In vitro synthesis of 9,10-dihydroxyhexadecanoic acid using recombinant Escherichia coli
© The Author(s) 2017
Received: 22 October 2016
Accepted: 5 May 2017
Published: 18 May 2017
Hydroxy fatty acids are widely used in food, chemical and cosmetic industries. A variety of dihydroxy fatty acids have been synthesized so far; however, no studies have been done on the synthesis of 9,10-dihydroxyhexadecanoic acid. In the present study recombinant E. coli has been used for the heterologous expression of fatty acid hydroxylating enzymes and the whole cell lysate of the induced culture was used for in vitro production of 9,10-dihydroxyhexadecanoic acid.
A first of its kind proof of principle has been successfully demonstrated for the production of 9,10-dihydroxyhexadecanoic acid using three different enzymes viz. fatty acid desaturase (FAD) from Saccharomyces cerevisiae, epoxide hydrolase (EH) from Caenorhabditis elegance and epoxygenase (EPOX) from Stokasia laevis. The genes for these proteins were codon-optimised, synthesised and cloned in pET 28a (+) vector. The culture conditions for induction of these three proteins in E. coli were optimised in shake flask. The induced cell lysates were used both singly and in combination along with the trans-supply of hexadecanoic acid and 9-hexadecenoic acid, followed by product profiling by GC–MS. Formation of 9,10-dihydroxyhexadecanoic acid was successfully achieved when combination of induced cell lysates of recombinant E. coli containing FAD, EH, and EPOX were incubated with 9-hexadecenoic acid.
The in vitro production of 9,10-dihydroxyhexadecanoic acid synthesis using three fatty acid modification genes from different sources has been successfully demonstrated. The strategy adopted can be used for the production of similar compounds.
KeywordsFatty acid desaturase Epoxygenase Epoxide hydrolase Fatty acid hydroxylation Recombinant E. coli
Hydroxy fatty acids are widely used in food, chemical and cosmetic industries as starting materials for the synthesis of polymers and as additives for the manufacture of lubricants, emulsifiers, and stabilizers. Hydroxy fatty acid producing enzymes can be broadly classified into two categories viz. fatty acid mono-hydroxylation enzymes and di-hydroxylation enzymes . Major classes of mono-hydroxylation enzymes are cytochrome P450 monooxygenases, hydratases, 12-hydroxylases and lipoxygenases. All these carry out mono-hydroxylation using different mechanisms of action. Cytochrome P450s catalyse the insertion of one oxygen atom from molecular oxygen into an organic substrate with NAD(P)H as a cofactor through electron transfer . Hydratases produce 10-hydroxy fatty acids, wherein it uses water molecule to add a hydrogen atom and a hydroxyl group at C9 and C10 positions, respectively, on to the carbon–carbon cis-double bond of unsaturated fatty acids. 12-Hydroxylases catalyze the NADH-dependent site-specific hydroxylation of the 12-position of oleic acid using oxygen to convert oleic acid to ricinoleic acid . Lipoxygenases are enzymes under family of dioxygenases, which catalyse the synthesis of hydroperoxy fatty acids of polyunsaturated fatty acids (PUFAs) having one or more cis,cis-pentadiene units by insertion of molecular oxygen [4, 5]. Fatty acid di-hydroxylation enzymes are generally known as diol synthases.
The synthesis of vicinal diol, in which hydoxyl groups are present at adjacent carbon atoms, was first reported in arachidonic acid metabolism. The reactions catalysed by cytochrome P450 monooxygenases in arachidonic acid metabolism are classified into three categories viz. epoxygenases, lipoxygenases and hydroxylases. Out of these three, epoxygenases are responsible for the formation of the epoxide derivative, which is a reduced form of hydroxy fatty acid . When a fatty acid is to be hydroxylated at adjacent carbon atoms, desaturation occurs at the corresponding C–C bond by fatty acid desaturase followed by epoxygenation of the unsaturated double bond by an epoxygenase and by hydrolysis of the epoxide by epoxide hydrolase to form vicinal diols.
In the present study, we have successfully synthesised 9,10-dihydroxyhexadecanoic acid by using three different types of fatty acid modifying enzymes, viz. fatty acid desaturase, epoxygenase and epoxide hydrolase. The sources of these enzymes were: fatty acid desaturase (FAD) from Saccharomyces cerevisiae, epoxide hydrolase (EH) from Caenorhabditis elegance and epoxygenase (EPOX) from Stokasia laevis. The genes for these enzymes were codon-optimised and synthesised for expression in E. coli and the whole cell lysate of the same was used for the synthesis of 9,10-dihydroxyhexadecanoic acid.
Synthesis of genes and cloning
Gene sequences of fatty acid desaturase (FAD) from S. cerevisiae (Accession number NP_011460), epoxide hydrolase (EH) from Caenorhabditis elegance (Accession number ABV45408), and epoxygenase (EPOX) from Stokasia laevis (Accession Number AAR23815) were codon-optimised for expression in E. coli and were chemically synthesised from GenScript® USA. The synthesised genes were having BamHI and HindIII sites at 5′ and 3′ ends respectively. The synthesised genes were confirmed by sequencing using ABI 3130 automated sequencer (Applied Biosystems, Inc. Carlsbad, CA, USA). These genes were cloned into pET 28a (+) vector after double digestion of the vector by BamHI and HindIII. The cloning was confirmed by restriction digestion with BamHI and HindIII.
Optimisation of expression
The optimisation of expression of all the three genes using isopropyl β-d-1-thiogalactopyranoside (IPTG, final concentration of 100 µM) was carried out by varying incubation temperature and by changing the expression host. Four different hosts viz. E. coli BL21(DE3), BL21(DE3)-pLysS (Novagen, USA), BL21(DE3)CodonPlus-RIL and BL21(DE3)-Gold (Stratagene, USA) were used for expression. Shake flask cultures (250 mL) at 250 rpm were used for this study wherein Luria both was supplemented with kanamycin (50 µg/mL). After reaching an OD600nm of 0.4, 250 µL IPTG (100 mM) was added and the cultures were incubated at 30, 37, 42 or 16 °C for obtaining the maximum expression of the proteins. The cultures were grown till the OD600nm of 2.0 was attained. The cultures after induction were centrifuged at 6000g for 10 min and the pellets were suspended in phosphate buffer (pH 7.4). It was mixed with equal volume of 2X Laemmli buffer and run on 12% SDS PAGE to check the induction of the enzymes .
In vitro synthesis of 9,10-dihydroxyhexadecanoic acid
Sample preparation for GC–MS
After the reaction was over, the products formed were extracted with hexane and ethyl acetate in series, and both the hexane and ethyl acetate fractions were pooled and concentrated to 1 mL volume using rotary vacuum evaporator. It was combined with 3 mL of BF3-Methanol in a 10 mL test tube and was heated at 60 °C for 10 min after capping. The contents were cooled and transferred to a separating funnel with 30 mL of hexane and ethyl acetate separately. It was washed two times with a saturated NaCl solution. Aqueous (bottom) layer was discarded after each wash. Hexane/ethyl acetate extracts were dried over anhydrous sodium sulphite and transferred to a clean, dry container. Hexane and ethyl acetate extracts were concentrated using vacuum concentrator to a final volume of 3 mL. Since the expected profile constitutes epoxides, a second round of methylation was carried out after acid hydrolysis of epoxy bonds. For this a modified protocol of Cahoon et al.  was adopted. Fatty acid methyl esters (500 µL) obtained after BF3-Methanol treatment (as described above) were mixed with 1 mL of 2.5% (v/v) sulfuric acid in methanol and heated at 70 °C for 20 min. After cooling, 1 mL of water was added, and fatty acid methyl esters were extracted with 2 mL of hexane. Hexane fraction was concentrated to 300 µL using vacuum concentrator. To this 3 mL of BF3-Methanol was added and heated at 60 °C for 10 min. The methylated product, which contains vicinal dimethyl derivatives was extracted using hexane as described in the first round of methylation.
Hexane and ethyl acetate extracts were mixed in equal proportion before subjecting to GC–MS. Fatty acid methyl esters (FAME) of the lac insect were estimated using Shimadzu GCMS-QP2010 Plus as per the following conditions. Column: RTX-5MS, 30 m; Column Oven Temp.: 140 °C; Injection Temp.: 260 °C; Injection Mode: Splitless; Carrier gas: Helium; Oven Programme: 140 °C hold for 5 min; 4 °C/min 240 °C hold for 5 min; Diluent: n-Hexane; Scan range: 40-650 m/z. GC–MS analysis of the methyl ester/trimethylsilyl derivative of 9,10-dihydroxyhexadecanoic acid (LGC standards, GmBH Cat. No. LA 14-1601-5-4) was carried out using a capillary column of 5% phenylmethylsiloxane (12 m, 0.33 µm film thickness, carrier gas: helium; The temperature was raised from 120 to 300 °C at a rate of 10 °C/min).
Gene synthesis and cloning
The codon optimised sequences of the three genes were cloned in pET 28(+) vector at BamHI–HindIII restriction site. The cloning was confirmed by restriction digestion with BamHI and HindIII. The insert release of ~1.5, ~1.2 and ~1.1 kb sizes were observed when recombinant plasmids containing genes encoding for FAD, EH and EPOX, respectively, were run on 1% agarose gel after restriction digestion (Additional file 1: Figure S2). Using these three enzymes a hypothetical pathway for producing 9,10-dihydroxyhexadecanoic acid was envisaged. Before these recombinant clones were used for the biosynthesis of 9,10-dihydroxyhexadecanoic acid, it was essential to optimise the protein expression protocol.
Optimisation of protein expression
Optimum culture conditions standardised for the induction of FAD, EPOX and EH proteins
Optimum temperature (°C)
Orbital shaking (rpm)
High-level expression of recombinant protein in E. coli is reported to often result in aggregation of the expressed protein molecules into inclusion bodies . Inclusion bodies are of two types viz. classical and non-classical. Biologically active inclusion bodies are known as non-classical inclusion bodies . Most of the non-classical inclusion bodies can be solubilized even at low concentration of denaturants as they are characterized by a loose arrangement of protein molecules . The biological activity of non-classical inclusion bodies can be utilised for the synthesis of the desired product using suitable substrates.
In vitro reconstitution of 9,10-dihydroxyhexadecanoic acid
The 9,10-dihydroxyhexadecanoic acid, which was successfully synthesised during in vitro reconstitution experiment, is the precursor for the following compounds: 9,10-dibromohexadecanoic acid, palmitelaidic acid and suberic acid. Suberic acid and its derivatives have a variety of industrial uses as lubricants, plasticizers, cosmetics, hydraulic fluids, and candles. It is used in the synthesis of polyamide and alkyd resins. It is also used as an intermediate for aromatics, antiseptics and painting materials. Preparation of reduction-sensitive micelles, having potential application in delivery of anticancer drugs has been reported using suberic acid. Its application for the formulations in the fluorescent detection of amidinium-carboxylate and amidinium formation has also been reported  Palmitelaidic (C16:1 trans-9) acid has been reported to have beneficial health effects. Circulating palmitelaidic acid in adult human leads to decreased adiposity, decreased triacylglycerol, decreased insulin resistance, greater high-density lipoprotein (HDL) cholesterol concentrations and reduced incidence of diabetes . 9,10-dibromohexadecanoic acid is used in organic synthetic chemistry reactions as related to fatty acid biohydrogenation . Currently, the source of 9,10-dihydroxyhexadecanoic acid for the synthesis of above three compounds is through organic synthesis. The present study will help to explore enzymatic route for the production of 9,10-dihydroxyhexadecanoic acid using cell conversion strategy.
Various types of biosynthetic strategies have been applied for the production of hydroxy fatty acids. Use of photolithotrophic cell suspension cultures of Laminariu saccharina was reported for the production of three different types of hydroxy fatty acids viz. 15-hydroxy-5,8,11,13-eicosatetraenoic acid (15-HETE), 13-hydroxy-6,9,11,15-octadecatetraenoic acid (13-HODTA), and 13-hydroxy-9,11-octadecadienoic acid (13-HODE) . Hydroxy fatty acid production using metabolically engineered microbes such as E. coli and S. cerevisiae has been successfully achieved . Due to higher reactivity, solvent miscibility, stability, and viscosity of hydroxy fatty acids as compared to non-hydroxylated fatty acids, their applications are unlimited . Different types of hydroxy fatty acid production has been reported with the help of microbes. Production of 10-hydroxystearic acid has been achieved by whole cell conversions using wild-type [19–21] and recombinant microorganisms . 10-Hydroxystearic acid is produced from oleic acid using whole cells of recombinant E. coli expressing the oleate hydratase gene of Stenotrophomonas maltophilia . Mono- and di-hydroxy fatty acids are synthesised by cell conversions using P. aeruginosa strains [23–26]. Production of tri-hydroxy fatty acids are reported from B. megaterium ALA2 strains . By expressing oleate 12-hydroxylase gene of C. purpurea in Schizosaccharomyces pombe, ricinoleic acid has been produced from oleic acid . Hydroperoxy fatty acids as precursors of hydroxy fatty acids have been produced from unsaturated fatty acids by lipoxygenases. Soybean 13-lipoxygenases and fungal Mn-lipoxygenase produce 13-hydropreroxyoctadecadinoic acid from linoleic acid [29–31]. Recently, Cao et al.  have, for the first time, used engineered E. coli for the production of hydroxy fatty acid. They could accumulate hydroxy fatty acids like 9-hydroxydecanoic acid, 11-hydroxydodecanoic acid, 10-hydroxyhexadecanoic acid and 12-hydroxyoctadecanoic acid through the introduction of fatty acid hydroxylase (CYP102A1) from Bacillus megaterium coupled with co-expression of the acetyl-CoA carboxylase (ACCase) and acyl-CoA thioesterase (TesA), and knockout of the endogenous acyl-CoA synthetase (FadD). This engineered E. coli strain accumulated up to 58.7 mg/L of total hydroxy fatty acids in culture broth. There are no report regarding the use of engineered E. coli for the production of 9,10-dihydroxyhexadecanoic acid. The production of 9,10-dihydroxyoctadecadienoic acid has been successfully achieved using bacterial diol synthases . Dihydroxy fatty acids with 16 carbon atoms is not reported for their production using microbial cultures. Among the dihydroxyhexadecanoids, 10,16-dihydroxyhexadecanoic acid is reported to be isolated from tomato peel and has been in use for synthesis of the 16-hydroxy-10-oxo-hexadecanoic acid (a monomer present in lime cuticle) and 7-oxohexadecanendioic acid, which are used as starting materials in the preparation of different aliphatic polyesters . Thus, the microbial synthesis of 9,10-dihydroxyhexadecanoic acid reported in the study will lead to the easy production of economically important downstream components like 9,10-dibromohexadecanoic acid, palmitelaidic acid and suberic acid. Since the approach applied in the present study has resulted in dihydroxy derivative of hexadecanoic acid as one of the major products, this system can be applied in future for the production of a variety of other dihydroxy fatty acids.
Hydroxy fatty acids are important compounds due to their wide variety of high end applications. The production of vicinal diols of fatty acids has not been reported in the literature. The current study is the first report for the production of 9,10-dihydroxy hexadecanoic acid using 9-hexadecanoic acid as substrate and three different heterologous genes. The approach used in the study demonstrated that the whole cell lysate of the induced cultures can be used for realizing the activity of the already known enzymes on new substrates. This strategy could be optimised for the production of 9,10-dihydroxyhexadecanoic acid for its large scale production. This type of combinatorial strategies could well be employed for the production of other novel hydroxy fatty acids.
AK, PS and VSB conceived and designed the experiments; AK performed the experiments. AK and PS analyzed experimental data. AK, PS and VSB wrote the manuscript. All authors read and approved the final manuscript.
The authors acknowledge the help of Mr. Ajai at Advanced Instrumentation Research Facility, Jawaharlal Nehru University, New Delhi for GC–MS analysis.
The authors declare that they have no competing interests.
Consent for publication
All authors agree to submit this work to MICF.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Kim KR, Oh DK. Production of hydroxy fatty acids by microbial fatty acid-hydroxylation enzymes. Biotechnol Adv. 2013;36:1473–85.View ArticleGoogle Scholar
- Van Bogaert INA, Groeneboer S, Saerens K, Soetaert W. The role of cytochrome P450 monooxygenases in microbial fatty acid metabolism. FEBS J. 2011;278:206–21.View ArticleGoogle Scholar
- James AT, Hadaway HC, Webb JP. The biosynthesis of ricinoleic acid. Biochem J. 1965;95:448–52.View ArticleGoogle Scholar
- Brash AR. Lipoxygenases: occurrence, functions, catalysis, and acquisition of substrate. J Biol Chem. 1999;274:23679–82.View ArticleGoogle Scholar
- Ivanov I, Heydeck D, Hofheinz K, Roffeis J, O’Donnell V, Kuhn H, Walther M. Molecular enzymology of lipoxygenases. Arch Biochem Biophys. 2010;503:161–74.View ArticleGoogle Scholar
- Capdevila JH, Falck JR, Harris RC. Cytochrome P450 and arachidonic acid bioactivation. Molecular and functional properties of the arachidonate monooxygenase. J Lipid Res. 2000;41:163–81.Google Scholar
- Sambrook J, Russell D. Molecular Cloning: A laboratory manual. New York: Cold Spring Harbor Laboratory Press; 2001.Google Scholar
- Cahoon EB, Ripp KG, Hall SE, McGonigle B. Transgenic production of epoxy fatty acids by expression of a cytochrome P450 enzyme from Euphorbia lagascae seed. Plant Physiol. 2002;28:615–24.View ArticleGoogle Scholar
- Singh A, Upadhyay V, Upadhyay AK, Singh SM, Panda AK. Protein recovery from inclusion bodies of Escherichia coli using mild solubilization process. Microb Cell Fact. 2015;14:41.View ArticleGoogle Scholar
- Jevsevar S, Gaberc-Porekar V, Fonda I, Podobnik B, Grdadolnik J, Menart V. Production of nonclassical inclusion bodies from which correctly folded protein can be extracted. Biotechnol Prog. 2005;21:632–9.View ArticleGoogle Scholar
- Ralston AW, Hoerr CW. The solubilities of the normal saturated fatty acids. J Org Chem. 1942;07:546–55.View ArticleGoogle Scholar
- Fialkov AB, Steiner U, Lehotay SJ, Amirav A. Sensitivity and noise in GC–MS: achieving low limits of detection for difficult analytes. Int J Mass Spectrom. 2007;260:31–48.View ArticleGoogle Scholar
- Von Wettstein-Knowles P. Plant waxes. In: Encyclopedia of life sciences. Chichester, UK: Wiley; 2012. p. 1–11.Google Scholar
- Kadegowda AKG, Burns TA, Miller MC, Duckett SK. Cis-9, trans-11 conjugated linoleic acid is endogenously synthesized from palmitelaidic (C16:1 trans-9) acid in bovine adipocytes. J Anim Sci. 2013;91:1614–23.View ArticleGoogle Scholar
- Mead JF, Nevenzel JC. The question of biohydrogenation of fatty acids. J Lipid Res. 1960;1:305–10.Google Scholar
- Rorrer GL, Yuan X, Huang YM, Hayden C, Gerwick WH. Production of hydroxy fatty acids by cell suspension cultures of the marine brown alga Laminaria saccharina. Phytochem. 1997;46:871–7.View ArticleGoogle Scholar
- Tian H, Lu Y, Shah SP, Hong S. 14S,21R-dihydroxydocosahexaenoic acid remedies impaired healing and mesenchymal stem cell functions in diabetic wounds. J Biol Chem. 2011;286:4443–53.View ArticleGoogle Scholar
- Metzger JO, Bornscheuer U. Lipids as renewable resources: current state of chemical and biotechnological conversion and diversification. Appl Microbiol Biotechnol. 2006;71:13–22.View ArticleGoogle Scholar
- Kuo TM, Lanser AC. Factors influencing the production of a novel compound, 7,10-dihydroxy-8(E)-octadecenoic acid, by Pseudomonas aeruginosa PR3 (NRRL B-18602) in batch cultures. Curr Microbiol. 2003;47:186–91.View ArticleGoogle Scholar
- Morvan B, Joblin KN. Hydration of Oleic Acid by Enterococcus gallinarum, Pediococcus acidilactici and Lactobacillus sp. isolated from the Rumen. Anaerobe. 1999;5:605–11.View ArticleGoogle Scholar
- Kim BN, Joo YC, Kim YS, Kim KR, Oh DK. Production of 10-hydroxystearic acid from oleic acid and olive oil hydrolyzate by an oleate hydratase from Lysinibacillus fusiformis. Appl Microbiol Biotechnol. 2012;95:929–37.View ArticleGoogle Scholar
- Joo YC, Seo ES, Kim YS, Kim KR, Park JB, Oh DK. Production of 10-hydroxystearic acid from oleic acid by whole cells of recombinant Escherichia coli containing oleate hydratase from Stenotrophomonas maltophilia. J Biotechnol. 2012;158:17–23.View ArticleGoogle Scholar
- Bae JH, Suh MJ, Kim BS, Hou CT, Lee IJ, Kim IH, Kim HR. Optimal production of 7,10-dihydroxy-8(E)-hexadecenoic acid from palmitoleic acid by Pseudomonas aeruginosa PR3. Nat Biotechnol. 2010;27:352–7.Google Scholar
- Chang IA, Bae JH, Suh MJ, Kim IH, Hou CT, Kim HR. Environmental optimization for bioconversion of triolein into 7,10-dihydroxy-8(E)-octadecenoic acid by Pseudomonas aeruginosa PR3. Appl Microbiol Biotechnol. 2008;78:581–6.View ArticleGoogle Scholar
- Culleré J, Durany O, Busquets M, Manresa A. Biotransformation of oleic acid into (E)-10-hydroxy-8-octadecenoic acid and (E)-7,10-dihydroxy-8-octadecenoic acid by Pseudomonas sp. 42A2 in an immobilized system. Biotechnol Lett. 2001;23:215–9.View ArticleGoogle Scholar
- Suh MJ, Baek KY, Kim BS, Hou CT, Kim HR. Production of 7,10-dihydroxy-8(E)-octadecenoic acid from olive oil by Pseudomonas aeruginosa PR3. Appl Microbiol Biotechnol. 2011;89:1721–7.View ArticleGoogle Scholar
- Kim BS, Kim HR, Hou CT. Effect of surfactant on the production of oxygenated unsaturated fatty acids by Bacillus megaterium ALA2. Nat Biotechnol. 2010;27:33–7.Google Scholar
- Holic R, Yazawa H, Kumagai H, Uemura H. Engineered high content of ricinoleic acid in fission yeast Schizosaccharomyces pombe. Appl Microbiol Biotechnol. 2012;95:179–87.View ArticleGoogle Scholar
- Drouet P, Thomas D, Legoy MD. Production of 13(S)-hydroperoxy-9(Z),11(E)-octadecadienoic acid using soybean lipoxygenase 1 in a biphasic octane-water system. Tetrahedron Lett. 1994;35:3923–6.View ArticleGoogle Scholar
- Iacazio G, Langrand G, Baratti J, Buono G, Triantaphylides C. Preparative, enzymic synthesis of linoleic acid (13S)-hydroperoxide using soybean lipoxygenase-1. J Org Chem. 1990;55:1690–1.View ArticleGoogle Scholar
- Villaverde JJ, vander Vlist V, Santos SAO, Haarmann T, Langfelder K, Pirttimaa M, Nyyssola A, Jylha S, Tamminen T, Kruus K, de Graaff L, Neto CP, Simoes MMQ, Domingues MRM, Silvestre AJD, Eidner J, Buchert J. Hydroperoxide production from linoleic acid by heterologous Gaeumannomyces graminis tritici lipoxygenase: optimization and scale-up. Chem Eng J. 2013;217:82–90.View ArticleGoogle Scholar
- Cao Y, Cheng T, Zhao G, Niu W, Guo J, Xian M, Liu H. Metabolic engineering of Escherichia coli for the production of hydroxy fatty acids from glucose. BMC Biotechnol. 2016;16:26.View ArticleGoogle Scholar
- Lang I, Göbel C, Porzel A, Heilmann I, Feussner I. A lipoxygenase with linoleate diol synthase activity from Nostoc sp. PCC 7120. Biochem J. 2008;410:347–57.View ArticleGoogle Scholar
- Arrieta-Baez D, Cruz-Carrillo M, Gmez-Patio MB, Zepeda-Vallejo LG. Derivatives of 10,16-dihydroxyhexadecanoic acid isolated from tomato (Solanum lycopersicum) as potential material for aliphatic polyesters. Molecules. 2011;20116:4923–36.View ArticleGoogle Scholar