Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

Membrane lipid homeostasis in bacteria

Key Points

  • The ability of bacteria to control the biophysical properties of their membrane phospholipids allows them to thrive in a wide range of physical environments.

  • Fatty acid biosynthesis is an energy-intensive biosynthetic pathway, and the production of the building blocks for membrane phospholipids is precisely regulated to match the rate of cell division.

  • Bacteria control the homeostasis of membrane lipid biophysical properties by altering the chain length of fatty acids, as well as the ratio of saturated:unsaturated fatty acids. The de novo, type II fatty acid biosynthetic pathway is a major focal point for the regulation of fatty acid composition.

  • Bacteria can alter the physical properties of existing phospholipids by introducing cis double bonds into saturated acyl chains, thereby converting cis double bonds into cyclopropane rings and catalysing the isomerization of cis fatty acids to their trans conformations.

  • Phospholipids are used as intermediates in the formation of major structural constituents of the cell. Managing the metabolism of the lipid by-products of these biosynthetic pathways is important to prevent the accumulation of membrane-disruptive lipids and to conserve the energy that is invested in the biosynthesis of the fatty acids.

  • Modification of phosphatidylglycerol by the attachment of a lysine residue to the glycerol head group is an adaptive response that is used by pathogens to increase their resistance to cationic antibacterial peptides (defensins) that are produced by the innate immune system.

Abstract

The ability of bacteria to control the biophysical properties of their membrane phospholipids allows them to thrive in a wide range of physical environments. Bacteria precisely adjust their membrane lipid composition by modifying the types of fatty acids that are produced by the biosynthetic pathway and altering the structures of pre-existing phospholipids. The recycling of phospholipids that are used as intermediates in the biosynthesis of other major membrane components is also crucial to bilayer stability in dividing cells. Here, the principal genetic and biochemical processes that are responsible for membrane lipid homeostasis in bacteria are reviewed.

This is a preview of subscription content, access via your institution

Access options

Buy this article

Prices may be subject to local taxes which are calculated during checkout

Figure 1: Conserved pathway for the formation of phosphatidic acid in bacteria.
Figure 2: Diversification of polar head groups.
Figure 3: Modification of existing phospholipid structures in bacteria.
Figure 4: Membrane phospholipid turnover.

Similar content being viewed by others

References

  1. Cronan, J. E. Jr & Gelmann, E. P. Physical properties of membrane lipids: biological relevance and regulation. Bacteriol. Rev. 39, 232–256 (1975).

    CAS  PubMed  PubMed Central  Google Scholar 

  2. Raetz, C. R., Reynolds, C. M., Trent, M. S. & Bishop, R. E. Lipid A modification systems in Gram-negative bacteria. Annu. Rev. Biochem. 76, 295–329 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Neuhaus, F. C. & Baddiley, J. A continuum of anionic charge: structures and functions of D-alanyl-teichoic acids in Gram-positive bacteria. Microbiol. Mol. Biol. Rev. 67, 686–723 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. White, S. W., Zheng, J., Zhang, Y. M. & Rock, C. O. The structural biology of type II fatty acid biosynthesis. Annu. Rev. Biochem. 74, 791–831 (2005).

    Article  CAS  PubMed  Google Scholar 

  5. Kim, Y. M. & Prestegard, J. H. A dynamic model for the structure of acyl carrier protein in solution. Biochemistry 28, 8792–8797 (1989).

    Article  CAS  PubMed  Google Scholar 

  6. Xu, G. Y. et al. Solution structure of B. subtilis acyl carrier protein. Structure 9, 277–287 (2001).

    Article  CAS  PubMed  Google Scholar 

  7. Roujeinikova, A. et al. Structural studies of fatty acyl-(acyl carrier protein) thioesters reveal a hydrophobic binding cavity that can expand to fit longer substrates. J. Mol. Biol. 365, 135–145 (2007).

    Article  CAS  PubMed  Google Scholar 

  8. Zhang, Y. M., Marrakchi, H., White, S. W. & Rock, C. O. The application of computational methods to explore the diversity and structure of bacterial fatty acid synthase. J. Lipid Res. 44, 1–10 (2003).

    Article  CAS  PubMed  Google Scholar 

  9. De Lay, N. R. & Cronan, J. E. In vivo functional analyses of the type II acyl carrier proteins of fatty acid biosynthesis. J. Biol. Chem. 282, 20319–20328 (2007).

    Article  CAS  PubMed  Google Scholar 

  10. Cronan, J. E. Jr & Waldrop, G. L. Multi-subunit acetyl-CoA carboxylases. Prog. Lipid Res. 41, 407–435 (2002).

    Article  CAS  PubMed  Google Scholar 

  11. Jackowski, S. & Rock, C. O. Acetoacetyl-acyl carrier protein synthase, a potential regulator of fatty acid biosynthesis in bacteria. J. Biol. Chem. 262, 7927–7931 (1987). Reports the discovery of FabH, the condensing enzyme that initiates fatty acid elongation.

    CAS  PubMed  Google Scholar 

  12. Choi, K. H., Heath, R. J. & Rock, C. O. β-ketoacyl-acyl carrier protein synthase III (FabH) is a determining factor in branched-chain fatty acid biosynthesis. J. Bacteriol. 182, 365–370 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Qiu, X. et al. Crystal structure and substrate specificity of the β-ketoacyl-acyl carrier protein synthase III (FabH) from Staphylococcus aureus. Protein Sci. 14, 2087–2094 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Choi, K. H., Kremer, L., Besra, G. S. & Rock, C. O. Identification and substrate specificity of β-ketoacyl (acyl carrier protein) synthase III (mtFabH) from Mycobacterium turberculosis. J. Biol. Chem. 275, 28201–28207 (2000).

    CAS  PubMed  Google Scholar 

  15. Musayev, F., Sachdeva, S., Neel Scarsdale, J., Reynolds, K. A. & Wright, H. T. Crystal structure of a substrate complex of Mycobacterium tuberculosis β-ketoacyl-acyl carrier protein synthase III (FabH) with lauroyl-coenzyme A. J. Mol. Biol. 346, 1313–1321 (2005).

    Article  CAS  PubMed  Google Scholar 

  16. Heath, R. J. & Rock, C. O. Roles of the FabA and FabZ β-hydroxyacyl-acyl carrier protein dehydratases in Escherichia coli fatty acid biosynthesis. J. Biol. Chem. 271, 27795–27801 (1996).

    Article  CAS  PubMed  Google Scholar 

  17. Heath, R. J. & Rock, C. O. Enoyl-acyl carrier protein reductase (fabI) plays a determinant role in completing cycles of fatty acid elongation in Escherichia coli. J. Biol. Chem. 270, 26538–26542 (1995).

    Article  CAS  PubMed  Google Scholar 

  18. Heath, R. J. & Rock, C. O. A triclosan-resistant bacterial enzyme. Nature 406, 145–146 (2000). Reports the discovery of FabK, a flavoprotein enoyl-ACP reductase that substitutes for FabI.

    Article  CAS  PubMed  Google Scholar 

  19. Marrakchi, H. et al. Characterization of Streptococcus pneumoniae enoyl-(acyl carrier protein) reductase (FabK). Biochem. J. 370, 1055–1062 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Massengo-Tiasse, R. P. & Cronan, J. E. Vibrio cholerae FabV defines a new class of enoyl acyl-carrier-protein reductase. J. Biol. Chem. 283, 1308–1316 (2008).

    Article  CAS  PubMed  Google Scholar 

  21. Lu, Y. J. et al. Acyl-phosphates initiate membrane phospholipid synthesis in Gram-positive pathogens. Mol. Cell 23, 765–772 (2006). Reports the discovery of the PlsX–PlsY pathway for glycerol-phosphate acylation, which involves an acyl-phosphate intermediate. This is the principal pathway that initiates membrane phospholipid synthesis in bacteria.

    Article  CAS  PubMed  Google Scholar 

  22. Paoletti, L., Lu, Y. J., Schujman, G. E., de Mendoza, D. & Rock, C. O. Coupling of fatty acid and phospholipid synthesis in Bacillus subtilis. J. Bacteriol. 189, 5816–5824 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Lu, Y. J., Zhang, F., Grimes, K. D., Lee, R. E. & Rock, C. O. Topology and active site of PlsY: the bacterial acylphosphate:glycerol-3-phosphate acyltransferase. J. Biol. Chem. 282, 11339–11346 (2007).

    Article  CAS  PubMed  Google Scholar 

  24. Cronan, J. E. Jr & Rock, C. O. Biosynthesis of Membrane Lipids in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (eds Neidhardt, F. C. et al.) 612–636 (American Society of Microbiology, Washington DC, 1996).

    Google Scholar 

  25. Coleman, J. Characterization of the Escherichia coli gene for 1-acyl-sn-glycerol-3-phosphate acyltransferase (plsC). Mol. Gen. Genet. 232, 295–303 (1992).

    CAS  PubMed  Google Scholar 

  26. Jorasch, P., Wolter, F. P., Zahringer, U. & Heinz, E. A UDP glucosyltransferase from Bacillus subtilis successively transfers up to four glucose residues to 1,2-diacylglycerol: expression of ypfP in Escherichia coli and structural analysis of its reaction products. Mol. Microbiol. 29, 419–430 (1998).

    Article  CAS  PubMed  Google Scholar 

  27. Weissenmayer, B., Gao, J. L., Lopez-Lara, I. M. & Geiger, O. Identification of a gene required for the biosynthesis of ornithine-derived lipids. Mol. Microbiol. 45, 721–733 (2002).

    Article  CAS  PubMed  Google Scholar 

  28. Gao, J. L. et al. Identification of a gene required for the formation of lyso-ornithine lipid, an intermediate in the biosynthesis of ornithine-containing lipids. Mol. Microbiol. 53, 1757–1770 (2004).

    Article  CAS  PubMed  Google Scholar 

  29. Agun-Sunar, S. et al. Ornithine lipid is required for optimal steady-state amounts of c-type cytochromes in Rhodobacter capsulatus. Mol. Microbiol. 61, 418–435 (2006).

    Article  CAS  Google Scholar 

  30. Voelker, T. A. & Davies, H. M. Alteration of the specificity and regulation of fatty acid synthesis of Escherichia coli by expression of a plant medium-chain acyl–acyl carrier protein thioesterase. J. Bacteriol. 176, 7320–7327 (1994).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Jiang, P. & Cronan, J. E. Jr. Inhibition of fatty acid synthesis in Escherichia coli in the absence of phospholipid synthesis and release of inhibition by thioesterase action. J. Bacteriol. 176, 2814–2821 (1994).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Cho, H. & Cronan, J. E. Jr. Defective export of a periplasmic enzyme disrupts regulation of bacterial fatty acid synthesis. J. Biol. Chem. 270, 4216–4219 (1995).

    Article  CAS  PubMed  Google Scholar 

  33. Davis, M. S. & Cronan, J. E. Jr. Inhibition of Eschericia coli acetyl coenzyme A carboxylase by acyl–acyl carrier protein. J. Bacteriol. 183, 1499–1503 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Heath, R. J. & Rock, C. O. Regulation of fatty acid elongation and initiation by acyl–acyl carrier protein in Escherichia coli. J. Biol. Chem. 271, 1833–1836 (1996).

    Article  CAS  PubMed  Google Scholar 

  35. Heath, R. J. & Rock, C. O. Inhibition of β-ketoacyl-acyl carrier protein synthase III (FabH) by acyl–acyl carrier protein in Escherichia coli. J. Biol. Chem. 271, 10996–11000 (1996).

    Article  CAS  PubMed  Google Scholar 

  36. Heath, R. J., Jackowski, S. & Rock, C. O. Guanosine tetraphosphate inhibition of fatty acid and phospholipid synthesis in Escherichia coli is relieved by overexpression of glycerol-3-phosphate acyltransferase (plsB). J. Biol. Chem. 269, 26584–26590 (1994).

    CAS  PubMed  Google Scholar 

  37. Magnusson, L. U., Farewell, A. & Nystrom, T. ppGpp: a global regulator in Escherichia coli. Trends Microbiol. 13, 236–242 (2005).

    Article  CAS  PubMed  Google Scholar 

  38. Seyfzadeh, M., Keener, J. & Nomura, M. spoT-dependent accumulation of guanosine tetraphosphate in response to fatty acid starvation in Escherichia coli. Proc. Natl Acad. Sci. USA 90, 11004–11008 (1993).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Battesti, A. & Bouveret, E. Acyl carrier protein/SpoT interaction, the switch linking SpoT-dependent stress response to fatty acid metabolism. Mol. Microbiol. 62, 1048–1063 (2006).

    Article  CAS  PubMed  Google Scholar 

  40. Schujman, G. E., Paoletti, L., Grossman, A. D. & de Mendoza, D. FapR, a bacterial transcription factor involved in global regulation of membrane lipid biosynthesis. Dev. Cell 4, 663–672 (2003).

    Article  CAS  PubMed  Google Scholar 

  41. Schujman, G. E. et al. Structural basis of lipid biosynthesis regulation in Gram-positive bacteria. EMBO J. 25, 4074–4083 (2006). Identifies malonyl-CoA as the ligand for FapR, a global transcriptional regulator of fatty acid biosynthetic genes in Gram-positive bacteria.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Xie, J., Bogdanov, M., Heacock, P. & Dowhan, W. Phosphatidylethanolamine and monoglucosyldiacylglycerol are interchangeable in supporting topogenesis and function of the polytopic membrane protein lactose permease. J. Biol. Chem. 281, 19172–19178 (2006).

    Article  CAS  PubMed  Google Scholar 

  43. Zhang, W., Campbell, H. A., King, S. C. & Dowhan, W. Phospholipids as determinants of membrane protein topology. Phosphatidylethanolamine is required for the proper topological organization of the γ-aminobutyric acid permease (GabP) of Escherichia coli. J. Biol. Chem. 280, 26032–26038 (2005).

    Article  CAS  PubMed  Google Scholar 

  44. Tsatskis, Y. et al. The osmotic activation of transporter ProP is tuned by both its C-terminal coiled-coil and osmotically induced changes in phospholipid composition. J. Biol. Chem. 280, 41387–41394 (2005).

    Article  CAS  PubMed  Google Scholar 

  45. Dowhan, W. Molecular basis for membrane phospholipid diversity: why are there so many lipids? Annu. Rev. Biochem. 66, 199–232 (1997).

    Article  CAS  PubMed  Google Scholar 

  46. Rilfors, L. et al. Reconstituted phosphatidylserine synthase from Escherichia coli is activated by anionic phospholipids and micelle-forming amphiphiles. Biochim. Biophys. Acta 1438, 281–294 (1999).

    Article  CAS  PubMed  Google Scholar 

  47. Raetz, C. R. H., Larson, T. J. & Dowhan, W. Gene cloning for the isolation of enzymes of membrane lipid synthesis: phosphatidylserine synthase overproduction in Escherichia coli. Proc. Natl Acad. Sci. USA 74, 1412–1416 (1977).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  48. Tyhach, R. J., Hawrot, E., Satre, M. & Kennedy, E. P. Increased synthesis of phosphatidylserine decarboxylase in a strain of Escherichia coli bearing a hybrid plasmid. Altered association of enzyme with the membrane. J. Biol. Chem. 254, 627–633 (1979).

    CAS  PubMed  Google Scholar 

  49. Salamon, Z., Lindblom, G., Rilfors, L., Linde, K. & Tollin, G. Interaction of phosphatidylserine synthase from E. coli with lipid bilayers: coupled plasmon–waveguide resonance spectroscopy studies. Biophys. J. 78, 1400–1412 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  50. Linde, K., Grobner, G. & Rilfors, L. Lipid dependence and activity control of phosphatidylserine synthase from Escherichia coli. FEBS Lett. 575, 77–80 (2004).

    Article  CAS  PubMed  Google Scholar 

  51. Saha, S. K., Nishijima, S., Matsuzaki, H., Shibuya, I. & Matsumoto, K. A regulatory mechanism for the balanced synthesis of membrane phospholipid species in Escherichia coli. Biosci. Biotechnol. Biochem. 60, 111–116 (1996).

    Article  CAS  PubMed  Google Scholar 

  52. Saha, S. K., Furukawa, Y., Matsuzaki, H., Shibuya, I. & Matsumoto, K. Directed mutagenesis, Ser-56 to Pro, of Bacillus subtilis phosphatidylserine synthase drastically lowers enzymatic activity and relieves amplification toxicity in Escherichia coli. Biosci. Biotechnol. Biochem. 60, 630–633 (1996).

    Article  CAS  PubMed  Google Scholar 

  53. Tropp, B. E. Cardiolipin synthase from Escherichia coli. Biochim. Biophys. Acta 1348, 192–200 (1997).

    Article  CAS  PubMed  Google Scholar 

  54. Bernal, P., Munoz-Rojas, J., Hurtado, A., Ramos, J. L. & Segura, A. A Pseudomonas putida cardiolipin synthesis mutant exhibits increased sensitivity to drugs related to transport functionality. Environ. Microbiol. 9, 1135–1145 (2007).

    Article  CAS  PubMed  Google Scholar 

  55. Tropp, B. E. et al. Identity of the Escherichia coli cls and nov genes. J. Bacteriol. 177, 5155–5157 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  56. Shibuya, I., Miyazaki, C. & Ohta, A. Alteration of phospholipid composition by combined defects in phosphatidylserine and cardiolipin synthases and physiological consequences in Escherichia coli. J. Bacteriol. 161, 1086–1092 (1985).

    CAS  PubMed  PubMed Central  Google Scholar 

  57. Lopez, C. S., Alice, A. F., Heras, H., Rivas, E. A. & Sanchez-Rivas, C. Role of anionic phospholipids in the adaptation of Bacillus subtilis to high salinity. Microbiology 152, 605–616 (2006).

    Article  CAS  PubMed  Google Scholar 

  58. Hiraoka, S., Matsuzaki, H. & Shibuya, I. Active increase in cardiolipin synthesis in the stationary growth phase and its physiological significance in Escherichia coli. FEBS Lett. 336, 221–224 (1993).

    Article  CAS  PubMed  Google Scholar 

  59. Ragolia, L. & Tropp, B. E. The effects of phosphoglycerides on Escherichia coli cardiolipin synthase. Biochim. Biophys. Acta 1214, 323–332 (1994).

    Article  PubMed  Google Scholar 

  60. Zhu, K., Bayles, D. O., Xiong, A., Jayaswal, R. K. & Wilkinson, B. J. Precursor and temperature modulation of fatty acid composition and growth of Listeria monocytogenes cold-sensitive mutants with transposon-interrupted branched-chain α-ketoacid dehydrogenase. Microbiology 151, 615–623 (2005).

    Article  CAS  PubMed  Google Scholar 

  61. Giotis, E. S., McDowell, D. A., Blair, I. S. & Wilkinson, B. J. Role of branched-chain fatty acids in pH stress tolerance in Listeria monocytogenes. Appl. Environ. Microbiol. 73, 997–1001 (2007).

    Article  CAS  PubMed  Google Scholar 

  62. Cronan, J. E. Jr & Subrahmanyam, S. FadR, transcriptional co-ordination of metabolic expediency. Mol. Microbiol. 29, 937–943 (1998).

    Article  CAS  PubMed  Google Scholar 

  63. DiRusso, C. C., Heimert, T. L. & Metzger, A. K. Characterization of FadR, a global transcriptional regulator of fatty acid metabolism in Escherichia coli. Interaction with the fadB promoter is prevented by long chain fatty acyl coenzyme A. J. Biol. Chem. 267, 8685–8691 (1992).

    CAS  PubMed  Google Scholar 

  64. Henry, M. F. & Cronan, J. E. Jr. A new mechanism of transcriptional regulation: release of an activator triggered by small molecule binding. Cell 70, 671–679 (1992). Reveals that FadR is an activator of fabA gene transcription.

    Article  CAS  PubMed  Google Scholar 

  65. Campbell, J. W. & Cronan, J. E. Jr. Escherichia coli FadR positively regulates transcription of the fabB fatty acid biosynthetic gene. J. Bacteriol. 183, 5982–5990 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Zhang, Y. M., Marrakchi, H. & Rock, C. O. The FabR (YijC) transcription factor regulates unsaturated fatty acid biosynthesis in Escherichia coli. J. Biol. Chem. 277, 15558–15565 (2002). Identifies FabR as a transcription factor that regulates UFA biosynthesis as a repressor of fabA and fabB gene expression.

    Article  CAS  PubMed  Google Scholar 

  67. Wang, H. & Cronan, J. E. Functional replacement of the FabA and FabB proteins of Escherichia coli fatty acid synthesis by Enterococcus faecalis FabZ and FabF homologues. J. Biol. Chem. 279, 34489–34495 (2004).

    Article  CAS  PubMed  Google Scholar 

  68. Lu, Y. J., White, S. W. & Rock, C. O. Domain swapping between Enterococcus faecalis FabN and FabZ proteins localizes the structural determinants for isomerase activity. J. Biol. Chem. 280, 30342–30348 (2005).

    Article  CAS  PubMed  Google Scholar 

  69. Lu, Y. J. & Rock, C. O. Transcriptional regulation of fatty acid biosynthesis in Streptococcus pneumoniae. Mol. Microbiol. 59, 551–566 (2006).

    Article  CAS  PubMed  Google Scholar 

  70. Garwin, J. L., Klages, A. L. & Cronan, J. E. Jr. β-ketoacyl-acyl carrier protein synthase II of Escherichia coli. Evidence for function in the thermal regulation of fatty acid synthesis. J. Biol. Chem. 255, 3263–3265 (1980). Identifies FabF as the condensing enzyme that is responsible for the temperature-dependent change in cis -vaccenate formation.

    CAS  PubMed  Google Scholar 

  71. Ulrich, A. K., de Mendoza, D., Garwin, J. L. & Cronan, J. E. Jr. Genetic and biochemical analyses of Escherichia coli mutants altered in the temperature-dependent regulation of membrane lipid composition. J. Bacteriol. 154, 221–230 (1983).

    CAS  PubMed  PubMed Central  Google Scholar 

  72. Aquilar, P. S., Hernandez-Arriaga, A. M., Cybulski, L. E., Erazo, A. C. & de Mendoza, D. Molecular basis of thermosensing: a two component signal transduction thermometer in Bacillus subtilis. EMBO J. 20, 1681–1691 (2001). Describes DesR as a molecular sensor of the biophysical properties of membrane phospholipids.

    Article  Google Scholar 

  73. Zhu, K., Choi, K. H., Schweizer, H. P., Rock, C. O. & Zhang, Y. M. Two aerobic pathways for the formation of unsaturated fatty acids in Pseudomonas aeruginosa. Mol. Microbiol. 60, 260–273 (2006).

    Article  CAS  PubMed  Google Scholar 

  74. Zhang, Y. M., Zhu, K., Frank, M. W. & Rock, C. O. A Pseudomonas aeruginosa transcription factor that senses fatty acid structure. Mol. Microbiol. 66, 622–632 (2007).

    Article  CAS  PubMed  Google Scholar 

  75. Grogan, D. W. & Cronan, J. E. Jr. Cyclopropane ring formation in membrane lipids of bacteria. Microbiol. Mol. Biol. Rev. 61, 429–441 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  76. Wang, A. Y. & Cronan, J. E. Jr. The growth phase dependent synthesis of cyclopropane fatty acids in Escherichia coli is due to an RpoS (KatF) dependent promoter plus enzyme instability. Mol. Microbiol. 11, 1009–1017 (1994).

    Article  PubMed  Google Scholar 

  77. Chang, Y. Y., Eichel, J. & Cronan, J. E. Jr. Metabolic instability of Escherichia coli cyclopropane fatty acid synthase is due to RpoH-dependent proteolysis. J. Bacteriol. 182, 4288–4294 (2000). Together with reference 76, describes the dual regulation of Cfa by RpoH- and RpoS-dependent mechanisms.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  78. Chang, Y. Y. & Cronan, J. E. Jr. Membrane cyclopropane fatty acid content is a major factor in acid resistance of Escherichia coli. Mol. Microbiol. 33, 249–259 (1999).

    Article  CAS  PubMed  Google Scholar 

  79. Munoz-Rojas, J. et al. Involvement of cyclopropane fatty acids in the response of Pseudomonas putida KT2440 to freeze-drying. Appl. Environ. Microbiol. 72, 472–477 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  80. Glickman, M. S., Cox, J. S. & Jacobs, W. R. Jr. A novel mycolic acid cyclopropane synthetase is required for cording, persistence, and virulence of Mycobacterium tuberculosis. Mol. Cell 5, 717–727 (2000). Shows that cyclopropane fatty acid formation is required for the persistence and virulence of M. tuberculosis.

    Article  CAS  PubMed  Google Scholar 

  81. Morita, N. et al. Evidence for cistrans isomerization of a double bond in the fatty acids of the psychrophilic bacterium Vibrio sp. strain ABE-1. J. Bacteriol. 175, 916–918 (1993).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Heipieper, H. J., Meulenbeld, G., van Oirschot, Q. & de Bont, J. A. M. Effect of environmental factors on the trans/cis ratio of unsaturated fatty acids in Pseudomonas putida S12. Appl. Environ. Microbiol. 62, 2773–2777 (1996).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. Junker, F. & Ramos, J. L. Involvement of the cis/trans isomerase Cti in solvent resistance of Pseudomonas putida DOT-T1E. J. Bacteriol. 181, 5693–5700 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. Holtwick, R., Meinhardt, F. & Keweloh, H. cistrans isomerization of unsaturated fatty acids: cloning and sequencing of the cti gene from Pseudomonas putida P8. Appl. Environ. Microbiol. 63, 4292–4297 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Kiran, M. D., Annapoorni, S., Suzuki, I., Murata, N. & Shivaji, S. cistrans isomerase gene in psychrophilic Pseudomonas syringae is constitutively expressed during growth and under conditions of temperature and solvent stress. Extremophiles 9, 117–125 (2005).

    Article  CAS  PubMed  Google Scholar 

  86. Halverson, L. J. & Firestone, M. K. Differential effects of permeating and nonpermeating solutes on the fatty acid composition of Pseudomonas putida. Appl. Environ. Microbiol. 66, 2414–2421 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Peschel, A. How do bacteria resist human antimicrobial peptides? Trends Microbiol. 10, 179–186 (2002).

    Article  CAS  PubMed  Google Scholar 

  88. Oku, Y., Kurokawa, K., Ichihashi, N. & Sekimizu, K. Characterization of the Staphylococcus aureus mprF gene, involved in lysinylation of phosphatidylglycerol. Microbiology 150, 45–51 (2004).

    Article  CAS  PubMed  Google Scholar 

  89. Peschel, A. et al. Staphylococcus aureus resistance to human defensins and evasion of neutrophil killing via the novel virulence factor MprF is based on modification of membrane lipids with L-lysine. J. Exp. Med. 193, 1067–1076 (2001). Connects the formation of lysyl-phosphatidylglycerol with the resistance of bacteria to defensins.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  90. Staubitz, P. & Peschel, A. MprF-mediated lysinylation of phospholipids in Bacillus subtilis — protection against bacteriocins in terrestrial habitats? Microbiology 148, 3331–3332 (2002).

    Article  CAS  PubMed  Google Scholar 

  91. Thedieck, K. et al. The MprF protein is required for lysinylation of phospholipids in listerial membranes and confers resistance to cationic antimicrobial peptides (CAMPs) on Listeria monocytogenes. Mol. Microbiol. 62, 1325–1339 (2006).

    Article  CAS  PubMed  Google Scholar 

  92. Nishi, H., Komatsuzawa, H., Fujiwara, T., McCallum, N. & Sugai, M. Reduced content of lysyl-phosphatidylglycerol in the cytoplasmic membrane affects susceptibility to moenomycin, as well as vancomycin, gentamicin, and antimicrobial peptides, in Staphylococcus aureus. Antimicrob. Agents Chemother. 48, 4800–4807 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  93. Goldberg, D. E., Rumley, M. K. & Kennedy, E. P. Biosynthesis of membrane-derived oligosaccharides: a periplasmic phosphoglyceroltransferase. Proc. Natl Acad. Sci. USA 78, 5513–5517 (1981).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Kennedy, E. P. Osmotic regulation and biosynthesis of membrane-derived oligosaccharides in Escherichia coli. Proc. Natl Acad. Sci. USA 79, 1092–1095 (1982).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  95. Raetz, C. R. H. & Newman, K. F. Diglyceride kinase mutants of Escherichia coli: inner membrane association of 1,2-diglyceride and its relation to synthesis of membrane-derived oligosaccharides. J. Bacteriol. 137, 860–868 (1979).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. Rumley, M. K., Therisod, H., Weissborn, A. C. & Kennedy, E. P. Mechanisms of regulation of the biosynthesis of membrane-derived oligosaccharides in Escherichia coli. J. Biol. Chem. 267, 11806–11810 (1992).

    CAS  PubMed  Google Scholar 

  97. Lacroix, J. M., Loubens, I., Tempete, M., Menichi, B. & Bohin, J. P. The mdoA locus of Escherichia coli consists of an operon under osmotic control. Mol. Microbiol. 5, 1745–1753 (1991).

    Article  CAS  PubMed  Google Scholar 

  98. Raetz, C. R. H. & Newman, K. F. Neutral lipid accumulation in the membranes of Escherichia coli mutants lacking diglyceride kinase. J. Biol. Chem. 253, 3882–3887 (1978).

    CAS  PubMed  Google Scholar 

  99. Bielawska, A., Perry, D. K. & Hannun, Y. A. Determination of ceramides and diglycerides by the diglyceride kinase assay. Anal. Biochem. 298, 141–150 (2001).

    Article  CAS  PubMed  Google Scholar 

  100. Jerga, A., Lu, Y. J., Schujman, G. E., de Mendoza, D. & Rock, C. O. Identification of a soluble diacylglycerol kinase required for lipoteichoic acid production in Bacillus subtilis. J. Biol. Chem. 282, 21738–21745 (2007).

    Article  CAS  PubMed  Google Scholar 

  101. Koch, H. U., Haas, R. & Fischer, W. The role of lipoteichoic acid biosynthesis in membrane lipid metabolism of growing Staphylococcus aureus. Eur. J. Biochem. 138, 357–363 (1984).

    Article  CAS  PubMed  Google Scholar 

  102. Taron, D. J., Childs, W. C. & Neuhaus, F. C. Biosynthesis of D-alanyl-lipoteichoic acid: role of diglyceride kinase in the synthesis of phosphatidylglycerol for chain elongation. J. Bacteriol. 154, 1110–1116 (1983).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. Jackowski, S., Jackson, P. D. & Rock, C. O. Sequence and function of the aas gene in Escherichia coli. J. Biol. Chem. 269, 2921–2928 (1994).

    CAS  PubMed  Google Scholar 

  104. Harvat, E. M. et al. Lysophospholipid flipping across the Escherichia coli inner membrane catalyzed by a transporter (LplT) belonging to the major facilitator superfamily. J. Biol. Chem. 280, 12028–12034 (2005).

    Article  CAS  PubMed  Google Scholar 

  105. Hsu, L., Jackowski, S. & Rock, C. O. Isolation and characterization of Escherichia coli K-12 mutants lacking both 2-acyl-glycerophosphoethanolamine acyltransferase and acyl–acyl carrier protein synthetase activity. J. Biol. Chem. 266, 13783–13788 (1991).

    CAS  PubMed  Google Scholar 

  106. Zhang, Y. M., White, S. W. & Rock, C. O. Inhibiting bacterial fatty acid synthesis. J. Biol. Chem. 281, 17541–17544 (2006).

    Article  CAS  PubMed  Google Scholar 

  107. Revill, W. P., Bibb, M. J. & Hopwood, D. A. Purification of a malonyltransferase from Streptomyces coelicolor A3(2) and analysis of its genetic determinant. J. Bacteriol. 177, 3946–3952 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  108. Zheng, C. J. et al. Cephalochromin, a FabI-directed antibacterial of microbial origin. Biochem. Biophys. Res. Commun. 362, 1107–1112 (2007).

    Article  CAS  PubMed  Google Scholar 

  109. McMurray, L. M., Oethinger, M. & Levy, S. Triclosan targets lipid synthesis. Nature 394, 531–532 (1998).

    Article  CAS  Google Scholar 

  110. Heath, R. J. et al. Mechanism of triclosan inhibition of bacterial fatty acid synthesis. J. Biol. Chem. 274, 11110–11114 (1999).

    Article  CAS  PubMed  Google Scholar 

  111. Zheng, C. J., Sohn, M. J. & Kim, W. G. Atromentin and leucomelone, the first inhibitors specific to enoyl-ACP reductase (FabK) of Streptococcus pneumoniae. J. Antibiot. 59, 808–812 (2006).

    Article  CAS  Google Scholar 

  112. Takahata, S. et al. AG205, a novel agent directed against FabK of Streptococcus pneumoniae. Antimicrob. Agents Chemother. 50, 2869–2871 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

Research in the authors' laboratory is supported by National Institute of General Medical Sciences grant GM34496, Cancer Center (CORE) Support grant CA21765 and the American Lebanese Syrian Associated Charities.

Author information

Authors and Affiliations

Authors

Related links

Related links

DATABASES

Entrez Genome Project

Bacillus subtilis

Enterococcus faecalis

Escherichia coli

Listeria monocytogenes

Mycobacterium tuberculosis

Pseudomonas putida

Streptococcus pneumoniae

FURTHER INFORMATION

Charles O. Rock's homepage

Glossary

Type II fatty acid biosynthetic pathway

The pathway by which fatty acids are synthesized in bacteria, chloroplasts and mitochondria. Distinct enzymes, each of which are encoded by an individual gene, catalyse the different steps of the pathway.

Lysophosphatidic acid

1-acyl-sn-glycero-3-phosphate; the first intermediate in membrane phospholipid formation.

Phosphatidic acid

1,2-diacyl-sn-glycero-3-phosphate; the key intermediate in the formation of most bacterial membrane phospholipids.

Zwitterionic

A compound that is electrically neutral, but that carries formal positive and negative charges on different atoms.

RpoS

A specific sigma factor that is induced as cells enter the stationary phase of growth.

Sigma factor

A transcription initiation factor that enables the binding of RNA polymerase to gene promoters.

RpoH

A specific sigma factor that is induced in response to heat shock.

Defensin

A cationic peptide that is produced by the innate immune system and that kills bacteria by disrupting the phospholipid bilayer.

Undecaprenol

A 55-carbon isoprenoid alcohol that functions as a carrier of sugars in the synthesis of peptidoglycan.

Major facilitator superfamily

A large family of transporters that has varied substrate specificity; members of this family possess 12–14 transmembrane segments.

Rights and permissions

Reprints and permissions

About this article

Cite this article

Zhang, YM., Rock, C. Membrane lipid homeostasis in bacteria. Nat Rev Microbiol 6, 222–233 (2008). https://doi.org/10.1038/nrmicro1839

Download citation

  • Issue Date:

  • DOI: https://doi.org/10.1038/nrmicro1839

This article is cited by

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing