Proteolytic Regulation of Stress Response Pathways in Escherichia coli

  • Dimce Micevski
  • David A. Dougan
Part of the Subcellular Biochemistry book series (SCBI, volume 66)


Maintaining correct cellular function is a fundamental biological process for all forms of life. A critical aspect of this process is the maintenance of protein homeostasis (proteostasis) in the cell, which is largely performed by a group of proteins, referred to as the protein quality control (PQC) network. This network of proteins, comprised of chaperones and proteases, is critical for maintaining proteostasis not only during favourable growth conditions, but also in response to stress. Indeed proteases play a crucial role in the clearance of unwanted proteins that accumulate during stress, but more importantly, in the activation of various different stress response pathways. In bacteria, the cells response to stress is usually orchestrated by a specific transcription factor (sigma factor). In Escherichia coli there are seven different sigma factors, each of which responds to a particular stress, resulting in the rapid expression of a specific set of genes. The cellular concentration of each transcription factor is tightly controlled, at the level of transcription, translation and protein stability. Here we will focus on the proteolytic regulation of two sigma factors (σ32 and σS), which control the heat and general stress response pathways, respectively. This review will also briefly discuss the role proteolytic systems play in the clearance of unwanted proteins that accumulate during stress.


Sigma Factor Heat Shock Response General Stress Response Protein Quality Control Stress Response Pathway 
These keywords were added by machine and not by the authors. This process is experimental and the keywords may be updated as the learning algorithm improves.



Work in the DAD laboratory is funded by the Australian Research Council.


  1. 1.
    Barchinger SE, Ades SE (2013) Regulated proteolysis: control of the Escherichia coli σE-dependent cell envelope stress response. In: Dougan DA (ed) Regulated proteolysis in microorganisms. Springer, Subcell Biochem 66:129–160Google Scholar
  2. 2.
    Sledjeski DD, Gupta A, Gottesman S (1996) The small RNA, DsrA, is essential for the low temperature expression of RpoS during exponential growth in Escherichia coli. EMBO J 15(15):3993–4000PubMedGoogle Scholar
  3. 3.
    Muffler A, Traulsen DD, Lange R, Hengge-Aronis R (1996) Posttranscriptional osmotic regulation of the sigma(s) subunit of RNA polymerase in Escherichia coli. J Bacteriol 178(6):1607–1613PubMedGoogle Scholar
  4. 4.
    Lange R, Hengge-Aronis R (1991) Growth phase-regulated expression of bolA and morphology of stationary-phase Escherichia coli cells are controlled by the novel sigma factor sigma S. J Bacteriol 173(14):4474–4481PubMedGoogle Scholar
  5. 5.
    Hengge-Aronis R (1993) Survival of hunger and stress: the role of rpoS in early stationary phase gene regulation in E. coli. Cell 72(2):165–168PubMedGoogle Scholar
  6. 6.
    McCann MP, Kidwell JP, Matin A (1991) The putative sigma factor KatF has a central role in development of starvation-mediated general resistance in Escherichia coli. J Bacteriol 173(13):4188–4194PubMedGoogle Scholar
  7. 7.
    Farewell A, Kvint K, Nystrom T (1998) Negative regulation by RpoS: a case of sigma factor competition. Mol Microbiol 29(4):1039–1051PubMedGoogle Scholar
  8. 8.
    Lange R, Hengge-Aronis R (1994) The cellular concentration of the sigma S subunit of RNA polymerase in Escherichia coli is controlled at the levels of transcription, translation, and protein stability. Genes Dev 8(13):1600–1612PubMedGoogle Scholar
  9. 9.
    Atlung T, Brondsted L (1994) Role of the transcriptional activator AppY in regulation of the cyx appA operon of Escherichia coli by anaerobiosis, phosphate starvation, and growth phase. J Bacteriol 176(17):5414–5422PubMedGoogle Scholar
  10. 10.
    Altuvia S, Almiron M, Huisman G, Kolter R et al (1994) The dps promoter is activated by OxyR during growth and by IHF and sigma S in stationary phase. Mol Microbiol 13(2):265–272PubMedGoogle Scholar
  11. 11.
    Dong T, Schellhorn HE (2009) Global effect of RpoS on gene expression in pathogenic Escherichia coli O157:H7 strain EDL933. BMC Genomics 10:349PubMedGoogle Scholar
  12. 12.
    Weber H, Polen T, Heuveling J, Wendisch VF et al (2005) Genome-wide analysis of the general stress response network in Escherichia coli: sigmaS-dependent genes, promoters, and sigma factor selectivity. J Bacteriol 187(5):1591–1603PubMedGoogle Scholar
  13. 13.
    Lacour S, Landini P (2004) SigmaS-dependent gene expression at the onset of stationary phase in Escherichia coli: function of sigmaS-dependent genes and identification of their promoter sequences. J Bacteriol 186(21):7186–7195PubMedGoogle Scholar
  14. 14.
    Patten CL, Kirchhof MG, Schertzberg MR, Morton RA et al (2004) Microarray analysis of RpoS-mediated gene expression in Escherichia coli K-12. Mol Genet Genomics 272(5):580–591PubMedGoogle Scholar
  15. 15.
    Bishop RE, Leskiw BK, Hodges RS, Kay CM et al (1998) The entericidin locus of Escherichia coli and its implications for programmed bacterial cell death. J Mol Biol 280(4):583–596PubMedGoogle Scholar
  16. 16.
    Dong T, Schellhorn HE (2010) Role of RpoS in virulence of pathogens. Infect Immun 78(3):887–897PubMedGoogle Scholar
  17. 17.
    Lange R, Fischer D, Hengge-Aronis R (1995) Identification of transcriptional start sites and the role of ppGpp in the expression of rpoS, the structural gene for the sigma S subunit of RNA polymerase in Escherichia coli. J Bacteriol 177(16):4676–4680PubMedGoogle Scholar
  18. 18.
    Lange R, Hengge-Aronis R (1994) The nlpD gene is located in an operon with rpoS on the Escherichia coli chromosome and encodes a novel lipoprotein with a potential function in cell wall formation. Mol Microbiol 13(4):733–743PubMedGoogle Scholar
  19. 19.
    Hengge-Aronis R (2002) Signal transduction and regulatory mechanisms involved in control of the sigma(S) (RpoS) subunit of RNA polymerase. Microbiol Mol Biol Rev 66(3):373–395PubMedGoogle Scholar
  20. 20.
    Muffler A, Fischer D, Hengge-Aronis R (1996) The RNA-binding protein HF-I, known as a host factor for phage Qbeta RNA replication, is essential for rpoS translation in Escherichia coli. Genes Dev 10(9):1143–1151PubMedGoogle Scholar
  21. 21.
    Argaman L, Hershberg R, Vogel J, Bejerano G et al (2001) Novel small RNA-encoding genes in the intergenic regions of Escherichia coli. Curr Biol 11(12):941–950PubMedGoogle Scholar
  22. 22.
    Eddy SR (2002) Computational genomics of noncoding RNA genes. Cell 109(2):137–140PubMedGoogle Scholar
  23. 23.
    Storz G (2002) An expanding universe of noncoding RNAs. Science 296(5571):1260–1263PubMedGoogle Scholar
  24. 24.
    Wassarman KM, Repoila F, Rosenow C, Storz G et al (2001) Identification of novel small RNAs using comparative genomics and microarrays. Genes Dev 15(13):1637–1651PubMedGoogle Scholar
  25. 25.
    Storz G, Opdyke JA, Zhang A (2004) Controlling mRNA stability and translation with small, noncoding RNAs. Curr Opin Microbiol 7(2):140–144PubMedGoogle Scholar
  26. 26.
    Updegrove T, Wilf N, Sun X, Wartell RM (2008) Effect of Hfq on RprA-rpoS mRNA pairing: Hfq-RNA binding and the influence of the 5′ rpoS mRNA leader region. Biochemistry 47(43):11184–11195PubMedGoogle Scholar
  27. 27.
    Waters LS, Storz G (2009) Regulatory RNAs in bacteria. Cell 136(4):615–628PubMedGoogle Scholar
  28. 28.
    Zhang A, Wassarman KM, Rosenow C, Tjaden BC et al (2003) Global analysis of small RNA and mRNA targets of Hfq. Mol Microbiol 50(4):1111–1124PubMedGoogle Scholar
  29. 29.
    Brown L, Elliott T (1996) Efficient translation of the RpoS sigma factor in Salmonella typhimurium requires host factor I, an RNA-binding protein encoded by the hfq gene. J Bacteriol 178(13):3763–3770PubMedGoogle Scholar
  30. 30.
    Moller T, Franch T, Hojrup P, Keene DR et al (2002) Hfq: a bacterial Sm-like protein that mediates RNA-RNA interaction. Mol Cell 9(1):23–30PubMedGoogle Scholar
  31. 31.
    Muffler A, Traulsen DD, Fischer D, Lange R et al (1997) The RNA-binding protein HF-I plays a global regulatory role which is largely, but not exclusively, due to its role in expression of the sigmaS subunit of RNA polymerase in Escherichia coli. J Bacteriol 179(1):297–300PubMedGoogle Scholar
  32. 32.
    Zhang A, Wassarman KM, Ortega J, Steven AC et al (2002) The Sm-like Hfq protein increases OxyS RNA interaction with target mRNAs. Mol Cell 9(1):11–22PubMedGoogle Scholar
  33. 33.
    Majdalani N, Chen S, Murrow J, St John K et al (2001) Regulation of RpoS by a novel small RNA: the characterization of RprA. Mol Microbiol 39(5):1382–1394PubMedGoogle Scholar
  34. 34.
    Sledjeski DD, Whitman C, Zhang A (2001) Hfq is necessary for regulation by the untranslated RNA DsrA. J Bacteriol 183(6):1997–2005PubMedGoogle Scholar
  35. 35.
    Ueguchi C, Misonou N, Mizuno T (2001) Negative control of rpoS expression by phosphoenolpyruvate: carbohydrate phosphotransferase system in Escherichia coli. J Bacteriol 183(2):520–527PubMedGoogle Scholar
  36. 36.
    Zhang A, Altuvia S, Tiwari A, Argaman L et al (1998) The OxyS regulatory RNA represses rpoS translation and binds the Hfq (HF-I) protein. EMBO J 17(20):6061–6068PubMedGoogle Scholar
  37. 37.
    Kawamoto H, Koide Y, Morita T, Aiba H (2006) Base-pairing requirement for RNA silencing by a bacterial small RNA and acceleration of duplex formation by Hfq. Mol Microbiol 61(4):1013–1022PubMedGoogle Scholar
  38. 38.
    Lease RA, Cusick ME, Belfort M (1998) Riboregulation in Escherichia coli: DsrA RNA acts by RNA:RNA interactions at multiple loci. Proc Natl Acad Sci U S A 95(21):12456–12461PubMedGoogle Scholar
  39. 39.
    Majdalani N, Cunning C, Sledjeski D, Elliott T et al (1998) DsrA RNA regulates translation of RpoS message by an anti-antisense mechanism, independent of its action as an antisilencer of transcription. Proc Natl Acad Sci U S A 95(21):12462–12467PubMedGoogle Scholar
  40. 40.
    Majdalani N, Hernandez D, Gottesman S (2002) Regulation and mode of action of the second small RNA activator of RpoS translation, RprA. Mol Microbiol 46(3):813–826PubMedGoogle Scholar
  41. 41.
    Altuvia S, Weinstein-Fischer D, Zhang A, Postow L et al (1997) A small, stable RNA induced by oxidative stress: role as a pleiotropic regulator and antimutator. Cell 90(1):43–53PubMedGoogle Scholar
  42. 42.
    Repoila F, Gottesman S (2001) Signal transduction cascade for regulation of RpoS: temperature regulation of DsrA. J Bacteriol 183(13):4012–4023PubMedGoogle Scholar
  43. 43.
    Sledjeski D, Gottesman S (1995) A small RNA acts as an antisilencer of the H-NS-silenced rcsA gene of Escherichia coli. Proc Natl Acad Sci U S A 92(6):2003–2007PubMedGoogle Scholar
  44. 44.
    Repoila F, Gottesman S (2003) Temperature sensing by the dsrA promoter. J Bacteriol 185(22):6609–6614PubMedGoogle Scholar
  45. 45.
    Lease RA, Woodson SA (2004) Cycling of the Sm-like protein Hfq on the DsrA small regulatory RNA. J Mol Biol 344(5):1211–1223PubMedGoogle Scholar
  46. 46.
    Cunning C, Brown L, Elliott T (1998) Promoter substitution and deletion analysis of upstream region required for rpoS translational regulation. J Bacteriol 180(17):4564–4570PubMedGoogle Scholar
  47. 47.
    Soper T, Mandin P, Majdalani N, Gottesman S et al (2010) Positive regulation by small RNAs and the role of Hfq. Proc Natl Acad Sci U S A 107(21):9602–9607PubMedGoogle Scholar
  48. 48.
    Soper TJ, Woodson SA (2008) The rpoS mRNA leader recruits Hfq to facilitate annealing with DsrA sRNA. RNA 14(9):1907–1917PubMedGoogle Scholar
  49. 49.
    Hwang W, Arluison V, Hohng S (2011) Dynamic competition of DsrA and rpoS fragments for the proximal binding site of Hfq as a means for efficient annealing. Nucleic Acids Res 39(12):5131–5139PubMedGoogle Scholar
  50. 50.
    Arluison V, Mutyam SK, Mura C, Marco S et al (2007) Sm-like protein Hfq: location of the ATP-binding site and the effect of ATP on Hfq– RNA complexes. Protein Sci 16(9):1830–1841PubMedGoogle Scholar
  51. 51.
    McCullen CA, Benhammou JN, Majdalani N, Gottesman S (2010) Mechanism of positive regulation by DsrA and RprA small noncoding RNAs: pairing increases translation and protects rpoS mRNA from degradation. J Bacteriol 192(21):5559–5571PubMedGoogle Scholar
  52. 52.
    Papenfort K, Said N, Welsink T, Lucchini S et al (2009) Specific and pleiotropic patterns of mRNA regulation by ArcZ, a conserved, Hfq-dependent small RNA. Mol Microbiol 74(1):139–158PubMedGoogle Scholar
  53. 53.
    Mandin P, Gottesman S (2010) Integrating anaerobic/aerobic sensing and the general stress response through the ArcZ small RNA. EMBO J 29(18):3094–3107PubMedGoogle Scholar
  54. 54.
    Schweder T, Lee KH, Lomovskaya O, Matin A (1996) Regulation of Escherichia coli starvation sigma factor (sigma s) by ClpXP protease. J Bacteriol 178(2):470–476PubMedGoogle Scholar
  55. 55.
    Klauck E, Lingnau M, Hengge-Aronis R (2001) Role of the response regulator RssB in sigma recognition and initiation of sigma proteolysis in Escherichia coli. Mol Microbiol 40(6):1381–1390PubMedGoogle Scholar
  56. 56.
    Muffler A, Fischer D, Altuvia S, Storz G et al (1996) The response regulator RssB controls stability of the sigma(S) subunit of RNA polymerase in Escherichia coli. EMBO J 15(6):1333–1339PubMedGoogle Scholar
  57. 57.
    Pratt LA, Silhavy TJ (1996) The response regulator SprE controls the stability of RpoS. Proc Natl Acad Sci U S A 93(6):2488–2492PubMedGoogle Scholar
  58. 58.
    Zhou Y, Gottesman S, Hoskins JR, Maurizi MR et al (2001) The RssB response regulator directly targets sigma(S) for degradation by ClpXP. Genes Dev 15(5):627–637PubMedGoogle Scholar
  59. 59.
    Becker G, Klauck E, Hengge-Aronis R (2000) The response regulator RssB, a recognition factor for sigmaS proteolysis in Escherichia coli, can act like an anti-sigmaS factor. Mol Microbiol 35(3):657–666PubMedGoogle Scholar
  60. 60.
    Becker G, Klauck E, Hengge-Aronis R (1999) Regulation of RpoS proteolysis in Escherichia coli: the response regulator RssB is a recognition factor that interacts with the turnover element in RpoS. Proc Natl Acad Sci U S A 96(11):6439–6444PubMedGoogle Scholar
  61. 61.
    Bouche S, Klauck E, Fischer D, Lucassen M et al (1998) Regulation of RssB-dependent proteolysis in Escherichia coli: a role for acetyl phosphate in a response regulator-controlled process. Mol Microbiol 27(4):787–795PubMedGoogle Scholar
  62. 62.
    Bougdour A, Cunning C, Baptiste PJ, Elliott T et al (2008) Multiple pathways for regulation of sigmaS (RpoS) stability in Escherichia coli via the action of multiple anti-adaptors. Mol Microbiol 68(2):298–313PubMedGoogle Scholar
  63. 63.
    Bougdour A, Wickner S, Gottesman S (2006) Modulating RssB activity: IraP, a novel regulator of sigma(S) stability in Escherichia coli. Genes Dev 20(7):884–897PubMedGoogle Scholar
  64. 64.
    Sauer RT, Baker TA (2011) AAA+  proteases: ATP-fueled machines of protein destruction. Annu Rev Biochem 80:587–612PubMedGoogle Scholar
  65. 65.
    Glynn SE, Martin A, Nager AR, Baker TA et al (2009) Structures of asymmetric ClpX hexamers reveal nucleotide-dependent motions in a AAA+  protein-unfolding machine. Cell 139(4):744–756PubMedGoogle Scholar
  66. 66.
    Kim DY, Kim KK (2003) Crystal structure of ClpX molecular chaperone from Helicobacter pylori. J Biol Chem 278(50):50664–50670PubMedGoogle Scholar
  67. 67.
    Iyer LM, Leipe DD, Koonin EV, Aravind L (2004) Evolutionary history and higher order classification of AAA+  ATPases. J Struct Biol 146(1–2):11–31PubMedGoogle Scholar
  68. 68.
    Kim YI, Levchenko I, Fraczkowska K, Woodruff RV et al (2001) Molecular determinants of complex formation between Clp/Hsp100 ATPases and the ClpP peptidase. Nat Struct Biol 8(3):230–233PubMedGoogle Scholar
  69. 69.
    Neuwald AF, Aravind L, Spouge JL, Koonin EV (1999) AAA+: a class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res 9(1):27–43PubMedGoogle Scholar
  70. 70.
    Walker JE, Saraste M, Gay NJ (1982) E. coli F1-ATPase interacts with a membrane protein component of a proton channel. Nature 298(5877):867–869PubMedGoogle Scholar
  71. 71.
    Gur E, Ottofuelling R, Dougan DA (2013) Machines of destruction – AAA+  proteases and the adaptors that control them. In: Dougan DA (ed) Regulated proteolysis in microorganisms. Springer, Subcell Biochem 66:3–33Google Scholar
  72. 72.
    Jennings LD, Lun DS, Medard M, Licht S (2008) ClpP hydrolyzes a protein substrate processively in the absence of the ClpA ATPase: mechanistic studies of ATP-independent proteolysis. Biochemistry 47(44):11536–11546PubMedGoogle Scholar
  73. 73.
    Wang J, Hartling JA, Flanagan JM (1997) The structure of ClpP at 2.3 A resolution suggests a model for ATP-dependent proteolysis. Cell 91(4):447–456PubMedGoogle Scholar
  74. 74.
    Grimaud R, Kessel M, Beuron F, Steven AC et al (1998) Enzymatic and structural similarities between the Escherichia coli ATP-dependent proteases, ClpXP and ClpAP. J Biol Chem 273(20):12476–12481PubMedGoogle Scholar
  75. 75.
    Maglica Z, Kolygo K, Weber-Ban E (2009) Optimal efficiency of ClpAP and ClpXP chaperone-proteases is achieved by architectural symmetry. Structure 17(4):508–516PubMedGoogle Scholar
  76. 76.
    Ortega J, Lee HS, Maurizi MR, Steven AC (2002) Alternating translocation of protein substrates from both ends of ClpXP protease. EMBO J 21(18):4938–4949PubMedGoogle Scholar
  77. 77.
    Joshi SA, Hersch GL, Baker TA, Sauer RT (2004) Communication between ClpX and ClpP during substrate processing and degradation. Nat Struct Mol Biol 11(5):404–411PubMedGoogle Scholar
  78. 78.
    Martin A, Baker TA, Sauer RT (2007) Distinct static and dynamic interactions control ATPase-peptidase communication in a AAA+  protease. Mol Cell 27(1):41–52PubMedGoogle Scholar
  79. 79.
    Dougan DA (2011) Chemical activators of ClpP: turning Jekyll into Hyde. Chem Biol 18(9):1072–1074PubMedGoogle Scholar
  80. 80.
    Kirstein J, Hoffmann A, Lilie H, Schmidt R et al (2009) The antibiotic ADEP reprogrammes ClpP, switching it from a regulated to an uncontrolled protease. EMBO Mol Med 1(1):37–49PubMedGoogle Scholar
  81. 81.
    Lee BG, Park EY, Lee KE, Jeon H et al (2010) Structures of ClpP in complex with acyldepsipeptide antibiotics reveal its activation mechanism. Nat Struct Mol Biol 17(4):471–478PubMedGoogle Scholar
  82. 82.
    Leung E, Datti A, Cossette M, Goodreid J et al (2011) Activators of cylindrical proteases as antimicrobials: identification and development of small molecule activators of ClpP protease. Chem Biol 18(9):1167–1178PubMedGoogle Scholar
  83. 83.
    Li DH, Chung YS, Gloyd M, Joseph E et al (2010) Acyldepsipeptide antibiotics induce the formation of a structured axial channel in ClpP: A model for the ClpX/ClpA-bound state of ClpP. Chem Biol 17(9):959–969PubMedGoogle Scholar
  84. 84.
    Gribun A, Kimber MS, Ching R, Sprangers R et al (2005) The ClpP double ring tetradecameric protease exhibits plastic ring-ring interactions, and the N termini of its subunits form flexible loops that are essential for ClpXP and ClpAP complex formation. J Biol Chem 280(16):16185–16196PubMedGoogle Scholar
  85. 85.
    Kang SG, Maurizi MR, Thompson M, Mueser T et al (2004) Crystallography and mutagenesis point to an essential role for the N-terminus of human mitochondrial ClpP. J Struct Biol 148(3):338–352PubMedGoogle Scholar
  86. 86.
    Jennings LD, Bohon J, Chance MR, Licht S (2008) The ClpP N-terminus coordinates substrate access with protease active site reactivity. Biochemistry 47(42):11031–11040PubMedGoogle Scholar
  87. 87.
    Martin A, Baker TA, Sauer RT (2008) Pore loops of the AAA+  ClpX machine grip substrates to drive translocation and unfolding. Nat Struct Mol Biol 15(11):1147–1151PubMedGoogle Scholar
  88. 88.
    Martin A, Baker TA, Sauer RT (2008) Diverse pore loops of the AAA+  ClpX machine mediate unassisted and adaptor-dependent recognition of ssrA-tagged substrates. Mol Cell 29(4):441–450PubMedGoogle Scholar
  89. 89.
    Flynn JM, Neher SB, Kim YI, Sauer RT et al (2003) Proteomic discovery of cellular substrates of the ClpXP protease reveals five classes of ClpX-recognition signals. Mol Cell 11(3):671–683PubMedGoogle Scholar
  90. 90.
    Kim YI, Burton RE, Burton BM, Sauer RT et al (2000) Dynamics of substrate denaturation and translocation by the ClpXP degradation machine. Mol Cell 5(4):639–648PubMedGoogle Scholar
  91. 91.
    Ortega J, Singh SK, Ishikawa T, Maurizi MR et al (2000) Visualization of substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 6(6):1515–1521PubMedGoogle Scholar
  92. 92.
    Thompson MW, Singh SK, Maurizi MR (1994) Processive degradation of proteins by the ATP-dependent Clp protease from Escherichia coli. Requirement for the multiple array of active sites in ClpP but not ATP hydrolysis. J Biol Chem 269(27):18209–18215PubMedGoogle Scholar
  93. 93.
    Schlieker C, Weibezahn J, Patzelt H, Tessarz P et al (2004) Substrate recognition by the AAA+  chaperone ClpB. Nat Struct Mol Biol 11(7):607–615PubMedGoogle Scholar
  94. 94.
    Siddiqui SM, Sauer RT, Baker TA (2004) Role of the processing pore of the ClpX AAA+  ATPase in the recognition and engagement of specific protein substrates. Genes Dev 18(4):369–374PubMedGoogle Scholar
  95. 95.
    Wang J, Song JJ, Franklin MC, Kamtekar S et al (2001) Crystal structures of the HslVU peptidase-ATPase complex reveal an ATP-dependent proteolysis mechanism. Structure 9(2):177–184PubMedGoogle Scholar
  96. 96.
    Yamada-Inagawa T, Okuno T, Karata K, Yamanaka K et al (2003) Conserved pore residues in the AAA protease FtsH are important for proteolysis and its coupling to ATP hydrolysis. J Biol Chem 278(50):50182–50187PubMedGoogle Scholar
  97. 97.
    Lum R, Tkach JM, Vierling E, Glover JR (2004) Evidence for an unfolding/threading mechanism for protein disaggregation by Saccharomyces cerevisiae Hsp104. J Biol Chem 279(28):29139–29146PubMedGoogle Scholar
  98. 98.
    Gonzalez M, Rasulova F, Maurizi MR, Woodgate R (2000) Subunit-specific degradation of the UmuD/D′ heterodimer by the ClpXP protease: the role of trans recognition in UmuD′ stability. EMBO J 19(19):5251–5258PubMedGoogle Scholar
  99. 99.
    Dougan DA, Weber-Ban E, Bukau B (2003) Targeted delivery of an ssrA-tagged substrate by the adaptor protein SspB to its cognate AAA+  protein ClpX. Mol Cell 12(2):373–380PubMedGoogle Scholar
  100. 100.
    Neher SB, Sauer RT, Baker TA (2003) Distinct peptide signals in the UmuD and UmuD′ subunits of UmuD/D′ mediate tethering and substrate processing by the ClpXP protease. Proc Natl Acad Sci U S A 100(23):13219–13224PubMedGoogle Scholar
  101. 101.
    Wah DA, Levchenko I, Rieckhof GE, Bolon DN et al (2003) Flexible linkers leash the substrate binding domain of SspB to a peptide module that stabilizes delivery complexes with the AAA+  ClpXP protease. Mol Cell 12(2):355–363PubMedGoogle Scholar
  102. 102.
    Galperin MY (2006) Structural classification of bacterial response regulators: diversity of output domains and domain combinations. J Bacteriol 188(12):4169–4182PubMedGoogle Scholar
  103. 103.
    Peterson CN, Ruiz N, Silhavy TJ (2004) RpoS proteolysis is regulated by a mechanism that does not require the SprE (RssB) response regulator phosphorylation site. J Bacteriol 186(21):7403–7410PubMedGoogle Scholar
  104. 104.
    Bren A, Welch M, Blat Y, Eisenbach M (1996) Signal termination in bacterial chemotaxis: CheZ mediates dephosphorylation of free rather than switch-bound CheY. Proc Natl Acad Sci U S A 93(19):10090–10093PubMedGoogle Scholar
  105. 105.
    Hess JF, Bourret RB, Simon MI (1988) Histidine phosphorylation and phosphoryl group transfer in bacterial chemotaxis. Nature 336(6195):139–143PubMedGoogle Scholar
  106. 106.
    Bachhawat P, Stock AM (2007) Crystal structures of the receiver domain of the response regulator PhoP from Escherichia coli in the absence and presence of the phosphoryl analog beryllofluoride. J Bacteriol 189(16):5987–5995PubMedGoogle Scholar
  107. 107.
    Bachhawat P, Swapna GV, Montelione GT, Stock AM (2005) Mechanism of activation for transcription factor PhoB suggested by different modes of dimerization in the inactive and active states. Structure 13(9):1353–1363PubMedGoogle Scholar
  108. 108.
    Lee SY, Cho HS, Pelton JG, Yan D et al (2001) Crystal structure of activated CheY. Comparison with other activated receiver domains. J Biol Chem 276(19):16425–16431PubMedGoogle Scholar
  109. 109.
    Toro-Roman A, Mack TR, Stock AM (2005) Structural analysis and solution studies of the activated regulatory domain of the response regulator ArcA: a symmetric dimer mediated by the alpha4-beta5-alpha5 face. J Mol Biol 349(1):11–26PubMedGoogle Scholar
  110. 110.
    Toro-Roman A, Wu T, Stock AM (2005) A common dimerization interface in bacterial response regulators KdpE and TorR. Protein Sci 14(12):3077–3088PubMedGoogle Scholar
  111. 111.
    Studemann A, Noirclerc-Savoye M, Klauck E, Becker G et al (2003) Sequential recognition of two distinct sites in sigma(S) by the proteolytic targeting factor RssB and ClpX. EMBO J 22(16):4111–4120PubMedGoogle Scholar
  112. 112.
    D’Souza C, Nakano MM, Zuber P (1994) Identification of comS, a gene of the srfA operon that regulates the establishment of genetic competence in Bacillus subtilis. Proc Natl Acad Sci U S A 91(20):9397–9401PubMedGoogle Scholar
  113. 113.
    Hamoen LW, Eshuis H, Jongbloed J, Venema G et al (1995) A small gene, designated comS, located within the coding region of the fourth amino acid-activation domain of srfA, is required for competence development in Bacillus subtilis. Mol Microbiol 15(1):55–63PubMedGoogle Scholar
  114. 114.
    Turgay K, Hamoen LW, Venema G, Dubnau D (1997) Biochemical characterization of a molecular switch involving the heat shock protein ClpC, which controls the activity of ComK, the competence transcription factor of Bacillus subtilis. Genes Dev 11(1):119–128PubMedGoogle Scholar
  115. 115.
    Persuh M, Turgay K, Mandic-Mulec I, Dubnau D (1999) The N- and C-terminal domains of MecA recognize different partners in the competence molecular switch. Mol Microbiol 33(4):886–894PubMedGoogle Scholar
  116. 116.
    Turgay K, Hahn J, Burghoorn J, Dubnau D (1998) Competence in Bacillus subtilis is controlled by regulated proteolysis of a transcription factor. EMBO J 17(22):6730–6738PubMedGoogle Scholar
  117. 117.
    Ogura M, Liu L, Lacelle M, Nakano MM et al (1999) Mutational analysis of ComS: evidence for the interaction of ComS and MecA in the regulation of competence development in Bacillus subtilis. Mol Microbiol 32(4):799–812PubMedGoogle Scholar
  118. 118.
    Bougdour A, Gottesman S (2007) ppGpp regulation of RpoS degradation via anti-adaptor protein IraP. Proc Natl Acad Sci U S A 104(31):12896–12901PubMedGoogle Scholar
  119. 119.
    Tissieres A, Mitchell HK, Tracy UM (1974) Protein synthesis in salivary glands of Drosophila melanogaster: relation to chromosome puffs. J Mol Biol 84(3):389–398PubMedGoogle Scholar
  120. 120.
    Gamer J, Bujard H, Bukau B (1992) Physical interaction between heat shock proteins DnaK, DnaJ, and GrpE and the bacterial heat shock transcription factor sigma 32. Cell 69(5):833–842PubMedGoogle Scholar
  121. 121.
    Guisbert E, Herman C, Lu CZ, Gross CA (2004) A chaperone network controls the heat shock response in E. coli. Genes Dev 18(22):2812–2821PubMedGoogle Scholar
  122. 122.
    Straus DB, Walter WA, Gross CA (1987) The heat shock response of E. coli is regulated by changes in the concentration of sigma 32. Nature 329(6137):348–351PubMedGoogle Scholar
  123. 123.
    Herman C, Thevenet D, D’Ari R, Bouloc P (1995) Degradation of sigma 32, the heat shock regulator in Escherichia coli, is governed by HflB. Proc Natl Acad Sci U S A 92(8):3516–3520PubMedGoogle Scholar
  124. 124.
    Kanemori M, Nishihara K, Yanagi H, Yura T (1997) Synergistic roles of HslVU and other ATP-dependent proteases in controlling in vivo turnover of sigma32 and abnormal proteins in Escherichia coli. J Bacteriol 179(23):7219–7225PubMedGoogle Scholar
  125. 125.
    Tomoyasu T, Gamer J, Bukau B, Kanemori M et al (1995) Escherichia coli FtsH is a membrane-bound, ATP-dependent protease which degrades the heat-shock transcription factor sigma 32. EMBO J 14(11):2551–2560PubMedGoogle Scholar
  126. 126.
    Sousa MC, Trame CB, Tsuruta H, Wilbanks SM et al (2000) Crystal and solution structures of an HslUV protease-chaperone complex. Cell 103(4):633–643PubMedGoogle Scholar
  127. 127.
    Okuno T, Ogura T (2013) FtsH protease-mediated regulation of various cellular functions. In: Dougan DA (ed) Regulated proteolysis in microorganisms. Springer, Subcell Biochem 66:53–69Google Scholar
  128. 128.
    Kanemori M, Yanagi H, Yura T (1999) Marked instability of the sigma(32) heat shock transcription factor at high temperature. Implications for heat shock regulation. J Biol Chem 274(31):22002–22007PubMedGoogle Scholar
  129. 129.
    Rodriguez F, Arsene-Ploetze F, Rist W, Rudiger S et al (2008) Molecular basis for regulation of the heat shock transcription factor sigma32 by the DnaK and DnaJ chaperones. Mol Cell 32(3):347–358PubMedGoogle Scholar
  130. 130.
    Horikoshi M, Yura T, Tsuchimoto S, Fukumori Y et al (2004) Conserved region 2.1 of Escherichia coli heat shock transcription factor sigma32 is required for modulating both metabolic stability and transcriptional activity. J Bacteriol 186(22):7474–7480PubMedGoogle Scholar
  131. 131.
    Obrist M, Milek S, Klauck E, Hengge R et al (2007) Region 2.1 of the Escherichia coli heat-shock sigma factor RpoH (sigma32) is necessary but not sufficient for degradation by the FtsH protease. Microbiology 153(Pt 8):2560–2571PubMedGoogle Scholar
  132. 132.
    Herman C, Prakash S, Lu CZ, Matouschek A et al (2003) Lack of a robust unfoldase activity confers a unique level of substrate specificity to the universal AAA protease FtsH. Mol Cell 11(3):659–669PubMedGoogle Scholar
  133. 133.
    Langklotz S, Baumann U, Narberhaus F (2012) Structure and function of the bacterial AAA protease FtsH. Biochim Biophys Acta 1823(1):40–48PubMedGoogle Scholar
  134. 134.
    Obrist M, Langklotz S, Milek S, Fuhrer F et al (2009) Region C of the Escherichia coli heat shock sigma factor RpoH (sigma 32) contains a turnover element for proteolysis by the FtsH protease. FEMS Microbiol Lett 290(2):199–208PubMedGoogle Scholar
  135. 135.
    Tomoyasu T, Arsene F, Ogura T, Bukau B (2001) The C terminus of sigma(32) is not essential for degradation by FtsH. J Bacteriol 183(20):5911–5917PubMedGoogle Scholar
  136. 136.
    Wickner S, Maurizi MR, Gottesman S (1999) Posttranslational quality control: folding, refolding, and degrading proteins. Science 286(5446):1888–1893PubMedGoogle Scholar
  137. 137.
    Gur E, Sauer RT (2008) Recognition of misfolded proteins by Lon, a AAA(+) protease. Genes Dev 22(16):2267–2277PubMedGoogle Scholar
  138. 138.
    Dougan DA, Reid BG, Horwich AL, Bukau B (2002) ClpS, a substrate modulator of the ClpAP machine. Mol Cell 9(3):673–683PubMedGoogle Scholar
  139. 139.
    Farrell CM, Grossman AD, Sauer RT (2005) Cytoplasmic degradation of ssrA-tagged proteins. Mol Microbiol 57(6):1750–1761PubMedGoogle Scholar
  140. 140.
    Mogk A, Tomoyasu T, Goloubinoff P, Rudiger S et al (1999) Identification of thermolabile Escherichia coli proteins: prevention and reversion of aggregation by DnaK and ClpB. EMBO J 18(24):6934–6949PubMedGoogle Scholar
  141. 141.
    Goloubinoff P, Mogk A, Zvi AP, Tomoyasu T et al (1999) Sequential mechanism of solubilization and refolding of stable protein aggregates by a bichaperone network. Proc Natl Acad Sci U S A 96(24):13732–13737PubMedGoogle Scholar
  142. 142.
    Schlothauer T, Mogk A, Dougan DA, Bukau B et al (2003) MecA, an adaptor protein necessary for ClpC chaperone activity. Proc Natl Acad Sci U S A 100(5):2306–2311PubMedGoogle Scholar
  143. 143.
    Wawrzynow A, Wojtkowiak D, Marszalek J, Banecki B et al (1995) The ClpX heat-shock protein of Escherichia coli, the ATP-dependent substrate specificity component of the ClpP-ClpX protease, is a novel molecular chaperone. EMBO J 14(9):1867–1877PubMedGoogle Scholar
  144. 144.
    Kock H, Gerth U, Hecker M (2004) The ClpP peptidase is the major determinant of bulk protein turnover in Bacillus subtilis. J Bacteriol 186(17):5856–5864PubMedGoogle Scholar
  145. 145.
    Miethke M, Hecker M, Gerth U (2006) Involvement of Bacillus subtilis ClpE in CtsR degradation and protein quality control. J Bacteriol 188(13):4610–4619PubMedGoogle Scholar
  146. 146.
    Dougan DA, Mogk A, Bukau B (2002) Protein folding and degradation in bacteria: to degrade or not to degrade? That is the question. Cell Mol Life Sci 59(10):1607–1616PubMedGoogle Scholar
  147. 147.
    Matuszewska M, Kuczynska-Wisnik D, Laskowska E, Liberek K (2005) The small heat shock protein IbpA of Escherichia coli cooperates with IbpB in stabilization of thermally aggregated proteins in a disaggregation competent state. J Biol Chem 280(13):12292–12298PubMedGoogle Scholar
  148. 148.
    Mogk A, Deuerling E, Vorderwulbecke S, Vierling E et al (2003) Small heat shock proteins, ClpB and the DnaK system form a functional triade in reversing protein aggregation. Mol Microbiol 50(2):585–595PubMedGoogle Scholar
  149. 149.
    Bissonnette SA, Rivera-Rivera I, Sauer RT, Baker TA (2010) The IbpA and IbpB small heat-shock proteins are substrates of the AAA+  Lon protease. Mol Microbiol 75(6):1539–1549PubMedGoogle Scholar

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© Springer Science+Business Media Dordrecht 2013

Authors and Affiliations

  1. 1.Department of Biochemistry, La Trobe Institute for Molecular Science (LIMS)La Trobe UniversityMelbourneAustralia

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