Enzymatic Transesterification of Coconut Oil Using Chitosan

Article pubs.acs.org/EF

Cite This: Energy Fuels XXXX, XXX, XXX-XXX

Enzymatic Transesterification of Coconut Oil Using ChitosanImmobilized Lipase Produced by Fluidized-Bed System T. A. Costa-Silva,*,† A. K. F. Carvalho,‡ C. R. F. Souza,† H. F. De Castro,‡ S. Said,† and W. P. Oliveira† †

Faculty of Pharmaceutical Sciences of Ribeirão Preto, University of São Paulo, 05508-000 São Paulo, Brazil Engineering School of Lorena, University of São Paulo, 05508-000 São Paulo, Brazil

‡

ABSTRACT: This work demonstrated the catalytic performance of Cercospora kikichii lipase immobilized onto chitosan acetate microparticles activated with different cross-linking agents and dried using a fluidized-bed system. The activating agent affected the lipase activity, and the highest immobilizing yield was achieved by the chitosan microparticles activated with 1.5% glutaraldehyde (93.7%) followed by activation with metaperiodate (78.9%) and epichlorohydrin (70.6%). The immobilized biocatalysts produced showed low moisture content and water activity, namely, 3.5% and 0.12, respectively; and high stability under storage conditions maintaining 77.7% of its initial activity after 6 months at 5 °C. The industrial applicability of the biocatalyst was assessed in the transesterification of coconut oil (Cocos nucifera oil) using ethanol as an acylant agent. The viscosity value for the coconut oil (29 mm2·s−1) sharply decreased to 3.2 mm2·s−1, upon the progress of transesterification reaction. This represents a final product containing high ester content (97.9%) and low levels of acylglycerides (0.7%). The stability of chitosan-immobilized lipase was also estimated, under successive batch runs, and after five reuse cycles the ester content remained above 96.5%. Therefore, the immobilization process developed and the immobilized derivative produced could represent an alternative route both for protein engineering (specially for enzyme stability) as for biodiesel production employing lipases.

1. INTRODUCTION Diesel is derived mainly from petroleum (fossil fuels) and, owing to population explosion and industrialization, is the fuel most consumed worldwide; the current demands and future trends in diesel fuel lead concerns are in the finite nature of fossil fuels, their negative impacts on the environment, and health and safety considerations, emphasizing the importance and necessity to develop renewable and environmentally friendly alternatives to fossil diesel.1 In addition, there is the critical impasse of global warming and atmospheric pollution generated by sulfur and carbon oxides and hydrocarbons, released on using these fuels. Therefore, the combined effect of the problems in a society whose development is strongly associated with fossil fuels use becomes an incentive for alternative energy sources to be of paramount relevance for policy makers, investors, and scientists worldwide.2 An appropriate source to replace petroleum-fueled engines is biodiesel, which is industrially obtained by chemical catalysts (especially homogeneous alkaline) due to their lower costs and faster kinetics. However, there are several limitations associated with this chemical route such as the need to use refined oils and excess reagents (alcohol and catalyst) to avoid reversible reactions, hardship in glycerol recuperation, and high energy cost.3 Hence, some alternative production routes have been studied, such as metallic complexes, oxides, organic bases, and enzymes. Biotechnology routes employing enzyme (heterogeneous catalysts) are highly attractive because of its exceptional selectivity, gentle operating requirement, absence of soap formation, simplified downstream processes, production of a cleaner glycerol, and environmental friendliness.4,5 The free enzymes utilization for biodiesel production results in technical limitations. Therefore, biocatalyst immobilization © XXXX American Chemical Society

plays a crucial role within applied enzymology, especially for biodiesel production, and the main reason is the ability to isolate the enzyme−support system from the reactive medium and to reuse it, increasing the rate yields and consequently reducing the production cost, easier downstream operations (easy product separation and simple glycerol recovery), the potential to run continuous processes via packed-bed reactors, and improvement of activity and stability in terms of mechanical, chemical, and thermal properties of the lipase.6,7 The enzymes’ properties would be affected depending on the type of immobilization technique and support used. Several methods for enzyme immobilization have been used by many researchers including adsorption onto solid supports, covalent attachment, and entrapment within sol−gel compounds.8−10 The choice of the support/carrier is a relevant aspect influencing the immobilized biocatalyst properties and consequently the feasibility and process cost.11 An overview of the recent scientific literature reveals that a wide range of materials have been applied for enzymes immobilization and the high cost of commercial matrices (synthetic polymers and silica-based supports) has led several researchers to look for cheaper substitutes such as sugar cane bagasse,11 CaCO3,12 rice husk,13,14 chitin,15 and chitosan.8 From these alternatives, the chitosan (a chitin derivative) appears to be more interesting since chitin is one of the most plentiful biopolymers on the planet. This material shows many benefits/advantages as enzyme immobilizing support: abundant and relatively inexpensive; several possibilities of chemical modification; Received: July 13, 2017 Revised: October 18, 2017

A

DOI: 10.1021/acs.energyfuels.7b02033 Energy Fuels XXXX, XXX, XXX−XXX

Article

Energy & Fuels

́ were of analytical grade acquired from Synth and Vetec Quimica Ltda. (Brazil). 2.2. Lipase Production and Enzymatic Activity. The plant Tithonia diversifolia known as the Mexican sunflower, an essential oil producer, was used in a screening of lipase endophytic microorganism producers, and C. kikuchii was isolated and selected as a good lipase source.11 For enzyme production, soybean oil was used as carbon source and purification steps were carried out using a butyl-sepharose column.9 The endophytic fungus lipase showed the following characteristics: optimum pH, 6.5; optimum temperature, 40 °C; specific activity, 223.6 U·mg−1; Vmax, 10.28 μmol·(min·mg)−1; KM, 0.0324 mM. The purified C. kikuchii lipase was used for the immobilization process and biodiesel production. Lipase activities (free and immobilized form) were measured using olive oil as substrate.19 One international unit of activity (1 IU) was defined as the amount of lipase that liberates 1 μmol of free fatty acid per minute. The protein content was carried out according to the method of Bradford.20 2.3. Chitosan Beads Preparation and Activation. For the immobilization process, a homogeneous chitosan acetate solution was obtained by stirring 4% chitosan in 5% acetic acid.21 The acid solution was dropped into a beaker containing 1 M NaOH solution with the aid of a syringe and peristaltic pump, and the beads formed were maintained in this alkaline solution for 24 h. The chitosan beads were washed using distilled water until neutral pH and dried in an air circulation oven. For the chitosan beads’ chemical modification, the formed acetylated product was added to phosphate buffer (50 mM, pH 6.5) containing the following cross-liking agents (1.5% (v/v)): glutaraldehyde, sodium metaperiodate, or epichlorohydrin. The suspensions were kept under 350 rpm stirring at 25 °C for 60 min. After this time the modified support was recovered (filtration) and washed with distilled water to remove the nonbound cross-linking agents. The activated supports were dried at 50 °C for 12 h, and the activated chitosan microparticles were used for lipase immobilization using a fluidized-bed dryer. 2.4. Lipase Immobilization and Drying Performance. The fungal lipase was immobilized by covalent binding onto chitosan beads previously activated with cross-linking agents using PEG-1500 (1.5% (m/v)) as a stabilizing agent. In an effort to save time and reduce costs, we associated the benefits of drying and immobilizing processes and performed them in a single step employing a fluidized-bed dryer. The immobilization and drying processes were conducted in a fluidized-bed system containing a cylindrical column (height of 300 mm and a diameter of 85 mm). Figure 1 shows a schematic diagram of the proposed fluidized-bed dryer system and the drying/immobilization mechanism. The lipase suspension was directly supplied into the fluidized-bed column where the activated chitosan beads were fluidized by hot air provided by a blower and air heater. The charged lipase suspension adhered covalently to the surface of activated chitosan particles. The fluidizing air flow rate was adjusted to 0.75 kg·min−1. When the drying system stabilized, the fluidizing air was heated to 60 °C (for all experimental runs) and the feeding of the lipase solution (2.0 mgprotein· gsupport−1) and the feed gas flow rate were regulated to 1.5 g·min−1. Table 2 presents the operating conditions of the dryer equipment. At the end, samples of the immobilized derivatives were collected and used to evaluate the fluidized-bed system performance and immobilized lipase properties. 2.5. Biodiesel Synthesis. The best immobilized derivative obtained (using glutaraldehyde as cross-linking agent) was evaluated in the synthesis of biodiesel. The transesterification reactions were performed in a screw-capped glass vessel (250 mL) filled with 12 g of coconut oil and ethanol at fixed oil to alcohol molar ratio of 1:12. The experiments were carried out with chitosan-immobilized lipase at proportions of 20% (w/w) in relation to the final weight of reagents. The biodiesel production was carried out for 120 h at 40 °C under continuous stirring (150 rpm). At the end of the reaction, the biocatalyst was recovered by centrifugation (4000g) at room temperature for 10 min. The liquid phase was transferred into a decanting funnel in which the same amount of distilled water was

high affinity toward the enzymes; biodegradability; and nontoxicity.16 During biodiesel production by the enzymatic route, the catalytic activity of the lipase can be changed by increasing or decreasing the water content of the reaction medium, mainly when shorter chain alcohols (methanol or ethanol) are used. The use of lipases in organic media requires a minimum amount of water, but its excess may favor substrate hydrolysis and promote diffusion limitations for hydrophobic substrates, thus decreasing the biofuel yield.5 So, a quantitative understanding on the interference of water content on biocatalyst activity/stability is fully justified. Lipase immobilization is normally performed in aqueous media, and the resulting immobilized derivative is frequently dehydrated before it is used in organic media, utilizing, for example, lyophilization process. However, it has been demonstrated that the dehydration step can induce irreversible enzyme inactivation.17 Thus, the development of new techniques for dehydrating the immobilized enzyme becomes highly relevant. Herein, the fluidized bed was used for drying of the immobilized enzyme in a single step. Therefore, the main objective of the present work was the obtainment of an immobilized form of Cercospora kikuchii (C. kikuchii) lipase, with advantageous biochemical properties and high stability. The enzyme was covalently immobilized onto a chitin derived support (chitosan) using cross-liking agents and dried by fluidized bed. The properties of the immobilized lipase were studied, and its catalytic activity was assessed in the biodiesel production using as the model system the transesterification of coconut oil.

2. MATERIALS AND METHODS 2.1. Materials. Bradford reagent, glutaraldehyde (25% solution), metaperiodate (78.9%), epichlorohydrin, and albumin were bought from Sigma-Aldrich (St. Louis, MO, USA); poly(ethylene glycol) (PEG-1500, Reagen, São Paulo, SP, Brazil) was used as a stabilizing agent. Olive oil (low acidity) from Carbonell (Spain) was purchased from a local market. Arabic gum was purchased from Synth (São Paulo, SP, Brazil). Coconut (Cocos nucifera (Co. nucifera)) oil was acquired from Frescoco (São Paulo, SP, Brazil), and its main properties are given in Table 1. The fatty acid composition and physicochemical properties were determined according to AOCS (American Oil Chemists’ Society) methods.18 The organic solvents were of standard laboratory grade from Synth. All the other reagents

Table 1. Properties and Fatty Acid Composition of Coconut Oil property

value

kinematic viscosity at 40 °C (mm /s) acidic value (mg of KOH/(g of oil)) saponification value (mg of KOH/(g of oil)) iodine value (g of I2/(100 g of oil)) peroxide value (mequiv/(kg of oil)) Fatty Acids Composition (wt %) 2

caprilic capric lauric myristic palmitic stearic oleic linoleic

C8:0 C10:0 C12:0 C14:0 C16:0 C18:0 C18:1 C18:2

29.0 ± 0.75 0.41 ± 0.81 238.0 ± 0.48 25.0 ± 0.92 0.36 ± 0.80 8.90 6.10 47.20 18.80 7.80 2.58 6.10 1.60

± ± ± ± ± ± ± ±

0.66 0.25 1.05 0.91 0.12 0.11 0.38 0.15 B


Article

Energy & Fuels

Aqua Lab 4Tev water activity meter (Decagon devices, USA) and an oven drying method for moisture.24,25 The morphology of the biocatalysts was analyzed by high-resolution scanning electron microscopy in a EVO Model 50 (Zeiss, Cambridge, U.K.), operating with accelerating voltage of 20 kV.12 The surface area and pore diameter analyses were performed by BET method.11 The operational stability of the selected biocatalyst was carried out in both hydrolysis and transesterification reactions by measuring the formed product (fatty acids or ethyl esters, respectively) at the end of each reaction in the batch systems (runs 1−5), taking the original activity as 100%.19 The recovered immobilized lipase was then washed with tert-butanol to remove any substrate or product eventually retained in the biocatalyst microenvironment as glycerol. 2.6.2. Purified Ethyl Ester Analysis. The composition of the biodiesel was analyzed by gas chromatography (Varian 3800, Varian, Inc. Palo Alto, CA, USA). Data collection and analyses were performed using Varian Star Data System software, version 6.0. Ethyl esters were identified and quantified by comparing their retention times to standard FAEE with ethyl tricosanoate as the internal standard.22 Total ester contents were also determined by nuclear magnetic resonance 1 H NMR spectroscopy (Varian spectrometer, model Mercury-300 MHz) taking the database generated using26

Figure 1. (A) Drying of lipase suspension in a fluidized bed of activated chitosan particles and (B) drying/immobilization mechanism.

%EE = [(A C4 × 8)/(Add + ee)] × 100

Table 2. Fluidized Bed Parameters Set for Immobilizing/ Drying of Enzyme−Support System fluidized-bed drying parameter

value

inlet gas temperature, Tgi (°C) fluidizing air flow rate, Wg (kg·min−1) enzyme composition feed flow rate, Ws (g·min−1) static bed height, H0 (cm) mass of inert material, Mpi (g) feed atomizer position

60.0 0.75 1.5 1.85 80.0 top spray

where AC4 = area of the quartet (fourth peak), Add+ee = area of all signals between 4.35 and 4.05, and %EE = FAEEs. According to eq eq 2, the ethoxyl hydrogen atoms signal of FAEEs split a quartet and AC4 was estimated by integrating the peak at 4.08 ppm.25 The area coincides to 1/8 of the complete ethoxy−-carbon hydrogen area (-OCH2 region varying from 4.05 to 4.20 ppm). The region nearby 4.08 ppm is the unique area where crossover does not appear, and this integrated signal can be attributed to FAEEs.28,29 The remaining amount of glycerides after the transesterification was determined in an Agilent 1200 Series liquid chromatograph (Agilent Technologies, USA) equipped with an Evaporative Light Scattering Detector and a Gemini C-18 (5 μm, 150 × 4.6 mm2, 110 Å) column at 40 °C.30 Viscosity and density values were analyzed by following the standard methods ASTM D 445 and ASTM D 4052, respectively. The concentration of water was measured by Karl Fischer methodology (AKF5000, K90365 model, Koehler Instrument Co., Inc.). 2.6.3. Biodiesel Properties Based on the Coconut Oil Fatty Acids Profile. In this study, biodiesel properties based on the fatty acids profile from coconut oil were estimated using the software “Biodiesel Analyzer version 1.1” (available on http://www.brteam.ir/ biodieselanalyzer).31

added. Then, vigorous stirring was carried out and the mixture was allowed to stand for 6 h, for phase separation. This procedure was performed three times, in sequence. The superior phase, consisting of fatty acid ethyl esters (biodiesel), was evaporated in a rotatory evaporator. Subsequently, the solution was dried sodium sulfate and the lower phase, consisting of glycerol and wastewater, was discharged.22,23 2.6. Analysis. 2.6.1. Immobilized Lipase Properties. Hydrolytic activities of free and immobilized lipase derivatives were assayed by the olive oil emulsion method.19 One unit (U) of enzyme activity was defined as the amount of enzyme that liberates 1 μmol of free fatty acid per min under the assay conditions (37 °C, pH 7.0, 150 rpm). Analyses of hydrolytic activities carried out on the lipase loading solution and immobilized preparation were used to determine the activity retention (RAE, %), as shown in eq 1.

RAE/% = 100 ×

immobilized enzyme activity/(U/mg) soluble enzyme activity/(U/mg)

(2)

3. RESULTS AND DISCUSSION 3.1. Biocatalysts’ Properties. The C. kikuchii lipase was immobilized onto chitosan by chemical attachment carried out in fluidized drying. Chitosan beads were first prepared and activated by cross-linking with glutaraldehyde, sodium metaperiodate, or epichlorohydrin. The activated chitosan beads were then used as a support material during the immobilization/drying process. The main purpose was to collaborate toward researching for a low-cost support capable of promoting

(1)

The storage stability of the immobilized lipases was estimated by determining the enzyme activity after 6 months of storage at 5 °C.16 The water content of immobilized derivative was determined using an

Table 3. Fluidized-Bed Performance and Product Properties: Residual Lipase Activity, Water Activity, Moisture Content, Activity after Reuse Cycles, and Stability after Storage cross-linking agent

Tins (°C)a

REA (%)b

Awc

moisture (%)

storage stability (%)d

operational stability (%)d,e

glutaraldehyde metaperiodate epichlorohydrin

42.1 41.5 42.5

93.87 ± 0.85 78.97 ± 0.50 70.60 ± 0.36

0.08 ± 0.07 0.14 ± 0.06 0.14 ± 0.04

5.74 ± 1.13 6.23 ± 1.07 6.08 ± 0.77

83.87 ± 0.15 77.27 ± 0.75 71.81 ± 0.80

92.0 ± 1.07 90.2 ± 0.97 91.1 ± 0.87

a

Tins, bed temperature. bREA, lipase recovered in the support. cAw, water activity. dStability tests: activity remained. The responses were calculated from triplicate analyses. eHydrolysis reaction using olive oil, 5 cycles. C


Article

Energy & Fuels

Figure 2. Scanning electron photomicrographs of the immobilized enzyme derivatives.

stabilization in the long term, mainly if it is taken into consideration that the enzyme in its free form (phosphate buffer) lost 83.4% of its initial activity under the same storage conditions. Lipase was immobilized onto a rice husk and dried using a spouted-bed system, and the residual enzyme activity of the immobilized derivative was an average of 75% of the initial activity, while for enzyme in phosphate buffer (free form) the final lipase activity was only 12%.14 The operational stability was also determined, and as can be observed in Table 3, the immobilized derivatives produced maintained an average of 91.1% of its initial lipase activity after 5 hydrolysis cycles using olive oil as substrate. These values of residual enzyme activity after 5 reuse cycles are interesting due to the practical and economical importance associated with biocatalysts’ use, mainly in biodiesel production. Chitosan from Syncephalastrum racemosum was used for lipase immobilization, and the derivative produced retained 47% of the catalytic activity after 4 reuse cycles.34 Chitosan was prepared for Ca. rugosa lipase immobilization, and the residual activity of immobilized derivative was about 46% at the end of 10 operation cycles.32 According to the results of the surface area and pore diameter analyses, the chitosan particles have the BET surface area of 325 m2·g−1 and total pore area 1.1 m2·g−1 with average pore diameter of 85 nm. After the activation with cross-linking agents a reduction of surface area to 301 m2·g−1 and pore volume to 0.87 m2·g−1 were verified, probably due to chemical modification in the chitosan surface responsible for covalent binding with C. kikucii lipase. Figure 2 shows the photomicrographs of the immobilized derivatives for supports activated with 1.5% glutaraldehyde (the procedure which gave the highest retention of enzyme activity). As expected, the beads showed high sphericity and smooth surface allowing a more homogeneous surface coated by the enzyme composition. The chitosan beads’ sizes varied from 50 to 100 μm in all experimental runs. Chitosan can be considered too fragile to use for support in the enzyme immobilization process, depending on the immobilization conditions and industrial application of immobilized derivative produced. However, the mechanical strength of the aforementioned material can be improved through irreversible chemical modification of its free amino groups with cross-linking agents. The chitosan chemical modifications have proven efficiency at improvement of mechanical properties and thermal stability.35 Herein, we use three cross-linking agents to activate the chitosan beads. These

excellent catalytic performance and stabilization (especially with low water content). Table 3 shows the effect of the three tested cross-linking agents used on fluidized-bed performance and product properties: residual lipase activity, water activity, moisture content, activity after reuse cycles, and stability during storage period. The residual enzymatic activities of all enzymatic formulations used in this study were in the range 70.6−93.8%. Among all formulations assessed, those prepared with 1.5% glutaraldehyde demonstrated the best performance because the final product maintained at least 93% of the initial enzyme activity. This result compared favorably with data reported in the literature; for example, Candida rugosa (Ca. rugosa) lipase was immobilized onto chitosan membrane by covalent binding (utilizing glutaraldehyde as binding reagent) and the final immobilized lipase activity was 49.8% enzyme in free form.32 Throughout the fluidized-bed drying process, the spray droplets and the immobilized lipase produced reached a maximum temperature value (Tgi) inferior to the fixed temperature used to warm the drying gas. Herein, the inlet temperature used was set at 60 °C and the average temperature inside the fluidized bed (Tins) was 42.0 °C. A cooling effect is reached as long as liquid evaporates from the system, and this phenomenon avoids the overheating of drying composition. This feature of a fluidized-bed dryer makes it a feasible method for dehydration of thermosensitive biomolecules such as enzymes. In general, excellent product stability and an acceptable protein shelf life are combined with small values of water content. In this study, the moisture content in the obtained biocatalysts ranged between 5.74 and 6.23%. An additional characteristic that is related to the lipase stability is water activity because higher values could provide suitable conditions for microorganism development and other undesirable reactions. The water activity values of the chitosan-immobilized lipase were in the range 0.08−0.14, which is considered safe to avoid the growth of microorganisms.33 Moreover, these moisture content and Aw values would favor the transesterification process yield. Based on the method of olive oil hydrolysis previously mentioned, Table 3 also displays the activity of the immobilized biocatalyst after 6 months storage at 5 °C. Depending on the activating agent, the resulting immobilized derivative shows slight differences, and on average 22.3% of activity lost was determined during the storage period. This result is evidence of the positive role of the immobilizing/drying process for enzyme D


Article

Energy & Fuels

Table 4. Ethyl Esters Profile Obtained in the Transesterification of the Coconut Oil with Ethanol Using C. kikuchii Lipase Immobilized onto Chitosan at 40 °C reaction time ethyl esters (%) ethyl ethyl ethyl ethyl ethyl ethyl ethyl ethyl

caprylate (C8:0) capronate (C10:0) laurate (C12:0) myristate (C14:0) palmitate (C16:0) stearate (C18:0) oleate (C18:1) linoleate (C18:2)

total (%)

24 h

48 h

72 h

96 h

120 h

6.23 ± 0.03 2.55 ± 0.02 24.06 ± 0.08 8.58 ± 0.09 2.15 ± 0.05 3.22 ± 0.09 4.57 ± 0.41 1.20 ± 0.16

7.57 ± 0.08 3.72 ± 0.04 30.74 ± 0.67 10.92 ± 0.13 2.5 ± 0.03 3.5 ± 0.03 6.37 ± 0.19 1.19 ± 0.01

7.51 ± 0.07 2.78 ± 0.24 37.39 ± 0.57 11.37 ± 0.17 3.19 ± 0.10 4.45 ± 0.18 6.58 ± 0.04 1.52 ± 0.04

7.63 ± 0.04 6.61 ± 0.41 38.29 ± 0.9 17.32 ± 0.43 4.15 ± 0.03 4.29 ± 0.02 6.57 ± 0.07 1.53 ± 0.02

7.73 ± 0.02 6.55 ± 047 42.17 ± 0.36 22.09 ± 0.03 6.45 ± 0.13 4.67 ± 0.21 6.41 ± 0.01 1.59 ± 0.09

52.56 ± 0.03

66.51 ± 0.02

74.79 ± 0.05

86.49 ± 0.01

97.96 ± 0.08

Figure 3. 1H NMR spectra triacylglycerol (a) and biodiesel (b) obtained by transesterification of coconut oil using lipase from C. kikuchii immobilized onto chitosan as catalyst.

evaluate the applicability of C. kikuchii immobilized lipase generated in this work. 3.2. Biodiesel Synthesis. In enzymatic biodiesel production, the ethanol utilization rather of methanol has conquered special interest. Ethyl alcohol has the quality of being produced from renewable raw material, differently from methanol which is petrochemically derived and, moreover, causes fewer lipase activity inactivation than methyl alcohol (low stearic effect). To make biodiesel more attractive and environmentally friendly, in

surface modifications may be responsible for the mechanical resistance of support during the drying process and biodiesel production since the beads’ size remained unchanged and no particle fragments were accumulated in the collection bottle. The fluidized-bed drying process produced immobilized derivatives with high enzyme activity and low water content, features suitable for transesterification reaction using organic media. So, the enzymatic synthesis of biodiesel was chosen to E


Article

Energy & Fuels

value of used oil: 29−3.2 mm2·s−1. High-viscosity values lead to weaker fuel spray atomization and less precise fuel injectors operation.37 Herein, the kinematic viscosity obtained for biodiesel was within the limit established by the standard agencies European Union (EN 14,214), USA (ASTM D6715), and Brazilian (ANP) specifications. The amounts of mono(0.3%) and diacylglycerol (0.4%) are also in accordance with the ASTM standard for biodiesel (ASTM D6751) that establishes values up to 1.0% for these compounds. Other proprieties for the biodiesel can be also verified to estimate its quality, taking into consideration the feedstock composition in fatty acids as shown in Table 6. In contrast with

addition of the use of ethanol as an acyl acceptor, the coconut oil was chosen for biodiesel production due to its high abundance in northeast Brazil and may improve the socioeconomic aspects of the coconut producer region due to generation of employment and revenues. Besides, coconut oil could be a residue of the production-grated coconut and coconut milk and thus would not compete with food production. The saturation and chain length are important features of the feedstock which could be associated with the properties and quality of biodiesel produced. The coconut oil used in this study showed a low composition of unsaturad fatty acids and the prevalence of lauric acidhigh concentration of short carbon chain fatty acids (Table 1). This fatty acids composition profile enables the use of this feedstock for enzymatic biodiesel production due to lower stearic effect which allows more effective interaction among biocatalyst and the acyl acceptor (ethanol) during transesterification process.22 The coconut oil was subjected to transesterification process applying as catalyst the C. kikuchii lipase immobilized onto chemically activated chitosan acetate beads.36 The ethanolysis reaction progress regarding ethyl esters (wt %) is shown in Table 4. The highest FAEE content (97.96 wt %) was attained at 120 h of reaction, and no reversible reaction was observed during the process. By comparing Table 1 and Table 4, a correlation between the fatty acids found in coconut oil and ethyl esters composition is clearly evidenced. Laurate and myristate esters showed the highest concentrations as expected, since lauric and myristic fatty acids are found in major proportion in coconut oil. This result acquiesced with studies performed by lipases from C. kikuchii and Mucor circinelloides in the transesterification of non-edible vegetable oils, such as coconut and/or macaw palm oils.14,29 This FAEE concentration was confirmed by carrying out the 1 H NMR analysis.26 Figure 3 shows the 1H NMR spectra of the used feedstock and the biodiesel sample. It can be observed that the transesterification process produced the characteristic spectra etoxyl hydrogen of ethyl ester molecule. When juxtaposed with the coconut oil spectra, it can be verified that there is no overlap with the signals of the triglyceride compounds, evidencing excellent ethyl esters production.27 The area of the integration peaks between 4.06 and 4.35 ppm showed an AC4 (area of the component fourth peakquartet), of 1.17, Add+ee (area of all signals between 4.35 and 4.05) of 0.14 and concentration reached 97.45%, corroborating with the results obtained through gas chromatography analysis. Important properties that are related to biodiesel quality, such as kinematic viscosity, density, and residual glycerides, are shown in Table 5. The sample of biodiesel showed a pronounced decrease in viscosity compared with the initial

Table 6. Biodiesel Estimated Properties: Comparison of the Predicted Value of Biodiesel Samples from Coconut and Soybean Oils predicted valuea

a b

coconut oil

soybean oilb

limitsc

specificationc

SFA (%) MUFA (%) PUFA (%) DU CN LCSF CFPP (°C) CP (°C) PP (°C) APE BAPE OS (h) HHV

91.38 6.1 1.6 9.3 65 2.1 −9.974 −0.889 −7.786 9.83 1.6 76.3 37.101

15.6 22.7 60 142 43 3.18 −6.486 0.952 −5.788 142.7 67.4 4.556 38.74

ns ns ns ns 47, min. ns 19, max. report −15−10 ns ns 3, min. report

ns ns ns ns ASTM D6751 ns NBR 15512 ASTM D6751 ASTM D6751 ns ns ASTM D6751 ASTM D6751

Calculated using the software Biodiesel Analyzer version 1.1.30 Sajjadi; Raman; Arandiyan, 2016. cns, not a specified limit.

soybean oil, a raw material normally used for biodiesel production, coconut oil has a high ratio of saturated ones (SFA) to unsaturated fatty acids (MUFA and PUFA). On the other hand, coconut oil has a profile similar to those of babassu and macaw palm oils, also used for biodiesel production, both having lauric acid as a predominant component in their compositions.14,38 Therefore, the low values for the degree of unsaturation (DU) and iodine value (IV) calculated for coconut oil are due to the low values of allylic position equivalent (APE) and bis-allylic position equivalent (BAPE). This corresponds to high values of oxidation stability (OS = 76.3 min), turning the coconut biodiesel into an advantageous resource to be used in hot climate regions such as Brazil, because its oxidative stability value is greater than the required specification value, suggesting less susceptibility to oxidation (biofuel depreciation) besides avoiding the use of antioxidants, providing a relatively cheap product with significant quality compared to biodiesel obtained from different raw materials. The immobilized biocatalyst was also evaluated, under consecutive batch runs, for operational stability (ability to reuse the biocatalyst) during the enzymatic transesterification reactions. Satisfactory performance of the biocatalyst was observed, confirming, after five consecutive batches, the FAEE levels remained above 96.5%. 3.3. Data Comparison with Reported Studies. The Table 7 displays reported data concerning biodiesel production using lipase immobilized with different procedures. Several

Table 5. Properties of Purified Biodiesel Obtained from Transesterification of Coconut Oil Using Lipase from C. kikuchii Immobilized onto Chitosan ester contents (wt %) monoacylglycerides (wt %) diacylglycerides (wt %) triacylglycerides (wt %) kinematic viscosity at 40 °C (mm2·s−1) density at 20 °C (kg·m−3) water content (%)

biodiesel property

97.96 ± 0.08 0.3 ± 0.1 0.4 ± 0.1 0 3.2 ± 0.1 886 ± 0.4 0.03 ± 0.002 F


Article

Energy & Fuels Table 7. Production of Biodiesel Using Immobilized Lipase Enzyme by Various Immobilization Techniques support

lipase source

oil source

esters (%)

ref

chitosan chitosan chitosan chitosan chitosan chitosan chitosan alginate olive pomace acrylic resin textile membrane sol−gel support celite Nb2O5 rice husk acrylic resin

Cercospora kikuchii Candida rugosa Candida rugosa porcine pancreatic Candida rugosa Candida rugosa Talaromyces thermophilus Pseudomonas f luorescens Thermomyces lanuginosus Candida antartica Candida sp. Pseudomonas cepacia porcine pancreatic Burkholderia cepacia Cercospora kikuchii Candida antarctica

coconut oil rapeseed soapstock salicornia oil coconut oil soybean oil cooking oil waste frying oil Jatropha oil pomace oil soybean oil lard soybean oil sunflower oil beef tallow coconut oil microbial oil

97.9 63.6 55.0 12.0 87.0 72.2 >92 72.0 93.0 >90 >87 60.0 >80 42.2 >96 93.0

this article 39 40 41 42 43 44 45 46 47 48 49 50 51 14 29

the immobilized derivative produced could represent an alternative route for biodiesel production employing lipases.

materials were evaluated as carrier for enzyme immobilization, and all immobilized derivatives showed a lesser conversion rate compared with that one achieved by chitosan-immobilized lipase produced by the fluidized-bed system. The biodiesel production by enzymatic route enables one to eliminate or at least decrease the disadvantages of the transesterification process performed by chemical route (alkalior acid-catalyzed biodiesel production). Notwithstanding, industrial scale commercialization of the enzyme-catalyzed biodiesel remains uncertain, due to the high cost of the biocatalyst. Thereby, the development of innovative strategies for the increment of inexpensive and efficient technologies for enzyme biodiesel synthesis is of crucial interest. In this study were used two different strategies that could reflect on the production cost and present multifold advantages: the enzyme immobilization process and drying technology. Using fluidizedbed drying for obtainment of stable immobilized biocatalyst with low water content could ensure continuous use of the enzyme, decrease the biodiesel contamination, and improve the glycerol recovery. The drying step of the immobilized derivative is very important for the following reasons: in the presence of water could occur hydrolysis of the substrate and promotion of diffusion limitations, decreasing the biodiesel yield; and lipase demands a very small quantity of water molecules to retain its active structural configuration.3 Finally, the lipase source is original and the support used is considered cheap, so the immobilized derivative produced has potential to become a cost-effective alternative for the commercial lipases currently in the market.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

T. A. Costa-Silva: 0000-0002-7814-9257 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This study was supported by the State of São Paulo Research Foundation (FAPESP). T.A.C.-S. received a Ph.D. fellowship from FAPESP (Grant No. 2011/00743-8).

■

REFERENCES

(1) D̵ urišić-Mladenović, N.; Kiss, F.; Škrbić, B.; Tomić, M.; Mićić, R.; Predojević, Z. Renewable Sustainable Energy Rev. 2018, 81, 280−291. (2) Nigam, S.; Mehrotra, S.; Vani, B.; Mehrotra, R. Inter. J. Renewable Energy & Biofuels 2014, 2014, 1−16. (3) Ribeiro, B. D.; Castro, A. M. d.; Coelho, M. A. Z.; Freire, D. M. G. Enzyme Res. 2011, 2011, 615803. (4) Christopher, L. P.; Kumar, H.; Zambare, V. P. Appl. Energy 2014, 119, 497−520. (5) Meunier, S. M.; Kariminia, H.-R.; Legge, R. L. Immobilized enzyme technology for biodiesel production. In Advances in biofeedstocks and biofuels: Production technologies for biofuels; Singh, L. K., Chaudhary, G., Eds.; John Wiley & Sons: Hoboken, NJ, USA, 2011; pp 67−106, DOI: 10.1002/9781119117551.ch3. (6) Stoytcheva, M.; Montero, G.; Toscano, L.; Gochev, V.; Valdez, B. The Immobilized Lipases in Biodiesel Production. In Biodiesel Feedstocks and Processing Technologies; Stoytcheva, M., Montero, G., Eds.; INTECH: Rijeka, Croatia, 2011; Chapter 19, DOI: 10.5772/ 25246. (7) Costa-Silva, T. A.; Cognette, R. C.; Souza, C. R. F.; Said, S.; Oliveira, W. P. Drying Technol. 2013, 31, 1756−1763. (8) Carneiro, L. A. B. C.; Costa-Silva, T. A.; Souza, C. R. F.; Bachmann, L.; Oliveira, W. P.; Said, S. Braz. Arch. Biol. Technol. 2014, 57, 578−586. (9) Costa-Silva, T. A.; Souza, C. R. F.; Oliveira, W. P.; Said, S. Braz. J. Chem. Eng. 2014, 31, 849−858. (10) Zanin, G. M.; de Moraes, F. F. In Enzymes as biotechnology agents; Said, S., Pietro, R. C. L. R., Eds.; Legis Summa: Ribeirão Preto, Brazil, 2004, Chapter 4.

4. CONCLUSION This study revealed the viability of use of the fluidized-bed dryer in dehydration of thermally sensitive biomaterial such as enzymes. The possibility to produce dried immobilized biocatalysts, with excellent maintenance of enzyme activity, in a single step, also was demonstrated. The immobilized derivative produced showed high stability during a shelf life period and low moisture content and water activity, both characteristics considered suitable for reactions carried out in organic media. The immobilized biocatalyst demonstrated feasibility for industrial biodiesel production, since it was able to convert 97.9% of the coconut oil into biodiesel of good quality. Therefore, the immobilization process developed and G


Article

Energy & Fuels (11) Costa-Silva, T. A.; Souza, C. R. F.; Said, S.; Oliveira, W. P. Afr. J. Biotechnol. 2015, 14, 3019−3026. (12) Rosu, R.; Iwasaki, Y.; Shimizu, N.; Doisaki, N.; Yamane, T. J. J. Biotechnol. 1998, 66, 51−59. (13) de Castro, H. F.; de Lima, R.; Roberto, I. C. Biotechnol. Prog. 2001, 17, 1061−1064. (14) Costa-Silva, T. A.; Carvalho, A. K. F.; Souza, C. R. F.; De Castro, H. F.; Said, S.; Oliveira, W. P. Energy Fuels 2016, 30, 4820−4824. (15) Gomes, F. M.; Pereira, E. B.; De Castro, H. F. Biomacromolecules 2004, 5, 17−23. (16) Krajewska, B. Enzyme Microb. Technol. 2004, 35, 126−39. (17) Samborska, K.; Witrowa-Rajchert, D.; Gonçalves, A. Drying Technol. 2005, 23, 941−953. (18) American Oil Chemists’ Society (AOCS). Official Methods and Recommended Practices of the AOCS, 5th ed.; AOCS Press: Champaign, IL, USA, 2004, . (19) Soares, C. M. F.; De Castro, H. F.; De Moraes, F. F.; Zanin, G. M. Appl. Biochem. Biotechnol. 1999, 79, 745−757. (20) Bradford, M. M. Anal. Biochem. 1976, 72, 248−254. (21) Rorrer, G. L.; Hsien, T. Y.; Way, J. D. Ind. Eng. Chem. Res. 1993, 32, 2170−2178. (22) Carvalho, A. K. F.; Da Rós, P. C. M.; Teixeira, L. F.; Andrade, G. S. S.; Zanin, G. M.; De Castro, H. F. Ind. Crops Prod. 2013, 50, 485− 493. (23) Urioste, D.; Castro, M. B. A.; Biaggio, F. C.; Castro, H. F. Quim. Nova 2008, 31, 407−412. (24) Norena, C. Z.; Hubinger, M. D.; Menegalli, F. C. Bol. Soc. Bras. Ciênc. Tecnol. Aliment. 1996, 30, 91−96. (25) World Health Organization (WHO). Quality Control Methods for Medical Plants Materials; World Health Organization: Geneva, Switzerland, 1998; pp 235−389. (26) Paiva, E. J. M.; da Silva, M. L. C. P.; Barboza, J. C. S.; de Oliveira, P. C.; de Castro, H. F.; Giordani, D. S. Ultrason. Sonochem. 2013, 20, 833−838. (27) Rosset, I. G.; Tavares, M. C. H.; Assaf, E. M.; Porto, A. L. M. Appl. Catal., A 2011, 392, 136−142. (28) Sarpal, A. S.; Teixeira, C. M. L. L.; Silva, P. R. M.; da Costa Monteiro, T. V.; da Silva, J. I.; da Cunha, V. S.; Daroda, R. J. Appl. Microbiol. Biotechnol. 2016, 100, 2471−2485. (29) Carvalho, A. K. F.; Rivaldi, J. D.; Barbosa, J. C.; De Castro, H. F. Bioresour. Technol. 2015, 181, 47−53. (30) Andrade, G. S. S.; Carvalho, A. K. F.; Romero, C. M.; Oliveira, P. C.; De Castro, H. F. Bioprocess Biosyst. Eng. 2014, 37, 2539−2548. (31) Talebi, A. F.; Tabatabaei, M.; Chisti, Y. Biofuel Res. J. 2014, 2, 55−57. (32) Huang, X.-J.; Ge, D.; Xu, Z.-K. Eur. Polym. J. 2007, 43, 3710− 3718. (33) Beauchat, L. R. Cereal Foods World 1981, 26, 345−349. (34) Amorim, R. V. S.; Melo, E. S.; Caneiro-da-Cunha, M. G.; Ledinghan, W. M.; Campos-Takaki, G. M. Bioresour. Technol. 2003, 89, 35−39. (35) Wafiroh, S.; Wathoniyyah, M.; Abdulloh, A.; Rahardjo, Y.; Zakki Fahmi, M. Chem. Chem. Technol. 2017, 11, 65−70. (36) Moreira, A. B. R.; Perez, V. H.; Zanin, G. M.; De Castro, H. F. Energy Fuels 2007, 21, 3689−3694. (37) Knothe, G. Fuel Process. Technol. 2005, 86, 1059−1070. (38) Da Rós, P. C. M.; Silva, W. C.; Grabauskas, D.; Perez, V. H.; De Castro, H. F. Ind. Crops Prod. 2014, 52, 313−320. (39) Shao, P.; Meng, X.; He, J.; Sun, P. Food Bioprod. Process. 2008, 86, 283−289. (40) Desai, P. D.; Dave, A. M.; Devi, S. Food Chem. 2006, 95, 193− 199. (41) Cubides-Roman, D. C.; Pérez, V. H.; De Castro, H. F.; Orrego, C. E.; Giraldo, O. H.; Silveira, E. G.; David, G. F. Fuel 2017, 196, 481− 487. (42) Xie, W.; Wang, J. Biomass Bioenergy 2012, 36, 373−380. (43) Nasratun, M.; Said, A.; Noraziah, A.; Abd Alla, A. N. Am. J. Appl. Sci. 2009, 6, 1653−1657.

(44) Romdhane, I. B-B.; Romdhane, Z. B.; Bouzid, M.; Gargouri, A.; Belghith, H. Appl. Biochem. Biotechnol. 2013, 171, 1986−2002. (45) Devanesan, M. G.; Viruthagiri, T.; Sugumar, N. Afr. J. Biotechnol. 2007, 6, 2497−2501. (46) Yücel, Y. Bioresour. Technol. 2011, 102, 3977−3980. (47) Du, W.; Xu, Y.; Liu, D.; Zeng, J. J. Mol. Catal. B: Enzym. 2004, 30, 125−9. (48) Lu, J.; Nie, K.; Xie, F.; Wang, F.; Tan, T. Process Biochem. 2007, 42, 1367−70. (49) Meunier, S. M.; Legge, R. L. J. Mol. Catal. B: Enzym. 2010, 62, 53−57. (50) Yesiloglu, Y. J. Am. Oil Chem. Soc. 2004, 81, 157−60. (51) Da Rós, P. C. M.; Silva, G. A. M.; Mendes, A. A.; Santos, J. C.; De Castro, H. F. Bioresour. Technol. 2010, 101, 5508−5516.

H


Enzymatic Transesterification of Coconut Oil Using Chitosan

Recommend Documents