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ASSESSING THE ENZYME ACTIVITY OF DIFFERENT PLANT EXTRACTS OF BIOMASSES FROM SUBSAHARAN AFRICA FOR ETHYL BIODIESEL PRODUCTION Redeo Wilfried Moussavou M., Christel Brunschwig, Bruno Barea, Pierre Villeneuve, and Joel Blin Energy Fuels, Just Accepted Manuscript • DOI: 10.1021/acs.energyfuels.5b01829 • Publication Date (Web): 26 Jan 2016 Downloaded from http://pubs.acs.org on February 1, 2016
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Graphical abstract
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ASSESSING THE ENZYME ACTIVITY OF DIFFERENT PLANT EXTRACTS OF BIOMASSES FROM SUBSAHARAN AFRICA FOR ETHYL BIODIESEL PRODUCTION
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Rédéo Wilfried Moussavou M.a, Christel Brunschwiga, Bruno Baréa b, Pierre Villeneuveb,
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Joël Blinab*
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a
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BP 594, Ouagadougou 01, Burkina Faso
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b
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(CIRAD),73 rue Jean-François Breton, 34398 Montpellier Cedex 5, France
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* Corresponding author
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Tel.: +33 4 67 61 59 05
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E-mail address:
[email protected] Page 2 of 34
Institut International d'Ingénierie de l'Eau et de l'Environnement (2iE), Rue de la Science 01
Centre de Coopération Internationale en Recherche Agronomique pour le Développement
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Abstract
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This work focused on identifying plant biomasses as potential sources of lipases and their use
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as biocatalysts for ethyl biodiesel production. To that end, plant extracts prepared from
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dormant and germinated seeds of eight local (Burkina Faso) oil-bearing biomasses were tested
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as a biocatalyst for the hydrolysis and ethanol transesterification of sunflower oil. The
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optimum transesterification conditions with the most active extracts were then determined.
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The results of the hydrolysis and transesterification tests showed that the plant extracts from
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germinated Jatropha curcas (JG) and Moringa oleifera (MG) seeds contained lipases whose
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catalytic activity could be utilized without prior purification. The highest hydrolysis yield
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obtained was 88% using the JG extracts and 62% with the MG extracts after 24 hours of
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reaction at 40°C and at pH 7. For transesterification, the highest ethyl ester yield obtained was
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94% with the JG extracts and 48% with the MG extracts after 144 hours at 37°C, following
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two successive transesterification reactions. Using the plant extracts in transesterification
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called for the stepwise addition of ethanol in a TAG:Ethanol molar ratio of (1:0.3) at 6-hour
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intervals to overcome the inhibiting effect of high ethanol concentrations on the catalytic
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activity of the plant extracts. In addition, a study on the influence of the reaction conditions
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showed that water had a negligible effect on transesterification yield when its content in the
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ethanol was lower than or equal to 5% v/v and that adding silica to the reaction medium led to
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an increase in transesterification yield.
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Keywords: Plant lipases, transesterification, enzyme catalysis, glycerol adsorption,
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biodiesel,
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I. Introduction
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Biodiesel is a mixture of Fatty Acid Alkyl Esters (FAAE) recognized as a relevant alternative
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fuel to fossil fuel in diesel engines. Either as an additive or as replacement, biodiesel is well-
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known to have environmental and economical benefits. Its merits include being non-toxic,
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biodegradable, domestically produced and renewable resource 1, 2. The most common way to
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produce biodiesel is the transesterification reaction of triacylglycerol (TAGs) with an alcohol.
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TAGs are the main constituents of vegetable oils or animal fats. The alcohol used may be
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methanol or ethanol to obtain Fatty Acid Methyl Esters (FAME) or Fatty Acid Ethyl Esters
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(FAEE), respectively. For health-safety and environmental reasons, the use of ethanol as
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alcohol in biodiesel production processes is preferred over methanol 3. Ethanol is less toxic
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than methanol and can be easily obtained by fermenting sugar- or starch-rich biomasses 4, 5. As
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ethanol is obtained from biomass conversion (also called bioethanol) its CO2 footprint is
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almost neutral compared to methanol, mostly derived from the petrochemical industry.
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TAGs transesterification is conventionally performed using alkaline (NaOH, KOH), acid (HCl,
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H2SO4) or enzyme catalysts (lipases) 2, 6. Alkaline catalysis is most investigated for biodiesel
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production because of its shorter reaction time and high productivity. However, alkaline
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catalysis presents drawbacks related to the inevitable production of soaps caused by the
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saponification of free fatty acids (FFA) 7. The use of alkaline catalysts is strictly limited to oils
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with low FFA and water content. Acid catalysts are rarely investigated because they are more
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corrosive and less efficient 5.
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An increasing number of research work published these last 10 years suggest the use of lipase-
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catalyzed transesterification methods for biodiesel production 8. Indeed, compared to chemical
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catalysts, lipases-catalyzed transesterification is performed in more gentle conditions, and with
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a large variety of triglyceride substrates, including waste oils and fats with high levels of FFA. 3 ACS Paragon Plus Environment
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Furthermore, biodiesel separation and purification is much easier, resulting in a more
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environmentally friendly process 9-11. Lipases are water tolerant and less demanding regarding
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feedstock quality than chemical catalysts. A minimum quantity of water is needed to maintain
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their active conformation when used in an organic medium 12, and lipases are able to catalyze
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both transesterification of TAGs and esterification of FFA avoiding the formation of soaps.
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This action specificity makes possible the production of biodiesel from vegetable oil
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containing high level of FFA 5, 11, 13-18. Furthermore, in the case of lipase-based synthesis of
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biodiesel, the purification of the produced esters are easily performed without wastewater
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generation 1.
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Despite the growing interest regarding the use of lipases for biodiesel production, alkaline
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catalysis is still the most important technological route for the industrial production 7. Only a
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few cases have been reported to use enzymatic process for the production of biodiesel on an
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industrial scale 19, 20. There are still obstacles to the effective use of lipases for industrial
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biodiesel production. Lipases have low stability in the presence of excess alcohol and low
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reaction velocity compared to the chemical catalysts. But the main obstacles to the effective
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use of lipases is the relatively high costs of commercial lipase production 1, 6. Most of
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commercial lipases are produced from microorganisms such as fungi, yeasts and bacteria 21
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and obtaining these microbial lipases often calls for complex biotechnological processes.
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Consequently, the high cost of these enzyme preparations has a direct impact on biodiesel
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production costs.
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To reduce biodiesel production costs by lipase catalysis, a strategy is the identification of
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others lipase sources, easy to use and to mobilize which should facilitate large-scale
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development of lipase-catalyzed transesterification. Villeneuve 22, Paques et al., 23 and Barros
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promising source of lipases for industrial application. Plant lipases are less expensive than
clearly pointed out in three interesting bibliographic reviews that plants represent a
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microbial lipases. They are widely available, depending on the availability of the biomass from
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which they are derived and they are easy to use with or without partial purification. Lipases
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activity of crude extracts of plants like oilseeds, cereal grains and latex has been investigated.
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And it was shown that crude extracts from papaya latex (Carica papaya) 25-27, babaco latex
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(Carica pentagona) 28, germinated rape seeds (Brassica napus), germinated Jatropha seeds
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(Jatropha curcas) 29, 30, black cumin seeds (Nigella sativa) 31 or castor beans (Ricinus
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communis) 32 are able to catalyze both hydrolysis and transesterification of TAGs.
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In the light of these results, the purpose of this work was to identify active plant lipases to
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produce biodiesel by ethanol transesterification from new biomass sources. To that end, plant
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extracts prepared from seeds of eight oil-bearing biomasses available in Burkina Faso were
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screened as biocatalysts in the hydrolysis of TAGs then in the transesterification of TAGs with
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ethanol for the most active ones. Optimum transesterification conditions such as TAG/EtOH
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ratio, EtOH/water ratios and biocatalyst load with the most active extracts were then
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determined. Lastly, the yields were improved by glycerol trapping then using a two-stage
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transesterification process.
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II. Material and methods
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II.1.Material
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All the chemicals, solvents and alcohols used were of analytical grade and purchased from
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Sigma, St Louis, MO, USA. The high performance silica plates (HPTLC plates, 20 × 10 cm
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Silica-gel 60 F254) for thin-layer chromatography (TLC) were purchased from Merck
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(Darmstadt, Germany). The vegetable oil used (sunflower oil, Lesieur®) was of food quality
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and was purchased in a local supermarket in Ouagadougou.
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The crude plant extracts used as biocatalysts for TAG hydrolysis and transesterification were
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obtained from dormant or germinated seeds of eight plant biomasses available in Burkina Faso 5 ACS Paragon Plus Environment
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and more generally in West Africa. They were shea nut (Vitellaria paradoxa), cashew nut
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(Anacardium occidentale), desert date (Balanites aegyptiaca), groundnut (Arachis hypogaea),
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moringa (Moringa oleifera), jatropha (Jatropha curcas), neem (Azadirachta indica) and
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mango (Mangifera indica). All of the seeds and mango stones were obtained on the local
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market in Ouagadougou.
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II.2.Methods
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II.2.1. Preparation of plant extracts from seeds
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The plant extracts were prepared from germinated seeds. To trigger germination, the seeds
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were placed between two sheets of moist absorbent paper for 3 to 4 days. Nevertheless, some
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plant extracts from dormant seeds were prepared in order to test whether the seeds were active
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all the same. The dormant and germinated seeds, from the different plant biomasses, were
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hulled and the kernels obtained were ground in an electric propeller grinder (Moulinex ®, 843
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model) until a uniform homogenate was obtained. The homogenate was dried in a ventilated
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oven (HERAEUS, UT6) at 35°C for 24 hours. In order to extract the lipids contained in the
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homogenate, 10 g aliquot fractions were weighed and placed in cellulose thimbles plugged
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with cotton wool and inserted into a Soxhlet apparatus. Extraction was carried out under reflux
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with around 200 mL of hexane for 6 to 8 hours. The defatted homogenate was dried in the
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open air under a hood for one night to eliminate all traces of solvent. The resulting enzyme
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extracts were ground again to a powder with a particle size under 600 µm. Each enzyme
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extract was then packed in hermetically sealed plastic boxes and stored in the refrigerator at
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4°C pending use.
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II.2.2. Hydrolysis tests
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The catalytic activity of the plant extracts in hydrolysis was tested on the sunflower oil in an
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aqueous medium. In a 250 mL flask, 3 g of plant extracts (i.e. 20% w/w compared to the oil) 6 ACS Paragon Plus Environment
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were added to a heterogeneous mixture containing 15 g of sunflower oil (i.e. 17.36 mmol) and
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150 mL of distilled water. The mixture was vigorously homogenized using a magnetic stirrer
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(IKA IKAMAG RCT model) at 400 rpm at room temperature for 7 days. A hydrolysis test
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without a catalyst was carried out in parallel under the same reaction conditions. Samples of 2
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mL of reaction medium were taken after 24, 48, 72 and 168 hours. The samples were washed
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in hexane, then centrifuged (Jouan CR412 model centrifuge) for 30 minutes at 2000 rpm. The
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hexane phases recovered were evaporated under reduced pressure (40°C, 400 mbar). Around
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40 µL of the lipid fraction collected was dissolved in 4 mL of hexane. 100 µL of the stock
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solution obtained was diluted in 900 µL of hexane. From 10 to 20 µL of the daughter solution
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was then analysed qualitatively by HPTLC. Each hydrolysis test was duplicated. The existence
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of hydrolytic activity was reflected in the appearance at the HPTLC plate development line of
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a black spot typical of FFA (Rf ≈ 0.4) resulting from TAG hydrolysis.
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In order to determine the optimum hydrolysis pH for the most active lipases extracts,
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hydrolysis tests were carried out on 4 g of sunflower oil (4.63 mmol) in 40 mL of a buffer
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solution whose pH varied between 2 and 10 (buffer sodium acetate pH4.0–5.0, buffer sodium
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phosphate pH6–8, and buffer sodium borate pH9-10). Oil hydrolysis was initiated by adding
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0.8 g of plant extracts (20% w/w compared to oil). The different reaction media were
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incubated in an oven (Stuart orbital incubator, Legallais company) at 40°C for 24 hours. After
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24 hours, the different reaction media were acidified with 37% hydrochloric acid (HCl) to halt
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the reaction by denaturing the enzyme.
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To determine the optimum hydrolysis temperature, the tests were carried out under the same
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reaction conditions as those described for the pH. The pH of the reaction medium was
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maintained fixed (pH=7.2 for the hydrolysis reactions using extracts of germinated jatropha
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and moringa seeds and pH=8 for the hydrolysis reactions using extracts of germinated shea nut
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seeds), and different reaction temperatures ranging from 30°C to 60°C were applied. The
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different reaction media were then incubated in an orbital incubator at 250 rpm for 6 hours. 7 ACS Paragon Plus Environment
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When determining the optimum pH and optimum temperature, a test was carried out without a
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biocatalyst for each of the conditions studied.
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II.2.3. Transesterification tests
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II.2.3.1.
Screening assay
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The transesterification reactions were initiated by adding 0.8 g of plant extract (i.e. 20% w/w
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compared to oil) to a 25-mL flask containing 4 g of sunflower oil (4.63 mmol) and 7.24 mL of
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ethanol (0.12 mol) corresponding to a TAG:EtOH molar ratio of (1:27). As it is known that a
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large amount of alcohol can induce inhibition effects on the enzyme activity, a second set of
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transesterification tests was carried out, this time with 0.13 mL of ethanol (i.e. 2.31 mmol)
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corresponding to a TAG:EtOH molar ratio of (2:1). The hermetically sealed flasks were
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incubated in an oven (Stuart orbital incubator, Legallais company) at 37°C for 48 hours with
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orbital stirring at 250 rpm. Two blank tests without enzyme extract were carried out under the
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two sets of reaction conditions. Some samples of around 40 µL of reaction medium were taken
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periodically after 2, 7, 24 and 48 hours for qualitative analysis by HPTLC, then quantitative
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analysis by HPTLC coupled to a densitometry system. The existence of catalytic activity was
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reflected in the appearance at the HPTLC plate development line of a black spot typical of
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FAEE (Rf ≈ 0.7) resulting from TAG transesterification with ethanol.
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II.2.3.2.
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Optimum transesterification conditions were determined solely with the plant extracts found to
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be most active in hydrolysis. An initial set of transesterification tests was carried out on 4 g of
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sunflower oil with 0.8 g of plant extracts (20% w/w compared to oil) using different initial
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quantities of anhydrous ethanol, i.e. 80, 135, 270, 440 and 800 µL of ethanol corresponding to
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respective TAG:EtOH molar ratios of (1:0.3), (1:0.5), (1:1), (1:2) and (1:3). Then, under the
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same operating conditions, three transesterification tests were carried out fixing the
Determination of optimum transesterification conditions
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TAG:EtOH molar ratio at (1:0.3) and the catalyst load at 20%, but this time varying the
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ethanol water content (v/v) by 5, 10 and 15%. Lastly, three final transesterification tests were
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carried out with anhydrous ethanol varying the catalyst load in an enzyme/substrate ratio by
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10, 20 and 30% w/w while maintaining the TAG:EtOH ratio at (1:0.3). The different reaction
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media were incubated in an oven (Stuart orbital incubator, Legallais company) at 37°C for 24
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hours under orbital stirring at 250 rpm. After 24 hours, some samples of around 40 µL of each
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reaction medium were taken and analysed by HPTLC coupled to a densitometry system to
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determine the amount of fatty acids ethyl ester (FAEE) formed.
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II.2.3.3.
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To have a complete transesterification of TAG, a stepwise addition method of ethanol was
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performed and the reaction was developed with a given quantity of silica-gel (20% w/w
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compared to the oil). Ethanol was added in the medium in fractions of 80 µl at 6-hour intervals
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for 72 hours. The reaction medium obtained with each of the plant extracts after carrying out
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the transesterification tests was roughly filtered through cotton to eliminate any suspended
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particles. The filtered medium was then used as the substrate for a second transesterification
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reaction adding 0.8 g of new biocatalyst and 0.8 g of new silica. For the first 24 hours, no
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alcohol was added to the medium to enable the incorporation of any ethanol that did not react
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during the first transesterification reaction. Then, 80 µL of ethanol was added to the medium
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every 6 hours for 48 hours (8 additions). Some samples of around 40 µL of each reaction
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medium were taken at the end of the reaction and analysed by HPTLC coupled to a
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densitometry system to determine the amount of FAEE formed.
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Two stages transesterification tests
II.2.4. Thin layer chromatography analysis coupled to a densitometry system
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FFA and FAEE compositions of the reaction medium were determined using a
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HPTLC/densitometry system (CAMAG) according to the literature 33 with the following
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modifications. 40 µL samples of each reaction medium were diluted in hexane so as to obtain a
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solution with 0.1 µg/µL of total lipids. Aliquots of 10 to 20 µL were analysed on silica plates
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by high performance thin layer chromatography (HPTLC) (20 × 10 cm, Silica-gel 60 F254,
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Merck, Darmstadt, Germany). The sample to be analysed was deposited using an automatic
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depositor (CAMAG, Linomat IV, Camag Ltd.) under a nitrogen atmosphere 15 mm from the
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lateral edge and 8 mm from the lower edge of the plate in bands 5 mm in length, 5 mm apart.
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Sixteen lanes were applied to each plate, at a rate of 0.2 µl/s. The HPTLC plates were eluted
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with a hexane-ethyl ether-acetic acid mixture (70:30:1 v/v/v) to analyse the FFAs or with a
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hexane-ethyl ether-acetic acid mixture (90:10:1, v/v/v) to analyse the FAEE. The lipid
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compounds were then visualized by carbonization at 180°C in an oven for 10 min after rapid
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immersion of the plate in a saturated copper sulphate solution, to which phosphoric acid,
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methanol and water had been added beforehand in the following proportions, 10:8:5:78,
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(v/v/v/v). Each constituent appeared as a black spot on the plate substrate which remained
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white. Some FFA and FAEE standards were prepared from the sunflower oil and eluted at the
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same time as the different reaction media to enable identification by comparison of the Rf
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values of the target compounds in the different reaction media. After development in a suitable
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elution solvent, HPTLC plates were analyzed by densitometry for FFA and FAEE
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quantification. Densitometric evaluations were performed with a TLC-scanner 3 (CAMAG,
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Switzerland) in absorbance mode at λ = 370 nm. Data were processed with the software
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winCATS.
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The hydrolysis yield was determined by the ratio between the number of moles of FFA
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measured and the number of hydrolysable functions initially present on the lipid substrate. The
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formula used to calculate the hydrolysis yield was as follows (Equation 1):
FFA% mol =
nFFA ∗ 100 1 3 ∗ n TAG + 2 ∗ nDAG + nMAG
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Where, nFFA is the number of moles of FFA. nTAG, nDAG and nMAG are respectively the
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initial number of moles of tri- , di and mono-acylglycerides initially contained in sunflower
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oil.
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The transesterification yields expressed as a percentage of FAEEs formed were obtained from
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the following equation (Equation 2):
Yld % =
n FAEE ∗ 100 2 3 ∗ n TAG + 2 ∗ nDAG + nMAG
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Where, nFAEE is the number of moles of FAEE. nTAG, nDAG and nMAG are respectively
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the initial number of moles of tri- , di and mono-acylglycerides initially contained in sunflower
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oil.
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III.
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In order to ascertain the lipase activities of the prepared plant extracts, the extracts were tested
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for their catalysis of sunflower oil hydrolysis and transesterification. We opted for refined
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sunflower oil as the substrate, due to the need to work with very clean oil, identical from one
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batch to the next, and especially whose composition was close to jatropha, cottonseed and
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rapeseed oils.
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III.1. Screening of hydrolytic activity
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The hydrolytic activity of 16 extracts obtained from dormant and germinated seeds of the eight
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oil-bearing biomasses selected was tested on sunflower oil in simple ways without using an
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emulsifying agent and at room temperature. As hydrolysis is the biological function of lipases,
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this test was used for an initial screening to identify catalytically active extracts. According to
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the hydrolysis test results obtained after a qualitative HPTLC analysis, six of the fourteen plant
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extracts displayed a hydrolytic activity on sunflower oil : the extracts obtained i) after
Results and discussion
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germination of shea nut, jatropha, desert date, moringa and groundnut seeds and ii) the extract
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of ungerminated shea nut seeds. The extracts obtained from the jatropha seeds having the
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strongest catalytic activity during TAG hydrolysis. Hydrolytic activity was found to be
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particularly present in the plant extracts prepared from germinated seeds, except from the
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extracts of dormant shea nut seeds. In compliance with the data in the literature, hydrolytic
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activity in the seeds was generally greater once the seeds had started to germinate. In seeds,
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lipases mobilize lipids during germination. Lipids are nutrient reserves in oilseeds; they are
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hydrolysed into fatty acids and glycerol, used to produce energy or for the synthesis of sugars,
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amino acids and carbon chains needed for embryo growth 34.
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III.2. Comparative hydrolytic activities
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To get a better view of the catalytic efficiency of each of the plant extracts in hydrolysing
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sunflower oil, the reaction was kinetically monitored for 7 days. The hydrolysis kinetics
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obtained by quantification of the FFAs by HPTLC coupled to a densitometry system are
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shown in figure 1.
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According to the Figure 1, kinetics of fatty acid release by the hydrolysis of sunflower oil
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catalysed by different plant extracts are characterized by a rapid release of fatty acids during
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the first 48 hours, followed by a slowing down of the kinetics, reaching a plateau. Depending
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on plant extract sources, the kinetics curves have different amplitudes, and based on these
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results, a pre-classification regarding the efficiency of the tested plant extracts for TAG
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hydrolysis could be carried out. The three most efficient extracts for sunflower oil hydrolysis
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were those obtained from germinated seeds of Jatropha curcas with 62 % of fatty acids
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released after 48 hours followed by those obtained from germinated seeds of Moringa oleifera
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with 29% then those obtained from germinated seeds of shea nut (Vitellaria paradoxa) with
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15%. The others plant extract were not able to yield more than 10% of fatty acids.
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To our knowledge, this is the first time that plant extracts from Moringa oleifera and shea nut
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(Vitellaria paradoxa) are identified as biomasses with hydrolytic activity, comparatively to
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Jatropha curcas seeds extract which has been previously reported as a source of lipases 29, 30,
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35
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Jatropha curcas. The authors reported that extract from germinated seed of jatropha curcas
279
was able to hydrolyze over than 90% of a wide range of vegetable oils and biodiesel raw
280
materials.
281
III.3. Optimisation of hydrolysis conditions
282
Using the three most active extracts, germinated seeds of Jatropha curcas (JG), germinated
283
seeds of Moringa oleifera (MG) germinated seeds of shea nut (SG) the optimum hydrolysis
284
conditions in terms of pH and temperature were determined.
285
. De Sousa et al., 29 also observed a strong hydrolytic activity from germinated seeds of
III.3.1.
Optimisation of hydrolysis pH
286
The FFA yield with the three most active extracts was measured in the range of pH 2–10 at
287
40°C during 48 hours. The results are shown in Figure 2. The maximum amount of FFA
288
released was obtained with the extracts of germinated jatropha seeds (JG), the extracts of
289
germinated moringa seeds (MG) and with the extracts of germinated shea nut seeds,
290
respectively 83%, 59% and 17%. TAG hydrolysis was maximum when the medium pH was
291
around 7 with the extracts of germinated jatropha seeds (JG) and the extracts of germinated
292
moringa seeds (MG), and at a pH of 8 with the extracts of germinated shea nut seeds. The
293
optimum pH for TAG hydrolysis using the extracts of JG and MG was a pH close to neutral.
294
The optimum pH observed with the extracts of JG tallied with that reported in the studies by
295
Staubmann et al 30 and Abigor et al 35, who found that partially purified lipase extracts of
296
germinated jatropha seed had a maximum hydrolytic activity around a pH of 7.5. To our
297
knowledge, there have not been any studies published on the hydrolytic activities of shea nut
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or moringa lipases. The hydrolytic activity existing in the extracts of germinated shea nut
299
seeds was optimum in an alkaline medium. Other plant lipases described in the literature
300
display optimum activity with an alkaline pH, such as those from Pachira aquatica seeds 36,
301
rice seeds and cottonseeds 24. Staubmann et al 30 reported that they had isolated a lipase from
302
seeds of Jatropha curcas with an optimum pH of 8.5. Thus, the high hydrolysis yield found
303
between pH 6 and pH 9 in our study with the JG extracts might reflect the presence of several
304
lipases in the extract, unlike what was seen with the extracts of germinated moringa and shea
305
nut seeds.
306
III.3.2.
Optimisation of hydrolysis temperature
307
The optimum TAG hydrolysis temperature when using the three most active plant extracts was
308
determined by fixing the pH of each reaction medium at that corresponding to the optimum pH
309
for each extract, i.e. a medium pH of 7.2 for the JG and MG extracts, and a pH of 8 for the SG
310
extracts. According to the literature, the range of temperatures at which plant lipases remain
311
active is between 30 and 55°C. In the great majority of cases, the maximum catalytic activity
312
of plant lipases is observed at around 40°C 37-40. The TAG hydrolysis rates for the sunflower
313
oil obtained for the different temperatures and quantitatively analysed by HPTLC coupled to a
314
densitometry system are presented in Figure 3.
315
Maximum lipase activity was reached at 45°C with the three extracts. At that temperature,
316
TAG hydrolysis was maximum and the FFA released was over 46%, 28% and 7% for the
317
reaction catalysed by the JG, MG and SG extracts respectively. An increase in temperatures
318
between 30 and 40°C led to an increase in the hydrolysis rate. Beyond the threshold
319
temperature (45°C), increasing the temperature to 50 then 55°C, there was a significant drop in
320
the hydrolysis rate in the three cases. The increase in temperature caused molecular stirring,
321
which reached the very structure of the enzyme, leading to the breakage of the hydrogen bonds
322
that govern protein folding 41. The result is a loss of active conformation of the enzyme 14 ACS Paragon Plus Environment
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323
molecule. The optimum hydrolysis temperature of 45°C found with the JG extracts tallied with
324
that reported by Staubmann et al 30, but was higher than that found by Abigor et al who
325
showed that the optimum hydrolysis temperature for hydrolysis with partially purified jatropha
326
extracts was 37°C 35.
327
III.4.
Screening of transesterification activity
328
The transesterification tests were carried out using two different molar ratios. First using a
329
molar excess of alcohol compared to the TAG, i.e. a TAG:EtOH molar ratio of 1:27, then
330
using an excess of TAGs compared to ethanol, with a TAG:EtOH molar ratio of 2:1. From the
331
six extracts active in hydrolysis, four of them displayed a catalytic activity in
332
transesterification as shown by the qualitative analysis of the reaction medium by HPTLC.
333
These were the extracts obtained from germinated jatropha seed, shea nut seeds in both forms
334
(dormant and germinated), and the extracts obtained from germinated moringa seeds. The
335
extracts from germinated groundnut and desert date seeds, while previously active in TAG
336
hydrolysis, were inactive in transesterification under the two sets of operational conditions
337
tested. Two hypotheses help to explain these results: i) either the lipases contained in the
338
extracts previously active in hydrolysis are not adapted to ethanol transesterification, ii) or
339
those lipases did not find conditions conducive to their action. The reaction medium composed
340
of vegetable oil and anhydrous ethanol did not contain enough water to enable flexibility and
341
molecular folding of the lipases in their active form. In order to have active conformation,
342
minimum hydration of the enzyme is essential 42.
343
The transesterification tests with a large excess of ethanol compared to TAGs (TAG:EtOH
344
molar ratio of 1:27) did not give FAEEs either with the extracts of shea nut seeds in both
345
forms (dormant and germinated), or with the extracts obtained from germinated moringa
346
seeds, whereas the presence of FAEEs was found with the same extracts during the
347
transesterification tests where the TAGs were in excess compared to the ethanol. Only the 15 ACS Paragon Plus Environment
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extracts of germinated jatropha seeds seemed to result in FAEEs whatever the TAG:EtOH
349
molar ratio.
350
These results show that high ethanol concentrations had an inhibiting effect on the plant
351
extracts obtained from dormant and germinated shea nut seeds and the germinated moringa
352
seeds. This inhibiting effect was also found using extracts of germinated jatropha seeds, for
353
which weak catalytic activity was observed in the presence of a large excess of ethanol.
354
Catalytic activity was much greater when the initial quantity of ethanol was low. A low
355
TAG:EtOH molar ratio is therefore more appropriate for converting TAGs into FAEEs using
356
plant extracts obtained from germinated jatropha, moringa or shea nut seeds. In order to
357
maximize TAG conversion into FAEEs, while maintaining a low concentration of ethanol in
358
the medium, the method describe by Shimada 43, with stepwise addition of alcohol, should be
359
considered in transesterification optimization tests. This stepwise additions method consists in
360
gradually adding small quantities of alcohol at regular time intervals throughout the duration
361
of the reaction, to limit the alcohol concentration in the medium and thereby limit deactivation
362
of the lipases contained in the plant extracts. In the studies in which this method was used, it
363
proved its efficiency in converting TAGs into vegetable oil alkyl esters 14, 28, 43.
364
III.5.
Comparative transesterification activities
365
In order to gain a precise idea of the catalytic efficiency of the plant extracts active in
366
transesterification, the kinetics of a transesterification reaction with a TAG:EtOH molar ratio
367
of (2:1) were monitored for 48 hours. The transesterification kinetics obtained by the
368
quantification of FAEEs by HPTLC coupled to a densitometry system is shown in Figure 4.
369
A comparison of the kinetic study results showed that the most efficient extracts in
370
transesterification were classed in the same order as those in the hydrolysis tests
371
(JG>MG>SG). This result tends to confirm the hypothesis that we put forward whereby in
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372
order for the extracts to be active in transesterification they have to be active in hydrolysis.
373
The FAEE yields obtained with extracts of jatropha and moringa at 34% and 10% respectively
374
were too low for them to be used as it is for biodiesel production.
375
III.6.
Optimisation of transesterification conditions
376
The FAEE yields obtain during previous transesterification reactions can be improved by
377
optimizing the transesterification conditions. According to the literature, TAG:EtOH ratio,
378
EtOH water content and catalyst load are the parameters that considerably affect the catalytic
379
activity of lipases with an impact on transesterification yield 2, 44, 45. The optimization of the
380
TAG:EtOH ratio, water content and catalyst load were carried out with the two most active
381
extracts (JG and MG).
382
III.6.1.
TAG:EtOH molar ratio
383
As high ethanol concentrations inhibited the activity of the plant extracts, it was important to
384
determine the optimum molar ratio to catalyse the reaction. Some transesterification tests were
385
therefore carried out with TAG:EtOH molar ratios of (3:1), (2:1), (1:1), (1:2) and (1:3). The
386
quantities of FAEEs obtained using the JG and MG extracts as catalysts for the different
387
TAG:EtOH molar ratios are shown in Figure 5.
388
The results in Figure 5 show that changes in the quantity of FAEEs formed depending on the
389
amount of alcohol in the medium followed the same trends for the JG and MG extracts.
390
Between the TAG:EtOH molar ratios of (3:1) and (2:1), the number of moles of FAEEs
391
formed increased, with a maximum being reached for the TAG:EtOH molar ratio of (2:1).
392
Beyond that ratio the number of moles of FAEEs decreased. There was no FAEE synthesis
393
with the MG extracts from the TAG:EtOH molar ratio of (1:2). The JG extracts seemed less
394
sensitive to the presence of ethanol than the MG extracts and continued to produce FAEEs
395
beyond the TAG:EtOH molar ratio of (1:2), though the quantity of FAEEs became very low. 17 ACS Paragon Plus Environment
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The MG and JG extracts were more active for a TAG:EtOH molar ratio below (3:1). As
397
alcohol inhibition starts at much lower content than the stoichiometric value, in order to have a
398
complete transesterification of TAG and to avoid alcohol inhibition phenomena, the so-called
399
‘stepwise addition of alcohol’ method is recommended 14, 46.
400
Short carbon chain alcohols are known to have an inhibiting effect on the catalytic activity of
401
lipases 46, 47. According to Robles-Medina 48, the solubility of short-chain alcohols (case of
402
methanol and ethanol) is the limiting factor in the enzyme transesterification reaction with
403
lipase catalysis. As these alcohols are not completely soluble in oil, contact with the alcohol
404
dispersed in fine droplets causes enzyme inhibition. Two mechanisms are put forward to
405
explain this phenomenon: i) either the elimination by alcohol of the enzyme’s
406
microenvironment, which modifies the active conformation of the enzyme by dehydration 28,
407
or ii) the formation around the lipase (polar) proteins of a hydrophilic barrier made up of
408
alcohol molecules, which prevents hydrophobic TAGs from gaining access to the active site of
409
the enzyme. This hydrophilic barrier may also lead to the agglomeration of lipase proteins and
410
thereby reduce the availability of active sites accessible to the substrates.
411
III.6.2.
Ethanol water content
412
As dehydrating ethanol beyond 95% is complicated and very energy-intensive 12, the most
413
widely available and cheapest commercial bioethanol is generally partially hydrated. To
414
determine the optimum level of water content in ethanol for transesterification reaction with
415
JG and MG extracts as biocatalyst, transesterification tests were therefore carried out with
416
ethanol water contents varying from 0% to 15% (v/v). The results of the FAEE yields obtained
417
are shown in Figure 6.
418
An ethanol water content up to 1% for MG and up to 5% v/v for JG extracts did have a
419
positive effect on the formation of FAEEs, while more than 5% v/v water content was
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detrimental to the FAEE yield using JG or MG extracts (Fig. 6). MG extracts appear to be
421
more sensitive to water content in ethanol than JG extracts.
422
This is consistent with the study of Al-Zuhair et al. 49 and Shah et al., 50 who found that lipases
423
generally need a minimum amount of water in their microenvironment in order for them to be
424
in active catalytic conformation..
425
Over the optimum levels of 5% v/v and 1% v/v of water content in ethanol, for JG and MG
426
extracts respectively, increasing the water content led to a decrease of transesterification yield
427
for both plant extracts. Two phenomena might cause this drop in activity: i) either the increase
428
in the amount of water is conducive to the formation of TAG or FAEE hydrolysis reactions,
429
leading to a foreseeable drop in transesterification yield ii) and/or water causes the aggregation
430
of enzymatic plant extracts particles, thereby limiting substrate accessibility to the active sites
431
of the lipases.
432
In order to check the hypothesis whereby the increase in water content would seem to promote
433
the formation of hydrolysis reactions, changes in the quantity of FFAs formed were studied
434
according to increasing ethanol water contents during the transesterification reaction. The
435
results are presented in Figure 7 and it shows that in the 1-5% v/v H2O range, the quantity of
436
FFAs resulting from hydrolysis linked to the addition of water was very small. This quantity of
437
FFAs was under 2% of the FFAs initially present in the medium. Similar results were obtained
438
with the MG extracts (data not shown).
439
This suggests that the relative decrease of the quantity of FAEEs produced in the presence of
440
water (
