The Various Roles of Fatty Acids

Carla C C R de Carvalho, Maria José Caramujo, Carla C C R de Carvalho, Maria José Caramujo

Abstract

Lipids comprise a large group of chemically heterogeneous compounds. The majority have fatty acids (FA) as part of their structure, making these compounds suitable tools to examine processes raging from cellular to macroscopic levels of organization. Among the multiple roles of FA, they have structural functions as constituents of phospholipids which are the "building blocks" of cell membranes; as part of neutral lipids FA serve as storage materials in cells; and FA derivatives are involved in cell signalling. Studies on FA and their metabolism are important in numerous research fields, including biology, bacteriology, ecology, human nutrition and health. Specific FA and their ratios in cellular membranes may be used as biomarkers to enable the identification of organisms, to study adaptation of bacterial cells to toxic compounds and environmental conditions and to disclose food web connections. In this review, we discuss the various roles of FA in prokaryotes and eukaryotes and highlight the application of FA analysis to elucidate ecological mechanisms. We briefly describe FA synthesis; analyse the role of FA as modulators of cell membrane properties and FA ability to store and supply energy to cells; and inspect the role of polyunsaturated FA (PUFA) and the suitability of using FA as biomarkers of organisms.

Keywords: biomarkers; cellular membranes; fatty acid synthesis; glycerophospholipids; lipid bodies; lipidomics; membrane remodelling; omega-3 fatty acids; specialized lipids; storage lipids.

Conflict of interest statement

The authors declare no conflict of interest.

Figures

Figure 1
Figure 1
Accumulation of storage lipids in prokaryotes shown in Nile Red stained cells: PHA production in B. megaterium (a), C. necator (b) and S. aureus (c); TAG production in R. erythropolis (d); (de Carvalho and Caramujo, unpublished data).
Figure 2
Figure 2
(a) In temporary Mediterranean water ponds, Daphnia sp. feeds on bacteria, fungus and algae—algal material visible as green mass inside the gut; (b) FA of auto and heterotrophic origin are incorporated into the phospholipids of Daphnia, as shown by PLFA analysis (de Carvalho and Caramujo, unpublished data).

References

    1. Berg J.M., Tymoczko J.L., Stryer L. Section 22.5–Acetyl Coenzyme A carboxylase plays a key role in controlling fatty acid metabolism. In: Berg J.M., Tymoczko J.L., Stryer L., editors. Biochemistry. 5th ed. W. H. Freeman; New York, NY, USA: 2002.
    1. Leibundgut M., Maier T., Jenni S., Ban N. The multienzyme architecture of eukaryotic fatty acid synthases. Curr. Opin. Struct. Biol. 2008;18:714–725. doi: 10.1016/j.sbi.2008.09.008.
    1. Schweizer E., Hofmann J. Microbial type I fatty acid synthases (FAS): Major players in a network of cellular FAS systems. Microbiol. Mol. Biol. Rev. 2004;68:501–517. doi: 10.1128/MMBR.68.3.501-517.2004.
    1. White S.W., Zheng J., Zhang Y.M., Rock C.O. The structural biology of type II fatty acid biosynthesis. Annu. Rev. Biochem. 2005;74:791–831. doi: 10.1146/annurev.biochem.74.082803.133524.
    1. Harwood J.L. Fatty acid metabolism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1988;39:101–138. doi: 10.1146/annurev.pp.39.060188.000533.
    1. Harwood J.L. Recent advances in the biosynthesis of plant fatty acids. Biochim. Biophys. Acta (BBA) Lipids Lipid Metab. 1996;1301:7–56. doi: 10.1016/0005-2760(95)00242-1.
    1. Bloch K., Vance D. Control mechanisms in the synthesis of saturated fatty acids. Annu. Rev. Biochem. 1977;46:263–298. doi: 10.1146/annurev.bi.46.070177.001403.
    1. Kikuchi S., Rainwater D.L., Kolattukudy P.E. Purification and characterization of an unusually large fatty acid synthase from Mycobacterium tuberculosis var. bovis BCG. Arch. Biochem. Biophys. 1992;295:318–326. doi: 10.1016/0003-9861(92)90524-Z.
    1. Kolattukudy P.E., Fernandes N.D., Azad A.K., Fitzmaurice A.M., Sirakova T.D. Biochemistry and molecular genetics of cell-wall lipid biosynthesis in mycobacteria. Mol. Microbiol. 2003;24:263–270. doi: 10.1046/j.1365-2958.1997.3361705.x.
    1. Boucher Y. Lipids: Biosynthesis, function and evolution. In: Cavicchioli R., editor. Archaea. ASM Press; Washington, DC, USA: 2007. pp. 341–353.
    1. Jain S., Caforio A., Driessen A.J.M. Biosynthesis of archaeal membrane ether lipids. Front. Microbiol. 2014;5:641. doi: 10.3389/fmicb.2014.00641.
    1. Gattinger A., Schloter M., Munch J.C. Phospholipid etherlipid and phospholipid fatty acid fingerprints in selected euryarchaeotal monocultures for taxonomic profiling. FEMS Microbiol. Lett. 2002;213:133–139. doi: 10.1111/j.1574-6968.2002.tb11297.x.
    1. Pugh E.L., Wassef M.K., Kates M. Inhibition of fatty acid synthetase in Halobacterium cutirubrum and Escherichia coli by high salt concentrations. Can. J. Biochem. 1971;49:953–958. doi: 10.1139/o71-138.
    1. Pugh E.L., Kates M. Acylation of proteins of the archaebacteria Halobacterium cutirubrum and Methanobacterium thermoautotrophicum. Biochim. Biophys. Acta (BBA) Biomembr. 1994;1196:38–44. doi: 10.1016/0005-2736(94)90292-5.
    1. Lombard J., López-García P., Moreira D. An ACP-independent fatty acid synthesis pathway in Archaea: Implications for the origin of phospholipids. Mol. Biol. Evol. 2012;29:3261–3265. doi: 10.1093/molbev/mss160.
    1. Pereira S.L., Leonard A.E., Mukerji P. Recent advances in the study of fatty acid desaturases from animals and lower eukaryotes. Prostaglandins Leukot. Essent. Fat. Acids. 2003;68:97–106. doi: 10.1016/S0952-3278(02)00259-4.
    1. Leonard A.E., Pereira S.L., Sprecher H., Huang Y.S. Elongation of long-chain fatty acids. Prog. Lipid Res. 2004;43:36–54. doi: 10.1016/S0163-7827(03)00040-7.
    1. Cook H.W. Chapter 5 Fatty acid desaturation and chain elongation in eukaryotes. In: Vance D.E., Vance J.E., editors. New Comprehensive Biochemistry. Volume 31. Elsevier; Amsterdam, The Netherlands: 1996. pp. 129–152.
    1. Sperling P., Ternes P., Zank T.K., Heinz E. The evolution of desaturases. Prostaglandins Leukot. Essent. Fat. Acids. 2003;68:73–95. doi: 10.1016/S0952-3278(02)00258-2.
    1. Hashimoto K., Yoshizawa A.C., Saito K., Yamada T., Kanehisa M. The repertoire of desaturases for unsaturated fatty acid synthesis in 397 genomes. Genome Inform. 2006;17:173–183. doi: 10.11234/gi1990.17.173.
    1. Harwood J.L., Guschina I.A. The versatility of algae and their lipid metabolism. Biochimie. 2009;91:679–684. doi: 10.1016/j.biochi.2008.11.004.
    1. Monroig Ó., Tocher D.R., Navarro J.C. Biosynthesis of polyunsaturated fatty acids in marine invertebrates: Recent advances in molecular mechanisms. Mar. Drugs. 2013;11:3998–4018. doi: 10.3390/md11103998.
    1. Kabeya N., Fonseca M.M., Ferrier D.E.K., Navarro J.C., Bay L.K., Francis D.S., Tocher D.R., Castro L.F.C., Monroig Ó. Genes for de novo biosynthesis of omega-3 polyunsaturated fatty acids are widespread in animals. Sci. Adv. 2018;4:eaar6849. doi: 10.1126/sciadv.aar6849.
    1. Cook H.W., McMaster C.R. New Comprehensive Biochemistry. Volume 36. Elsevier; Amsterdam, The Netherlands: 2002. Chapter 7 Fatty acid desaturation and chain elongation in eukaryotes; pp. 181–204.
    1. Tocher D.R. Metabolism and functions of lipids and fatty acids in teleost fish. Rev. Fish. Sci. 2003;11:107–184. doi: 10.1080/713610925.
    1. Parrish C.C. Essential fatty acids in aquatic food webs. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 309–326.
    1. Castro L.F.C., Tocher D.R., Monroig O. Long-chain polyunsaturated fatty acid biosynthesis in chordates: Insights into the evolution of Fads and Elovl gene repertoire. Prog. Lipid Res. 2016;62:25–40. doi: 10.1016/j.plipres.2016.01.001.
    1. Bell M.V., Tocher D.R. Biosynthesis of polyunsaturated fatty acids in aquatic ecosystems: General pathways and new directions. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 211–236.
    1. Tocher D.R. Fatty acid requirements in ontogeny of marine and freshwater fish. Aquacult. Res. 2010;41:717–732. doi: 10.1111/j.1365-2109.2008.02150.x.
    1. Brenna J.T. Efficiency of conversion of α-linolenic acid to long chain n-3 fatty acids in man. Curr. Opin. Clin. Nutr. Metab. Care. 2002;5:127–132. doi: 10.1097/00075197-200203000-00002.
    1. Barceló-Coblijn G., Murphy E.J. Alpha-linolenic acid and its conversion to longer chain n−3 fatty acids: Benefits for human health and a role in maintaining tissue n−3 fatty acid levels. Prog. Lipid Res. 2009;48:355–374. doi: 10.1016/j.plipres.2009.07.002.
    1. Scott B.L., Bazan N.G. Membrane docosahexaenoate is supplied to the developing brain and retina by the liver. Proc. Natl. Acad. Sci. USA. 1989;86:2903–2907. doi: 10.1073/pnas.86.8.2903.
    1. Brossard N., Croset M., Normand S., Pousin J., Lecerf J., Laville M., Tayot J.L., Lagarde M. Human plasma albumin transports [13C]docosahexaenoic acid in two lipid forms to blood cells. J. Lipid Res. 1997;38:1571–1582.
    1. Rapoport S.I., Rao J.S., Igarashi M. Brain metabolism of nutritionally essential polyunsaturated fatty acids depends on both the diet and the liver. Prostaglandins Leukot. Essent. Fat. Acids. 2007;77:251–261. doi: 10.1016/j.plefa.2007.10.023.
    1. Kang J.X. The importance of omega-6/omega-3 fatty acid ratio in cell function. In: Simopoulos A.P., editor. Omega-6/Omega-3 essential Fatty Acid Ratio: The Scientific Evidence. Volume 92. Karger; Basel, Switzerland: 2003. pp. 23–36.
    1. Simopoulos A.P. The importance of the omega-6/omega-3 fatty acid ratio in cardiovascular disease and other chronic diseases. Exp. Biol. Med. 2008;233:674–688. doi: 10.3181/0711-MR-311.
    1. Simopoulos A.P. An increase in the omega-6/omega-3 fatty acid ratio increases the risk for obesity. Nutrients. 2016;8:128. doi: 10.3390/nu8030128.
    1. Weaver K.L., Ivester P., Seeds M., Case L.D., Arm J.P., Chilton F.H. Effect of dietary fatty acids on inflammatory gene expression in healthy humans. J. Biol. Chem. 2009;284:15400–15407. doi: 10.1074/jbc.M109.004861.
    1. Donahue S.M.A., Rifas-Shiman S.L., Gold D.R., Jouni Z.E., Gillman M.W., Oken E. Prenatal fatty acid status and child adiposity at age 3 y: Results from a US pregnancy cohort. Am. J. Clin. Nutri. 2011;93:780–788. doi: 10.3945/ajcn.110.005801.
    1. EFSA Panel on Dietetic Products, Nutrition, and Allergies (NDA) Scientific Opinion on Dietary Reference Values for fats, including saturated fatty acids, polyunsaturated fatty acids, monounsaturated fatty acids, trans fatty acids and cholesterol. EFSA J. 2010;8:1461. doi: 10.2903/j.efsa.2010.1461.
    1. Alberts B., Johnson A., Lewis J., Morgan D., Raff M., Roberts K., Walter P. Molecular Biology of the Cell. Garland Science; New York, NY, USA: Abingdon, UK: 2015. p. 1464.
    1. Hamerly T., Tripet B., Wurch L., Hettich R.L., Podar M., Bothner B., Copié V. Characterization of fatty acids in Crenarchaeota by GC-MS and NMR. Archaea. 2015;2015:9. doi: 10.1155/2015/472726.
    1. Koga Y. Thermal adaptation of the archaeal and bacterial lipid membranes. Archaea. 2012;2012:6. doi: 10.1155/2012/789652.
    1. Weijers J.W.H., Schouten S., Hopmans E.C., Geenevasen J.A.J., David O.R.P., Coleman J.M., Pancost R.D., Sinninghe Damsté J.S. Membrane lipids of mesophilic anaerobic bacteria thriving in peats have typical archaeal traits. Environ. Microbiol. 2006;8:648–657. doi: 10.1111/j.1462-2920.2005.00941.x.
    1. de Carvalho C.C.C.R., Fernandes P. Production of metabolites as bacterial responses to the marine environment. Mar. Drugs. 2010;8:705–727. doi: 10.3390/md8030705.
    1. de Carvalho C.C.C.R., Caramujo M.J. Carotenoids in aquatic ecosystems and aquaculture: A colorful business with implications for human health. Front. Mar. Sci. 2017;4:93. doi: 10.3389/fmars.2017.00093.
    1. Hazel J.R., Eugene Williams E. The role of alterations in membrane lipid composition in enabling physiological adaptation of organisms to their physical environment. Prog. Lipid Res. 1990;29:167–227. doi: 10.1016/0163-7827(90)90002-3.
    1. Konings W.N., Albers S.-V., Koning S., Driessen A.J.M. The cell membrane plays a crucial role in survival of bacteria and archaea in extreme environments. Antonie Van Leeuwenhoek. 2002;81:61–72. doi: 10.1023/A:1020573408652.
    1. Yazawa K. Production of eicosapentaenoic acid from marine bacteria. Lipids. 1996;31:S297–S300. doi: 10.1007/BF02637095.
    1. Okuyama H., Orikasa Y., Nishida T., Watanabe K., Morita N. Bacterial genes responsible for the biosynthesis of eicosapentaenoic and docosahexaenoic acids and their heterologous expression. Appl. Environ. Microbiol. 2007;73:665–670. doi: 10.1128/AEM.02270-06.
    1. Fang J., Barcelona M.J., Nogi Y., Kato C. Biochemical implications and geochemical significance of novel phospholipids of the extremely barophilic bacteria from the Marianas Trench at 11,000 m. Deep Sea Res. Part I Oceanogr. Res. Pap. 2000;47:1173–1182. doi: 10.1016/S0967-0637(99)00080-1.
    1. Yano Y., Nakayama A., Ishihara K., Saito H. Adaptive changes in membrane lipids of barophilic bacteria in response to changes in growth pressure. Appl. Environ. Microbiol. 1998;64:479–485.
    1. de Carvalho C.C.C.R., da Fonseca M.M.R. Preventing biofilm formation: Promoting cell separation with terpenes. FEMS Microbiol. Ecol. 2007;61:406–413. doi: 10.1111/j.1574-6941.2007.00352.x.
    1. Kaye J.Z., Baross J.A. Synchronous effects of temperature, hydrostatic pressure and salinity on growth, phospholipid profiles and protein patterns of four Halomonas species isolated from deep-sea hydrothermal-vent and sea surface environments. Appl. Environ. Microbiol. 2004;70:6220–6229. doi: 10.1128/AEM.70.10.6220-6229.2004.
    1. de Carvalho C.C.C.R., Wick L.Y., Heipieper H.J. Cell wall adaptations of planktonic and biofilm Rhodococcus erythropolis cells to growth on C5 to C16 n-alkane hydrocarbons. Appl. Microbiol. Biotechnol. 2009;82:311–320. doi: 10.1007/s00253-008-1809-3.
    1. Wirsen C.O., Jannasch H.W., Wakeham S.G., Canuel E.A. Membrane lipids of a psychrophilic and barophilic deep-sea bacterium. Curr. Microbiol. 1986;14:319–322. doi: 10.1007/BF01568697.
    1. Nichols D.S., Nichols P.D., Russell N.J., Davies N.W., McMeekin T.A. Polyunsaturated fatty acids in the psychrophilic bacterium Shewanella gelidimarina ACAM 456T: Molecular species analysis of major phospholipids and biosynthesis of eicosapentaenoic acid. Biochim. Biophys. Acta (BBA) Lipids Lipid Metab. 1997;1347:164–176. doi: 10.1016/S0005-2760(97)00068-4.
    1. Yoshida K., Hashimoto M., Hori R., Adachi T., Okuyama H., Orikasa Y., Nagamine T., Shimizu S., Ueno A., Morita N. Bacterial long-chain polyunsaturated fatty acids: Their biosynthetic genes, functions and practical use. Mar. Drugs. 2016;14:94. doi: 10.3390/md14050094.
    1. de Carvalho C.C.C.R., Marques M.P.C., Hachicho N., Heipieper H.J. Rapid adaptation of Rhodococcus erythropolis cells to salt stress by synthesizing polyunsaturated fatty acids. Appl. Microbiol. Biotechnol. 2014;98:5599–5606. doi: 10.1007/s00253-014-5549-2.
    1. de Carvalho C.C.C.R. Adaptation of Rhodococcus erythropolis cells for growth and bioremediation under extreme conditions. Res. Microbiol. 2012;163:125–136. doi: 10.1016/j.resmic.2011.11.003.
    1. Heipieper H.J., Fischer J. Bacterial solvent responses and tolerance: Cis-trans isomerization. In: Timmis K.N., editor. Handbook of Hydrocarbon and Lipid Microbiology. Springer; Berlin/Heidelberg, Germany: 2010. pp. 4203–4211.
    1. Eberlein C., Baumgarten T., Starke S., Heipieper H.J. Immediate response mechanisms of Gram-negative solvent-tolerant bacteria to cope with environmental stress: Cis-trans isomerization of unsaturated fatty acids and outer membrane vesicle secretion. Appl. Microbiol. Biotechnol. 2018;102:2583–2593. doi: 10.1007/s00253-018-8832-9.
    1. Saunders L.P., Sen S., Wilkinson B.J., Gatto C. Insights into the mechanism of homeoviscous adaptation to low temperature in branched-chain fatty acid-containing bacteria through modeling FabH kinetics from the foodborne pathogen Listeria monocytogenes. Front. Microbiol. 2016;7:1386. doi: 10.3389/fmicb.2016.01386.
    1. Gonçalves F.D.A., de Carvalho C.C.C.R. Phenotypic modifications in Staphylococcus aureus cells exposed to high concentrations of vancomycin and teicoplanin. Front. Microbiol. 2016;7:13. doi: 10.3389/fmicb.2016.00013.
    1. Unell M., Kabelitz N., Jansson J.K., Heipieper H.J. Adaptation of the psychrotroph Arthrobacter chlorophenolicus A6 to growth temperature and the presence of phenols by changes in the anteiso/iso ratio of branched fatty acids. FEMS Microbiol. Lett. 2006;266:138–143. doi: 10.1111/j.1574-6968.2006.00502.x.
    1. 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. 2000;5:717–727. doi: 10.1016/S1097-2765(00)80250-6.
    1. Monteoliva-Sanchez M., Ramos-Cormenzana A., Russell N.J. The effect of salinity and compatible solutes on the biosynthesis of cyclopropane fatty acids in Pseudomonas halosaccharolytica. Microbiology. 1993;139:1877–1884. doi: 10.1099/00221287-139-8-1877.
    1. Álvarez-Ordóñez A., Fernández A., López M., Arenas R., Bernardo A. Modifications in membrane fatty acid composition of Salmonella typhimurium in response to growth conditions and their effect on heat resistance. Int. J. Food Microbiol. 2008;123:212–219. doi: 10.1016/j.ijfoodmicro.2008.01.015.
    1. Vinçon-Laugier A., Cravo-Laureau C., Mitteau I., Grossi V. Temperature-dependent alkyl glycerol ether lipid composition of mesophilic and thermophilic sulfate-reducing bacteria. Front. Microbiol. 2017;8:1532. doi: 10.3389/fmicb.2017.01532.
    1. Ernst R., Ejsing C.S., Antonny B. Homeoviscous adaptation and the regulation of membrane lipids. J. Mol. Biol. 2016;428:4776–4791. doi: 10.1016/j.jmb.2016.08.013.
    1. Melchior D.L. Lipid phase transitions and regulation of membrane fluidity in prokaryotes. In: Bronner F., Kleinteller A., editors. Current Topics in Membranes and Transport. Volume 17. Academic Press; Cambridge, MA, USA: 1982. pp. 263–316.
    1. Siliakus M.F., van der Oost J., Kengen S.W.M. Adaptations of archaeal and bacterial membranes to variations in temperature, pH and pressure. Extremophiles. 2017;21:651–670. doi: 10.1007/s00792-017-0939-x.
    1. Eze M.O. Phase transitions in phospholipid bilayers: Lateral phase separations play vital roles in biomembranes. Biochem. Educ. 1991;19:204–208. doi: 10.1016/0307-4412(91)90103-F.
    1. Edidin M. Rotational and translational diffusion in membranes. Annu. Rev. Biophys. Bioeng. 1974;3:179–201. doi: 10.1146/annurev.bb.03.060174.001143.
    1. Wan C., Kiessling V., Tamm L.K. Coupling of cholesterol-rich lipid phases in asymmetric bilayers. Biochemistry. 2008;47:2190–2198. doi: 10.1021/bi7021552.
    1. Mouritsen O.G. Phase transitions in biological membranes. Ann. N. Y. Acad. Sci. 1987;491:166–169. doi: 10.1111/j.1749-6632.1987.tb30051.x.
    1. Phadtare S. Recent developments in bacterial cold-shock response. Curr. Issues Mol. Biol. 2004;6:125–136.
    1. Diomandé S., Nguyen-The C., Guinebretière M.H., Broussolle V., Brillard J. Role of fatty acids in Bacillus environmental adaptation. Front. Microbiol. 2015;6:813. doi: 10.3389/fmicb.2015.00813.
    1. Aguilar P.S., Cronan J.E., de Mendoza D. A Bacillus subtilis gene induced by cold shock encodes a membrane phospholipid desaturase. J. Bacteriol. 1998;180:2194–2200.
    1. Klein W., Weber M.H.W., Marahiel M.A. Cold shock response of Bacillus subtilis: Isoleucine-dependent switch in the fatty acid branching pattern for membrane adaptation to low temperatures. J. Bacteriol. 1999;181:5341–5349.
    1. Graumann P.L., Marahiel M.A. Cold shock response in Bacillus subtilis. J. Mol. Microbiol. Biotechnol. 1999;1:203–209.
    1. Yamanaka K. Cold shock response in Escherichia coli. J. Mol. Microbiol. Biotechnol. 1999;1:193–202.
    1. Annous B.A., Becker L.A., Bayles D.O., Labeda D.P., Wilkinson B.J. Critical role of anteiso-C15:0 fatty acid in the growth of Listeria monocytogenes at low temperatures. Appl. Environ. Microbiol. 1997;63:3887–3894.
    1. Kaneda T. Iso- and anteiso-fatty acids in bacteria: Biosynthesis, function and taxonomic significance. Microbiol. Rev. 1991;55:288–302.
    1. Suutari M., Laakso S. Microbial fatty acids and thermal adaptation. Crit. Rev. Microbiol. 1994;20:285–328. doi: 10.3109/10408419409113560.
    1. Mantsch H.H., Madec C., Lewis R.N.A.H., McElhaney R.N. Thermotropic phase behavior of model membranes composed of phosphatidylcholines containing dl-methyl anteisobranched fatty acids. 2. An infrared spectroscopy study. Biochemistry. 1987;26:4045–4049. doi: 10.1021/bi00387a045.
    1. Diomandé S.E., Guinebretière M.H., De Sarrau B., Nguyen-the C., Broussolle V., Brillard J. Fatty acid profiles and desaturase-encoding genes are different in thermo- and psychrotolerant strains of the Bacillus cereus Group. BMC Res. Notes. 2015;8:329. doi: 10.1186/s13104-015-1288-4.
    1. Suutari M., Laakso S. Unsaturated and branched chain-fatty acids in temperature adaptation of Bacillus subtilis and Bacillus megaterium. Biochim. Biophys. Acta (BBA) Lipids Lipid Metab. 1992;1126:119–124. doi: 10.1016/0005-2760(92)90281-Y.
    1. Heipieper H.J., Meinhardt F., Segura A. The cis-trans isomerase of unsaturated fatty acids in Pseudomonas and Vibrio: Biochemistry, molecular biology and physiological function of a unique stress adaptive mechanism. FEMS Microbiol. Lett. 2003;229:1–7. doi: 10.1016/S0378-1097(03)00792-4.
    1. Heipieper H.J., Fischer J., Meinhardt F. Cis–trans isomerase of unsaturated fatty acids: An immediate bacterial adaptive mechanism to cope with emerging membrane perturbation caused by toxic hydrocarbons. In: Timmis K.N., editor. Handbook of Hydrocarbon and Lipid Microbiology. Springer; Berlin/Heidelberg, Germany: 2010. pp. 1605–1614.
    1. Okuyama H., Okajima N., Sasaki S., Higashi S., Murata N. The cis/trans isomerization of the double bond of a fatty acid as a strategy for adaptation to changes in ambient temperature in the psychrophilic bacterium, Vibrio sp. strain ABE-1. Biochim.e Biophys. Acta (BBA) Lipids Lipid Metab. 1991;1084:13–20. doi: 10.1016/0005-2760(91)90049-N.
    1. Junker F., Ramos J.L. Involvement of the cis/trans isomerase Cti in solvent resistance of Pseudomonas putida DOT-T1E. J. Bacteriol. 1999;181:5693–5700.
    1. Pinkart H.C., White D.C. Phospholipid biosynthesis and solvent tolerance in Pseudomonas putida strains. J. Bacteriol. 1997;179:4219–4226. doi: 10.1128/jb.179.13.4219-4226.1997.
    1. Atashgahi S., Sánchez-Andrea I., Heipieper H.J., van der Meer J.R., Stams A.J.M., Smidt H. Prospects for harnessing biocide resistance for bioremediation and detoxification. Science. 2018;360:743–746. doi: 10.1126/science.aar3778.
    1. Kadurugamuwa J.L., Beveridge T.J. Virulence factors are released from Pseudomonas aeruginosa in association with membrane vesicles during normal growth and exposure to gentamicin: A novel mechanism of enzyme secretion. J. Bacteriol. 1995;177:3998–4008. doi: 10.1128/jb.177.14.3998-4008.1995.
    1. Baumgarten T., Sperling S., Seifert J., von Bergen M., Steiniger F., Wick L.Y., Heipieper H.J. Membrane vesicle formation as a multiple-stress response mechanism enhances Pseudomonas putida DOT-T1E cell surface hydrophobicity and biofilm formation. Appl. Environ. Microbiol. 2012;78:6217–6224. doi: 10.1128/AEM.01525-12.
    1. DeLong E.F., Yayanos A.A. Biochemical function and ecological significance of novel bacterial lipids in deep-sea procaryotes. Appl. Environ. Microbiol. 1986;51:730–737.
    1. Nichols D.S. Prokaryotes and the input of polyunsaturated fatty acids to the marine food web. FEMS Microbiol. Lett. 2003;219:1–7. doi: 10.1016/S0378-1097(02)01200-4.
    1. Marsh D. Handbook of Lipid Bilayers. CRC Press; Boca Raton, FL, USA: 2013. p. 1174.
    1. Allen E.E., Facciotti D., Bartlett D.H. Monounsaturated but not polyunsaturated fatty acids are required for growth of the deep-sea bacterium Photobacterium profundum SS9 at high pressure and low temperature. Appl. Environ. Microbiol. 1999;65:1710–1720.
    1. Presentato A., Cappelletti M., Sansone A., Ferreri C., Piacenza E., Demeter M.A., Crognale S., Petruccioli M., Milazzo G., Fedi S., et al. Aerobic growth of Rhodococcus aetherivorans BCP1 using selected naphthenic acids as the sole carbon and energy sources. Front. Microbiol. 2018;9:672. doi: 10.3389/fmicb.2018.00672.
    1. Marsh D. Structural and thermodynamic determinants of chain-melting transition temperatures for phospholipid and glycolipids membranes. Biochim. Biophys. Acta (BBA)-Biomembr. 2010;1798:40–51. doi: 10.1016/j.bbamem.2009.10.010.
    1. Poger D., Caron B., Mark A.E. Effect of methyl-branched fatty acids on the structure of lipid bilayers. J. Phys. Chem. B. 2014;118:13838–13848. doi: 10.1021/jp503910r.
    1. Chang Y.Y., Cronan J.E. Membrane cyclopropane fatty acid content is a major factor in acid resistance of Escherichia coli. Mol. Microbiol. 1999;33:249–259. doi: 10.1046/j.1365-2958.1999.01456.x.
    1. Kim B.H., Kim S., Kim H.G., Lee J., Lee I.S., Park Y.K. The formation of cyclopropane fatty acids in Salmonella enterica serovar Typhimurium. Microbiology. 2005;151:209–218. doi: 10.1099/mic.0.27265-0.
    1. Grogan D.W., Cronan J.E., Jr. Cyclopropane ring formation in membrane lipids of bacteria. Microbiol. Mol. Biol. Rev. 1997;61:429–441.
    1. Desbois A.P., Smith V.J. Antibacterial free fatty acids: Activities, mechanisms of action and biotechnological potential. Appl. Microbiol. Biotechnol. 2010;85:1629–1642. doi: 10.1007/s00253-009-2355-3.
    1. Yoon B., Jackman J., Valle-González E., Cho N.J. Antibacterial free fatty acids and monoglycerides: Biological activities, experimental testing and therapeutic applications. Int. J. Mo. Sci. 2018;19:1114. doi: 10.3390/ijms19041114.
    1. Harayama T., Riezman H. Understanding the diversity of membrane lipid composition. Nat. Rev. Mol. Cell Biol. 2018;19:281. doi: 10.1038/nrm.2017.138.
    1. Simons K., Ikonen E. Functional rafts in cell membranes. Nature. 1997;387:569. doi: 10.1038/42408.
    1. Van Meer G., Voelker D.R., Feigenson G.W. Membrane lipids: Where they are and how they behave. Nat. Rev. Mol. Cell Biol. 2008;9:112. doi: 10.1038/nrm2330.
    1. Ibarguren M., López D.J., Escribá P.V. The effect of natural and synthetic fatty acids on membrane structure, microdomain organization, cellular functions and human health. Biochim. Biophys. Acta (BBA) Biomembr. 2014;1838:1518–1528. doi: 10.1016/j.bbamem.2013.12.021.
    1. Guschina I.A., Harwood J.L. Mechanisms of temperature adaptation in poikilotherms. FEBS Lett. 2006;580:5477–5483. doi: 10.1016/j.febslet.2006.06.066.
    1. Klose C., Surma M.A., Gerl M.J., Meyenhofer F., Shevchenko A., Simons K. Flexibility of a eukaryotic lipidome–insights from yeast lipidomics. PLoS ONE. 2012;7:e35063. doi: 10.1371/journal.pone.0035063.
    1. Suutari M., Liukkonen K., Laakso S. Temperature adaptation in yeasts: The role of fatty acids. Microbiology. 1990;136:1469–1474. doi: 10.1099/00221287-136-8-1469.
    1. Pearcy R.W. Effect of growth temperature on the fatty acid composition of the leaf lipids in Atriplex lentiformis (Torr.) Wats. Plant Physiol. 1978;61:484–486. doi: 10.1104/pp.61.4.484.
    1. Berry J., Bjorkman O. Photosynthetic response and adaptation to temperature in higher plants. Annu. Rev. Plant Physiol. 1980;31:491–543. doi: 10.1146/annurev.pp.31.060180.002423.
    1. Williams J.P., Khan M.U., Mitchell K., Johnson G. The effect of temperature on the level and biosynthesis of unsaturated fatty acids in diacylglycerols of Brassica napus leaves. Plant Physiol. 1988;87:904–910. doi: 10.1104/pp.87.4.904.
    1. Harwood J.L. Environmental factors which can alter lipid metabolism. Prog. Lipid Res. 1994;33:193–202. doi: 10.1016/0163-7827(94)90022-1.
    1. Falcone D.L., Ogas J.P., Somerville C.R. Regulation of membrane fatty acid composition by temperature in mutants of Arabidopsis with alterations in membrane lipid composition. BMC Plant Biol. 2004;4:17. doi: 10.1186/1471-2229-4-17.
    1. Mizusawa N., Wada H. The role of lipids in photosystem II. Biochim. Biophys. Acta (BBA) Bioenerg. 2012;1817:194–208. doi: 10.1016/j.bbabio.2011.04.008.
    1. Farkas T., Fodor E., Kitajka K., Halver J.E. Response of fish membranes to environmental temperature. Aquacult. Res. 2001;32:645–655. doi: 10.1046/j.1365-2109.2001.00600.x.
    1. Hachicho N., Reithel S., Miltner A., Heipieper H.J., Küster E., Luckenbach T. Body mass parameters, lipid profiles and protein contents of zebrafish embryos and effects of 2,4-dinitrophenol exposure. PLoS ONE. 2015;10:e0134755. doi: 10.1371/journal.pone.0134755.
    1. Gurr M.I., James A.T. Phospholipids. In: Gurr M.I., James A.T., editors. Lipid Biochemistry: An Introduction. Springer; Dordrecht, The Netherlands: 1980. pp. 129–154.
    1. Gurr M.I., James A.T. Fatty acids. In: Gurr M.I., James A.T., editors. Lipid Biochemistry: An Introduction. Springer; Dordrecht, The Netherlands: 1980. pp. 18–89.
    1. Huster D., Jin A.J., Arnold K., Gawrisch K. Water permeability of polyunsaturated lipid membranes measured by 17O NMR. Biophys. J. 1997;73:855–864. doi: 10.1016/S0006-3495(97)78118-9.
    1. Koenig B.W., Strey H.H., Gawrisch K. Membrane lateral compressibility determined by NMR and X-ray diffraction: Effect of acyl chain polyunsaturation. Biophys. J. 1997;73:1954–1966. doi: 10.1016/S0006-3495(97)78226-2.
    1. Smaby J.M., Momsen M.M., Brockman H.L., Brown R.E. Phosphatidylcholine acyl unsaturation modulates the decrease in interfacial elasticity induced by cholesterol. Biophys. J. 1997;73:1492–1505. doi: 10.1016/S0006-3495(97)78181-5.
    1. Williams E.E., Jenski L.J., Stillwell W. Docosahexaenoic acid (DHA) alters the structure and composition of membranous vesicles exfoliated from the surface of a murine leukemia cell line. Biochim. Biophys. Acta (BBA) Biomembr. 1998;1371:351–362. doi: 10.1016/S0005-2736(98)00039-X.
    1. Calder P.C. Immunomodulation by omega-3 fatty acids. Prostaglandins Leukot. Essent. Fat. Acids. 2007;77:327–335. doi: 10.1016/j.plefa.2007.10.015.
    1. Stillwell W., Wassall S.R. Docosahexaenoic acid: Membrane properties of a unique fatty acid. Chem. Phys. Lipids. 2003;126:1–27. doi: 10.1016/S0009-3084(03)00101-4.
    1. Armstrong V.T., Brzustowicz M.R., Wassall S.R., Jenski L.J., Stillwell W. Rapid flip-flop in polyunsaturated (docosahexaenoate) phospholipid membranes. Arch. Biochem. Biophys. 2003;414:74–82. doi: 10.1016/S0003-9861(03)00159-0.
    1. Ma D.W.L., Seo J., Switzer K.C., Fan Y.Y., McMurray D.N., Lupton J.R., Chapkin R.S. n−3 PUFA and membrane microdomains: A new frontier in bioactive lipid research. J. Nutri. Biochem. 2004;15:700–706. doi: 10.1016/j.jnutbio.2004.08.002.
    1. Wassall S.R., Stillwell W. Polyunsaturated fatty acid-cholesterol interactions: Domain formation in membranes. Biochim. Biophys. Acta (BBA) Biomembr. 2009;1788:24–32. doi: 10.1016/j.bbamem.2008.10.011.
    1. Stulnig T.M., Huber J., Leitinger N., Imre E.M., Angelisová P., Nowotny P., Waldhäusl W. Polyunsaturated eicosapentaenoic acid displaces proteins from membrane rafts by altering raft lipid composition. J. Biol. Chem. 2001;276:37335–37340. doi: 10.1074/jbc.M106193200.
    1. Chen W., Jump D.B., Esselman W.J., Busik J.V. Inhibition of cytokine signaling in human retinal endothelial cells through modification of Caveolae/Lipid Rafts by docosahexaenoic acid. Investig. Ophthalmol. Vis. Sci. 2007;48:18–26. doi: 10.1167/iovs.06-0619.
    1. Ye S., Tan L., Ma J., Shi Q., Li J. Polyunsaturated docosahexaenoic acid suppresses oxidative stress induced endothelial cell calcium influx by altering lipid composition in membrane caveolar rafts. Prostaglandins Leukot. Essent. Fat. Acids. 2010;83:37–43. doi: 10.1016/j.plefa.2010.02.002.
    1. Schumann J., Leichtle A., Thiery J., Fuhrmann H. Fatty acid and peptide profiles in plasma membrane and membrane rafts of PUFA supplemented RAW264.7 macrophages. PLoS ONE. 2011;6:e24066. doi: 10.1371/journal.pone.0024066.
    1. Salmon A., Dodd S.W., Williams G.D., Beach J.M., Brown M.F. Configurational statistics of acyl chains in polyunsaturated lipid bilayers from deuterium NMR. J. Am. Chem. Soc. 1987;109:2600–2609. doi: 10.1021/ja00243a010.
    1. Tanguy E., Kassas N., Vitale N. Protein-phospholipid interaction motifs: A focus on phosphatidic acid. Biomolecules. 2018;8:20. doi: 10.3390/biom8020020.
    1. Pinot M., Vanni S., Pagnotta S., Lacas-Gervais S., Payet L.A., Ferreira T., Gautier R., Goud B., Antonny B., Barelli H. Polyunsaturated phospholipids facilitate membrane deformation and fission by endocytic proteins. Science. 2014;345:693. doi: 10.1126/science.1255288.
    1. Barelli H., Antonny B. Lipid unsaturation and organelle dynamics. Curr. Opin. Cell Biol. 2016;41:25–32. doi: 10.1016/j.ceb.2016.03.012.
    1. Rawicz W., Olbrich K.C., McIntosh T., Needham D., Evans E. Effect of chain length and unsaturation on elasticity of lipid bilayers. Biophys. J. 2000;79:328–339. doi: 10.1016/S0006-3495(00)76295-3.
    1. Hashidate-Yoshida T., Harayama T., Hishikawa D., Morimoto R., Hamano F., Tokuoka S.M., Eto M., Tamura-Nakano M., Yanobu-Takanashi R., Mukumoto Y., et al. Fatty acid remodeling by LPCAT3 enriches arachidonate in phospholipid membranes and regulates triglyceride transport. eLife. 2015;4:e06328. doi: 10.7554/eLife.06328.
    1. Rustan A.C., Drevon C.A. eLS. John Wiley & Sons Ltd.; Chischester, UK: 2005. Fatty acids: Structures and properties.
    1. Weijers R.N.M. Lipid composition of cell membranes and its relevance in Type 2 diabetes Mellitus. Curr. Diabetes Rev. 2012;8:390–400. doi: 10.2174/157339912802083531.
    1. Koehrer P., Saab S., Berdeaux O., Isaïco R., Grégoire S., Cabaret S., Bron A.M., Creuzot-Garcher C.P., Bretillon L., Acar N. Erythrocyte phospholipid and polyunsaturated fatty acid composition in diabetic retinopathy. PLoS ONE. 2014;9:e106912. doi: 10.1371/journal.pone.0106912.
    1. Weijers R.N.M. Membrane flexibility, free fatty acids and the onset of vascular and neurological lesions in type 2 diabetes. J. Diabetes Metab. Disord. 2016;15:13. doi: 10.1186/s40200-016-0235-9.
    1. Olukoshi E.R., Packter N.M. Importance of stored triacylglycerols in Streptomyces: Possible carbon source for antibiotics. Microbiology. 1994;140:931–943. doi: 10.1099/00221287-140-4-931.
    1. Alvarez H., Steinbüchel A. Triacylglycerols in prokaryotic microorganisms. Appl. Microbiol. Biotechnol. 2002;60:367–376. doi: 10.1007/s00253-002-1135-0.
    1. Lemoigne M. Produit de deshydratation et de polymerisation de l’acide β-oxybutyrique. Bull. Soc. Chim. Biol. 1926;8:770–782.
    1. Koller M. Advances in polyhydroxyalkanoate (PHA) production. Bioengineering. 2017;4:88. doi: 10.3390/bioengineering4040088.
    1. Anderson A.J., Dawes E.A. Occurrence, metabolism, metabolic role and industrial uses of bacterial polyhydroxyalkanoates. Microbiol. Rev. 1990;54:450–472.
    1. Bresan S., Sznajder A., Hauf W., Forchhammer K., Pfeiffer D., Jendrossek D. Polyhydroxyalkanoate (PHA) granules have no phospholipids. Sci. Rep. 2016;6:26612. doi: 10.1038/srep26612.
    1. Cavalheiro J.M.B.T., Almeida M.C.M.D., da Fonseca M.M.R., de Carvalho C.C.C.R. Adaptation of Cupriavidus necator to conditions favoring polyhydroxyalkanoate production. J. Biotechnol. 2013;164:309–317. doi: 10.1016/j.jbiotec.2013.01.009.
    1. Koller M. Production of polyhydroxyalkanoate (PHA) biopolyesters by extremophiles. MOJ Polym. Sci. 2017;1:69–85. doi: 10.15406/mojps.2017.01.00011.
    1. Obruca S., Sedlacek P., Mravec F., Samek O., Marova I. Evaluation of 3-hydroxybutyrate as an enzyme-protective agent against heating and oxidative damage and its potential role in stress response of poly(3-hydroxybutyrate) accumulating cells. Appl. Microbiol. Biotechnol. 2016;100:1365–1376. doi: 10.1007/s00253-015-7162-4.
    1. Wältermann M., Hinz A., Robenek H., Troyer D., Reichelt R., Malkus U., Galla H.J., Kalscheuer R., Stöveken T., Von Landenberg P., et al. Mechanism of lipid-body formation in prokaryotes: How bacteria fatten up. Mol. Microbiol. 2005;55:750–763. doi: 10.1111/j.1365-2958.2004.04441.x.
    1. Barney B.M., Wahlen B.D., Garner E., Wei J., Seefeldt L.C. Differences in substrate specificities of five bacterial wax ester synthases. Appl. Environ. Microbiol. 2012;78:5734–5745. doi: 10.1128/AEM.00534-12.
    1. Wältermann M., Steinbüchel A. Neutral lipid bodies in prokaryotes: Recent insights into structure, formation and relationship to eukaryotic lipid depots. J. Bacteriol. 2005;187:3607–3619. doi: 10.1128/JB.187.11.3607-3619.2005.
    1. Sirakova T.D., Deb C., Daniel J., Singh H.D., Maamar H., Dubey V.S., Kolattukudy P.E. Wax ester synthesis is required for Mycobacterium tuberculosis to enter in vitro dormancy. PLoS ONE. 2012;7:e51641. doi: 10.1371/journal.pone.0051641.
    1. 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. 2011;7:e1002093. doi: 10.1371/journal.ppat.1002093.
    1. Daniel J., Deb C., Dubey V.S., Sirakova T.D., Abomoelak B., Morbidoni H.R., Kolattukudy P.E. 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. 2004;186:5017–5030. doi: 10.1128/JB.186.15.5017-5030.2004.
    1. de Carvalho C.C.C.R., da Fonseca M.M.R. The remarkable Rhodococcus erythropolis. Appl. Microbiol. Biotechnol. 2005;67:715–726. doi: 10.1007/s00253-005-1932-3.
    1. Cortes M.A.L.R.M., de Carvalho C.C.C.R. Effect of carbon sources on lipid accumulation in Rhodococcus cells. Biochem. Eng. J. 2015;94:100–105. doi: 10.1016/j.bej.2014.11.017.
    1. Wältermann M., Luftmann H., Baumeister D., Kalscheuer R., Steinbüchel A. Rhodococcus opacus strain PD630 as a new source of high-value single-cell oil? Isolation and characterization of triacylglycerols and other storage lipids. Microbiology. 2000;146:1143–1149. doi: 10.1099/00221287-146-5-1143.
    1. Alvarez H., Silva R., Herrero M., Hernández M., Villalba M.S. Metabolism of triacylglycerols in Rhodococcus species: Insights from physiology and molecular genetics. J. Mol. Biochem. 2013;2:69–78.
    1. Alvarez H.M. Relationship between β-oxidation pathway and the hydrocarbon-degrading profile in actinomycetes bacteria. Int. Biodeterior. Biodegrad. 2003;52:35–42. doi: 10.1016/S0964-8305(02)00120-8.
    1. Alvarez H.M., Mayer F., Fabritius D., Steinbüchel A. Formation of intracytoplasmic lipid inclusions by Rhodococcus opacus strain PD630. Arch. Microbiol. 1996;165:377–386. doi: 10.1007/s002030050341.
    1. Castro A.R., Rocha I., Alves M.M., Pereira M.A. Rhodococcus opacus B4: A promising bacterium for production of biofuels and biobased chemicals. AMB Express. 2016;6:35. doi: 10.1186/s13568-016-0207-y.
    1. Farese R.V., Walther T.C. Lipid droplets finally get a little R-E-S-P-E-C-T. Cell. 2009;139:855–860. doi: 10.1016/j.cell.2009.11.005.
    1. Walther T.C., Farese R.V. Lipid droplets and cellular lipid metabolism. Annu. Rev. Biochem. 2012;81:687–714. doi: 10.1146/annurev-biochem-061009-102430.
    1. Greenberg A.S., Coleman R.A., Kraemer F.B., McManaman J.L., Obin M.S., Puri V., Yan Q.W., Miyoshi H., Mashek D.G. The role of lipid droplets in metabolic disease in rodents and humans. J. Clin. Investig. 2011;121:2102–2110. doi: 10.1172/JCI46069.
    1. Welte M.A. Expanding roles for lipid droplets. Curr. Biol. CB. 2015;25:R470–R481. doi: 10.1016/j.cub.2015.04.004.
    1. Borel P., Grolier P., Armand M., Partier A., Lafont H., Lairon D., Azais-Braesco V. Carotenoids in biological emulsions: Solubility, surface-to-core distribution and release from lipid droplets. J. Lipid Res. 1996;37:250–261.
    1. Goodrich H.B., Hill G.A., Arrick M.S. The chemical identification of gene-controlled pigments in Platypoecilus and Xiphophorus and comparisons with other tropical fish. Genetics. 1941;26:573–586.
    1. Matsumoto J. Studies on fine structure and cytochemical properties of erythrophores in swordtail, Xiphophorus helleri, with special reference to their pigment granules (pterinosomes) J. Cell Biol. 1965;27:493–504. doi: 10.1083/jcb.27.3.493.
    1. Krinsky N.I. The protective function of carotenoid pigments. In: Giese A.C., editor. Photophysiology. Academic Press; New York, NY, USA: 1968. pp. 123–195.
    1. Cogdell R.J. Carotenoids in photosynthesis. Philos. Transac. R. Soc. Lond. B Biol. Sci. 1978;284:569–579. doi: 10.1098/rstb.1978.0090.
    1. Maoka T. Carotenoids in marine animals. Mar. Drugs. 2011;9:278. doi: 10.3390/md9020278.
    1. Schneider T., Grosbois G., Vincent W.F., Rautio M. Carotenoid accumulation in copepods is related to lipid metabolism and reproduction rather than to UV-protection. Limnol. Oceanogr. 2016;61:1201–1213. doi: 10.1002/lno.10283.
    1. Schneider T., Grosbois G., Vincent W.F., Rautio M. Saving for the future: Pre-winter uptake of algal lipids supports copepod egg production in spring. Freshw. Biol. 2017;62:1063–1072. doi: 10.1111/fwb.12925.
    1. Goodman D.S. Overview of current knowledge of metabolism of vitamin A and carotenoids12. JNCI J. Nat. Cancer Inst. 1984;73:1375–1379. doi: 10.1093/jnci/73.6.1375.
    1. Kainz M.J., Fisk A.T. Integrating lipids and contaminants in aquatic ecology and ecotoxicology. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 93–114.
    1. Ansari G.A.S., Kaphalia B.S., Khan M.F. Fatty acid conjugates of xenobiotics. Toxicol. Lett. 1995;75:1–17. doi: 10.1016/0378-4274(94)03171-3.
    1. D’Adamo R., Pelosi S., Trotta P., Sansone G. Bioaccumulation and biomagnification of polycyclic aromatic hydrocarbons in aquatic organisms. Mar. Chem. 1997;56:45–49. doi: 10.1016/S0304-4203(96)00042-4.
    1. Wong C.S., Mabury S.A., Whittle D.M., Backus S.M., Teixeira C., DeVault D.S., Bronte C.R., Muir D.C.G. Organochlorine compounds in Lake Superior: chiral polychlorinated biphenyls and biotransformation in the aquatic food web. Environ. Sci. Technol. 2004;38:84–92. doi: 10.1021/es0346983.
    1. Murphy G., Rouse R.L., Polk W.W., Henk W.G., Barker S.A., Boudreaux M.J., Floyd Z.E., Penn A.L. Combustion-derived hydrocarbons localize to lipid droplets in respiratory cells. Am. J. Respir. Cell Mol. Biol. 2008;38:532–540. doi: 10.1165/rcmb.2007-0204OC.
    1. Rowan-Carroll A., Halappanavar S., Williams A., Somers C.M., Yauk C.L. Mice exposed in situ to urban air pollution exhibit pulmonary alterations in gene expression in the lipid droplet synthesis pathways. Environ. Mol. Mutag. 2013;54:240–249. doi: 10.1002/em.21768.
    1. Olofsson S.O., Boström P., Andersson L., Rutberg M., Perman J., Borén J. Lipid droplets as dynamic organelles connecting storage and efflux of lipids. Biochim. Biophys. Acta (BBA) Mol. Cell Biol. Lipids. 2009;1791:448–458. doi: 10.1016/j.bbalip.2008.08.001.
    1. Henne W.M., Reese M.L., Goodman J.M. The assembly of lipid droplets and their roles in challenged cells. EMBO J. 2018 doi: 10.15252/embj.201898947.
    1. Hugo W.B., Stretton R.J. The role of cellular lipid in the resistance of Gram-positive bacteria to penicillins. Microbiology. 1966;42:133–138. doi: 10.1099/00221287-42-1-133.
    1. Chang W., Zhang M., Zheng S., Li Y., Li X., Li W., Li G., Lin Z., Xie Z., Zhao Z., et al. Trapping toxins within lipid droplets is a resistance mechanism in fungi. Sci. Rep. 2015;5:15133. doi: 10.1038/srep15133.
    1. Alvarez H.M., Luftmann H., Silva R.A., Cesari A.C., Viale A., Wältermann M., Steinbüchel A. Identification of phenyldecanoic acid as a constituent of triacylglycerols and wax ester produced by Rhodococcus opacus PD630. Microbiology. 2002;148:1407–1412. doi: 10.1099/00221287-148-5-1407.
    1. Alvarez H.M., Souto M.F., Viale A., Pucci O.H. Biosynthesis of fatty acids and triacylglycerols by 2,6,10,14-tetramethyl pentadecane-grown cells of Nocardia globerula 432. FEMS Microbiol. Lett. 2001;200:195–200. doi: 10.1111/j.1574-6968.2001.tb10715.x.
    1. Uauy R., Hoffman D.R., Peirano P., Birch D.G., Birch E.E. Essential fatty acids in visual and brain development. Lipids. 2001;36:885–895. doi: 10.1007/s11745-001-0798-1.
    1. Glomset J.A. Role of docosahexaenoic acid in neuronal plasma membranes. Sci. STKE. 2006;2006:pe6. doi: 10.1126/stke.3212006pe6.
    1. Cao D., Kevala K., Kim J., Moon H.S., Jun S.B., Lovinger D., Kim H.Y. Docosahexaenoic acid promotes hippocampal neuronal development and synaptic function. J. Neurochem. 2009;111:510–521. doi: 10.1111/j.1471-4159.2009.06335.x.
    1. Marcheselli V.L., Hong S., Lukiw W.J., Tian X.H., Gronert K., Musto A., Hardy M., Gimenez J.M., Chiang N., Serhan C.N., et al. Novel docosanoids inhibit brain ischemia-reperfusion-mediated leukocyte infiltration and pro-inflammatory gene expression. J. Biol. Chem. 2003;278:43807–43817. doi: 10.1074/jbc.M305841200.
    1. Bazan N.G., Molina M.F., Gordon W.C. Docosahexaenoic acid signalolipidomics in nutrition: Significance in aging, neuroinflammation, macular degeneration, Alzheimer’s and other neurodegenerative diseases. Annu. Rev. Nutr. 2011;31:321–351. doi: 10.1146/annurev.nutr.012809.104635.
    1. Orr S.K., Trépanier M.O., Bazinet R.P. n-3 Polyunsaturated fatty acids in animal models with neuroinflammation. Prostaglandins Leukot. Essent. Fat. Acids. 2013;88:97–103. doi: 10.1016/j.plefa.2012.05.008.
    1. Bang H.O., Dyerberg J. Plasma lipids and lipoproteins in Greenlandic west coast Eskimos. Acta Med. Scand. 1972;192:85–94. doi: 10.1111/j.0954-6820.1972.tb04782.x.
    1. Dyerberg J., Bang H.O., Stoffersen E., Moncada S., Vane J.R. Eicosapentaenoic acid and prevention of thrombosis and atheroscloerosis? Lancet. 1978;312:117–119. doi: 10.1016/S0140-6736(78)91505-2.
    1. Dyerberg J. Linolenate-derived polyunsaturated fatty acids and prevention of atherosclerosis. Nutr. Rev. 1986;44:125–134. doi: 10.1111/j.1753-4887.1986.tb07603.x.
    1. Leaf A., Kang J.X., Xiao Y.F., Billman G. Clinical prevention of sudden cardiac death by n-3 polyunsaturated fatty acids and mechanism of prevention of arrhythmias by n-3 fish oils. Circulation. 2003;107:2646–2652. doi: 10.1161/01.CIR.0000069566.78305.33.
    1. Tong M., Wang J., Ji Y., Chen X., Wang J., Wang S., Ruan L., Cui H., Zhou Y., Zhang Q., et al. Effect of eicosapentaenoic acid and pitavastatin on electrophysiology and anticoagulant gene expression in mice with rapid atrial pacing. Exp. Ther. Med. 2017;14:2310–2316. doi: 10.3892/etm.2017.4741.
    1. Harris W.S. n-3 fatty acids and serum lipoproteins: Human studies. Am. J. Clin. Nutr. 1997;65:1645S–1654S. doi: 10.1093/ajcn/65.5.1645S.
    1. Gladine C., Newman J.W., Durand T., Pedersen T.L., Galano J.M., Demougeot C., Berdeaux O., Pujos-Guillot E., Mazur A., Comte B. Lipid profiling following intake of the omega 3 fatty acid DHA identifies the peroxidized metabolites F4-neuroprostanes as the best predictors of atherosclerosis prevention. PLoS ONE. 2014;9:e89393. doi: 10.1371/journal.pone.0089393.
    1. Phillips M.C. Molecular mechanisms of cellular cholesterol efflux. J. Biol. Chem. 2014;289:24020–24029. doi: 10.1074/jbc.R114.583658.
    1. Yamagata K. Docosahexaenoic acid regulates vascular endothelial cell function and prevents cardiovascular disease. Lipids Health Dis. 2017;16:118. doi: 10.1186/s12944-017-0514-6.
    1. Knapp H.R. Dietary fatty acids in human thrombosis and hemostasis. Am. J. Clin. Nutri. 1997;65:1687S–1698S. doi: 10.1093/ajcn/65.5.1687S.
    1. Calder P.C. Polyunsaturated fatty acids, inflammation and immunity. Lipids. 2001;36:1007–1024. doi: 10.1007/s11745-001-0812-7.
    1. Ferreri C., Masi A., Sansone A., Giacometti G., Larocca V.A., Menounou G., Scanferlato R., Tortorella S., Rota D., Conti M., et al. Fatty acids in membranes as homeostatic, metabolic and nutritional biomarkers: Recent advancements in analytics and diagnostics. Diagnostics. 2017;7 doi: 10.3390/diagnostics7010001.
    1. Bergström S., Danielsson H., Samuelsson B. The enzymatic formation of prostaglandin E2 from arachidonic acid prostaglandins and related factors 32. Biochim. Biophys. Acta (BBA) Gen. Subj. 1964;90:207–210. doi: 10.1016/0304-4165(64)90145-X.
    1. Van Dorp D.A., Beerthuis R.K., Nugteren D.H., Vonkeman H. The biosynthesis of prostaglandins. Biochim. Biophys. Acta (BBA) Gen. Subj. 1964;90:204–207. doi: 10.1016/0304-4165(64)90144-8.
    1. Corey E.J., Albright J.O., Barton A.E., Hashimoto S. Chemical and enzymic syntheses of 5-HPETE, a key biological precursor of slow-reacting substance of anaphylaxis (SRS) and 5-HETE. J. Am. Chem. Soc. 1980;102:1435–1436. doi: 10.1021/ja00524a044.
    1. Von Euler U.S. On the specific vaso-dilating and plain muscle stimulating substances from accessory genital glands in man and certain animals (prostaglandin and vesiglandin) J. Physiol. 1936;88:213–234. doi: 10.1113/jphysiol.1936.sp003433.
    1. Samuelsson B. From studies of biochemical mechanism to novel biological mediators: Prostaglandin endoperoxides, thromboxanes and leukotrienes (Nobel Lecture) Angew. Chem. Int. Ed. Engl. 1983;22:805–815. doi: 10.1002/anie.198308053.
    1. Ricciotti E., FitzGerald G.A. Prostaglandins and inflammation. Atertio. Thromb. Vasc. Biol. 2011;31:986–1000. doi: 10.1161/ATVBAHA.110.207449.
    1. Duvall M.G., Levy B.D. DHA- and EPA-derived resolvins, protectins and maresins in airway inflammation. Eur. J. Pharmacol. 2016;785:144–155. doi: 10.1016/j.ejphar.2015.11.001.
    1. Levy B.D., Clish C.B., Schmidt B., Gronert K., Serhan C.N. Lipid mediator class switching during acute inflammation: Signals in resolution. Nat. Immunol. 2001;2:612. doi: 10.1038/89759.
    1. Serhan C.N. Novel pro-resolving lipid mediators in inflammation are leads for resolution physiology. Nature. 2014;510:92–101. doi: 10.1038/nature13479.
    1. Stanley-Samuelson D.W. The biological significance of prostaglandins and related eicosanoids in invertebrates. Am. Zool. 1994;34:589–598. doi: 10.1093/icb/34.6.589.
    1. Stanley D.W., Howard R.W. The biology of prostaglandins and related eicosanoids in invertebrates: Cellular, organismal and ecological actions. Am. Zool. 1998;38:369–381. doi: 10.1093/icb/38.2.369.
    1. Miralto G.B., Romano G., Sa P., Ianora A., Russo G.L., Buttino I., Mazzarella G., Laabir M., Cabrini M., Mg G. The insidious effect of diatoms on copepod reproduction. Nature. 1999;402:173–176. doi: 10.1038/46023.
    1. Eckardt N.A. Oxylipin signaling in plant stress responses. Plant Cell. 2008;20:495–497. doi: 10.1105/tpc.108.059485.
    1. Okazaki Y., Saito K. Roles of lipids as signaling molecules and mitigators during stress response in plants. Plant J. 2014;79:584–596. doi: 10.1111/tpj.12556.
    1. Watson S.B., Caldwell G., Pohnert G. Fatty acids and oxylipins as semiochemicals. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 65–92.
    1. Pohl H.C., Kock L.J. Oxidized fatty acids as inter-kingdom signaling molecules. Molecules. 2014;19:1273–1285. doi: 10.3390/molecules19011273.
    1. Okamoto T., Katoh S. Linolenic acid binding by chloroplasts. Plant Cell Physiol. 1977;18:539–550. doi: 10.1093/oxfordjournals.pcp.a075466.
    1. Wu J.T., Chiang Y.R., Huang W.Y., Jane W.N. Cytotoxic effects of free fatty acids on phytoplankton algae and cyanobacteria. Aquat. Toxicol. 2006;80:338–345. doi: 10.1016/j.aquatox.2006.09.011.
    1. Murakami M., Makabe K., Yamaguchi K., Konosu S. Cytotoxic polyunsaturated fatty acid from Pediastrum. Phytochemistry. 1989;28:625–626. doi: 10.1016/0031-9422(89)80065-2.
    1. Rukmini C. Reproductive toxicology and nutritional studies on mahua oil (Madhuca latifolia) Food Chem. Toxicol. 1990;28:601–605. doi: 10.1016/0278-6915(90)90166-K.
    1. Sellem F., Pesando D., Bodennec G., El Abed A., Girard J.P. Toxic effects of Gymnodinium cf. mikimotoi unsaturated fatty acids to gametes and embryos of the sea urchin Paracentrotus lividus. Water Res. 2000;34:550–556. doi: 10.1016/S0043-1354(99)00181-5.
    1. Reinikainen M., Meriluoto J.A.O., Spoof L., Harada K.-i. The toxicities of a polyunsaturated fatty acid and a microcystin to Daphnia magna. Environ. Toxicol. 2001;16:444–448. doi: 10.1002/tox.10003.
    1. Hombeck M., Pohnert G., Boland W. Biosynthesis of dictyopterene A: Stereoselectivity of a lipoxygenase/hydroperoxide lyase from Gomphonema parvulum (Bacillariophyceae) Chem. Commun. 1999:243–244. doi: 10.1039/a808409b.
    1. Barofsky A., Pohnert G. Biosynthesis of polyunsaturated short chain aldehydes in the diatom Thalassiosira rotula. Org. Lett. 2007;9:1017–1020. doi: 10.1021/ol063051v.
    1. Wichard T., Pohnert G. Formation of halogenated medium chain hydrocarbons by a lipoxygenase/hydroperoxide halolyase-mediated transformation in planktonic microalgae. J. Am. Chem. Soc. 2006;128:7114–7115. doi: 10.1021/ja057942u.
    1. Tosti E., Romano G., Buttino I., Cuomo A., Ianora A., Miralto A. Bioactive aldehydes from diatoms block the fertilization current in ascidian oocytes. Mol. Reprod. Dev. 2003;66:72–80. doi: 10.1002/mrd.10332.
    1. Adolph S., Poulet S.A., Pohnert G. Synthesis and biological activity of α,β,γ,δ-unsaturated aldehydes from diatoms. Tetrahedron. 2003;59:3003–3008. doi: 10.1016/S0040-4020(03)00382-X.
    1. Lewis C., Caldwell G.S., Bentley M.G., Olive P.J.W. Effects of a bioactive diatom-derived aldehyde on developmental stability in Nereis virens (Sars) larvae: An analysis using fluctuating asymmetry. J. Exp. Mar. Biol. Ecol. 2004;304:1–16. doi: 10.1016/j.jembe.2003.11.018.
    1. Caldwell G.S. The influence of bioactive oxylipins from marine diatoms on invertebrate reproduction and development. Mar. Drugs. 2009;7:367–400. doi: 10.3390/md7030367.
    1. Ianora A., Poulet S.A., Miralto A. The effects of diatoms on copepod reproduction: A review. Phycologia. 2003;42:351–363. doi: 10.2216/i0031-8884-42-4-351.1.
    1. de Carvalho C.C.C.R., Caramujo M.J. Bacterial diversity assessed by cultivation-based techniques shows predominance of Staphylococccus species on coins collected in Lisbon and Casablanca. FEMS Microbiol. Ecol. 2014;88:26–37. doi: 10.1111/1574-6941.12266.
    1. Dijkman N.A., Kromkamp J.C. Phospholipid-derived fatty acids as chemotaxonomic markers for phytoplankton: Application for inferring phytoplankton composition. Mar. Ecol. Prog. Ser. 2006;324:113–125. doi: 10.3354/meps324113.
    1. Graeve M., Kattner G., Hagen W. Diet-induced changes in the fatty acid composition of Arctic herbivorous copepods: Experimental evidence of trophic markers. J. Exp. Mar. Biol. Ecol. 1994;182:97–110. doi: 10.1016/0022-0981(94)90213-5.
    1. Pollierer M.M., Scheu S., Haubert D. Taking it to the next level: Trophic transfer of marker fatty acids from basal resource to predators. Soil Biol. Biochem. 2010;42:919–925. doi: 10.1016/j.soilbio.2010.02.008.
    1. Kunitsky C., Osterhout G., Sasser M. Identification of microorganisms using fatty acid methyl ester (FAME) analysis and the MIDI Sherlock Microbial Identification System. In: Miller M.J., editor. Encyclopedia of Rapid Microbiological Methods. Volume 3. PDA Bookstore; Baltimore, MD, USA: 2006. pp. 1–18.
    1. Ifkovits R.W., Ragheb H.S. Cellular fatty acid composition and identification of rumen bacteria. Appl. Microbiol. 1968;16:1406–1413.
    1. Tang Y.W., Ellis N.M., Hopkins M.K., Smith D.H., Dodge D.E., Persing D.H. Comparison of phenotypic and genotypic techniques for identification of unusual aerobic pathogenic Gram-negative bacilli. J. Clin. Microbiol. 1998;36:3674–3679.
    1. Abel K., deSchmertzing H., Peterson J.I. Classification of microorganisms by analysis of chemical composition I. Feasibility of utilizing gas chromatography. J. Bacteriol. 1963;85:1039–1044.
    1. Osterhout G.J., Shull V.H., Dick J.D. Identification of clinical isolates of gram-negative nonfermentative bacteria by an automated cellular fatty acid identification system. J. Clin. Microbiol. 1991;29:1822–1830.
    1. Sasser M. Bacterial Identification by Gas Chromatographic Analysis of Fatty Acid Methyl Esters (GC-FAME) MIDI, Inc.; Newark, DE, USA: 1990. Technical Note #101.
    1. Stead D.E., Sellwood J.E., Wilson J., Viney I. Evaluation of a commercial microbial identification system based on fatty acid profiles for rapid, accurate identification of plant pathogenic bacteria. J. Appl. Bacteriol. 1992;72:315–321. doi: 10.1111/j.1365-2672.1992.tb01841.x.
    1. Leonard R.B., Mayer J., Sasser M., Woods M.L., Mooney B.R., Brinton B.G., Newcomb-Gayman P.L., Carroll K.C. Comparison of MIDI Sherlock system and pulsed-field gel electrophoresis in characterizing strains of methicillin-resistant Staphylococcus aureus from a recent hospital outbreak. J. Clin. Microbiol. 1995;33:2723–2727.
    1. Birnbaum D., Herwaldt L., Low D.E., Noble M., Pfaller M., Sherertz R., Chow A.W. Efficacy of microbial identification system for epidemiologic typing of coagulase-negative staphylococci. J. Clin. Microbiol. 1994;32:2113–2119.
    1. Ash C., Farrow J.A.E., Wallbanks S., Collins M.D. Phylogenetic heterogeneity of the genus Bacillus revealed by comparative analysis of small-subunit-ribosomal RNA sequences. Lett. Appl. Microbiol. 1991;13:202–206. doi: 10.1111/j.1472-765X.1991.tb00608.x.
    1. Priest F.G., Kaji D.A., Rosato Y.B., Canhos V.P. Characterization of Bacillus thuringiensis and related bacteria by ribosomal RNA gene restriction fragment length polymorphisms. Microbiology. 1994;140:1015–1022. doi: 10.1099/13500872-140-5-1015.
    1. Bourque S.N., Valero J.R., Lavoie M.C., Levesque R.C. Comparative analysis of the 16S to 23S ribosomal intergenic spacer sequences of Bacillus thuringiensis strains and subspecies and of closely related species. Appl. Environ. Microbiol. 1995;61:1623–1626.
    1. Wintzingerode F., Rainey F.A., Kroppenstedt R.M., Stackebrandt E. Identification of environmental strains of Bacillus mycoides by fatty acid analysis and species-specific 16S rDNA oligonucleotide probe. FEMS Microbiol. Ecol. 1997;24:201–209. doi: 10.1016/S0168-6496(97)00057-3.
    1. Slabbinck B., De Baets B., Dawyndt P., De Vos P. Genus-wide Bacillus species identification through proper artificial neural network experiments on fatty acid profiles. Antonie Van Leeuwenhoek. 2008;94:187–198. doi: 10.1007/s10482-008-9229-z.
    1. Abraham W.R., Lünsdorf H., Vancanneyt M., Smit J. Cauliform bacteria lacking phospholipids from an abyssal hydrothermal vent: Proposal of Glycocaulis abyssi gen. nov., sp. nov., belonging to the family Hyphomonadaceae. Int. J. Syst. Evol. Microbiol. 2013;63:2207–2215. doi: 10.1099/ijs.0.047894-0.
    1. Torreblanca M., Rodriguez-Valera F., Juez G., Ventosa A., Kamekura M., Kates M. Classification of non-alkaliphilic halobacteria based on numerical taxonomy and polar lipid composition and description of Haloarcula gen. nov. and Haloferax gen. nov. Syst. Appl. Microbiol. 1986;8:89–99. doi: 10.1016/S0723-2020(86)80155-2.
    1. Oren A., Ventosa A. Subcommittee on the taxonomy of Halobacteriaceae and Subcommittee on the taxonomy of Halomonadaceae. Int. J. Syst. Evol. Microbiol. 2013;63:3540–3544. doi: 10.1099/ijs.0.055988-0.
    1. de la Haba R.R., Corral P., Sánchez-Porro C., Infante-Domínguez C., Makkay A.M., Amoozegar M.A., Ventosa A., Papke R.T. Genotypic and lipid analyses of strains from the Archaeal genus Halorubrum reveal insights into their taxonomy, divergence and population structure. Front. Microbiol. 2018;9:512. doi: 10.3389/fmicb.2018.00512.
    1. White D.C., Davis W.M., Nickels J.S., King J.D., Bobbie R.J. Determination of the sedimentary microbial biomass by extractible lipid phosphate. Oecologia. 1979;40:51–62. doi: 10.1007/BF00388810.
    1. Pinkart H., Ringelberg D., Piceno Y., Macnaughton S., White D.C. Biochemical Approaches to Biomass Measurements and Community Structure. American Society for Microbiology Press; Washington, DC, USA: 2002. pp. 101–113.
    1. Frostegård Å., Bååth E., Tunlio A. Shifts in the structure of soil microbial communities in limed forests as revealed by phospholipid fatty acid analysis. Soil Biol. Biochem. 1993;25:723–730. doi: 10.1016/0038-0717(93)90113-P.
    1. Feinstein L.M., Sul W.J., Blackwood C.B. Assessment of bias associated with incomplete extraction of microbial DNA from soil. Appl. Environ. Microbiol. 2009;75:5428–5433. doi: 10.1128/AEM.00120-09.
    1. Zhang Z., Qu Y., Li S., Feng K., Wang S., Cai W., Liang Y., Li H., Xu M., Yin H., et al. Soil bacterial quantification approaches coupling with relative abundances reflecting the changes of taxa. Sci. Rep. 2017;7:4837. doi: 10.1038/s41598-017-05260-w.
    1. Ramsey P.W., Rillig M.C., Feris K.P., Holben W.E., Gannon J.E. Choice of methods for soil microbial community analysis: PLFA maximizes power compared to CLPP and PCR-based approaches. Pedobiologia. 2006;50:275–280. doi: 10.1016/j.pedobi.2006.03.003.
    1. White D.C., Ringelberg D.B. Signature lipid biomarker analysis. In: Burlage R.S., Atlas R., Stahl D., Geesey G., Sayler G., editors. Techniques in Microbial Ecology. Oxford University Press; New York, NY, USA: 1998. pp. 255–272.
    1. Ackman R.G., Tocher C.S., McLachlan J. Marine phytoplankter fatty acids. J. Fish. Res. Board Can. 1968;25:1603–1620. doi: 10.1139/f68-145.
    1. Ahlgren G., Gustafsson I.B., Boberg M. Fatty acid content and chemical composition of freshwater microalgae. J. Phycol. 1992;28:37–50. doi: 10.1111/j.0022-3646.1992.00037.x.
    1. Bourdier G.G., Amblard C.A. Lipids in Acanthodiaptomus denticomis during starvation and fed on three different algae. J. Plankton Res. 1989;11:1201–1212. doi: 10.1093/plankt/11.6.1201.
    1. Nichols P.D., Jones G.J., De Leeuw J.W., Johns R.B. The fatty acid and sterol composition of two marine dinoflagellates. Phytochemistry. 1984;23:1043–1047. doi: 10.1016/S0031-9422(00)82605-9.
    1. Viso A.-C., Marty J.C. Fatty acids from 28 marine microalgae. Phytochemistry. 1993;34:1521–1533. doi: 10.1016/S0031-9422(00)90839-2.
    1. Volkman J.K., Jeffrey S.W., Nichols P.D., Rogers G.I., Garland C.D. Fatty acid and lipid composition of 10 species of microalgae used in mariculture. J. Exp. Mar. Biol. Ecol. 1989;128:219–240. doi: 10.1016/0022-0981(89)90029-4.
    1. Dunstan G.A., Volkman J.K., Barrett S.M., Leroi J.M., Jeffrey S.W. Essential polyunsaturated fatty acids from 14 species of diatom (Bacillariophyceae) Phytochemistry. 1993;35:155–161. doi: 10.1016/S0031-9422(00)90525-9.
    1. Zhukova N.V., Aizdaicher N.A. Fatty acid composition of 15 species of marine microalgae. Phytochemistry. 1995;39:351–356. doi: 10.1016/0031-9422(94)00913-E.
    1. Caramujo M.J., Boschker H.T.S., Admiraal W.I.M. Fatty acid profiles of algae mark the development and composition of harpacticoid copepods. Freshw. Biol. 2008;53:77–90. doi: 10.1111/j.1365-2427.2007.01868.x.
    1. Napolitano G.E. The relationship of lipids with light and chlorophyll measurements in freshwater algae and periphyton. J. Phycol. 1994;30:943–950. doi: 10.1111/j.0022-3646.1994.00943.x.
    1. Guschina I.A., Harwood J.L. Algal lipids and effect of the environment on their biochemistry. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 1–24.
    1. Reuss N., Poulsen L. Evaluation of fatty acids as biomarkers for a natural plankton community. A field study of a spring bloom and a post-bloom period off West Greenland. Mar. Biol. 2002;141:423–434. doi: 10.1007/s00227-002-0841-6.
    1. Cañavate J.P. Advancing assessment of marine phytoplankton community structure and nutritional value from fatty acid profiles of cultured microalgae. Rev. Aquacult. 2018 doi: 10.1111/raq.12244.
    1. Bec A., Perga M.E., Desvilettes C., Bourdier G. How well can the fatty acid content of Lake Seston be predicted from its taxonomic composition? Freshw. Biol. 2010;55:1958–1972. doi: 10.1111/j.1365-2427.2010.02429.x.
    1. Desvilettes C., Bec A. Formation and transfer of fatty acids in aquatic microbial food webs: Role of heterotrophic protists. In: Kainz M., Brett M.T., Arts M.T., editors. Lipids in Aquatic Ecosystems. Springer; New York, NY, USA: 2009. pp. 25–42.
    1. Copeman L.A., Parrish C.C., Gregory R.S., Jamieson R.E., Wells J., Whiticar M.J. Fatty acid biomarkers in coldwater eelgrass meadows: Elevated terrestrial input to the food web of age-0 Atlantic cod Gadus morhua. Mar. Ecol. Prog. Ser. 2009;386:237–251. doi: 10.3354/meps08063.
    1. McMeans B.C., Koussoroplis A.M., Kainz M.J. Effects of seasonal seston and temperature changes on lake zooplankton fatty acids. Limnol. Oceanogr. 2015;60:573–583. doi: 10.1002/lno.10041.
    1. Budge S., Springer A., Iverson S., Sheffield G. Fatty acid biomarkers reveal niche separation in an Arctic benthic food web. Mar. Ecol. Prog. Ser. 2007;336:305–309. doi: 10.3354/meps336305.
    1. James Henderson R., Tocher D.R. The lipid composition and biochemistry of freshwater fish. Prog. Lipid Res. 1987;26:281–347. doi: 10.1016/0163-7827(87)90002-6.
    1. Dalsgaard J., St John M., John G., Kattner D.C., Muller-Navarra D.C., Hagen W. Fatty acid trophic markers in the pelagic marine food environment. Adv. Mar. Biol. 2003;46:226–340. doi: 10.1016/S0065-2881(03)46005-7.
    1. Iverson S.J., Field C., Don Bowen W., Blanchard W. Quantitative fatty acid signature analysis: A new method of estimating predator diets. Ecol. Monogr. 2004;74:211–235. doi: 10.1890/02-4105.
    1. Falk-Petersen S., Haug T., Nilssen K.T., Wold A., Dahl T.M. Lipids and trophic linkages in harp seal (Phoca groenlandica) from the eastern Barents Sea. Polar Res. 2006;23:43–50. doi: 10.3402/polar.v23i1.6265.
    1. Martin-Creuzburg D., Kowarik C., Straile D. Cross-ecosystem fluxes: Export of polyunsaturated fatty acids from aquatic to terrestrial ecosystems via emerging insects. Sci. Total Environ. 2017;577:174–182. doi: 10.1016/j.scitotenv.2016.10.156.
    1. Ludvigsen L., Albrechtsen H.J., Holst H., Christensen T.H. Correlating phospholipid fatty acids (PLFA) in a landfill leachate polluted aquifer with biogeochemical factors by multivariate statistical methods. FEMS Microbiol. Rev. 1997;20:447–460. doi: 10.1111/j.1574-6976.1997.tb00329.x.
    1. Green C.T., Scow K.M. Analysis of phospholipid fatty acids (PLFA) to characterize microbial communities in aquifers. Hydrogeol. J. 2000;8:126–141. doi: 10.1007/s100400050013.
    1. de Carvalho C.C.C.R., Caramujo M.-J. Fatty acids as a tool to understand microbial diversity and their role in food webs of Mediterranean temporary ponds. Molecules. 2014;19:5570–5598. doi: 10.3390/molecules19055570.
    1. Sundh I., Nilsson M., Borga P. Variation in microbial community structure in two boreal peatlands as determined by analysis of phospholipid fatty acid profiles. Appl. Environ. Microbiol. 1997;63:1476–1482.
    1. Quezada M., Buitrón G., Moreno-Andrade I., Moreno G., López-Marín L.M. The use of fatty acid methyl esters as biomarkers to determine aerobic, facultatively aerobic and anaerobic communities in wastewater treatment systems. FEMS Microbiol. Lett. 2007;266:75–82. doi: 10.1111/j.1574-6968.2006.00509.x.
    1. Torres-Ruiz M., Wehr J.D., Perrone A.A. Trophic relations in a stream food web: Importance of fatty acids for macroinvertebrate consumers. J. N. Am. Benthol. Soc. 2007;26:509–522. doi: 10.1899/06-070.1.
    1. de Carvalho C.C.C.R., Caramujo M. Lipids of prokaryotic origin at the base of marine food webs. Mar. Drugs. 2012;10:2698–2714. doi: 10.3390/md10122698.
    1. Middelburg J.J. Stable isotopes dissect aquatic food webs from the top to the bottom. Biogeosciences. 2014;11:2357–2371. doi: 10.5194/bg-11-2357-2014.
    1. De Troch M., Boeckx P., Cnudde C., Van Gansbeke D., Vanreusel A., Vincx M., Caramujo M.J. Bioconversion of fatty acids at the basis of marine food webs: Insights from a compound-specific stable isotope analysis. Mar. Ecol. Prog. Ser. 2012;465:53–67. doi: 10.3354/meps09920.
    1. Dijkman N.A., Boschker H.T.S., Stal L.J., Kromkamp J.C. Composition and heterogeneity of the microbial community in a coastal microbial mat as revealed by the analysis of pigments and phospholipid-derived fatty acids. J. Sea Res. 2010;63:62–70. doi: 10.1016/j.seares.2009.10.002.
    1. Olsson P.A. Signature fatty acids provide tools for determination of the distribution and interactions of mycorrhizal fungi in soil. FEMS Microbiol. Ecol. 1999;29:303–310. doi: 10.1111/j.1574-6941.1999.tb00621.x.
    1. Costello A.M., Auman A.J., Macalady J.L., Scow K.M., Lidstrom M.E. Estimation of methanotroph abundance in a freshwater lake sediment. Environ. Microbiol. 2002;4:443–450. doi: 10.1046/j.1462-2920.2002.00318.x.
    1. Bowman J.P., Skerratt J.H., Nichols P.D., Sly L.I. Phospholipid fatty acid and lipopolysaccharide fatty acid signature lipids in methane-utilizing bacteria. FEMS Microbiol. Lett. 1991;85:15–21. doi: 10.1111/j.1574-6968.1991.tb04693.x.
    1. Fang J., Chan O., Joeckel R.M., Huang Y., Wang Y., Bazylinski D.A., Moorman T.B., Ang Clement B.J. Biomarker analysis of microbial diversity in sediments of a saline groundwater seep of Salt Basin, Nebraska. Org. Geochem. 2006;37:912–931. doi: 10.1016/j.orggeochem.2006.04.007.
    1. Elvert M., Boetius A., Knittel K., Jørgensen B.B. Characterization of specific membrane fatty acids as chemotaxonomic markers for sulfate-reducing bacteria involved in anaerobic oxidation of methane. Geomicrobiol. J. 2003;20:403–419. doi: 10.1080/01490450303894.
    1. Zelles L. Identification of single cultured micro-organisms based on their whole-community fatty acid profiles, using an extended extraction procedure. Chemosphere. 1999;39:665–682. doi: 10.1016/S0045-6535(99)00131-9.
    1. Lee A.K.Y., Chan C.K., Fang M., Lau A.P.S. The 3-hydroxy fatty acids as biomarkers for quantification and characterization of endotoxins and Gram-negative bacteria in atmospheric aerosols in Hong Kong. Atmos. Environ. 2004;38:6307–6317. doi: 10.1016/j.atmosenv.2004.08.013.
    1. Frostegård A., Bååth E. The use of phospholipid fatty acid analysis to estimate bacterial and fungal biomass in soil. Biol. Fertil. Soils. 1996;22:59–65. doi: 10.1007/BF00384433.
    1. Fernandes M.F., Saxena J., Dick R.P. Comparison of whole-cell fatty acid (MIDI) or phospholipid fatty acid (PLFA) extractants as biomarkers to profile soil microbial communities. Microb. Ecol. 2013;66:145–157. doi: 10.1007/s00248-013-0195-2.
    1. Rajendran N., Matsuda O., Imamura N., Urushigawa Y. Microbial community structure analysis of euxinic sediments using phospholipid fatty acid biomarkers. J. Oceanogr. 1995;51:21–38. doi: 10.1007/BF02235934.
    1. Jiang L., Cai C., Zhang Y., Mao S., Sun Y., Li K., Xiang L., Zhang C. Lipids of sulfate-reducing bacteria and sulfur-oxidizing bacteria found in the Dongsheng uranium deposit. Chin. Sci. Bull. 2012;57:1311–1319. doi: 10.1007/s11434-011-4955-4.
    1. Frostegård Å., Tunlid A., Bååth E. Phospholipid fatty acid composition, biomass and activity of microbial communities from two soil types experimentally exposed to different heavy metals. Appl. Environ. Microbiol. 1993;59:3605–3617.
    1. White D.C., Geyer R., Peacock A.D., Hedrick D.B., Koenigsberg S.S., Sung Y., He J., Löffler F.E. Phospholipid furan fatty acids and ubiquinone-8: Lipid biomarkers that may protect Dehalococcoides strains from free radicals. Appl. Environ. Microbiol. 2005;71:8426–8433. doi: 10.1128/AEM.71.12.8426-8433.2005.
    1. Stahl P.D., Klug M.J. Characterization and differentiation of filamentous fungi based on fatty acid composition. Appl. Environ. Microbiol. 1996;62:4136–4146.
    1. Sud M., Fahy E., Cotter D., Brown A., Dennis E., Glass C., Murphy R., Raetz C., Russell D., Subramaniam S. LIPID MAPS structure database. Nucleic Acids Res. 2006;35:D527–D532. doi: 10.1093/nar/gkl838.

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