Lipase Activity of Tropical Oilseed Plants for Ethyl Biodiesel Synthesis

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Lipase activity of tropical oilseed plants for ethyl biodiesel synthesis and their typo-and regioselectivity Paul Alain Nanssou Kouteu, Bruno Baréa, Nathalie Barouh, Joel Blin, and Pierre Villeneuve J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.6b03674 • Publication Date (Web): 31 Oct 2016 Downloaded from http://pubs.acs.org on November 1, 2016

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Journal of Agricultural and Food Chemistry

Lipase activity of tropical oilseed plants for ethyl biodiesel synthesis and their typo-and regioselectivity Paul A. Nanssou Kouteu†‡, Bruno Barea§, Nathalie Barouh§, Joël Blin§†, Pierre Villeneuve§*.

†Institut International d’Ingénierie de l’Eau et de l’Environnement (2iE), Laboratoire Biomasse Energie et Biocarburants (LBEB), Rue de la Science, 01 BP 594, Ouagadougou 01, Burkina Faso. ‡Montpellier SupAgro, UMR 1208 Ingénierie des Agro-polymères et Technologies Emergentes, 2 Place Viala, F-34060 Montpellier, France. §

Centre de Coopération Internationale en Recherche Agronomique pour le Développement

(CIRAD), 73 rue Jean-François Breton, 34393 Cedex 5 Montpellier, France.

*Corresponding Author: Pierre Villeneuve, Phone: +33 (0) 4 99 61 20 29 Email: [email protected]

1 ACS Paragon Plus Environment


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ABSTRACT

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The aim of this work was to investigate lipase activities in crude extracts from Adansonia

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suarezensis, Adansonia grandidieri, Moringa drouhardii, Moringa oleifera, Jatropha

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mahafalensis and Jatropha curcas seeds in ethanolysis and hydrolysis reactions. All crude

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extracts from germinated seeds showed both ethanolysis and hydrolysis activities. The

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influence of germination, of the delipidation procedure and of the triacylglycerol: ethanol

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molar ratio on their ethanolysis activity was studied. Crude extracts of Jatropha and

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Adansonia seeds showed optimal activity at pH 8 with an optimum temperature of 30 °C and

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40 °C, respectively. The study of the regioselectivity of crude extracts from J. mahafalensis

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and A. grandidieri seeds, which had the most active hydrolysis reaction, showed 1,3

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regioselectivity in the hydrolysis reaction of vegetable oils. The crude extract from A.

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grandidieri seeds showed no typoselectivity, whereas the typoselectivity of the crude extract

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of J. mahafalensis seeds depended on the type of reaction.

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KEYWORDS Seed lipases, Ethanolysis, Biodiesel, Regioselectivity, Typoselectivity


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INTRODUCTION

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Lipases, triacylglycerol acylhydrolases (E.C.3.1.1.3), are enzymes that catalyze the hydrolysis

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of ester bonds in triacylglycerols to glycerol and the corresponding free fatty acids. As lipases

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are essential to living bodies for the hydrolysis of biological lipids, they are ubiquitous and

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can be isolated from many sources including microorganisms, animals and plants1–3. To date,

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a very large majority of lipases used in biotechnologies are of microbial origin3,4. However,

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they are produced using complex biotechnological processes that are not mastered all over

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the globe1,2, 4. For these reasons, the study and use of plant lipases for the modification of

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lipids have many advantages because of the wide availability of plants and their simplicity of

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use as crude enzyme extracts. Apart from the hydrolysis reaction, lipases can catalyze

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reactions such as esterification, inter-esterification and transesterification2,3. Today, industrial

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production of biodiesel by transesterification is mainly by chemical catalysis6,7. Increasing

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interest has been shown in the enzymatic production of biodiesel, because of the advantages

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they have over conventional chemical catalysts5–7. Using lipases as biocatalysts in such

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reactions enables the use of mild reaction conditions and facilitates recovery of glycerol4–7. To

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date, very little research has focused on plant lipases for the transesterification of vegetable

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oils, particularly when ethanol is used as alcohol, which is even more advantageous, because

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ethanol can be produced through fermentation of plant biomass, whereas methanol is toxic

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and is produced from petroleum.

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Plant lipases can be found in leaves, stems, latex, oilseeds, etc.3 Crude enzyme extracts can be

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obtained directly from oil cake produced by oil mills. The lipase activity of crude extracts

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from these seeds is mostly influenced by seed germination although some crude extracts from

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dormant seeds have been shown to be active2,3,5. The preparation and characterization of

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crude lipase extracts from seeds require drying, grinding and delipidation2,5 of the seed kernel.

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Delipidation is a key step5 to remove all endogenous lipids in the seed capable of interfering 3 ACS Paragon Plus Environment


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in the evaluation of hydrolytic activity of the extracts. Depending on the procedure and on the

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nature of the seed, the delipidation step can increase or decrease the activity of the crude

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extract5. It is thus important not only to evaluate lipase activity in crude extracts, but also to

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find the best way to prepare the crude extracts to obtain the highest activity possible.

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The aim of this work was to study potential lipase activity in ethanolysis and hydrolysis

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reactions of crude extracts from Adansonia suarezensis, Adansonia grandidieri, Moringa

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oleifera, Moringa drouhardii, Jatropha mahafalensis and Jatropha curcas seeds. The

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influence of the state of the seed (germinated or dormant), the mode of delipidation, and the

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triacylglycerol: ethanol molar ratio on ethanolysis activity were analyzed in the different

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extracts. The most active extracts in hydrolysis were selected to study their regioselectivity

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with different vegetable oils and their chain-length selectivity in a mixture of triacylglycerols

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or fatty acid ethyl esters at their optimum pH and temperature.

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MATERIALS AND METHODS

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Materials. The seeds were carefully selected based on quality. M. oleifera and J. curcas seeds

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were obtained from a plantation belonging to Belwet Biofuels SA in Ouagadougou (Burkina

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Faso) in May 2014, J. mahafalensis, A. grandidieri, A. suarezensis and M. drouhardii seeds

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from PhileoL in the region of Androy (Madagascar) during the same period. The proximal

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composition of these seeds is listed in table 1. Sunflower, olive, palm and groundnut oils were

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purchased in a supermarket in Montpellier (France). Tristearin, tripalmitin, trimyristin,

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trilaurin, tricaprin and ethyl esters of capric, lauric, myristic, palmitic and stearic acids (all

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>99% purity) were purchased from Sigma-Aldrich (Saint Quentin, France). High-

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performance thin layer chromatography silica plates (HPTLC, 20 × 10 cm, silica gel 60 F254)

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were purchased from Merck (Darmstadt, Germany). All other chemicals and solvents used in

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this study were of analytical or higher grade and were purchased from Sigma Aldrich (Saint

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Quentin, France). 4 ACS Paragon Plus Environment

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Preparation of crude lipase extract: J. mafahalensis, J. curcas, M. oleifera, M. drouhardii,

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A. grandidieri and A. suarezensis seeds were used to prepare crude lipase extracts. Each type

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of seed was divided into two batches (Figure 1). The seeds in one batch were germinated

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before being used to prepare crude lipase extracts, the seeds in the other batch were dormant

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(non germinated).

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Seed Germination: Before germination, all the seeds were thoroughly cleaned and left to

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steep at room temperature in a beaker of water for six hours (Figure 1). A. grandidieri and A.

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suarezensis seeds were left to steep for two days because of the hardness of their coats. For

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germination, 1 kg of each variety was spread out in a single layer between two moistened

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sheets of paper in a thermostatically controlled oven (25 °C) with controlled humidity (90%)

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(Fischer Scientific, Memmert, HPP108L, Germany).

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Preparation of non-delipidated crude enzyme extracts. The seeds harvested four days after

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germination and the dormant seeds were dehulled by hand, then crushed in an electric

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propeller grinder (Waring, France). All the different crushed seeds were dried under vacuum

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at 35 °C in an oven (Bioblock Scientific, 45001, France) for 24 h. They were then sieved and

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particles less than 635 µm in size were collected and defined as non-delipidated crude extract

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(Figure 1). They were stored in sealed jars at 4 ° C until use.

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Preparation of delipidated crude lipase extracts. Delipidation was carried out using

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acetone or hexane as solvent. Acetone powders were prepared according to the protocol

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described by Hassanien and Mukherjee8 with some modifications. Typically, 30 g of crushed

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germinated or dormant seeds were homogenized in 45 mL of acetone at 4 °C in a blender

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(Carrefour home HBL615-12, France). Acetone was separated from the residue by

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centrifugation (Jouan CR412 Legallais, France) at 4,500 rpm for 15 minutes at 4 ° C. The



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extraction process was repeated four times. Traces of solvent left in the powder were

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eliminated by drying at room temperature overnight.

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Delipidation with hexane was carried out according to French Standard NF V 03-924 using a

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Soxhlet equipped with a cooling system. Hexane (175 mL) was used for the delipidation of 15

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g of crude extract in a cellulose cartridge (Macherey-Nagel, Germany). After eight hours of

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extraction, the residue was dried at room temperature overnight to remove traces of hexane.

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The acetone and hexane powders were stored in sealed jars at 4 °C until use.

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Protein content. The protein content of the different crude extracts was determined using the

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Dumas method9 on a CHN analyzer (Vario MACRO cube ELEMENTAR, Germany). The

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nitrogen-protein conversion factor used was 6.25.

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Screening of non-delipidated crude extracts for ethanolysis activity. To 4.56 mmol of

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sunflower oil, 1.14 mmol of anhydrous ethanol (corresponding to a triacylglycerol:ethanol

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molar ratio of 4:1) were added with or without 12.5 % (w/w of the oil) of non-delipidated

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crude extract. All experiments were performed in an incubator (IKA KS 4000i control,

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Legallais, France) at 40 °C with orbital shaking (250 rpm). After 72 hours of reaction, the

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reaction media were filtered (Millipore, 0.45 µm), diluted with hexane and analyzed by high-

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performance thin-layer chromatography (HPTLC) combined with densitometry.

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The specific activity of the crude extract (µmol.min-1.g-1) was defined as the number of µmol

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of fatty acid ethyl esters (FAEE) released per minute per gram of protein (g) of the crude

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extract.

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Yields were calculated based on the limiting substrate (ethanol).

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ℎ (%) =

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× 100

[1]

where nFAEE and nethanol are the number of moles of ethyl esters and ethanol, respectively.


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The effect of temperature on the transesterification activity of the most active crude extracts

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was evaluated by varying the temperature from 25 to 80 °C.

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Determination of hydrolysis activity. The hydrolysis activities of the crude extracts were

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determined using a pH-stat (Metrohm 736 Titrino GP, Switzerland) equipped with a pH

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electrode and a thermostated vessel. The reaction was started by adding 12.5% (m/m of the

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oil) of the delipidated crude extract to 45 mL of a sunflower oil water emulsion at 10% (w/w)

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previously prepared by homogenization in an Ultra-Turrax IKA T18 (Staufen, Germany). The

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fatty acids derived from lipolysis at 37 °C and pH 7 were continuously titrated using a 20 mM

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sodium hydroxide solution. One unit (U) of lipase activity was defined as the micromoles of

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free fatty acid released per minute in the experimental conditions. Enzymatic activity is

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expressed as units per milligram of protein (U.mg-1).

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Influence of pH on hydrolysis activity. The optimum pH of the crude extracts in hydrolysis

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was determined by varying the pH between 4 and 10 using different buffers at a concentration

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of 100 mM (citrate-phosphate buffer 4-8; Borate-HCl buffer 9 and borate-NaOH buffer 10).

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Sunflower oil (4 g) was emulsified with 40 mL of buffer. The reactions were carried out

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under orbital stirring (250 rpm) for 24 h with 12.5% (w/w of the oil) of delipidated crude

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extract. Next, 10 mL of hexane were added to the reaction medium. The organic phase was

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recovered by centrifugation (Jouan, CR412, Legallais, France) at 4,000 rpm for 15 min at 4

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°C. Free fatty acids were separated by HPTLC and quantified by densitometry. Data are

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expressed as relative activity, compared with activity at the optimum pH defined as 100%.

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Determination of regioselectivity. Regioselectivity during the hydrolysis of purified

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triacylglycerols of sunflower, olive, coconut, groundnut and palm oils by crude extracts was

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determined according to the protocol developed by Akil et al.10 with some modifications. The

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triacylglycerols of the different oils were purified on an alumina column according to



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AFNOR standard T60-241. The reaction medium consisted of 200 µL of the emulsion of

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purified triacylglycerols (10 µmol) in polyvinyl alcohol (20 g.L-1), 1 mL of citrate-phosphate

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buffer (100 mM) and 25 mg of crude lipase extract. The reactions were carried out at the

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optimum temperature and pH of the different crude lipase extracts for 20, 40 and 60 minutes.

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One milliliter of ethanol: sulfuric acid (100: 0.8, v/v) solution was added to stop the reaction.

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The lipids were extracted with 2 mL of hexane and analyzed by HPTLC/densitometry. The

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ratio of 1,2 (2,3) diacylglycerols to that of 1,3 diacylglycerol obtained after partial hydrolysis

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of triacylglycerols was used to determine the crude lipase extract regioselectivity.

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Determination of typoselectivity. Typoselectivity was determined in an emulsion of an

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equimolar mixture of five saturated fatty acid ethyl esters (FAEE) whose chain lengths ranged

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from C10 to C18 or with an equimolar mixture of the five corresponding triacylglycerols in an

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aqueous solution of polyvinyl alcohol (20 g/L). The reaction medium was composed of 200

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µL of the emulsion of lipids (either as ethyl ester or triacylglycerols (10 µmol) in polyvinyl

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alcohol (20 g/L), 1 mL of phosphate buffer (100 mM) and 25 mg of crude extract. The

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reactions were performed at the optimum temperature and pH of the different crude extracts

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for 20, 40 and 60 minutes. One milliliter of ethanol: sulfuric acid solution (100: 0.8, v/v) was

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added to stop the reaction. Next, 500 µL of the hexane phase containing a known amount of

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heptadecanoic acid (internal standard) were spotted on a TLC silica plate (20 × 10 cm, silica

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gel 60 F254) using a semi-automatic Linomat 4 (Camag, Muttenz, Switzerland). Once

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developed in a hexane/diethyl ether/acetic acid (70: 30: 1, v/v/v) solvent, the plates were dried

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at room temperature, and the free fatty acid band (Rf ≈ 0.43) was scraped and transferred into

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50 mL vials for methylation and subsequent analysis by GC-FID

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Determination of fatty acid profiles by GC-FID. The fatty acid methyl esters were prepared

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according to the French standard NF T60-233: 20-30 mg of lipids were diluted in 3 mL of

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sodium methoxide (0.05 M) containing a few drops of phenolphthalein. After 10 min of 8 ACS Paragon Plus Environment

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refluxing, 3 mL of methanol/ acetyl chloride (1:1, v/v) were added until phenolphthalein

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discoloration occurred. The mixture was refluxed again for 10 min. After cooling at room

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temperature, 10 mL of distilled water and 6 mL of hexane were added to the flask. The

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recovered organic phase was dried with anhydrous Na2SO4, filtered and analyzed by GC (HP

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6890 series, Agilent Technologies, USA) using a Supelcowax 10 capillary column (30 m ×

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0.32 mm × 0.250 µm) and a flame ionization detector (270 °C) with helium as carrier gas (2

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mL/min). The column temperature was maintained at 185 °C for 2 min, then raised at a rate of

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4 ° C/min to 225 °C and maintained at this temperature for 5 min. The injector temperature

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was 250 ° C in split mode 1/100. For the analysis of the FAME during the study of the

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typoselectivity, the oven temperature was maintained at 70 ° C for 2 min, then raised to 225 °

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C at a rate of 5 °C/min and maintained at this temperature for 10 min. The FAME peaks were

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identified by comparing retention times with standards. The result given for each analysis is

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the average of three determinations.

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High performance thin layer chromatography coupled with densitometry. These

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measurements were performed according to the protocol established by Moussavou et al.4

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with some modifications. Using an automatic ATS4 (Camag, Muttenz, Switzerland), different

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amounts of standards (ethyl esters of sunflower oil or fatty acids) and test samples were

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applied on a silica plate (HPTLC, 20 × 10 cm, silica gel 60 F254). The plates were eluted with

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hexane/diethyl ether/acetic acid (80: 20: 2, v/v/v), (or 70: 30: 1 (v/v/v) for the regioselectivity

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study) and then revealed by immersion in a copper sulfate: phosphoric acid: methanol: water

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solution (10: 8: 5: 78, v/v/v/v) followed by carbonization of the plates at 180 °C for 10 min.

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The amounts of different lipid species were determined by scanning the plates at 550 with by

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a TLC3 scanner (CAMAG, Muttenz, Switzerland). The amount of ethyl esters (µg) in the

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different reaction media was derived from the calibration curve.



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Statistical analyses. The results are expressed as the mean ± standard deviations of at least

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three repetitions. One way analysis of variance (ANOVA) was performed using Statgraphics

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Centurion XVI (Statpoint Technologies, USA). A difference was considered significant when

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p ˂ 0.05.

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RESULTS AND DISCUSSION

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Screening of crude lipase extract for ethanolysis activity

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Influence of germination on the ethanolysis activity of non-delipidated crude

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extracts

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It is known that most lipases are highly sensitive to ethanol and organic solvents. Ethanolysis

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activity was screened using crude extracts of non-delipidated seeds with a low concentration

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of ethanol in the reaction medium (triacylglycerol: ethanol molar ratio of 4:1) to minimize

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possible inhibition of lipases contained in the extracts. The ethanolysis activities of non-

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delipidated crude extracts from germinated and dormant seeds are listed in Table 2.

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Twelve powders were tested for ethanolysis activity, six from germinated seeds and six from

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dormant seeds. All non-delipidated crude extracts from germinated seeds showed ethanolysis

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activity, whereas among the dormant seeds, only the two species of Adansonia showed

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activity. Results obtained with crude extracts from M. oleifera and J. curcas seeds are in

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agreement with those obtained by Moussavou et al.4 who reported ethanolysis activity in these

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germinated seeds. It is important to note that a negative test for an ethanolysis activity with

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crude extract from dormant seeds of J. curcas, J. mahafalensis, M. drouhardii, and M.

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oleifera does not mean these seeds do not contain lipases. It is possible that the experimental

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conditions, the age or maturity of seeds and environmental conditions of cultivation of the

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plant contributed to the negative result. For example, although both Staubmann et al. (1999)

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and Abigor et al. (2002) worked on J. curcas seeds, they obtained conflicting results.


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Staubmann et al. reported lipase activity only in germinated seeds whereas Abigor et al.

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detected lipase activity in both the dormant and germinated seeds.

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Although crude extract of dormant Adansonia seeds showed some activity, it was less than

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that of germinated seeds. Germination thus appears to be a key to significant activity. This is

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similar to other seeds including castor bean in which lipase activity was reported to increase

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tenfold when germinated seeds were used11. During germination, the plant needs energy to

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grow, and thus activates the necessary lipases to hydrolyze the triacylglycerols in their seeds,

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in fatty acids and glycerol. The fatty acids may then feed the glyoxylate cycle to release

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glucose that will be used to generate the energy necessary for the growth of the plant.

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Among all the active crude extracts, the crude extracts of Adansonia seeds produced the

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highest yields of ethanolysis. Their yields were close (A. suarezensis 65.4 % and A.

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grandidieri 61.4 %) in contrast to other species of the same genus in which a significant

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difference was observed (M. drouhardii 47.5% and M. oleifera 24.8%; J. mahafelnsis 32.2%

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and J. curcas 40%).

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Influence of the delipidation procedure on the ethanolysis activity of crude

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extracts

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Germination has proved to be essential for high level activity or activity in crude extracts

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from non-delipidated seeds. To obtain crude extracts with a high concentration of lipases and

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avoid wrong interpretation of the results due to the presence of native lipids during the

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evaluation of their hydrolytic activity, a preliminary delipidation step is usually performed.

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The influence of this step on the activity of crude extracts from germinated seed was studied.

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In this context, delipidation was carried out either with hexane or acetone. Yields of ethyl

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ester and specific activities of the crude extracts from delipidated seeds were compared with

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those of crude extracts from non-delipidated seeds (Figure 2 and Figure 3).



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Whatever the triacylglycerol: ethanol molar ratio used with the crude extracts from M.

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oleifera and M. drouhardii seeds, the highest yields and specific activities were obtained with

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the crude extracts from non-delipidated seeds. Delipidation of Moringa seeds with acetone

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resulted in the absence of any activity whatever the triacylglycerol:ethanol molar ratio (Figure

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2). Hexane powders from A. suarezensis seeds and from Jatropha seeds led to higher yields

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than their other types of extracts regardless of the triacylglycerol:ethanol molar ratio. This

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behavior resembled that of crude extracts from A. grandidieri seeds, except with a

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triacylglycerol:ethanol molar ratio of 4: 1 whereas acetone powder produced a higher yield.

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Results obtained with crude extracts from M. oleifera seeds resemble those obtained by Dahot

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and Memon12 who described inhibition of lipase from M. oleifera seeds with acetone. These

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authors also reported low heat stability of this lipase. This could explain the low yields and

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specific activities obtained with hexane powder from this seed compared to their non-

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delipidated crude extracts due to the conditions of Soxhlet delipidation. In most cases, acetone

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powder leads to lower yields and specific activities. During extraction with acetone,

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dehydration and delipidation of the powder may occur simultaneously because acetone is a

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strong hydrophilic and polar solvent5. The level of hydration of the enzyme influences the

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spatial structure of the enzyme and its flexibility, thus affecting their catalytic activity13.

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Influence of the triacylglycerol: ethanol molar ratio on the ethanolysis activity of

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crude extracts

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Stoichiometrically, complete conversion of triacylglycerols into ethyl esters requires three

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moles of alcohol per mole of triacylglycerol. To determine the influence of the

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triacylglycerol: ethanol molar ratio on the activity of different crude lipase extracts, other

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experiments were performed with different triacylglycerol: ethanol molar ratios (2:1, 1:1,

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1:1.5, 1:2 and 1:3).


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A decrease in the activity of all acetone powders and in the non-delipidated crude extracts

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from Jatropha seeds was observed when triacylglycerol: ethanol molar ratios higher than 4:1

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were used. The activities of non-delipidated crude extracts from Moringa and Adansonia

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seeds and hexane powders from Jatropha and Adansonia seeds increased when the ratio

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varied from 4: 1 to 2: 1. With a molar ratio of 2:1 to 1:1, only the hexane powders from

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Adansonia seeds showed activity, which increased with the triacylglycerol: ethanol molar

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ratio. However, beyond a molar ratio of 1:1, a decrease was observed. The influence of the

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molar ratio on the ethanolysis activity of crude extracts therefore appears to be a function of

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the delipidation process and of the type of seed. The observed harmful effect of ethanol on the

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activities of crude lipase extracts has also been reported in other lipases in many studies5,6, but

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its mechanism is clearly not yet fully understood14,15, given the different terms used to

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describe it in the literature such as inactivation, deactivation, inhibition and denaturation15.

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However, Li et al.14 estimated that this effect is due to an interaction between the alcohol

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molecules and the hydrophobic and amphipathic amino acid residues of the enzyme. At high

270

concentrations, the alcohol induce a destruction of the original hydrophobic interactions of the

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protein, leading to changes in the tertiary structure of the lipase. This effect is hypothesized to

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be a function of the nature of the interactions involved in the stabilization of the protein and

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explain the observed differences in behavior between the different types of crude lipase

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extracts. Hexane powders are more stable to ethanol than acetone powder. One possible

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reason could be the state of hydration of the enzyme powder as mentioned above. Indeed,

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Kulschewski et al.16 found a loss of catalytic activity of Candida antarctica lipase B when

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incubated in a 60% alcohol solution and a water activity of 0.02, whereas the enzyme was

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stable when water activities were high. At high concentrations of ethanol, non-delipidated

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crude extracts from Moringa and Adansonia seeds maintained their specific activities better,

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although they provided lower yields than their hexane powders (Figure 3). This phenomenon



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was very marked with A. grandidieri seeds: the non-delipidated crude extracts retained nearly

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50% of their specific activity at a triacylglycerol: ethanol molar ratio of 1:3 compared to at a

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molar ratio of 2:1, when its activity was maximum. Delipidation of these seeds is thus

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hypothesized to lead to a reduction in their resistance to ethanol. One possible hypothesis is

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the loss of certain cofactors during delipidation, such as minerals, which are required for

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enzyme stability. Lu et al.17 showed that the incorporation of certain salts (CaCl2, MgCl2,

287

KCl) in the lipase from Candida sp. 99-125 increased its tolerance to methanol. These salts

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formed a stable complex with the protein to resist the conformational changes caused by high

289

concentrations of methanol.

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To summarize screening, crude lipase extracts of species of the same genus appear to have

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similar sensitivity to ethanol and to the delipidation solvent, even though they lead to different

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specific activities and yields. The crude lipase extracts from J. curcas, M. drouhardii and A.

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grandidieri seeds were the species of Jatropha, Moringa and Adansonia that were most active

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in ethanolysis, respectively. Among the crude lipase extracts studied, hexane powders from A.

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grandidieri and A. suarezensis seeds were seen to be the best candidates to produce ethyl

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esters from sunflower oil since their tolerance to ethanol and yields was close to 100% with a

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triacylglycerol: ethanol molar ratio of 1:1. For complete conversion of sunflower oil to ethyl

298

esters with different extracts, strategies such as pretreatment of lipase (salts, etc.), stepwise

299

addition of ethanol, immobilization on a support, or a co-solvent should be used to minimize

300

the effect of ethanol on these lipases. Yields obtained with crude lipase extracts from Moringa

301

seeds were the lowest of the crude lipase extracts we studied. For this reason, they were not

302

retained for subsequent experiments.


Page 15 of 38

303 304


Effects of temperature on the ethanolysis activity of the crude extracts from Adansonia and Jatropha seeds.

305

An increase in temperature in the ethanolysis reaction generally leads to an increase in

306

enzymatic activity up to an optimum temperature beyond which the denaturation of the

307

enzyme resulted in a reduction in its activity. The optimum operational temperature is an

308

important parameter to investigate for optimization of the enzyme-catalyzed reaction. Figure

309

4 shows the effect of temperature on the relative activity of hexane powders from J. curcas, J.

310

mahafalensis, A. grandidieri and A. suarezensis seeds in ethanolysis at temperatures ranging

311

from 25 °C to 80 °C at the optimum triacylglycerol: ethanol molar ratio for each enzymatic

312

powder tested, which was previously determined at 40 °C (triacylglycerol: ethanol molar ratio

313

of 2:1 for the species of Jatropha and triacylglycerol: ethanol molar ratio of 1:1 for the

314

species of Adansonia).

315

Our results showed that 30 °C was the optimum temperature for hexane powders from J.

316

mahafalensis and J. curcas seeds whereas 40 °C was optimum temperature for the hexane

317

powders from A. grandidieri and A. suarezensis seeds. The results we obtained with J. curcas

318

seeds are close to those reported by Jiang et al.18 and Su et al.19 during an in-situ

319

transesterification of germinated seeds from J. curcas with dimethyl carbonate and methanol,

320

respectively. Other plant lipases extracted from seeds, including Pachira aquatica20,

321

Pentaclethra macrophylla Benth21 and passion fruit1 displayed their maximum catalytic

322

activity between 30 °C and 40 °C. The hexane powders of Adansonia seeds showed higher

323

relative activity than 50% at temperatures above 60 °C. These hexane powders appeared to be

324

less prone to thermal denaturation than the two hexane powders from Jatropha seeds whose

325

relative activity was less than 20% at these temperatures. Thus the hexane powders of two

326

species of Adansonia appear to be more thermostable than those from two species of

327

Jatropha. One possible explanation for this behavior is mineral content. Minerals are known 15 ACS Paragon Plus Environment


Page 16 of 38

328

to influence the thermal stability of lipase, and the mineral content of Adansonia is higher

329

than that of Jatropha seeds (Table 1). For example, Inverzini et al.

330

calcium influences the thermal stability of Burkholderia glumae lipase.

331

22

demonstrated that

Hydrolysis activity of delipidated crude lipase extracts

332

The hydrolytic activities of delipidated crude lipase extracts that exhibited ethanolysis activity

333

were assessed. As shown in Figure 5, all extracts that exhibited ethanolysis activity also

334

exhibited hydrolysis activity. The activities of acetone powders were lower than those of

335

hexane powders. The hexane powders from J. mahafalensis seeds (2.89 U/mg) produced the

336

highest activity followed by the hexane powder from A. grandidieri seeds (2.35 U/mg). The

337

most active crude lipase extract in ethanolysis was not necessarily the most active in

338

hydrolysis. For example, hexane powder from J. mahafalensis produced low ethanolysis

339

activity whereas in hydrolysis, it produced the highest hydrolytic activity. Similarly,

340

Tongboriboon et al.23 studied the potential of lipase AK from P. fluoresscens, lipase PS from

341

P. cepacia, lipase AY from C. rugosa, Novozym 435 from C. antartica and Lipozyme TL IM

342

of T. lanuginosa for hydrolysis reactions, esterification and transesterification. These authors

343

demonstrated that the lipase AK had the highest hydrolytic activity, but with the

344

transesterification and esterification reactions the yields obtained were virtually zero. Our

345

results are complementary to this work and confirm that the hydrolytic activity of lipase

346

cannot be used to predict its activity in synthesis reaction and vice versa. Of all the reactions

347

studied with delipidated crude lipase extract, hexane powders produced the highest yields. We

348

thus retained them for subsequent experiments on the effects of pH on hydrolytic activity and

349

for the characterization of the selectivity of crude lipase extracts.


Page 17 of 38


350

Effect of pH on the hydrolytic activity of hexane powders from Adansonia and

351

Jatropha seeds.

352

To determine the optimum pH for hydrolysis, the activity of hexane powder from A.

353

grandidieri, A. suarezensis, J. curcas and J. mahafalensis seeds was evaluated at different pH

354

(4 to 10) relative to their respective optimum temperature. The results are shown in Figure 6.

355

All the enzymes were very active in the pH range 7 to 8 with maximum catalytic activity at

356

pH 8. Staubmann et al.24 and Abigor et al.25 found maximum activity at pH 7.5 and 8.5 with

357

purified and partially purified lipase from J. curcas seed, respectively. The 0.5 unit difference

358

between their results and ours may be due to the potential interaction of the enzyme with

359

other compounds present in the extracts1. Working on the lipase extracted from rapeseeds,

360

Hoppe and Theimer 26 found an optimum pH of 9 for triolein emulsion stabilized with gum

361

arabic alone, but when gum arabic and deoxycholate were used concomitantly, the optimum

362

pH was 8. The presence or absence of an emulsifier in the reaction medium may also explain

363

the differences between our values and those obtained by Staubmann et al.24 and Abigor et

364

al.25. To our knowledge, no study has reported the optimum pH in hydrolysis of crude lipase

365

extracts from J. mahafalensis, A. suarezensis and A. grandidieri seeds. Lipase extracted from

366

the seeds of Pachira aquatica20, which belongs to the same family (Malvaceae) as the species

367

of Adansonia studied here, has an optimum pH of 8, like other lipases extracted from seeds

368

such as Nigella damascena 27 and rubberwood 28.

369 370

Characterization of the selectivity of crude lipase extracts from A. grandidieri and J. mahafalensis seeds.

371

During the enzymatic hydrolysis of triacylglycerols, certain lipases may show specificity of

372

action that can be divided into different groups: typoselectivity, stereoselectivity or

373

regioselectivity. The determination of this specificity could be the first step towards



Page 18 of 38

374

understanding the functioning of these lipases. It is with this goal in view, that we studied the

375

typoselectivy and regioselectivy of the most active extracts in hydrolysis.

376

Regioselectivity of crude lipase extracts from A. grandidieri and J. mahafalensis

377

seeds during the hydrolysis of different vegetable oils

378

Lipase is known to be regioselective during hydrolysis if it has the ability to distinguish the

379

external positions (sn-1 and sn-3) of the triacylglycerol from the internal position (sn-2) 10,29.

380

To investigate the regioselectivity of crude lipase extracts from A. grandidieri and J.

381

mahafalensis seeds, the proportions of 1,2 (2,3) diacylglycerol and 1,3 diacylglycerols of the

382

reaction medium were determined after partial hydrolysis of purified triacylglycerols of olive,

383

coconut, sunflower, peanut and palm oil by these crude extracts (Table 3).

384

Concerning preliminary purification of oils, it is worth noting that it was incomplete for palm

385

and sunflowers oils and that traces of remaining partial glycerides were slightly present. The

386

two crude lipase extracts had the ability to hydrolyze the triacylglycerols in all the oils

387

studied. A lipase that hydrolyzes equally the sn-1, sn-2 and sn-3 positions of a triacyglycerol

388

leads to a ratio of 1,2 (2,3) diacylglycerols to 1,3 diacylglycerols close to 2. After 20 minutes

389

of reaction, this ratio ranged from 82.3 to 2.5 and 199 to 3.2 for the crude lipase extract from

390

A. grandidieri and J. mahafalensis seeds, respectively, depending on the nature of the oil.

391

This means that, regardless of the oil used, crude lipase extracts from A. grandidieri and J.

392

mahafalensis seeds preferentially hydrolyze the ester bonds in sn-1,3 positions of the

393

triacylglycerol and are therefore 1,3 regioselective. Other plant lipases such as those extracted

394

from papaya latex and babaco are also 1,3 regioselective 29. With the crude lipase extract from

395

A. grandidieri seeds, this ratio tended to decrease over time with values that fluctuated from

396

10.8 to 3.8 and 14.4 to 3.4 at respectively, 40 and 60 min, depending on the nature of the oil.

397

On the other hand, with the crude lipase extract from J. mahafalensis seeds, this ratio varied

398

very little (82.30 to 4.1 and 82.3 to 3.9 at 40 and 60 min, respectively, depending on the 18 ACS Paragon Plus Environment

Page 19 of 38


399

nature of the oil). This difference in the behavior of the two crude lipase extracts can be

400

explained by the different hydrolysis percentage of each lipase. Cambon et al.29 estimated that

401

for better expression of lipase specificities, percentage hydrolysis should be less than 15%.

402

The percentage hydrolysis with crude lipase extracts from A. grandidieri seeds was greater

403

than 15% after 20 minutes of reaction for all oils except coconut oil, whereas with the crude

404

extract from J. mahafalensis seeds, percentage hydrolysis remained below 15%, whatever the

405

time or the oil.

406

Typoselectivity of crude lipase extracts from A. grandidieri and J. mahafalensis

407

seeds in a triacylglycerol or ethyl ester mixture.

408

Competitive substrate reactions were used to investigate the typoselectivity of the lipase

409

contained in crude extracts from J. mahafalensis and A. grandidieri seeds to an equimolar

410

emulsified mixture of saturated triacylglycerols or ethyl ester (C10 to C18). Figure 6 shows the

411

time course of the relative molar percentage of each fatty acid obtained after the hydrolysis of

412

the mixture of triacylglycerols and ethyl esters. High relative molar percentage of a fatty acid

413

indicates a preference of the lipase for the corresponding ethyl ester or triacylglycerol30.

414

With the mixture of ethyl esters, both enzyme extracts were capable of hydrolyzing all the

415

substrates. With the crude lipase extract from A. grandidieri seeds (Figure 7-B), at 20 and 40

416

minutes of reaction, capric acid (24.7% and 25.8%) and lauric acid (24.5% and 25.6%) were

417

the fatty acids with the highest relative molar percentages, indicating a preference of this

418

crude extract for ethyl caprate and ethyl laurate. However, at 60 min of reaction, their relative

419

molar percentage decreased in favor of myristic (23.4%) and palmitic (22.5%) acids. The

420

crude extract from A. grandidieri seeds therefore has no pronounced preference for a

421

particularly ethyl ester. The crude lipase extract from J. mahafalensis seeds (Figure 7-D)

422

showed preference for ethyl caprate and ethyl laurate. At 20, 40 and 60 min, capric and lauric

423

acid represented, respectively, 40.8% and 30.7%, 45.1% and 26.2%, 50.7% and 26.1% of the 19 ACS Paragon Plus Environment


Page 20 of 38

424

released fatty acids. When crude lipase extracts from A. grandidieri seeds were used to

425

hydrolyze the mixture of triacylglycerols, at 20 minutes of reaction, all the fatty acids were in

426

relatively equal relative molar proportions (≈ 21%), except capric acid (14.6%). At 40 and 60

427

min of reaction, a preference for tristearin (24.8% and 21.4%), tripalmitin (23.1% and 23.9%)

428

and trimyristin (23.8% and 23.5%) was observed. Although with the longest reaction times, a

429

preference for myristic, palmitic and stearic acids was observed, it was not sufficiently

430

pronounced to confirm the specificity of the crude lipase extract from A. grandidieri seeds for

431

a triacylglycerol. The crude lipase extract from J. mahafalensis seeds (Figure 7-C) showed a

432

preference for long-chain triacylglycerol (tristearin and tripalmitin), whereas the other

433

triacylglycerols were hydrolyzed at almost identical rates (˂ 5.5 %).

434

Whatever the equimolar mixture of substrates used, the crude lipase extract from A.

435

grandidieri seeds did not display clear typoselectivity, unlike the crude lipase extract from J.

436

mahafalensis seeds, which displayed a preference for short chains with ethyl esters and long-

437

chain with triacylglycerols. The selectivity of the crude lipase extract from J. mahafalensis

438

seeds has been shown to be influenced by the type of reaction. Several studies have shown

439

that the type of reaction affects the selectivity of a lipase due to the combined influence of the

440

enzyme and substrate properties31,32. Romero et al.31 explained the selectivity of induced

441

lyophilized supernatant of Aspergillus Niger NYA 135 which is influenced by the type of

442

reaction by the hypothesis that it could contain more than one lipase. Staubmann et al.24 also

443

reported that, after purification of the crude extracts from J. curcas, these crude extracts

444

contained both esterase and lipase. Given the similarity of the properties of the crude lipase

445

extracts from J. curcas and J. mahafalensis seeds previously observed in this study, crude

446

lipase extract from J. mahafalensis seeds could also contain more lipolytic enzymes.

447

In conclusion, this work highlighted the potential of the use of crude lipase extracts from

448

germinated seeds of A. suarezensis, A. grandidieri, J. mahafalensis and M. drouhardii to 20 ACS Paragon Plus Environment

Page 21 of 38


449

produce ethyl esters and fatty acid concentrates from vegetable oils. To our knowledge, no

450

study has previously reported ethanolysis or hydrolysis activity in these seeds. Their activities

451

were shown to be conditioned by their method of preparation, namely the need for seed

452

germination before preparation of the extract, the nature of the solvent used when delipidation

453

is required, and the molar ratios of the reaction medium between oil and ethanol. The crude

454

lipase extracts from A. grandidieri and J. mahafalensis seeds were shown to be 1,3

455

regioselective and can therefore be used for the synthesis of new structured

456

Concerning their selectivity with respect to the carbon chain length of an ethyl ester and a

457

triacylglycerol, crude lipase extract from A. grandidieri seeds showed no selectivity, while

458

that of J. mahafalensis displayed selectivity as a function of the nature of the substrate used.

459

ABBREVIATIONS AND NOMENCLATURE

460

HPTLC, High performance thin layer chromatography; TLC, thin layer chromatography;

461

FAEE, Fatty acid ethyl ester; FAME, Fatty acid methyl ester; TAG, triacylglycerol; NDCE,

462

non delipidated crude extract; RPM, revolution per minute; GC, Gas chromatography; FID,

463

Flame ionization detector.

464

ACKNOWLEDGMENTS

465

This work was conducted in the framework of the PRONOVABIO project, with the financial

466

assistance of the European Union. The contents of this publication are the sole responsibility

467

of the partners and can under no circumstances be regarded as reflecting the position of the

468

European Union. The authors thank the Agence Universitaire de la Francophonie (AUF) and

469

the Centre de Coopération Internationale en Recherche Agronomique pour le Développement

470

(CIRAD) for grants to Paul Alain KOUTEU NANSSOU. We would also like to acknowledge

471

PhileoL and Belwet Biocarburant S.A. for providing the seeds used in this study.

lipids.



472

REFERENCES

473

(1)

Page 22 of 38

Santos, K. C.; Cassimiro, D. M. J.; Avelar, M. H. M.; Hirata, D. B.; de Castro, H. F.;

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Fernández-Lafuente, R.; Mendes, A. A. Characterization of the catalytic properties of

475

lipases from plant seeds for the production of concentrated fatty acids from different

476

vegetable oils. Ind. Crops Prod. 2013, 49, 462–470.

477

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– challenges encountered. Protein Expr. Purif. 2014, 95, 13–21.

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Villeneuve, P. Plant lipases and their applications in oils and fats modification. Eur. J. Lipid Sci. Technol. 2003, 105 (6), 308–317.

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Seth, S.; Chakravorty, D.; Dubey, V. K.; Patra, S. An insight into plant lipase research

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Moussavou, R. W.; Brunschwig, C.; Baréa, B.; Villeneuve, P.; Blin, J. Assessing the

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Enzyme Activity of Different Plant Extracts of Biomasses from Sub-Saharan Africa for

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Ethyl Biodiesel Production. Energy Fuels 2016, 30 (3), 2356–2364.

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(5)

Moussavou, M. R. W.; Brunschwig, C.; Baréa, B.; Villeneuve, P.; Blin, J. Are plant

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lipases a promising alternative to catalyze transesterification for biodiesel production?

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Prog. Energy Combust. Sci. 2013, 39 (5), 441–456.

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Challenges and opportunities. Appl. Energy 2014, 119, 497–520.

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Hassanien, F. R.; Mukherjee, K. D. Isolation of lipase from germinating oilseeds for biotechnological processes. J. Am. Oil Chem. Soc. 1986, 63 (7), 893–897.

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Hama, S.; Kondo, A. Enzymatic biodiesel production: An overview of potential feedstocks and process development. Bioresour. Technol. 2013, 135, 386–395.

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Christopher, L. P.; Hemanathan Kumar; Zambare, V. P. Enzymatic biodiesel:

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Yi, L.; Lakemond, C. M. M.; Sagis, L. M. C.; Eisner-Schadler, V.; van Huis, A.; van

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Boekel, M. A. J. S. Extraction and characterisation of protein fractions from five insect

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species. Food Chem. 2013, 141 (4), 3341–3348.


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(10) Akil, E.; Carvalho, T.; Bárea, B.; Finotelli, P.; Lecomte, J.; Torres, A. G.; Amaral, P.;

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Villeneuve, P. Accessing regio-and typo-selectivity of Yarrowia lipolytica lipase in its

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free form and immobilized onto magnetic nanoparticles. Biochem. Eng. J. 2016, 109,

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101–111.

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(11) Su, E.; Zhou, Y.; You, P.; Wei, D. Lipases in the castor bean seed of Chinese varieties:

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Activity comparison, purification and characterization. J. Shanghai Univ. Engl. Ed.

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2010, 14 (2), 137–144.

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(12) Dahot, M. U.; Memon, A. R. Properties of Moringa oleifera seed lipase. Pak. J. Sci. Ind. Res. 1987, 30 (11), 832–835. (13) Adlercreutz, P. On the importance of the support material for enzymatic synthesis in organic media. Eur. J. Biochem. 1991, 199 (3), 609–614.

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(14) Li, L.; Jiang, Y.; Zhang, H.; Feng, W.; Chen, B.; Tan, T. Theoretical and Experimental

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Studies on Activity of Yarrowia lipolytica Lipase in Methanol/Water Mixtures. J. Phys.

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Chem. B 2014, 118 (8), 1976–1983.

510 511

(15) Lotti, M.; Pleiss, J.; Valero, F.; Ferrer, P. Effects of methanol on lipases: Molecular, kinetic and process issues in the production of biodiesel. Biotechnol. J. 2014, n/a-n/a.

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(16) Kulschewski, T.; Sasso, F.; Secundo, F.; Lotti, M.; Pleiss, J. Molecular mechanism of

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deactivation of C. antarctica lipase B by methanol. J. Biotechnol. 2013, 168 (4), 462–

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(17) Lu, J.; Deng, L.; Zhao, R.; Zhang, R.; Wang, F.; Tan, T. Pretreatment of immobilized

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Candida sp. 99-125 lipase to improve its methanol tolerance for biodiesel production. J.

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Mol. Catal. B Enzym. 2010, 62 (1), 15–18.

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(18) Jiang, Y.; Li, D.; Li, Y.; Gao, J.; Zhou, L.; He, Y. In situ self-catalyzed reactive

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extraction of germinated oilseed with short-chained dialkyl carbonates for biodiesel

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production. Bioresour. Technol. 2013, 150, 50–54.



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(19) Su, E.; You, P.; Wei, D. In situ lipase-catalyzed reactive extraction of oilseeds with

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short-chained dialkyl carbonates for biodiesel production. Bioresour. Technol. 2009,

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100 (23), 5813–5817.

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(20) Polizelli, P. P.; Facchini, F. D. A.; Cabral, H.; Bonilla-Rodriguez, G. O. A New Lipase

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Isolated from Oleaginous Seeds from Pachira aquatica (Bombacaceae). Appl. Biochem.

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Biotechnol. 2008, 150 (3), 233–242.

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(21) Enujiugha, V. N.; Thani, F. A.; Sanni, T. M.; Abigor, R. D. Lipase activity in dormant

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seeds of the African oil bean (Pentaclethra macrophylla Benth). Food Chem. 2004, 88

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(3), 405–410.

530

(22) Invernizzi, G.; Papaleo, E.; Grandori, R.; De Gioia, L.; Lotti, M. Relevance of metal

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ions for lipase stability: Structural rearrangements induced in the Burkholderia glumae

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lipase by calcium depletion. J. Struct. Biol. 2009, 168 (3), 562–570.

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(23) Tongboriboon, K.; Cheirsilp, B.; H-Kittikun, A. Mixed lipases for efficient enzymatic

534

synthesis of biodiesel from used palm oil and ethanol in a solvent-free system. J. Mol.

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Catal. B Enzym. 2010, 67 (1–2), 52–59.

536 537

(24) Staubmann, R.; Ncube, I.; Gübitz, G. M.; Steiner, W.; Read, J. S. Esterase and lipase activity in Jatropha curcas L. Seeds. J. Biotechnol. 1999, 75 (2), 117–126.

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(25) Abigor, R. D.; Uadia, P. O.; Foglia, T. A.; Haas, M. J.; Scott, K.; Savary, B. J. Partial

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purification and properties of lipase from germinating seeds of Jatropha curcas L. J.

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Am. Oil Chem. Soc. 2002, 79 (11), 1123–1126.

541

(26) Hoppe, A.; Theimer, R. R. Rapeseed lipase—pH dependent specificity for native lipid

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body autolysis and lipolysis of artifical oil droplets. J. Plant Physiol. 1997, 151 (4),

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390–398.

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(27) Rudyuk, V. F.; Korchagina, L. N. A study of some properties of the lipase from the seeds ofNigella damascena. Chem. Nat. Compd. 1975, 11 (5), 673–676.


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(28) Weerasooriya, M. K. B.; Kumarasinghe, A. A. N. Isolation of alkaline lipase from

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rubber seed — Partial purification, characterization and its potential applications as a

548

detergent additive. Indian J. Chem. Technol. 2012, 19, 244–249.

549

(29) Cambon, E.; Rodriguez, J. A.; Pina, M.; Arondel, V.; Carriere, F.; Turon, F.; Ruales, J.;

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Villeneuve, P. Characterization of typo-, regio-, and stereo-selectivities of babaco latex

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lipase in aqueous and organic media. Biotechnol. Lett. 2007, 30 (4), 769–774.

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(30) Rangheard, M.-S.; Langrand, G.; Triantaphylides, C.; Baratti, J. Multi-competitive

553

enzymatic reactions in organic media: a simple test for the determination of lipase fatty

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acid specificity. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1989, 1004 (1), 20–

555

28.

556

(31) Romero, C. M.; Pera, L. M.; Olivaro, C.; Vazquez, A.; Baigori, M. D. Tailoring chain

557

length selectivity of a solvent-tolerant lipase activity from Aspergillus niger MYA 135

558

by submerged fermentation. Fuel Process. Technol. 2012, 98, 23–29.

559

(32) Vaysse, L.; Ly, A.; Moulin, G.; Dubreucq, E. Chain-length selectivity of various

560

lipases during hydrolysis, esterification and alcoholysis in biphasic aqueous medium.

561

Enzyme Microb. Technol. 2002, 31 (5), 648–655.

562

(33) AOAC International. AOAC: Official Methods of Analysis (Volume 1); 1990.

563

(34) Folch, J.; Lees, M.; Stanley, G. H. S. A Simple Method for the Isolation and

564

Purification of Total Lipides from Animal Tissues. J. Biol. Chem. 1957, 226 (1), 497–

565

509.



Page 26 of 38


Page 27 of 38


566

Figure captions

567

Figure 1. Schematic diagram of the preparation of the different crude lipase extracts.

568

Figure 2. Yield of ethanolysis reaction (%) catalyzed by crude lipase extracts from A.

569

grandidieri, A. suarezensis, M. oleifera, M. drouhardii, J. curcas and J. mahafalensis seeds

570

depending on the TAG:ethanol molar ratio and delipidation procedure. Reaction conditions:

571

40 °C under orbital agitation at 250 rpm for 72 h with 12.5% (m/m of oil) of crude extract.

572

.Figure 3. Ethanolysis activity of crude lipase extracts from A. grandidieri, A. suarezsensis,

573

M. oleifera, M. drouhardii, J. curcas and J. mahafalensis seeds depending on the delipidation

574

procedure and on the TAG:ethanol molar ratio. Reaction conditions: 40 °C under orbital

575

agitation at 250 rpm for 72 h with 12.5% (m/m of oil) of crude extract.

576

Figure 4. Influence of temperature on the ethanolysis activity of crude lipase extracts from A.

577

grandidieri, A. suarezensis, J. mahafalensis and J. curcas seeds. Reaction conditions: 72 h of

578

orbital agitation at 250 rpm at 12.5% (m/m of oil) of crude lipase extract with a TAG:ethanol

579

molar ratio of 2:1 and 1:1 for Jatropha and Adansonia seeds, respectively.

580

.Figure 5. Hydrolytic activity (U.mg-1 of protein) of delipided crude extract in the hydrolysis

581

of sunflower oil. Reaction conditions: oil:water mass ratio of 10% (m/m) at pH 7 with 12.5%

582

(m/m of oil) of crude extract.

583

.Figure 6. Influence of pH on the hydrolytic activity of crude lipase extracts from A.

584

grandidieri, A. suarezensis, J. mahafalensis and J. curcas seeds. Reaction conditions:

585

oil:buffer mass ratio of 10 % (m/m) under orbital agitation at 250 rpm for 24 h with 12.5% of

586

crude extract.

587

.Figure 7. Relative molar content of free fatty acids released after the hydrolysis of an

588

equimolar mixture of ethyl ester (B and D) and triacylglycerol (A and C) by crude lipase

589

extract from J. mahafalensis (C and D) and A. grandidieri (A and B) seeds at 0, 20, 40 and 60

590

min. Reaction conditions: 200 µl of the emulsion of mixture of triacylglycerol or ethyl ester



Page 28 of 38

591

(10 µmol) in polyvinyl alcohol (20 g/l), 1 ml of citrate-phospahte buffer and 25 mg of crude

592

lipase extract under orbital agitation (250 rpm) at 40 °C and 30 °C for the crude lipase extract

593

from A. grandidieri and J. mahafalensis seeds, respectively.


Page 29 of 38


Table 1. Proximal composition of J. mafahalensis, J. curcas, M. oleifera, M. drouhardii, A. grandidieri and A. suarezensis seeds. Asha

Proteinb Lipidc Carbohydratesd Moisture (%) (g/100 g dry matter) Seeds 4.7 ±0.1 4.2 ±0.1 24.4 ±0.7 50.6 ± 0.5 20.8±0.3 J. curcas 4.5 ± 0.1 2.7 ±0.5 31.3 ±0.4 52.3±0.3 13.7±0.3 J. mahafalensis 4.3 ±0.1 2.7 ±0.1 33.6 ±0.6 44.5±0.1 19.2±0.6 M. drouhardii 7.1 ±0.0 4.4± 0.1 40.2 ±0.3 34.5± 0.7 20.9±0.4 M. oleifera 4.8± 0.4 5.5 ±0.5 19.1 ±0.1 61..8± 0.7 13.6±0.3 A. grandidierii 7.5 ±0.4 8.4 ±0.2 36.9 ±0.1 35.3± 0.5 19.3±0.3 A. suarezensis a,b,c,d based on the standard method recommended by AOAC33, Dumas method9, Folch procedure34 and by difference (100- (sum of percentage of ash, protein and lipid), respectively.



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Table 2. Ester yields from ethanolysis of sunflower oil catalyzed by a non-delipidated crude extract. Reaction conditions: triacylglycerol: ethanol molar ratio of 4:1 at 40 °C using 12.5 % of non-delipidated crude extract under orbital agitation at 250 rpm for 72 h. Seed

Germinated Dormant

J. curcas

40.0 ± 1.6

-

J. mahafalensis 32.2 ± 0.9

-

M. drouhardii

47.5 ± 3.1

-

M. oleifera

24.8 ± 0.5

-

A. grandidierii

61.4 ± 3.0

˂ 0.1

A. suarezensis

65.4 ± 2.1

˂ 0.1


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Table 3. Relative percentage of diacylglycerol obtained during the partial hydrolysis of vegetables oils with the crude lipase extract from A. grandidieri and J. mahafalensis seeds after 20, 40 and 60 min of reaction. Reaction conditions: 200 µl of the emulsion of oil (10 µmol) in polyvinyl alcohol, 1 mL of citrate-phosphate buffer and 25 mg of crude lipase extract under orbital agitation at 250 rpm at 40 °C and 30 °C for the crude lipase extract from A. grandidieri and J. mahafalensis seeds, respectively.

Olive 1,2(2,3) 1,3 DAG DAG A. grandidieri 0 0 0 20 0.5 ± 0.0 99.5 ± 0.0 40 10.0 ±1.7 90.0 ± 1.7 60 17.1 ±0.2 82.9±0.2 J.mahafalensis 0 0 0 20 1.2 ±0.1 98.8 ± 0.1 40 1.2 ±0.1 98.8 ± 0.1 60 1.2 ±0.1 98.8 ± 0.1

Copra 1,2(2,3) 1,3 DAG DAG

Peanut 1,2(2,3) 1,3 DAG DAG

Palm 1,2(2,3) 1,3 DAG DAG

Sunflower 1,2(2,3) 1,3 DAG DAG

0 5.1 ± 0.0 8.5 ± 0.2 6.5 ± 0.7

0 94.9±0.0 91.5±0.2 93.5±0.7

0 8.9 ± 0.7 15.7 ± 0.2 22.6± 0.3

0 91.1 ± 0.7 84.3±0.2 77.4±0.3

76.4 ± 3.2 10.4 ± 0.8 9.9 ± 0.0 6.9 ± 0.6

23.6 ±3.2 89.6 ± 0.8 90.1 ± 0.0 93.1 ± 0.6

63.9 ± 3.6 23.6 ± 1.0 20.8 ± 0.7 15.0 ± 1.8

36.1 ± 3.6 76.4 ± 1.0 79.2 ±0.7 85.0 ± 1.8

0 10.4 ±0.4 9.6 ± 0.2 10.4 ±1.0

0 89.6 ± 0.4 90.4 ± 0.2 89.6 ± 1.0

0 1.7 ± 0.0 1.6 ± 0.2 1.4 ± 0.0

0 98.3 ± 0.0 98.4 ± 0.2 98.6 ± 0.0

76.4 ± 3.2 14.6 ± 1.2 12.6 ± 1.5 10.8 ± 0.6

23.6 ±3.2 85.4 ± 1.2 87.4 ± 1.5 89.2 ± 0.6

63.9 ± 3.6 28.5 ± 2.0 19.7 ± 3.2 20.6 ± 0.1

36.1 ± 3.6 71.5 ± 2 80.3 ± 3.2 79.4 ± 0.1

31

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Seeds

Sorting Soaking Germination Dehulling

Dehulling

Crushing

Crushing

Drying

Drying

Non-delipidated crude extract from germinated seeds Delipidation (Acetone)

Delipidation (Hexane)

Acetone powder from Hexane powder from germinated seeds germinated seeds

Non-delipidated crude extract from dormant seeds Delipidation (Acetone) Acetone powder from dormant seeds

Delipidation (Hexane) Hexane powder from dormant seeds

Figure 1.


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J. curcas

J. mahafalensis

M. oleifera

M. drouhardii

A. suarezensis

A. grandidieri

Figure 2.



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J. mahafalensis

J. curcas

M. oleifera

M. drouhardii

A. grandidieri

A. suarezensis

Figure 3.


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100

Relative activity (%)

80

60

40

A. grandidierii A. suazeriensis J. curcas J. mahafaliensis

20

0 25

30

40

50

60

70

80

Temperature (°C)

Specific activity (U/mg protein)

Figure 4.

3.5

Hexane powder

Acetone powder

3 2.5 2 1.5 1 0.5 0 J. curcas

J. mahafalensis A. suarezensis A. grandidieri

Figure 5.



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100

Relative activity (%)

80

60

40

A. grandidieri A. suarezensis J. curcas J. mahafalensis

20

0 4

5

6

7

8

9

10

11

pH

Figure 6.


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A

B

D C

Figure 7. 37

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Table of contents graphics


Lipase Activity of Tropical Oilseed Plants for Ethyl Biodiesel Synthesis

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