Enzymatic Production of Biodiesel from Nannochloropsis gaditana

Jul 3, 2015 - The commercial production of biodiesel is based on the transesterification of triglycerides and/or the esterification of free fatty acid...
14 downloads 6 Views 6MB Size
Download PDF
Article pubs.acs.org/EF

Enzymatic Production of Biodiesel from Nannochloropsis gaditana Microalgae Using Immobilized Lipases in Mesoporous Materials Luis Fernando Bautista,* Gemma Vicente, Á lvaro Mendoza, Sara González, and Victoria Morales Department of Chemical and Energy Technology, Chemical and Environmental Technology, Mechanical Technology and Analytical Chemistry, Universidad Rey Juan Carlos, C/Tulipán s/n, E-28933 Móstoles, Madrid, Spain S Supporting Information *

ABSTRACT: A study of the production of fatty acid methyl and ethyl esters (FAMEs and FAEEs) to be used as biodiesel was carried out with Nannochloropsis gaditana oil using three fungal lipases (from Thermomyces lanuginosus, Candida antarctica B, and Mucor miehei) both free and immobilized in hexagonal (SBA-15 and MCM-41) and cubic (FDU-12 and SBA-16) mesoporous supports. The operating variables were optimized using free lipase (ethanol, 40 °C, 24 h, 500:1 oil/lipase mass ratio, and 8:1 ethanol/oil mass ratio). Higher FAEE yields were obtained with supported lipase than with free lipase because of the protection and stability given by the supports. The FAEE yields achieved were higher for lipase immobilized in hexagonal mesostructured materials because lipase molecules are more accessible than lipase immobilized in the three-dimensional cubic supports. C. antarctica lipase B immobilized in functionalized SBA-15 showed the better performance and reusability among the biocatalysts used. biofuels.8,9 For this study, we used the microalga Nannochloropsis gaditana, which is an autotrophic unicellular alga, having an ellipsoidal shape of 3.5−4 × 2.5−3 μm, approximately. Its main use is in aquaculture, but it has been of great interest for the production of biodiesel in recent years.9,10 The most commonly used catalysts for first-generation biodiesel production are basic compounds, such as NaOH or KOH. The problem of using these catalysts is the neutralization of FFAs when the amount of FFAs in the oil is greater than 3 wt %,11 which is very common in microalgal oil. An alternative is the use of acid catalysts, such as sulfuric or hydrochloric acid and acidic organic resins,12 because these catalysts can be used with a higher concentration of FFAs in the oil. However, the reaction rate is slower than with a basic catalyst. In addition, potential problems with this kind of catalyst are the corrosion of the equipment in which the reaction is carried out, the larger energy consumption because of high-temperature conditions, or the production of water, which shifts the equilibrium of the reaction. Furthermore, the above two types of catalysts have the additional problem of the formation of secondary products, such as inorganic salts, which must be treated, thus increasing the costs and energy consumption of the process. Currently, enzymes (mainly lipases) are being used as an alternative to some industrial conventional processes.13 Lipases have been used for hydrolysis, alcoholysis, esterification, and transesterification processes because of their excellent catalytic activity and stability in non-aqueous media, which is beneficial for these reactions. As advantages, lipases are highly selective and capable of working with high amounts of water14 and FFAs,15 reduce energy costs because of the mild temperature

1. INTRODUCTION There is worldwide interest in biofuels as a renewable energy source for long-term fuel sustainability because of the predictable depletion of fossil fuels. The world is increasingly accepting the fact that conventional sources of fuel and energy are being rapidly depleted and cannot be renewed. In this sense, biomass stands out as a fully renewable resource that is used for the production of biofuels, power, chemicals, and materials, generating virtually no net greenhouse gases. What is needed is a safe, reliable, and efficient process of generating renewable biofuels that can potentially reduce fossil fuel dependence. One of these alternatives is the production of biodiesel, a common term for long-chain fatty acid methyl and ethyl esters (FAMEs and FAEEs, respectively). The commercial production of biodiesel is based on the transesterification of triglycerides and/or the esterification of free fatty acids (FFAs) in the presence of an alcohol and a catalyst. Biodiesel can be produced from different organic materials, such as edible seed oils (sunflower, rapeseed, palm, peanut, etc.),1 inedible oils (castor, Jatropha, etc.),2,3 used oils and fats, waste sludge, or animal remains.4 These materials have some drawbacks, such as competition with human food, land use, lack of a constant supply, and heterogeneity in the composition. Oleaginous microorganisms have started to be used to produce biodiesel in recent years.5 These organisms do not compete with human food and have a relatively stable composition,6 and their availability can be constant along time. Microalgae are photosynthetic microorganisms, prokaryotes or eukaryotes, which exist in all ecosystems on Earth. It is estimated that there are around 50 000 microalgal species, about 30 000 of which are known at the moment.7 They have the advantage of a larger biomass production than other agricultural resources; their life cycle is short; and they can reach concentrations up to 85 wt % of lipids based on dry weight, making them very suitable for the production of © 2015 American Chemical Society

Received: December 17, 2014 Revised: July 3, 2015 Published: July 3, 2015 4981

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels required (20−50 °C), and avoid process wastes. The biggest bottlenecks for the industrial application of lipases are their higher cost and reaction times required in comparison to conventional catalysts.16 Immobilization methods have been introduced to improve lipase stability, allowing for repeated utilization. After the pioneering work of Chibata and coworkers,17 the immobilization of enzymes has attracted worldwide interest to develop industrial applications. One of the main drawbacks in the use of immobilized enzymes is the mass-transfer limitation because of internal diffusion of substrates and products through the solid matrix. The use of ordered mesoporous silicas as support may help to overcome this problem because they have good connectivity throughout the pore network18 as well as an optimum control of the textural properties, allowing for the tailoring of the mesostructure for specific applications.19−22 These mesoporous silica materials have been recently used for the immobilization of lipases for biotransformations of triglycerides from refined vegetable oils.23 However, immobilization of lipases on mesoporous silica supports has not been reported thus far for biodiesel production using more complex oils, such as those extracted from microalgae. According to the interaction between enzyme and support, methods for enzyme immobilization are classified as adsorption, covalent bonding, entrapment, and cross-linking. The selection of the method and the support is a key factor for obtaining an efficient lipase activity. Comparative studies revealed that the same lipase molecule can show very different catalytic activities depending upon the supports used for the immobilization.18 The use of oleaginous microorganisms as raw materials for FAEE production by enzymatic catalysis is quite recent. Thus, Li et al. reached yields of 98.1 wt % using Chlorella protothecoides oil and lipase from Candida sp.24 Lai et al. reached a 90.7 and 86.2 wt % yield with Chlorella sp. oil in ionic liquids by methanolysis with Candida antarctica and Penicillium expansum lipases, respectively.25 Tran et al., using Chlorella vulgaris oil in n-hexane and Burkholderia sp. lipase as a catalyst, achieved yields of 97.3 wt %.26 Zhao et al. achieved 88.4 wt % yield in tert-butanol media27 with the yeast Rhodosporidium toruloides oil using commercial Thermomyces lanuginosus lipase. In this work, we studied the production of FAEEs from N. gaditana oil using three different fungal lipases from C. antarctica B (CalB), Mucor miehei, and T. lanuginosus. The enzymes were immobilized, through covalent binding to avoid denaturation in the highly polar alcoholic media, in four different mesoporous silica materials (SBA-15, SBA-16, MCM41, and FDU-12) to know the effect of the pore structure morphology on the overall lipase-based biocatalyst activity. This work reports, for the first time, the production of FAEE from microalgal lipids using immobilized lipases on mesoporous silicas. Specifically, we studied the production of FAEE from the marine oleaginous microalga N. gaditana using three different fungal lipases from C. antarctica B (CalB), M. miehei, and T. lanuginosus.

propyltriethoxysilane (APTES) was purchased from ABCR. Bradford reagent was acquired from Bio-Rad. C. antarctica lipase B (L3170), M. miehei lipase (L04277), and T. lanuginosus lipase (L0777) were supplied by Sigma-Aldrich (Madrid, Spain) in the form of aqueous solution containing the following protein concentration measured by the Bradford assay: 20.5, 11.2, and 16.6 mg/mL, respectively. 2.2. Lipid Extraction and Characterization. Lipid extraction from the N. gaditana microalga was carried out under reflux with methanol for 2 h in a nitrogen atmosphere.10 The sample was filtered, and the solvent was evaporated in a Laborota 4000 rotary evaporator (Heidolph, Schwabach, Germany). The composition of the oil was determined by thin-layer chromatography (TLC) using a method described elsewhere.28 The fatty acid profile was calculated according to ISO 5509 and EN 14103 standards. The iodine value was determined according to the EN ISO 3961 standard. 2.3. Synthesis of Mesoporous Supports. 2.3.1. SBA-15. The method used for the synthesis of SBA-15 is described by Zhao and coworkers.29 In a typical synthesis, 8 g of block copolymer surfactant Pluronic 123 were dissolved at room temperature under stirring in 250 mL of 1.9 M HCl. The solution was heated to 40 °C, and 17.8 g of TEOS was added to the solution. The resultant mixture was then stirred at 40 °C for 20 h and hydrothermally aged at 110 °C for a further 24 h. The template was removed by calcination at 550 °C for 5 h at a heating rate of 1.8 °C min−1 from room temperature. 2.3.2. MCM-41. The synthesis of MCM-41 was accomplished following the method described by Lin and co-workers,30 in which CTAB (2.53 g), dimethylamine (7.83 g), and H2O (95.3 g) were mixed and stirred in a beaker at room temperature until complete dissolution. Then, TEOS (9.84 g) was dropped into the transparent solution, and the resultant gel was stirred for 4 h at room temperature. The mixture was transferred to an autoclave for aging under static conditions at 110 °C and autogenous pressure for 48−72 h. The resultant materials were then recovered by filtration and air-dried. The template was removed by calcination at 550 °C for 5 h at a heating rate of 1.8 °C min−1 from room temperature. 2.3.3. SBA-16. SBA-16 silica was synthesized by the method described by Kim and co-workers31 using poly(ethylene oxide)− poly(propylene oxide)−poly(ethylene oxide) triblock F127 copolymer as a supramolecular template. Briefly, F127 copolymer was dissolved into a solution of hydrochloric acid in distilled water. TEOS was then added. The starting molar composition was 0.0040 F127/1.0 TEOS/ 4.0 HCl/130 H2O. The mixture was magnetically stirred for about 15 min at 35 °C until TEOS was completely dissolved. The mixture was placed in an oven for 24 h under static conditions at 35 °C for precipitation of the product. The mixture was kept at 100 °C for hydrothermal treatment during 24 h. The white solid precipitate was isolated by filtration and dried at 100 °C. The solid material was washed with methanol and, subsequently, calcined in air at 550 °C to remove the copolymer template. 2.3.4. FDU-12. The synthesis of FDU-12 material was accomplished using the method described by Kruck and co-workers.32 The molar ratios TEOS/Pluronic F127/TMB/KCl/HCl/H2O in the synthesis medium were 1:0.0037:0.5:3.36:6.08:165. The synthesis was carried out as follows: 3 g of Pluronic F127 copolymer were placed in a polypropylene (PP) bottle, and 185 mL of 1.97 M HCl was added. The mixture was placed in an open PP bottle submerged in a water bath at 15 °C and mechanically stirred. Then, 4.2 mL (3.6 g) of TMB and 15.0 g of KCl were introduced. After 2 h of constant stirring, 13.3 mL (12.5 g) of TEOS were added and the mixture was again stirred for 24 h in an open container at 15 °C. Afterward, the PP bottle with the reaction mixture was closed and placed in an oven at 100 °C for further 24 h. As-synthesized material was subjected to an additional hydrothermal treatment in an acid solution. Thus, 0.5 g of the material was placed in 30 mL of 1.97 M HCl solution kept at 110 °C (in a PP bottle) for 48 h. The resulting material was filtered, dried, and calcined under air at 550 °C for 5 h with a heating ramp of 2 °C min−1. 2.4. Immobilization of Lipase in Mesoporous Materials. APTES was used in the amine functionalization process to enhance the loading and activity of the immobilized lipase. The immobilization procedure was adapted from Wang methodology.33 Thus, 2 g of

2. EXPERIMENTAL SECTION 2.1. Chemicals and Enzymes. Lyophilized N. gaditada microalga was purchased from Easyalgae (Spain). CalB, phosphate buffer pH 7.0, Pluronic P123, Pluronic F127, tetraethyl orthosilicate (TEOS), dimethylamine, cetyltrimethylammonium bromide (CTAB), 1,3,5triisopropylbenzene, 1,3,5- trimethylbenzene (TMB), and glutaraldehyde were all supplied by Sigma-Aldrich. Methanol, ethanol, n-hexane, and anhydrous toluene were provided by Scharlab. N-Methylamino4982

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels

Figure 1. TEM images: (a) SBA-15, (b) SBA-16, (c) MCM-41, and (d) FDU-12. calcined mesoporous material was immersed into a solution of 1 g of APTES in 30 mL of anhydrous toluene. The flask was refluxed at 110 °C for 24 h under an inert nitrogen atmosphere. Then, the mixture was filtered and washed 3 times with anhydrous toluene. The solid was placed in glass vials and vacuum-dried for 24 h at 110 °C. Then, 1 g of amine-functionalized support was blended with 1 mL of 25 vol % aqueous glutaraldehyde and 9 mL of 0.1 M phosphate buffer (pH 7) for 2 h. The solid material was filtered and washed with phosphate buffer. The functionalized materials were used as carriers for lipase immobilization. A total of 100 mg of functionalized mesoporous material was blended with 1 mL of lipase in 4 mL of 0.1 M phosphate buffer (pH 7), and the mixture was shaken at 200 rpm and 25 °C for 3 h. Then, the material was filtered and washed 3 times with 5 mL of pH 7 phosphate buffer. The lipase concentration in the filtrate was measured by the Bradfordś assay, and the amount of enzyme loaded was estimated by mass balance. 2.5. Mesoporous Material Characterization. Adsorption/ desorption isotherms of nitrogen at 77 K were obtained using a Tristar 3000 (Micromeritics Instrument Co., Norcross, GA). Different methods were used for textural characterization: the Brunauer− Emmett−Teller (BET) method for surface area, the Barrett−Joyner− Halenda (BJH) method for pore diameter, and the Harkins−Jura method for pore volume. A Tecnai F20 transmission electron microscope (Philips. Amsterdam, Netherlands) with a resolution of 0.27 nm equipped with an energy-dispersive X-ray (EDX) detector was used to obtain the TEM images. 2.6. Enzyme Loading. The amount of enzyme immobilized was calculated by mass balance measuring the enzyme concentration in the supernatant solution using the Bradfords assay, with bovine serum albumin (BSA) as the standard. 2.7. FAME and FAEE Production. Reaction experiments were carried out in 50 mL closed glass flasks with magnetic stirring in a multi-reactor system (Radleys. Essex, U.K.). In all cases, 0.1 g of oil was used. A sample of 100 μL was withdrawn at different times and

centrifuged at 6500g for 5 min to remove the solid biocatalyst. The supernatant was placed in a vial, and ethanol or methanol was evaporated in an oven at 110 °C. The resulting oil was dissolved in nhexane, and a 1 μL aliquot was analyzed by TLC.28 The reaction was first performed with free lipase to optimize the reaction conditions. The variables studied were the type of alcohol (methanol or ethanol), the lipase concentration, the alcohol/oil ratio at different times (4, 8, 24, and 48 h), and the temperature of the reaction. Standard error for fatty acid ester yield was ±0.9 wt %, as determined after replication of experiments (data not shown).

3. RESULTS AND DISCUSSION 3.1. Mesoporous Material Characterization and Lipase Immobilization. Siliceous materials used as enzyme carriers were obtained using different methods and surfactants. Thus, MCM-41 and SBA-15 were produced using cationic CTAB and triblock copolymer P123, respectively. These structures display hexagonally closed packed cylindrical pore channels showing p6mm symmetry. Transmission electron microscopy (TEM) images (panels a and c of Figure 1) show the typical hexagonal structures along the channel system and parallel stripes when viewed perpendicular to channel directions. SBA-16 and FDU-12 were synthesized using acidic triblock copolymer F127, obtaining large cages that are connected by smaller windows and arranged into different cubic structures (panels b and d of Figure 1). FDU-12 presented a cage-like face-centered cubic mesostructure following the Fm3m symmetry with a large cavity size, while SBA-16 had a body-centered cubic mesostructure following the Im3m symmetry. The X-ray diffraction (XRD) pattern and N2 adsorption isotherms of the mesoporous materials (not shown) indicated highly ordered two-dimensional (2D) mesostructured 4983

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels Table 1. Textural Properties of Mesoporous Materials and Lipase (CalB) Loading textural properties

a

support

SBET (m2 g−1)a

Dp (Å)b

Vp (cm3 g−1)c

−NH2 loading (mmol N gsupport−1)d

lipase loading (mglipase/gsupport)e

SBA-15 SBA-16 MCM-41 FDU-12

573.14 801.41 1027.24 553.64

89.4 45.1 31.4 111.4

0.97 0.55 0.76 0.74

1.57 1.65 1.60 1.50

36.1 26.6 28.8 22.8

BET surface area. bBJH pore diameter. cPore volume (calculated at P/P0 = 0.904). dElemental analysis. eBradford assay.

diameter and pore volume. Among cubic cage-like materials, both showed lower enzyme loading likely caused by the larger tortuosity of the pore network, in comparison to the unidirectional channels of SBA-15, which could block or hinder the enzyme diffusion along the pores. One-dimensional (1D) pore structures have shown to facilitate internal diffusion of enzymes to reach the immobilization sites. Conversely, cagelike pore structures provided the highest diffusional restriction, leading to lower enzyme loading.34 When the amount of all three lipases loaded in SBA-15 was analyzed and compared, the results showed that 36.1, 40.2, and 15.7 mg of CalB, T. lanuginosus, and M. miehei lipases were covalently immobilized per gram of support. 3.2. Lipid Extraction and Characterization. Prior to FAME and FAEE production, lipids from N. gaditana microalga were extracted with methanol. The lipid content was 35.5 ± 1.2 wt %, as shown in Table 2, and the composition, analyzed by

materials with hexagonal symmetry and mesoporous materials with three-dimensional (3D) cage-type pores. N2 adsorption/desorption isotherms of all materials were of type IV. MCM-41 and SBA-15 showed a H1 hysteresis cycle, which is associated with mesoporous materials with large pores. SBA-16 and FDU-12 presented a H2 hysteresis cycle that is usually present in cage-like porous materials. Table 1 summarizes the textural properties of the four synthesized carriers, where surface area and pore diameter are crucial for the enzyme immobilization process. Among hexagonal materials, SBA-15 had a higher pore size than MCM-41 (89.4 and 31.4 Å, respectively). Focusing on cubic materials, FDU-12 had a pore size 3 times larger than SBA-16 as well as a lower surface area. Both pore size and surface area have a key effect on both the immobilization process and enzyme activity. Prior to enzyme immobilization, mesoporous materials were modified via grafting of APTES and further addition of glutaraldehyde as a cross-linking agent. Nitrogen content in APTES-grafted materials were measured by elemental analysis. The results showed that the aminosilane content was very similar for all supports (Table 1). The proper incorporation of amino groups after the grafting process was confirmed by solid-state 29Si magic-angle spinning (MAS) nuclear magnetic resonance (NMR). The spectra (see Figures S1−S4 of the Supporting Information) display two regions of major intensity, centered from about −100 to −110 ppm and about −70 ppm, respectively. These two regions correspond to Si(−O−)4 and R−Si(−O−)3 species, respectively, where R is the amino group. The pattern from −100 to −110 ppm is composed of at least three contributions, at about −92, −100, and −110 ppm, corresponding to (≡SiO)2Si(OR′)2 (Q2), (≡SiO)3-Si(OR′) (Q3), and (≡SiO)4Si (Q4) species, respectively, where R′ is (−CH2−CH2−) or H. The lower shielding pattern is formed by two main contributions, at −60 and −69 ppm, associated with species containing the amino ligand groups, R−Si(OSi≡)n (Tn), T2, and T3, respectively. As the 29Si NMR spectrum was obtained by a direct one-pulse experiment, the quantification of the species associated with the respective intensities was performed by Gaussian deconvolution of the spectra. Thus, the incorporation of the amino organic moiety in the silica framework was calculated, corroborating those obtained through elemental analysis. In terms of amino group incorporation, there were no significant differences between hexagonal and cubic mesostructured materials used as enzyme supports. Once the structure and amino group incorporation were confirmed, all materials were used for CalB immobilization. Table 1 summarizes the CalB loading for each support. According to the results, it can be seen that the hexagonal materials, SBA-15 and MCM-41, led to higher enzyme incorporation than cubic cage-like pore materials, SBA-16 and FDU-12. SBA-15 showed a significantly higher enzyme loading compared to the rest of the supports because of its higher pore

Table 2. Lipids Extracted from N. gaditana Using Methanol as a Solvent lipids (wt %)

biomass free of lipids (wt %)

extraction losses (wt %)

35.5 ± 1.2

58.0 ± 0.5

7.0 ± 0.6

Table 3. Composition of the Lipids Extracted from N. gaditana classification FFA and saponifiable lipids

non-saponifiable lipids

other non-lipidic compounds a

concentration (wt %)

type of lipid FFAs triglycerides diglycerides monoglycerides sterol esters polar lipids total saponifiable lipids carotenoids sterols and tocopherols retinoids total non-saponifiable lipids

40.1 ± 2.4 nda nda nda 4.6 ± 0.2 42.0 ± 1.4 86.7 ± 4.0 1.10 ± 0.04 nda nda 1.10 ± 0.04 12.2 ± 4.04

nd = not detected.

TLC, is summarized in Table 3. The extracted lipids are mainly composed of saponifiable matter (86.7 ± 4.0 wt %), most of which are polar lipids (phospholipids, sphingolipids, and saccharolipids) and FFAs that can be transformed into methyl or ethyl esters. The high content of FFAs and absence of triglycerides, diglycerides, and monoglycerides are due to the hydrolysis of the last three during microalgae biomass lyophilization. Table 4 shows the fatty acid profile for the 4984

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels Table 4. Fatty Acid Composition of Saponifiable Lipids and Fatty Acids Extracted from N. gaditana fatty acid lauric acid myristic acid myristoleic acid pentadecanoic acid palmitic acid palmitoleic acid stearic acid oleic acid linoleic acid linolenic acid arachidic acid gadoleic acid arachidonic acid eicosapentaenoic acid behenic acid erucic acid lignoceric acid nervonic acid other a

concentration (wt %) (C12:0) (C14:0) (C14:1) (C15:0) (C16:0) (C16:1) (C18:0) (C18:1) (C18:2) (C18:3) (C20:0) (C20:1) (C20:4) (C20:5) (C22:0) (C22:1) (C24:0) (C24:1)

nda 4.40 2.14 nda 24.59 32.42 0.78 3.95 2.35 2.12 0.00 nda 2.43 16.81 0.00 7.96 nda nda 0.08

Figure 2. Effect of the type of alcohol (■, ethanol; ▲, methanol) on the ester content in the reaction with free C. antarctica lipase B (40 °C temperature, 5:1 alcohol/oil mass ratio, and 500:1 oil/lipase mass ratio).

obtained with this lipase was very sensitive to the enzyme concentration, with the activity decreasing dramatically when the oil/lipase mass ratio increased above 50:1. However, the decrease in the FAEE yield was similar for 50:1 and 500:1 oil/ lipase mass ratios when CalB (Figure 3B) and M. miehei lipase (Figure 3C) were used. These are remarkable results from the practical application point of view because they allow for the reduction of the lipase concentration by 10 times. Therefore, the above results indicate that the reaction rate and conversion increased with enzyme loading up to a certain value where a higher lipase content had no further effect on the enzymatic activity, which is now controlled by diffusional processes inside the pores. To check the influence of the excess ethanol, FAEE production was carried out at different ethanol/oil mass ratios (8:1, 40:1, and 80:1) using free T. lanuginosus, CalB, and M. miehei lipases (panels A, B, and C of Figure 4, respectively). This figure shows that, at short times, the lower the ethanol excess, the higher the ester yield for the three enzymes because of a better contact between the oil and lipase, so that the use of low ethanol excess will help to reduce operational costs associated with alcohol recycling. Longer reaction times, using the three lipases, lead to closer yield values for all ethanol/oil mass ratios used because the final yield is controlled by the reaction equilibrium. The effect of the temperature on enzymatic FAEE production was evaluated with free lipases using a 8:1 ethanol/oil mass ratio and 500:1 oil/lipase mass ratio. The results (Figure 5) showed the presence of an optimum region within the range of 40−50 °C for CalB. At a higher temperature, the activity decreased because of thermal deactivation caused by denaturation of the enzyme. It can be observed that the maximum ester production was around 75 wt % at 50 °C. A similar pattern was observed for T. lanuginosus lipase, although the activity was lower than that for CalB within the temperature range studied. However, the activity of M. miehei lipase at 30−30 °C was very low, showing a marked maximum at 40 °C and a steep decrease to zero at 60 °C. This behavior agrees with previous studies about transesterification of refined vegetable oils.37 The immobilization is expected to provide a protected microenvironment within the mesoporous walls, increasing the thermal stability of the enzyme.38

nd = not detected.

saponifiable lipids extracted from N. gaditana. Microalgae oils are usually rich in polyunsaturated fatty acids. Therefore, FAMEs derived from these fatty acids would provide evidence for good fuel properties at low temperatures, which is an important advantage in winter operation. However, they would show a low oxidative stability.5,10 In the analyzed N. gaditana oil, the contents of saturated, monounsaturated, and polyunsaturated fatty acids were 29.8, 46.5, and 23.8 wt %, respectively. These values supported a iodine value of 161 g of I2/100 g for the saponifiable lipids, which is higher than the limit of 120 g of I2/100 g specified in the EN 14214 standards. This parameter is a measure of the unsaturation level providing high values because of the high concentration of eicosapentaenoic acid (20:5) in the microbial oil. 3.3. FAME and FAEE Production with Free Lipase. The reaction conditions were optimized using free CalB. Conversion of the extracted crude microalgal oil to FAMEs or FAEEs was performed with different alcohols, lipase concentrations, molar ratios of ethanol/lipase, reaction times, and temperatures. The influence of the type of alcohol using free CalB is shown in Figure 2. It can be seen that the ester content in the reaction with ethanol was significantly higher than that with methanol, along with the reaction time. This effect is in agreement with results reported in previous studies,35 because the enzyme activity increased with alcohol chain length. Coupled with this, a higher enzyme deactivation was described in the reaction with methanol.36 Although methanol acts as a lipase inhibitor, it must be used in excess to shift the reversible esterification and transesterification reactions for the production of FAME. For these reasons, the methanol concentration must be optimized to balance both opposite effects.26 The influence of the enzyme concentration on the FAEE production was the second variable studied. Three different oil/ enzyme mass ratios (50:1, 500:1, and 5000:1) were tested in the reactions using three types of free lipases (T. lanuginosus, CalB, and M. miehei). Figure 3A shows that the FAEE content increased faster for T. lanuginosus lipase. Thus, the yield 4985

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels

Figure 3. Effect of the oil/lipase mass ratio (■, 50:1; ▲, 500:1; ◆, 5000:1) on the FAEE content in the reaction with free lipase from (A) T. lanuginosus, (B) C. antarctica B, and (C) M. miehei. Operating conditions: 40 °C temperature and 15:1 alcohol/oil mass ratio.

Figure 4. Effect of the ethanol/oil mass ratio (▲, 8:1; ◆, 40:1; ●, 80:1) on the FAEE content in the reaction with free lipase from (A) T. lanuginosus, (B) C. antarctica B, and (C) M. miehei. Operating conditions: 40 °C temperature and 500:1 oil/lipase mass ratio.

Figure 6 shows the ester content in the reactions with CalB, T. lanuginosus, and M. miehei lipases immobilized in SBA-15. The FAEE content increased with time for the CalB- and T. lanuginosus-supported lipases, but this increase was more significant for the CalB lipase, which achieved a FAEE content of 74.1 wt % at 24 h. The FAEE content also rose with time in the reactions with M. miehei lipase immobilized in SBA-15 up to 8 h. At longer reaction times, the activity of this catalyst was practically kept constant. In this sense, CalB immobilized in SBA-15 produce the best catalytic activity in the FAEE production using N. gaditana oil. To evaluate the effect of the support on the enzymatic activity of lipases, new experiments with free CalB, T. lanuginosus, and M. miehei lipases were

The optimal reaction conditions obtained are summarized in Table 5 and will be used for the FAEE production reactions with supported lipases. 3.4. FAEE Production Reaction with Supported Lipase. The oil extracted from N. gaditana with methanol under reflux and magnetic stirring was used in the FAEE production with CalB, T. lanuginosus, and M. miehei lipases immobilized in SBA-15, SBA-16, MCM-41, and FDU-12. The optimal operating conditions for the free lipase (Table 5) were selected for the supported lipase experiments. In addition, for a better comparison of the activity of all biocatalysts, the amount of lipase in all reactions was set constant, regardless the support used. 4986

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels

Figure 7 shows the FAEE content in the reactions with CalB immobilized in different mesoporous materials with both

Figure 5. Effect of the temperature on the FAEE content in the reaction with free lipases (▲, C. antarctica B; ◆, T. lanuginosus; ●, M. miehei). Operating conditions: 15:1 alcohol/oil mass ratio, 500:1 oil/ lipase mass ratio, and ethanol as the type of alcohol.

Figure 7. Effect of the carrier on the FAEE content using supported C. antarctica lipase B (reaction conditions are shown in Table 5).

Table 5. Optimal Reaction Conditions with Free C. antarctica Lipase B variables

optimal conditions

time (h) alcohol ethanol/oil (m/m) oil/lipase (m/m) temperature (°C)

24 ethanol 8:1 500:1 40

hexagonal and cubic structures. High FAEE contents were obtained within the reaction media with all biocatalysts, although enzyme activities were largely affected by the morphology of the siliceous support. A noticeable result that must be highlighted was the higher ester yield obtained when using immobilized CalB in hexagonal structure materials, reaching ester contents of 74.1 and 71.8 wt % for SBA-15 and MCM-41, respectively. For both cubic supports, FAEE production was lower than that obtained with hexagonal siliceous supports, 62.3 and 65.8 wt % for FDU-12 and SBA-16, respectively. Therefore, CalB immobilized in 1D pores of hexagonal materials seems to be more accessible than lipase immobilized in the 3D cubic supports because of the textural properties of both types of carriers.39 Regular morphology of SBA-15 and MCM-41 in hexagonally close packed cylindrical pore channels allows for easier diffusion than SBA-16 and FDU-12. The 3D geometry generates an inherent tortuosity, hindering the microalgal oil diffusion along the channels. Keeping in mind the similar enzyme loading for every support, with both hexagonal and cubic structures, the tridimensional cubic network led to non-accessible enzymes inside the cages, which induced lower FAEE production (Figure 7). Among hexagonal materials, SBA-15 provided higher FAEE production because of its larger pore size favoring the diffusion of the lipids through the inner channels. The larger MCM-41 surface area (1027 m2/g) may partially offset the low pore diameter effect because this is the material with the smallest pore size (31.4 Å), which could hamper the enzyme entering the pores. Because lipase loading in MCM-41 was similar to that in SBA-15, a high amount of enzyme could be in the external surface of the MCM-41 support. The enzyme anchored on the external surface is accessible to the microalga oil favoring the FAEE production and counteracting the diffusional problems because the MCM-41 pore size is smaller than the lipase molecule whose approximate dimensions are 52 × 47 × 52 Å40 (evaluated from the molecular structure available in the Protein Data Bank; PDB ID: 1TCA; http://www.rcsb. org). Higher yields were obtained with the supported lipase than with the free lipase because of the protection and stability given by the siliceous support to the enzyme, avoiding denaturation and loss of activity.

Figure 6. Effect of the lipase type (▲, C. antarctica B; ◆, T. lanuginosus; ●, M. miehei) on the FAEE content in the reaction with supported lipase in SBA-15 compared to the equilibrium FAEE content with free lipases (dotted lines) at optimum reaction conditions (shown in Table 5).

carried out at the optimum conditions (Table 5). The results (Figure 6) showed that, for CalB, only an additional 4.1% of ester (78.2 versus 74.1%) was obtained with free lipase compared to that immobilized in SBA-15. This proved that the immobilization process for this enzyme slightly affected its activity. For M. miehei lipase, a similar decay in the ester content was measured (63.3% for free versus 59.2% for immobilized lipase). However, T. lanuginosus lipase showed a dramatic decrease in enzymatic activity after immobilization, achieving 68.4% of ester after 24 h for free lipase but only 28.5% when this enzyme was supported in SBA-15. 4987

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels

microalgae. These noticeable results demonstrate again the robustness of the CalB-based biocatalyst prepared and its promising application to microalgae biodiesel production.

To evaluate the stability of covalently immobilized lipases for FAEE production from N. gaditana oil, several recycling tests were carried out with CalB, M. miehei, and T. lanuginosus lipases on SBA-15. Figure 8 displays the yield results achieved after

4. CONCLUSION The present work demonstrates for the first time the feasibility of the enzymatic production of FAEEs from the microalga N. gaditana using T. lanuginosus, CalB, and M. miehei lipases, both free and immobilized in mesostructured silica materials. A FAEE content of 71 and 74.1 wt % were achieved with free and immobilized CalB, respectively, in SBA-15 at the following operating conditions: 40 °C and 8:1 ethanol/oil and 500:1 oil/ lipase mass ratios. The immobilization of CalB in SBA-15 produces an enhancement of the lipase activity without detecting any leaching of the lipase.

■

ASSOCIATED CONTENT

S Supporting Information *

29

Si MAS NMR solid-state spectra of MCM-41 material (Figure S1), 29Si MAS NMR solid-state spectra of SBA-16 material (Figure S2), 29Si MAS NMR solid-state spectra of SBA-15 material (Figure S3), and 29Si MAS NMR solid-state spectra of FDU-12 material (Figure S4) (PDF). The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/ef502838h.

Figure 8. Reuse of biocatalysts: (black bars) C. antarctica lipase B, (white bars) M. miehei lipase, and (gray bars) T. lanuginosus lipase supported on SBA-15.

two consecutive runs with intermediate washing with ethanol. A marked decrease (>50 wt %) in the FAEE yield can be observed in the second reaction cycle for M. miehei and T. lanuginosus lipases. However, when immobilized CalB was used, the FAEE yield was not reduced in the second use. This is due to the stronger resistance to structural denaturation of CalB against nucleophiles, such as low-molecular-weight alcohols, compared to M. miehei and T. lanuginosus lipases.41 A recycling test using CalB lipase supported on SBA-15 corroborated these results (Figure 9). Thus, the FAEE yield

■

AUTHOR INFORMATION

Corresponding Author

*Telephone: 34-91-4888501. Fax: 34-91-4887068. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank financial support from Universidad Rey Juan Carlos and Oficina de Cooperación Universitaria (Project PRIN13-CC05).

■

REFERENCES

(1) Mofijur, M.; Masjuki, H. H.; Kalam, M. A.; Atabani, A. E.; Shahabuddin, M.; Palash, S. M.; Hazrat, M. A. Renewable Sustainable Energy Rev. 2013, 28, 441−455. (2) Balat, M. Energy Convers. Manage. 2011, 52, 1479−1492. (3) Atabani, A. E.; Silitonga, A. S.; Ong, H. C.; Mahlia, T. M. I.; Masjuki, H. H.; Badruddin, I. A.; Fayaz, H. Renewable Sustainable Energy Rev. 2013, 18, 211−245. (4) Singh, B.; Guldhe, A.; Rawat, I.; Bux, F. Renewable Sustainable Energy Rev. 2014, 29, 216−245. (5) Meng, X.; Yang, J.; Xu, X.; Zhang, L.; Nie, Q.; Xian, M. Renewable Energy 2009, 34, 1−5. (6) Rawat, I.; Ranjith Kumar, R.; Mutanda, T.; Bux, F. Appl. Energy 2013, 103, 444−467. (7) Richmond, A.; Hu, Q. Handbook of Microalgal Culture: Applied Phycology and Biotechnology, 2nd ed.; Richmond, A., Hu, Q., Eds.; Wiley-Blackwell: Chichester, U.K., 2011. (8) Chisti, Y. Trends Biotechnol. 2008, 26, 126−131. (9) Oncel, S. S. Renewable Sustainable Energy Rev. 2013, 26, 241−264. (10) Carrero, A.; Vicente, G.; Rodríguez, R.; Linares, M.; del Peso, G. L. Catal. Today 2011, 167, 148−153. (11) Atadashi, I. M.; Aroua, M. K.; Abdul Aziz, A. R.; Sulaiman, N. M. N. Renewable Sustainable Energy Rev. 2012, 16, 3275−3285. (12) Sani, Y. M.; Daud, W. M. A. W.; Abdul Aziz, A. R. J. Environ. Chem. Eng. 2013, 1, 113−121.

Figure 9. Recycling test for the production of FAEE from N. gaditana using C. antarctica lipase B supported on SBA-15.

was slightly reduced between the first and seventh reaction cycle from an initial value from 74.1 to 71.5%. After 10 reaction cycles, the FAEE content decreased to a final value of 61.1%. Although the production of biodiesel from microalgae using lipases immobilized in mesoporous silicas has not been reported thus far, the above behavior can be considered an improvement compared to other biocatalytic systems, both commercial42,43 or based on amorphous silicas,26 for the enzymatic production of a biodiesel from ethanol and 4988

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989

Article

Energy & Fuels (13) Bajaj, A.; Lohan, P.; Jha, P. N.; Mehrotra, R. J. Mol. Catal. B: Enzym. 2010, 62, 9−14. (14) Szczęsna Antczak, M.; Kubiak, A.; Antczak, T.; Bielecki, S. Renewable Energy 2009, 34, 1185−1194. (15) Souza, M. S.; Aguieiras, E. C. G.; da Silva, M. A. P.; Langone, M. A. P. Appl. Biochem. Biotechnol. 2009, 154, 74−88. (16) Gog, A.; Roman, M.; Toşa, M.; Paizs, C.; Irimie, F. D. Renewable Energy 2012, 39, 10−16. (17) Tosa, T.; Mori, T.; Fuse, N.; Chibata, I. Enzymologia 1966, 31, 214−224. (18) Santalla, E.; Serra, E.; Mayoral, A.; Losada, J.; Blanco, R. M.; Díaz, I. Solid State Sci. 2011, 13, 691−697. (19) Moelans, D.; Cool, P.; Baeyens, J.; Vansant, E. F. Catal. Commun. 2005, 6, 307−311. (20) Hartmann, M. Chem. Mater. 2005, 17, 4577−4593. (21) Lee, C. H.; Lin, T. S.; Mou, C. Y. Nano Today 2009, 4, 165− 179. (22) Ciesla, U.; Schüth, F. Microporous Mesoporous Mater. 1999, 27, 131−149. (23) Canilho, N.; Jacoby, J.; Pasc, A.; Carteret, C.; Dupire, F.; Stébé, M. J.; Blin, J. L. Colloids Surf., B 2013, 112, 139−145. (24) Li, X.; Xu, H.; Wu, Q. Biotechnol. Bioeng. 2007, 98, 764−771. (25) Lai, J.-Q.; Hu, Z.-L.; Wang, P.-W.; Yang, Z. Fuel 2012, 95, 329− 333. (26) Tran, D.-T.; Yeh, K.-L.; Chen, C.-L.; Chang, J.-S. Bioresour. Technol. 2012, 108, 119−127. (27) Zhao, X.; Peng, F.; Du, W.; Liu, C.; Liu, D. Bioprocess Biosyst. Eng. 2012, 35, 993−1004. (28) Vicente, G.; Bautista, L. F.; Rodríguez, R.; Gutiérrez, F. J.; Sádaba, I.; Ruiz-Vázquez, R. M.; Torres-Martínez, S.; Garre, V. Biochem. Eng. J. 2009, 48, 22−27. (29) Zhao, D.; Feng, J.; Huo, Q.; Melosh, N.; Fredrickson, G. H.; Chmelka, B. F.; Stucky, G. D. Science 1998, 279, 548−552. (30) Lin, W.; Cai, Q.; Pang, W.; Yue, Y.; Zou, B. Microporous Mesoporous Mater. 1999, 33, 187−196. (31) Kim, T.-W.; Ryoo, R.; Kruk, M.; Gierszal, K. P.; Jaroniec, M.; Kamiya, S.; Terasaki, O. J. Phys. Chem. B 2004, 108, 11480−11489. (32) Kruk, M.; Hui, C. M. Microporous Mesoporous Mater. 2008, 114, 64−73. (33) Wang, C.; Zhou, G.; Li, Y.-J.; Lu, N.; Song, H.; Zhang, L. Colloids Surf., A 2012, 406, 75−83. (34) Serra, E.; Mayoral, Á .; Sakamoto, Y.; Blanco, R. M.; Díaz, I. Microporous Mesoporous Mater. 2008, 114, 201−213. (35) Kulschewski, T.; Sasso, F.; Secundo, F.; Lotti, M.; Pleiss, J. J. Biotechnol. 2013, 168, 462−469. (36) Adlercreutz, P. Chem. Soc. Rev. 2013, 42, 6406−6436. (37) Gulyamova, K. A.; Davranov, K. D. Chem. Nat. Compd. 1996, 31, 372−375. (38) Abdullah, A. Z.; Sulaiman, N. S.; Kamaruddin, A. H. Biochem. Eng. J. 2009, 44, 263−270. (39) Barrón Cruz, A. E.; Melo Banda, J. A.; Mendoza, H.; RamosGalvan, C. E.; Meraz Melo, M. A.; Esquivel, D. Catal. Today 2011, 166, 111−115. (40) Uppenberg, J.; Hansen, M. T.; Patkar, S.; Jones, T. A. Structure 1994, 2, 293−308. (41) Hernández-Martín, E.; Otero, C. Bioresour. Technol. 2008, 99, 277−286. (42) Guldhe, A.; Singh, B.; Rawat, I.; Permaul, K.; Bux, F. Fuel 2015, 147, 117−124. (43) Alavijeh, R. S.; Tabandeh, F.; Tavakoli, O.; Karkhane, A.; Shariati, P. J. Oleo Sci. 2015, 64, 69−74.

4989

DOI: 10.1021/ef502838h Energy Fuels 2015, 29, 4981−4989