Science304, 983–987 (2004). CASPubMedGoogle Scholar
  99. Wu, X. et al. Structural insights into the unique mechanism of transcription activation by Caulobacter crescentus GcrA. Nucleic Acids Res.46, 3245–3256 (2018). CASPubMedPubMed CentralGoogle Scholar
  100. Gaal, T., Mandel, M. J., Silhavy, T. J. & Gourse, R. L. Crl facilitates RNA polymerase holoenzyme formation. J. Bacteriol.188, 7966–7970 (2006). CASPubMedPubMed CentralGoogle Scholar
  101. Banta, A. B. et al. Key features of σ S required for specific recognition by Crl, a transcription factor promoting assembly of RNA polymerase holoenzyme. Proc. Natl Acad. Sci. USA110, 15955–15960 (2013). CASPubMedPubMed CentralGoogle Scholar
  102. Cartagena, A. J. et al. Structural basis for transcription activation by Crl through tethering of σS and RNA polymerase. Proc. Natl Acad. Sci. USA116, 18923–18927 (2019). CASPubMedPubMed CentralGoogle Scholar
  103. Xu, J. et al. Crl activates transcription by stabilizing active conformation of the master stress transcription initiation factor. eLife8, e50928 (2019). CASPubMedPubMed CentralGoogle Scholar
  104. Wassarman, K. M. & Storz, G. 6S RNA regulates E. coli RNA polymerase activity. Cell101, 613–623 (2000). CASPubMedGoogle Scholar
  105. Trotochaud, A. E. & Wassarman, K. M. A highly conserved 6S RNA structure is required for regulation of transcription. Nat. Struct. Mol. Biol.12, 313–319 (2005). CASPubMedGoogle Scholar
  106. Barrick, J. E. 6S RNA is a widespread regulator of eubacterial RNA polymerase that resembles an open promoter. RNA11, 774–784 (2005). CASPubMedPubMed CentralGoogle Scholar
  107. Wassarman, K. M. in Regulating with RNA in Bacteria and Archaea Ch. 20 (eds Storz, G. & Papenfort, K.) 355–367 (ASM Press, 2019).
  108. Wassarman, K. M. & Saecker, R. M. Synthesis-mediated release of a small RNA inhibitor of RNA polymerase. Science314, 1601–1603 (2006). Discovery that 6S RNA serves as a functional template for transcription initiation by RNAP. CASPubMedGoogle Scholar
  109. Chen, J. et al. 6S RNA mimics B-form DNA to regulate Escherichia coli RNA polymerase. Mol. Cell68, 388–397.e6 (2017). CASPubMedPubMed CentralGoogle Scholar
  110. Lane, W. J. & Darst, S. A. Molecular evolution of multisubunit RNA polymerases: sequence analysis. J. Mol. Biol.395, 671–685 (2010). CASPubMedGoogle Scholar
  111. Lane, W. J. & Darst, S. A. Molecular evolution of multisubunit RNA polymerases: structural analysis. J. Mol. Biol.395, 686–704 (2010). CASPubMedGoogle Scholar
  112. Artsimovitch, I., Svetlov, V., Murakami, K. S. & Landick, R. Co-overexpression of Escherichia coli RNA polymerase subunits allows isolation and analysis of mutant enzymes lacking lineage-specific sequence insertions. J. Biol. Chem.278, 12344–12355 (2003). CASPubMedGoogle Scholar
  113. Chen, J. et al. E. coli TraR allosterically regulates transcription initiation by altering RNA polymerase conformation. eLife8, e49375 (2019). PubMedPubMed CentralGoogle Scholar
  114. Lin, W. et al. Structural basis of Mycobacterium tuberculosis transcription and transcription inhibition. Mol. Cell66, 169–179.e8 (2017). CASPubMedPubMed CentralGoogle Scholar
  115. Browning, D. F. & Busby, S. J. W. Local and global regulation of transcription initiation in bacteria. Nat. Rev. Microbiol.14, 638 (2016). CASPubMedGoogle Scholar
  116. Bergkessel, M. et al. The dormancy-specific regulator, SutA, is intrinsically disordered and modulates transcription initiation in Pseudomonas aeruginosa. Mol. Microbiol.112, 992–1009 (2019). CASPubMedPubMed CentralGoogle Scholar
  117. Molodtsov, V. et al. Allosteric effector ppGpp potentiates the inhibition of transcript initiation by DksA. Mol. Cell69, 828–839.e5 (2018). CASPubMedPubMed CentralGoogle Scholar
  118. Rutherford, S. T., Villers, C. L., Lee, J.-H., Ross, W. & Gourse, R. L. Allosteric control of Escherichia coli rRNA promoter complexes by DksA. Genes Dev.23, 236–248 (2009). CASPubMedPubMed CentralGoogle Scholar
  119. Gopalkrishnan, S., Ross, W., Chen, A. Y. & Gourse, R. L. TraR directly regulates transcription initiation by mimicking the combined effects of the global regulators DksA and ppGpp. Proc. Natl Acad. Sci. USA114, E5539–E5548 (2017). CASPubMedPubMed CentralGoogle Scholar
  120. Srivastava, D. B. et al. Structure and function of CarD, an essential mycobacterial transcription factor. Proc. Natl Acad. Sci. USA110, 12619–12624 (2013). CASPubMedPubMed CentralGoogle Scholar
  121. Bae, B. et al. CarD uses a minor groove wedge mechanism to stabilize the RNA polymerase open promoter complex. eLife4, e08505 (2015). PubMed CentralGoogle Scholar
  122. Venugopal, A. A. & Johnson, S. Fidaxomicin: a novel macrocyclic antibiotic approved for treatment of Clostridium difficile infection. Clin. Infect. Dis.54, 568–574 (2012). PubMedGoogle Scholar
  123. Srivastava, A. et al. New target for inhibition of bacterial RNA polymerase: ‘switch region’. Curr. Opin. Microbiol.14, 532–543 (2011). CASPubMedPubMed CentralGoogle Scholar
  124. Lin, W. et al. Structural basis of transcription inhibition by fidaxomicin (lipiarmycin A3). Mol. Cell70, 60–70.e16 (2018). CASPubMedPubMed CentralGoogle Scholar
  125. Belogurov, G. A. et al. Transcription inactivation through local refolding of the RNA polymerase structure. Nature457, 332–335 (2009). CASPubMedGoogle Scholar
  126. Mukhopadhyay, J. et al. The RNA polymerase “switch region” is a target for inhibitors. Cell135, 295–307 (2008). CASPubMedPubMed CentralGoogle Scholar
  127. Peek, J. et al. Rifamycin congeners kanglemycins are active against rifampicin-resistant bacteria via a distinct mechanism. Nat. Commun.9, 4147 (2018). PubMedPubMed CentralGoogle Scholar
  128. Mosaei, H. et al. Mode of action of kanglemycin A, an ansamycin natural product that is active against rifampicin-resistant Mycobacterium tuberculosis. Mol. Cell72, 263–274.e5 (2018). CASPubMedPubMed CentralGoogle Scholar
  129. Maffioli, S. I. et al. Antibacterial nucleoside-analog inhibitor of bacterial RNA polymerase. Cell169, 1240–1248.e23 (2017). CASPubMedPubMed CentralGoogle Scholar
  130. Zhang, Y. et al. GE23077 binds to the RNA polymerase ‘i’ and ‘i+1’ sites and prevents the binding of initiating nucleotides. eLife3, e02450 (2014). PubMedPubMed CentralGoogle Scholar
  131. McClure, W. R. On the mechanism of streptolydigin inhibition of Escherichia coli RNA polymerase. J. Biol. Chem.255, 1610–1616 (1980). CASPubMedGoogle Scholar
  132. Cassani, G., Burgess, R. R., Goodman, H. M. & Gold, L. Inhibition of RNA polymerase by streptolydigin. Nat. New Biol.230, 197–200 (1971). CASPubMedGoogle Scholar
  133. Siddhikol, C., Erbstoeszer, J. W. & Weisblum, B. Mode of action of streptolydigin. J. Bacteriol.99, 151–155 (1969). CASPubMedPubMed CentralGoogle Scholar
  134. Temiakov, D. et al. Structural basis of transcription inhibition by antibiotic streptolydigin. Mol. Cell19, 655–666 (2005). CASPubMedGoogle Scholar
  135. Tuske, S. et al. Inhibition of bacterial RNA polymerase by streptolydigin: stabilization of a straight-bridge-helix active-center conformation. Cell122, 541–552 (2005). CASPubMedPubMed CentralGoogle Scholar
  136. Artsimovitch, I., Chu, C., Lynch, A. S. & Landick, R. A new class of bacteria RNA polymerase inhibitor affects nucleotide addition. Science302, 650–654 (2003). CASPubMedGoogle Scholar
  137. Bae, B. et al. CBR antimicrobials inhibit RNA polymerase via at least two bridge-helix cap-mediated effects on nucleotide addition. Proc. Natl Acad. Sci. USA112, E4178 (2015). CASPubMedPubMed CentralGoogle Scholar
  138. Feng, Y. et al. Structural basis of transcription inhibition by CBR hydroxamidines and CBR pyrazoles. Structure23, 1470–1481 (2015). CASPubMedPubMed CentralGoogle Scholar
  139. Degen, D. et al. Transcription inhibition by the depsipeptide antibiotic salinamide A. eLife3, e02451 (2014). PubMedPubMed CentralGoogle Scholar
  140. McClure, W. R. & Cech, C. L. On the mechanism of rifampicin inhibition of RNA synthesis. J. Biol. Chem.253, 8949–8956 (1978). CASPubMedGoogle Scholar
  141. Campbell, E. a. E. A. et al. Structural mechanism for rifampicin inhibition of bacterial RNA polymerase. Cell104, 901–912 (2001). Structure of RNAP bound to the clinically used antibiotic rifampicin revealing the basis of inhibition. CASPubMedGoogle Scholar
  142. Campbell, E. A. et al. Structural, functional, and genetic analysis of sorangicin inhibition of bacterial RNA polymerase. EMBO J.24, 674–682 (2005). CASPubMedPubMed CentralGoogle Scholar
  143. Darst, S. A., Kubalek, E. W. & Kornberg, R. D. Three-dimensional structure of Escherichia coli RNA polymerase holoenzyme determined by electron crystallography. Nature340, 730–732 (1989). CASPubMedGoogle Scholar
  144. Gnatt, A. L., Cramer, P., Fu, J., Bushnell, D. A. & Kornberg, R. D. Structural basis of transcription: an RNA polymerase II elongation complex at 3.3 Å resolution. Science292, 1876–1882 (2001). CASPubMedGoogle Scholar
  145. Murakami, K. S. X-ray crystal structure of Escherichia coli RNA polymerase σ70 holoenzyme. J. Biol. Chem.288, 9126–9134 (2013). CASPubMedPubMed CentralGoogle Scholar
  146. Bae, B. et al. Phage T7 Gp2 inhibition of Escherichia coli RNA polymerase involves misappropriation of 70 domain 1.1. Proc. Natl Acad. Sci. USA110, 19772–19777 (2013). CASPubMedPubMed CentralGoogle Scholar
  147. Zuo, Y., Wang, Y. & Steitz, T. A. The mechanism of E. coli RNA polymerase regulation by ppGpp is suggested by the structure of their complex. Mol. Cell50, 430–436 (2013). CASPubMedPubMed CentralGoogle Scholar
  148. Plaschka, C. et al. Transcription initiation complex structures elucidate DNA opening. Nature533, 353–358 (2016). CASPubMedGoogle Scholar
  149. Kang, J. Y. et al. Structural basis of transcription arrest by coliphage HK022 Nun in an Escherichia coli RNA polymerase elongation complex. eLife6, e25478 (2017). PubMedPubMed CentralGoogle Scholar
  150. Liu, B., Hong, C., Huang, R. K., Yu, Z. & Steitz, T. A. Structural basis of bacterial transcription activation. Science358, 947–951 (2017). CASPubMedGoogle Scholar
  151. Stevens, A. An inhibitor of host sigma-stimulated core enzyme activity that purified with DNA-dependent RNA polymerase of E. coli following T4 phage infection. Biochem. Biophys. Res. Commun.54, 488–493 (1973). CASPubMedGoogle Scholar
  152. Wiggs, J. L., Gilman, M. Z. & Chamberlin, M. J. Heterogeneity of RNA polymerase in Bacillus subtilis: evidence for an additional sigma factor in vegetative cells. Proc. Natl Acad. Sci. USA78, 2762–2766 (1981). CASPubMedPubMed CentralGoogle Scholar
  153. Haldenwang, W. G. & Losick, R. A modified RNA polymerase transcribes a cloned gene under sporulation control in Bacillus subtilis. Nature282, 256–260 (1979). CASPubMedGoogle Scholar
  154. Buttner, M. J., Smith, A. M. & Bibb, M. J. At least three different RNA polymerase holoenzymes direct transcription of the agarase gene (dagA) of Streptomyces coelicolor A3(2). Cell52, 599–607 (1988). CASPubMedGoogle Scholar
  155. Erickson, J. W. & Gross, C. A. Identification of the sigma E subunit of Escherichia coli RNA polymerase: a second alternate sigma factor involved in high-temperature gene expression. Genes Dev.3, 1462–1471 (1989). CASPubMedGoogle Scholar
  156. Lonetto, M. A., Brown, K. L., Rudd, K. E. & Buttner, M. J. Analysis of the Streptomyces coelicolor sigE gene reveals the existence of a subfamily of eubacterial RNA polymerase σ factors involved in the regulation of extracytoplasmic functions. Proc. Natl Acad. Sci. USA91, 7573–7577 (1994). CASPubMedPubMed CentralGoogle Scholar
  157. Malhotra, A., Severinova, E. & Darst, S. A. Crystal structure of a σ70 subunit fragment from E. coli RNA polymerase. Cell87, 127–136 (1996). CASPubMedGoogle Scholar
  158. Rhodius, V. A. et al. Design of orthogonal genetic switches based on a crosstalk map of σs, anti-σs, and promoters. Mol. Syst. Biol.9, 702 (2013). CASPubMedPubMed CentralGoogle Scholar
  159. Campagne, S., Marsh, M. E., Capitani, G., Vorholt, J. A. & Allain, F. H. T. Structural basis for -10 promoter element melting by environmentally induced sigma factors. Nat. Struct. Mol. Biol.21, 269–276 (2014). CASPubMedGoogle Scholar
  160. Li, L., Fang, C., Zhuang, N., Wang, T. & Zhang, Y. Structural basis for transcription initiation by bacterial ECF σ factors. Nat. Commun.10, 1153 (2019). PubMedPubMed CentralGoogle Scholar
  161. Lin, W. et al. Structural basis of ECF-σ-factor-dependent transcription initiation. Nat. Commun.10, 710 (2019). CASPubMedPubMed CentralGoogle Scholar
  162. Li, S. et al. Structural basis for the recognition of MucA by MucB and AlgU in Pseudomonas aeruginosa. FEBS J.286, 4982–4994 (2019). CASPubMedGoogle Scholar

Acknowledgements

The authors thank R. Landick, S. Darst and R. Froom for helpful discussions and copyediting. They apologize to colleagues whose work could not be cited owing to the scope and space limits of the Review. The authors are grateful for support from NIH grant 2-R01 GM114450 (E.A.C.) and the Charles H. Revson Foundation award CEN5650030 (H.B.).