Protein post-translational modifications in bacteria

Article metrics

Abstract

Over the past decade the number and variety of protein post-translational modifications that have been detected and characterized in bacteria have rapidly increased. Most post-translational protein modifications occur in a relatively low number of bacterial proteins in comparison with eukaryotic proteins, and most of the modified proteins carry low, substoichiometric levels of modification; therefore, their structural and functional analysis is particularly challenging. The number of modifying enzymes differs greatly among bacterial species, and the extent of the modified proteome strongly depends on environmental conditions. Nevertheless, evidence is rapidly accumulating that protein post-translational modifications have vital roles in various cellular processes such as protein synthesis and turnover, nitrogen metabolism, the cell cycle, dormancy, sporulation, spore germination, persistence and virulence. Further research of protein post-translational modifications will fill current gaps in the understanding of bacterial physiology and open new avenues for treatment of infectious diseases.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Fig. 1: Protein modifications in bacteria.
Fig. 2: Phosphorylation, lysine acetylation and succinylation.
Fig. 3: The prokaryotic ubiquitin-like protein (Pup)–proteasome system.
Fig. 4: Role of Hanks-type kinases in cell division and morphogenesis and developmental behaviours.
Fig. 5: Summary of known bacterial PTMs and their involvement in cellular physiology.

References

  1. 1.

    Ree, R., Varland, S. & Arnesen, T. Spotlight on protein N-terminal acetylation. Exp. Mol. Med. 50, 90 (2018).

  2. 2.

    Latousakis, D. & Juge, N. How sweet are our gut beneficial bacteria? A focus on protein glycosylation in Lactobacillus. Int. J. Mol. Sci. 19, 136 (2018).

  3. 3.

    Faridmoayer, A., Fentabil, M. A., Mills, D. C., Klassen, J. S. & Feldman, M. F. Functional characterization of bacterial oligosaccharyltransferases involved in O-linked protein glycosylation. J. Bacteriol. 189, 8088–8098 (2007).

  4. 4.

    Nita-Lazar, M., Wacker, M., Schegg, B., Amber, S. & Aebi, M. The N-X-S/T consensus sequence is required but not sufficient for bacterial N-linked protein glycosylation. Glycobiology 15, 361–367 (2005).

  5. 5.

    Pearce, M. J., Mintseris, J., Ferreyra, J., Gygi, S. P. & Darwin, K. H. Ubiquitin-like protein involved in the proteasome pathway of Mycobacterium tuberculosis. Science 322, 1104–1107 (2008). This is the first report of Pup in bacteria, linking its action to proteasome PafA and protein degradation in M. tuberculosis.

  6. 6.

    Greer, E. L. & Shi, Y. Histone methylation: a dynamic mark in health, disease and inheritance. Nat. Rev. Genet. 13, 343–357 (2012).

  7. 7.

    Koyano, F. et al. Ubiquitin is phosphorylated by PINK1 to activate parkin. Nature 510, 162–166 (2014).

  8. 8.

    Loi, V. V., Rossius, M. & Antelmann, H. Redox regulation by reversible protein S-thiolation in bacteria. Front. Microbiol. 6, 187 (2015).

  9. 9.

    Imber, M., Pietrzyk-Brzezinska, A. J. & Antelmann, H. Redox regulation by reversible protein S-thiolation in Gram-positive bacteria. Redox Biol. 20, 130–145 (2019).

  10. 10.

    Olsen, J. V. & Mann, M. Status of large-scale analysis of post-translational modifications by mass spectrometry. Mol. Cell Proteomics 12, 3444–3452 (2013).

  11. 11.

    Macek, B. & Mijakovic, I. Site-specific analysis of bacterial phosphoproteomes. Proteomics 11, 3002–3011 (2011).

  12. 12.

    Macek, B. et al. Phosphoproteome analysis of E. coli reveals evolutionary conservation of bacterial Ser/Thr/Tyr phosphorylation. Mol. Cell Proteomics 7, 299–307 (2008). This article reports one of the early applications of shot-gun proteomics to global analysis of bacterial phosphoproteins.

  13. 13.

    Potel, C. M., Lin, M. H., Heck, A. J. R. & Lemeer, S. Widespread bacterial protein histidine phosphorylation revealed by mass spectrometry-based proteomics. Nat. Methods 15, 187–190 (2018).

  14. 14.

    Schmidt, A. et al. Quantitative phosphoproteomics reveals the role of protein arginine phosphorylation in the bacterial stress response. Mol. Cell. Proteomics 13, 537–550 (2014).

  15. 15.

    Trentini, D. B. et al. Arginine phosphorylation marks proteins for degradation by a Clp protease. Nature 539, 48–53 (2016). This landmark article demonstrates that in Gram-positive bacteria phosphoarginine functions as a degradation tag for the ClpC–ClpP protease.

  16. 16.

    Elsholz, A. K. et al. Global impact of protein arginine phosphorylation on the physiology of Bacillus subtilis. Proc. Natl Acad. Sci. USA 109, 7451–7456 (2012).

  17. 17.

    Junker, S. et al. Spectral library based analysis of arginine phosphorylations in Staphylococcus aureus. Mol. Cell. Proteomics 17, 335–348 (2018).

  18. 18.

    Suskiewicz, M. J. et al. Structure of McsB, a protein kinase for regulated arginine phosphorylation. Nat. Chem. Biol. 5, 510–518 (2019).

  19. 19.

    Mijakovic, I., Grangeasse, C. & Turgay, K. Exploring the diversity of protein modifications: special bacterial phosphorylation systems. FEMS Microbiol. Rev. 40, 398–417 (2016).

  20. 20.

    Whitmore, S. E. & Lamont, R. J. Tyrosine phosphorylation and bacterial virulence. Int. J. Oral Sci. 4, 1–6 (2012).

  21. 21.

    Rajagopalan, K. & Dworkin, J. Identification and biochemical characterization of a novel protein phosphatase 2C-like Ser/Thr phosphatase in Escherichia coli. J. Bacteriol. 200, e00225–18 (2018).

  22. 22.

    Gross, R., Arico, B. & Rappuoli, R. Families of bacterial signal-transducing proteins. Mol. Microbiol. 3, 1661–1667 (1989).

  23. 23.

    Yuan, J., Jin, F., Glatter, T. & Sourjik, V. Osmosensing by the bacterial PhoQ/PhoP two-component system. Proc. Natl Acad. Sci. USA 114, E10792–E10798 (2017).

  24. 24.

    Deng, L. et al. Characterization of a two-component system transcriptional regulator LtdR that impacts group B streptococcal colonization and disease. Infect. Immun. https://doi.org/10.1128/IAI.00822-17 (2018).

  25. 25.

    Namugenyi, S. B., Aagesen, A. M., Elliott, S. R. & Tischler, A. D. Mycobacterium tuberculosis PhoY proteins promote persister formation by mediating Pst/SenX3-RegX3 phosphate sensing. MBio 8, e00494–17 (2017).

  26. 26.

    Vo, C. D. et al. Repurposing Hsp90 inhibitors as antibiotics targeting histidine kinases. Bioorg. Med. Chem. Lett. 27, 5235–5244 (2017).

  27. 27.

    Bae, H. J. et al. Inhibition of the DevSR Two-component system by overexpression of Mycobacterium tuberculosis PknB in Mycobacterium smegmatis. Mol. Cells 40, 632–642 (2017).

  28. 28.

    Libby, E. A., Goss, L. A. & Dworkin, J. The eukaryotic-like Ser/Thr kinase PrkC regulates the essential WalRK two-component system in Bacillus subtilis. PLOS Genet. 11, e1005275 (2015).

  29. 29.

    Fuhs, S. R. et al. Monoclonal 1- and 3-phosphohistidine antibodies: new tools to study histidine phosphorylation. Cell 162, 198–210 (2015).

  30. 30.

    Kee, J. M., Oslund, R. C., Perlman, D. H. & Muir, T. W. A pan-specific antibody for direct detection of protein histidine phosphorylation. Nat. Chem. Biol. 9, 416–421 (2013).

  31. 31.

    Stancik, I. A. et al. Serine/threonine protein kinases from bacteria, archaea and eukarya share a common evolutionary origin deeply rooted in the tree of life. J. Mol. Biol. 430, 27–32 (2018). This article clarifies the classification and terminology of the Hanks-type family of protein kinases on the basis of phylogenomic evidence (phylostratigraphy).

  32. 32.

    Pereira, S. F., Goss, L. & Dworkin, J. Eukaryote-like serine/threonine kinases and phosphatases in bacteria. Microbiol. Mol. Biol. Rev. 75, 192–212 (2011).

  33. 33.

    Mijakovic, I. & Macek, B. Impact of phosphoproteomics on studies of bacterial physiology. FEMS Microbiol. Rev. 36, 877–892 (2012).

  34. 34.

    Amin, R. et al. Post-translational serine/threonine phosphorylation and lysine acetylation: a novel regulatory aspect of the global nitrogen response regulator GlnR in S. coelicolor M145. Front. Mol. Biosci. 3, 38 (2016).

  35. 35.

    Yadav, G. S., Ravala, S. K., Malhotra, N. & Chakraborti, P. K. Phosphorylation modulates catalytic activity of mycobacterial sirtuins. Front. Microbiol. 7, 677 (2016).

  36. 36.

    Canova, M. J. & Molle, V. Bacterial serine/threonine protein kinases in host-pathogen interactions. J. Biol. Chem. 289, 9473–9479 (2014).

  37. 37.

    Lee, W. L. et al. Mechanisms of Yersinia YopO kinase substrate specificity. Sci. Rep. 7, 39998 (2017).

  38. 38.

    Fuhrmann, J. et al. McsB is a protein arginine kinase that phosphorylates and inhibits the heat-shock regulator CtsR. Science 324, 1323–1327 (2009).

  39. 39.

    Fuhrmann, J., Subramanian, V. & Thompson, P. R. Targeting the arginine phosphatase YwlE with a catalytic redox-based inhibitor. ACS Chem. Biol. 8, 2024–2032 (2013).

  40. 40.

    Fuhrmann, J., Subramanian, V., Kojetin, D. J. & Thompson, P. R. Activity-based profiling reveals a regulatory link between oxidative stress and protein arginine phosphorylation. Cell. Chem. Biol. 23, 967–977 (2016).

  41. 41.

    Zhou, B. et al. Arginine dephosphorylation propels spore germination in bacteria. Proc. Natl Acad. Sci. USA https://doi.org/10.1073/pnas.1817742116 (2019).

  42. 42.

    Weinert, B. T. et al. Acetyl-phosphate is a critical determinant of lysine acetylation in E. coli. Mol. Cell 51, 265–272 (2013). This landmark study detects widespread lysine acetylation of bacterial proteins and proves the non-enzymatic origin of most lysine acetylation events.

  43. 43.

    Weinert, B. T. et al. Lysine succinylation is a frequently occurring modification in prokaryotes and eukaryotes and extensively overlaps with acetylation. Cell Rep. 4, 842–851 (2013).

  44. 44.

    Schilling, B. et al. Protein acetylation dynamics in response to carbon overflow in Escherichia coli. Mol. Microbiol. 98, 847–863 (2015).

  45. 45.

    Ghosh, S., Padmanabhan, B., Anand, C. & Nagaraja, V. Lysine acetylation of the Mycobacterium tuberculosis HU protein modulates its DNA binding and genome organization. Mol. Microbiol. 100, 577–588 (2016).

  46. 46.

    Tu, S. et al. YcgC represents a new protein deacetylase family in prokaryotes. eLife 4, e05322 (2015).

  47. 47.

    Pan, J., Chen, R., Li, C., Li, W. & Ye, Z. Global analysis of protein lysine succinylation profiles and their overlap with lysine acetylation in the marine bacterium Vibrio parahemolyticus. J. Proteome Res. 14, 4309–4318 (2015).

  48. 48.

    Colak, G. et al. Identification of lysine succinylation substrates and the succinylation regulatory enzyme CobB in Escherichia coli. Mol. Cell. Proteomics 12, 3509–3520 (2013).

  49. 49.

    Wolfe, A. J. Bacterial protein acetylation: new discoveries unanswered questions. Curr. Genet. 62, 335–341 (2016).

  50. 50.

    Weinert, B. T. et al. Accurate quantification of site-specific acetylation stoichiometry reveals the impact of sirtuin deacetylase CobB on the E. coli acetylome. Mol. Cell. Proteomics 16, 759–769 (2017).

  51. 51.

    Carabetta, V. J. & Cristea, I. M. Regulation, function, and detection of protein acetylation in bacteria. J. Bacteriol. 199, e00107–17 (2017).

  52. 52.

    Ouidir, T., Cosette, P., Jouenne, T. & Hardouin, J. Proteomic profiling of lysine acetylation in Pseudomonas aeruginosa reveals the diversity of acetylated proteins. Proteomics 15, 2152–2157 (2015).

  53. 53.

    Ouidir, T., Jarnier, F., Cosette, P., Jouenne, T. & Hardouin, J. Extracellular Ser/Thr/Tyr phosphorylated proteins of pseudomonas aeruginosa PA14 strain. Proteomics 14, 2017–2030 (2014).

  54. 54.

    Ouidir, T., Jarnier, F., Cosette, P., Jouenne, T. & Hardouin, J. Potential of liquid-isoelectric-focusing protein fractionation to improve phosphoprotein characterization of Pseudomonas aeruginosa PA14. Anal. Bioanal. Chem. 406, 6297–6309 (2014).

  55. 55.

    Soares, N. C. et al. Ser/Thr/Tyr phosphoproteome characterization of Acinetobacter baumannii: comparison between a reference strain and a highly invasive multidrug-resistant clinical isolate. J. Proteomics 102, 113–124 (2014).

  56. 56.

    Kentache, T., Jouenne, T., De, E. & Hardouin, J. Proteomic characterization of Nα- and Nε-acetylation in Acinetobacter baumannii. J. Proteomics 144, 148–158 (2016).

  57. 57.

    Gaviard, C. et al. Lysine succinylation and acetylation in pseudomonas aeruginosa. J. Proteome Res. 17, 2449–2459 (2018).

  58. 58.

    Amato, S. M. et al. The role of metabolism in bacterial persistence. Front. Microbiol. 5, 70 (2014).

  59. 59.

    Striebel, F. et al. Bacterial ubiquitin-like modifier Pup is deamidated and conjugated to substrates by distinct but homologous enzymes. Nat. Struct. Mol. Biol. 16, 647–651 (2009).

  60. 60.

    Festa, R. A. et al. Prokaryotic ubiquitin-like protein (Pup) proteome of mycobacterium tuberculosis [corrected]. PLOS ONE 5, e8589 (2010).

  61. 61.

    Cerda-Maira, F. A. et al. Molecular analysis of the prokaryotic ubiquitin-like protein (Pup) conjugation pathway in mycobacterium tuberculosis. Mol. Microbiol. 77, 1123–1135 (2010).

  62. 62.

    Imkamp, F. et al. Dop functions as a depupylase in the prokaryotic ubiquitin-like modification pathway. EMBO Rep. 11, 791–797 (2010).

  63. 63.

    Ozcelik, D. et al. Structures of Pup ligase PafA and depupylase dop from the prokaryotic ubiquitin-like modification pathway. Nat. Commun. 3, 1014, (2012) 10.1038/ncomms2009 (2012).

  64. 64.

    Bolten, M. et al. Depupylase Dop requires inorganic phosphate in the active site for catalysis. J. Biol. Chem. 292, 4044–4053 (2017).

  65. 65.

    Guth, E., Thommen, M. & Weber-Ban, E. Mycobacterial ubiquitin-like protein ligase pafa follows a two-step reaction pathway with a phosphorylated Pup intermediate. J. Biol. Chem. 286, 4412–4419 (2011).

  66. 66.

    Striebel, F., Hunkeler, M., Summer, H. & Weber-Ban, E. The mycobacterial Mpa-proteasome unfolds and degrades pupylated substrates by engaging Pup’s N-terminus. EMBO J. 29, 1262–1271 (2010).

  67. 67.

    Wang, T. et al. Structural insights on the Mycobacterium tuberculosis proteasomal ATPase Mpa. Structure 17, 1377–1385 (2009).

  68. 68.

    Schaffer, C. & Messner, P. Emerging facets of prokaryotic glycosylation. FEMS Microbiol. Rev. 41, 49–91 (2017).

  69. 69.

    Charbonneau, M. E. et al. O-linked glycosylation ensures the normal conformation of the autotransporter adhesin involved in diffuse adherence. J. Bacteriol. 189, 8880–8889 (2007).

  70. 70.

    Logan, S. M. Flagellar glycosylation - a new component of the motility repertoire? Microbiology 152, 1249–1262 (2006).

  71. 71.

    Just, I. et al. Glucosylation of Rho proteins by clostridium difficile toxin B. Nature 375, 500–503 (1995).

  72. 72.

    Wacker, M. et al. N-linked glycosylation in campylobacter jejuni and its functional transfer into E. coli. Science 298, 1790–1793 (2002). Discovery of an N-linked glycosylation system in C. jejuni and pathway transfer to E. coli : a basis for glycol-engineering.

  73. 73.

    Kowarik, M. et al. Definition of the bacterial N-glycosylation site consensus sequence. EMBO J. 25, 1957–1966 (2006).

  74. 74.

    Feldman, M. F. et al. Engineering N-linked protein glycosylation with diverse O antigen lipopolysaccharide structures in Escherichia coli. Proc. Natl Acad. Sci. USA 102, 3016–3021 (2005).

  75. 75.

    Cain, J. A. et al. Proteomics reveals multiple phenotypes associated with N-linked glycosylation in campylobacter jejuni. Mol. Cell. Proteomics 18, 715–734 (2019).

  76. 76.

    Lassak, J. et al. Arginine-rhamnosylation as new strategy to activate translation elongation factor P. Nat. Chem. Biol. 11, 266–270 (2015).

  77. 77.

    Eichler, J. & Koomey, M. Sweet new roles for protein glycosylation in prokaryotes. Trends Microbiol. 25, 662–672 (2017).

  78. 78.

    Guerry, P. et al. Changes in flagellin glycosylation affect campylobacter autoagglutination and virulence. Mol. Microbiol. 60, 299–311 (2006).

  79. 79.

    Champasa, K., Longwell, S. A., Eldridge, A. M., Stemmler, E. A. & Dube, D. H. Targeted identification of glycosylated proteins in the gastric pathogen Helicobacter pylori (Hp). Mol. Cell. Proteomics 12, 2568–2586 (2013).

  80. 80.

    Hanuszkiewicz, A. et al. Identification of the flagellin glycosylation system in Burkholderia cenocepacia and the contribution of glycosylated flagellin to evasion of human innate immune responses. J. Biol. Chem. 289, 19231–19244 (2014).

  81. 81.

    Iwashkiw, J. A. et al. Identification of a general O-linked protein glycosylation system in Acinetobacter baumannii and its role in virulence and biofilm formation. PLOS Pathog. 8, e1002758 (2012).

  82. 82.

    Steinemann, M., Schlosser, A., Jank, T. & Aktories, K. The chaperonin TRiC/CCT is essential for the action of bacterial glycosylating protein toxins like Clostridium difficile toxins A and B. Proc. Natl Acad. Sci. USA 115, 9580–9585 (2018).

  83. 83.

    Parker, J. L. et al. Maf-dependent bacterial flagellin glycosylation occurs before chaperone binding and flagellar T3SS export. Mol. Microbiol. 92, 258–272 (2014).

  84. 84.

    Vik, A. et al. Insights into type IV pilus biogenesis and dynamics from genetic analysis of a C-terminally tagged pilin: a role for O-linked glycosylation. Mol. Microbiol. 85, 1166–1178 (2012).

  85. 85.

    Sankaran, K. & Wu, H. C. Lipid modification of bacterial prolipoprotein. Transfer of diacylglyceryl moiety from phosphatidylglycerol. J. Biol. Chem. 269, 19701–19706 (1994).

  86. 86.

    Nakayama, H., Kurokawa, K. & Lee, B. L. Lipoproteins in bacteria: structures and biosynthetic pathways. FEBS J. 279, 4247–4268 (2012).

  87. 87.

    Issartel, J. P., Koronakis, V. & Hughes, C. Activation of Escherichia coli prohaemolysin to the mature toxin by acyl carrier protein-dependent fatty acylation. Nature 351, 759–761 (1991).

  88. 88.

    Sobocinska, J., Roszczenko-Jasinska, P., Ciesielska, A. & Kwiatkowska, K. Protein palmitoylation and its role in bacterial and viral infections. Front. Immunol. 8, 2003 (2017).

  89. 89.

    Bray, B. A., Sutcliffe, I. C. & Harrington, D. J. Impact of lgt mutation on lipoprotein biosynthesis and in vitro phenotypes of Streptococcus agalactiae. Microbiology 155, 1451–1458 (2009).

  90. 90.

    Spera, J. M., Guaimas, F., Corvi, M. M. & Ugalde, J. E. Brucella hijacks host-mediated palmitoylation to stabilize and localize PrpA to the plasma membrane. Infect. Immun. 86, e00402–18 (2018).

  91. 91.

    Hang, H. C. et al. Chemical probes for the rapid detection of fatty-acylated proteins in mammalian cells. J. Am. Chem. Soc. 129, 2744–2745 (2007).

  92. 92.

    Roth, A. F. et al. Global analysis of protein palmitoylation in yeast. Cell 125, 1003–1013 (2006).

  93. 93.

    Charlton, T. M., Kovacs-Simon, A., Michell, S. L., Fairweather, N. F. & Tate, E. W. quantitative lipoproteomics in Clostridium difficile reveals a role for lipoproteins in sporulation. Chem. Biol. 22, 1562–1573 (2015).

  94. 94.

    Rosenberg, A. et al. Phosphoproteome dynamics mediate revival of bacterial spores. BMC Biol. 13, 76 (2015).

  95. 95.

    Petrickova, K. & Petricek, M. Eukaryotic-type protein kinases in Streptomyces coelicolor: variations on a common theme. Microbiology 149, 1609–1621 (2003).

  96. 96.

    Manteca, A., Ye, J., Sanchez, J. & Jensen, O. N. Phosphoproteome analysis of streptomyces development reveals extensive protein phosphorylation accompanying bacterial differentiation. J. Proteome Res. 10, 5481–5492 (2011).

  97. 97.

    Ladwig, N. et al. Control of morphological differentiation of Streptomyces coelicolor A3(2) by phosphorylation of MreC and PBP2. PLOS ONE 10, e0125425 (2015).

  98. 98.

    Hempel, A. M. et al. The Ser/Thr protein kinase AfsK regulates polar growth and hyphal branching in the filamentous bacteria Streptomyces. Proc. Natl Acad. Sci USA 109, E2371–E2379 (2012). This study, together with Kang et al. (1999), highlights that DivIVA phosphorylation is a conserved key feature of the bacterial cell cycle.

  99. 99.

    Kang, C. M., Nyayapathy, S., Lee, J. Y., Suh, J. W. & Husson, R. N. Wag31, a homologue of the cell division protein DivIVA, regulates growth, morphology and polar cell wall synthesis in mycobacteria. Microbiology 154, 725–735 (2008). This study shows that DivIVA phosphorylation regulates cell shape and cell wall synthesis.

  100. 100.

    Fleurie, A. et al. Mutational dissection of the S/T-kinase StkP reveals crucial roles in cell division of Streptococcus pneumoniae. Mol. Microbiol. 83, 746–758 (2012).

  101. 101.

    Stein, E. A., Cho, K., Higgs, P. I. & Zusman, D. R. Two Ser/Thr protein kinases essential for efficient aggregation and spore morphogenesis in Myxococcus xanthus. Mol. Microbiol. 60, 1414–1431 (2006).

  102. 102.

    Inouye, S. & Nariya, H. Dual regulation with Ser/Thr kinase cascade and a His/Asp TCS in Myxococcus xanthus. Adv. Exp. Med. Biol. 631, 111–121 (2008).

  103. 103.

    Kimura, Y., Kato, T. & Mori, Y. Function analysis of a bacterial tyrosine kinase, BtkB, in Myxococcus xanthus. FEMS Microbiol. Lett. 336, 45–51 (2012).

  104. 104.

    Kimura, Y., Yamashita, S., Mori, Y., Kitajima, Y. & Takegawa, K. A Myxococcus xanthus bacterial tyrosine kinase, BtkA, is required for the formation of mature spores. J. Bacteriol. 193, 5853–5857 (2011).

  105. 105.

    Bragg, J. et al. Identification and characterization of a putative arginine kinase homolog from Myxococcus xanthus required for fruiting body formation and cell differentiation. J. Bacteriol. 194, 2668–2676 (2012).

  106. 106.

    Giefing, C., Jelencsics, K. E., Gelbmann, D., Senn, B. M. & Nagy, E. The pneumococcal eukaryotic-type serine/threonine protein kinase StkP co-localizes with the cell division apparatus and interacts with FtsZ in vitro. Microbiology 156, 1697–1707 (2010).

  107. 107.

    Sureka, K. et al. Novel role of phosphorylation-dependent interaction between FtsZ and FipA in mycobacterial cell division. PLOS ONE 5, e8590 (2010).

  108. 108.

    Kieser, K. J. et al. Phosphorylation of the peptidoglycan synthase PonA1 governs the rate of polar elongation in mycobacteria. PLOS Pathog. 11, e1005010 (2015).

  109. 109.

    Morlot, C. et al. Interaction of penicillin-binding protein 2x and Ser/Thr protein kinase StkP, two key players in Streptococcus pneumoniae R6 morphogenesis. Mol. Microbiol. 90, 88–102 (2013).

  110. 110.

    Zucchini, L. et al. PASTA repeats of the protein kinase StkP interconnect cell constriction and separation of Streptococcus pneumoniae. Nat. Microbiol. 3, 197–209 (2018). This study illustrates that a Hanks-type kinase can affect cell division without necessarily catalysing the phosphorylation of the interacting partner.

  111. 111.

    Fleurie, A. et al. MapZ marks the division sites and positions FtsZ rings in Streptococcus pneumoniae. Nature 516, 259–262 (2014).

  112. 112.

    Fenton, A. K. et al. Phosphorylation-dependent activation of the cell wall synthase PBP2a in Streptococcus pneumoniae by MacP. Proc. Natl Acad. Sci. USA 115, 2812–2817 (2018).

  113. 113.

    Stamsas, G. A. et al. Identification of EloR (Spr1851) as a regulator of cell elongation in Streptococcus pneumoniae. Mol. Microbiol. 105, 954–967 (2017).

  114. 114.

    Corte, L. et al. A conserved cysteine residue of Bacillus subtilis SpoIIIJ is important for endospore development. PLOS ONE 9, e99811 (2014).

  115. 115.

    Baronian, G. et al. Phosphorylation of Mycobacterium tuberculosis ParB participates in regulating the ParABS chromosome segregation system. PLOS ONE 10, e0119907 (2015).

  116. 116.

    Nourikyan, J. et al. Autophosphorylation of the bacterial tyrosine-kinase cpsd connects capsule synthesis with the cell cycle in Streptococcus pneumoniae. PLOS Genet. 11, e1005518 (2015).

  117. 117.

    Mercy, C. et al. Rocs drives chromosome segregation and nucleoid protection in Streptococcus pneumoniae. Nat. Microbiol. https://doi.org/10.1038/s41564-019-0472-z (2019).

  118. 118.

    Zhou, P., Wong, D., Li, W., Xie, J. & Av-Gay, Y. Phosphorylation of Mycobacterium tuberculosis protein tyrosine kinase A PtkA by Ser/Thr protein kinases. Biochem. Biophys. Res. Commun. 467, 421–426 (2015).

  119. 119.

    Nicolas, P. et al. Condition-dependent transcriptome reveals high-level regulatory architecture in Bacillus subtilis. Science 335, 1103–1106 (2012).

  120. 120.

    Bidnenko, V. et al. Bacillus subtilis serine/threonine protein kinase YabT is involved in spore development via phosphorylation of a bacterial recombinase. Mol. Microbiol. 88, 921–935 (2013).

  121. 121.

    Garcia Garcia, T. et al. Phosphorylation of the Bacillus subtilis replication controller YabA plays a role in regulation of sporulation and biofilm formation. Front. Microbiol. 9, 486 (2018).

  122. 122.

    Shi, L. et al. Cross-phosphorylation of bacterial serine/threonine and tyrosine protein kinases on key regulatory residues. Front. Microbiol. 5, 495 (2014).

  123. 123.

    Arigoni, F., Duncan, L., Alper, S., Losick, R. & Stragier, P. SpoIIE governs the phosphorylation state of a protein regulating transcription factor sigma F during sporulation in Bacillus subtilis. Proc. Natl Acad. Sci. USA 93, 3238–3242 (1996).

  124. 124.

    Pereira, S. F., Gonzalez, R. L. Jr. & Dworkin, J. Protein synthesis during cellular quiescence is inhibited by phosphorylation of a translational elongation factor. Proc. Natl Acad. Sci. USA 112, E3274–3281 (2015).

  125. 125.

    Shah, I. M., Laaberki, M. H., Popham, D. L. & Dworkin, J. A eukaryotic-like Ser/Thr kinase signals bacteria to exit dormancy in response to peptidoglycan fragments. Cell 135, 486–496 (2008).

  126. 126.

    Yeats, C., Finn, R. D. & Bateman, A. The PASTA domain: a beta-lactam-binding domain. Trends Biochem. Sci. 27, 438 (2002).

  127. 127.

    Pompeo, F. et al. Phosphorylation of CpgA protein enhances both its GTPase activity and its affinity for ribosome and is crucial for Bacillus subtilis growth and morphology. J. Biol. Chem. 287, 20830–20838 (2012).

  128. 128.

    Pompeo, F., Foulquier, E., Serrano, B., Grangeasse, C. & Galinier, A. Phosphorylation of the cell division protein GpsB regulates PrkC kinase activity through a negative feedback loop in Bacillus subtilis. Mol. Microbiol. 97, 139–150 (2015).

  129. 129.

    Foulquier, E. et al. PrkC-mediated phosphorylation of overexpressed YvcK protein regulates PBP1 protein localization in Bacillus subtilis mreB mutant cells. J. Biol. Chem. 289, 23662–23669 (2014).

  130. 130.

    Kobir, A. et al. Phosphorylation of Bacillus subtilis gene regulator AbrB modulates its DNA-binding properties. Mol. Microbiol. 92, 1129–1141 (2014).

  131. 131.

    Jers, C., Kobir, A., Sondergaard, E. O., Jensen, P. R. & Mijakovic, I. Bacillus subtilis two-component system sensory kinase DegS is regulated by serine phosphorylation in its input domain. PLOS ONE 6, e14653 (2011).

  132. 132.

    Garcia-Garcia, T. et al. Role of protein phosphorylation in the regulation of cell cycle and DNA-related processes in bacteria. Front. Microbiol. 7, 184 (2016).

  133. 133.

    Nguyen, K. B. et al. Phosphorylation of spore coat proteins by a family of atypical protein kinases. Proc. Natl Acad. Sci. USA 113, E3482–3491 (2016). This study, providing evidence that spore germination is influenced by the phosphorylation of spore coat protein, illustrates the diversity of protein kinases playing a role in a bacterial developmental behaviour.

  134. 134.

    Compton, C. L., Fernandopulle, M. S., Nagari, R. T. & Sello, J. K. Genetic and proteomic analyses of pupylation in Streptomyces coelicolor. J. Bacteriol. 197, 2747–2753 (2015).

  135. 135.

    Fimlaid, K. A. et al. Identification of a novel lipoprotein regulator of Clostridium difficile spore germination. PLOS Pathog. 11, e1005239 (2015).

  136. 136.

    Pisithkul, T., Patel, N. M. & Amador-Noguez, D. Post-translational modifications as key regulators of bacterial metabolic fluxes. Curr. Opin. Microbiol. 24, 29–37 (2015).

  137. 137.

    Jedrzejas, M. J., Chander, M., Setlow, P. & Krishnasamy, G. Mechanism of catalysis of the cofactor-independent phosphoglycerate mutase from Bacillus stearothermophilus. crystal structure of the complex with 2-phosphoglycerate. J. Biol. Chem. 275, 23146–23153 (2000).

  138. 138.

    Satishchandran, C., Hickman, Y. N. & Markham, G. D. Characterization of the phosphorylated enzyme intermediate formed in the adenosine 5′-phosphosulfate kinase reaction. Biochemistry 31, 11684–11688 (1992).

  139. 139.

    Kochanowski, K., Sauer, U. & Noor, E. Posttranslational regulation of microbial metabolism. Curr. Opin. Microbiol. 27, 10–17 (2015).

  140. 140.

    Brunk, E. et al. Characterizing posttranslational modifications in prokaryotic metabolism using a multiscale workflow. Proc. Natl Acad. Sci. USA 115, 11096–11101 (2018).

  141. 141.

    Deutscher, J., Francke, C. & Postma, P. W. How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria. Microbiol. Mol. Biol. Rev. 70, 939–1031 (2006).

  142. 142.

    Gorke, B. & Stulke, J. Carbon catabolite repression in bacteria: many ways to make the most out of nutrients. Nat. Rev. Microbiol. 6, 613–624 (2008).

  143. 143.

    Forchhammer, K. & Lüddecke, J. Sensory properties of the PII signalling protein family. FEBS J. 283, 425–437 (2016).

  144. 144.

    Huergo, L. F. & Dixon, R. The emergence of 2-oxoglutarate as a master regulator metabolite. Microbiol. Mol. Biol. Rev. 79, 419–435 (2015).

  145. 145.

    Merrick, M. Post-translational modification of P II signal transduction proteins. Front. Microbiol. 5, 763 (2014).

  146. 146.

    Elharar, Y. et al. Survival of mycobacteria depends on proteasome-mediated amino acid recycling under nutrient limitation. EMBO J. 33, 1802–1814 (2014).

  147. 147.

    Samanovic, M. I. et al. Proteasomal control of cytokinin synthesis protects Mycobacterium tuberculosis against nitric oxide. Mol. Cell 57, 984–994 (2015).

  148. 148.

    Fascellaro, G. et al. Comprehensive proteomic analysis of nitrogen-starved Mycobacterium smegmatis Δpup reveals the impact of pupylation on nitrogen stress response. J. Proteome Res. 15, 2812–2825 (2016).

  149. 149.

    Muller, A. U., Imkamp, F. & Weber-Ban, E. The mycobacterial LexA/RecA-independent DNA damage response is controlled by PafBC and the Pup-proteasome system. Cell Rep. 23, 3551–3564 (2018).

  150. 150.

    Kuberl, A., Polen, T. & Bott, M. The pupylation machinery is involved in iron homeostasis by targeting the iron storage protein ferritin. Proc. Natl Acad. Sci. USA 113, 4806–4811 (2016).

  151. 151.

    Thao, S., Chen, C. S., Zhu, H. & Escalante-Semerena, J. C. Nε-Lysine acetylation of a bacterial transcription factor inhibits its DNA-binding activity. PLOS ONE 5, e15123 (2010).

  152. 152.

    Yang, H. et al. Lysine acetylation of DosR regulates the hypoxia response of mycobacterium tuberculosis. Emerg. Microbes Infect. 7, 34 (2018).

  153. 153.

    Munita, J. M. & Arias, C. A. Mechanisms of antibiotic resistance. Microbiol. Spectr. https://doi.org/10.1128/microbiolspec.VMBF-0016-2015 (2016).

  154. 154.

    Balaban, N. Q. Persistence: mechanisms for triggering and enhancing phenotypic variability. Curr. Opin. Genet. Dev. 21, 768–775 (2011).

  155. 155.

    Lewis, K. Persister cells. Annu. Rev. Microbiol. 64, 357–372 (2010).

  156. 156.

    Moyed, H. S. & Bertrand, K. P. hipA, a newly recognized gene of Escherichia coli K-12 that affects frequency of persistence after inhibition of murein synthesis. J. Bacteriol. 155, 768–775 (1983).

  157. 157.

    Schumacher, M. A. et al. HipBA–promoter structures reveal the basis of heritable multidrug tolerance. Nature 524, 59 (2015).

  158. 158.

    Germain, E., Castro-Roa, D., Zenkin, N. & Gerdes, K. Molecular mechanism of bacterial persistence by HipA. Mol. Cell 52, 248–254 (2013). This study, together with Kaspy et al. (2013), characterizes the molecular mechanism of the serine/threonine kinase HipA in bacterial persistence and identifies GltX as its main substrate.

  159. 159.

    Kaspy, I. et al. HipA-mediated antibiotic persistence via phosphorylation of the glutamyl-tRNA-synthetase. Nat. Commun. 4, 3001 (2013).

  160. 160.

    Semanjski, M. et al. The kinases HipA and HipA7 phosphorylate different substrate pools in eEcherichia coli to promote multidrug tolerance. Sci. Signal. 11, eaat5750 (2018).

  161. 161.

    Vang Nielsen, S. et al. Serine-threonine kinases encoded by split hipA homologs inhibit tryptophanyl-tRNA synthetase. MBio 10, e01138–19 (2019).

  162. 162.

    Veyron, S. et al. A Ca2+-regulated deAMPylation switch in human and bacterial FIC proteins. Nat. Commun. 10, 1142 (2019).

  163. 163.

    da Silva, R. A. G. et al. The role of apolipoprotein N-acyl transferase, Lnt, in the lipidation of factor H binding protein of Neisseria meningitidis strain MC58 and its potential as a drug target. Br. J. Pharmacol. 174, 2247–2260 (2017).

  164. 164.

    Nguyen, J. Q., Gilley, R. P., Zogaj, X., Rodriguez, S. A. & Klose, K. E. Lipidation of the FPI protein IglE contributes to Francisella tularensis ssp. novicida intramacrophage replication and virulence. Pathog. Dis. 72, 10–18 (2014).

  165. 165.

    Wenzel, M. et al. Influence of lipidation on the mode of action of a small RW-rich antimicrobial peptide. Biochim. Biophys. Acta 1858, 1004–1011 (2016).

  166. 166.

    Standish, A. J. et al. Unprecedented abundance of protein tyrosine phosphorylation modulates shigella flexneri virulence. J. Mol. Biol. 428, 4197–4208 (2016).

  167. 167.

    Wong, D. et al. Protein tyrosine kinase, PtkA, is required for Mycobacterium tuberculosis growth in macrophages. Sci. Rep. 8, 155 (2018).

  168. 168.

    Rieck, B. et al. PknG senses amino acid availability to control metabolism and virulence of Mycobacterium tuberculosis. PLOS Pathog. 13, e1006399 (2017).

  169. 169.

    Pensinger, D. A. et al. The listeria monocytogenes PASTA kinase PrkA and its substrate YvcK are required for cell wall homeostasis, metabolism, and virulence. PLOS Pathog. 12, e1006001 (2016).

Download references

Acknowledgements

The authors thank A. Velic, P. Spät and M. Semanjski for help with the preparation of the manuscript. B.M. was supported by grants from the Deutsche Forschungsgemeinschaft (German Research Foundation Cluster of Excellence EXC 2124, SFB 766, FOR 2816, TRR 261) and the German–Israeli Foundation (I-1464–416.13/2018). I.M. was supported by grants from the Swedish Research Council and the Novo Nordisk Foundation (NNF10CC1016517). K.F. was supported by grants from the Deutsche Forschungsgemeinschaft (EXC 2124, SFB 766, GRK 1708, FOR 2816). C.G. was supported by grants from the CNRS, the ANR (ANR-15-CE32–01, ANR-18-CE11–0017–02) and the Bettencourt Schueller Foundation. E.W.-B. was supported by the Swiss National Science Foundation (31003A, 163314).

Author information

B.M. researched data for the article, designed the outline and reviewed and edited the manuscript before submission. B.M., K.F., J.H, E.W.-B., C.G. and I.M. contributed substantially to the discussion of the content and wrote the article.

Correspondence to Boris Macek.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information

Nature Reviews Microbiology thanks T. Clausen and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Glossary

Phosphoproteome

All proteins of an organism, tissue or a cell that contain at least one phosphate group.

Two-component systems

(TCS). Stimulus–response coupling mechanisms that enable organisms to sense and respond to changes in different environmental conditions. They typically consist of a membrane-bound histidine kinase that senses a specific environmental stimulus and a corresponding response regulator that mediates the cellular response.

Sirtuin

Protein deacetylases that couple lysine deacetylation to NAD hydrolysis, yielding O-acetyl-ADP-ribose, the deacetylated substrate and nicotinamide.

Fruiting bodies

Aerial structures composed of aligned chains of attached Bacillus subtilis cells that function as preferential sites for sporulation.

Divisome

A contractile ring of proteins forming around the circumference of the midpoint of the cell at the time of division and mediating the formation of a septum.

Muropeptides

Polymers of glycan and peptides found in bacterial cell walls (also termed ‘peptidoglycans’).

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark