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Formation and function of bacterial organelles


Advances in imaging technologies have revealed that many bacteria possess organelles with a proteomically defined lumen and a macromolecular boundary. Some are bound by a lipid bilayer (such as thylakoids, magnetosomes and anammoxosomes), whereas others are defined by a lipid monolayer (such as lipid bodies), a proteinaceous coat (such as carboxysomes) or have a phase-defined boundary (such as nucleolus-like compartments). These diverse organelles have various metabolic and physiological functions, facilitating adaptation to different environments and driving the evolution of cellular complexity. This Review highlights that, despite the diversity of reported organelles, some unifying concepts underlie their formation, structure and function. Bacteria have fundamental mechanisms of organelle formation, through which conserved processes can form distinct organelles in different species depending on the proteins recruited to the luminal space and the boundary of the organelle. These complex subcellular compartments provide evolutionary advantages as well as enabling metabolic specialization, biogeochemical processes and biotechnological advances. Growing evidence suggests that the presence of organelles is the rule, rather than the exception, in bacterial cells.

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Fig. 1: Structural and functional diversity of bacterial organelles.
Fig. 2: Formation of membrane-bound organelles.
Fig. 3


  1. 1.

    Saier, M. H. Jr. Microcompartments and protein machines in prokaryotes. J. Mol. Microbiol. Biotechnol. 23, 243–269 (2013).

    CAS  Google Scholar 

  2. 2.

    Cheng, S., Liu, Y., Crowley, C. S., Yeates, T. O. & Bobik, T. A. Bacterial microcompartments: their properties and paradoxes. Bioessays 30, 1084–1095 (2008).

    CAS  Google Scholar 

  3. 3.

    Yeates, T. O., Thompson, M. C. & Bobik, T. A. The protein shells of bacterial microcompartment organelles. Curr. Opin. Struct. Biol. 21, 223–231 (2011).

    CAS  Google Scholar 

  4. 4.

    Kerfeld, C. A. & Erbilgin, O. Bacterial microcompartments and the modular construction of microbial metabolism. Trends Microbiol. 23, 22–34 (2015).

    CAS  Google Scholar 

  5. 5.

    Weber, S. C., Spakowitz, A. J. & Theriot, J. A. Bacterial chromosomal loci move subdiffusively through a viscoelastic cytoplasm. Phys. Rev. Lett. 104, 238102 (2010).

    Google Scholar 

  6. 6.

    Wang, W., Li, G.-W., Chen, C., Xie, X. S. & Zhuang, X. Chromosome organization by a nucleoid-associated protein in live bacteria. Science 333, 1445–1449 (2011).

    CAS  Google Scholar 

  7. 7.

    Coltharp, C. & Xiao, J. Superresolution microscopy for microbiology. Cell. Microbiol. 14, 1808–1818 (2012).

    CAS  Google Scholar 

  8. 8.

    Chaikeeratisak, V. et al. Assembly of a nucleus-like structure during viral replication in bacteria. Science 355, 194–197 (2017).

    CAS  Google Scholar 

  9. 9.

    Fishov, I. & Norris, V. Membrane heterogeneity created by transertion is a global regulator in bacteria. Curr. Opin. Microbiol. 15, 724–730 (2012).

    CAS  Google Scholar 

  10. 10.

    Kannaiah, S. & Amster-Choder, O. Protein targeting via mRNA in bacteria. Biochim. Biophys. Acta 1843, 1457–1465 (2014).

    CAS  Google Scholar 

  11. 11.

    Redder, P. How does sub-cellular localization affect the fate of bacterial mRNA? Curr. Genet. 62, 687–690 (2016).

    CAS  Google Scholar 

  12. 12.

    Hay, I. D., Belousoff, M. J., Dunstan, R. A., Bamert, R. S. & Lithgow, T. Structure and membrane topography of the Vibrio-type secretin complex from the type 2 secretion system of enteropathogenic Escherichia coli. J. Bacteriol. 200, e00521-17 (2018).

    Google Scholar 

  13. 13.

    Fei, J. & Sharma, C. M. RNA localization in bacteria. Microbiol. Spectr. (2018).

    Article  Google Scholar 

  14. 14.

    Konorty, M., Kahana, N., Linaroudis, A., Minsky, A. & Medalia, O. Structural analysis of photosynthetic membranes by cryo-electron tomography of intact Rhodopseudomonas viridis cells. J. Struct. Biol. 161, 393–400 (2008).

    CAS  Google Scholar 

  15. 15.

    Adams, P. G. et al. Comparison of the physical characteristics of chlorosomes from three different phyla of green phototrophic bacteria. Biochim. Biophys. Acta 1827, 1235–1244 (2013).

    CAS  Google Scholar 

  16. 16.

    Frain, K. M., Gangl, D., Jones, A., Zedler, J. A. Z. & Robinson, C. Protein translocation and thylakoid biogenesis in cyanobacteria. Biochim. Biophys. Acta 1857, 266–273 (2016).

    CAS  Google Scholar 

  17. 17.

    LaSarre, B. et al. Restricted localization of photosynthetic intracytoplasmic membranes (ICMs) in multiple genera of purple nonsulfur bacteria. mBio 9, e00780-18 (2018).

    Google Scholar 

  18. 18.

    Day, P. M. & Theg, S. M. Evolution of protein transport to the chloroplast envelope membranes. Photosynth. Res. 138, 315–326 (2018).

    CAS  Google Scholar 

  19. 19.

    Cornejo, E., Subramanian, P., Li, Z., Jensen, G. J. & Komeili, A. Dynamic remodeling of the magnetosome membrane is triggered by the initiation of biomineralization. mBio 7, e01898-15 (2016).

    Google Scholar 

  20. 20.

    Stoeger, T., Battich, N. & Pelkmans, L. Passive noise filtering by cellular compartmentalization. Cell 164, 1151–1161 (2016).

    CAS  Google Scholar 

  21. 21.

    Grommet, A. B., Feller, M. & Klajn, R. Chemical reactivity under nanoconfinement. Nat. Nanotechnol. 15, 256–271 (2020).

    CAS  Google Scholar 

  22. 22.

    Dworkin, M. & Gutnick, D. Sergei Winogradsky: a founder of modern microbiology and the first microbial ecologist. FEMS Microbiol. Rev. 36, 364–379 (2012).

    CAS  Google Scholar 

  23. 23.

    Saier, M. H. Jr. & Bogdanov, M. V. Membranous organelles in bacteria. J. Mol. Microbiol. Biotechnol. 23, 5–12 (2013).

    CAS  Google Scholar 

  24. 24.

    Grant, C. R., Wan, J. & Komeili, A. Organelle formation in bacteria and Archaea. Annu. Rev. Cell Dev. Biol. 34, 217–238 (2018).

    CAS  Google Scholar 

  25. 25.

    Santarella-Mellwig, R. et al. The compartmentalized bacteria of the planctomycetes-verrucomicrobia-chlamydiae superphylum have membrane coat-like proteins. PLoS Biol. 8, e1000281 (2010).

    Google Scholar 

  26. 26.

    Boedeker, C. et al. Determining the bacterial cell biology of Planctomycetes. Nat. Commun. 8, 14853 (2017).

    CAS  Google Scholar 

  27. 27.

    Muñoz-Gómez, S. A., Wideman, J. G., Roger, A. J. & Slamovits, C. H. The origin of mitochondrial cristae from alphaproteobacteria. Mol. Biol. Evol. 34, 943–956 (2017).

    Google Scholar 

  28. 28.

    Nickelsen, J. et al. Biogenesis of the cyanobacterial thylakoid membrane system — an update. FEMS Microbiol. Lett. 315, 1–5 (2011).

    CAS  Google Scholar 

  29. 29.

    Uebe, R. & Schüler, D. Magnetosome biogenesis in magnetotactic bacteria. Nat. Rev. Microbiol. 14, 621–637 (2016).

    CAS  Google Scholar 

  30. 30.

    Dobro, M. J. et al. Uncharacterized bacterial structures revealed by electron cryotomography. J. Bacteriol. 199, e00100-17 (2017).

    Google Scholar 

  31. 31.

    Martijn, J., Vosseberg, J., Guy, L., Offre, P. & Ettema, T. J. G. Deep mitochondrial origin outside the sampled alphaproteobacteria. Nature 557, 101–105 (2018).

    CAS  Google Scholar 

  32. 32.

    Spang, A. et al. Proposal of the reverse flow model for the origin of the eukaryotic cell based on comparative analyses of Asgard archaeal metabolism. Nat. Microbiol. 4, 1138–1148 (2019).

    CAS  Google Scholar 

  33. 33.

    Brantner, C. A., Remsen, C. C., Owen, H. A., Buchholz, L. A. & Perille Collins, M. L. Intracellular localization of the particulate methane monooxygenase and methanol dehydrogenase in Methylomicrobium album BG8. Arch. Microbiol. 178, 59–64 (2002).

    CAS  Google Scholar 

  34. 34.

    Tucker, J. D. et al. Membrane invagination in Rhodobacter sphaeroides is initiated at curved regions of the cytoplasmic membrane, then forms both budded and fully detached spherical vesicles. Mol. Microbiol. 76, 833–847 (2010).

    CAS  Google Scholar 

  35. 35.

    Noble, J. M. et al. Connectivity of centermost chromatophores in Rhodobacter sphaeroides bacteria. Mol. Microbiol. 109, 812–825 (2018).

    CAS  Google Scholar 

  36. 36.

    Van De Meene, A. M. L., Hohmann-Marriott, M. F., Vermaas, W. F. J. & Roberson, R. W. The three-dimensional structure of the cyanobacterium Synechocystis sp. PCC 6803. Arch. Microbiol. 184, 259–270 (2006).

    CAS  Google Scholar 

  37. 37.

    Nevo, R. et al. Thylakoid membrane perforations and connectivity enable intracellular traffic in cyanobacteria. EMBO J. 26, 1467–1473 (2007).

    CAS  Google Scholar 

  38. 38.

    Ting, C. S., Hsieh, C., Sundararaman, S., Mannella, C. & Marko, M. Cryo-electron tomography reveals the comparative three-dimensional architecture of Prochlorococcus, a globally important marine cyanobacterium. J. Bacteriol. 189, 4485–4493 (2007).

    CAS  Google Scholar 

  39. 39.

    Gonzalez-Esquer, C. R. et al. Cyanobacterial ultrastructure in light of genomic sequence data. Photosynth. Res. 129, 147–157 (2016).

    CAS  Google Scholar 

  40. 40.

    Strous, M. et al. Missing lithotroph identified as new planctomycete. Nature 400, 446–449 (1999).

    CAS  Google Scholar 

  41. 41.

    Lindsay, M. R. et al. Cell compartmentalisation in planctomycetes: novel types of structural organisation for the bacterial cell. Arch. Microbiol. 175, 413–429 (2001).

    CAS  Google Scholar 

  42. 42.

    Strous, M. et al. Deciphering the evolution and metabolism of an anammox bacterium from a community genome. Nature 440, 790–794 (2006).

    Google Scholar 

  43. 43.

    van Niftrik, L. et al. Linking ultrastructure and function in four genera of anaerobic ammonium-oxidizing bacteria: cell plan, glycogen storage, and localization of cytochrome c proteins. J. Bacteriol. 190, 708–717 (2008).

    Google Scholar 

  44. 44.

    van Niftrik, L. et al. Combined structural and chemical analysis of the anammoxosome: a membrane-bounded intracytoplasmic compartment in anammox bacteria. J. Struct. Biol. 161, 401–410 (2008).

    Google Scholar 

  45. 45.

    Kartal, B., van Niftrik, L., Keltjens, J. T., Op den Camp, H. J. M. & Jetten, M. S. M. Anammox — growth physiology, cell biology, and metabolism. Adv. Microb. Physiol. 60, 211–262 (2012).

    CAS  Google Scholar 

  46. 46.

    Neumann, S. et al. Isolation and characterization of a prokaryotic cell organelle from the anammox bacterium Kuenenia stuttgartiensis. Mol. Microbiol. 94, 794–802 (2014).

    CAS  Google Scholar 

  47. 47.

    de Almeida, N. M. et al. Immunogold localization of key metabolic enzymes in the anammoxosome and on the tubule-like structures of Kuenenia stuttgartiensis. J. Bacteriol. 197, 2432–2441 (2015).

    Google Scholar 

  48. 48.

    Damsté, J. S. S. et al. Linearly concatenated cyclobutane lipids form a dense bacterial membrane. Nature 419, 708–712 (2002).

    Google Scholar 

  49. 49.

    van Niftrik, L. A. et al. The anammoxosome: an intracytoplasmic compartment in anammox bacteria. FEMS Microbiol. Lett. 233, 7–13 (2004).

    Google Scholar 

  50. 50.

    Moss, F. R. et al. Ladderane phospholipids form a densely packed membrane with normal hydrazine and anomalously low proton/hydroxide permeability. Proc. Natl Acad. Sci. USA 115, 9098–9103 (2018).

    CAS  Google Scholar 

  51. 51.

    Fuerst, J. A. & Webb, R. I. Membrane-bounded nucleoid in the eubacterium Gemmatata obscuriglobus. Proc. Natl Acad. Sci. USA 88, 8184–8188 (1991).

    CAS  Google Scholar 

  52. 52.

    Lindsay, M. R., Webb, R. I. & Fuerst, J. A. Pirellulosomes: a new type of membrane-bounded cell compartment in planctomycete bacteria of the genus Pirellula. Microbiology 143, 739–748 (1997).

    CAS  Google Scholar 

  53. 53.

    Gottshall, E. Y., Seebart, C., Gatlin, J. C. & Ward, N. L. Spatially segregated transcription and translation in cells of the endomembrane-containing bacterium Gemmata obscuriglobus. Proc. Natl Acad. Sci. USA 111, 11067–11072 (2014).

    CAS  Google Scholar 

  54. 54.

    Fuerst, J. A. & Sagulenko, E. Beyond the bacterium: planctomycetes challenge our concepts of microbial structure and function. Nat. Rev. Microbiol. 9, 403–413 (2011).

    CAS  Google Scholar 

  55. 55.

    Santarella-Mellwig, R., Pruggnaller, S., Roos, N., Mattaj, I. W. & Devos, D. P. Three-dimensional reconstruction of bacteria with a complex endomembrane system. PLoS Biol. 11, e1001565 (2013).

    CAS  Google Scholar 

  56. 56.

    Okuda, Y., Denda, K. & Fukumori, Y. Cloning and sequencing of a gene encoding a new member of the tetratricopeptide protein family from magnetosomes of magnetospirillum magnetotacticum. Gene 171, 99–102 (1996).

    CAS  Google Scholar 

  57. 57.

    Gorby, Y. A., Beveridge, T. J. & Blakemore, R. P. Characterization of the bacterial magnetosome membrane. J. Bacteriol. 170, 834–841 (1988).

    CAS  Google Scholar 

  58. 58.

    Grünberg, K., Wawer, C., Tebo, B. M. & Schüler, D. A large gene cluster encoding several magnetosome proteins is conserved in different species of magnetotactic bacteria. Appl. Environ. Microbiol. 67, 4573–4582 (2001).

    Google Scholar 

  59. 59.

    Grünberg, K. et al. Biochemical and proteomic analysis of the magnetosome membrane in Magnetospirillum gryphiswaldense. Appl. Environ. Microbiol. 70, 1040–1050 (2004).

    Google Scholar 

  60. 60.

    Guo, Y., Sirkis, D. W. & Schekman, R. Protein sorting at the trans-Golgi network. Annu. Rev. Cell Dev. Biol. 30, 169–206 (2014).

    CAS  Google Scholar 

  61. 61.

    Wickner, W. & Schekman, R. Protein translocation across biological membranes. Science 310, 1452–1456 (2005).

    CAS  Google Scholar 

  62. 62.

    Komeili, A., Vali, H., Beveridge, T. J. & Newman, D. K. Magnetosome vesicles are present before magnetite formation, and MamA is required for their activation. Proc. Natl Acad. Sci. USA 101, 3839–3844 (2004).

    CAS  Google Scholar 

  63. 63.

    Byrne, M. E. et al. Desulfovibrio magneticus RS-1 contains an iron-and phosphorus-rich organelle distinct from its bullet-shaped magnetosomes. Proc. Natl Acad. Sci. USA 107, 12263–12268 (2010).

    CAS  Google Scholar 

  64. 64.

    Glasauer, S., Langley, S. & Beveridge, T. J. Intracellular iron minerals in a dissimilatory iron-reducing bacterium. Science 295, 117–119 (2002).

    CAS  Google Scholar 

  65. 65.

    Glasauer, S. et al. Mixed-valence cytoplasmic iron granules are linked to anaerobic respiration. Appl. Environ. Microbiol. 73, 993–996 (2007).

    CAS  Google Scholar 

  66. 66.

    Rahn-Lee, L. et al. A genetic strategy for probing the functional diversity of magnetosome formation. PLoS Genet. 11, e1004811 (2015).

    Google Scholar 

  67. 67.

    Grant, C. R. & Komeili, A. Ferrosomes are iron storage organelles formed by broadly conserved gene clusters in bacteria and archaea. Preprint at bioRxiv (2020).

    Article  Google Scholar 

  68. 68.

    Salman, V., Bailey, J. V. & Teske, A. Phylogenetic and morphologic complexity of giant sulphur bacteria. Antonie Van Leeuwenhoek 104, 169–186 (2013).

    CAS  Google Scholar 

  69. 69.

    Docampo, R., de Souza, W., Miranda, K., Rohloff, P. & Moreno, S. N. J. Acidocalcisomes? Conserved from bacteria to man. Nat. Rev. Microbiol. 3, 251–261 (2005).

    CAS  Google Scholar 

  70. 70.

    Seufferheld, M. et al. Identification of organelles in bacteria similar to acidocalcisomes of unicellular eukaryotes. J. Biol. Chem. 278, 29971–29978 (2003).

    CAS  Google Scholar 

  71. 71.

    Dolezal, P., Likic, V., Tachezy, J. & Lithgow, T. Evolution of the molecular machines for protein import into mitochondria. Science 313, 314–318 (2006).

    CAS  Google Scholar 

  72. 72.

    Michels, P. A. M. et al. Peroxisomes, glyoxysomes and glycosomes. Mol. Membr. Biol. 22, 133–145 (2005).

    CAS  Google Scholar 

  73. 73.

    Liu, X., Ma, C. & Subramani, S. Recent advances in peroxisomal matrix protein import. Curr. Opin. Cell Biol. 24, 484–489 (2012).

    CAS  Google Scholar 

  74. 74.

    Low, H. H. & Löwe, J. A bacterial dynamin-like protein. Nature 444, 766–769 (2006).

    CAS  Google Scholar 

  75. 75.

    Heidrich, J., Thurotte, A. & Schneider, D. Specific interaction of IM30/Vipp1 with cyanobacterial and chloroplast membranes results in membrane remodeling and eventually in membrane fusion. Biochim. Biophys. Acta 1859, 537–549 (2017).

    CAS  Google Scholar 

  76. 76.

    Van Niftrik, L. et al. Cell division ring, a new cell division protein and vertical inheritance of a bacterial organelle in anammox planctomycetes. Mol. Microbiol. 73, 1009–1019 (2009).

    Google Scholar 

  77. 77.

    McMahon, H. T. & Gallop, J. L. Membrane curvature and mechanisms of dynamic cell membrane remodelling. Nature 438, 590–596 (2005).

    CAS  Google Scholar 

  78. 78.

    McMahon, H. T. & Boucrot, E. Membrane curvature at a glance. J. Cell Sci. 128, 1065–1070 (2015).

    CAS  Google Scholar 

  79. 79.

    Tavano, C. L. & Donohue, T. J. Development of the bacterial photosynthetic apparatus. Curr. Opin. Microbiol. 9, 625–631 (2006).

    CAS  Google Scholar 

  80. 80.

    Chandler, D. E., Hsin, J., Harrison, C. B., Gumbart, J. & Schulten, K. Intrinsic curvature properties of photosynthetic proteins in chromatophores. Biophys. J. 95, 2822–2836 (2008).

    CAS  Google Scholar 

  81. 81.

    Qian, P., Bullough, P. A. & Hunter, C. N. Three-dimensional reconstruction of a membrane-bending complex the RC-LH1-PufX core dimer of Rhodobacter sphaeroides. J. Biol. Chem. 283, 14002–14011 (2008).

    CAS  Google Scholar 

  82. 82.

    Zeng, X. et al. Proteomic characterization of the Rhodobacter sphaeroides 2.4.1 photosynthetic membrane: identification of new proteins. J. Bacteriol. 189, 7464–7474 (2007).

    CAS  Google Scholar 

  83. 83.

    Muñoz-Gómez, S. A. et al. Ancient homology of the mitochondrial contact site and cristae organizing system points to an endosymbiotic origin of mitochondrial cristae. Curr. Biol. 25, 1489–1495 (2015).

    Google Scholar 

  84. 84.

    Muñoz-Gómez, S. A., Slamovits, C. H., Dacks, J. B. & Wideman, J. G. The evolution of MICOS: ancestral and derived functions and interactions. Commun. Integr. Biol. 8, e1094593 (2015).

    Google Scholar 

  85. 85.

    Arechaga, I. et al. Characterisation of new intracellular membranes in Escherichia coli accompanying large scale over-production of the b subunit of F1Fo ATP synthase. FEBS Lett. 482, 215–219 (2000).

    CAS  Google Scholar 

  86. 86.

    Blum, T. B., Hahn, A., Meier, T., Davies, K. M. & Kühlbrandt, W. Dimers of mitochondrial ATP synthase induce membrane curvature and self-assemble into rows. Proc. Natl Acad. Sci. USA 116, 4250–4255 (2019).

    CAS  Google Scholar 

  87. 87.

    Walser, P. J. et al. Constitutive formation of caveolae in a bacterium. Cell 150, 752–763 (2012).

    CAS  Google Scholar 

  88. 88.

    Arechaga, I. Membrane invaginations in bacteria and mitochondria: common features and evolutionary scenarios. J. Mol. Microbiol. Biotechnol. 23, 13–23 (2013).

    CAS  Google Scholar 

  89. 89.

    Zak, E. et al. The initial steps of biogenesis of cyanobacterial photosystems occur in plasma membranes. Proc. Natl Acad. Sci. USA 98, 13443–13448 (2001).

    CAS  Google Scholar 

  90. 90.

    Medema, M. H. et al. A predicted physicochemically distinct sub-proteome associated with the intracellular organelle of the anammox bacterium Kuenenia stuttgartiensis. BMC Genomics 11, 299 (2010).

    Google Scholar 

  91. 91.

    Raschdorf, O. et al. Genetic and ultrastructural analysis reveals the key players and initial steps of bacterial magnetosome membrane biogenesis. PLoS Genet. 12, e1006101 (2016).

    Google Scholar 

  92. 92.

    Alcock, F., Clements, A., Webb, C. & Lithgow, T. Tinkering inside the organelle. Science 327, 649–650 (2010).

    CAS  Google Scholar 

  93. 93.

    Aldridge, C., Spence, E., Kirkilionis, M. A., Frigerio, L. & Robinson, C. Tat-dependent targeting of Rieske iron-sulphur proteins to both the plasma and thylakoid membranes in the cyanobacterium Synechocystis PCC6803. Mol. Microbiol. 70, 140–150 (2008).

    CAS  Google Scholar 

  94. 94.

    Rajalahti, T. et al. Proteins in different Synechocystis compartments have distinguishing N-terminal features: a combined proteomics and multivariate sequence analysis. J. Proteome Res. 6, 2420–2434 (2007).

    CAS  Google Scholar 

  95. 95.

    Nevo-Dinur, K., Nussbaum-Shochat, A., Ben-Yehuda, S. & Amster-Choder, O. Translation-independent localization of mRNA in E. coli. Science 331, 1081–1084 (2011).

    CAS  Google Scholar 

  96. 96.

    Binenbaum, Z., Parola, A. H., Zaritsky, A. & Fishov, I. Transcription-and translation-dependent changes in membrane dynamics in bacteria: testing the transertion model for domain formation. Mol. Microbiol. 32, 1173–1182 (1999).

    CAS  Google Scholar 

  97. 97.

    Bakshi, S., Choi, H., Mondal, J. & Weisshaar, J. C. Time-dependent effects of transcription-and translation-halting drugs on the spatial distributions of the Escherichia coli chromosome and ribosomes. Mol. Microbiol. 94, 871–887 (2014).

    CAS  Google Scholar 

  98. 98.

    Matsumoto, K., Hara, H., Fishov, I., Mileykovskaya, E. & Norris, V. The membrane: transertion as an organizing principle in membrane heterogeneity. Front. Microbiol. 6, 572 (2015).

    Google Scholar 

  99. 99.

    Roggiani, M. & Goulian, M. Chromosome-membrane interactions in bacteria. Annu. Rev. Genet. 49, 115–129 (2015).

    CAS  Google Scholar 

  100. 100.

    Schübbe, S. et al. Characterization of a spontaneous nonmagnetic mutant of Magnetospirillum gryphiswaldense reveals a large deletion comprising a putative magnetosome island. J. Bacteriol. 185, 5779–5790 (2003).

    Google Scholar 

  101. 101.

    Ullrich, S., Kube, M., Schübbe, S., Reinhardt, R. & Schüler, D. A hypervariable 130-kilobase genomic region of Magnetospirillum gryphiswaldense comprises a magnetosome island which undergoes frequent rearrangements during stationary growth. J. Bacteriol. 187, 7176–7184 (2005).

    CAS  Google Scholar 

  102. 102.

    Murat, D., Quinlan, A., Vali, H. & Komeili, A. Comprehensive genetic dissection of the magnetosome gene island reveals the step-wise assembly of a prokaryotic organelle. Proc. Natl Acad. Sci. USA 107, 5593–5598 (2010).

    CAS  Google Scholar 

  103. 103.

    Lohße, A. et al. Functional analysis of the magnetosome island in Magnetospirillum gryphiswaldense: the mamAB operon is sufficient for magnetite biomineralization. PLoS ONE 6, e25561 (2011).

    Google Scholar 

  104. 104.

    Murat, D. et al. The magnetosome membrane protein, MmsF, is a major regulator of magnetite biomineralization in Magnetospirillum magneticum AMB-1. Mol. Microbiol. 85, 684–699 (2012).

    CAS  Google Scholar 

  105. 105.

    Lohße, A. et al. Genetic dissection of the mamAB and mms6 operons reveals a gene set essential for magnetosome biogenesis in Magnetospirillum gryphiswaldense. J. Bacteriol. 196, 2658–2669 (2014).

    Google Scholar 

  106. 106.

    Yamamoto, D. et al. Visualization and structural analysis of the bacterial magnetic organelle magnetosome using atomic force microscopy. Proc. Natl Acad. Sci. USA 107, 9382–9387 (2010).

    CAS  Google Scholar 

  107. 107.

    Cornejo, E., Abreu, N. & Komeili, A. Compartmentalization and organelle formation in bacteria. Curr. Opin. Cell Biol. 26, 132–138 (2014).

    CAS  Google Scholar 

  108. 108.

    Zeytuni, N. et al. Self-recognition mechanism of MamA, a magnetosome-associated TPR-containing protein, promotes complex assembly. Proc. Natl Acad. Sci. USA 108, E480–E487 (2011).

    CAS  Google Scholar 

  109. 109.

    Uebe, R. et al. The cation diffusion facilitator proteins MamB and MamM of Magnetospirillum gryphiswaldense have distinct and complex functions, and are involved in magnetite biomineralization and magnetosome membrane assembly. Mol. Microbiol. 82, 818–835 (2011).

    CAS  Google Scholar 

  110. 110.

    Scheffel, A., Gärdes, A., Grünberg, K., Wanner, G. & Schüler, D. The major magnetosome proteins MamGFDC are not essential for magnetite biomineralization in Magnetospirillum gryphiswaldense but regulate the size of magnetosome crystals. J. Bacteriol. 190, 377–386 (2008).

    CAS  Google Scholar 

  111. 111.

    Raschdorf, O., Müller, F. D., Pósfai, M., Plitzko, J. M. & Schüler, D. The magnetosome proteins MamX, MamZ and MamH are involved in redox control of magnetite biomineralization in Magnetospirillum gryphiswaldense. Mol. Microbiol. 89, 872–886 (2013).

    CAS  Google Scholar 

  112. 112.

    Komeili, A., Li, Z., Newman, D. K. & Jensen, G. J. Magnetosomes are cell membrane invaginations organized by the actin-like protein MamK. Science 311, 242–245 (2006).

    CAS  Google Scholar 

  113. 113.

    Scheffel, A. et al. An acidic protein aligns magnetosomes along a filamentous structure in magnetotactic bacteria. Nature 440, 110–114 (2006).

    CAS  Google Scholar 

  114. 114.

    Taoka, A. et al. Tethered magnets are the key to magnetotaxis: direct observations of Magnetospirillum magneticum AMB-1 show that MamK distributes magnetosome organelles equally to daughter cells. mBio 8, e00679-17 (2017).

    Google Scholar 

  115. 115.

    Toro-Nahuelpan, M. et al. Segregation of prokaryotic magnetosomes organelles is driven by treadmilling of a dynamic actin-like MamK filament. BMC Biol. 14, 88 (2016).

    Google Scholar 

  116. 116.

    Toro-Nahuelpan, M. et al. MamY is a membrane-bound protein that aligns magnetosomes and the motility axis of helical magnetotactic bacteria. Nat. Microbiol. 4, 1978–1989 (2019).

    CAS  Google Scholar 

  117. 117.

    Nott, T. J., Craggs, T. D. & Baldwin, A. J. Membraneless organelles can melt nucleic acid duplexes and act as biomolecular filters. Nat. Chem. 8, 569 (2016).

    CAS  Google Scholar 

  118. 118.

    Banani, S. F., Lee, H. O., Hyman, A. A. & Rosen, M. K. Biomolecular condensates: organizers of cellular biochemistry. Nat. Rev. Mol. Cell Biol. 18, 285–298 (2017).

    CAS  Google Scholar 

  119. 119.

    Abbondanzieri, E. A. & Meyer, A. S. More than just a phase: the search for membraneless organelles in the bacterial cytoplasm. Curr. Genet. 65, 691–694 (2019).

    CAS  Google Scholar 

  120. 120.

    Ladouceur, A.-M. et al. Clusters of bacterial RNA polymerase are biomolecular condensates that assemble through liquid-liquid phase separation. Preprint at bioRxiv (2020).

    Article  Google Scholar 

  121. 121.

    Hondele, M. et al. DEAD-box ATPases are global regulators of phase-separated organelles. Nature 573, 144–148 (2019).

    CAS  Google Scholar 

  122. 122.

    Al-Husini, N., Tomares, D. T., Bitar, O., Childers, W. S. & Schrader, J. M. α-Proteobacterial RNA degradosomes assemble liquid-liquid phase-separated RNP bodies. Mol. Cell 71, 1027–1039 (2018).

    CAS  Google Scholar 

  123. 123.

    Boisvert, F.-M., van Koningsbruggen, S., Navascués, J. & Lamond, A. I. The multifunctional nucleolus. Nat. Rev. Mol. Cell Biol. 8, 574–585 (2007).

    CAS  Google Scholar 

  124. 124.

    Jin, D. J., Mata Martin, C., Sun, Z., Cagliero, C. & Zhou, Y. N. Nucleolus-like compartmentalization of the transcription machinery in fast-growing bacterial cells. Crit. Rev. Biochem. Mol. Biol. 52, 96–106 (2017).

    CAS  Google Scholar 

  125. 125.

    Uversky, V. N. Intrinsically disordered proteins in overcrowded milieu: Membrane-less organelles, phase separation, and intrinsic disorder. Curr. Opin. Struct. Biol. 44, 18–30 (2017).

    CAS  Google Scholar 

  126. 126.

    Mendoza, S. D. et al. A bacteriophage nucleus-like compartment shields DNA from CRISPR nucleases. Nature 577, 244–248 (2020).

    CAS  Google Scholar 

  127. 127.

    Malone, L. M. et al. A jumbo phage that forms a nucleus-like structure evades CRISPR–Cas DNA targeting but is vulnerable to type III RNA-based immunity. Nat. Microbiol. 5, 48–55 (2020).

    CAS  Google Scholar 

  128. 128.

    Chaikeeratisak, V. et al. Viral capsid trafficking along treadmilling tubulin filaments in bacteria. Cell 177, 1771–1780 (2019).

    CAS  Google Scholar 

  129. 129.

    Kerfeld, C. A., Aussignargues, C., Zarzycki, J., Cai, F. & Sutter, M. Bacterial microcompartments. Nat. Rev. Microbiol. 16, 277–290 (2018).

    CAS  Google Scholar 

  130. 130.

    Chowdhury, C., Sinha, S., Chun, S., Yeates, T. O. & Bobik, T. A. Diverse bacterial microcompartment organelles. Microbiol. Mol. Biol. Rev. 78, 438–468 (2014).

    CAS  Google Scholar 

  131. 131.

    Kerfeld, C. A. et al. Protein structures forming the shell of primitive bacterial organelles. Science 309, 936–938 (2005).

    CAS  Google Scholar 

  132. 132.

    Tanaka, S. et al. Atomic-level models of the bacterial carboxysome shell. Science 319, 1083–1086 (2008).

    CAS  Google Scholar 

  133. 133.

    Sutter, M., Greber, B., Aussignargues, C. & Kerfeld, C. A. Assembly principles and structure of a 6.5-MDa bacterial microcompartment shell. Science 356, 1293–1297 (2017).

    CAS  Google Scholar 

  134. 134.

    Kalnins, G. et al. Encapsulation mechanisms and structural studies of GRM2 bacterial microcompartment particles. Nat. Commun. 11, 388 (2020).

    CAS  Google Scholar 

  135. 135.

    Axen, S. D., Erbilgin, O. & Kerfeld, C. A. A taxonomy of bacterial microcompartment loci constructed by a novel scoring method. PLoS Comput. Biol. 10, e1003898 (2014).

    Google Scholar 

  136. 136.

    Chowdhury, C. et al. Selective molecular transport through the protein shell of a bacterial microcompartment organelle. Proc. Natl Acad. Sci. USA 112, 2990–2995 (2015).

    CAS  Google Scholar 

  137. 137.

    Yang, M. et al. Decoding the stoichiometric composition and organisation of bacterial metabolosomes. Nat. Commun. 11, 1976 (2020).

    CAS  Google Scholar 

  138. 138.

    Niederhuber, M. J., Lambert, T. J., Yapp, C., Silver, P. A. & Polka, J. K. Superresolution microscopy of the β-carboxysome reveals a homogeneous matrix. Mol. Biol. Cell 28, 2734–2745 (2017).

    CAS  Google Scholar 

  139. 139.

    Cameron, J. C., Wilson, S. C., Bernstein, S. L. & Kerfeld, C. A. Biogenesis of a bacterial organelle: the carboxysome assembly pathway. Cell 155, 1131–1140 (2013).

    CAS  Google Scholar 

  140. 140.

    Wang, H. et al. Rubisco condensate formation by CcmM in β-carboxysome biogenesis. Nature 566, 131–135 (2019).

    CAS  Google Scholar 

  141. 141.

    Ryan, P. et al. The small RbcS-like domains of the β-carboxysome structural protein CcmM bind RubisCO at a site distinct from that binding the RbcS subunit. J. Biol. Chem. 294, 2593–2603 (2019).

    CAS  Google Scholar 

  142. 142.

    Kerfeld, C. A. & Melnicki, M. R. Assembly, function and evolution of cyanobacterial carboxysomes. Curr. Opin. Plant. Biol. 31, 66–75 (2016).

    CAS  Google Scholar 

  143. 143.

    Oltrogge, L. M. et al. Multivalent interactions between CsoS2 and Rubisco mediate α-carboxysome formation. Nat. Struct. Mol. Biol. 27, 281–287 (2020).

    CAS  Google Scholar 

  144. 144.

    Savage, D. F., Afonso, B., Chen, A. H. & Silver, P. A. Spatially ordered dynamics of the bacterial carbon fixation machinery. Science 327, 1258–1261 (2010).

    CAS  Google Scholar 

  145. 145.

    MacCready, J. S. et al. Protein gradients on the nucleoid position the carbon-fixing organelles of cyanobacteria. eLife 7, e39723 (2018).

    Google Scholar 

  146. 146.

    MacCready, J. S., Basalla, J. L. & Vecchiarelli, A. G. Origin and evolution of carboxysome positioning systems in Cyanobacteria. Mol. Biol. Evol. 37, 1434–1451 (2020).

    Google Scholar 

  147. 147.

    Sutter, M. et al. Structural basis of enzyme encapsulation into a bacterial nanocompartment. Nat. Struct. Mol. Biol. 15, 939–947 (2008).

    CAS  Google Scholar 

  148. 148.

    McHugh, C. A. et al. A virus capsid-like nanocompartment that stores iron and protects bacteria from oxidative stress. EMBO J. 33, 1896–1911 (2014).

    CAS  Google Scholar 

  149. 149.

    Giessen, T. W. et al. Large protein organelles form a new iron sequestration system with high storage capacity. eLife 8, e46070 (2019).

    Google Scholar 

  150. 150.

    Nichols, R. J. et al. Discovery and characterization of a novel family of prokaryotic nanocompartments involved in sulfur metabolism. Preprint at bioRxiv (2020).

    Article  Google Scholar 

  151. 151.

    Giessen, T. W. & Silver, P. A. Widespread distribution of encapsulin nanocompartments reveals functional diversity. Nat. Microbiol. 2, 17029 (2017).

    Google Scholar 

  152. 152.

    He, D. et al. Conservation of the structural and functional architecture of encapsulated ferritins in bacteria and archaea. Biochem. J. 476, 975–989 (2019).

    CAS  Google Scholar 

  153. 153.

    Contreras, H. et al. Characterization of a Mycobacterium tuberculosis nanocompartment and its potential cargo proteins. J. Biol. Chem. 289, 18279–18289 (2014).

    CAS  Google Scholar 

  154. 154.

    Xing, C.-Y. et al. A self-assembled nanocompartment in anammox bacteria for resisting intracelluar hydroxylamine stress. Sci. Total Environ. 717, 137030 (2020).

    CAS  Google Scholar 

  155. 155.

    Rahmanpour, R. & Bugg, T. D. H. Assembly in vitro of Rhodococcus jostii RHA 1 encapsulin and peroxidase DypB to form a nanocompartment. FEBS J. 280, 2097–2104 (2013).

    CAS  Google Scholar 

  156. 156.

    Rurup, W. F., Snijder, J., Koay, M. S. T., Heck, A. J. R. & Cornelissen, J. J. L. M. Self-sorting of foreign proteins in a bacterial nanocompartment. J. Am. Chem. Soc. 136, 3828–3832 (2014).

    CAS  Google Scholar 

  157. 157.

    Lončar, N., Rozeboom, H. J., Franken, L. E., Stuart, M. C. A. & Fraaije, M. W. Molecular packaging of biocatalysts using a robust protein cage. Preprint at ChemRxiv (2020).

    Article  Google Scholar 

  158. 158.

    Pfeifer, F. Distribution, formation and regulation of gas vesicles. Nat. Rev. Microbiol. 10, 705–715 (2012).

    CAS  Google Scholar 

  159. 159.

    Uchino, K., Saito, T., Gebauer, B. & Jendrossek, D. Isolated poly (3-hydroxybutyrate) (PHB) granules are complex bacterial organelles catalyzing formation of PHB from acetyl coenzyme A (CoA) and degradation of PHB to acetyl-CoA. J. Bacteriol. 189, 8250–8256 (2007).

    CAS  Google Scholar 

  160. 160.

    Jendrossek, D. Polyhydroxyalkanoate granules are complex subcellular organelles (carbonosomes). J. Bacteriol. 191, 3195–3202 (2009).

    CAS  Google Scholar 

  161. 161.

    Jendrossek, D. & Pfeiffer, D. New insights in the formation of polyhydroxyalkanoate granules (carbonosomes) and novel functions of poly (3-hydroxybutyrate). Environ. Microbiol. 16, 2357–2373 (2014).

    CAS  Google Scholar 

  162. 162.

    Griebel, R., Smith, Z. & Merrick, J. M. Metabolism of poly (β-hydroxybutyrate). I. Purification, composition, and properties of native poly (β-hydroxybutyrate) granules from Bacillus megaterium. Biochemistry 7, 3676–3681 (1968).

    CAS  Google Scholar 

  163. 163.

    Bresan, S. et al. Polyhydroxyalkanoate (PHA) granules have no phospholipids. Sci. Rep. 6, 26612 (2016).

    CAS  Google Scholar 

  164. 164.

    Cohen-Bazire, G., Pfennig, N. & Kunisawa, R. The fine structure of green bacteria. J. Cell Biol. 22, 207–225 (1964).

    CAS  Google Scholar 

  165. 165.

    Bryant, D. A. et al. Candidatus Chloracidobacterium thermophilum: an aerobic phototrophic acidobacterium. Science 317, 523–526 (2007).

    CAS  Google Scholar 

  166. 166.

    Oostergetel, G. T., van Amerongen, H. & Boekema, E. J. The chlorosome: a prototype for efficient light harvesting in photosynthesis. Photosynth. Res. 104, 245–255 (2010).

    CAS  Google Scholar 

  167. 167.

    Günther, L. M. et al. Structure of light-harvesting aggregates in individual chlorosomes. J. Phys. Chem. B 120, 5367–5376 (2016).

    Google Scholar 

  168. 168.

    Manske, A. K., Glaeser, J., Kuypers, M. M. M. & Overmann, J. Physiology and phylogeny of green sulfur bacteria forming a monospecific phototrophic assemblage at a depth of 100 meters in the Black Sea. Appl. Environ. Microbiol. 71, 8049–8060 (2005).

    CAS  Google Scholar 

  169. 169.

    Beatty, J. T. et al. An obligately photosynthetic bacterial anaerobe from a deep-sea hydrothermal vent. Proc. Natl Acad. Sci. USA 102, 9306–9310 (2005).

    CAS  Google Scholar 

  170. 170.

    Frigaard, N.-U. & Bryant, D. A. In Complex Intracellular Structures in Prokaryotes Microbiology Monographs vol. 2, (ed Shively, J.M.) 79–114 (Springer, 2006).

  171. 171.

    Martinez-Planells, A. et al. Determination of the topography and biometry of chlorosomes by atomic force microscopy. Photosynth. Res. 71, 83–90 (2002).

    CAS  Google Scholar 

  172. 172.

    Pšenčík, J. et al. Structure of chlorosomes from the green filamentous bacterium Chloroflexus aurantiacus. J. Bacteriol. 191, 6701–6708 (2009).

    Google Scholar 

  173. 173.

    Bína, D., Gardian, Z., Vácha, F. & Litvín, R. Native FMO-reaction center supercomplex in green sulfur bacteria: an electron microscopy study. Photosynth. Res. 128, 93–102 (2016).

    Google Scholar 

  174. 174.

    Nielsen, J. T. et al. In situ high-resolution structure of the baseplate antenna complex in Chlorobaculum tepidum. Nat. Commun. 7, 12454 (2016).

    CAS  Google Scholar 

  175. 175.

    Staehelin, L. A., Golecki, J. R. & Drews, G. Supramolecular organization of chlorosomes (Chlorobium vesicles) and of their membrane attachment sites in Chlorobium limicola. Biochim. Biophys. Acta 589, 30–45 (1980).

    CAS  Google Scholar 

  176. 176.

    Holo, H., Broch-Due, M. & Ormerod, J. G. Glycolipids and the structure of chlorosomes in green bacteria. Arch. Microbiol. 143, 94–99 (1985).

    CAS  Google Scholar 

  177. 177.

    Sørensen, P. G., Cox, R. P. & Miller, M. Chlorosome lipids from Chlorobium tepidum: characterization and quantification of polar lipids and wax esters. Photosynth. Res. 95, 191–196 (2008).

    Google Scholar 

  178. 178.

    Kudryashev, M., Aktoudianaki, A., Dedoglou, D., Stahlberg, H. & Tsiotis, G. The ultrastructure of Chlorobaculum tepidum revealed by cryo-electron tomography. Biochim. Biophys. Acta 1837, 1635–1642 (2014).

    CAS  Google Scholar 

  179. 179.

    Fetisova, Z. G., Freiberg, A. M. & Timpmann, K. E. Long-range molecular order as an efficient strategy for light harvesting in photosynthesis. Nature 334, 633–634 (1988).

    CAS  Google Scholar 

  180. 180.

    Saga, Y., Shibata, Y., Itoh, S. & Tamiaki, H. Direct counting of submicrometer-sized photosynthetic apparatus dispersed in medium at cryogenic temperature by confocal laser fluorescence microscopy: estimation of the number of bacteriochlorophyll c in single light-harvesting antenna complexes chlorosomes of green photosynthetic bacteria. J. Phys. Chem. B 111, 12605–12609 (2007).

    CAS  Google Scholar 

  181. 181.

    Ganapathy, S. et al. Alternating syn-anti bacteriochlorophylls form concentric helical nanotubes in chlorosomes. Proc. Natl Acad. Sci. USA 106, 8525–8530 (2009).

    CAS  Google Scholar 

  182. 182.

    Borrego, C. M., Gerola, P. D., Miller, M. & Cox, R. P. Light intensity effects on pigment composition and organisation in the green sulfur bacterium Chlorobium tepidum. Photosynth. Res. 59, 159–166 (1999).

    CAS  Google Scholar 

  183. 183.

    Tang, J. K.-H. et al. Temperature and carbon assimilation regulate the chlorosome biogenesis in green sulfur bacteria. Biophys. J. 105, 1346–1356 (2013).

    CAS  Google Scholar 

  184. 184.

    Smith, K. M., Kehres, L. A. & Fajer, J. Aggregation of the bacteriochlorophylls c, d, and e. Models for the antenna chlorophylls of green and brown photosynthetic bacteria. J. Am. Chem. Soc. 105, 1387–1389 (1983).

    CAS  Google Scholar 

  185. 185.

    Jochum, T. et al. The supramolecular organization of self-assembling chlorosomal bacteriochlorophyll c, d, or e mimics. Proc. Natl Acad. Sci. USA 105, 12736–12741 (2008).

    CAS  Google Scholar 

  186. 186.

    Hohmann-Marriott, M. F. & Blankenship, R. E. Hypothesis on chlorosome biogenesis in green photosynthetic bacteria. FEBS Lett. 581, 800–803 (2007).

    CAS  Google Scholar 

  187. 187.

    Frigaard, N.-U., Li, H., Milks, K. J. & Bryant, D. A. Nine mutants of Chlorobium tepidum each unable to synthesize a different chlorosome protein still assemble functional chlorosomes. J. Bacteriol. 186, 646–653 (2004).

    CAS  Google Scholar 

  188. 188.

    Wullink, W., Knudsen, J., Olson, J. M., Redlinger, T. E. & Van Bruggen, E. F. J. Localization of polypeptides in isolated chlorosomes from green phototrophic bacteria by immuno-gold labeling electron microscopy. Biochim. Biophys. Acta 1060, 97–105 (1991).

    CAS  Google Scholar 

  189. 189.

    Chung, S. & Bryant, D. A. Characterization of the csmD and csmE genes from Chlorobium tepidum. The CsmA, CsmC, CsmD, and CsmE proteins are components of the chlorosome envelope. Photosynth. Res. 50, 41–59 (1996).

    CAS  Google Scholar 

  190. 190.

    Vassilieva, E. V. et al. Subcellular localization of chlorosome proteins in Chlorobium tepidum and characterization of three new chlorosome proteins: CsmF, CsmH, and CsmX. Biochemistry 41, 4358–4370 (2002).

    CAS  Google Scholar 

  191. 191.

    Li, H., Frigaard, N.-U. & Bryant, D. A. Molecular contacts for chlorosome envelope proteins revealed by cross-linking studies with chlorosomes from Chlorobium tepidum. Biochemistry 45, 9095–9103 (2006).

    CAS  Google Scholar 

  192. 192.

    Costas, A. M. G. et al. Ultrastructural analysis and identification of envelope proteins of “Candidatus Chloracidobacterium thermophilum” chlorosomes. J. Bacteriol. 193, 6701–6711 (2011).

    CAS  Google Scholar 

  193. 193.

    Li, H. & Bryant, D. A. Envelope proteins of the CsmB/CsmF and CsmC/CsmD motif families influence the size, shape, and composition of chlorosomes in Chlorobaculum tepidum. J. Bacteriol. 191, 7109–7120 (2009).

    CAS  Google Scholar 

  194. 194.

    Li, H., Frigaard, N.-U. & Bryant, D. A. [2Fe-2S] proteins in chlorosomes: CsmI and CsmJ participate in light-dependent control of energy transfer in chlorosomes of Chlorobaculum tepidum. Biochemistry 52, 1321–1330 (2013).

    CAS  Google Scholar 

  195. 195.

    Murphy, D. J. The biogenesis and functions of lipid bodies in animals, plants and microorganisms. Prog. Lipid Res. 40, 325–438 (2001).

    CAS  Google Scholar 

  196. 196.

    Alvarez, H. & Steinbüchel, A. Triacylglycerols in prokaryotic microorganisms. Appl. Microbiol. Biotechnol. 60, 367–376 (2002).

    CAS  Google Scholar 

  197. 197.

    Low, K. L. et al. Lipid droplet-associated proteins are involved in the biosynthesis and hydrolysis of triacylglycerol in Mycobacterium bovis bacillus Calmette-Guerin. J. Biol. Chem. 285, 21662–21670 (2010).

    CAS  Google Scholar 

  198. 198.

    Chen, Y. et al. Integrated omics study delineates the dynamics of lipid droplets in Rhodococcus opacus PD630. Nucleic Acids Res. 42, 1052–1064 (2013).

    Google Scholar 

  199. 199.

    Packter, N. M. & Olukoshi, E. R. Ultrastructural studies of neutral lipid localisation in Streptomyces. Arch. Microbiol. 164, 420–427 (1995).

    CAS  Google Scholar 

  200. 200.

    Alvarez, H. M., Mayer, F., Fabritius, D. & Steinbüchel, A. Formation of intracytoplasmic lipid inclusions by Rhodococcus opacus strain PD630. Arch. Microbiol. 165, 377–386 (1996).

    CAS  Google Scholar 

  201. 201.

    Daniel, J., Maamar, H., Deb, C., Sirakova, T. D. & Kolattukudy, P. E. Mycobacterium tuberculosis uses host triacylglycerol to accumulate lipid droplets and acquires a dormancy-like phenotype in lipid-loaded macrophages. PLoS Pathog. 7, e1002093 (2011).

    CAS  Google Scholar 

  202. 202.

    Zhang, C. et al. Bacterial lipid droplets bind to DNA via an intermediary protein that enhances survival under stress. Nat. Commun. 8, 15979 (2017).

    CAS  Google Scholar 

  203. 203.

    Leung, P. M. et al. Energetic basis of microbial growth and persistence in desert ecosystems. mSystems 5, e00495-19 (2020).

    Google Scholar 

  204. 204.

    Daniel, J. et al. Induction of a novel class of diacylglycerol acyltransferases and triacylglycerol accumulation in Mycobacterium tuberculosis as it goes into a dormancy-like state in culture. J. Bacteriol. 186, 5017–5030 (2004).

    CAS  Google Scholar 

  205. 205.

    Fujimoto, T., Ohsaki, Y., Cheng, J., Suzuki, M. & Shinohara, Y. Lipid droplets: a classic organelle with new outfits. Histochem. Cell Biol. 130, 263–279 (2008).

    CAS  Google Scholar 

  206. 206.

    Hänisch, J., Wältermann, M., Robenek, H. & Steinbüchel, A. Eukaryotic lipid body proteins in oleogenous actinomycetes and their targeting to intracellular triacylglycerol inclusions: impact on models of lipid body biogenesis. Appl. Environ. Microbiol. 72, 6743–6750 (2006).

    Google Scholar 

  207. 207.

    Wältermann, M. et al. Mechanism of lipid-body formation in prokaryotes: how bacteria fatten up. Mol. Microbiol. 55, 750–763 (2005).

    Google Scholar 

  208. 208.

    Wältermann, M. & Steinbüchel, A. Neutral lipid bodies in prokaryotes: recent insights into structure, formation, and relationship to eukaryotic lipid depots. J. Bacteriol. 187, 3607–3619 (2005).

    Google Scholar 

  209. 209.

    Fossing, H. et al. Concentration and transport of nitrate by the mat-forming sulphur bacterium Thioploca. Nature 374, 713–715 (1995).

    CAS  Google Scholar 

  210. 210.

    Walter, T. & Erdmann, R. Current advances in protein import into peroxisomes. Protein J. 38, 351–362 (2019).

    CAS  Google Scholar 

  211. 211.

    Hjort, K., Goldberg, A. V., Tsaousis, A. D., Hirt, R. P. & Embley, T. M. Diversity and reductive evolution of mitochondria among microbial eukaryotes. Philos. Trans. R. Soc. B Biol. Sci. 365, 713–727 (2010).

    CAS  Google Scholar 

  212. 212.

    Strahl, H. & Errington, J. Bacterial membranes: structure, domains, and function. Annu. Rev. Microbiol. 71, 519–538 (2017).

    CAS  Google Scholar 

  213. 213.

    Gunasinghe, S. D. et al. The WD40 protein BamB mediates coupling of BAM complexes into assembly precincts in the bacterial outer membrane. Cell Rep. 23, 2782–2794 (2018).

    CAS  Google Scholar 

  214. 214.

    Buskila, A. A., Kannaiah, S. & Amster-Choder, O. RNA localization in bacteria. RNA Biol. 11, 1051–1060 (2014).

    Google Scholar 

  215. 215.

    Lau, Y. H., Giessen, T. W., Altenburg, W. J. & Silver, P. A. Prokaryotic nanocompartments form synthetic organelles in a eukaryote. Nat. Commun. 9, 1311 (2018).

    Google Scholar 

  216. 216.

    Moon, H., Lee, J., Min, J. & Kang, S. Developing genetically engineered encapsulin protein cage nanoparticles as a targeted delivery nanoplatform. Biomacromolecules 15, 3794–3801 (2014).

    CAS  Google Scholar 

  217. 217.

    Bourdeau, R. W. et al. Acoustic reporter genes for noninvasive imaging of microorganisms in mammalian hosts. Nature 553, 86–90 (2018).

    CAS  Google Scholar 

  218. 218.

    Schüler, D. & Frankel, R. B. Bacterial magnetosomes: microbiology, biomineralization and biotechnological applications. Appl. Microbiol. Biotechnol. 52, 464–473 (1999).

    Google Scholar 

  219. 219.

    Kolinko, I. et al. Biosynthesis of magnetic nanostructures in a foreign organism by transfer of bacterial magnetosome gene clusters. Nat. Nanotechnol. 9, 193–197 (2014).

    CAS  Google Scholar 

  220. 220.

    Giessen, T. W. Encapsulins: microbial nanocompartments with applications in biomedicine, nanobiotechnology and materials science. Curr. Opin. Chem. Biol. 34, 1–10 (2016).

    CAS  Google Scholar 

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Work in the authors’ labs is supported by NHMRC EL2 Fellowship 1178715 (to C.G.) and NHMRC Program Grant 1092262 (to T.L.). The authors thank Christopher Stubenrauch and Chaille Webb for critical comments on the manuscript and Rhys Grinter for assistance with figure preparation.

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Correspondence to Chris Greening or Trevor Lithgow.

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In cases of high concentration of nucleic acids (or specific proteins) a phase separation can be observed in the cytoplasm that results in a partitioning of specific proteins that either prefer or disfavour the conditions of each phase. A notable example is the nucleolus in eukaryotes and the ribonucleoprotein granules found in bacteria such as Caulobacter crescentus.


Anabolic bacterial microcompartments that have a polyhedral protein shell and a lumen filled with the CO2-fixing enzyme RuBisCO. Examples include the β-carboxysomes of the phototroph Synechococcus elongatus and the α-carboxysomes of the lithotroph Acidithiobacillus spp.


Membrane-bound organelle that mediates light-harvesting and energy transduction in photosynthetic cyanobacteria. Through evolutionary links, thylakoids are also one of the three membrane systems found in the chloroplasts of eukaryotes.


Catabolic bacterial microcompartments bound by polyhedral protein shells and housing enzymes to oxidize a specific metabolite. Notable examples include the organelles that metabolize propanediol or ethanolamine in various bacteria and archaea, including Salmonella enterica.


First identified in single-celled eukaryotes, these organelles have a single membrane boundary. A pH gradient across the membrane is generated by a proton-translocating pyrophosphatase, providing an acidic, phosphate-rich lumen that can accommodate excess cellular calcium. Typical examples are found in Alphaproteobacteria such as Agrobacterium tumefaciens.


Eukaryotic organelles with two membrane systems that primarily mediate ATP synthesis through aerobic respiration. Derived through endosymbiosis of an alphaproteobacterial cell.


Plastids are organelles with three membrane systems found in eukaryotes, with the chloroplast being defined as a type of plastid mediating oxygenic photosynthesis in plants and algae. Derived through endosymbiosis of a cyanobacterial cell.


Membrane-bound organelles that contain a lumen filled with magnetic iron oxide or iron sulfide crystals, which enable bacteria, such as Magnetospirillum gryphiswaldense, to orient towards magnetic fields and, in turn, mediate aerotaxis.


Membrane-bound organelles that function in light harvesting and energy transduction during anoxygenic photosynthesis. Examples are found in Alphaproteobacteria such as Rhodobacter sphaeroides.


Large membrane-bound organelles containing enzymes for anaerobic ammonium oxidation in the lumen and associated energy transduction in the membrane. Exclusively found in anammox Planctomycetes such as Kuenenia stuttgartiensis.


Region within bacterial cells defined by the compacted genome. These regions are often phase-separated from the surrounding cytoplasm and therefore contain a distinct population of proteins.

Nitrate vacuoles

Giant membrane-bound organelles that store the anaerobic electron acceptor nitrate and found in various sulfur-oxidizing, nitrate-reducing Gammaproteobacteria such as Thioploca araucae.


A recently discovered group of small membrane-bound organelles that store iron, including in iron-reducing bacteria such as Shewanella putrefaciens.


Organelles with two membrane systems found in some single-celled eukaryotes, notably in the genera Giardia and Entamoeba, containing luminal enzymes required for the biosynthesis of iron–sulfur clusters. These organelles are functionally specialized derivatives of mitochondria.


Organelles with two membrane systems found in some anaerobic protists, fungi and animals. They contain luminal enzymes that produce ATP via fermentation, resulting in the production of end products such as hydrogen gas. These organelles are functionally specialized derivatives of mitochondria.


Organelles found in many animals and fungi, containing luminal enzymes required for oxidative reactions such as lipid biosynthesis and detoxification reactions. These single-membrane-bound organelles are evolutionarily related to glycosomes and glyoxysomes.


Organelles found in trypanosomes and other single-celled eukaryotes, containing enzymes required for glycolysis in their lumen. These single-membrane-bound organelles are functionally specialized derivatives of peroxisomes.


Organelles found in many plants and some other eukaryotes, containing in the lumen the enzymes required for the glyoxylate cycle. These single-membrane-bound organelles are functionally specialized derivatives of peroxisomes.


Minimalistic organelles containing a polyhedral shell made of the protein encapsulin and a lumen containing targeted cargo proteins. These organelles are widespread in bacteria and archaea, for example, supporting oxidative stress responses in Mycobacterium tuberculosis.

Gas vesicles

Protein-bound compartments that function in buoyancy for bacterial and archaeal cells, for example, to position photosynthetic bacteria in a water column to access optimal light levels; also known as gas vacuoles.


A recently defined organelle that stores polyhydroxyalkanoates and potentially other carbon storage compounds; thought to be bound by proteins rather than phospholipids.


Efficient light-harvesting organelles found in green sulfur bacteria and some other anoxygenic phototrophs. They contain a protein-coated lipid monolayer boundary and are attached to the inner membrane through a proteinaceous baseplate. Chlorosomes have been extensively studied in Chlorobaculum tepidum.

Lipid bodies

Organelles found in eukaryotes and bacteria that function in the storage and hydrolysis of triacylglycerols, wax esters and other neutral lipids. They have a boundary formed by a phospholipid monolayer that also includes protein components. Notable examples are found in Mycobacterium tuberculosis and Rhodococcus opacus; also known as lipid droplets or oil bodies.

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Greening, C., Lithgow, T. Formation and function of bacterial organelles. Nat Rev Microbiol 18, 677–689 (2020).

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