Mechanistic insights into bacterial AAA+ proteases and protein-remodelling machines

Nature Reviews Microbiology

Volumn 14, Issue 1, 2015, Pages 33-44

  Olivares, Adrian O ^a   Baker, Tania A ^a   Sauer, Robert T ^a  

^a MASSACHUSETTS INSTITUTE OF TECHNOLOGY   (United States)

Author keywords

[No Author keywords available]

Indexed keywords

BACTERIAL AAA+ PROTEASE; BACTERIAL PROTEIN; CLPXP PROTEIN; ENDOPEPTIDASE CLPX; PROTEINASE; UNCLASSIFIED DRUG; ADENOSINE TRIPHOSPHATE; ENDOPEPTIDASE CLP; MACROMOLECULE;

COMPLEX FORMATION; ENZYME MECHANISM; ENZYME RELEASE; ENZYME SUBSTRATE; NONHUMAN; PRIORITY JOURNAL; PROTEIN DEGRADATION; PROTEIN DENATURATION; PROTEIN DOMAIN; PROTEIN FUNCTION; PROTEIN INTERACTION; PROTEIN STRUCTURE; PROTEIN UNFOLDING; REVIEW; BACTERIUM; BIOLOGICAL MODEL; CHEMISTRY; ENZYMOLOGY; GENETICS; MACROMOLECULE; METABOLISM; MOLECULAR MODEL; PROTEIN FOLDING;

ADENOSINE TRIPHOSPHATE; BACTERIA; ENDOPEPTIDASE CLP; MACROMOLECULAR SUBSTANCES; MODELS, BIOLOGICAL; MODELS, MOLECULAR; PROTEIN FOLDING; PROTEOLYSIS;

EID: 84951569973     PISSN: 17401526     EISSN: 17401534     Source Type: Journal    
DOI: 10.1038/nrmicro.2015.4     Document Type: Review

References (108)

1. Oldfield, C. J. & Dunker, A. K. Intrinsically disordered proteins and intrinsically disordered protein regions. Annu. Rev. Biochem. 83, 553-584 (2014).
2. Sauer, R. T. & Baker, T. A. AAA+ proteases: ATP-fueled machines of protein destruction. Annu. Rev. Biochem. 80, 587-612 (2011).
3. Striebel, F., Kress, W. & Weber-Ban, E. Controlled destruction: AAA+ ATPases in protein degradation from bacteria to eukaryotes. Curr. Opin. Struct. Biol. 19, 209-217 (2009).
4. Matyskiela, M. E. & Martin, A. Design principles of a universal protein degradation machine. J. Mol. Biol. 425, 199-213 (2013).
5. Barthelme, D. & Sauer, R. T. Identification of the Cdc48-20S proteasome as an ancient AAA+ proteolytic machine. Science 337, 843-846 (2012).
6. Meyer, A. S. & Baker, T. A. Proteolysis in the Escherichia coli heat shock response: a player at many levels. Curr. Opin. Microbiol. 14, 194-199 (2011).
7. Gur, E. The Lon AAA+ protease. Subcell. Biochem. 66, 35-51 (2013).
8. Flynn, J. M., Levchenko, I., Sauer, R. T. & Baker, T. A. Modulating substrate choice: the SspB adaptor delivers a regulator of the extracytoplasmic-stress response to the AAA+ protease ClpXP for degradation. Genes Dev. 18, 2292-2301 (2004).
9. Moore, S. D. & Sauer, R. T. The tmRNA system for translational surveillance and ribosome rescue. Annu. Rev. Biochem. 76, 101-124 (2007).
10. Keiler, K. C. Mechanisms of ribosome rescue in bacteria. Nat. Rev. Microbiol. 13, 285-297 (2015).
11. Zhou, Y. & Gottesman, S. Regulation of proteolysis of the stationary-phase sigma factor RpoS. J. Bacteriol. 180, 1154-1158 (1998).
12. Peterson, C. N., Levchenko, I., Rabinowitz, J. D., Baker, T. A. & Silhavy, T. J. RpoS proteolysis is controlled directly by ATP levels in Escherichia coli. Genes Dev. 26, 548-553 (2012).
13. Mizusawa, S. & Gottesman, S. Protein degradation in Escherichia coli: the lon gene controls the stability of SulA protein. Proc. Natl Acad. Sci. USA 80, 358-362 (1983).
14. Jenal, U. & Fuchs, T. An essential protease involved in bacterial cell-cycle control. EMBO J. 17, 5658-5669 (1998).
15. Ingmer, H. & Brondsted, L. Proteases in bacterial pathogenesis. Res. Microbiol. 160, 704-710 (2009).
16. Konovalova, A., Sogaard-Andersen, L. & Kroos, L. Regulated proteolysis in bacterial development. FEMS Microbiol. Rev. 38, 493-522 (2014).
17. Gur, E. & Sauer, R. T. Evolution of the ssrA degradation tag in Mycoplasma: specificity switch to a different protease. Proc. Natl Acad. Sci. USA 105, 16113-16118 (2008).
18. Ge, Z. & Karzai, A. W. Co-evolution of multipartite interactions between an extended tmRNA tag and a robust Lon protease in Mycoplasma. Mol. Microbiol. 74, 1083-1099 (2009).
19. Baker, T. A. & Sauer, R. T. ClpXP, an ATP-powered unfolding and protein-degradation machine. Biochim. Biophys. Acta 1823, 15-28 (2012).
20. Singh, S. K. et al. Functional domains of the ClpA and ClpX molecular chaperones identified by limited proteolysis and deletion analysis. J. Biol. Chem. 276, 29420-29429 (2001).
21. Wojtyra, U. A., Thibault, G., Tuite, A. & Houry, W. A. The N-terminal zinc binding domain of ClpX is a dimerization domain that modulates the chaperone function. J. Biol. Chem. 278, 48981-48990 (2003).
22. Wang, J., Hartling, J. A. & Flanagan, J. M. The structure of ClpP at 2.3 resolution suggests a model for ATP-dependent proteolysis. Cell 91, 447-456 (1997).
23. Yu, A. Y. H. & Houry, W. A. ClpP: a distinctive family of cylindrical energy-dependent serine proteases. FEBS Lett. 581, 3749-3757 (2007).
24. Alexopoulos, J. A., Guarn, A. & Ortega, J. ClpP: a structurally dynamic protease regulated by AAA+ proteins. J. Struct. Biol. 179, 202-210 (2012).
25. Lee, M. E., Baker, T. A. & Sauer, R. T. Control of substrate gating and translocation into ClpP by channel residues and ClpX binding. J. Mol. Biol. 399, 707-718 (2010).
26. Grimaud, R., Kessel, M., Beuron, F., Steven, A. C. & Maurizi, M. R. Enzymatic and structural similarities between the Escherichia coli ATP-dependent proteases, ClpXP and ClpAP. J. Biol. Chem. 273, 12476-12481 (1998).
27. Kim, Y. I. et al. Molecular determinants of complex formation between Clp/Hsp100 ATPases and the ClpP peptidase. Nat. Struct. Biol. 8, 230-233 (2001).
28. Martin, A., Baker, T. A. & Sauer, R. T. Distinct static and dynamic interactions control ATPase-peptidase communication in a AAA+ protease. Mol. Cell 27, 41-52 (2007).
29. Akopian, T. et al. The active ClpP protease from M. tuberculosis is a complex composed of a heptameric ClpP1 and a ClpP2 ring. EMBO J. 31, 1529-1541 (2012).
30. Schmitz, K. R. & Sauer, R. T. Substrate delivery by the AAA+ ClpX and ClpC1 unfoldases activates the mycobacterial ClpP1P2 peptidase. Mol. Microbiol. 93, 617-628 (2014).
31. Stanne, T. M., Pojidaeva, E., Andersson, F. I. & Clarke, A. K. Distinctive types of ATP-dependent Clp proteases in cyanobacteria. J. Biol. Chem. 282, 14394-14402 (2007).
32. Olinares, P. D. B., Kim, J., Davis, J. I. & van Wijk, K. J. Subunit stoichiometry, evolution, and functional implications of an asymmetric plant plastid ClpP/R protease complex in Arabidopsis. Plant Cell 23, 2348-2361 (2011).
33. Compton, C. L., Schmitz, K. R., Sauer, R. T. & Sello, J. K. Antibacterial activity of and resistance to small molecule inhibitors of the ClpP peptidase. ACS Chem. Biol. 8, 2669-2677 (2013).
34. Kang, S. G., Dimitrova, M. N., Ortega, J., Ginsburg, A. & Maurizi, M. R. Human mitochondrial ClpP is a stable heptamer that assembles into a tetradecamer in the presence of ClpX. J. Biol. Chem. 280, 35424-35432 (2005).
35. Schmitz, K. R., Carney, D. W., Sello, J. K. & Sauer, R. T. Crystal structure of Mycobacterium tuberculosis ClpP1P2 suggests a model for peptidase activation by AAA+ partner binding and substrate delivery. Proc. Natl Acad. Sci. USA 111, E4587-E4595 (2014).
36. Sprangers, R., Gribun, A., Hwang, P. M., Houry, W. A. & Kay, L. E. Quantitative NMR spectroscopy of supramolecular complexes: dynamic side pores in ClpP are important for product release. Proc. Natl Acad. Sci. USA 102, 16678-16683 (2005).
37. Brotz-Oesterhelt, H. et al. Dysregulation of bacterial proteolytic machinery by a new class of antibiotics. Nat. Med. 11, 1082-1087 (2005). This article details the discovery that acyldepsipeptides (ADEPs) kill bacteria by targeting the ClpP peptidase and activating the degradation of unstructured proteins in the absence of a AAA+ unfoldase partner.
38. Kirstein, J. et al. The antibiotic ADEP reprogrammes ClpP, switching it from a regulated to an uncontrolled protease. EMBO Mol. Med. 1, 37-49 (2009).
39. Lee, B. G. et al. Structures of ClpP in complex with acyldepsipeptide antibiotics reveal its activation mechanism. Nat. Struct. Mol. Biol. 17, 471-478 (2010).
40. Li, D. H. et al. 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, 959-969 (2010).
41. Zeiler, E., Korotkov, V. S., Lorenz-Baath, K., Bottcher, T. & Sieber, S. A. Development and characterization of improved β-lactone-based anti-virulence drugs targeting ClpP. Bioorg. Med. Chem. 20, 583-591 (2012).
42. Conlon, B. P. et al. Activated ClpP kills persisters and eradicates a chronic biofilm infection. Nature 503, 365-370 (2013). This study demonstrates that ADEPs, in combination with a traditional antibiotic, can effectively eliminate dormant persister cells in biofilms, which are responsible for many drug-resistant chronic infections.
43. Vasudevan, D., Rao, S. P. S. & Noble, C. G. Structural basis of mycobacterial inhibition by cyclomarin A. J. Biol. Chem. 288, 30883-30891 (2013).
44. Carney, D. W., Schmitz, K. R., Truong, J. V., Sauer, R. T. & Sello, J. K. Restriction of the conformational dynamics of the cyclic acyldepsipeptide antibiotics improves their antibacterial activity. J. Am. Chem. Soc. 136, 1922-1929 (2014).
45. Gavrish, E. et al. Lassomycin, a ribosomally synthesized cyclic peptide, kills Mycobacterium tuberculosis by targeting the ATP-dependent protease ClpC1P1P2. Chem. Biol. 21, 509-518 (2014).
46. Thompson, M. W., Singh, S. K. & Maurizi, M. R. Processive degradation of proteins by the ATPdependent Clp protease from Escherichia coli. Requirement for the multiple array of active sites in ClpP but not ATP hydrolysis. J. Biol. Chem. 269, 18209-18215 (1994).
47. Jennings, L. D., Lun, D. S., Mdard, M. & Licht, S. ClpP hydrolyzes a protein substrate processively in the absence of the ClpA ATPase: mechanistic studies of ATP-independent proteolysis. Biochemistry 47, 11536-11546 (2008).
48. Finley, D. Recognition and processing of ubiquitinprotein conjugates by the proteasome. Annu. Rev. Biochem. 78, 477-513 (2009).
49. Striebel, F., Imkamp, F., Ozcelik, D. & Weber-Ban, E. Pupylation as a signal for proteasomal degradation in bacteria. Biochim. Biophys. Acta 1843, 103-113 (2014).
50. Elharar, Y. et al. Survival of mycobacteria depends on proteasome-mediated amino acid recycling under nutrient limitation. EMBO J. 33, 1802-1814 (2014).
51. Bougdour, A., Cunning, C., Baptiste, P. J., Elliott, T. & Gottesman, S. Multiple pathways for regulation of S (RpoS) stability in Escherichia coli via the action of multiple anti-adaptors. Mol. Microbiol. 68, 298-313 (2008).
52. Abel, S. et al. Regulatory cohesion of cell cycle and cell differentiation through interlinked phosphorylation and second messenger networks. Mol. Cell 43, 550-560 (2011).
53. Rood, K. L., Clark, N. E., Stoddard, P. R., Garman, S. C. & Chien, P. Adaptor-dependent degradation of a cellcycle regulator uses a unique substrate architecture. Structure 20, 1223-1232 (2012).
54. Glynn, S. E., Martin, A., Nager, A. R., Baker, T. A. & Sauer, R. T. Structures of asymmetric ClpX hexamers reveal nucleotide-dependent motions in a AAA+ protein-unfolding machine. Cell 139, 744-756 (2009).
55. Stinson, B. M. et al. Nucleotide binding and conformational switching in the hexameric ring of a AAA+ machine. Cell 153, 628-639 (2013).
56. Matyskiela, M. E., Lander, G. C. & Martin, A. Conformational switching of the 26S proteasome enables substrate degradation. Nat. Struct. Mol. Biol. 20, 781-788 (2013). This article reports a cryo-EM structure of the 26S proteasome, which provides important mechanistic insights into substrate recognition, deubiquitylation, unfolding and translocation.
57. Stinson, B. M., Baytshtok, V., Schmitz, K. R., Baker, T. A. & Sauer, R. T. Subunit asymmetry and roles of conformational switching in the hexameric AAA+ ring of ClpX. Nat. Struct. Mol. Biol. 22, 411-416 (2015).
58. Glynn, S. E., Nager, A. R., Baker, T. A. & Sauer, R. T. Dynamic and static components power unfolding in topologically closed rings of a AAA+ proteolytic machine. Nat. Struct. Mol. Biol. 19, 616-622 (2012). This study probes ClpX using structure-guided crosslinking across the rigid-body interfaces, which reveals that a topologically closed ring is mechanically active and assumes different conformations by altering the geometry of the hinges between the large and small AAA+ domains of each subunit.
59. Martin, A., Baker, T. A. & Sauer, R. T. Rebuilt AAA+ motors reveal operating principles for ATP-fuelled machines. Nature 437, 1115-1120 (2005). In this study, engineering and characterization of single-chain ClpX hexamers with different combinations of active and inactive subunits support a probabilistic model of AAA+ ring function in which ATP hydrolysis in a single subunit generates a power stroke.
60. Gai, D., Zhao, R., Li, D., Finkielstein, C. V. & Chen, X. S. Mechanisms of conformational change for a replicative hexameric helicase of SV40 large tumor antigen. Cell 119, 47-60 (2004).
61. Smith, D., Fraga, H., Reis, C. & Kafri, G. ATP binds to proteasomal ATPases in pairs with distinct functional effects, implying an ordered reaction cycle. Cell 144, 526-538 (2011).
62. Aubin-Tam, M. E., Olivares, A. O., Sauer, R. T., Baker, T. A. & Lang, M. J. Single-molecule protein unfolding and translocation by an ATP-fueled proteolytic machine. Cell 145, 257-267 (2011).
63. Maillard, R. A. et al. ClpX(P) generates mechanical force to unfold and translocate its protein substrates. Cell 145, 459-469 (2011).
64. Sen, M. et al. The ClpXP protease unfolds substrates using a constant rate of pulling but different gears. Cell 155, 636-646 (2013).
65. Cordova, J. C. et al. Stochastic but highly coordinated protein unfolding and translocation by the ClpXP proteolytic machine. Cell 158, 647-658 (2014).
66. Kenniston, J. A., Baker, T. A., Fernandez, J. M. & Sauer, R. T. Linkage between ATP consumption and mechanical unfolding during the protein processing reactions of an AAA+ degradation machine. Cell 114, 511-520 (2003).
67. Lee, C., Schwartz, M. P., Prakash, S., Iwakura, M. & Matouschek, A. ATP-dependent proteases degrade their substrates by processively unraveling them from the degradation signal. Mol. Cell 7, 627-637 (2001). This is the first study to demonstrate that AAA+ proteases catalyse unfolding by processively unravelling substrates from the engagement tag, with the stability of adjacent local secondary structure having an important role in degradation susceptibility.
68. Martin, A., Baker, T. A. & Sauer, R. T. Diverse pore loops of the AAA+ ClpX machine mediate unassisted and adaptor-dependent recognition of ssrA-tagged substrates. Mol. Cell 29, 441-450 (2008).
69. Martin, A., Baker, T. A. & Sauer, R. T. Pore loops of the AAA+ ClpX machine grip substrates to drive translocation and unfolding. Nat. Struct. Mol. Biol. 15, 1147-1151 (2008).
70. Iosefson, O., Nager, A. R., Baker, T. A. & Sauer, R. T. Coordinated gripping of substrate by subunits of an AAA+ proteolytic machine. Nat. Chem. Biol. 11, 201-206 (2015).
71. Iosefson, O., Olivares, A. O., Baker, T. A. & Sauer, R. T. Dissection of axial-pore loop function during unfolding and translocation by a AAA+ proteolytic machine. Cell Rep. 12, 1032-1041 (2015).
72. Barkow, S. R., Levchenko, I., Baker, T. A. & Sauer, R. T. Polypeptide translocation by the AAA+ ClpXP protease machine. Chem. Biol. 16, 605-612 (2009).
73. Kenniston, J. A., Baker, T. A. & Sauer, R. T. Partitioning between unfolding and release of native domains during ClpXP degradation determines substrate selectivity and partial processing. Proc. Natl Acad. Sci. USA 102, 1390-1395 (2005).
74. Flynn, J. M., Neher, S. B., Kim, Y. I., Sauer, R. T. & Baker, T. A. Proteomic discovery of cellular substrates of the ClpXP protease reveals five classes of ClpXrecognition signals. Mol. Cell 11, 671-683 (2003).
75. Burton, R. E., Siddiqui, S. M., Kim, Y. I., Baker, T. A. & Sauer, R. T. Effects of protein stability and structure on substrate processing by the ClpXP unfolding and degradation machine. EMBO J. 20, 3092-3100 (2001).
76. Bolon, D. N., Grant, R. A., Baker, T. A. & Sauer, R. T. Nucleotide-dependent substrate handoff from the SspB adaptor to the AAA+ ClpXP protease. Mol. Cell 16, 343-350 (2004).
77. Martin, A., Baker, T. A. & Sauer, R. T. Protein unfolding by a AAA+ protease is dependent on ATPhydrolysis rates and substrate energy landscapes. Nat. Struct. Mol. Biol. 15, 139-145 (2008).
78. Koodathingal, P. et al. ATP-dependent proteases differ substantially in their ability to unfold globular proteins. J. Biol. Chem. 284, 18674-18684 (2009).
79. Vass, R. H. & Chien, P. Critical clamp loader processing by an essential AAA+ protease in Caulobacter crescentus. Proc. Natl Acad. Sci. USA 110, 18138-18143 (2013). This study reports the discovery of an unexpected mode of partial proteolytic processing by ClpXP that generates DNA-clamp loader isoforms required for C. crescentus viability.
80. Bachmair, A., Finley, D. & Varshavsky, A. In vivo half-life of a protein is a function of its amino-terminal residue. Science 234, 179-186 (1986).
81. Hoppe, T. et al. Activation of a membrane-bound transcription factor by regulated ubiquitin/ proteasome-dependent processing. Cell 102, 577-586 (2000).
82. Tian, L., Holmgren, R. A. & Matouschek, A. A conserved processing mechanism regulates the activity of transcription factors Cubitus interruptus and NF-κB. Nat. Struct. Mol. Biol. 12, 1045-1053 (2005).
83. Nager, A. R., Baker, T. A. & Sauer, R. T. Stepwise unfolding of a β barrel protein by the AAA+ ClpXP protease. J. Mol. Biol. 413, 4-16 (2011).
84. Kress, W., Mutschler, H. & Weber-Ban, E. Both ATPase domains of ClpA are critical for processing of stable protein structures. J. Biol. Chem. 284, 31441-31452 (2009).
85. Hinnerwisch, J. et al. Loops in the central channel of ClpA chaperone mediate protein binding, unfolding, and translocation. Cell 121, 1029-1041 (2005).
86. Baytshtok, V., Baker, T. A. & Sauer, R. T. Assaying the kinetics of protein denaturation catalyzed by AAA+ unfolding machines and proteases. Proc. Natl Acad. Sci. USA 112, 5377-5382 (2015).
87. Olivares, A. O., Nager, A. R., Iosefson, O., Sauer, R. T. & Baker, T. A. Mechanochemical basis of protein degradation by a double-ring AAA+ machine. Nat. Struct. Mol. Biol. 21, 871-875 (2014). This study used optical-trapping experiments to reveal similarities and differences in the mechanical unfolding and translocation activities of single-ring and double-ring AAA+ partners of ClpP.
88. Flynn, J. M. et al. Overlapping recognition determinants within the ssrA degradation tag allow modulation of proteolysis. Proc. Natl Acad. Sci. USA 98, 10584-10589 (2001).
89. Ito, K. & Akiyama, Y. Cellular functions, mechanism of action, and regulation of FtsH protease. Annu. Rev. Microbiol. 59, 211-231 (2005).
90. Gerdes, F., Tatsuta, T. & Langer, T. Mitochondrial AAA proteases-towards a molecular understanding of membrane-bound proteolytic machines. Biochim. Biophys. Acta 1823, 49-55 (2012).
91. Vieux, E. F., Wohlever, M. L., Chen, J. Z., Sauer, R. T. & Baker, T. A. Distinct quaternary structures of the AAA+ Lon protease control substrate degradation. Proc. Natl Acad. Sci. USA 110, E2002-E2008 (2013).
92. Gur, E. & Sauer, R. T. Degrons in protein substrates program the speed and operating efficiency of the AAA+ Lon proteolytic machine. Proc. Natl Acad. Sci. USA 106, 18503-18508 (2009).
93. Schmitt, E. K. et al. The natural product cyclomarin kills Mycobacterium tuberculosis by targeting the ClpC1 subunit of the caseinolytic protease. Angew. Chem. Int. Ed. Engl. 50, 5889-5891 (2011).
94. Burton, R. E., Baker, T. A. & Sauer, R. T. Nucleotidedependent substrate recognition by the AAA+ HslUV protease. Nat. Struct. Mol. Biol. 12, 245-251 (2005).
95. Wu, W. F., Zhou, Y. & Gottesman, S. Redundant in vivo proteolytic activities of Escherichia coli Lon and the ClpYQ (HslUV) protease. J. Bacteriol. 181, 3681-3687 (1999).
96. Schrader, E. K., Harstad, K. G. & Matouschek, A. Targeting proteins for degradation. Nat. Chem. Biol. 5, 815-822 (2009).
97. Burton, B. M. & Baker, T. A. Remodeling protein complexes: insights from the AAA+ unfoldase ClpX and Mu transposase. Protein Sci. 14, 1945-1954 (2005).
98. Ling, L., Montao, S. P., Sauer, R. T., Rice, P. A. & Baker, T. A. Deciphering the roles of multicomponent recognition signals by the AAA+ unfoldase ClpX. J. Mol. Biol. 427, 2966-2982 (2015).
99. Kapitein, N. et al. ClpV recycles VipA/VipB tubules and prevents non-productive tubule formation to ensure efficient type VI protein secretion. Mol. Microbiol. 87, 1013-1028 (2013).
100. Winkler, J., Tyedmers, J., Bukau, B. & Mogk, A. Chaperone networks in protein disaggregation and prion propagation. J. Struct. Biol. 179, 152-160 (2012).
101. Roll-Mecak, A. & McNally, F. J. Microtubule-severing enzymes. Curr. Opin. Cell Biol. 22, 96-103 (2010).
102. Camberg, J. L., Viola, M. G., Rea, L., Hoskins, J. R. & Wickner, S. Location of dual sites in E. coli FtsZ important for degradation by ClpXP; one at the C-terminus and one in the disordered linker. PLoS ONE 9, e94964 (2014).
103. Kardon, J. R. et al. Mitochondrial ClpX activates a key enzyme for heme biosynthesis and erythropoiesis. Cell 161, 858-867 (2015). This study shows that mitochondrial ClpX remodels an enzyme required for haem biosynthesis to accelerate the rate of cofactor insertion and regulate activity.
104. Zhao, M. et al. Mechanistic insights into the recycling machine of the SNARE complex. Nature 518, 61-67 (2015). This paper reports the cryo-EM structures of NSF in different nucleotide states that reveal dramatic changes in conformation that may explain protein complex disassembly.
105. Ryu, J.-K. et al. Spring-loaded unraveling of a single SNARE complex by NSF in one round of ATP turnover. Science 347, 1485-1489 (2015).
106. Li, T. et al. Escherichia coli ClpB is a non-processive polypeptide translocase. Biochem. J. 470, 39-52 (2015).
107. Sassetti, C. M., Boyd, D. H. & Rubin, E. J. Genes required for mycobacterial growth defined by high density mutagenesis. Mol. Microbiol. 48, 77-84 (2003).
108. Raju, R. M. et al. Mycobacterium tuberculosis ClpP1 and ClpP2 function together in protein degradation and are required for viability in vitro and during infection. PLoS Pathog. 8, e1002511 (2012).