1887
No metrics data to plot.
The attempt to load metrics for this article has failed.
The attempt to plot a graph for these metrics has failed.

Carbohydrate Utilization in Bacteria: Making the Most Out of Sugars with the Help of Small Regulatory RNAs

MyBook is a cheap paperback edition of the original book and will be sold at uniform, low price.
  • Authors: Svetlana Durica-Mitic*1, Yvonne Göpel*2, Boris Görke3
  • Editors: Gisela Storz4, Kai Papenfort5
  • VIEW AFFILIATIONS HIDE AFFILIATIONS
    Affiliations: 1: Department of Microbiology, Immunobiology and Genetics, Max F. Perutz Laboratories, University of Vienna, Vienna Biocenter, Vienna, Austria; 2: Department of Microbiology, Immunobiology and Genetics, Max F. Perutz Laboratories, University of Vienna, Vienna Biocenter, Vienna, Austria; 3: Department of Microbiology, Immunobiology and Genetics, Max F. Perutz Laboratories, University of Vienna, Vienna Biocenter, Vienna, Austria; 4: Division of Molecular and Cellular Biology, Eunice Kennedy Shriver National Institute of Child Health and Human Development, Bethesda, MD; 5: Department of Biology I, Microbiology, LMU Munich, Martinsried, Germany
    • Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
  • Received 27 November 2017 Accepted 03 January 2018 Published 23 March 2018
  • Boris Görke, [email protected]
image of Carbohydrate Utilization in Bacteria: Making the Most Out of Sugars with the Help of Small Regulatory RNAs
    Preview this microbiology spectrum article:
    Preview Magnify
    Zoom in
    Zoomout

    Carbohydrate Utilization in Bacteria: Making the Most Out of Sugars with the Help of Small Regulatory RNAs, Page 1 of 2

    | /docserver/preview/fulltext/microbiolspec/6/2/RWR-0013-2017-1.gif /docserver/preview/fulltext/microbiolspec/6/2/RWR-0013-2017-2.gif

  • Abstract:

    Survival of bacteria in ever-changing habitats with fluctuating nutrient supplies requires rapid adaptation of their metabolic capabilities. To this end, carbohydrate metabolism is governed by complex regulatory networks including posttranscriptional mechanisms that involve small regulatory RNAs (sRNAs) and RNA-binding proteins. sRNAs limit the response to substrate availability and set the threshold or time required for induction and repression of carbohydrate utilization systems. Carbon catabolite repression (CCR) also involves sRNAs. In , sRNA Spot 42 cooperates with the transcriptional regulator cyclic AMP (cAMP)-receptor protein (CRP) to repress secondary carbohydrate utilization genes when a preferred sugar is consumed. In pseudomonads, CCR operates entirely at the posttranscriptional level, involving RNA-binding protein Hfq and decoy sRNA CrcZ. Moreover, sRNAs coordinate fluxes through central carbohydrate metabolic pathways with carbohydrate availability. In Gram-negative bacteria, the interplay between RNA-binding protein CsrA and its cognate sRNAs regulates glycolysis and gluconeogenesis in response to signals derived from metabolism. Spot 42 and cAMP-CRP jointly downregulate tricarboxylic acid cycle activity when glycolytic carbon sources are ample. In addition, bacteria use sRNAs to reprogram carbohydrate metabolism in response to anaerobiosis and iron limitation. Finally, sRNAs also provide homeostasis of essential anabolic pathways, as exemplified by the hexosamine pathway providing cell envelope precursors. In this review, we discuss the manifold roles of bacterial sRNAs in regulation of carbon source uptake and utilization, substrate prioritization, and metabolism.

  • Citation: Durica-Mitic* S, Göpel* Y, Görke B. 2018. Carbohydrate Utilization in Bacteria: Making the Most Out of Sugars with the Help of Small Regulatory RNAs. Microbiol Spectrum 6(2):RWR-0013-2017. doi:10.1128/microbiolspec.RWR-0013-2017.

References

1. Chubukov V, Gerosa L, Kochanowski K, Sauer U. 2014. Coordination of microbial metabolism. Nat Rev Microbiol 12:327–340. [PubMed]
2. Georg J, Hess WR. 2011. cis-Antisense RNA, another level of gene regulation in bacteria. Microbiol Mol Biol Rev 75:286–300. [PubMed]
3. Storz G, Vogel J, Wassarman KM. 2011. Regulation by small RNAs in bacteria: expanding frontiers. Mol Cell 43:880–891. [PubMed]
4. Sedlyarova N, Shamovsky I, Bharati BK, Epshtein V, Chen J, Gottesman S, Schroeder R, Nudler E. 2016. sRNA-mediated control of transcription termination in E. coli. Cell 167:111–121.e113. doi:10.1016/j.cell.2016.09.004.
5. Vogel J, Luisi BF. 2011. Hfq and its constellation of RNA. Nat Rev Microbiol 9:578–589. [PubMed]
6. Kavita K, de Mets F, Gottesman S. 2017. New aspects of RNA-based regulation by Hfq and its partner sRNAs. Curr Opin Microbiol 42:53–61. [PubMed]
7. Göpel Y, Görke B. 2012. Rewiring two-component signal transduction with small RNAs. Curr Opin Microbiol 15:132–139. [PubMed]
8. Mandin P, Guillier M. 2013. Expanding control in bacteria: interplay between small RNAs and transcriptional regulators to control gene expression. Curr Opin Microbiol 16:125–132. [PubMed]
9. Arraiano CM, Andrade JM, Domingues S, Guinote IB, Malecki M, Matos RG, Moreira RN, Pobre V, Reis FP, Saramago M, Silva IJ, Viegas SC. 2010. The critical role of RNA processing and degradation in the control of gene expression. FEMS Microbiol Rev 34:883–923. [PubMed]
10. Ziebuhr W, Vogel J. 2015. The end is not the end: remnants of tRNA precursors live on to sponge up small regulatory RNAs. Mol Cell 58:389–390. [PubMed]
11. Baumler DJ, Peplinski RG, Reed JL, Glasner JD, Perna NT. 2011. The evolution of metabolic networks of E. coli. BMC Syst Biol 5:182. doi:10.1186/1752-0509-5-182.
12. Müller-Hill B. 1996. The lac Operon: a Short History of a Genetic Paradigm. de Gruyter, Berlin, Germany.
13. Jacob F, Monod J. 1961. Genetic regulatory mechanisms in the synthesis of proteins. J Mol Biol 3:318–356. [PubMed]
14. Mizuno T, Chou MY, Inouye M. 1984. A unique mechanism regulating gene expression: translational inhibition by a complementary RNA transcript (micRNA). Proc Natl Acad Sci U S A 81:1966–1970. [PubMed]
15. Monod J. 1942. Recherches sur la Croissance des Cultures Bacteriennes. Hermann et Cie, Paris, France.
16. Deutscher J, Francke C, Postma PW. 2006. How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria. Microbiol Mol Biol Rev 70:939–1031. [PubMed]
17. Eisenreich W, Rudel T, Heesemann J, Goebel W. 2017. To eat and to be eaten: mutual metabolic adaptations of immune cells and intracellular bacterial pathogens upon infection. Front Cell Infect Microbiol 7:316. doi:10.3389/fcimb.2017.00316.
18. Jahreis K, Pimentel-Schmitt EF, Brückner R, Titgemeyer F. 2008. Ins and outs of glucose transport systems in eubacteria. FEMS Microbiol Rev 32:891–907. [PubMed]
19. Irani MH, Maitra PK. 1977. Properties of Escherichia coli mutants deficient in enzymes of glycolysis. J Bacteriol 132:398–410. [PubMed]
20. Kadner RJ, Murphy GP, Stephens CM. 1992. Two mechanisms for growth inhibition by elevated transport of sugar phosphates in Escherichia coli. J Gen Microbiol 138:2007–2014. [PubMed]
21. Kimata K, Tanaka Y, Inada T, Aiba H. 2001. Expression of the glucose transporter gene, ptsG, is regulated at the mRNA degradation step in response to glycolytic flux in Escherichia coli. EMBO J 20:3587–3595. [PubMed]
22. Morita T, El-Kazzaz W, Tanaka Y, Inada T, Aiba H. 2003. Accumulation of glucose 6-phosphate or fructose 6-phosphate is responsible for destabilization of glucose transporter mRNA in Escherichia coli. J Biol Chem 278:15608–15614. [PubMed]
23. Vanderpool CK, Gottesman S. 2004. Involvement of a novel transcriptional activator and small RNA in post-transcriptional regulation of the glucose phosphoenolpyruvate phosphotransferase system. Mol Microbiol 54:1076–1089. [PubMed]
24. Vanderpool CK, Gottesman S. 2007. The novel transcription factor SgrR coordinates the response to glucose-phosphate stress. J Bacteriol 189:2238–2248. [PubMed]
25. Kawamoto H, Morita T, Shimizu A, Inada T, Aiba H. 2005. Implication of membrane localization of target mRNA in the action of a small RNA: mechanism of post-transcriptional regulation of glucose transporter in Escherichia coli. Genes Dev 19:328–338. [PubMed]
26. Morita T, Maki K, Aiba H. 2005. RNase E-based ribonucleoprotein complexes: mechanical basis of mRNA destabilization mediated by bacterial noncoding RNAs. Genes Dev 19:2176–2186. [PubMed]
27. Morita T, Mochizuki Y, Aiba H. 2006. Translational repression is sufficient for gene silencing by bacterial small noncoding RNAs in the absence of mRNA destruction. Proc Natl Acad Sci U S A 103:4858–4863. [PubMed]
28. Wadler CS, Vanderpool CK. 2007. A dual function for a bacterial small RNA: SgrS performs base pairing-dependent regulation and encodes a functional polypeptide. Proc Natl Acad Sci U S A 104:20454–20459. [PubMed]
29. Lloyd CR, Park S, Fei J, Vanderpool CK. 2017. The small protein SgrT controls transport activity of the glucose-specific phosphotransferase system. J Bacteriol 199:e00869-16. doi:10.1128/JB.00869-16.
30. Kosfeld A, Jahreis K. 2012. Characterization of the interaction between the small regulatory peptide SgrT and the EIICB Glc of the glucose-phosphotransferase system of E. coli K-12. Metabolites 2:756–774. [PubMed]
31. Bobrovskyy M, Vanderpool CK. 2016. Diverse mechanisms of post-transcriptional repression by the small RNA regulator of glucose-phosphate stress. Mol Microbiol 99:254–273. [PubMed]
32. Wright PR, Richter AS, Papenfort K, Mann M, Vogel J, Hess WR, Backofen R, Georg J. 2013. Comparative genomics boosts target prediction for bacterial small RNAs. Proc Natl Acad Sci U S A 110:E3487–E3496. [PubMed]
33. Rice JB, Vanderpool CK. 2011. The small RNA SgrS controls sugar-phosphate accumulation by regulating multiple PTS genes. Nucleic Acids Res 39:3806–3819. [PubMed]
34. Rice JB, Balasubramanian D, Vanderpool CK. 2012. Small RNA binding-site multiplicity involved in translational regulation of a polycistronic mRNA. Proc Natl Acad Sci U S A 109:E2691–E2698. [PubMed]
35. Papenfort K, Sun Y, Miyakoshi M, Vanderpool CK, Vogel J. 2013. Small RNA-mediated activation of sugar phosphatase mRNA regulates glucose homeostasis. Cell 153:426–437. [PubMed]
36. Sun Y, Vanderpool CK. 2013. Physiological consequences of multiple-target regulation by the small RNA SgrS in Escherichia coli. J Bacteriol 195:4804–4815. [PubMed]
37. Maleki S, Hrudikova R, Zotchev SB, Ertesvåg H. 2016. Identification of a new phosphatase enzyme potentially involved in the sugar phosphate stress response in Pseudomonas fluorescens. Appl Environ Microbiol 83:e02361-16. doi:10.1128/AEM.02361-16.
38. Tchieu JH, Norris V, Edwards JS, Saier MH, Jr. 2001. The complete phosphotransferase system in Escherichia coli. J Mol Microbiol Biotechnol 3:329–346. [PubMed]
39. Reichenbach B, Breustedt DA, Stülke J, Rak B, Görke B. 2007. Genetic dissection of specificity determinants in the interaction of HPr with enzymes II of the bacterial phosphoenolpyruvate:sugar phosphotransferase system in Escherichia coli. J Bacteriol 189:4603–4613. [PubMed]
40. Richards GR, Patel MV, Lloyd CR, Vanderpool CK. 2013. Depletion of glycolytic intermediates plays a key role in glucose-phosphate stress in Escherichia coli. J Bacteriol 195:4816–4825. [PubMed]
41. Wadler CS, Vanderpool CK. 2009. Characterization of homologs of the small RNA SgrS reveals diversity in function. Nucleic Acids Res 37:5477–5485. [PubMed]
42. Horler RS, Vanderpool CK. 2009. Homologs of the small RNA SgrS are broadly distributed in enteric bacteria but have diverged in size and sequence. Nucleic Acids Res 37:5465–5476. [PubMed]
43. Görke B, Stülke J. 2008. Carbon catabolite repression in bacteria: many ways to make the most out of nutrients. Nat Rev Microbiol 6:613–624. [PubMed]
44. Lindner SN, Petrov DP, Hagmann CT, Henrich A, Krämer R, Eikmanns BJ, Wendisch VF, Seibold GM. 2013. Phosphotransferase system-mediated glucose uptake is repressed in phosphoglucoisomerase-deficient Corynebacterium glutamicum strains. Appl Environ Microbiol 79:2588–2595. [PubMed]
45. Landmann JJ, Busse RA, Latz JH, Singh KD, Stülke J, Görke B. 2011. Crh, the paralogue of the phosphocarrier protein HPr, controls the methylglyoxal bypass of glycolysis in Bacillus subtilis. Mol Microbiol 82:770–787. [PubMed]
46. Klancher CA, Hayes CA, Dalia AB. 2017. The nucleoid occlusion protein SlmA is a direct transcriptional activator of chitobiose utilization in Vibrio cholerae. PLoS Genet 13:e1006877. doi:10.1371/journal.pgen.1006877.
47. Meibom KL, Li XB, Nielsen AT, Wu CY, Roseman S, Schoolnik GK. 2004. The Vibrio cholerae chitin utilization program. Proc Natl Acad Sci U S A 101:2524–2529. [PubMed]
48. Dalia AB, Lazinski DW, Camilli A. 2014. Identification of a membrane-bound transcriptional regulator that links chitin and natural competence in Vibrio cholerae. mBio 5:e01028-e13. doi:10.1128/mBio.01028-13. [PubMed]
49. Yamamoto S, Izumiya H, Mitobe J, Morita M, Arakawa E, Ohnishi M, Watanabe H. 2011. Identification of a chitin-induced small RNA that regulates translation of the tfoX gene, encoding a positive regulator of natural competence in Vibrio cholerae. J Bacteriol 193:1953–1965. [PubMed]
50. Borgeaud S, Metzger LC, Scrignari T, Blokesch M. 2015. The type VI secretion system of Vibrio cholerae fosters horizontal gene transfer. Science 347:63–67. [PubMed]
51. Göpel Y, Görke B. 2014. Lies and deception in bacterial gene regulation: the roles of nucleic acid decoys. Mol Microbiol 92:641–647. [PubMed]
52. Bossi L, Figueroa-Bossi N. 2016. Competing endogenous RNAs: a target-centric view of small RNA regulation in bacteria. Nat Rev Microbiol 14:775–784. [PubMed]
53. Plumbridge J, Bossi L, Oberto J, Wade JT, Figueroa-Bossi N. 2014. Interplay of transcriptional and small RNA-dependent control mechanisms regulates chitosugar uptake in Escherichia coli and Salmonella. Mol Microbiol 92:648–658. [PubMed]
54. Figueroa-Bossi N, Valentini M, Malleret L, Fiorini F, Bossi L. 2009. Caught at its own game: regulatory small RNA inactivated by an inducible transcript mimicking its target. Genes Dev 23:2004–2015. [PubMed]
55. Rasmussen AA, Johansen J, Nielsen JS, Overgaard M, Kallipolitis B, Valentin-Hansen P. 2009. A conserved small RNA promotes silencing of the outer membrane protein YbfM. Mol Microbiol 72:566–577. [PubMed]
56. Bossi L, Schwartz A, Guillemardet B, Boudvillain M, Figueroa-Bossi N. 2012. A role for Rho-dependent polarity in gene regulation by a noncoding small RNA. Genes Dev 26:1864–1873. [PubMed]
57. Overgaard M, Johansen J, Møller-Jensen J, Valentin-Hansen P. 2009. Switching off small RNA regulation with trap-mRNA. Mol Microbiol 73:790–800. [PubMed]
58. Suzuki K, Shimizu M, Sasaki N, Ogawa C, Minami H, Sugimoto H, Watanabe T. 2016. Regulation of the chitin degradation and utilization system by the ChiX small RNA in Serratia marcescens 2170. Biosci Biotechnol Biochem 80:376–385. [PubMed]
59. Mustachio LM, Aksit S, Mistry RH, Scheffler R, Yamada A, Liu JM. 2012. The Vibrio cholerae mannitol transporter is regulated posttranscriptionally by the MtlS small regulatory RNA. J Bacteriol 194:598–606. [PubMed]
60. Chang H, Replogle JM, Vather N, Tsao-Wu M, Mistry R, Liu JM. 2015. A cis-regulatory antisense RNA represses translation in Vibrio cholerae through extensive complementarity and proximity to the target locus. RNA Biol 12:136–148. [PubMed]
61. Byer T, Wang J, Zhang MG, Vather N, Blachman A, Visser B, Liu JM. 2017. MtlR negatively regulates mannitol utilization by Vibrio cholerae. Microbiology doi:10.1099/mic.0.000559.
62. Ymele-Leki P, Houot L, Watnick PI. 2013. Mannitol and the mannitol-specific enzyme IIB subunit activate Vibrio cholerae biofilm formation. Appl Environ Microbiol 79:4675–4683. [PubMed]
63. White BA, Lamed R, Bayer EA, Flint HJ. 2014. Biomass utilization by gut microbiomes. Annu Rev Microbiol 68:279–296. [PubMed]
64. Grondin JM, Tamura K, Déjean G, Abbott DW, Brumer H. 2017. Polysaccharide utilization loci: fueling microbial communities. J Bacteriol 199:e00860-16. doi:10.1128/JB.00860-16.
65. Cao Y, Förstner KU, Vogel J, Smith CJ. 2016. cis-Encoded small RNAs, a conserved mechanism for repression of polysaccharide utilization in Bacteroides. J Bacteriol 198:2410–2418. [PubMed]
66. Pudlo NA, Urs K, Kumar SS, German JB, Mills DA, Martens EC. 2015. Symbiotic human gut bacteria with variable metabolic priorities for host mucosal glycans. mBio 6:e01282-e15. doi:10.1128/mBio.01282-15.
67. Aidelberg G, Towbin BD, Rothschild D, Dekel E, Bren A, Alon U. 2014. Hierarchy of non-glucose sugars in Escherichia coli. BMC Syst Biol 8:133. doi:10.1186/s12918-014-0133-z. [PubMed]
68. Rothe FM, Bahr T, Stülke J, Rak B, Görke B. 2012. Activation of Escherichia coli antiterminator BglG requires its phosphorylation. Proc Natl Acad Sci U S A 109:15906–15911. [PubMed]
69. Borirak O, Rolfe MD, de Koning LJ, Hoefsloot HC, Bekker M, Dekker HL, Roseboom W, Green J, de Koster CG, Hellingwerf KJ. 2015. Time-series analysis of the transcriptome and proteome of Escherichia coli upon glucose repression. Biochim Biophys Acta 1854(10 Pt A) :1269–1279.
70. Sonnleitner E, Bläsi U. 2014. Regulation of Hfq by the RNA CrcZ in Pseudomonas aeruginosa carbon catabolite repression. PLoS Genet 10:e1004440. doi:10.1371/journal.pgen.1004440.
71. Beisel CL, Storz G. 2011. Discriminating tastes: physiological contributions of the Hfq-binding small RNA Spot 42 to catabolite repression. RNA Biol 8:766–770. [PubMed]
72. Rice PW, Dahlberg JE. 1982. A gene between polA and glnA retards growth of Escherichia coli when present in multiple copies: physiological effects of the gene for spot 42 RNA. J Bacteriol 152:1196–1210. [PubMed]
73. Polayes DA, Rice PW, Garner MM, Dahlberg JE. 1988. Cyclic AMP-cyclic AMP receptor protein as a repressor of transcription of the spf gene of Escherichia coli. J Bacteriol 170:3110–3114. [PubMed]
74. Møller T, Franch T, Udesen C, Gerdes K, Valentin-Hansen P. 2002. Spot 42 RNA mediates discoordinate expression of the E. coli galactose operon. Genes Dev 16:1696–1706. [PubMed]
75. Beisel CL, Storz G. 2011. The base-pairing RNA spot 42 participates in a multioutput feedforward loop to help enact catabolite repression in Escherichia coli. Mol Cell 41:286–297. [PubMed]
76. Beisel CL, Updegrove TB, Janson BJ, Storz G. 2012. Multiple factors dictate target selection by Hfq-binding small RNAs. EMBO J 31:1961–1974. [PubMed]
77. Melamed S, Peer A, Faigenbaum-Romm R, Gatt YE, Reiss N, Bar A, Altuvia Y, Argaman L, Margalit H. 2016. Global mapping of small RNA-target interactions in bacteria. Mol Cell 63:884–897. [PubMed]
78. Conway T, Cohen PS. 2015. Commensal and pathogenic Escherichia coli metabolism in the gut. Microbiol Spectr 3:MBP-0006-2014. doi:10.1128/microbiolspec.MBP-0006-2014.
79. Wu J, Li Y, Cai Z, Jin Y. 2014. Pyruvate-associated acid resistance in bacteria. Appl Environ Microbiol 80:4108–4113. [PubMed]
80. Møller T, Franch T, Højrup P, Keene DR, Bächinger HP, Brennan RG, Valentin-Hansen P. 2002. Hfq: a bacterial Sm-like protein that mediates RNA-RNA interaction. Mol Cell 9:23–30. [PubMed]
81. Wang X, Ji SC, Jeon HJ, Lee Y, Lim HM. 2015. Two-level inhibition of galK expression by Spot 42: degradation of mRNA mK2 and enhanced transcription termination before the galK gene. Proc Natl Acad Sci U S A 112:7581–7586. [PubMed]
82. Desnoyers G, Massé E. 2012. Noncanonical repression of translation initiation through small RNA recruitment of the RNA chaperone Hfq. Genes Dev 26:726–739. [PubMed]
83. Ellis MJ, Trussler RS, Haniford DB. 2015. Hfq binds directly to the ribosome-binding site of IS 10 transposase mRNA to inhibit translation. Mol Microbiol 96:633–650. [PubMed]
84. Salvail H, Caron MP, Bélanger J, Massé E. 2013. Antagonistic functions between the RNA chaperone Hfq and an sRNA regulate sensitivity to the antibiotic colicin. EMBO J 32:2764–2778. [PubMed]
85. Sharma CM, Papenfort K, Pernitzsch SR, Mollenkopf HJ, Hinton JC, Vogel J. 2011. Pervasive post-transcriptional control of genes involved in amino acid metabolism by the Hfq-dependent GcvB small RNA. Mol Microbiol 81:1144–1165. [PubMed]
86. Durand S, Storz G. 2010. Reprogramming of anaerobic metabolism by the FnrS small RNA. Mol Microbiol 75:1215–1231. [PubMed]
87. Bækkedal C, Haugen P. 2015. The Spot 42 RNA: a regulatory small RNA with roles in the central metabolism. RNA Biol 12:1071–1077. [PubMed]
88. Hansen GA, Ahmad R, Hjerde E, Fenton CG, Willassen NP, Haugen P. 2012. Expression profiling reveals Spot 42 small RNA as a key regulator in the central metabolism of Aliivibrio salmonicida. BMC Genomics 13:37. doi:10.1186/1471-2164-13-37.
89. Livny J, Zhou X, Mandlik A, Hubbard T, Davis BM, Waldor MK. 2014. Comparative RNA-Seq based dissection of the regulatory networks and environmental stimuli underlying Vibrio parahaemolyticus gene expression during infection. Nucleic Acids Res 42:12212–12223. [PubMed]
90. Tanabe T, Miyamoto K, Tsujibo H, Yamamoto S, Funahashi T. 2015. The small RNA Spot 42 regulates the expression of the type III secretion system 1 (T3SS1) chaperone protein VP1682 in Vibrio parahaemolyticus. FEMS Microbiol Lett 362:fnv173. doi:10.1093/femsle/fnv173.
91. Rojo F. 2010. Carbon catabolite repression in Pseudomonas: optimizing metabolic versatility and interactions with the environment. FEMS Microbiol Rev 34:658–684. [PubMed]
92. Moreno R, Hernández-Arranz S, La Rosa R, Yuste L, Madhushani A, Shingler V, Rojo F. 2015. The Crc and Hfq proteins of Pseudomonas putida cooperate in catabolite repression and formation of ribonucleic acid complexes with specific target motifs. Environ Microbiol 17:105–118. [PubMed]
93. Sonnleitner E, Wulf A, Campagne S, Pei XY, Wolfinger MT, Forlani G, Prindl K, Abdou L, Resch A, Allain FH, Luisi BF, Urlaub H, Bläsi U. 2017. Interplay between the catabolite repression control protein Crc, Hfq and RNA in Hfq-dependent translational regulation in Pseudomonas aeruginosa. Nucleic Acids Res doi:10.1093/nar/gkx1245.
94. Sonnleitner E, Abdou L, Haas D. 2009. Small RNA as global regulator of carbon catabolite repression in Pseudomonas aeruginosa. Proc Natl Acad Sci U S A 106:21866–21871. [PubMed]
95. Liu Y, Gokhale CS, Rainey PB, Zhang XX. 2017. Unravelling the complexity and redundancy of carbon catabolic repression in Pseudomonas fluorescens SBW25. Mol Microbiol 105:589–605. [PubMed]
96. Filiatrault MJ, Stodghill PV, Wilson J, Butcher BG, Chen H, Myers CR, Cartinhour SW. 2013. CrcZ and CrcX regulate carbon source utilization in Pseudomonas syringae pathovar tomato strain DC3000. RNA Biol 10:245–255. [PubMed]
97. Valentini M, García-Mauriño SM, Pérez-Martínez I, Santero E, Canosa I, Lapouge K. 2014. Hierarchical management of carbon sources is regulated similarly by the CbrA/B systems in Pseudomonas aeruginosa and Pseudomonas putida. Microbiology 160:2243–2252. [PubMed]
98. Sonnleitner E, Prindl K, Bläsi U. 2017. The Pseudomonas aeruginosa CrcZ RNA interferes with Hfq-mediated riboregulation. PLoS One 12:e0180887. doi:10.1371/journal.pone.0180887.
99. Kochanowski K, Gerosa L, Brunner SF, Christodoulou D, Nikolaev YV, Sauer U. 2017. Few regulatory metabolites coordinate expression of central metabolic genes in Escherichia coli. Mol Syst Biol 13:903. doi:10.15252/msb.20167402.
100. Shimada T, Yamamoto K, Ishihama A. 2011. Novel members of the Cra regulon involved in carbon metabolism in Escherichia coli. J Bacteriol 193:649–659. [PubMed]
101. Saier MH, Jr, Ramseier TM. 1996. The catabolite repressor/activator (Cra) protein of enteric bacteria. J Bacteriol 178:3411–3417. [PubMed]
102. Hogema BM, Arents JC, Bader R, Eijkemans K, Yoshida H, Takahashi H, Aiba H, Postma PW. 1998. Inducer exclusion in Escherichia coli by non-PTS substrates: the role of the PEP to pyruvate ratio in determining the phosphorylation state of enzyme IIA Glc. Mol Microbiol 30:487–498. [PubMed]
103. Bettenbrock K, Sauter T, Jahreis K, Kremling A, Lengeler JW, Gilles ED. 2007. Correlation between growth rates, EIIA Crr phosphorylation, and intracellular cyclic AMP levels in Escherichia coli K-12. J Bacteriol 189:6891–6900. [PubMed]
104. Matsuoka Y, Shimizu K. 2011. Metabolic regulation in Escherichia coli in response to culture environments via global regulators. Biotechnol J 6:1330–1341. [PubMed]
105. Vakulskas CA, Potts AH, Babitzke P, Ahmer BM, Romeo T. 2015. Regulation of bacterial virulence by Csr (Rsm) systems. Microbiol Mol Biol Rev 79:193–224. [PubMed]
106. Potts AH, Vakulskas CA, Pannuri A, Yakhnin H, Babitzke P, Romeo T. 2017. Global role of the bacterial post-transcriptional regulator CsrA revealed by integrated transcriptomics. Nat Commun 8:1596. doi:10.1038/s41467-017-01613-1.
107. Van Assche E, Van Puyvelde S, Vanderleyden J, Steenackers HP. 2015. RNA-binding proteins involved in post-transcriptional regulation in bacteria. Front Microbiol 6:141. doi:10.3389/fmicb.2015.00141.
108. Sabnis NA, Yang H, Romeo T. 1995. Pleiotropic regulation of central carbohydrate metabolism in Escherichia coli via the gene csrA. J Biol Chem 270:29096–29104. [PubMed]
109. Morin M, Ropers D, Letisse F, Laguerre S, Portais JC, Cocaign-Bousquet M, Enjalbert B. 2016. The post-transcriptional regulatory system CSR controls the balance of metabolic pools in upper glycolysis of Escherichia coli. Mol Microbiol 100:686–700. [PubMed]
110. Timmermans J, Van Melderen L. 2009. Conditional essentiality of the csrA gene in Escherichia coli. J Bacteriol 191:1722–1724. [PubMed]
111. Weilbacher T, Suzuki K, Dubey AK, Wang X, Gudapaty S, Morozov I, Baker CS, Georgellis D, Babitzke P, Romeo T. 2003. A novel sRNA component of the carbon storage regulatory system of Escherichia coli. Mol Microbiol 48:657–670. [PubMed]
112. Liu MY, Gui G, Wei B, Preston JF, III, Oakford L, Yüksel U, Giedroc DP, Romeo T. 1997. The RNA molecule CsrB binds to the global regulatory protein CsrA and antagonizes its activity in Escherichia coli. J Biol Chem 272:17502–17510. [PubMed]
113. Chavez RG, Alvarez AF, Romeo T, Georgellis D. 2010. The physiological stimulus for the BarA sensor kinase. J Bacteriol 192:2009–2012. [PubMed]
114. Lawhon SD, Maurer R, Suyemoto M, Altier C. 2002. Intestinal short-chain fatty acids alter Salmonella typhimurium invasion gene expression and virulence through BarA/SirA. Mol Microbiol 46:1451–1464. [PubMed]
115. Vakulskas CA, Leng Y, Abe H, Amaki T, Okayama A, Babitzke P, Suzuki K, Romeo T. 2016. Antagonistic control of the turnover pathway for the global regulatory sRNA CsrB by the CsrA and CsrD proteins. Nucleic Acids Res 44:7896–7910. [PubMed]
116. Suzuki K, Babitzke P, Kushner SR, Romeo T. 2006. Identification of a novel regulatory protein (CsrD) that targets the global regulatory RNAs CsrB and CsrC for degradation by RNase E. Genes Dev 20:2605–2617. [PubMed]
117. Leng Y, Vakulskas CA, Zere TR, Pickering BS, Watnick PI, Babitzke P, Romeo T. 2016. Regulation of CsrB/C sRNA decay by EIIA Glc of the phosphoenolpyruvate: carbohydrate phosphotransferase system. Mol Microbiol 99:627–639. [PubMed]
118. Morin M, Ropers D, Cinquemani E, Portais JC, Enjalbert B, Cocaign-Bousquet M. 2017. The Csr system regulates Escherichia coli fitness by controlling glycogen accumulation and energy levels. mBio 8:e01628-17. doi:10.1128/mBio.01628-17.
119. Pannuri A, Vakulskas CA, Zere T, McGibbon LC, Edwards AN, Georgellis D, Babitzke P, Romeo T. 2016. Circuitry linking the catabolite repression and Csr global regulatory systems of Escherichia coli. J Bacteriol 198:3000–3015. [PubMed]
120. You C, Okano H, Hui S, Zhang Z, Kim M, Gunderson CW, Wang YP, Lenz P, Yan D, Hwa T. 2013. Coordination of bacterial proteome with metabolism by cyclic AMP signalling. Nature 500:301–306. [PubMed]
121. Nuss AM, Schuster F, Kathrin Heroven A, Heine W, Pisano F, Dersch P. 2014. A direct link between the global regulator PhoP and the Csr regulon in Y. pseudotuberculosis through the small regulatory RNA CsrC. RNA Biol 11:580–593. [PubMed]
122. Pannekoek Y, Huis In ’t Veld RA, Schipper K, Bovenkerk S, Kramer G, Brouwer MC, van de Beek D, Speijer D, van der Ende A. 2017. Neisseria meningitidis uses sibling small regulatory RNAs to switch from cataplerotic to anaplerotic metabolism. mBio 8:e02293-16. doi:10.1128/mBio.02293-16.
123. Bauer S, Helmreich J, Zachary M, Kaethner M, Heinrichs E, Rudel T, Beier D. 2017. The sibling sRNAs NgncR_162 and NgncR_163 of Neisseria gonorrhoeae participate in the expression control of metabolic, transport and regulatory proteins. Microbiology 163:1720–1734. [PubMed]
124. Oglesby-Sherrouse AG, Murphy ER. 2013. Iron-responsive bacterial small RNAs: variations on a theme. Metallomics 5:276–286. [PubMed]
125. Salvail H, Massé E. 2012. Regulating iron storage and metabolism with RNA: an overview of posttranscriptional controls of intracellular iron homeostasis. Wiley Interdiscip Rev RNA 3:26–36. [PubMed]
126. Massé E, Gottesman S. 2002. A small RNA regulates the expression of genes involved in iron metabolism in Escherichia coli. Proc Natl Acad Sci U S A 99:4620–4625. [PubMed]
127. Seo SW, Kim D, Latif H, O’Brien EJ, Szubin R, Palsson BO. 2014. Deciphering Fur transcriptional regulatory network highlights its complex role beyond iron metabolism in Escherichia coli. Nat Commun 5:4910. doi:10.1038/ncomms5910.
128. Porcheron G, Dozois CM. 2015. Interplay between iron homeostasis and virulence: Fur and RyhB as major regulators of bacterial pathogenicity. Vet Microbiol 179:2–14. [PubMed]
129. Friedman DB, Stauff DL, Pishchany G, Whitwell CW, Torres VJ, Skaar EP. 2006. Staphylococcus aureus redirects central metabolism to increase iron availability. PLoS Pathog 2:e87. doi:10.1371/journal.ppat.0020087.
130. Jung YS, Kwon YM. 2008. Small RNA ArrF regulates the expression of sodB and feSII genes in Azotobacter vinelandii. Curr Microbiol 57:593–597. [PubMed]
131. de Graef MR, Alexeeva S, Snoep JL, Teixeira de Mattos MJ. 1999. The steady-state internal redox state (NADH/NAD) reflects the external redox state and is correlated with catabolic adaptation in Escherichia coli. J Bacteriol 181:2351–2357. [PubMed]
132. Green J, Paget MS. 2004. Bacterial redox sensors. Nat Rev Microbiol 2:954–966. [PubMed]
133. Sawers G. 1999. The aerobic/anaerobic interface. Curr Opin Microbiol 2:181–187. [PubMed]
134. Constantinidou C, Hobman JL, Griffiths L, Patel MD, Penn CW, Cole JA, Overton TW. 2006. A reassessment of the FNR regulon and transcriptomic analysis of the effects of nitrate, nitrite, NarXL, and NarQP as Escherichia coli K12 adapts from aerobic to anaerobic growth. J Biol Chem 281:4802–4815. [PubMed]
135. Myers KS, Yan H, Ong IM, Chung D, Liang K, Tran F, Keleş S, Landick R, Kiley PJ. 2013. Genome-scale analysis of Escherichia coli FNR reveals complex features of transcription factor binding. PLoS Genet 9:e1003565. doi:10.1371/journal.pgen.1003565.
136. Liu X, De Wulf P. 2004. Probing the ArcA-P modulon of Escherichia coli by whole genome transcriptional analysis and sequence recognition profiling. J Biol Chem 279:12588–12597. [PubMed]
137. Park DM, Akhtar MS, Ansari AZ, Landick R, Kiley PJ. 2013. The bacterial response regulator ArcA uses a diverse binding site architecture to regulate carbon oxidation globally. PLoS Genet 9:e1003839. doi:10.1371/journal.pgen.1003839.
138. Boysen A, Møller-Jensen J, Kallipolitis B, Valentin-Hansen P, Overgaard M. 2010. Translational regulation of gene expression by an anaerobically induced small non-coding RNA in Escherichia coli. J Biol Chem 285:10690–10702.
139. van der Rest ME, Frank C, Molenaar D. 2000. Functions of the membrane-associated and cytoplasmic malate dehydrogenases in the citric acid cycle of Escherichia coli. J Bacteriol 182:6892–6899. [PubMed]
140. Mandin P, Gottesman S. 2010. Integrating anaerobic/aerobic sensing and the general stress response through the ArcZ small RNA. EMBO J 29:3094–3107. [PubMed]
141. Papenfort K, Said N, Welsink T, Lucchini S, Hinton JC, Vogel J. 2009. Specific and pleiotropic patterns of mRNA regulation by ArcZ, a conserved, Hfq-dependent small RNA. Mol Microbiol 74:139–158. [PubMed]
142. Parker A, Cureoglu S, De Lay N, Majdalani N, Gottesman S. 2017. Alternative pathways for Escherichia coli biofilm formation revealed by sRNA overproduction. Mol Microbiol 105:309–325. [PubMed]
143. Chen J, Gottesman S. 2017. Hfq links translation repression to stress-induced mutagenesis in E. coli. Genes Dev 31:1382–1395. [PubMed]
144. Xia H, Yang X, Tang Q, Ye J, Wu H, Zhang H. 2017. EsrE—a yigP locus-encoded transcript—is a 3′ UTR sRNA involved in the respiratory chain of E. coli. Front Microbiol 8:1658. doi:10.3389/fmicb.2017.01658.
145. Isabella VM, Clark VL. 2011. Deep sequencing-based analysis of the anaerobic stimulon in Neisseria gonorrhoeae. BMC Genomics 12:51. doi:10.1186/1471-2164-12-51.
146. Fantappiè L, Oriente F, Muzzi A, Serruto D, Scarlato V, Delany I. 2011. A novel Hfq-dependent sRNA that is under FNR control and is synthesized in oxygen limitation in Neisseria meningitidis. Mol Microbiol 80:507–523. [PubMed]
147. Knapp JS, Clark VL. 1984. Anaerobic growth of Neisseria gonorrhoeae coupled to nitrite reduction. Infect Immun 46:176–181. [PubMed]
148. Barth KR, Isabella VM, Clark VL. 2009. Biochemical and genomic analysis of the denitrification pathway within the genus Neisseria. Microbiology 155:4093–4103. [PubMed]
149. Phillips NJ, Steichen CT, Schilling B, Post DM, Niles RK, Bair TB, Falsetta ML, Apicella MA, Gibson BW. 2012. Proteomic analysis of Neisseria gonorrhoeae biofilms shows shift to anaerobic respiration and changes in nutrient transport and outermembrane proteins. PLoS One 7:e38303. doi:10.1371/journal.pone.0038303.
150. Tanwer P, Bauer S, Heinrichs E, Panda G, Saluja D, Rudel T, Beier D. 2017. Post-transcriptional regulation of target genes by the sRNA FnrS in Neisseria gonorrhoeae. Microbiology 163:1081–1092. [PubMed]
151. Durand S, Braun F, Lioliou E, Romilly C, Helfer AC, Kuhn L, Quittot N, Nicolas P, Romby P, Condon C. 2015. A nitric oxide regulated small RNA controls expression of genes involved in redox homeostasis in Bacillus subtilis. PLoS Genet 11:e1004957. doi:10.1371/journal.pgen.1004957.
152. Geissmann T, Chevalier C, Cros MJ, Boisset S, Fechter P, Noirot C, Schrenzel J, François P, Vandenesch F, Gaspin C, Romby P. 2009. A search for small noncoding RNAs in Staphylococcus aureus reveals a conserved sequence motif for regulation. Nucleic Acids Res 37:7239–7257. [PubMed]
153. Bohn C, Rigoulay C, Chabelskaya S, Sharma CM, Marchais A, Skorski P, Borezée-Durant E, Barbet R, Jacquet E, Jacq A, Gautheret D, Felden B, Vogel J, Bouloc P. 2010. Experimental discovery of small RNAs in Staphylococcus aureus reveals a riboregulator of central metabolism. Nucleic Acids Res 38:6620–6636. [PubMed]
154. Durand S, Braun F, Helfer AC, Romby P, Condon C. 2017. sRNA-mediated activation of gene expression by inhibition of 5′-3′ exonucleolytic mRNA degradation. eLife 6:e23602. doi:10.7554/eLife.23602.
155. Kinkel TL, Roux CM, Dunman PM, Fang FC. 2013. The Staphylococcus aureus SrrAB two-component system promotes resistance to nitrosative stress and hypoxia. mBio 4:e00696-e13. doi:10.1128/mBio.00696-13.
156. Fechter P, Caldelari I, Lioliou E, Romby P. 2014. Novel aspects of RNA regulation in Staphylococcus aureus. FEBS Lett 588:2523–2529. [PubMed]
157. Durand P, Golinelli-Pimpaneau B, Mouilleron S, Badet B, Badet-Denisot MA. 2008. Highlights of glucosamine-6P synthase catalysis. Arch Biochem Biophys 474:302–317. [PubMed]
158. Winkler WC, Nahvi A, Roth A, Collins JA, Breaker RR. 2004. Control of gene expression by a natural metabolite-responsive ribozyme. Nature 428:281–286. [PubMed]
159. Bingaman JL, Zhang S, Stevens DR, Yennawar NH, Hammes-Schiffer S, Bevilacqua PC. 2017. The GlcN6P cofactor plays multiple catalytic roles in the glmS ribozyme. Nat Chem Biol 13:439–445. [PubMed]
160. Collins JA, Irnov I, Baker S, Winkler WC. 2007. Mechanism of mRNA destabilization by the glmS ribozyme. Genes Dev 21:3356–3368. [PubMed]
161. Göpel Y, Lüttmann D, Heroven AK, Reichenbach B, Dersch P, Görke B. 2011. Common and divergent features in transcriptional control of the homologous small RNAs GlmY and GlmZ in Enterobacteriaceae. Nucleic Acids Res 39:1294–1309. [PubMed]
162. Khan MA, Göpel Y, Milewski S, Görke B. 2016. Two small RNAs conserved in Enterobacteriaceae provide intrinsic resistance to antibiotics targeting the cell wall biosynthesis enzyme glucosamine-6-phosphate synthase. Front Microbiol 7:908. doi:10.3389/fmicb.2016.00908.
163. Kalamorz F, Reichenbach B, März W, Rak B, Görke B. 2007. Feedback control of glucosamine-6-phosphate synthase GlmS expression depends on the small RNA GlmZ and involves the novel protein YhbJ in Escherichia coli. Mol Microbiol 65:1518–1533. [PubMed]
164. Reichenbach B, Maes A, Kalamorz F, Hajnsdorf E, Görke B. 2008. The small RNA GlmY acts upstream of the sRNA GlmZ in the activation of glmS expression and is subject to regulation by polyadenylation in Escherichia coli. Nucleic Acids Res 36:2570–2580. [PubMed]
165. Urban JH, Vogel J. 2008. Two seemingly homologous noncoding RNAs act hierarchically to activate glmS mRNA translation. PLoS Biol 6:e64. doi:10.1371/journal.pbio.0060064.
166. Göpel Y, Papenfort K, Reichenbach B, Vogel J, Görke B. 2013. 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. [PubMed]
167. Göpel Y, Khan MA, Görke B. 2016. Domain swapping between homologous bacterial small RNAs dissects processing and Hfq binding determinants and uncovers an aptamer for conditional RNase E cleavage. Nucleic Acids Res 44:824–837. [PubMed]
168. Gonzalez GM, Durica-Mitic S, Hardwick SW, Moncrieffe MC, Resch M, Neumann P, Ficner R, Görke B, Luisi BF. 2017. Structural insights into RapZ-mediated regulation of bacterial amino-sugar metabolism. Nucleic Acids Res 45:10845–10860. [PubMed]
169. Görke B, Vogel J. 2008. Noncoding RNA control of the making and breaking of sugars. Genes Dev 22:2914–2925. [PubMed]
170. Zeng L, Choi SC, Danko CG, Siepel A, Stanhope MJ, Burne RA. 2013. Gene regulation by CcpA and catabolite repression explored by RNA-seq in Streptococcus mutans. PLoS One 8:e60465. doi:10.1371/journal.pone.0060465.
171. Erickson DW, Schink SJ, Patsalo V, Williamson JR, Gerland U, Hwa T. 2017. A global resource allocation strategy governs growth transition kinetics of Escherichia coli. Nature 551:119–123. [PubMed]
172. Westermayer SA, Fritz G, Gutiérrez J, Megerle JA, Weißl MP, Schnetz K, Gerland U, Rädler JO. 2016. Single-cell characterization of metabolic switching in the sugar phosphotransferase system of Escherichia coli. Mol Microbiol 100:472–485. [PubMed]
173. Holmqvist E, Wright PR, Li L, Bischler T, Barquist L, Reinhardt R, Backofen R, Vogel J. 2016. Global RNA recognition patterns of post-transcriptional regulators Hfq and CsrA revealed by UV crosslinking in vivo. EMBO J 35:991–1011. [PubMed]
174. Waters SA, McAteer SP, Kudla G, Pang I, Deshpande NP, Amos TG, Leong KW, Wilkins MR, Strugnell R, Gally DL, Tollervey D, Tree JJ. 2017. Small RNA interactome of pathogenic E. coli revealed through crosslinking of RNase E. EMBO J 36:374–387. [PubMed]
175. Heroven AK, Dersch P. 2014. Coregulation of host-adapted metabolism and virulence by pathogenic yersiniae. Front Cell Infect Microbiol 4:146. doi:10.3389/fcimb.2014.00146.
176. Papenfort K, Vogel J. 2014. Small RNA functions in carbon metabolism and virulence of enteric pathogens. Front Cell Infect Microbiol 4:91. doi:10.3389/fcimb.2014.00091.
177. Nuss AM, Beckstette M, Pimenova M, Schmühl C, Opitz W, Pisano F, Heroven AK, Dersch P. 2017. Tissue dual RNA-seq allows fast discovery of infection-specific functions and riboregulators shaping host-pathogen transcriptomes. Proc Natl Acad Sci U S A 114:E791–E800. [PubMed]
178. Heroven AK, Nuss AM, Dersch P. 2016. RNA-based mechanisms of virulence control in Enterobacteriaceae. RNA Biol 14:471–487. [PubMed]
179. Papenfort K, Podkaminski D, Hinton JC, Vogel J. 2012. The ancestral SgrS RNA discriminates horizontally acquired Salmonella mRNAs through a single G-U wobble pair. Proc Natl Acad Sci U S A 109:E757–E764. [PubMed]
180. Gruber CC, Sperandio V. 2015. Global analysis of posttranscriptional regulation by GlmY and GlmZ in enterohemorrhagic Escherichia coli O157:H7. Infect Immun 83:1286–1295. [PubMed]
181. Chen J, Gottesman S. 2016. Spot 42 sRNA regulates arabinose-inducible araBAD promoter activity by repressing synthesis of the high-affinity low-capacity arabinose transporter. J Bacteriol 199:e00691-16. doi:10.1128/JB.00691-16.
182. Romeo T, Babitzke P. 2018. Global regulation by CsrA and its RNA antagonists. Microbiol Spectrum 6(2) :RWR-0009-2017.
183. Bandyra KJ, Luisi BF. 2018. RNase E and the high fidelity orchestration of RNA metabolism. Microbiol Spectrum 6(2) :RWR-0008-2017.
184. Westermann A. 2018. Regulatory RNAs in virulence and host-microbe interactions. Microbiol Spectrum 6(2) :RWR-0002-2017.
Loading

Article metrics loading...

/content/journal/microbiolspec/10.1128/microbiolspec.RWR-0013-2017
2018-03-23
2020-08-10

Abstract:

Survival of bacteria in ever-changing habitats with fluctuating nutrient supplies requires rapid adaptation of their metabolic capabilities. To this end, carbohydrate metabolism is governed by complex regulatory networks including posttranscriptional mechanisms that involve small regulatory RNAs (sRNAs) and RNA-binding proteins. sRNAs limit the response to substrate availability and set the threshold or time required for induction and repression of carbohydrate utilization systems. Carbon catabolite repression (CCR) also involves sRNAs. In , sRNA Spot 42 cooperates with the transcriptional regulator cyclic AMP (cAMP)-receptor protein (CRP) to repress secondary carbohydrate utilization genes when a preferred sugar is consumed. In pseudomonads, CCR operates entirely at the posttranscriptional level, involving RNA-binding protein Hfq and decoy sRNA CrcZ. Moreover, sRNAs coordinate fluxes through central carbohydrate metabolic pathways with carbohydrate availability. In Gram-negative bacteria, the interplay between RNA-binding protein CsrA and its cognate sRNAs regulates glycolysis and gluconeogenesis in response to signals derived from metabolism. Spot 42 and cAMP-CRP jointly downregulate tricarboxylic acid cycle activity when glycolytic carbon sources are ample. In addition, bacteria use sRNAs to reprogram carbohydrate metabolism in response to anaerobiosis and iron limitation. Finally, sRNAs also provide homeostasis of essential anabolic pathways, as exemplified by the hexosamine pathway providing cell envelope precursors. In this review, we discuss the manifold roles of bacterial sRNAs in regulation of carbon source uptake and utilization, substrate prioritization, and metabolism.

Highlighted Text: Show | Hide
Loading full text...

Full text loading...

Figures

Carbohydrate Utilization in Bacteria: Making the Most Out of Sugars with the Help of Small Regulatory RNAs
[com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-1,webId=/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-1,properties={foaf_givenname=Svetlana, foaf_name=Svetlana Durica-Mitic*, foaf_surname=Durica-Mitic*, pub_isAffiliatedWith=[com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/affiliation/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-aff1]]}], com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-2,webId=/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-2,properties={foaf_givenname=Yvonne, foaf_name=Yvonne Göpel*, foaf_surname=Göpel*, pub_isAffiliatedWith=[com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/affiliation/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-aff2]]}], com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-3,webId=/content/author/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-3,properties={foaf_givenname=Boris, foaf_name=Boris Görke, foaf_surname=Görke, pub_isAffiliatedWith=[com.pub2web.rdf.cci.facet.ContentItem[id=http://asm.metastore.ingenta.com/content/affiliation/microbiolspec/10.1128/microbiolspec.RWR-0013-2017-aff3]]}]]
Citation: Durica-Mitic* S, Göpel* Y, Görke B. 2018. Carbohydrate utilization in bacteria: making the most out of sugars with the help of small regulatory rnas. microbiolspec 6(2): doi:10.1128/microbiolspec.RWR-0013-2017
/content/microbiolspec/10.1128/microbiolspec.RWR-0013-2017.fig1
FIGURE 1
Manifold roles of sRNAs in regulation of carbohydrate metabolism in bacteria. The figure summarizes the major roles of sRNAs (depicted in red) in regulation of carbohydrate metabolism in bacteria. (A) Regulation of uptake and utilization of particular carbohydrates by sRNAs in various species. In , the -encoded sRNA SgrS counteracts phosphosugar stress through repression of glucose transporters and activation of the sugar phosphatase YigL. sRNA ChiX downregulates the chitosugar-specific porin ChiP, setting the threshold concentration for induction of degrading enzymes. Further examples include regulation of host glycan and mannitol uptake by -encoded sRNAs in and species, respectively. (B) Role of sRNAs in CCR. In and , the sRNA Spot 42 represses genes for utilization of secondary carbon sources. Spot 42 is repressed by cAMP-CRP and therefore only active in the presence of preferred sugars generating low cAMP levels. In , translation of mRNAs for utilization of secondary carbon sources is repressed by Hfq. In the absence of preferred substrates, the CbrA/CbrB TCS activates expression of the decoy sRNA CrcZ, titrating Hfq from target transcripts. (C) sRNAs coordinate carbohydrate metabolism with carbohydrate, oxygen, and iron availability. The RNA-binding protein CsrA activates glycolysis and represses gluconeogenesis by binding to corresponding RNAs. CsrA activity is counteracted through sequestration by sRNAs CsrB/CsrC, whose levels are regulated by signals from metabolism. In the absence of oxygen, sRNAs such as FnrS in and RoxS in redirect metabolism from oxidative phosphorylation to anaerobic respiration or fermentation. Upon iron starvation, sRNA RyhB represses TCA cycle enzymes to save iron for essential processes. (D) Example of an anabolic pathway regulated by sRNAs. In , two homologous sRNAs regulate the key enzyme GlmS to achieve homeostasis of glucosamine-6-phosphate (glucosamine-6-P), an essential precursor for cell envelope synthesis.
microbiolspec/6/2/RWR-0013-2017-fig1_thmb.gif
microbiolspec/6/2/RWR-0013-2017-fig1.gif
Image of FIGURE 1
FIGURE 1

Manifold roles of sRNAs in regulation of carbohydrate metabolism in bacteria. The figure summarizes the major roles of sRNAs (depicted in red) in regulation of carbohydrate metabolism in bacteria. (A) Regulation of uptake and utilization of particular carbohydrates by sRNAs in various species. In , the -encoded sRNA SgrS counteracts phosphosugar stress through repression of glucose transporters and activation of the sugar phosphatase YigL. sRNA ChiX downregulates the chitosugar-specific porin ChiP, setting the threshold concentration for induction of degrading enzymes. Further examples include regulation of host glycan and mannitol uptake by -encoded sRNAs in and species, respectively. (B) Role of sRNAs in CCR. In and , the sRNA Spot 42 represses genes for utilization of secondary carbon sources. Spot 42 is repressed by cAMP-CRP and therefore only active in the presence of preferred sugars generating low cAMP levels. In , translation of mRNAs for utilization of secondary carbon sources is repressed by Hfq. In the absence of preferred substrates, the CbrA/CbrB TCS activates expression of the decoy sRNA CrcZ, titrating Hfq from target transcripts. (C) sRNAs coordinate carbohydrate metabolism with carbohydrate, oxygen, and iron availability. The RNA-binding protein CsrA activates glycolysis and represses gluconeogenesis by binding to corresponding RNAs. CsrA activity is counteracted through sequestration by sRNAs CsrB/CsrC, whose levels are regulated by signals from metabolism. In the absence of oxygen, sRNAs such as FnrS in and RoxS in redirect metabolism from oxidative phosphorylation to anaerobic respiration or fermentation. Upon iron starvation, sRNA RyhB represses TCA cycle enzymes to save iron for essential processes. (D) Example of an anabolic pathway regulated by sRNAs. In , two homologous sRNAs regulate the key enzyme GlmS to achieve homeostasis of glucosamine-6-phosphate (glucosamine-6-P), an essential precursor for cell envelope synthesis.

Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
Permissions and Reprints Request Permissions
Download as Powerpoint
/content/microbiolspec/10.1128/microbiolspec.RWR-0013-2017.fig2
FIGURE 2
The transcriptional regulator cAMP-CRP and sRNA Spot 42 cooperate to trigger CCR in . (A) CRP and Spot 42 participate in coherent feedforward loops to prevent utilization of the indicated secondary carbon sources when the preferred carbon source glucose is present. In addition to cAMP-CRP, Spot 42 is regulated by base-pairing with the sponge RNA PspH. (B) The validated Spot 42 regulon to date. Target genes that are also positively controlled by cAMP-CRP at the level of transcription are boxed. Microarray analysis of Spot 42 pulse expression ( 75 ) and improved software prediction algorithms ( 32 , 76 ) fostered the identification of most targets. Additional targets were identified by human inference or by a CLIP-seq approach mapping Hfq binding sites on a global scale ( 74 , 82 , 173 , 181 ). Several of these targets were recovered by RIL-seq ( 77 ).
microbiolspec/6/2/RWR-0013-2017-fig2_thmb.gif
microbiolspec/6/2/RWR-0013-2017-fig2.gif
Image of FIGURE 2
FIGURE 2

The transcriptional regulator cAMP-CRP and sRNA Spot 42 cooperate to trigger CCR in . (A) CRP and Spot 42 participate in coherent feedforward loops to prevent utilization of the indicated secondary carbon sources when the preferred carbon source glucose is present. In addition to cAMP-CRP, Spot 42 is regulated by base-pairing with the sponge RNA PspH. (B) The validated Spot 42 regulon to date. Target genes that are also positively controlled by cAMP-CRP at the level of transcription are boxed. Microarray analysis of Spot 42 pulse expression ( 75 ) and improved software prediction algorithms ( 32 , 76 ) fostered the identification of most targets. Additional targets were identified by human inference or by a CLIP-seq approach mapping Hfq binding sites on a global scale ( 74 , 82 , 173 , 181 ). Several of these targets were recovered by RIL-seq ( 77 ).

Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
Permissions and Reprints Request Permissions
Download as Powerpoint
/content/microbiolspec/10.1128/microbiolspec.RWR-0013-2017.fig3
FIGURE 3
Posttranscriptional regulation of central carbon metabolic pathways in . Effects of sRNAs (depicted in red) and of the RNA-binding protein CsrA (blue) on synthesis of enzymes involved in glycolysis, gluconeogenesis, and the TCA cycle. A green asterisk and bold letters indicate direct regulation by CsrA ( 106 ). Anabolic pathways directing synthesis of glycogen, UDP-sugars, and the biofilm compound PGA are also shown.
microbiolspec/6/2/RWR-0013-2017-fig3_thmb.gif
microbiolspec/6/2/RWR-0013-2017-fig3.gif
Image of FIGURE 3
FIGURE 3

Posttranscriptional regulation of central carbon metabolic pathways in . Effects of sRNAs (depicted in red) and of the RNA-binding protein CsrA (blue) on synthesis of enzymes involved in glycolysis, gluconeogenesis, and the TCA cycle. A green asterisk and bold letters indicate direct regulation by CsrA ( 106 ). Anabolic pathways directing synthesis of glycogen, UDP-sugars, and the biofilm compound PGA are also shown.

Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
Permissions and Reprints Request Permissions
Download as Powerpoint
/content/microbiolspec/10.1128/microbiolspec.RWR-0013-2017.fig4
FIGURE 4
Model of the interconnection of the CsrA system with central carbon metabolism. Decoy sRNAs CsrB and CsrC regulate CsrA activity by sequestering the protein from its target mRNAs. CsrA indirectly activates transcription, creating a negative feedback loop. In fast-growing cells, when CsrB/C levels are low, CsrA activates glycolytic genes and represses the TCA cycle, gluconeogenesis, and glycogen synthesis. Metabolism of glycolytic carbon sources causes accumulation of FBP, which activates pyruvate kinase, thereby reducing the PEP/pyruvate ratio. Intake of PTS substrates and a low PEP/pyruvate ratio trigger dephosphorylation of EIIA, leading to activation of CsrD, which triggers degradation of CsrB/CsrC by RNase E. Upon accumulation of short carboxylic acids (R-COOH) as metabolic end products, expression of / is induced by the BarA/UvrY TCS. Deceleration of glycolytic activity elevates the PEP/pyruvate level and increases EIIA phosphorylation, leading to stabilization of CsrB/CsrC and titration of CsrA. EIIA∼P stimulates adenylate cyclase CyaA, which converts ATP to cAMP. The cAMP-CRP complex inhibits transcription of . Involvement of EIIA in regulation of CsrB/CsrC synthesis as well as decay allows integration of further cues.
microbiolspec/6/2/RWR-0013-2017-fig4_thmb.gif
microbiolspec/6/2/RWR-0013-2017-fig4.gif
Image of FIGURE 4
FIGURE 4

Model of the interconnection of the CsrA system with central carbon metabolism. Decoy sRNAs CsrB and CsrC regulate CsrA activity by sequestering the protein from its target mRNAs. CsrA indirectly activates transcription, creating a negative feedback loop. In fast-growing cells, when CsrB/C levels are low, CsrA activates glycolytic genes and represses the TCA cycle, gluconeogenesis, and glycogen synthesis. Metabolism of glycolytic carbon sources causes accumulation of FBP, which activates pyruvate kinase, thereby reducing the PEP/pyruvate ratio. Intake of PTS substrates and a low PEP/pyruvate ratio trigger dephosphorylation of EIIA, leading to activation of CsrD, which triggers degradation of CsrB/CsrC by RNase E. Upon accumulation of short carboxylic acids (R-COOH) as metabolic end products, expression of / is induced by the BarA/UvrY TCS. Deceleration of glycolytic activity elevates the PEP/pyruvate level and increases EIIA phosphorylation, leading to stabilization of CsrB/CsrC and titration of CsrA. EIIA∼P stimulates adenylate cyclase CyaA, which converts ATP to cAMP. The cAMP-CRP complex inhibits transcription of . Involvement of EIIA in regulation of CsrB/CsrC synthesis as well as decay allows integration of further cues.

Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
Permissions and Reprints Request Permissions
Download as Powerpoint
/content/microbiolspec/10.1128/microbiolspec.RWR-0013-2017.fig5
FIGURE 5
Role of RNase E adaptor protein RapZ in feedback regulation of GlmS synthesis in . When GlcN6P is plentiful in the cell, RapZ prevents upregulation by targeting its activating sRNA GlmZ to cleavage by RNase E. Within the tripartite complex formed, the sRNA is envisioned to be sandwiched between the tetrameric RapZ protein and the N-terminal domain (NTD) of RNase E, which also forms a tetramer ( 168 ). Processing results in functional inactivation of GlmZ and subsequent decline in GlmS levels. Conversely, under GlcN6P depletion, RapZ is predominantly sequestered in complexes with the homologous sRNA GlmY, whose levels increase under this condition. Consequently, GlmZ remains in its active full-length form and stimulates expression. Higher levels of GlmS replenish GlcN6P levels in the cell. Whether RapZ has an active role in sensing GlcN6P via direct binding of the metabolite within its C-terminal domain (CTD) is currently under investigation. The unusual tetrameric structure of RapZ is schematically depicted in the box in the upper left. Each monomer is represented by one color and consists of two globular domains, NTD and CTD, connected via flexible linkers. Three distinct surfaces involved in self-interaction can be discerned: CTD-CTD, NTD-NTD, as well as CTD-NTD ( 168 ).
microbiolspec/6/2/RWR-0013-2017-fig5_thmb.gif
microbiolspec/6/2/RWR-0013-2017-fig5.gif
Image of FIGURE 5
FIGURE 5

Role of RNase E adaptor protein RapZ in feedback regulation of GlmS synthesis in . When GlcN6P is plentiful in the cell, RapZ prevents upregulation by targeting its activating sRNA GlmZ to cleavage by RNase E. Within the tripartite complex formed, the sRNA is envisioned to be sandwiched between the tetrameric RapZ protein and the N-terminal domain (NTD) of RNase E, which also forms a tetramer ( 168 ). Processing results in functional inactivation of GlmZ and subsequent decline in GlmS levels. Conversely, under GlcN6P depletion, RapZ is predominantly sequestered in complexes with the homologous sRNA GlmY, whose levels increase under this condition. Consequently, GlmZ remains in its active full-length form and stimulates expression. Higher levels of GlmS replenish GlcN6P levels in the cell. Whether RapZ has an active role in sensing GlcN6P via direct binding of the metabolite within its C-terminal domain (CTD) is currently under investigation. The unusual tetrameric structure of RapZ is schematically depicted in the box in the upper left. Each monomer is represented by one color and consists of two globular domains, NTD and CTD, connected via flexible linkers. Three distinct surfaces involved in self-interaction can be discerned: CTD-CTD, NTD-NTD, as well as CTD-NTD ( 168 ).

Source: microbiolspec March 2018 vol. 6 no. 2 doi:10.1128/microbiolspec.RWR-0013-2017
Permissions and Reprints Request Permissions
Download as Powerpoint

Supplemental Material

No supplementary material available for this content.

Back to top
This is a required field
Please enter a valid email address
Please check the format of the address you have entered.
Approval was a Success
Invalid data
An Error Occurred
Approval was partially successful, following selected items could not be processed due to error