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Renewable Energy 68 (2014) 1e5

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Renewable Energy

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Biodiesel production via lipase catalysed transesterification ofmicroalgae lipids from Tetraselmis sp.

Chee Loong Teo a, Haryati Jamaluddin b, Nur Azimah Mohd Zain b, Ani Idris a,*

aDepartment of Bioprocess Engineering, Faculty of Chemical Engineering, c/o Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia,81310, UTM, Johor Bahru, Johor, Malaysiab Faculty of Biosciences and Bioengineering, Universiti Teknologi Malaysia, 81310, UTM, Johor Bahru, Johor, Malaysia

a r t i c l e i n f o

Article history:Received 6 July 2013Accepted 22 January 2014Available online

Keywords:MicroalgaeTetraselmis sp.BiodieselImmobilized lipaseTransesterificaton

* Corresponding author. Tel.: þ60 75535603; fax: þE-mail address: ani@cheme.utm.my (A. Idris).

0960-1481/$ e see front matter � 2014 Elsevier Ltd.http://dx.doi.org/10.1016/j.renene.2014.01.027

a b s t r a c t

Tetraselmis sp. is a green marine microalgae and known to produce lipids that can be transformed intobiodiesel. The influence of nitrate concentration (0.00 g/L, 0.10 g/L, 0.14 g/L and 0.18 g/L) on the growthrate of Tetraselmis sp. was investigated. The marine microalgae were harvested during the exponentialphase and lipid was extracted by chloroform-methanol solvent and quantified using Nile Red method.The conversion of lipid to biodiesel was performed via i) alkali-based transesterification reaction whichutilized sodium hydroxide (NaOH) and ii) enzyme catalysed transesterification process which utilizedimmobilized lipase. The fatty acid methyl esters (FAME) components were identified using gas chro-matography (GC) and then compared with the FAME standard. The results revealed that 0.18 g/L nitrateconcentration was the optimal for cultivation of microalgae. However, the highest lipid content wasachieved in the absence of nitrate (0.0 g/L). The biodiesel yield from the lipase catalysed trans-esterification process was 7 folds higher compared to the alkaline based transesterification.

� 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Recently, biodiesel from oil crops such as soy and palm, animalfat, waste cooking oil [1,2], bioethanol and other alcohols fromsugarcane and corn starch, hydrogen long-chain hydrocarbons, andbiogas [3] have attracted much attention as an important fuel op-tion and coined as biofuel. However biofuel from food crops willrequire large areas of arable land and has to compete with thecultivation of food crops and thus is currently a huge controversialissue [4]. The biofuel utilization in the European Union is growingrapidly and great efforts were made in developing the technology[5]. For the last few decades, microalgae have been recognized as apotential source of sustainable substrate to produce biofuel due tothe numerous advantages such as rapid growth potential andbiomass generation, higher photosynthesis efficiency and norequirement for large arable land area [6e8]. Microalgae can becultivated near to sewage or next to power plant smokestacks so asto digest the pollutants and generate lipid. In the late 1990’s, somemicroalgae were discovered to produce neutral lipids from carbondioxide during photosynthesis process with efficiency of 30 timesmore than that of plants in terms of the amount of neutral oil

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produced per unit area of the land allocated [9]. In the cultivation ofmicroalgae the amount of nitrogen is an important factor to beaddressed because it is an important component of amino acids, aswell as being vital for chlorophyll hence is likely to promotemicroalgae growth. Previous studies have shown that under ni-trogen (nitrate) limitation conditions, nitrogen metabolism in thealgal cells decreases, contributing to the enhancement of the syn-thetic activities of secondary carotenoids and other non-nitrogencompounds. The nitrogen-deficient condition increases lipid con-tent due to the lack of NaNO3 which limits protein biosynthesis,reduce biomass of microalgae but promotes lipid/protein ratio [10].

Biodiesel is a mixture of fatty acid methyl esters (FAMEs) whichis produced from the transesterification of lipids and is rapidlygaining acceptance by the majority as an alternative source of en-ergy for the future [11]. In production of biodiesel from marinemicroalgae, the low yield from the alkali-catalysed trans-esterification is one of the major challenges faced.

It is also believed that the lipase- catalysed transesterificationcan improve the yield of biodiesel. Lipases can be categorisedinto three classes based on their specificity or selectivity such asregio- or positional specific lipases; fatty acid type specific lipasesand specific lipases for a certain class of acylglycerols (mono-, di- ortriglycerides). Several examples are lipases from Pseudomonas flu-orescens, Pseudomonas cepacia, Candida rugosa, Candida antarctica,Candida cylindracea, Rhizopus oryzae and Mhizomucor miehei.

C.L. Teo et al. / Renewable Energy 68 (2014) 1e52

Enzymes can be linked to insoluble matrices by a variety ofmethods such as adsorption, covalent coupling and entrapment; aprocess called immobilization which can provide stability to en-zymes. Stability has always been an issue during transesterification,especially when the media is predominantly non-aqueous. Inaqueous media, where enzymes are freely soluble, immobilizationprovides a method to recover the biocatalyst by centrifugation orfiltration. In anhydrous media, enzyme molecules though mostlyinsoluble, tend to clump together. Thus, immobilization is a suitablemethod to increase the surface area of the biocatalyst.

For the last few decades, there were many reports on biodieselproduction using lipase as biocatalyst [12e17] but most of thesestudies investigated the use of lipase catalysed transesterificationfor biodiesel production from vegetable oils such as sunflower,jatropha, soybean and palm oil. In some of these applications thelipases were immobilized in various supports such as acrylic resin,textile membrane, polypropylene, celite, diatomaceous earth, pol-yglycidylmethacrylate beads [15,16]. Enzymatic catalysed trans-esterification requires mild environment and can tolerate thepresence of water and free acids in crude oil. In addition the re-covery of glycerol and fuel purification is minimized and lowamounts of waste is produced which overcomes the negative effectof alkaline catalysed transesterification [14]. Nearly all of theresearch focused on the use of lipase catalysed transesterificationfor vegetable oils for biodiesel production and to the best of ourknowledge no work has been reported on the use of lipase cata-lysed transesterification for microalgal oils to biodiesel.

Thus in this study biodiesel from microalgae (Tetraselmis sp.)lipids was synthesized using lipase- catalysed transesterification. Inorder to reduce cost and increase life span of enzymes the lipaseswere immobilised in alginate beads thus allowing recycling of theenzymes. Alginate is a common immobilization matrix and hasbeen used successfully for the immobilization of lipases [18e21]and in biodiesel production [22]. In addition, biodiesel was alsosynthesized by conventional alkali transesterification method un-der the same temperature and pH so as to compare their produc-tion yields.

2. Materials and methods

2.1. Microalgae cultures and medium

The marine microalgae strains were bought originally fromAlgae Tech (Ltd). Tetraselmis sp. was maintained in F/2 mediumwhich contains 0.075 g NaNO3; 0.005 g NaH2PO4.H2O; 0.023 gZnSO4.7H2O; 0.152 g MnSO4.H20; 0.0073 g Na2M0O4.2H2O; 0.014 gCOSO4.7H2O; 0.0068 g CuCl2.2H2O; 4.6 g Fe(NH4)2(SO4)2.6H2O and4.4 g Na2EDTA.2H2O per litre; 0.0005 g Biotin; 0.022 g Thiamineand 0.0027 g Vitamin B12 per 20 ml. Tetraselmis sp. was stored in a2 L conical flask as a batch culture at 25 �C� 0.5 �C, pH 7.8� 0.2 andwas exposed to continuous illumination.

2.2. Cultivation of marine microalgae under different nitrateconcentration

The marine microalgae, Tetraselmis sp. was cultivated in F/2medium with 10% starting inoculum taken from stock culture. Themarine microalgae, Tetraselmis sp. was cultivated at ambient tem-perature (27 �C) and pH 7 with aeration condition as the controlgrowth environment. The marine microalgae were also cultivatedunder different nitrate concentrations (0.00 g/L, 0.10 g/L, 0.14 g/Land 0.18 g/L). The marine microalgae growth was monitored interms of turbidity using the optical density (OD) method. Sampleswere taken every two days for twenty-two days where OD readingswere taken using a UVevis spectrophotometer (Shimadzu UVmini-

1240) at a wavelength of 620 nm. The lipid content was measuredusing the Nile Red method [23]. The biomass measured by thespectrophotometer only indicated an absorbance value, thus inorder to obtain a more accurate Tetraselmis sp. growth data, cell dryweight measurement was performed simultaneously. 2 ml of thesample was collected from each different nitrate concentrationcultures and was placed onto aluminium foil. The sample was thenplaced in the incubator at 60 �C and then left overnight. After oneday, the sample weight was measured. Each experiment was per-formed in triplicates for the different nitrate concentrations so as toensure reproducibility of data.

2.3. Measurement of lipid e Nile red staining method

Lipid content was determined rapidly using the Nile red stainingmethod by Perkin Elmer LS-55 fluorescence spectrophotometer.Nile red (NR, 9-diethylamino-5H-benzo[a] phenoxa-phenoxazine-5-one) was diluted to 0.5 mg/mL in acetone [23]. An aliquot 1 mLof the culture broth was taken from the conical flasks and centri-fuged at 2000 rpm for 5 min. The supernatant was discarded andthe pellet was washed with 1 mL of phosphate buffer saline (PBS)followed by centrifugation at 2000 rpm for 5 min. This process wasrepeated for 3 times. The sample was diluted for 1000 fold (103

dilution) and an aliquot of 2.8mLwas withdrawn from the test tubeand addedwith 1.8 mL of Nile Red solution and then transferred intoa cuvette. The solution was kept in the dark for 20 min to achievefull-staining. The excitation and emission wavelength was set at566 and 600 nm respectively; whereas excitation and emission slitwidth were set at 10.0 nm. The fluorescent intensity was thenrecorded.

2.4. Lipid extraction

100 mL of sample was withdrawn from the culture bottle whenthe growth of cultivated microalgae reached the exponential phaseand the lipid extractionwas performed according to Bligh and Dyermethod [24]. The sample was centrifuged repetitively at 4000 rpmfor 15 min. The biomass was washed with 20 mL of distilled waterwas freeze-dried overnight at �40 �C under vacuum. 1 g of thedried sample was grinded by a mortar and pestled into powderform and dissolved in 100 mL distilled water. Cell disruption wasperformed by microwave oven at 100 �C, 2450 MHz for 5 min andsubsequently dissolved in 30mL solvent mixture of chloroform andmethanol (ratio of 2: 1). 10 mL of lysozyme (1 %w/v) was thenadded into the solution so as to disrupt the extracellular cell walland then sonicated for 10 min at (4 �C, pulse 0.7, time 2 min andamplitude 5%) so as to ensure complete lysis of the cell walls. Thesonicated mixture was left for 24 h to allow the formation of bi-layers which consists of a methanol upper layer and chloroformlower layer. The chloroform layer containing the extracted lipid wasthen passed through a filter paper for 2 or 3 times so as to ensurethat any residual algal biomass from the extract was totallyremoved. The lipid was then placed in a rotary evaporator at 50 �Cunder vacuum for 15 min so as to remove any remaining solvent.The crude microalgae oil was stored at 4 �C for the followingtransesterification process.

2.5. Alkali-based transesterification

Methanol was mixed with 0.5 g of NaOH and stirred for 20 minat 400 rpm at approximately 65 �C. The ratio of methanol to oil inthe mixture was kept to 6:1. The mixture of catalyst and methanolwas then poured into the conical flask containing the algae oil toinitiate the transesterification process. The conical flask was stirredcontinuously for 3 h at 300 rpm and then allowed to settle for 16 h

Fig. 1. Cell growth profile of Tetraselmis sp. in terms of dry weight under differentnitrogen concentrations.

Fig. 2. Nile red fluorescence intensity plot over time (day) at different nitrogenconcentrations.

C.L. Teo et al. / Renewable Energy 68 (2014) 1e5 3

so as to obtain 2 separate layers; the supernatant layer (biodiesel)and sediment layers (methanol). The biodiesel was separatedcarefully from the sediment layer by a flask separator and washedusing 5% water until the entire methanol was removed. The bio-diesel was dried using dryer and kept under running fan for 12 h[25]. Biodiesel was stored and the amount of FAME was thendetermined using a Gas Chromatography (Agilent Technologies6890N Network GC system; G2085AA).

2.6. Enzymatic transesterification

2.6.1. Material for enzymatic transesterificationLipase (EC 3.1.1.3) from C. rugosa (Type VII, 700 units/mg) was

purchased from Sigma Aldrich (Germany). The immobilizationmatrix, sodium alginate was purchased from Fluka Chemie GmbH,Buchs, and sodium sulphate from GCE Laboratory Chemicals andcalcium chloride from R&M Marketing, Essex, UK. Iso-octane with99.84% assay was purchased from Fisher Chemicals (UK). All otherreagents were of analytical reagent grade and used without furtherpurification including phosphate buffer solution pH 7.5 or other-wise stated.

2.6.2. Preparation of potassium phosphate bufferThe Candida sp. lipase was dissolved in potassium phosphate

buffer. 0.1 M potassium phosphate buffer was prepared as follows:3.4015 g of potassium dihydrogen phosphate (KH2PO4) and4.3545 g of dipotassium phosphate (K2HPO4) were dissolved indistilled water separately. Then,1.9 mL of KH2PO4, 8.1 mL of K2HPO4and 40 mL distilled water were mixed together to form 0.1 M po-tassium phosphate buffer.

2.6.3. Immobilization of lipase in alginate beads30 g of sodium alginate was dissolved in 1 L distilled water to

make a 3% solution. 0.25 g of lipase was dissolved in 5 mL of po-tassium phosphate buffer and then mixed in 45 mL of sodiumalginate solution. This procedure was performed in aseptic condi-tions. The beads were formed by dropping the biopolymer solutionthrough a syringe and needle into an excess 100 ml of stirred 0.2 MCaCl2 solution at room temperature. The beads with diameter in therange of 1e2 mmwere left in the calcium chloride solution to curefor half an hour.

2.6.4. Lipase-based transesterificationThe reaction was performed in 10 mL of oil, 5 mL of iso-octane

and 1 g immobilized Candida sp. lipase. 60 mL methanol was thenadded to the mixture. The mixture was incubated in an orbitalshaker at 40 �C and 180 rpm for 10 h [17]. The sample was thenfiltered using 0.45 mm nylon syringe filter to remove any unwantedparticles and then placed in a conical flask was then stirredcontinuously for 3 h at 300 rpm. The mixture was then allowed tosettle for 16 h so as to allow the formation of 2 layers. The biodiesellayer was separated from the sediment layer by a flask separatorand washed using 5% water a couple of times so as to remove anyremaining impurities. The obtained biodiesel was dried using dryerand kept under running fan for 12 h [25]. Biodiesel was stored andthe lipid analysis was analysed using Gas Chromatography (GC).

2.7. Gas chromatography (GC) analysis of fatty acid methyl esters(FAME)

Separation and identification of FAME were performed andanalysed in a Gas Chromatography (GC) DB-23 capillary column(30 m � 0.25 mm), using helium as the carrier gas at 20 cm/s. Thecolumn temperature was set at 325 �C as maximum temperature.Both the injector and flame ionization detector (FID) temperature

were set at 300 �C. The front inlet was set at splitless mode andinitial temperature was 250 �C. The column initial temperature wasset at 90 �C during 6 min; the thermal gradient to 210 �C was at arate of 18 �C per min, post temperature at 50 �C in 2 min.

3. Results and discussion

Fig. 1 shows the growth profile of Tetraselmis sp. under differentnitrate concentrations in terms of cell dry weight. It was observedthat the microalgae cultivated in 0.18 g/L nitrate concentrationdisplayed the highest cell dry weight on day 18 of cultivation. Thecell dry weights were low in the absence of nitrate but increasedwhen nitrate concentrationwas increased from 0.1 g/L and 0.18 g/L.

Nile Red (NR) analysis was used to determine the lipid contentin microalgae under different nitrate concentrations. The highervalues of fluorescent intensity (a.u.) indicated higher lipid contentin the microalgae. Fig. 2 shows the growth of Tetraselmis sp. underdifferent nitrate concentrations reached the highest peak at day 20.Thus, the best time to harvest Tetraselmis sp. and extract the lipidwas at day 20 since the highest amount of lipid was attained duringthis time. Fig. 2 also shows the microalgae culture in the absence ofnitrate has the highest fluorescence intensity indicating highestlipid content. Although cultivating microalgae in 0.18 g/L nitrateconcentration exhibited highest dry cell weight, the fluorescenceintensity was lower than those cultivated in the absence of nitrateindicating that highest lipid accumulation occurred in culturesgrown in the absence of nitrate. Absence of nitrate increased the

Fig. 4. Volume of biodiesel under different transesterification methods.

Fig. 3. Volume of total lipids over cell dry weight.

C.L. Teo et al. / Renewable Energy 68 (2014) 1e54

lipid content due to limited protein biosynthesis but cannot sup-port sustainable growth of microalgae. The nitrate deficient con-dition increased lipid/protein ratio and absorbance (620 nm) ofmicroalgae and similar findings were reported by Illman et al. [10]for chlorella sp. Microalgae growth under stress condition whichwas the absence of nitrate source led to the termination of celldivision. Lipid started to accumulate rapidly in the microalgaeduring moment of environmental stress. However, the microalgaeentered the dead phase due to lack of nitrate condition.

In conclusion, although cultivating microalgae in the absence ofnitrate produced highest lipid concentration, the microalgae could

Fig. 5. Chromatogram of FAME found in Tetraselmis sp. (i) Palmitric acid, (ii) Stearic acid, (iii)and (viii) Behenic acid.

not be productive enough to sustain its growth. For long term lipidproduction, cultivating microalgae under total absence of nitratecould not be growth sustainable. In order to estimate productivityof lipid with respect to the amount of microalgae biomass, the totallipid/cell dry weights at various nitrate concentrations wereplotted. Fig. 3 clearly illustrates that the highest ratio is achieved inthe absence of nitrate which reached 508.42 mL/g at harvest day.

Fig. 4 shows higher biodiesel yields were obtained from theenzyme based method compared to the alkali-based trans-esterification. Two layers were formed after 16 h and the volume ofbiodiesel produced was higher than the alkali-based trans-esterificationmethod due to efficiency of enzyme to convert lipid tobiodiesel. Enzyme based transesterification method producedhigher biodiesel yield than alkali based method because lipasecatalysed reactions are very specific; in this case producing higheramounts of FAMEs (biodiesel) and less byproducts. Thus lipasecatalysed transesterification is very attractive because the reactioncan be performed under mild condition at 40 �C compared to alkali(NaOH) catalysed reaction which requires high temperature of65 �C. Compared to the alkali based transesterification, utilizationof immobilised lipases for biodiesel production has some advan-tages such as more compatibility with variations in the quality ofthe raw material and reusability of the enzymes. The ability of li-pases to produce higher amounts of specific products such as bio-diesel is an advantage, because the number of purification steps canbe reduced which translates to less energy usage and thus reducecost. It can significantly decrease the total amount of waste watergenerated and improve the product separation and glycerol qualityas reported by Minodora et al. [14]. Some studies have reportedhigh conversion rates of waste cooking oil (77.87%) [26] and plantoil 80e90% [27] to biodiesel using lipases. Others reportedconsiderable high conversion rates of approximately 78% [28] and73% [29] for biodiesel from waste plant oils and jatropha oilsrespectively using lipase catalysed transesterification.

Biodiesel components were confirmed using the gas chroma-tography. Gas chromatography analysis of the fatty acid methylesters (FAME) was performed in order to determine the fatty acidcomponents of microalgae lipid. The graph in Fig. 5 shows eachpeak representing individual FAME components such as (i) Palmi-tric acid, (ii) Stearic acid, (iii) Oleic acid, (iv) Linoleic acid, (v)Linolelaidic acid, (vi) Linolenic acid, (vii) Arochidic acid and (viii)Behenic acid. The results revealed that C. rugosa. produced lipaseproved to be suitable for biodiesel production; converting

Oleic acid, (iv) Linoleic acid, (v) Linolelaidic acid, (vi) Linolenic acid, (vii) Arochidic acid

C.L. Teo et al. / Renewable Energy 68 (2014) 1e5 5

microalgae lipids to fatty acid methyl ester (FAME). In previousstudies [15e17] conversion ratio of waste and plant oil to biodieselthrough immobilized lipase could reach up to 96%. Velasquez et al.[30] reported that palmitric stearic, linoleic and linolelaidic acidsare major components in FAMEs obtained from Chlorella vulgarislipid using alkali transesterification. In the case of soybean [31] andrapeseed oil [32] the FAMEs mostly consists of palmitric, stearic,linoleic and linolelaidic acids almost similar to that obtained frommicroalgae oils.

4. Conclusion

Tetraselmis sp. was successfully cultivated in F/2 mediumwithin22 days. The marine microalgae were cultured under aerated con-dition for better mixing of the marine microalgae in medium. Theresults revealed that exponential phase for Tetraselmis sp. growthwas achieved at day 18 of cultivation and highest biomass growthin nitrate concentration of 0.18 g/L. However, Nile red quantifica-tion of lipid content analysis showed the highest lipid content wasachieved in the absence of nitrate. The absence of nitrate limitsprotein biosynthesis, promotes lipid over protein ratio but biomassof marine microalgae is reduced. In the conversion of lipid to bio-diesel it was revealed that enzyme based transesterificationmethod was more superior than alkali based transesterificationwith a yield of 7 folds.

Acknowledgement

Financial support from Universiti Teknologi Malaysia (ResearchUniversity Grant/QJ1300.7125.00H03) for this research is gratefullyacknowledged.

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