Ultrasonics Sonochemistry xxx (2014) xxx–xxx
Contents lists available at ScienceDirect
Ultrasonics Sonochemistry journal homepage: www.elsevier.com/locate/ultson
Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass Raquel Rezende dos Santos a,⇑, Daniel Mendonça Moreira a, Claudete Norie Kunigami b, Donato Alexandre Gomes Aranda c, Cláudia Maria Luz Lapa Teixeira a a
Laboratório de Biotecnologia de Microalgas, Divisão de Energia, Instituto Nacional de Tecnologia, CEP: 20081-312, Rio de Janeiro, RJ, Brazil Laboratório de Análise Orgânica Instrumental, Divisão de Química Analítica, Instituto Nacional de Tecnologia, CEP: 20081-312, Rio de Janeiro, RJ, Brazil Laboratório de Tecnologia Verde, Departamento de Engenharia Química, Escola de Química, Centro de Tecnologia, Universidade Federal do Rio de Janeiro, CEP: 21941-590, Rio de Janeiro, RJ, Brazil b c
a r t i c l e
i n f o
Article history: Received 19 August 2013 Received in revised form 2 April 2014 Accepted 19 May 2014 Available online xxxx Keywords: Microalgae Biodiesel Lipid extraction Potter homogenizer Solvent Ultrasound
a b s t r a c t The use of lipids obtained from microalgae biomass has been described as a promising alternative for production of biodiesel to replace petro-diesel. It involves steps such as the cultivation of microalgae, biomass harvesting, extraction and transesterification of lipids. The purpose of the present study was to compare different methods of extracting total lipids. These methods were tested in biomass of Chlorella vulgaris with the solvents ethanol, hexane and a mixture of chloroform:methanol in ratios 1:2 and 2:1. The solvents were associated with other mechanisms of cell disruption such as use of a Potter homogenizer and ultrasound treatment. The percentage of triglycerides in the total lipids was determinated by the glycerol-3-phosphate oxidase-p-chlorophenol method (triglycerides monoreagent K117; Bioclin). Among the tested methods, the mixture of chloroform:methanol (2:1) assisted by ultrasound was most efficient, extracting an average of 19% of total lipids, of which 55% were triglycerides. The gas chromatographic analysis did not show differences in methyl ester profiles of oils extracted under the different methods. Ó 2014 Elsevier B.V. All rights reserved.
1. Introduction Given the current technological process and increased exploitation of new unconventional reserves (i.e., natural gas), it is probable that fossil fuels will continue to be available for a considerable period of time, although there may be variations in the supply and in the cost arising from geopolitical developments over time [1]. The steadily growing costs of petroleum, a desire for energy security in countries with limited petroleum resources and the inevitable depletion of fossil fuels are common concerns that have increased worldwide interest in biofuels [2]. In addition to these concerns, burning fossil fuels causes numerous environmental problems, including greenhouse gas (GHG) effects, which significantly contribute to global warming [3]. Microalgae are prokaryotic or eukaryotic photosynthetic microorganisms that can grow rapidly and live in harsh conditions due ⇑ Corresponding author. Tel.: +55 21 2123 1262. E-mail addresses: [email protected] (R.R. dos Santos), dm-moreira@ hotmail.com (D.M. Moreira), [email protected] (C.N. Kunigami), [email protected] (D.A.G. Aranda), [email protected] (Cláudia Maria Luz Lapa Teixeira).
to their simple cellular structure [4]. Known as one of the oldest life forms on the Earth, these microorganisms have a diversity of forms and ecological functions [5]. This diversity creates the capability for microalgae to be a valuable source in a multitude of products, such as value-added products for pharmaceutical purposes, food crops for human or animal consumption and as energy sources [4]. Microalgae biomass is considered a promising feedstock for producing a variety of renewable fuels, such as biodiesel, bioethanol, biohydrogen and methane [6,7]. Microalgae lipids have attracted attention as future raw materials for biodiesel synthesis, among others, because (1) microalgae have potential to attain higher lipid productivity in relation to oilseed crops [8]; (2) the biochemical composition of the microalgae biomass can be modulated by varying growth conditions, so the oil yield can be significantly increased [9]; (3) the microalgae biomass production can result in biofixation of waste CO2 (1 kg of dry microalgae biomass utilizes about 1.83 kg of CO2) [10]; (4) microalgae can be cultivated in brackish water or on non-arable land [11]; (5) the microalgae cultivation does not require application of herbicides or pesticides [12].
http://dx.doi.org/10.1016/j.ultsonch.2014.05.015 1350-4177/Ó 2014 Elsevier B.V. All rights reserved.
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
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R.R. dos Santos et al. / Ultrasonics Sonochemistry xxx (2014) xxx–xxx
The key processes involved in biodiesel production from microalgae are cultivation, harvesting, lipid extraction (cell disruption) and the transesterification of the lipids. Although all of these steps are essential, extraction is particularly important, as the contents of the extracted lipids are determined according to the disruption method and device. Therefore, the appropriate cell disruption method and device are keys to increasing the lipid extraction efficiency [13]. There are several reports of different methods for extracting lipids from microalgae, such as mechanical pressing, milling, supercritical fluid extraction, enzymatic extraction, microwave-assisted extraction, osmotic shock, homogenization, solvent extraction and ultrasonic-assisted extraction. The last three are evaluated here. While homogenization essentially involves using pressures to rupture cell walls, the solvent extraction entails extracting lipids by repeated washing or percolation with an organic solvent [14]. Some methods are usually used in combination with some kind of organic solvent. The application of ultrasound can enhance the extraction process due to a cavitation phenomenon. Ultrasonic waves create bubbles in the solvent, the bubbles burst near the cell walls, which produce shock waves, causing the release of lipid in the solvent [15]. All of these methods have their individual benefits and drawbacks. According to numerous reports in the literature about lipid extraction from microalgae biomass, the method’s efficiency depends on the species studied. But due to small number of studies with comparative analysis between these different methods, there have been no reports of the most efficient method of lipid extraction from Chlorella vulgaris biomass. The objective of the present study was to compare different methods of lipid extraction in relation to the total lipids and triglycerides. 2. Materials and methods 2.1. Microalga strain The microalga C. vulgaris was kindly donated by the Dr. Armando Augusto Henriques of São Carlos University Federal (UFSCar). Brazil. The strain was preserved in tubes containing 8 mL of sterile WC medium [16] for each 2 drops of culture, which was removed with a sterile Pasteur pipette. The tubes were kept in a germination chamber under 20 lmol of photons m 2 s 1 and 21 ± 1 °C manually shaken every 48 h. The WC medium was composed of TRIS buffer (0.5 g L 1), NaNO3 (0.085 g L 1), NaHCO3 (0.0126 g L 1), CaCl22H2O (0.03676 g L 1), MgSO47H2O (0.03697 g L 1), K2HPO4 (0.00871 g L 1), 1% H3BO3 (0.1 mL L 1), vitamin solution (1 mL L 1) and trace metals solution (1 mL L 1). The initial pH was adjusted to 8.5 with HCl 1 M. The vitamin solution was composed of thiamine (0.1 g L 1), cyanocobalamin (0.0005 g L 1) and biotin (0.0005 g L 1), being filtered through a 0.22-lm membrane. The trace metals solution was composed of CuSO45H2O (0.0098 g), ZnSO47H2O (0.022 g), CoCl26H2O (0.01 g), MnCl24H2O (0.18 g), NaMoO42H2O (0.0063 g) and chelated iron (1 L). The chelated iron was composed of Na2EDTA (4.36 g L 1) and FeCl36H2O (3.5 g L 1). 2.2. Obtaining microalgae biomass To obtain the inoculum, the cells were grow in 500-mL Erlenmeyer flasks containing 300 mL of WC medium. The flasks were kept under constant agitation of 180 rpm, 100 lmol of photons m 2 s 1 and 25 ± 2 °C. The culture was cultivated until it achieved an optical density at 730 nm (OD730nm) of approximately 0.8 (exponential phase). The inoculum obtained was transferred for clear 6-L bottles containing 5 L of WC medium. The cultures the feed batch were kept
under pneumatic stirring, light intensity of 100 lmol of photons m 2 s 1 and room temperature of 25 ± 2 °C until they achieved an OD730nm around 1.0 (stationary phase). The biomass obtained was harvested by centrifugation at 3500 rpm for 10 min, then lyophilized and stored at 4 °C until extractions. 2.2.1. Monitoring of biomass in culture medium The microalgae grown in culture medium were monitored by the optical density of the culture at 730 nm. The growth curve was plotted from the biomass value measurement in OD730nm and the growth phase determined when the growth curve was constructed in logarithmic scale. In order to maintain the inoculum, the microalgae were kept in exponential phase while the biomass obtained from extraction was harvested in stationary phase (about 25 days). 2.3. Lipid extraction Four methods of extraction were tested: ethanol [17], hexane [3], chloroform:methanol (1:2) [18] and chloroform:methanol (2:1) [19]. All solvents used in the extractions were of HPLC grade and were obtained from commercial source (Tedia Brazil). In the methods using ethanol and hexane, we added a mass of 0.5 g of dry microalgae for each 20 mL of solvent (ethanol or hexane) at room temperature (25 °C). This mixture was submitted to ultrasonic bath working at 40 kHz and producing an ultrasonic intensity of 34.74 W/L (Unique model 1800 USC – Indaiatuba, Brazil, 3.8 L, internal dimensions: 30 15.1 10 cm) during 20 min. The flask containing the mixture was submitted to ultrasonic bath with the help of a metal support to be centralized and not touch in the bottom tank. Later, the sample was centrifuged at 2000 rpm for 5 min. The organic phase was carefully collected and the solvent evaporated with a rotary evaporator at 60 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. In the method using chloroform:methanol (1:2) it was necessary to add 2 mL of distilled water for each 0.5 g of dry microalgae. A volume of 7.5 mL of the solvent mixture chloroform:methanol (1:2) was added to the wet biomass, and then 2.5 mL of chloroform and 2.5 mL of distilled water were added. This mixture was manually shaken during 3 min at room temperature. The biomass was harvested by centrifugation at 3500 rpm for 8 min at 4 °C. The organic phase was carefully collected and the solvent evaporated with a rotary evaporator at 50 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. The extraction using chloroform:methanol (1:2) was realized again replacing the manual agitation by a Potter homogenizer or ultrasound at room temperature. In the first case, the mixture was processed in a Potter homogenizer (Tecnal model TE-099 – Piracicaba, Brazil, internal dimensions: 30 35 54 cm) at medium speed during 3 min, while in the second case the mixture was submitted to ultrasonic bath during 20 min. In the methods using chloroform:methanol (2:1), we added a mass of 0.5 g of dry microalgae for each 36 mL of the solvent mixture at room temperature. This mixture was processed in a Potter homogenizer at medium speed during 3 min at room temperature. Later the sample was centrifuged at 3500 rpm for 8 min at 4 °C. The organic phase was carefully collected and transferred for another tube to which 9 mL of 0.88% KCl was added. At this moment, there were two phases and the upper phase was discarded with a pipette. Then 4.5 mL of chloroform:methanol:water (3:48:47) was added to the lower phase, so that two phases formed again. Again the upper phase was discarded with pipette. The washing with chloroform:methanol:water was repeated two times. The organic phase was carefully collected and the solvent evaporated
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
R.R. dos Santos et al. / Ultrasonics Sonochemistry xxx (2014) xxx–xxx
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with a rotary evaporator at 40 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. The extraction using chloroform:methanol (2:1) was realized again replacing the processing in the Potter homogenizer with ultrasound treatment at room temperature. In this case, the mixture was submitted to ultrasonic bath during 20 min. The lipid mass was determined by gravimetric analysis. All experiments were carried out in triplicate and the data are reported as mean ± standard deviation. The determination of the triglycerides was carried out in all of the lipids samples obtained (using different methods) but the fatty acid profiles were only determined in the lipid sample obtained in the most efficient extraction method. 2.4. Quantification of triglycerides The mass of triglycerides in the total lipids solubilized in isopropyl alcohol was determined by the glycerol-3-phosphate oxidasep-chlorophenol method [20] (monoreagent triglycerides K117; Bioclin, Brazil) using triolein as the standard.
Fig. 1. Total lipid percentage in the dry biomass using different extraction methods. 1 – ethanol, 2 – hexane, 3 – chloroform:methanol (1:2), 4 – chloroform:methanol (1:2) assisted by a Potter homogenizer, 5 – chloroform:methanol (1:2) assisted by ultrasound, 6 – chloroform:methanol (2:1) assisted by a Potter homogenizer, 7 – chloroform:methanol (2:1) assisted by ultrasound.
2.5. Fatty acid profiles The transesterification reaction in the total lipid fraction was carried out according to the ASTM D 3457 standard method. The analysis by gas chromatography and detection by mass spectrometry (Agilent- model 6890N gas chromatograph and model 5975 model mass spectrometer, USA) were performed in an HP-5MS column (30 m length 0.25 mm i.d. 0.25 lm, stationary phase) with split injection of 10:1 and injection volume of 1 lL. The initial temperature of the oven was 40 °C, which was increased until 300 °C at a temperature gradient of 10 °C min 1. Helium was used as the carrier gas with flow of 1 mL min 1. The fatty acids were identified by consulting the Wiley 7 Nist 05 digital library’s mass spectral database. 2.6. Statistical analysis
Fig. 2. Triglyceride percentage in the total lipids using different extraction methods. 1 – ethanol, 2 – hexane, 3 – chloroform:methanol (1:2), 4 – chloroform:methanol (1:2) assisted by a Potter homogenizer, 5 – chloroform:methanol (1:2) assisted by ultrasound, 6 – chloroform:methanol (2:1) assisted by a Potter homogenizer, 7 – chloroform:methanol (2:1) assisted by ultrasound.
The results obtained were evaluated using a one-way ANOVA and the Turkey test. The level of significance adopted was p < 0.05. 3. Results and discussion Seven methods for lipid extraction were evaluated in C. vulgaris biomass: ethanol assisted by ultrasound; hexane assisted by ultrasound; chloroform:methanol (1:2) and chloroform:methanol (1:2) assisted by Potter homogenizer treatment; chloroform:methanol (1:2) assisted by ultrasound; chloroform:methanol (2:1) assisted by Potter homogenizer treatment; and chloroform:methanol (2:1) assisted by ultrasound. Fig. 1 shows the comparative analysis of different lipid extraction methods in relation to the total lipid content in the dry biomass. After total lipid quantification, we evaluated the triglyceride content in each total lipid extract. Among the tested methods, the mixture of chloroform:methanol (2:1) assisted by ultrasound was the most efficient, obtaining approximately 19% of total lipids. Of this total lipid content, approximately 55% was in the form of triglycerides, as indicated by Fig. 2. When a microalga cell is exposed to a non-polar organic solvent, such as hexane and chloroform, the solvent penetrates the cell membrane and interacts with neutral lipids (i.e., triglycerides) present in the cytoplasm. The solvent interacts with lipids because of van der Walls forces to form an organic solvent-neutral lipid complex. This complex, driven by a concentration gradient, diffuses across the cell membrane and the static organic solvent film surrounding the cell, entering the organic solvent. As a result,
the neutral lipids are extracted from the cells and remain dissolved in the non-polar organic solvent. Some neutral lipids are found in the cytoplasm as a complex with polar lipids (i.e., phospholipids, glycolipids, sterols, carotenoids). This complex is strongly linked via hydrogen bonds to proteins in the cell membrane. The van der Waals interactions formed between non-polar organic solvents and neutral lipids are inadequate to disrupt these membranebased lipid-protein associations, making it necessary to use a polar organic solvent, such as methanol or ethanol [21]. Mixtures of chloroform:methanol are used to ensure complete extraction of all the neutral lipids, both in the form of free globules and in the form of complex associated proteins. Thus, these mixtures extract both neutral lipids by chloroform and polar lipids by methanol. For this reason, mixtures of chloroform:methanol tend to be more efficient in the extraction of total lipids. Mixtures of chloroform:methanol extract hydrocarbons, carotenoids, chlorophyll, sterols, triglycerides, waxes, aldehydes, fatty acids, phospholipids and glycolipids. The ultrasonic-assisted extraction has been recognized as an efficient extraction technique that reduces working time and increases lipid yields from oilseed and microalgae. During extraction assisted by ultrasound, the cavitation, mechanical function and thermal function together enhance the efficiency of extraction. Fig. 3 illustrates the effect of ultrasonic-assisted extraction in the C. vulgaris cell.
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
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R.R. dos Santos et al. / Ultrasonics Sonochemistry xxx (2014) xxx–xxx
(A)
(B)
Fig. 3. Chlorella vulgaris cell before and after extraction. (A) Control - cells before being subjected to extraction. Arrow indicates the intact cell. (B) Disrupted cells - extracted with chloroform: methanol (2:1) assisted by ultrasound during 20 minutes. Arrow indicates the disrupted cell.
Under the ultrasound irradiation, micro-bubbles are created when the negative pressure is high enough. Once created, the bubbles grow during the period of negative pressure and shrink during the period of positive pressure. The expansion and compression can cause constant pulsation or violent collapse of micro-bubbles. When collapse occurs near the solid surface, it can damage the cell walls to facilitate the release of the contents. Meanwhile, promotion of solvent penetration into cell walls by mechanical function and decrease of solvent viscosity by thermal function can also improve mass transfer [15]. This effect is much stronger at low frequencies (18–40 kHz). In case of dry cells, the ultrasound accelerates their rehydration and swelling [22]. The result obtained in the present study corroborates the finding of related study by Converti et al. [23] The authors tested several extraction methods in Nannochloropsis oculata biomass at different times. After 6 h of extraction, the method using chloroform:methanol (2:1) assisted by ultrasound produced the best results, obtaining 24.5% of total lipids. Montes D’Oca et al. [17] evaluated the efficiency of some lipid extraction methods too. They used Chlorella pyrenoidosa biomass and all methods assisted by ultrasound during 20 min. The extraction with chloroform:methanol (2:1) extracted the highest percentage of total lipids (12.29%). Although lipid extraction has been evaluated mostly through the action of organic solvents (due to the hydrophobicity of lipids), extraction assisted by ultrasound has also received attention; not only with regard to lipids from microalgae, but from oilseed and animal tissues [22]. Known as a method of cold extraction (i.e., room temperature), the use of ultrasound has received attention because it does not change the quality of the lipid fraction and it reduces the amount of solvent used. Many times when comparing different lipid extraction methods, authors have evaluated whether different protocols generate differences in fatty acid profiles. Usually, the gas chromatographic analysis shows only slight or negligible differences in methyl ester profiles of extracted oils [22,24]. Here we only reported the fatty
Table 1 Fatty acid profile (%) in Chlorella vulgaris cells extracted with chloroform: methanol (2:1) with ultrasound during 20 minutes. Acid name
Composition
Content (%)
Myristic Palmitic Palmitoleic Stearic Linoleic c Linolenic Arachidonic Others
C14:0 C16:0 C16:1 C18:0 C18:2 C18:3 C20:4
2,10 20,69 8,39 5,77 35,72 1,15 1,38 24,81
acid profile of the most efficient protocol. Table 1 summarizes the fatty acid composition of the C. vulgaris oil extracted with chloroform:methanol (2:1) assisted by ultrasound. Unsaturated oils produce a biodiesel that is more susceptible to oxidation due to their instability when stored for long periods. Therefore, the fatty acid profile is mainly evaluated to detect polyunsaturated fatty acids (PUFA) [25]. The oil extracted contains the highest percentage of unsaturated fatty acids and lowest percentage of saturated fatty acids. However, it is worth remembering that the content and lipid profile can change depending on the variations in growing conditions and species evaluated. Thus, by determining the most efficient extraction method to obtain total lipids from C. vulgaris, this study contributes to finding the best growing conditions for this microalga. 4. Conclusions The use of the mixture of organic solvents chloroform:methanol (2:1) assisted by ultrasound is the most efficient method for extraction of lipids and TAGs from C. vulgaris biomass. The use of this mixture may facilitate extraction of neutral lipids (TAGs), both in free form and in the form of complexes. The use of ultrasound favors cell disruption and increases the extraction yield. The methyl ester profiles obtained by gas chromatography were very similar for the oils extracted under different methods. Acknowledgements We thank the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) and Financiadora de Estudos e Projetos (FINEP) for their financial support of this work. References [1] L. Brennam, P. Owende, Biofuels from microalgae-A review of technologies for production, processing, and extractions of biofuels and co-products, Renewable Sustainable Energy Rev. 14 (2010) 557–577. [2] C.K. Westbrook, Biofuels combustion, Annu. Rev. Phys. Chem. 64 (2013) 201–219. [3] M.K. Lam, K.T. Lee, Microalgae biofuels: a critical review of issues, problems and the way forward, Biotechnol. Adv. 30 (2012) 673–690. [4] T.M. Mata, A.A. Martins, N.S. Caetano, Microalgae for biodiesel production and other applications: a review, Renewable Sustainable Energy Rev. 14 (2010) 217–232. [5] D.B. Stengel, S. Connan, Z.A. Popper, Algal chemodiversity and bioactivity: sources of natural variability and implications for commercial application, Biotechnol. Adv. 29 (2011) 483–501. [6] C. Posten, G. Schaub, Microalgae and terrestrial biomass as source for fuels-A process view, J. Biotechnol. 142 (2009) 64–69. [7] J.O.B. Carioca, Biofuels: problems, challenges and perspectives, Biotechnol. J. 5 (2010) 260–273.
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Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
Contents lists available at ScienceDirect
Ultrasonics Sonochemistry journal homepage: www.elsevier.com/locate/ultson
Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass Raquel Rezende dos Santos a,⇑, Daniel Mendonça Moreira a, Claudete Norie Kunigami b, Donato Alexandre Gomes Aranda c, Cláudia Maria Luz Lapa Teixeira a a
Laboratório de Biotecnologia de Microalgas, Divisão de Energia, Instituto Nacional de Tecnologia, CEP: 20081-312, Rio de Janeiro, RJ, Brazil Laboratório de Análise Orgânica Instrumental, Divisão de Química Analítica, Instituto Nacional de Tecnologia, CEP: 20081-312, Rio de Janeiro, RJ, Brazil Laboratório de Tecnologia Verde, Departamento de Engenharia Química, Escola de Química, Centro de Tecnologia, Universidade Federal do Rio de Janeiro, CEP: 21941-590, Rio de Janeiro, RJ, Brazil b c
a r t i c l e
i n f o
Article history: Received 19 August 2013 Received in revised form 2 April 2014 Accepted 19 May 2014 Available online xxxx Keywords: Microalgae Biodiesel Lipid extraction Potter homogenizer Solvent Ultrasound
a b s t r a c t The use of lipids obtained from microalgae biomass has been described as a promising alternative for production of biodiesel to replace petro-diesel. It involves steps such as the cultivation of microalgae, biomass harvesting, extraction and transesterification of lipids. The purpose of the present study was to compare different methods of extracting total lipids. These methods were tested in biomass of Chlorella vulgaris with the solvents ethanol, hexane and a mixture of chloroform:methanol in ratios 1:2 and 2:1. The solvents were associated with other mechanisms of cell disruption such as use of a Potter homogenizer and ultrasound treatment. The percentage of triglycerides in the total lipids was determinated by the glycerol-3-phosphate oxidase-p-chlorophenol method (triglycerides monoreagent K117; Bioclin). Among the tested methods, the mixture of chloroform:methanol (2:1) assisted by ultrasound was most efficient, extracting an average of 19% of total lipids, of which 55% were triglycerides. The gas chromatographic analysis did not show differences in methyl ester profiles of oils extracted under the different methods. Ó 2014 Elsevier B.V. All rights reserved.
1. Introduction Given the current technological process and increased exploitation of new unconventional reserves (i.e., natural gas), it is probable that fossil fuels will continue to be available for a considerable period of time, although there may be variations in the supply and in the cost arising from geopolitical developments over time [1]. The steadily growing costs of petroleum, a desire for energy security in countries with limited petroleum resources and the inevitable depletion of fossil fuels are common concerns that have increased worldwide interest in biofuels [2]. In addition to these concerns, burning fossil fuels causes numerous environmental problems, including greenhouse gas (GHG) effects, which significantly contribute to global warming [3]. Microalgae are prokaryotic or eukaryotic photosynthetic microorganisms that can grow rapidly and live in harsh conditions due ⇑ Corresponding author. Tel.: +55 21 2123 1262. E-mail addresses: [email protected] (R.R. dos Santos), dm-moreira@ hotmail.com (D.M. Moreira), [email protected] (C.N. Kunigami), [email protected] (D.A.G. Aranda), [email protected] (Cláudia Maria Luz Lapa Teixeira).
to their simple cellular structure [4]. Known as one of the oldest life forms on the Earth, these microorganisms have a diversity of forms and ecological functions [5]. This diversity creates the capability for microalgae to be a valuable source in a multitude of products, such as value-added products for pharmaceutical purposes, food crops for human or animal consumption and as energy sources [4]. Microalgae biomass is considered a promising feedstock for producing a variety of renewable fuels, such as biodiesel, bioethanol, biohydrogen and methane [6,7]. Microalgae lipids have attracted attention as future raw materials for biodiesel synthesis, among others, because (1) microalgae have potential to attain higher lipid productivity in relation to oilseed crops [8]; (2) the biochemical composition of the microalgae biomass can be modulated by varying growth conditions, so the oil yield can be significantly increased [9]; (3) the microalgae biomass production can result in biofixation of waste CO2 (1 kg of dry microalgae biomass utilizes about 1.83 kg of CO2) [10]; (4) microalgae can be cultivated in brackish water or on non-arable land [11]; (5) the microalgae cultivation does not require application of herbicides or pesticides [12].
http://dx.doi.org/10.1016/j.ultsonch.2014.05.015 1350-4177/Ó 2014 Elsevier B.V. All rights reserved.
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
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R.R. dos Santos et al. / Ultrasonics Sonochemistry xxx (2014) xxx–xxx
The key processes involved in biodiesel production from microalgae are cultivation, harvesting, lipid extraction (cell disruption) and the transesterification of the lipids. Although all of these steps are essential, extraction is particularly important, as the contents of the extracted lipids are determined according to the disruption method and device. Therefore, the appropriate cell disruption method and device are keys to increasing the lipid extraction efficiency [13]. There are several reports of different methods for extracting lipids from microalgae, such as mechanical pressing, milling, supercritical fluid extraction, enzymatic extraction, microwave-assisted extraction, osmotic shock, homogenization, solvent extraction and ultrasonic-assisted extraction. The last three are evaluated here. While homogenization essentially involves using pressures to rupture cell walls, the solvent extraction entails extracting lipids by repeated washing or percolation with an organic solvent [14]. Some methods are usually used in combination with some kind of organic solvent. The application of ultrasound can enhance the extraction process due to a cavitation phenomenon. Ultrasonic waves create bubbles in the solvent, the bubbles burst near the cell walls, which produce shock waves, causing the release of lipid in the solvent [15]. All of these methods have their individual benefits and drawbacks. According to numerous reports in the literature about lipid extraction from microalgae biomass, the method’s efficiency depends on the species studied. But due to small number of studies with comparative analysis between these different methods, there have been no reports of the most efficient method of lipid extraction from Chlorella vulgaris biomass. The objective of the present study was to compare different methods of lipid extraction in relation to the total lipids and triglycerides. 2. Materials and methods 2.1. Microalga strain The microalga C. vulgaris was kindly donated by the Dr. Armando Augusto Henriques of São Carlos University Federal (UFSCar). Brazil. The strain was preserved in tubes containing 8 mL of sterile WC medium [16] for each 2 drops of culture, which was removed with a sterile Pasteur pipette. The tubes were kept in a germination chamber under 20 lmol of photons m 2 s 1 and 21 ± 1 °C manually shaken every 48 h. The WC medium was composed of TRIS buffer (0.5 g L 1), NaNO3 (0.085 g L 1), NaHCO3 (0.0126 g L 1), CaCl22H2O (0.03676 g L 1), MgSO47H2O (0.03697 g L 1), K2HPO4 (0.00871 g L 1), 1% H3BO3 (0.1 mL L 1), vitamin solution (1 mL L 1) and trace metals solution (1 mL L 1). The initial pH was adjusted to 8.5 with HCl 1 M. The vitamin solution was composed of thiamine (0.1 g L 1), cyanocobalamin (0.0005 g L 1) and biotin (0.0005 g L 1), being filtered through a 0.22-lm membrane. The trace metals solution was composed of CuSO45H2O (0.0098 g), ZnSO47H2O (0.022 g), CoCl26H2O (0.01 g), MnCl24H2O (0.18 g), NaMoO42H2O (0.0063 g) and chelated iron (1 L). The chelated iron was composed of Na2EDTA (4.36 g L 1) and FeCl36H2O (3.5 g L 1). 2.2. Obtaining microalgae biomass To obtain the inoculum, the cells were grow in 500-mL Erlenmeyer flasks containing 300 mL of WC medium. The flasks were kept under constant agitation of 180 rpm, 100 lmol of photons m 2 s 1 and 25 ± 2 °C. The culture was cultivated until it achieved an optical density at 730 nm (OD730nm) of approximately 0.8 (exponential phase). The inoculum obtained was transferred for clear 6-L bottles containing 5 L of WC medium. The cultures the feed batch were kept
under pneumatic stirring, light intensity of 100 lmol of photons m 2 s 1 and room temperature of 25 ± 2 °C until they achieved an OD730nm around 1.0 (stationary phase). The biomass obtained was harvested by centrifugation at 3500 rpm for 10 min, then lyophilized and stored at 4 °C until extractions. 2.2.1. Monitoring of biomass in culture medium The microalgae grown in culture medium were monitored by the optical density of the culture at 730 nm. The growth curve was plotted from the biomass value measurement in OD730nm and the growth phase determined when the growth curve was constructed in logarithmic scale. In order to maintain the inoculum, the microalgae were kept in exponential phase while the biomass obtained from extraction was harvested in stationary phase (about 25 days). 2.3. Lipid extraction Four methods of extraction were tested: ethanol [17], hexane [3], chloroform:methanol (1:2) [18] and chloroform:methanol (2:1) [19]. All solvents used in the extractions were of HPLC grade and were obtained from commercial source (Tedia Brazil). In the methods using ethanol and hexane, we added a mass of 0.5 g of dry microalgae for each 20 mL of solvent (ethanol or hexane) at room temperature (25 °C). This mixture was submitted to ultrasonic bath working at 40 kHz and producing an ultrasonic intensity of 34.74 W/L (Unique model 1800 USC – Indaiatuba, Brazil, 3.8 L, internal dimensions: 30 15.1 10 cm) during 20 min. The flask containing the mixture was submitted to ultrasonic bath with the help of a metal support to be centralized and not touch in the bottom tank. Later, the sample was centrifuged at 2000 rpm for 5 min. The organic phase was carefully collected and the solvent evaporated with a rotary evaporator at 60 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. In the method using chloroform:methanol (1:2) it was necessary to add 2 mL of distilled water for each 0.5 g of dry microalgae. A volume of 7.5 mL of the solvent mixture chloroform:methanol (1:2) was added to the wet biomass, and then 2.5 mL of chloroform and 2.5 mL of distilled water were added. This mixture was manually shaken during 3 min at room temperature. The biomass was harvested by centrifugation at 3500 rpm for 8 min at 4 °C. The organic phase was carefully collected and the solvent evaporated with a rotary evaporator at 50 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. The extraction using chloroform:methanol (1:2) was realized again replacing the manual agitation by a Potter homogenizer or ultrasound at room temperature. In the first case, the mixture was processed in a Potter homogenizer (Tecnal model TE-099 – Piracicaba, Brazil, internal dimensions: 30 35 54 cm) at medium speed during 3 min, while in the second case the mixture was submitted to ultrasonic bath during 20 min. In the methods using chloroform:methanol (2:1), we added a mass of 0.5 g of dry microalgae for each 36 mL of the solvent mixture at room temperature. This mixture was processed in a Potter homogenizer at medium speed during 3 min at room temperature. Later the sample was centrifuged at 3500 rpm for 8 min at 4 °C. The organic phase was carefully collected and transferred for another tube to which 9 mL of 0.88% KCl was added. At this moment, there were two phases and the upper phase was discarded with a pipette. Then 4.5 mL of chloroform:methanol:water (3:48:47) was added to the lower phase, so that two phases formed again. Again the upper phase was discarded with pipette. The washing with chloroform:methanol:water was repeated two times. The organic phase was carefully collected and the solvent evaporated
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
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with a rotary evaporator at 40 °C. The lipid fraction was dried to constant weight in an oven with air circulation at 30 °C. The extraction using chloroform:methanol (2:1) was realized again replacing the processing in the Potter homogenizer with ultrasound treatment at room temperature. In this case, the mixture was submitted to ultrasonic bath during 20 min. The lipid mass was determined by gravimetric analysis. All experiments were carried out in triplicate and the data are reported as mean ± standard deviation. The determination of the triglycerides was carried out in all of the lipids samples obtained (using different methods) but the fatty acid profiles were only determined in the lipid sample obtained in the most efficient extraction method. 2.4. Quantification of triglycerides The mass of triglycerides in the total lipids solubilized in isopropyl alcohol was determined by the glycerol-3-phosphate oxidasep-chlorophenol method [20] (monoreagent triglycerides K117; Bioclin, Brazil) using triolein as the standard.
Fig. 1. Total lipid percentage in the dry biomass using different extraction methods. 1 – ethanol, 2 – hexane, 3 – chloroform:methanol (1:2), 4 – chloroform:methanol (1:2) assisted by a Potter homogenizer, 5 – chloroform:methanol (1:2) assisted by ultrasound, 6 – chloroform:methanol (2:1) assisted by a Potter homogenizer, 7 – chloroform:methanol (2:1) assisted by ultrasound.
2.5. Fatty acid profiles The transesterification reaction in the total lipid fraction was carried out according to the ASTM D 3457 standard method. The analysis by gas chromatography and detection by mass spectrometry (Agilent- model 6890N gas chromatograph and model 5975 model mass spectrometer, USA) were performed in an HP-5MS column (30 m length 0.25 mm i.d. 0.25 lm, stationary phase) with split injection of 10:1 and injection volume of 1 lL. The initial temperature of the oven was 40 °C, which was increased until 300 °C at a temperature gradient of 10 °C min 1. Helium was used as the carrier gas with flow of 1 mL min 1. The fatty acids were identified by consulting the Wiley 7 Nist 05 digital library’s mass spectral database. 2.6. Statistical analysis
Fig. 2. Triglyceride percentage in the total lipids using different extraction methods. 1 – ethanol, 2 – hexane, 3 – chloroform:methanol (1:2), 4 – chloroform:methanol (1:2) assisted by a Potter homogenizer, 5 – chloroform:methanol (1:2) assisted by ultrasound, 6 – chloroform:methanol (2:1) assisted by a Potter homogenizer, 7 – chloroform:methanol (2:1) assisted by ultrasound.
The results obtained were evaluated using a one-way ANOVA and the Turkey test. The level of significance adopted was p < 0.05. 3. Results and discussion Seven methods for lipid extraction were evaluated in C. vulgaris biomass: ethanol assisted by ultrasound; hexane assisted by ultrasound; chloroform:methanol (1:2) and chloroform:methanol (1:2) assisted by Potter homogenizer treatment; chloroform:methanol (1:2) assisted by ultrasound; chloroform:methanol (2:1) assisted by Potter homogenizer treatment; and chloroform:methanol (2:1) assisted by ultrasound. Fig. 1 shows the comparative analysis of different lipid extraction methods in relation to the total lipid content in the dry biomass. After total lipid quantification, we evaluated the triglyceride content in each total lipid extract. Among the tested methods, the mixture of chloroform:methanol (2:1) assisted by ultrasound was the most efficient, obtaining approximately 19% of total lipids. Of this total lipid content, approximately 55% was in the form of triglycerides, as indicated by Fig. 2. When a microalga cell is exposed to a non-polar organic solvent, such as hexane and chloroform, the solvent penetrates the cell membrane and interacts with neutral lipids (i.e., triglycerides) present in the cytoplasm. The solvent interacts with lipids because of van der Walls forces to form an organic solvent-neutral lipid complex. This complex, driven by a concentration gradient, diffuses across the cell membrane and the static organic solvent film surrounding the cell, entering the organic solvent. As a result,
the neutral lipids are extracted from the cells and remain dissolved in the non-polar organic solvent. Some neutral lipids are found in the cytoplasm as a complex with polar lipids (i.e., phospholipids, glycolipids, sterols, carotenoids). This complex is strongly linked via hydrogen bonds to proteins in the cell membrane. The van der Waals interactions formed between non-polar organic solvents and neutral lipids are inadequate to disrupt these membranebased lipid-protein associations, making it necessary to use a polar organic solvent, such as methanol or ethanol [21]. Mixtures of chloroform:methanol are used to ensure complete extraction of all the neutral lipids, both in the form of free globules and in the form of complex associated proteins. Thus, these mixtures extract both neutral lipids by chloroform and polar lipids by methanol. For this reason, mixtures of chloroform:methanol tend to be more efficient in the extraction of total lipids. Mixtures of chloroform:methanol extract hydrocarbons, carotenoids, chlorophyll, sterols, triglycerides, waxes, aldehydes, fatty acids, phospholipids and glycolipids. The ultrasonic-assisted extraction has been recognized as an efficient extraction technique that reduces working time and increases lipid yields from oilseed and microalgae. During extraction assisted by ultrasound, the cavitation, mechanical function and thermal function together enhance the efficiency of extraction. Fig. 3 illustrates the effect of ultrasonic-assisted extraction in the C. vulgaris cell.
Please cite this article in press as: R.R. dos Santos et al., Comparison between several methods of total lipid extraction from Chlorella vulgaris biomass, Ultrason. Sonochem. (2014), http://dx.doi.org/10.1016/j.ultsonch.2014.05.015
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(A)
(B)
Fig. 3. Chlorella vulgaris cell before and after extraction. (A) Control - cells before being subjected to extraction. Arrow indicates the intact cell. (B) Disrupted cells - extracted with chloroform: methanol (2:1) assisted by ultrasound during 20 minutes. Arrow indicates the disrupted cell.
Under the ultrasound irradiation, micro-bubbles are created when the negative pressure is high enough. Once created, the bubbles grow during the period of negative pressure and shrink during the period of positive pressure. The expansion and compression can cause constant pulsation or violent collapse of micro-bubbles. When collapse occurs near the solid surface, it can damage the cell walls to facilitate the release of the contents. Meanwhile, promotion of solvent penetration into cell walls by mechanical function and decrease of solvent viscosity by thermal function can also improve mass transfer [15]. This effect is much stronger at low frequencies (18–40 kHz). In case of dry cells, the ultrasound accelerates their rehydration and swelling [22]. The result obtained in the present study corroborates the finding of related study by Converti et al. [23] The authors tested several extraction methods in Nannochloropsis oculata biomass at different times. After 6 h of extraction, the method using chloroform:methanol (2:1) assisted by ultrasound produced the best results, obtaining 24.5% of total lipids. Montes D’Oca et al. [17] evaluated the efficiency of some lipid extraction methods too. They used Chlorella pyrenoidosa biomass and all methods assisted by ultrasound during 20 min. The extraction with chloroform:methanol (2:1) extracted the highest percentage of total lipids (12.29%). Although lipid extraction has been evaluated mostly through the action of organic solvents (due to the hydrophobicity of lipids), extraction assisted by ultrasound has also received attention; not only with regard to lipids from microalgae, but from oilseed and animal tissues [22]. Known as a method of cold extraction (i.e., room temperature), the use of ultrasound has received attention because it does not change the quality of the lipid fraction and it reduces the amount of solvent used. Many times when comparing different lipid extraction methods, authors have evaluated whether different protocols generate differences in fatty acid profiles. Usually, the gas chromatographic analysis shows only slight or negligible differences in methyl ester profiles of extracted oils [22,24]. Here we only reported the fatty
Table 1 Fatty acid profile (%) in Chlorella vulgaris cells extracted with chloroform: methanol (2:1) with ultrasound during 20 minutes. Acid name
Composition
Content (%)
Myristic Palmitic Palmitoleic Stearic Linoleic c Linolenic Arachidonic Others
C14:0 C16:0 C16:1 C18:0 C18:2 C18:3 C20:4
2,10 20,69 8,39 5,77 35,72 1,15 1,38 24,81
acid profile of the most efficient protocol. Table 1 summarizes the fatty acid composition of the C. vulgaris oil extracted with chloroform:methanol (2:1) assisted by ultrasound. Unsaturated oils produce a biodiesel that is more susceptible to oxidation due to their instability when stored for long periods. Therefore, the fatty acid profile is mainly evaluated to detect polyunsaturated fatty acids (PUFA) [25]. The oil extracted contains the highest percentage of unsaturated fatty acids and lowest percentage of saturated fatty acids. However, it is worth remembering that the content and lipid profile can change depending on the variations in growing conditions and species evaluated. Thus, by determining the most efficient extraction method to obtain total lipids from C. vulgaris, this study contributes to finding the best growing conditions for this microalga. 4. Conclusions The use of the mixture of organic solvents chloroform:methanol (2:1) assisted by ultrasound is the most efficient method for extraction of lipids and TAGs from C. vulgaris biomass. The use of this mixture may facilitate extraction of neutral lipids (TAGs), both in free form and in the form of complexes. The use of ultrasound favors cell disruption and increases the extraction yield. The methyl ester profiles obtained by gas chromatography were very similar for the oils extracted under different methods. Acknowledgements We thank the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) and Financiadora de Estudos e Projetos (FINEP) for their financial support of this work. References [1] L. Brennam, P. Owende, Biofuels from microalgae-A review of technologies for production, processing, and extractions of biofuels and co-products, Renewable Sustainable Energy Rev. 14 (2010) 557–577. [2] C.K. Westbrook, Biofuels combustion, Annu. Rev. Phys. Chem. 64 (2013) 201–219. [3] M.K. Lam, K.T. Lee, Microalgae biofuels: a critical review of issues, problems and the way forward, Biotechnol. Adv. 30 (2012) 673–690. [4] T.M. Mata, A.A. Martins, N.S. Caetano, Microalgae for biodiesel production and other applications: a review, Renewable Sustainable Energy Rev. 14 (2010) 217–232. [5] D.B. Stengel, S. Connan, Z.A. Popper, Algal chemodiversity and bioactivity: sources of natural variability and implications for commercial application, Biotechnol. Adv. 29 (2011) 483–501. [6] C. Posten, G. Schaub, Microalgae and terrestrial biomass as source for fuels-A process view, J. Biotechnol. 142 (2009) 64–69. [7] J.O.B. Carioca, Biofuels: problems, challenges and perspectives, Biotechnol. J. 5 (2010) 260–273.
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