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Hopanoid lipids: from membranes to plant–bacteria interactions

Key Points

  • Hopanes were discovered by petroleum geologists as ubiquitous molecular fossils in ancient sedimentary rocks. Later, bacterial hopanoids were identified as their progenitors.

  • Today, phylogenetically diverse bacteria make hopanoids using machinery encoded by a conserved set of genes. The rhizosphere appears to be a niche that is common to many hopanoid-producing bacteria, and some hopanoid producers are known plant symbionts.

  • Bacteria make structurally distinct types of hopanoids, including ones that can covalently bind lipid A. Different hopanoid classes exhibit different properties and likely have specific biological functions.

  • Hopanoids share similar biophysical properties with sterols, such as tuning membrane rigidity and permeability. Though some evidence suggests that hopanoids help order membranes, how such ordering impacts cells and whether hopanoids interact with particular membrane proteins remain to be determined.

  • In vitro, hopanoids contribute to bacterial stress resistance, which may help explain their ability to facilitate beneficial plant–bacteria interactions. However, given that hopanoids can also serve as carriers for plant hormones and that plants themselves make hopanoid-like compounds, it is likely that other mechanisms are additionally at play.


Lipid research represents a frontier for microbiology, as showcased by hopanoid lipids. Hopanoids, which resemble sterols and are found in the membranes of diverse bacteria, have left an extensive molecular fossil record. They were first discovered by petroleum geologists. Today, hopanoid-producing bacteria remain abundant in various ecosystems, such as the rhizosphere. Recently, great progress has been made in our understanding of hopanoid biosynthesis, facilitated in part by technical advances in lipid identification and quantification. A variety of genetically tractable, hopanoid-producing bacteria have been cultured, and tools to manipulate hopanoid biosynthesis and detect hopanoids are improving. However, we still have much to learn regarding how hopanoid production is regulated, how hopanoids act biophysically and biochemically, and how their production affects bacterial interactions with other organisms, such as plants. The study of hopanoids thus offers rich opportunities for discovery.

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Figure 1: Pathways of sterol and hopanoid biosynthesis.
Figure 2: Regulation of lipid raft formation by hopanoids.
Figure 3: Hopanoid-rich vesicles in Frankia spp.
Figure 4: Chemical structure of hopanoid-lipid A from Bradyrhizobium spp.


  1. 1

    Fahy, E., Sud, M., Cotter, D. & Subramaniam, S. LIPID MAPS online tools for lipid research. Nucleic Acids Res. 35, W606–W612 (2007).

    PubMed  PubMed Central  Google Scholar 

  2. 2

    Rohmer, M., Bouvier, P. & Ourisson, G. Molecular evolution of biomembranes: structural equivalents and phylogenetic precursors of sterols. Proc. Natl Acad. Sci. USA 76, 847–851 (1979).

    CAS  PubMed  Google Scholar 

  3. 3

    Burlingame, A. L., Haug, P. & Belsky, T. Calvin, M. Occurrence of biogenic steranes and pentacyclic triterpanes in an Eocene shale (52 millions years) and in an early Pre-Cambrian shale (2.7 billion years): a preliminary report. Proc. Natl Acad. Sci. USA 54, 1406–1412 (1965).

    CAS  PubMed  Google Scholar 

  4. 4

    Brocks, J. J. & Banfield, J. Biomarker evidence for green and purple sulphur bacteria in a stratified Paleoproterozoic sea. Nature 437, 866–870 (2005).

    CAS  PubMed  Google Scholar 

  5. 5

    Ourisson, G. & Albrecht, P. Geohopanoids: the most abundant natural products on Earth? Acc. Chem. Res. 25, 398–402 (1992).

    CAS  Google Scholar 

  6. 6

    Bird, C. W., Lynch, J. M., Pirt, S. J. & Reid, W. W. The identification of hop-22(29)-ene in prokaryotic organisms. Tetrahedron Lett. 12, 3189–3190 (1971).

    Google Scholar 

  7. 7

    De Rosa, M., Gambacorta, A., Minale, L. & Bu'Lock, J. D. Bacterial triterpenes. J. Chem. Soc. D Chem. Commun., 619–620 (1971). References 6 and 7 were the first papers to discover hopanoids in bacteria.

  8. 8

    Newman, D. K., Neubauer, C., Ricci, J. N., Wu, C.-H. & Pearson, A. Cellular and molecular biological approaches to interpreting ancient biomarkers. Annu. Rev. Earth Planet. Sci. 44, 493–522 (2016).

    CAS  Google Scholar 

  9. 9

    Pearson, A. in Treatise on Geochemistry 2nd edn (eds Falkowski, P. G. & Freeman, K. H.) 291–336. (Elsevier, London, 2014).

    Google Scholar 

  10. 10

    Vranová, E., Coman, D. & Gruissem, W. Network analysis of the MVA and MEP pathways for isoprenoid synthesis. Annu. Rev. Plant Biol. 64, 665–700 (2013).

    PubMed  Google Scholar 

  11. 11

    Abe, I. Enzymatic synthesis of cyclic triterpenes. Nat. Prod. Rep. 24, 1311–1321 (2007).

    CAS  PubMed  Google Scholar 

  12. 12

    Christianson, D. W. Structural biology and chemistry of the terpenoid cyclases. Chem. Rev. 106, 3412–3442 (2006).

    CAS  PubMed  Google Scholar 

  13. 13

    Barton, D. H. et al. Biosynthesis of 3beta-hydroxy-triterpenoids and -steroids from (3S)-2,3-epoxy-2,3-dihydrosqualene. J. Chem. Soc. Perkin Trans. 1 12, 1134–1138 (1975).

    Google Scholar 

  14. 14

    Wei, J. H., Yin, X. & Welander, P. V. Sterol synthesis in diverse Bacteria. Front. Microbiol. 7, 259–219 (2016).

    Google Scholar 

  15. 15

    Rohmer, M., Anding, C. & Ourisson, G. Non-specific biosynthesis of hopane triterpenes by a cell-free system from Acetobacter pasteurianum. Eur. J. Biochem. 112, 541–547 (1980).

    CAS  PubMed  Google Scholar 

  16. 16

    Poralla, K. The possible role of a repetitive amino acid motif in evolution of triterpenoid cyclases. Bioorg. Med. Chem. Lett. 4, 285–290 (1994).

    CAS  Google Scholar 

  17. 17

    Ourisson, G. & Nakatani, Y. The terpenoid theory of the origin of cellular life: the evolution of terpenoids to cholesterol. Chem. Biol. 1, 11–23 (1994).

    CAS  PubMed  Google Scholar 

  18. 18

    Syrén, P.-O., Henche, S., Eichler, A., Nestl, B. M. & Hauer, B. Squalene-hopene cyclases — evolution, dynamics and catalytic scope. Curr. Opin. Struct. Biol. 41, 73–82 (2016).

    PubMed  Google Scholar 

  19. 19

    Ourisson, G., Albrecht, P. & Rohmer, M. Predictive microbial biochemistry — from molecular fossils to procaryotic membranes. Trends Biochem. Sci. 7, 236–239 (1982).

    CAS  Google Scholar 

  20. 20

    Fischer, W. W. & Pearson, A. Hypotheses for the origin and early evolution of triterpenoid cyclases. Geobiology 5, 19–34 (2007).

    CAS  Google Scholar 

  21. 21

    Banta, A. B., Wei, J. H. & Welander, P. V. A distinct pathway for tetrahymanol synthesis in bacteria. Proc. Natl Acad. Sci. USA 112, 13478–13483 (2015).

    CAS  PubMed  Google Scholar 

  22. 22

    Fischer, W. W., Summons, R. E. & Pearson, A. Targeted genomic detection of biosynthetic pathways: anaerobic production of hopanoid biomarkers by a common sedimentary microbe. Geobiology 3, 33–40 (2005).

    CAS  Google Scholar 

  23. 23

    Ourisson, G., Albrecht, P. & Rohmer, M. The hopanoids: palaeochemistry and biochemistry of a group of natural products. Pure Appl. Chem. 51, 709–729 (1979). This paper is a concise compendium of key early references that describe the discovery of fossil hopanes in sedimentary rocks and their origin as bacterial hopanoids.

    CAS  Google Scholar 

  24. 24

    Racolta, S., Juhl, P. B., Sirim, D. & Pleiss, J. The triterpene cyclase protein family: a systematic analysis. Proteins 80, 2009–2019 (2012).

    CAS  PubMed  Google Scholar 

  25. 25

    Devon, T. K. & Scott, A. I. in Handbook of Naturally Occurring Compounds Vol. 2 281–389 (Academic Press, New York, 1972).

    Google Scholar 

  26. 26

    Inayama, S., Hori, H. & Pang, G. M. Isolation of a hopane-type triterpenoid, zeorin, from a higher plant, Tripterygium regelii. Chem. Pharm. Bull. 37, 2836–2837 (1989). This work provides a short review of hopanoid-like triterpenoids found in plants and lichens.

    CAS  Google Scholar 

  27. 27

    Shinozaki, J., Shibuya, M., Masuda, K. & Ebizuka, Y. Squalene cyclase and oxidosqualene cyclase from a fern. FEBS Lett. 582, 310–318 (2008).

    CAS  PubMed  Google Scholar 

  28. 28

    Frickey, T. & Kannenberg, E. Phylogenetic analysis of the triterpene cyclase protein family in prokaryotes and eukaryotes suggests bidirectional lateral gene transfer. Environ. Microbiol. 11, 1224–1241 (2009).

    CAS  PubMed  Google Scholar 

  29. 29

    Lopes, D., Villela, C. T., Mac Kaplan & Carauta, J. Moretenolactone, a β-lactone hopanoid from Ficus insipida. Phytochem 34, 279–280 (1993).

    CAS  Google Scholar 

  30. 30

    Perzl, M. et al. Cloning of conserved genes from Zymomonas mobilis and Bradyrhizobium japonicum that function in the biosynthesis of hopanoid lipids. Biochim. Biophys. Acta 1393, 108–118 (1998).

    CAS  PubMed  Google Scholar 

  31. 31

    Bradley, A. S., Pearson, A., Sáenz, J. P. & Marx, C. J. Adenosylhopane: The first intermediate in hopanoid side chain biosynthesis. Org. Geochem. 41, 1075–1081 (2010).

    CAS  Google Scholar 

  32. 32

    Flesch, G. & Rohmer, M. Prokaryotic hopanoids: the biosynthesis of the bacteriohopane skeleton. Formation of isoprenic units from two distinct acetate pools and a novel type of carbon/carbon linkage between a triterpene and D-ribose. Eur. J. Biochem. 175, 405–411 (1988).

    CAS  PubMed  Google Scholar 

  33. 33

    Rohmer, M., Sutter, B. & Sahm, H. Bacterial sterol surrogates. Biosynthesis of the side-chain of bacteriohopanetetrol and of a carbocyclic pseudopentose from 13C-labelled glucose in Zymomonas mobilis. J. Chem. Soc. Chem. Commun. 19, 1471–1472 (1989).

    Google Scholar 

  34. 34

    Welander, P. V. et al. Identification and characterization of Rhodopseudomonas palustris TIE-1 hopanoid biosynthesis mutants. Geobiology 10, 163–177 (2012). This work shows that hpnH is essential for formation of all extended hopanoids.

    CAS  PubMed  PubMed Central  Google Scholar 

  35. 35

    Liu, W. et al. Ribosylhopane, a novel bacterial hopanoid, as precursor of C35 bacteriohopanepolyols in Streptomyces coelicolor A3(2). ChemBioChem 15, 2156–2161 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. 36

    Schmerk, C. L. et al. Elucidation of the Burkholderia cenocepacia hopanoid biosynthesis pathway uncovers functions for conserved proteins in hopanoid-producing bacteria. Environ. Microbiol. 17, 735–750 (2014). This work identifies hopanoid biosynthesis-associated glycosyltransferase protein HpnI, hopanoid biosynthesis-associated radical SAM protein HpnJ and hopanoid biosynthesis-associated protein HpnK as enzymes that modify extended hopanoid tails.

    PubMed  Google Scholar 

  37. 37

    Welander, P. V., Coleman, M. L., Sessions, A. L., Summons, R. E. & Newman, D. K. Identification of a methylase required for 2-methylhopanoid production and implications for the interpretation of sedimentary hopanes. Proc. Natl Acad. Sci. USA 19, 8537–8542 (2010).

    Google Scholar 

  38. 38

    Welander, P. V. & Summons, R. E. Discovery, taxonomic distribution, and phenotypic characterization of a gene required for 3-methylhopanoid production. Proc. Natl Acad. Sci. USA 109, 12905–12910 (2012).

    CAS  PubMed  Google Scholar 

  39. 39

    Ricci, J. N., Michel, A. J. & Newman, D. K. Phylogenetic analysis of HpnP reveals the origin of 2-methylhopanoid production in Alphaproteobacteria. Geobiology 13, 267–277 (2015).

    CAS  PubMed  Google Scholar 

  40. 40

    Rohmer, M., Bouvier, P. & Ourisson, G. Non-specific lanosterol and hopanoid biosynthesis by a cell-free system from the bacterium Methylococcus capsulatus. Eur. J. Biochem. 112, 557–560 (1980).

    CAS  PubMed  Google Scholar 

  41. 41

    Herrmann, D. A non-extractable triterpenoid of the hopane series in Acetobacter xylinum. FEMS Microbiol. Lett. 135, 323–326 (1996).

    CAS  Google Scholar 

  42. 42

    Cvejic, J. et al. Bacterial triterpenoids of the hopane series as biomarkers for the chemotaxonomy of Burkholderia. Pseudomonas and Ralstonia spp. FEMS Microbiol. Lett. 183, 295–299 (2000).

    CAS  PubMed  Google Scholar 

  43. 43

    Cvejic, J. et al. Bacterial triterpenoids of the hopane series from the methanotrophic bacteria Methylocaldum spp.: phylogenetic implications and first evidence for an unsaturated aminobacteriohopanepolyol. FEMS Microbiol. Lett. 182, 361–365 (2000).

    CAS  PubMed  Google Scholar 

  44. 44

    Rohmer, M. & Ourisson, G. Unsaturated bacteriohopanepolyols from Acetobacter aceti ssp. xylinum. J. Chem. Res. (S), 356–357; (M), 3037–3059 (1976).

  45. 45

    Bligh, E. & Dyer, W. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917 (1959).

    CAS  Google Scholar 

  46. 46

    Sessions, A. L. et al. Identification and quantification of polyfunctionalized hopanoids by high temperature gas chromatography-mass spectrometry. Org. Geochem. 56, 120–130 (2013). This work establishes an optimized protocol for identification of structurally diverse hopanoids by gas chromatography.

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 47

    Talbot, H. et al. Analysis of intact bacteriohopanepolyols from methanotrophic bacteria by reversed-phase high performance liquid chromatography-atmospheric pressure chemical ionization mass spectrometry. J. Chromatogr. A 921, 175–185 (2001). This work establishes an optimized protocol for identification of various extended hopanoids by high-performance liquid chromatography.

    CAS  PubMed  Google Scholar 

  48. 48

    Wu, C.-H. et al. Quantitative hopanoid analysis enables robust pattern detection and comparison between laboratories. Geobiology 13, 391–407 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. 49

    Tahara, Y., Yamashita, T., Kondo, M. & Yamada, Y. Isolation of Zymomonas mobilis mutant deficient in tetrahydroxybacteriohopane biosynthesis. Agr. Biol. Chem. 52, 3189–3190 (1988).

    CAS  Google Scholar 

  50. 50

    Jürgens, U. J., Simonin, P. & Rohmer, M. Localization and distribution of hopanoids in membrane systems of the cyanobacterium Synechocystis PCC 6714. FEMS Microbiol. Lett. 1097, 90723–90722 (1992).

    Google Scholar 

  51. 51

    Kleemann, G., Alskog, G., Berry, A. M. & Huss-Danell, K. Lipid composition and nitrogenase activity of symbiotic Frankia (Alnus incana) in response to different oxygen concentrations. Protoplasma 183, 107–115 (1994).

    CAS  Google Scholar 

  52. 52

    Doughty, D. M., Hunter, R. C., Summons, R. E. & Newman, D. K. 2-Methylhopanoids are maximally produced in akinetes of Nostoc punctiforme: geobiological implications. Geobiology 7, 524–532 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. 53

    Wu, C.-H., Bialecka-Fornal, M. & Newman, D. K. Methylation at the C-2 position of hopanoids increases rigidity in native bacterial membranes. eLife 4, e05663 (2015). This work describes the specific effects of 2Me-hopanoids on the fluidity of membranes and compares the contribution of hopanoids to fluidity among inner, outer and native membranes.

    PubMed Central  Google Scholar 

  54. 54

    Doughty, D. M. et al. The RND-family transporter, HpnN, is required for hopanoid localization to the outer membrane of Rhodopseudomonas palustris TIE-1. Proc. Natl Acad. Sci. USA 108, E1045–1051 (2011).

    CAS  PubMed  Google Scholar 

  55. 55

    Kumar, N. et al. Crystal structures of the Burkholderia multivorans hopanoid transporter HpnN. Proc. Natl Acad. Sci. USA 114, 6557–6562 (2017).

    CAS  PubMed  Google Scholar 

  56. 56

    Sandhu, P. & Akhter, Y. Evolution of structural fitness and multifunctional aspects of mycobacterial RND family transporters. Arch. Microbiol. (2017).

    PubMed  Google Scholar 

  57. 57

    Kannenberg, E., Poralla, K. & Blume, A. A hopanoid from the thermo-acidophilic Bacillus acidocaldarius condenses membranes. Naturwissenschaften 67, 458 (1980).

    CAS  Google Scholar 

  58. 58

    Poralla, K., Kannenberg, E. & Blume, A. A glycolipid containing hopane isolated from the acidophilic, thermophilic Bacillus acidocaldarius, has a cholesterol-like function in membranes. FEBS Lett. 113, 107–110 (1980). This work provides the first evidence that hopanoids can condense and increase the stability of lipid monolayers similarly to cholesterol.

    CAS  PubMed  Google Scholar 

  59. 59

    Kannenberg, E., Blume, A. McElhaney, R. N. & Poralla, K. Monolayer and calorimetric studies of phosphatidylcholines containing branched-chain fatty acids and of their interactions with cholesterol and with a bacterial hopanoid in model membranes. Biochim. Biophys. Acta Biomembr. 733, 111–116 (1983).

    CAS  Google Scholar 

  60. 60

    Leonard, A. & Milon, A. Modulation of membrane hydrophobic thickness by cholesterol, cycloartenol and hopanoid. A solid state 2H-NMR Study. Bull. Magn. Reson. 15, 124–127 (1993).

    CAS  Google Scholar 

  61. 61

    Nagimo, A., Sato, Y. & Suzuki, Y. Electron spin resonance studies of phosphatidylcholine interacted with cholesterol and with a hopanoid in liposomal membrane. Chem. Pharm. Bull. 39, 3071–3074 (1991).

    CAS  PubMed  Google Scholar 

  62. 62

    Chen, Z., Tanno, N. & Takenaka, S. Effects of bacteriohopane-32-ol on the stability of various kinds of liposomal membranes. Biol. Pharm. Bull. 18, 600–604 (1995).

    CAS  PubMed  Google Scholar 

  63. 63

    Sato, Y., Chen, Z. & Suzuki, Y. Thermodynamic effects of hopanoids on synthetic and bacterial phospholipid membranes. Chem. Pharm. Bull. 43, 1241–1244 (1995).

    CAS  Google Scholar 

  64. 64

    Chen, Z., Sato, Y., Nakazawa, I. & Suzuki, Y. Interactions between bacteriohopane-32, 33, 34, 35-tetrol and liposomal membranes composed of dipalmitoylphosphatidylcholine. Biol. Pharm. Bull. 18, 477–480 (1995).

    CAS  PubMed  Google Scholar 

  65. 65

    Kannenberg, E. & Poralla, K. The influence of hopanoids on growth of Mycoplasma mycoides. Arch. Microbiol. 133, 100 (1983).

    Google Scholar 

  66. 66

    Stroeve, P., Pettit, C. D., Vasquez, V., Kim, I. & Berry, A. M. Surface active behavior of hopanoid lipids: bacteriohopanetetrol and phenylacetate monoester bacteriohopanetetrol. Langmuir 14, 4261–4265 (1998).

    CAS  Google Scholar 

  67. 67

    Poger, D. & Mark, A. E. The relative effect of sterols and hopanoids on lipid bilayers: when comparable is not identical. J. Phys. Chem. B. 117, 16129–16140 (2013).

    CAS  PubMed  Google Scholar 

  68. 68

    Neubauer, C. et al. Lipid remodeling in Rhodopseudomonas palustris TIE-1 upon loss of hopanoids and hopanoid methylation. Geobiology 13, 443–453 (2015).

    CAS  PubMed  Google Scholar 

  69. 69

    Bradley, A. S. et al. Hopanoid-free Methylobacterium extorquens DM4 overproduces carotenoids and has widespread growth impairment. PLoS ONE 12, e0173323 (2017).

    PubMed  PubMed Central  Google Scholar 

  70. 70

    Gruszecki, W. I. & Strzalka, K. Carotenoids as modulators of lipid membrane physical properties. Biochim. Biophys. Acta 1740, 108–115 (2005).

    CAS  PubMed  Google Scholar 

  71. 71

    Pike, L. J. The challenge of lipid rafts. J. Lipid Res. 50 (Suppl.), S323–S328 (2009).

    PubMed  PubMed Central  Google Scholar 

  72. 72

    Gumí- Audenis, B. et al. Structure and nanomechanics of model membranes by atomic force microscopy and spectroscopy: insights into the role of cholesterol and sphingolipids. Membranes 6, 58 (2016).

    Google Scholar 

  73. 73

    Brown, D. A. & London, E. Structure and function of sphingolipid- and cholesterol-rich membrane rafts. J. Biol. Chem. 275, 17221–17224 (2000).

    CAS  PubMed  Google Scholar 

  74. 74

    Sezgin, E., Levental, I., Mayor, S. & Eggeling, C. The mystery of membrane organization: composition, regulation, and roles of lipid rafts. Nat. Rev. Mol. Cell Biol. 18, 361–374 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. 75

    Farnoud, A. M., Toledo, A. M., Konopka, J. B., Del Poeta, M. & London, E. Raft-like membrane domains in pathogenic microorganisms. Curr. Top. Membr. 75, 233–268 (2015).

    PubMed  PubMed Central  Google Scholar 

  76. 76

    Bramkamp, M. & Lopez, D. Exploring the existence of lipid rafts in bacteria. Microbiol. Mol. Biol. Rev. 9, 81–100 (2015).

    Google Scholar 

  77. 77

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

    CAS  PubMed  Google Scholar 

  78. 78

    Sáenz, J. P., Sezgin, E., Schwille, P. & Simons, K. Functional convergence of hopanoids and sterols in membrane ordering. Proc. Natl Acad. Sci. USA 109, 14236–14240 (2012). This work demonstrates that hopanoids can confer membrane order via formation of liquid-ordered states in synthetic liposomes.

    PubMed  Google Scholar 

  79. 79

    Sáenz, J. P. et al. Hopanoids as functional analogues of cholesterol in bacterial membranes. Proc. Natl Acad. Sci. USA, 112, 11971–11976 (2015). This work demonstrates that hopanoids facilitate ordering of bacterial outer membranes via interaction with saturated lipids such as lipid A as well as via enhancing multidrug efflux.

    Google Scholar 

  80. 80

    Sáenz, J. P. Hopanoid enrichment in a detergent resistant membrane fraction of Crocosphaera watsonii: implications for bacterial lipid raft formation. Org. Geochem. 41, 853–856 (2010).

    Google Scholar 

  81. 81

    Doughty, D. M., Dieterle, M., Sessions, A. L., Fischer, W. W. & Newman, D. K. Probing the subcellular localization of hopanoid lipids in bacteria using nanoSIMS. PLoS ONE 9, e84455 (2014).

    PubMed  PubMed Central  Google Scholar 

  82. 82

    Flesch, G. & Rohmer, M. Growth inhibition of hopanoid synthesizing bacteria by squalene cyclase inhibitors. Arch. Microbiol. 147, 100–104 (1987).

    CAS  Google Scholar 

  83. 83

    Horbach, S., Neuss, B. & Sahm, H. Effect of azasqualene on hopanoid biosynthesis and ethanol tolerance of Zymomonas mobilis. FEMS Microbiol. Lett. 79, 347–350 (1991).

    CAS  Google Scholar 

  84. 84

    Schmerk, C. L., Bernards, M. A. & Valvano, M. A. Hopanoid production is required for low-pH tolerance, antimicrobial resistance, and motility in Burkholderia cenocepacia. J. Bacteriol. 193, 6712–6723 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. 85

    Seipke, R. F. & Loria, R. Hopanoids are not essential for growth of Streptomyces scabies 87–22. J. Bacteriol. 191, 5216–5223 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  86. 86

    Kulkarni, G. & Busset, N. et al. Specific hopanoid classes differentially affect free-living and symbiotic states of Bradyrhizobium diazoefficiens. mBio 6, e01251-15 (2015). This work identifies extended hopanoids as the most important hopanoid structural class for stress resistance and symbiosis of Bradyrhizobium diazoefficiens.

    PubMed  PubMed Central  Google Scholar 

  87. 87

    Welander, P. V. et al. Hopanoids play a role in membrane integrity and pH homeostasis in Rhodopseudomonas palustris TIE-1. J. Bacteriol. 191, 6145–6156 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  88. 88

    Poralla, K., Härtner, T. & Kannenberg, E. Effect of temperature and pH on the hopanoid content of Bacillus acidocaldarius. FEMS Microbiol. Lett. 23, 253–256 (1984).

    CAS  Google Scholar 

  89. 89

    Kulkarni, G., Wu, C.-H. & Newman, D. K. The general stress response factor EcfG regulates expression of the C-2 hopanoid methylase HpnP in Rhodopseudomonas palustris TIE-1. J. Bacteriol. 195, 2490–2498 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. 90

    Ricci, J. N., Morton, R., Kulkarni, G., Summers, M. L. & Newman, D. K. Hopanoids play a role in stress tolerance and nutrient storage in the cyanobacterium Nostoc punctiforme. Geobiology 15, 173–183 (2016).

    PubMed  Google Scholar 

  91. 91

    Garby, T. J. et al. Lack of methylated hopanoids renders the cyanobacterium Nostoc punctiforme sensitive to osmotic and pH Stress. Appl. Environ. Microbiol. 83, e00777-17 (2017).

    PubMed  PubMed Central  Google Scholar 

  92. 92

    Poralla, K., Muth, G. & Hartner, T. Hopanoids are formed during transition from substrate to aerial hyphae in Streptomyces coelicolor A3(2). FEMS Microbiol. Lett. 189, 93–95 (2000).

    CAS  PubMed  Google Scholar 

  93. 93

    Bosak, T., Losick, R. M. & Pearson, A. A polycyclic terpenoid that alleviates oxidative stress. Proc. Natl Acad. Sci. USA 105, 6725–6729 (2008).

    CAS  PubMed  Google Scholar 

  94. 94

    Caron, B., Mark, A. E. & Poger, D. Some like it hot: the effect of sterols and hopanoids on lipid ordering at high temperature. J. Phys. Chem. Lett. 5, 3953–3957 (2014).

    CAS  PubMed  Google Scholar 

  95. 95

    Lopez, D. & Koch, G. Exploring functional membrane microdomains in bacteria: an overview. Curr. Opin. Microbiol. 36, 76–84 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. 96

    Vilcheze, C., Llopiz, P. & Neunlist, S. Prokaryotic triterpenoids: new hopanoids from the nitrogen-fixing bacteria Azotobacter vinelandii. Beijerinckia indica and Beijerinckia mobilis. Microbiology 140, 2749–2753 (1994).

    CAS  Google Scholar 

  97. 97

    Rohmer, M., Bouvier-Nave, P. & Ourisson, G. Distribution of hopanoid triterpenes in prokaryotes. J. Gen. Microbiol. 130, 1137 (1984).

    CAS  Google Scholar 

  98. 98

    Kannenberg, E., Perzl, E. & Härtner, T. The occurrence of hopanoid lipids in Bradyrhizobium bacteria. FEMS Microbiol. Lett. 127, 255–261 (1995).

    CAS  Google Scholar 

  99. 99

    Rosa-Putra, S., Nalin, R., Domenach, A.-M. & Rohmer, M. Novel hopanoids from Frankia spp. and related soil bacteria: squalene cyclization and significance of geological biomarkers revisited. Eur. J. Biochem. 268, 4300 (2001).

    CAS  PubMed  Google Scholar 

  100. 100

    Hakoyama, T. et al. Host plant genome overcomes the lack of a bacterial gene for symbiotic nitrogen fixation. Nature 462, 514–517 (2009).

    CAS  PubMed  Google Scholar 

  101. 101

    Appleby, C. A. Leghemoglobin and rhizobium respiration. Annu. Rev. Plant Physiol. 35, 443–478 (1984).

    CAS  Google Scholar 

  102. 102

    Sabra, W., Zeng, A. P. & Lünsdorf, H. Effect of oxygen on formation and structure of Azotobacter vinelandii alginate and its role in protecting nitrogenase. Appl. Environ. Microbiol. 66, 4037–4044 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. 103

    Parsons, R., Silvester, W., Harris, S., Gruijters, W. & Bullivant, S. Frankia vesicles provide inducible and absolute oxygen protection for nitrogenase. Plant Physiol. 83, 728–731 (1987).

    CAS  PubMed  PubMed Central  Google Scholar 

  104. 104

    Abeysekera, R. M., Newcomb, W., Silvester, W. B. & Torrey, J. G. A freeze-fracture electron microscopy study of Frankia in root nodules of Alnus incana grown at three oxygen tensions. Can. J. Microbiol. 36, 97–108 (1990).

    Google Scholar 

  105. 105

    Berry, A. M., Moreau, R. A. & Jones, A. D. Bacteriohopanetetrol: abundant lipid in Frankia cells and in nitrogen-fixing nodule tissue. Plant Physiol. 95, 111–115 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  106. 106

    Fries, L. Growth regulating effects of phenylacetic acid and phydroxy-phenylacetic acid on Fucus spiralis L. (Phaecophyceae. Fucales) in axenic culture. Phycology 16, 451–455 (1977).

    CAS  Google Scholar 

  107. 107

    Hammad, Y. et al. A possible role for phenyl acetic acid (PAA) on Alnus glutinosa nodulation by Frankia. Plant Soil 254, 193 (2003). This work finds that the auxinomimetic plant hormone PAA can be released from PAA-conjugated hopanoid lipids to regulate nodulation in Frankia symbioses.

    CAS  Google Scholar 

  108. 108

    Ricci, J. N. et al. Diverse capacity for 2-methylhopanoid production correlates with a specific ecological niche. ISME J. 8, 675–684 (2013). This work identifies a important association between 2Me-hopanoids and plant-associated environments.

    PubMed  PubMed Central  Google Scholar 

  109. 109

    Kloepper, J. W., Ryu, C. M. & Zhang, S. Induced systemic resistance and promotion of plant growth by Bacillus spp. Phytopathology 94, 1259–1266 (2004).

    CAS  PubMed  Google Scholar 

  110. 110

    Lemaire, B. et al. Biogeographical patterns of legume-nodulating Burkholderia spp.: from African Fynbos to continental scales. Appl. Environ. Microbiol. 82, 5099–5115 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. 111

    Moulin, L., James, E. K., Klonowska, A., Daria, S. M. & Simon, M. F. in Biological Nitrogen Fixation (ed. de Bruijn, F. J.) 177–190 (John Wiley & Sons, Hoboken, NJ, USA, 2015).

    Google Scholar 

  112. 112

    Sy, A. et al. Methylotrophic Methylobacterium bacteria nodulate and fix nitrogen in symbiosis with legumes. J. Bacteriol. 183, 214–220 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. 113

    Silipo, A. et al. Covalently linked hopanoid-lipid A improves outer-membrane resistance of a Bradyrhizobium symbiont of legumes. Nat. Commun. 5, 5106 (2014). This work describes a novel HoLA structure and provides the first evidence that hopanoids affect legume–rhizobia symbioses.

    CAS  PubMed  Google Scholar 

  114. 114

    Delamuta, J. R. M. et al. Polyphasic evidence supporting the reclassification of Bradyrhizobium japonicum group Ia strains as Bradyrhizobium diazoefficiens sp. nov. Int. J. Syst. Evol. Microbiol. 63, 3342–3351 (2013).

    CAS  PubMed  Google Scholar 

  115. 115

    Komaniecka, I. et al. Occurrence of an unusual hopanoid-containing lipid A among lipopolysaccharides from Bradyrhizobium species. J. Biol. Chem. 289, 35644–35655 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  116. 116

    Busset, N. et al. The very long chain fatty acid (C26:25OH) linked to the lipid A Is important for the fitness of the photosynthetic Bradyrhizobium strain ORS278 and the establishment of a successful symbiosis with Aeschynomene legumes. Front. Microbiol. 8, 1821 (2017).

    PubMed  PubMed Central  Google Scholar 

  117. 117

    Shevchenko, A., Schwille, P. & Simons, K. Yeast lipids can phase-separate into micrometer-scale membrane domains. J. Biol. Chem. 285, 30224–30232 (2010).

    PubMed  PubMed Central  Google Scholar 

  118. 118

    Boyd, E. S., Hamilton, T. L., Wang, J., He, L. & Zhang, C. L. The role of tetraether lipid composition in the adaptation of thermophilic archaea to acidity. Front. Microbiol. 4, 62 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. 119

    Cooper, J. E. Early interactions between legumes and rhizobia: disclosing complexity in a molecular dialogue. J. Appl. Microbiol. 103, 1355–1365 (2007).

    CAS  PubMed  Google Scholar 

  120. 120

    D'Haeze, W. et al. Reactive oxygen species and ethylene play a positive role in lateral root base nodulation of a semiaquatic legume. Proc. Natl Acad. Sci. USA 100, 11789–11794 (2003).

    CAS  PubMed  Google Scholar 

  121. 121

    Pauly, N. et al. Reactive oxygen and nitrogen species and glutathione: key players in the legume-Rhizobium symbiosis. J. Exp. Bot. 57, 1769–1776 (2006).

    CAS  PubMed  Google Scholar 

  122. 122

    Pierre, O. et al. Peribacteroid space acidification: a marker of mature bacteroid functioning in Medicago truncatula nodules. Plant Cell Environ. 36, 2059–2070 (2013).

    CAS  PubMed  Google Scholar 

  123. 123

    Miller, K. J. & Wood, J. M. Osmoadaptation by rhizosphere bacteria. Annu. Rev. Microbiol. 50, 01–36 (1996).

    Google Scholar 

  124. 124

    Czernic, P. et al. Convergent evolution of endosymbiont differentiation in Dalbergoid and IRLC legumes mediated by nodule-specific cysteine-rich peptides. Plant Physiol. 169, 1254–1265 (2015).

    PubMed  PubMed Central  Google Scholar 

  125. 125

    Haag, A. F. et al. Molecular insights into bacteroid development during Rhizobium-legume symbiosis. FEMS Microbiol. Rev. 37, 364–383 (2013).

    CAS  PubMed  Google Scholar 

  126. 126

    Kondorosi, E., Mergaert, P. & Kereszt, A. A paradigm for endosymbiotic life: cell differentiation of Rhizobium bacteria provoked by host plant factors. Annu. Rev. Microbiol. 67, 611–628 (2013).

    CAS  PubMed  Google Scholar 

  127. 127

    Sprent, J., Ardley, J. & James, E. Biogeography of nodulated legumes and their nitrogen fixing symbionts. New Phytol. 215, 40–56 (2017).

    CAS  PubMed  Google Scholar 

  128. 128

    Parker, M. A. The spread of Bradyrhizobium lineages across host legume clades: from Abarema to Zygia. Microb. Ecol. 69, 630–640 (2015).

    PubMed  Google Scholar 

  129. 129

    Fowler, D. et al. Effects of global change during the 21st century on the nitrogen cycle. Atmos. Chem. Phys. 15, 13849–13893 (2015).

    CAS  Google Scholar 

  130. 130

    Volkman, J. K. Sterols and other triterpenoids: source specificity and evolution of biosynthetic pathways. Org. Geochem. 36, 139 (2005).

    CAS  Google Scholar 

  131. 131

    Killops, S. D. & Killops, V. in Introduction to Organic Geochemistry 2nd edn Ch. 2 (Blackwell Publishing, Oxford, 2005).

    Google Scholar 

  132. 132

    Sohlenkamp, C. & Geiger, O. Bacterial membrane lipids: Diversity in structures and pathways. FEMS Microbiol. Rev. 40, 133–159 (2015).

    PubMed  Google Scholar 

  133. 133

    Raetz, C. R. & Whitfield, C. Lipopolysaccharide endotoxins. Annu. Rev. Biochem. 71, 635–700 (2002).

    CAS  Google Scholar 

  134. 134

    Silipo, A., De Castro, C., Lanzetta, R., Parrilli, M. & Molinaro, A. in Prokaryotic Cell Wall Compounds: Structure and Biochemistry (eds König, H., Claus, H. & Varma, A.) 133–153 (Springer, Berlin and Heidelberg, Germany, 2010).

    Google Scholar 

  135. 135

    Molinaro, A. et al. Chemistry of lipid A: at the heart of innate immunity. Chemistry 21, 500–519 (2015).

    CAS  PubMed  Google Scholar 

  136. 136

    Róg, T., Pasenkiewicz-Gierula, M., Vattulainen, I. & Karttunen, M. Ordering effects of cholesterol and its analogues. Biochim. Biophys. Acta 1788, 97–121 (2009).

    PubMed  Google Scholar 

  137. 137

    Van Meer, G., Voelker, D. R. & Feigenson, G. W. Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 9, 112–124 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  138. 138

    Harriott, O. T., Khairallah, L. & Benson, D. R. Isolation and structure of the lipid envelopes from the nitrogen-fixing vesicles of Frankia sp. Strain CpI1. J. Bacteriol. 173, 2061–2067 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  139. 139

    Berry, A. M. et al. Hopanoid lipids compose the Frankia vesicle envelope, presumptive barrier of oxygen diffusion to nitrogenase. Proc. Natl Acad. Sci. USA 90, 6091–6094 (1993). This work identifies hopanoids as the primary component of vesicles in Frankia spp., suggesting that hopanoids limit oxygen diffusion during nitrogen fixation.

    CAS  PubMed  Google Scholar 

  140. 140

    Delgado-Baquerizo, M. et al. A global atlas of the dominant bacteria found in soil. Science 359, 320–325 (2018).

    CAS  PubMed  Google Scholar 

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The authors thank A. Session, P. Normand and the reviewers for constructive comments on the manuscript. We appreciate permission from D. Benson, A. Berry and J. Sáenz to reproduce images from their work. Grants from the Howard Hughes Medical Institute (HHMI; D.K.N.), National Aeronautics and Space Administration (NASA; NNX12AD93G, D.K.N.), the Jane Coffin Childs Memorial Fund (B.J.B.), the US National Institutes of Health (NIH; K99GM126141, B.J.B.), H2020- MSCA-ITN-2014-ETN TOLLerant (A.S.), Progetto Galileo G14-23 (A.S.), Mizutani Foundation for Glycoscience 2014 (A.M.) and the French National Research Agency (ANR-BugsInaCell-13-BSV7-0013) have sustained our research on this problem.

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B.J.B, E.G., A.S. and D.K.N. researched data for the article. B.J.B., E.G., A.M., A.S. and D.K.N. substantially contributed to the discussion of content. B.J.B., N.B., E.G., A.S. and D.K.N. wrote the article. All authors reviewed and edited the manuscript before submission.

Corresponding authors

Correspondence to Alba Silipo or Dianne K. Newman.

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PowerPoint slides



Pentacyclic lipids with C6 A-D rings and a C5 E ring, among which six core methyl groups are distributed. Hopanoids are also characterized by the formation of accessory groups at their C2, C3 and C30 positions.


Tetracyclic lipids with C6 A-C rings and a C5 D ring. The parent compounds for all sterols contain oxygen groups at the C3 position.


Molecules assembled from two or more C5 isoprene units that share the core formula (C5H8)n. They are also known as isoprenoids.


Molecules derived from assemblies of six isoprene units with the core formula (C5H8)6 or C30H48. They may be acyclic or cyclic.


An acyclic triterpene with an irregular (tail-to-tail) linkage between two 3-isoprene units; a parent molecule of cyclic triterpenoids.

Triterpenoid cyclase

A superfamily of enzymes that convert acyclic triterpenoids, including squalene and 2,3-oxidosqualene, into various cyclic products.

Oxidosqualene cyclases

(OSCs). A family of predominantly eukaryotic 3-β-hydroxytriterpene cyclases that transform 2,3-oxidosqualene into sterols and diverse other cyclic triterpenoids.

Squalene-hopene cyclases

A family of predominantly bacterial 3-deoxytriterpene cyclases that cyclize squalene to primarily form hopanoids.

C30 hopanoids

Short hopanoids containing no additional carbon atoms that are not derived from squalene.


A hopanoid-like compound with a C6 E-ring. In bacteria, it is made from E-ring expansion of the C30 hopanoid diploptene.


Legumes are flowering plants of the Fabaceae (previously known as Leguminosae) family, which includes economically important crops such as soybeans, common beans and peanuts.

C35 hopanoids

Extended hopanoids that contain ribose-derived hydrocarbon side chains at their C30 position.


(2Me-hopanoids). Hopanoids containing an accessory methyl group at the C2 position.


(3Me-hopanoids). Hopanoids containing an accessory methyl group at the C3 position.


Spherical vesicles surrounded by one or more lipid bilayers that can be produced from cellular membranes in vivo or synthetically by sonication or extrusion of lipids into aqueous solution.


(BHT). A common C35 hopanoid containing a tetra-hydroxylated C5 side chain.

Membrane fluidity

The rotational and diffusional freedom of movement of molecules within a membrane.

Lipid rafts

Membrane microdomains with high stability that are thought to recruit specific membrane-associated proteins to spatially regulate their functions.


(LPSs). Complex, heat-stable amphiphilic lipids that are the main component of the external leaflet of the outer membrane of Gram-negative bacteria.

Lipid A

The lipophilic moiety of lipopolysaccharides.


An environment with a low concentration of oxygen (usually less than 30% saturation).

Nitrogen fixation

The conversion of dinitrogen gas into fixed or bioavailable nitrogen sources such as ammonia.


A bacterial enzyme complex that performs the following reaction: N2 + 16ATP + 10H+ + 8e → 2NH4+ + H2 + 16ADP−Pi

Root nodule

A specialized root organ that is generated by most legume plants to house nitrogen-fixing symbionts and create a specialized microenvironment to support bacterial nitrogen fixation.


A paraphyletic group of nitrogen-fixing soil bacteria that can engage in symbioses with legumes.


An oxygen-carrying haem protein expressed in the root nodules of rhizobial host plants.


A genus of tropical legumes that is broadly distributed globally and is used for livestock grazing.

Hopanoid-lipid A

(HoLa). An extended hopanoid that is covalently attached to lipid A and appears to be unique to the Bradyrhizobiaceae.

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Belin, B., Busset, N., Giraud, E. et al. Hopanoid lipids: from membranes to plant–bacteria interactions. Nat Rev Microbiol 16, 304–315 (2018).

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