Selectivity of lipases for estolides synthesis

Anamaria Todea, Linda G. Otten 3 , August E. Frissen 2 , Isabel W.C.E. Arends 3 , Francisc Peter 1  and Carmen G. Boeriu 2
  • 1 Faculty of Industrial Chemistry and Environmental Engineering, University Politehnica of Timişoara, C. Telbisz 6, 300001 Timişoara, Romania
  • 2 Wageningen UR Food and Biobased Research, Bornse Weilanden 9, 6708 WG Wageningen, The Netherlands
  • 3 Biotechnology Department, Biocatalysis group, Delft University of Technology, Julianalaan 136, 2628 BL Delft, The Netherlands
Anamaria Todea, Linda G. Otten, August E. Frissen, Isabel W.C.E. Arends, Francisc Peter and Carmen G. Boeriu

Abstract

Lipase-catalyzed synthesis of estolides starting from different saturated (C16 16OH, C18 12OH) and unsaturated (C18:1 9 cis 12-OH) hydroxy-fatty acids was investigated. For this reason, the catalytic efficiency of several native and immobilized lipases in different organic reaction media at temperatures up to 75 °C was studied. The formation of mono- and di-lactone as well as estolide’s chain elongation depends on the type and source of lipase. The lipase from Pseudomonas stutzeri immobilized by cross-linking as cross-linked enzymes aggregates (CLEAs) was the best biocatalyst in terms of chain elongation. Estolides with polymerization degree up to 10 were obtained at substrate conversions higher than 80 %.

Introduction

Hydroxy fatty acids (HFA) have been intensively investigated due to their many applications especially in cosmetics, paints, lubricants, food industry, medicine as well as intermediates in the synthesis of fine chemicals and pharmaceuticals [1, 2]. For example, 12-hydroxystearic acid is mainly used as a thickening agent for the manufacture of lubricating greases, ricinoleic acid is used in soaps manufacturing, as viscosity regulator agent and as emulsifier in margarine, and 9-hydroxystearic acid has been reported as stereo-selective inhibitor against human histone deacetylase I (colon cancer) [3].

Polyesters based on hydroxy fatty acids from vegetable oils, called estolides, present a variety of potential applications due to their biodegradability and improved characteristics including the higher thermal oxidative stability compared to the vegetable oils [4]. The chemical route for estolides synthesis involves high temperatures (205–210 °C) or strong acids as catalysts that can lead to colored and malodorous products, together with undesired products that are not particularly suitable for pharma and food industry and would require supplementary costs for purification [5].

The activity of lipases towards HFA in the esterification/transesterification reactions is well established [6, 7]. One of the most investigated HFA is 12-hydroxy-9-cis-octadecenoic acid, known as ricinoleic acid that is available in large amounts in castor oil. Waxes esters based on ricinoleic acid were synthesized using lipases from different sources, such as Rhizomucor miehei (R. miehei) [7, 8]. The lipase from Candida rugosa (C. rugosa) was used for the synthesis of estolides of ricinoleic acid, having possible applications as a viscosity regulator agent and an emulsifier in margarine [9]. Estolides formation starting from ricinoleic acid have been successfully synthesized by using non-specific lipases such as C. rugosa, Chromobacterium viscosum (C. viscosum), Pseudomonas sp, Geothricum candidum (G. candidum) [10]. Less information is available about the activity of lipases towards estolides formation and the products formed upon reaction.

16-Hydroxyhexadecanoic acid (16HHDA) has been previously reported as substrate for several lipases, but the studies were focused only on the ability of different lipases, from Ps. species, porcine pancreas [11] and C. viscosum [12, 13] to synthesize monolactones. Even though the estolides or dilactones have been considered as possible reaction products based on TLC analysis their formation was not proved [14]. In this work, we accomplished a more comprehensive study, by extending the number of the tested lipases (including, lipase from T. lanuginosus, P. stutzeri, R. arrhizus, and P. cepacia) and identifying all individual products formed in the reaction besides monolactones.

The main objectives of the present work were:

  • To compare the selectivity of several native lipases for three structurally different hydroxy fatty acids: 12-hydroxy-9-cis-octadecenoic acid (ricinoleic acid, RCA), 12-hydroxyoctadecanoic acid (12HSA) and 16-hydroxyhexadecanoic acid (16HHDA).
  • To optimize the estolides synthesis, by studying the influence of the solvent, temperature, and substrate concentration.
  • To fully characterize the reaction products by different mass spectrometry techniques, to distinguish between cyclic and linear esters, chain length and dispersity.

Materials and methods

Materials

12-Hydroxystearic acid (99 %), 16-hydroxyhexadecanoic acid (98 %), ricinoleic acid (98 %), Bis(trimethylsilyl)trifluoroacetamide-trimethylchlorosilane (BSTFA + TMCS = 99:1), trans-2-[3-(4-t-butyl-phenyl)-2-methyl-2-propenylidene]malononitrile (DCTB), potassium trifluoroacetate (KTFA), Amano lipase from Pseudomonas fluorescens (P. fluorescens), Pseudomonas cepacia (P. cepacia), Thermomyces lanuginosus (T. lanuginosus), and CLEA Alcalase were purchased from Sigma Aldrich. The lipases from C. rugosa, Aspergillus oryzae (A. oryzae), Candida antarctica A (C. antarctica A), Rhizopus arrhizus (R. arrhizus), Penicillium roqueforti (P. roqueforti), Mucor javanicus (M. javanicus), Candida lipolytica (C. lipolytica), Thermomyces brockii (T. brockii), and hog pancreas were obtained from Fluka. Novozyme 435, Lipozyme TL, Candida antarctica B (C. antarctica B) lipase were purchased from Novozymes. The lipases from Alcaligenes sp (lipase TL), Pseudomonas stutzeri (P. stutzeri) (Lipase TL), were products of Meito Sangyo Co. Ltd., Japan. The CLEA from C. antarctica B and P. stutzeri were purchased from CLEA Technologies. The organic solvents used (heptane, toluene, cyclohexane, methyl ethyl ketone and N,N-dimethyl formamide) were from Merck and their water content (determined by Karl Fischer titration) was below 0.01 %.

Lipases activity assay

The hydrolytic activity of lipases was determined using p-nitro-phenyl palmitate as substrate, as reported elsewhere [15].

Biocatalytic synthesis of estolides

The reactions were performed by adding native and immobilized lipases, 50 U/mmol substrate, to different amounts of substrate (1–5 mM), brought to a final volume of 1 mL in the selected organic solvent. The reactions have been carried out at temperatures between 40 °C and 75 °C, while stirring at 350 rpm for 24 h. At the end of the reaction the enzyme was removed by centrifugation (13 000 rpm, 3 min), and the reaction mixture was analyzed by GC-MS and MALDI-TOF MS.

The HFA conversion was determined based on the GC-MS analysis, after derivatization with BSTFA + TMCS (99:1), at 2:1 reagent: sample ratio (w/w), for 1 h at 95 °C, as previously described [16]. Hexadecane was used as internal standard. The separation was carried out using an Interscience Trace GC Ultra GC+ PTV instrument, equipped with AS3000 II autosampler and Restek Rxi-5ms 30 m × 0.25 mm × 0.25 μm capillary column. The analysis conditions were: oven temperature 100–300 °C with 10 °C/min heating rate, injector temperature 300 °C, carrier gas (helium) flow 1.0 mL/min. Mass spectra were obtained from Interscience Trace DSQ II XL quadrupole mass selective detector (EI, mass range 35–500 Dalton, 150 ms sampling speed), the mass spectrometer being operated at 70 eV. For the calculation of conversions at a defined reaction time, calibration curves were built up for each substrate in the range from 0.05 mM to 3 mM, using hexadecane as internal standard.

MALDI-TOF MS analysis was carried out for composition analysis of estolides, using an Ultraflex Workstation with Flex Control and Flex Analysis software packages for acquisition and processing of the data (Bruker Daltonics, Germany), at an acceleration voltage of 25 kV, using DCTB as matrix and KTFA as ionization agent. 10 μL of the sample was mixed with 10 μL of matrix solution (40 mg/mL DCTB solubilized in THF) and 3 μL of KTFA (5 mg/mL). About 0.3 μL of the mixture have been applied on the plate and measured in the positive mode. Detection was performed in the reflector mode. The number average molecular weight Mn, weight average molecular weight Mw and the polydispersity index PDI = Mw/Mn have been calculated as described elsewhere [17].

Results and discussion

The HFA obtained from natural sources or from their corresponding fatty acids could be valuable starting materials for bio-products with a variety of applications. In this work we have chosen three structurally different HFAs, namely RCA, the main component of castor oil comprising a cis double bond, 12HSA, the stearic acid derivative normally obtained from castor oil upon hydrogenation [18] and 16HHDA, an ω-hydroxyl derivative of palmitic acid, produced by ω-hydroxylation of palmitic acid by cytochrome P450 in plants and animals [19]. Upon the catalytic action of lipases, HFA can be involved in intra- and intermolecular esterification with formation of cyclic and (branched) open chain esters, as illustrated in Fig. 1.

Fig. 1
Fig. 1

Synthesis of macrolactones/estolides based on hydroxy fatty acids, catalyzed by lipases.

Citation: Pure and Applied Chemistry 87, 1; 10.1515/pac-2014-0716

Specificity of lipases in the synthesis of estolides

The substrate specificity study of lipases was carried out to establish the differences in terms of product formationand substrate selectivity of enzymes from different sources. The esterification reactions of HFA were performed by using the same enzyme activity related to the substrate, i.e., 50 units/mmol substrate, in every reaction. In order to monitor the monolactone formation, the reactions have been performed at 1 mM substrate concentration, since previous works showed that lipase-catalyzed synthesis of cyclic esters is favored by lower substrate concentrations [11, 20]. The temperature was set at 40 °C, the optimum temperature for most lipases, using toluene as reaction media. After 24 h reaction time, the processes were stopped by centrifugation of the enzyme and the samples were derivatized and analyzed by GC-MS, as described in the Methods section. The GC-MS results together with the qualitative analysis performed by MALDI-TOF MS are presented in Table 1. Unexpectedly, monolactones were not identified in all reaction products (Table 1). The formation of the monolactone (easily monitored by GC-MS) was observed for all three tested substrates only when the lipases from P. fluorescens and P. stutzeri were used as catalyst. For the substrate bearing primary OH group, 16HHDA the monolactone was formed also by P. cepacia and Alcaligenes PL catalysis. The chain elongation for 16HHDA was favored by the lipases from P. cepacia, C. antarctica B, hog pancreas and P. stuzeri when the presence of the species as trimer and tetramer were identified in the MALDI-TOF MS spectra.

Table 1

Lipase ability to catalyze lactonization/oligomerization reaction of 12HSA, 16HHDA and RCA in the presence of toluene as solvent and 50 U enzyme/mmol substrate.

Lipase sourceConversion, % (GC-MS)/Identified product (GC-MS/MALDI TOF)
12HSA16HHDARCA
Alcaligenes PL52.3D62ML+D0n.d.
A. oryzae0n.d.9.8DL25D
C. antarctica A0n.d.0n.d15.1D
C. antarctica B27.5D91.6D+T+Tr33.2D
C. lipolytica9.1D0n.d.0n.d.
C. rugosa0n.d.0n.d.25.5DL
M. javanicus0n.d.12.5DL10.1D
P. cepacia0n.d.54.5ML+D+T6.6D
P. fluorescens15.2ML+D+T91.6ML+DL31.5ML+D+T
P. stutzeri32.8ML+D62.1ML+D+T10ML+D
P. roqueforti0n.d.0n.d.16.6D
R. arrhizus0n.d.48.9DL10.9D
T. brockii7.7D11.5D0n.d.
T. lanuginosus0n.d.91.6DL20.1D
Hog pancreas0n.d.91.6D+T+Tr16.5D

n.d., not determined; ML, monolactone; DL, dilactone; D, dimer; T, trimer; Tr, tetramer.

The highest conversion was achieved for 16HHDA. These results are in accordance with the results reported by Makita et al. [11], and can be explained by the regioselectivity of lipases for the primary alcohol groups, compared to the secondary alcohol groups due to the size of the active site and the orientation of the active site Ser in the middle. Secondary alcohols need to be bend to fit into the active site resulting in less degrees of freedom, and therefore more difficult binding. There were 5 lipases that led to singular products, the most interesting were the lipases from R. arrhizus and T. lanuginosus, that were able to form the dilactone with conversions higher than 40 %. The native lipase from C. antarctica B also showed to be promising for estolide synthesis, particularly from 16HHDA.

In the case of C18 substrates (12HSA and RCA), important differences were noticed. Even though each of these substrates contains a secondary OH group in the same position (C12), several lipases, such as C. rugosa, A. oryzae, M. javanicus, C. antarctica A, R. arrhizus, P. roqueforti, P. cepacia, T. langinosus and hog pancreas were not at all active against the saturated substrate (12HSA), but led to conversions up to 35 % for ricinoleic acid. Moreover, there were lipases that presented no activity for RCA, namely C. lipolytica, Alcaligenes PL and T. brockii, but led to 12HSA based monoestolide (referred as dimer D in Table 1) formation. Another important conclusion for these two substrates is that monoestolide was synthesized as unique product by all tested lipases, excepting those from Pseudomonas.

Even though such substrates have been already tested, especially ricinoleic acid, alone [10] or in combination with different polyols or fatty acids [21], there are no reports concerning the selectivity of several commercially available lipases, as we performed in this study. Moreover, in previous reports substrate conversion or chain elongation have been monitored solely by titration [9] or thin layer chromatography [14]. In our study, the MALDI-TOF technique allowed the identification of all the species present in the reaction mixture.

Effect of temperature and substrate concentration

To investigate the effect of temperature and substrate concentration on conversion, the concentration of the substrate has been increased 5 times, up to 5 mM, and the temperature raised up to 75 °C.

As preliminary experiments for the thermal stability study several lipases, selected according to their capability to favor the chain elongation reaction and/or higher conversion, were incubated up to 24 h, at temperatures up to 80 °C (Figure 1S). Excepting the lipase from T. lanuginosus the thermal stabilities were excellent, as they preserved more than 85 % of the initial hydrolytic activities.

The results presented in Fig. 2 indicate that the conversions were about 2-fold higher when the temperature and substrate concentration have been increased for 12HSA and RCA (excepting CalB). Nevertheless, the chain elongation for the C18 substrates was favored only by the lipases from Ps. species (data not shown). Because the lipase from P. fluorescens proved to be efficient for all three substrates, further experiments were performed with this lipase.

Fig. 2
Fig. 2

Influence of temperature and substrate concentration article image 1 mM 40 °C, article image 5 mM 75 °C on the conversion of HFA catalyzed by lipases from P. fluorescens, P. stutzeri and C. antarctica B.

Citation: Pure and Applied Chemistry 87, 1; 10.1515/pac-2014-0716

Influence of the reaction medium

Solvents with different polarity spanning logP [22] values between –1.0 and 4.3 were investigated to disclose the influence of the reaction medium polarity on the formation of estolides. The reactions were performed at 75 °C, 5 mM substrate concentration, using as catalyst 50 U/mmol substrate lipase from P. fluorescens. The results presented in Table 2 indicate that the substrate conversion and estolide formation were favored by the increase of the logP value of the solvent, meaning decreasing polarity. The results are in concordance with previous reports, where apolar solvents have been successfully used for the synthesis of monolactones [11].

Table 2

Influence of reaction medium on the total conversion of HFA and the formation of oligomers, catalyzed by P. fluorescens lipase.

SolventLogP [7] Conversion, % (GC-MS)/Identified product (MALDI TOF)
16HHDA12HSARCA
N,N-dimethyl formamide–1.07.6n.d.0.0n.d.0.0n.d.
Methyl ethyl ketone0.30.0n.d.0.0n.d.0.0n.d.
Toluene2.592.8D+T+Tr55.5ML+D65.7ML+D+T
Cyclohexane3.291.7D+T+Tr80.1D+T+Tr25.5D+DL
Heptane4.3n.c.ML+D+…n.c.ML+D+…Hn.c.D+DL

n.d., not determined; n.c., calculated; D, dimer (monoestolide); DL, dilactone; T, trimer; Tr, tetramer; P, pentamer; H, hexamer.

Products were not detected when the most polar solvents, N,N-dimethyl formamide and methyl ethyl ketone, were used as reaction medium. The highest conversion values were achieved, as expected, for the 16HHDA substrate, due to the high selectivity of the lipase for the primary OH group, as previously discussed. Even though the chain elongation was favored by heptane, leading to oligomers with polymerization degrees up to 6, identified in the MALDI-TOF MS spectrum, the solubilization of the substrate into heptane was not complete. Consequently, a real conversion in this solvent could not be calculated. The higher boiling point of toluene allows more elevated reaction temperatures, which can increase the reaction rate. Considering both conversion values and formation of products with higher chain elongation, toluene can be considered the best reaction medium to produce estolides.

Synthesis of estolides by immobilized lipases

In order to compare the catalytic efficiency and selectivity for the chain elongation reaction of different immobilized enzymes in estolides synthesis, several commercially available immobilized lipases and one protease have been tested. The reactions were performed as previously described, using 5 mM substrate concentration, toluene as solvent and 24 h reaction time at 75 °C. CLEA lipases from P. stutzeri (CLEA-P. stutzeri) C. antarctica B (CLEA CalB), CLEA Alcalase, as well as T. lanuginosus lipase immobilized on granulated silica (Lipozyme TL) and C. antarctica immobilized on acrylic resin (Novozyme 435) were tested. Mn and Mw values were calculated based on the MALDI-TOF MS spectra, as described elsewhere [17]. The results indicate that although conversions higher than 70 % were obtained for 16HHDA, the higher polymerization degree (up to 10) was achieved when RCA was used as substrate and CLEA- P. stutzeri lipase as catalyst (Table 3).

Table 3

The results obtained for HFA based reaction product when different immobilized lipases have been used as catalyst.

SubstrateLipozyme
MnMwPDIDPmax
16HHDA98610401.055
12HSA6596591.002
RCA6306311.002
SubstrateNovozyme
MnMwPDIDPmax
16HHDA7147791.095
12HSA6596591.002
RCA6556551.002
SubstrateCLEA CalB
MnMwPDIDPmax
16HHDA106711901.119
12HSA6596591.002
RCA6556551.002
SubstrateCLEA P. stutzeri
MnMwPDIDPmax
16HHDA96610341.077
12HSA7497951.067
RCA114013261.1610
SubstrateCLEA Alcalase
MnMwPDIDPmax
16HHDA5956121.023
12HSA6596591.002
RCA6556551.002

DPmax, maximal polymerization degree achieved in the reaction conditions; PDI = Mw/Mn.

The formation of estolides with polymerization degree ranging from 2 to 10 could be observed in the MALDI-TOF MS spectrum of the RCA-based reaction product (as potassium adduct), as presented in Fig. 3. Lower molecular weight estolides were previously reported for ricinoleic acid as substrate when the immobilized lipase from Staphylococcus xylosus has been used [23].

Fig. 3
Fig. 3

MALDI-TOF MS spectrum of ricinoleic-based estolides catalyzed by CLEA- P. stutzeri lipase.

Citation: Pure and Applied Chemistry 87, 1; 10.1515/pac-2014-0716

CLEA P. stutzeri lipase was the most efficient lipase with respect to chain elongation, for all three substrates tested. In the case of 12HSA substrate, formation of the monoestolide was observed in most of the cases, while formation of the monoestolide together with the dilactone was favored by the CLEA- P. stutzeri lipase. However, considering the highest selectivity of lipase for the dimer formation without any by-product and the usefulness of the monoestolide for cosmetic applications [24], this synthesis route is very promising, particularly in the perspective of further development for reuse of the catalyst and operation in continuous systems.

Interesting results have been obtained with CLEA Alcalase as well, proving the ability of this enzyme to use HFA as substrates for estolides synthesis in good yields, even though it is a protease.

Conclusion

Estolides based on C16 and C18 hydroxy fatty acids, having the Mn molecular weights in the range 795–1326, were synthesized successfully by native and immobilized enzymes. Based on the overall conversion and the product profile, the selectivity of lipases for the tested substrates decreased in the following order:

C16(16OH)>C18(12OH:9)>C18(12OH).

The chain elongation was favored by nonpolar solvents. The results obtained allow the selection of the best commercially available enzyme to obtain a required product.

Acknowledgments

The authors are grateful to Dr. Maurice Franssen for the CLEA lipases. This work was partially supported by the strategic grant POSDRU/159/1.5/S/137070 (2014) of the Ministry of National Education, Romania, co-financed by the European Social Fund – Investing in People, within the Sectoral Operational Programme Human Resources Development 2007–2013.

References

  • [1]

    H. M. Kim, H. R. Kim, C. T. Hou, B. S. Kim. J. Am. Oil Chem. Soc. 87, 1451 (2010).

  • [2]

    I. Martin-Arjol, M. Busquets, A. Manresa. Process Biochem. 48, 224 (2013).

  • [3]

    K.R. Kim, D. K. Oh. Biotechnol. Adv. 31, 1473 (2013).

  • [4]

    T. A. Isbell. Grasas Aceites, 62, 8 (2011).

  • [5]

    E. C. G. Aguieiras, C. O. Veloso, J. V. Bevilaqua, D. O. Rosas, M. A. P. da Silva, M.A. P. Langone. Enzyme Res. 20, 1 (2011).

  • [6]

    D. G. Hayes. J. Am. Oil Chem. Soc. 81, 1077 (2004).

    • Crossref
  • [7]

    M. Ghosh, D. K. Bhattacharyya. J. Am. Oil Chem. Soc. 75, 1057 (1998).

  • [8]

    D. Mukesh, R. S. Iyer, J. S. Wagh, A. A. Mokashi, A. A. Banerji, R. V. Newadkar, H. S. Bevinakatti. Biotechnol. Lett. 15, 251 (1993).

  • [9]

    A. Bodalo-Santoyo, J. Bastida-Rodriguez, M. F. Maximo-Martin, M. C. Montiel-Morte, M. D. Murcia-Almagro. Biochem. Eng. J. 26, 155 (2005).

  • [10]

    A. Bodalo, J. Bastida, M. F. Maximo, M. C. Montiel, M. Gomez, M. D. Murcia. Biochem. Eng. J. 39, 450 (2008).

  • [11]

    A. Makita, T. Nihira, Y. Yamada. Tetrahedron Lett. 28, 805 (1987).

  • [12]

    S. Matsumura, J. Takahashi. Makromol. Chem. Rapid Commun. 7, 369 (1986).

  • [13]

    M. J. Alston, R. B. Freedman. Biotechnol. Bioeng. 77, 641 (2002).

  • [14]

    G. K. Robinson, M. J. Alston, C. J. Knowles, P. S. J. Cheetham, K. R. Motion. Enzyme Microb. Technol. 16, 855 (1994).

  • [15]

    N. D. Mahadik, U. S. Puntambekar, K. B. Bastawde, J. M. Khire, D. V. Gokhale. Process Biochem. 38, 715 (2002).

  • [16]

    J. A. Hudson, C. A. M. MacKenzie, K. N. Joblin. Appl. Microbiol. Biotechnol. 44, 1 (1995).

    • PubMed
  • [17]

    H. J. Rader, W. Schrepp. Acta Polym. 49, 272 (1998).

  • [18]

    A. K. Maskaev, N. K. Mankovskaya, I. V. Lendel,V. T. Fedorovskii, E. I. Simurova,V. N. Terenteva. Chem. Technol. Fuels Oils 7, 109 (1971).

    • Crossref
  • [19]

    I. Benveniste, T. Saito, Y. Wang, S. Kandel, H. Huang, F. Pinot,R. A. Kahn, J. P. Salaun, M. Shimoji. Plant Sci. 170, 326 (2006).

  • [20]

    U. Antczak, J. Gora, T. Antczakt, E. Galas. Enzyme Microb. Technol. 13, 589 (1991).

  • [21]

    D. G. Hayes, V. K. Mannam, R. Ye, H. Zhao, S. Ortega, M. C. Montiel. Polymers 4, 1037 (2012).

    • Crossref
  • [22]

    C. M. Du, K. Valko, C. Bevan, D. Reynolds, M. H. Abraham. J. Chromatogr. Sci. 38, 503 (2000).

  • [23]

    H. Horchani, A. Bouaziz, Y. Gargouri, A. Sayari. J. Mol. Catal. B: Enzym. 75, 35 (2012).

  • [24]

    S. A. Madison, T. Moaddel, B. Harichian, J. G. Rosa, H. Meldrum, J. Lee. U. S. Patent 2012/0122936 A1, filed 11 Nov 2010, issued 17 May 2012.

Footnotes

Supplemental Material

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  • [1]

    H. M. Kim, H. R. Kim, C. T. Hou, B. S. Kim. J. Am. Oil Chem. Soc. 87, 1451 (2010).

  • [2]

    I. Martin-Arjol, M. Busquets, A. Manresa. Process Biochem. 48, 224 (2013).

  • [3]

    K.R. Kim, D. K. Oh. Biotechnol. Adv. 31, 1473 (2013).

  • [4]

    T. A. Isbell. Grasas Aceites, 62, 8 (2011).

  • [5]

    E. C. G. Aguieiras, C. O. Veloso, J. V. Bevilaqua, D. O. Rosas, M. A. P. da Silva, M.A. P. Langone. Enzyme Res. 20, 1 (2011).

  • [6]

    D. G. Hayes. J. Am. Oil Chem. Soc. 81, 1077 (2004).

    • Crossref
  • [7]

    M. Ghosh, D. K. Bhattacharyya. J. Am. Oil Chem. Soc. 75, 1057 (1998).

  • [8]

    D. Mukesh, R. S. Iyer, J. S. Wagh, A. A. Mokashi, A. A. Banerji, R. V. Newadkar, H. S. Bevinakatti. Biotechnol. Lett. 15, 251 (1993).

  • [9]

    A. Bodalo-Santoyo, J. Bastida-Rodriguez, M. F. Maximo-Martin, M. C. Montiel-Morte, M. D. Murcia-Almagro. Biochem. Eng. J. 26, 155 (2005).

  • [10]

    A. Bodalo, J. Bastida, M. F. Maximo, M. C. Montiel, M. Gomez, M. D. Murcia. Biochem. Eng. J. 39, 450 (2008).

  • [11]

    A. Makita, T. Nihira, Y. Yamada. Tetrahedron Lett. 28, 805 (1987).

  • [12]

    S. Matsumura, J. Takahashi. Makromol. Chem. Rapid Commun. 7, 369 (1986).

  • [13]

    M. J. Alston, R. B. Freedman. Biotechnol. Bioeng. 77, 641 (2002).

  • [14]

    G. K. Robinson, M. J. Alston, C. J. Knowles, P. S. J. Cheetham, K. R. Motion. Enzyme Microb. Technol. 16, 855 (1994).

  • [15]

    N. D. Mahadik, U. S. Puntambekar, K. B. Bastawde, J. M. Khire, D. V. Gokhale. Process Biochem. 38, 715 (2002).

  • [16]

    J. A. Hudson, C. A. M. MacKenzie, K. N. Joblin. Appl. Microbiol. Biotechnol. 44, 1 (1995).

    • PubMed
  • [17]

    H. J. Rader, W. Schrepp. Acta Polym. 49, 272 (1998).

  • [18]

    A. K. Maskaev, N. K. Mankovskaya, I. V. Lendel,V. T. Fedorovskii, E. I. Simurova,V. N. Terenteva. Chem. Technol. Fuels Oils 7, 109 (1971).

    • Crossref
  • [19]

    I. Benveniste, T. Saito, Y. Wang, S. Kandel, H. Huang, F. Pinot,R. A. Kahn, J. P. Salaun, M. Shimoji. Plant Sci. 170, 326 (2006).

  • [20]

    U. Antczak, J. Gora, T. Antczakt, E. Galas. Enzyme Microb. Technol. 13, 589 (1991).

  • [21]

    D. G. Hayes, V. K. Mannam, R. Ye, H. Zhao, S. Ortega, M. C. Montiel. Polymers 4, 1037 (2012).

    • Crossref
  • [22]

    C. M. Du, K. Valko, C. Bevan, D. Reynolds, M. H. Abraham. J. Chromatogr. Sci. 38, 503 (2000).

  • [23]

    H. Horchani, A. Bouaziz, Y. Gargouri, A. Sayari. J. Mol. Catal. B: Enzym. 75, 35 (2012).

  • [24]

    S. A. Madison, T. Moaddel, B. Harichian, J. G. Rosa, H. Meldrum, J. Lee. U. S. Patent 2012/0122936 A1, filed 11 Nov 2010, issued 17 May 2012.

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Pure and Applied Chemistry is the official monthly Journal of the International Union of Pure and Applied Chemistry (IUPAC), with responsibility for publishing works arising from those international scientific events and projects that are sponsored and undertaken by the Union.

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    Synthesis of macrolactones/estolides based on hydroxy fatty acids, catalyzed by lipases.

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    Influence of temperature and substrate concentration article image 1 mM 40 °C, article image 5 mM 75 °C on the conversion of HFA catalyzed by lipases from P. fluorescens, P. stutzeri and C. antarctica B.

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    MALDI-TOF MS spectrum of ricinoleic-based estolides catalyzed by CLEA- P. stutzeri lipase.