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Characterization and identification of lipid-producing microalgae species isolated from a freshwater lake
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Characterization and identification of lipid-producing microalgae species isolated from a freshwater lake Reda A.I. Abou-Shanab a,b, Ibrahim A. Matter a, Su-Nam Kim c, You-Kwan Oh d, Jaeyoung Choi c, Byong-Hun Jeon a,* a
Department of Environmental Engineering, Yonsei University, Wonju, Gangwon-do 220-710, South Korea Department of Environmental Biotechnology, Mubarak City for Scientific Research and Technology Applications, New Borg El Arab City, Alexandria 21934, Egypt c KIST Gangneung Institute, Gangneung 210-340, South Korea d Bioenergy Center, Korea Institute of Energy Research, Daejeon 305-343, South Korea b
article info
abstract
Article history:
Microalgal lipids are the oils of the future for sustainable biodiesel production. One of the
Received 31 March 2010
most important decisions in obtaining oil from microalgae is the choice of species. A total
Received in revised form
of 45 algal cultures were isolated from a freshwater lake at Wonju, South Korea. Five
10 April 2011
microalgal isolates were selected based on their morphology and ease of cultivation
Accepted 15 April 2011
under our test conditions. These cultures were identified as strains of Scenedesmus obli-
Available online 5 May 2011
quus YSL02, Chlamydomonas pitschmannii YSL03, Chlorella vulgaris YSL04, S. obliquus YSL05, and Chlamydomonas mexicana YSL07 based on microscopic examination and LSU rDNA
Keywords:
(D1-D2) sequence analysis. S. obliquus YSL02 reached a higher biomass concentration
Biodiesel
(1.84 0.30 g L1) with a lower lipid content (29% w/w), than did Chla. pitschmannii YSL03
Freshwater
(maximum biomass concentration of 1.04 0.09 with a 51% lipid content). Our results
Isolation
suggest that Chla. pitschmannii YSL03 is appropriate for producing biodiesel based on its
LSU rDNA (D1-D2)
high lipid content and oleic acid proportion. ª 2011 Elsevier Ltd. All rights reserved.
Microalgae
1.
Introduction
The basic resources currently exploited to obtain energy are petroleum, natural gas, coal, hydropower, and nuclear power. Continued use of petroleum-based fuels is now widely recognized as unsustainable because of limited supplies and the contribution of these fuels to atmospheric pollution. Fossil fuel combustion is also a major source of greenhouse gases responsible for global warming. Renewable, carbon-neutral, economically viable alternatives to fossil fuels are urgently needed to avert the impending oil crisis and the dramatic consequences of climate change [1].
Biomass is one of the better sources of energy to mitigate greenhouse gas emissions and to function as a substitute for fossil fuels [2]. Large-scale introduction of biomass energy could contribute to sustainable development on environmental, social, and economic fronts. Biodiesel (monoalkyl esters) is one such alternative fuel, obtained by the transesterification of triglyceride oil with monohydric alcohols. Commercial biodiesel has been obtained successfully from rapeseed, soybean, sunflower, corn, palm, and waste cooking oil, as well as from animal fat [3]. However, large-scale production of biodiesel from those resources cannot realistically satisfy the existing demand for transport fuels [1]. Biodiesel has received considerable
* Corresponding author. Tel.: þ82 33 760 2446; fax: þ82 33 760 2571. E-mail address: [email protected] (B.-H. Jeon). 0961-9534/$ e see front matter ª 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biombioe.2011.04.021
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attention in recent years, as it is a biodegradable, renewable, and non-toxic fuel. It contributes no net carbon dioxide or sulfur to the atmosphere and emits less gaseous pollutants than does normal diesel [3]. While biodiesel is a desirable product, the significant economic and environmental impacts of using agricultural crops, especially food crops, as a feedstock for biofuels has raised crucial sustainability issues. For example, the transformation of primary food resources into bio-fuels has led to a clash of interests, as using food crops to derived biodiesel has resulted in a reduced supply to poor countries and an increase in food costs [4]. All of these factors have stimulated the search for other sources of biodiesel production that are both sustainable and economical. Microalgae are microscopic heterotrophiceautotrophic photosynthesizing organisms that are able to use solar energy to combine water with carbon dioxide to create biomass. Microalgae are present in all existing earth ecosystems, both aquatic and terrestrial, and can flourish under a wide range of environmental conditions, including freshwater, brackish water, seawater, and even wastewater [5]. Microalgae have been suggested as good candidates for fuel production because of their higher photosynthetic efficiency, higher biomass production and faster growth compared to those of other energy crops [6]. Microalgae systems also use far less water than do traditional oilseed crops. For these reasons, microalgae are capable of producing more oil per unit area of land compared to terrestrial oilseed crops [7]. According to some estimates, the yield (per acre) of oil from algae is over 200 times the yield from the bestperforming plant/vegetable oils [8]. Hundreds of microalgal strains capable of producing large quantities of lipids have been screened, and their lipid production metabolisms characterized and reported. Most of these organisms are marine microalgae [8]. Some species of algae produce large quantities of oil as a storage product, regularly achieving 50%e60% of their dry weights as lipid [7]. Biosynthesis of fatty acids can vary significantly according to the external environmental conditions [9] with temperature, composition in the culture medium and bubbling gas concentration being important in microalgal growth. Microalgae with high contents of fatty acids, neutral lipids, and polar lipids as well as a high growth rate in the natural environment have yet to be exploited for biodiesel production, and the isolation and characterization of microalgae with the potential for more efficient oil production remain the focus of continuing research [10]. In this study, the growth rate, algal biomass, and lipid content of some environmental microalgal isolates were determined. Furthermore, these naturally isolated microalgal species were subjected to fatty acid profile analysis.Moreover, the large subunit (LSU) ribosomal DNA (D1-D2)-encoding gene of the isolates was sequenced to confirm the identities of the microalgal species.
2.
Materials and methods
2.1. Isolation, purification and identification of microalgae Water samples used to isolate microalgae were collected aseptically from sites that appeared to contain algal growth in
a freshwater lake at Yonsei University, Wonju, South Korea. Bold basal medium (BBM) was used in this study [11]. The media was autoclaved at 1.2 atm for 15 min before use. Ten milliliters of water sample was inoculated into 200 mL media in a 500 mL conical flask, and then incubated on a rotary shaker at 27 C and 150 rpm under continuous illumination with white fluorescent light for three weeks. Every two days, the flasks were examined for algal growth using an optical microscope, and serial dilutions were prepared in BBM from flasks showing growth. Subcultures were made by inoculating 50 mL onto petri plates containing BBM solidified with 1.5% (w/v) of bacteriological agar. Further, 50 mL aliquots of the same dilution were placed into wells of a 96-well microtiter plate containing 200 mL BBM. These procedures were repeated for each of the original flasks. Both the Petri and microtiter plates were incubated at 27 C under continuous illumination using white fluorescent light for two weeks. The purities of the culture were ensured by repeated plating and regular observation under a microscope. Microscopic identification was performed [12] and was confirmed using molecular markers.
2.2. DNA extraction, PCR amplification, sequencing, and phylogenetic analysis An aliquot of cultured cells (1 mL) was harvested in the mid- to late exponential growth phase (10e14 days) by centrifugation (13,000 g for 3 min at 4 C) in a sterile microcentrifuge tube. Genomic DNA was extracted using a Plant Genomic DNA extraction kit (SolGent, Daejeon, S. Korea) according to the manufacturer’s instructions and protocols. The DNA concentration of the extracted DNA was measured at 260 nm using a spectrophotometer (HACH, DR/4000v, USA). To amplify the D1-D2 (LSU) coding region of the rDNA, amplification reactions were performed on a T-Gradient thermocycler (Biometra GmbH, Gottingen, Germany) using the universal eukaryotic primers 50 -AGCGGAGGAAAAGAAACTA’3 as forward and 50 -TACTAGAAGGTTCGATTAGTC-’3 as reverse, according to the PCR protocol described by Sonnenberg et al. [13]. Aliquots (10 mL) of the reaction mixtures were analyzed by 1% horizontal agarose gel electrophoresis to confirm the presence of product. The PCR products were purified using the Gel PCR Clean-Up System (Applied Biosystems, Foster, CA). Sequencing reactions were performed using a Dye Deoxy Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, Foster City, CA), and sequencing fragments were analyzed on a ABI Prism 377 DNA Sequencer. Ribosomal RNA gene sequences from the isolates were searched against GenBank using BLAST [14]. Aligned sequences were checked manually and were edited with Genedoc [15]. Sequences containing fewer than 200 nucleotides or in excess of 1000 nucleotides were removed, and sequences not belonging to green microalgal species were also discarded from our study. A phylogenetic tree was constructed using the neighbor-joining (NJ) algorithm using Kimura’s two-parameter model of sequence evolution, as implemented in the MEGA4 program package [16].
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2.3.
Microalgae cultivation and biomass
One hundred milliliters BBM in a 250 mL Erlenmeyer flask was inoculated with the cells (OD680 0.05) and incubated at 27 C with shaking at 150 rpm under continuous illumination for three weeks. Algal growth was monitored by measuring daily changes in optical density at 680 nm with a spectrophotometer. If the optical density of the undiluted sample was greater than 1.0, the sample was diluted to give an absorbance in the range of 0.1e1.0. Microalgae dry weight per liter (g L1) was measured according to a method previously reported [17]. Microalgal cells were harvested by centrifugation and washed twice with deionized water. Microalgal pellets were dried overnight at 105 C for dry weight measurement [18]. Experiments were carried out in triplicate, and data are expressed as mean SD.
2.4.
Extraction of total lipids
Total lipids were extracted from fresh microalgal biomass using a slightly modified method of Bligh and Dyer [19]. The lipids were extracted with chloroform-methanol (2:1, v/v) and then separated into chloroform and aqueous methanol layers by the addition of methanol and water to give a final solvent ratio of chloroform: methanol: water of 1:1:0.9. The chloroform layer was washed with 20 mL of a 5% NaCl solution, and evaporated to dryness. Thereafter, the weight of the crude lipid obtained from each sample was measured gravimetrically. Experiments were carried out in triplicate, and data are expressed as mean SD.
2.5.
Fatty acid composition analysis
The fatty acids were analyzed using the modified method of Lepage and Roy [20]. The crude lipid (w10 mg) was dissolved using 2 mL of a freshly prepared chloroform-methanol mixture (2: 1, v/v) and transferred into capped test tube. One mL of chloroform containing nonadecanoic acid (500 mg L1) as internal standard, 1 mL methanol, and 300 mL of sulfuric acid as transmethylation reagents were added to the tube, mixed for 5 min and then incubated at 100 C for 10 min. The fatty acid-containing phase was separated by adding 1 ml distilled water and was then recovered. The organic phase was filtered using a hypodermic 0.22 mm PVDF syringe filter (Millex-GV, Millipore, USA). Methyl esters of fatty acids were analyzed
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using a gas chromatograph (GC-7890, Agilent, USA) equipped with a flame ionization detector anda HP-INNO wax capillary column (Agilent Technologies, USA). The temperatures of injector and detector were set at 250 C and 275 C, respectively. Oven temperature conditions were maintained at 50 C for 1 min, 200 C for 12 min, and 250 C for 2 min. Mix RM3, Mix RM5, GLC50, and GLC70 (Supelco Co., USA), and a-linolenic acid (Sigma Chemical Co. USA) were used as standard materials. All reagents were of analytical grade. The components were identified by comparing their retention times and fragmentation patterns with those of the standards [21].
3.
Results and discussion
3.1.
Isolation and identification of microalgae
A total of 45 algal cultures were isolated from a freshwater lake at Yonsei University, Wonju, S. Korea. Out of 45 cultures, five green microalgal isolates (YSL02, YSL03, YSL05, YSL04 and YSL07) were selected based on their morphologies (i.e., cell shape and size) and because they could be successfully cultivated in pure form under our test conditions. Light microscopic images of the new species isolated in this study are shown in Fig. 1. Microscopic observation of algal isolates revealed their colonial existences and purities. Microscopic analysis of the samples allowed preliminary identification of isolates YSL02, YSL03, YSL04, YSL05, and YSL07 as genus Scenedesmus, Chlamydomonas, Chlorella, Scenedesmus, and Chlamydomonas, respectively. Komarek and Marvan [22] proposed the existence of at least 13 species of Botryococcus on the basis of morphological differences by omitting the chemical analyses. Metzger and Largeau [23] reported that for algae, within each chemical race and for the same strain, morphology could vary in relation to age and culture conditions. The morphological heterogeneity of algae makes microscopic identification difficult. Therefore, we isolated total DNA and PCR-amplified rRNA (LSU) to confirm our morphology-based species identifications.
3.1.1. LSU-rRNA (D1-D2) coding region amplification and sequencing PCR amplification of the genomic DNA of the algal isolates with the universal forward and reverse primers revealed efficient amplification. A single band of amplified LSU rDNA
Fig. 1 e Light microscope (40x) pictures of the tested microalgal isolates.
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(D1-D2) product with a size of w 850 bp was recorded for all isolates. The LSU rRNA gene has a higher evolutionary rate than does the SSU rRNA gene [13] and should therefore allow for better discrimination between closely-related species using short diagnostic sequences. Based on the LSU rDNA (D1D2) sequences, we concluded that microalgal isolates YSL02, YSL03, YSL04, YSL05 and YSL07 were closely related to Scenedesmus obliquus, Chlamydomonas pitschmannii, Chlorella vulgaris, S. obliquus and Chlamydomonas mexicana, based on 97%, 99%, 97%, 97% and 99% sequence similarities, respectively. The DNA sequences were published in the NCBI databases (accession numbers are provided in Table 1). The lengths of the LSU rDNA (D1-D2) regions of the five species of microalgae, their specific accession numbers and the nearest identifiable match present in the GenBank nucleotide database are shown in Table 1. Identification of the five microalgal strains was also supported by the results from the phylogenetic analysis of the LSU rDNA D1-D2 sequence. In the phylogram (Fig. 2), YSL02 and YSL05 which were identified as S. obliquus clearly group with the microalgal strain S. obliquus AF183482 (Table 1). The LSU rDNA sequences of isolates YSL03, YSL04, and YSL07 confirmed their identification as Chla. pitschmannii, C. vulgaris, and Chla. mexicana; they had sequence similarities of 99%, 97%, and 99% to Chla. pitschmannii AF183462, C. vulgaris AB237642, and Chla. mexicana AF395501, respectively (Fig. 2 and Table 1).
3.2.
Fig. 2 e Phylogenetic tree showing the relationships among LSU rDNA D1-D2 sequences of isolates YSL02, YSL03, YSL04, YSL05, and YSL07 and the most similar sequences retrieved from the NCBI nucleotide database.
Growth rates of the microalgal strains
Under suitable conditions and sufficient nutrients, microalgae can grow profusely. Their biomass usually doubles within 3.5e24 h during the exponential growth phase [1]. The net growth rates differed among the examined species under similar environmental conditions (Fig. 3). The average growth rates of C. vulgaris YSL04, Chla. mexicana YSL07, S. obliquus YSL05, S. obliquus YSL02, and Chla. pitschmannii YSL03, were 1.83 0.07, 1.65 0.04, 1.24 0.07, 1.07 0.17, and 0.61 0.11 days1, respectively. The growth rate of C. vulgaris YSL04 after 20 days of incubation was 3.29 0.20 compared with an initial reading of 0.69 0.19 at OD 680 nm. This result indicates that C. vulgaris YSL04 is suitable for high-density culture. Algal growth is directly affected by the availability of nutrients, light, the stability of pH, temperature and the initial inoculum density [24]. An increase in the initial inoculum density leads to better algal growth and increases the nutrient removal efficiency [25].
3.3.
Biomass, lipid content, and lipid productivity
The five microalgal species were tested for lipid production by evaluating biomass productivity and lipid content in 250 mL flask laboratory cultures under the same conditions after 21 days of incubation (Table 2). Biomass productivities (g dwt L1) of 1.84 0.30; 1.71 0.53; 1.65 0.07, and 1.53 0.30 were found for S. obliquus YSL02; S. obliquus YSL05; C. vulgaris YSL04 and Chla. mexicana YSL07, respectively. S. obliquus YSL02 showed the highest biomass productivity at 1.84 0.30 g dwt L1, while Chla. pitschmannii YSL03 had the lowest biomass productivity at 1.04 0.09 g dwt L1. Under our experimental growth conditions, the total lipid contents of the microalgae cultured in this study ranged from
Table 1 e The accession numbers, lengths in base pairs, similarities between amplified sequences, and the closest relative sequences for five strains of environmentally isolated microalgae. Microalgal strain S. obliquus YSL02 Chla. pitschmannii YSL03 C. vulgarisYSL04 S. obliquus YSL05 Chla. mexicana YSL07
Accession number
Length (nt)
GU732415 GU732416 GU732417 GU732418 GU732420
864 874 881 864 868
S ¼ Scenedesmus, Chla ¼ Chlamydomonas, C ¼ Chlorella and (nt) ¼ nucleotides.
Closest relative and GenBank accession number S. obliquus AF183482 Chla. pitschmannii AF183462 C. vulgarisAB237642 S. obliquus AF183482 Chla. mexicana AF395501
% Similarity 97 99 97 97 99
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Fig. 3 e Growth curves of five microalgal species cultivated in a batch experiment on a rotary shaker at 27 C and 150 rpm under continuous illumination for 21 days. Experiments were carried out in triplicate.
26% to 51% of their dry weight. The lipid content of Chla. pitschmannii YSL03 was 51% of its dry weight, which was about two-fold higher than that from C. vulgaris YSL04. S. obliquus YSL02, S. obliquus YSL05, and Chla. mexicana YSL07 had largely similar lipid contents (Table 2). The lipid productivity of Chla. pitschmannii YSL03 was the highest at 0.54 0.9 g L1 (Table 2). Lipid content data for different algal species are readily available in the literature [26]. Many microalgae species can be induced to accumulate substantial quantities of lipids [8], thus contributing to a high oil yield. In previous studies [27,28], a total lipid content of 20e50% of the dry biomass weight was quite common, and some microalgae had a total lipid content of 90% of the dry biomass in response to different culture conditions. However, previous studies have demonstrated that some Chlorella and Scenedesmus species can produce more lipids under certain conditions [29,30].
3.4.
Fatty acids in the five strains of microalgae (S. obliquus YSL02, Chla. pitschmannii YSL03, C. vulgaris YSL04, S. obliquus YSL05,
Table 2 e Biomass productivities, lipid contents and lipid productivities of five microalgal strains.
S. obliquus YSL02 Chla. pitschmannii YSL03 C. vulgaris YSL04 S. obliquus YSL05 Chla. mexicana YSL07
Table 3 e Fatty acid composition of the different microalgal species. Fatty acid
Fatty acid composition
Microalgal strain
and Chla. mexicana YSL07) were primarily esterified and the major fatty acid composition of each isolate was determined using GC analysis (Table 3). Biodiesel consists largely of fatty acid methyl esters, which are produced by the transesterification of biologically-derived lipids [31], and the quality of biodiesel is considerably affected by the composition of the fatty acids in the biodiesel. In a previous report [32], palmitic, stearic, oleic, linoleic and linolenic acid were recognized as the most common fatty acids in biodiesel. The fatty acids profiles of the isolates (Table 3), indicated the presences of lauric (C12:0), myristic (C14:0), palmitic (C16:0), heptadecanoic (C17:0), stearic (C18:0), palmitoleic (C16:1), oleic (C18:1n9c), alinolenic (C18:3n3), and g-linolenic acid (C18:3n6). The major fatty acids in the five isolates were palmitic, lauric, and stearic acids comprising 22%e50%, 5%e34% and 5%e23% of the total fatty acids, respectively, whereas palmitoleic, heptadecanoic, and a-linolenic acid existed as minor fatty acids. Oleic and palmitic acid comprised 53% and 50% of total fatty acids in C. vulgaris YSL04 and Chal. mexicana YSL07, respectively. Oleic acid, which is an ideal component of biodiesel, occupied up to 53% of the total fatty acids in C. vulgaris YSL04, which is a much higher proportion than the 24.8% reported by Ranga Rao et al. [33]. For the green alga Chlorella, the fatty acid compositions of 14:0, 16:0, 16:1, 16:2, 16:3, 18:0, 18:1, 18:2, a-183 have been confirmed under many conditions including photoautotrophic and heterotrophic cultivation, nitrogen starvation, and outdoors in a photobioreactor [34]. In particular, oils with high oleic acid content have been reported to have reasonable ignition quality, combustion heat, cold filter plugging point (CFPP), oxidative stability, viscosity, and lubricity. Biodiesel fuels enriched in methyl oleate are desirable, relatively small percentages of saturated fatty esters can wreck the cold flow properties of biodiesel. Our finding shows
Lipid Lipid content Biomass productivity productivity (% biomass) (g L1) (g dwt L1) 1.84 0.30 1.04 0.09
0.53 0.04 0.54 0.13
29 51
1.65 0.07 1.71 0.53 1.53 0.30
0.44 0.05 0.48 0.01 0.45 0.05
26 28 29
Experiments were carried out in triplicate and data are expressed as mean SD.
Fatty acid composition (wt %) S. Chla. C. S. Chla. obliquus pitschmannii vulgaris obliquus mexicana YSL02 YSL03 YSL04 YSL05 YSL07
Lauric acid (C12:0) Myristic acid (C14:0) Palmitic acid (C16:0) Palmitoleic acid (C16:1) Heptadecanoic acid (C17:0) Stearic acid (C18:0) Oleic acid (C18:1n9c) a-Linolenic acid (C18:3n3) g-Linolenic acid (C18:3n6) Total
11
10
5
7
34
0
8
2
2
10
29
26
22
25
50
0
0
5
4
0
0
0
0
2
0
17
20
5
23
6
20
13
53
8
0
0
0
0
4
0
23
23
8
25
0
100
100
100
100
100
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that most of these strains contain w 20% of saturated fatty acids (C16 and C18). Certainly this amount would negatively impact cold flow properties such as CFPP. Higher oleic acid content increases the oxidative stability of fuel, enabling longer storage [32], and decreases the CFPP of the fuel, allowing it to be used in cold regions [35]. Among the tested microalgal species, C. vulgaris RAI04 showed the highest oleic acid content, making it the most suitable isolate for the production of good quality biodiesel.
4.
Conclusions
Biomass productivity, lipid cell content, and overall lipid productivity are some of the key parameters that determine the economic feasibility of algal oil for biodiesel production. To select microalgae with a high biomass and lipid productivity, five microalgal cultures were selected based on their ease of cultivation and were identified as S. obliquus YSL02, Chla. pitschmannii YSL03, C. vulgaris YSL04, S. obliquus YSL05, and Chla. mexicana YSL07. Under similar environmental conditions, the average growth rates (OD680 nm) of C. vulgaris YSL04, Chla. mexicana YSL07, S. obliquus YSL05, S. obliquus YSL02, and Chla. pitschmannii YSL03 were 1.83 0.071, 1.65 0.04, 1.24 0.07, 1.07 0.17 and 0.61 0.11 days1, respectively. These results indicated that the C. vulgaris YSL04 strain was the most suitable of the five strains for high-density culture. The total lipid contents of the algae were 51%, 29%, 29%, 28%, and 26% for Chla. pitschmannii YSL03, Chla. mexicana YSL07, S. obliquus YSL02, S. obliquus YSL05, and C. vulgaris YSL04, respectively. The composition of fatty acids in the studied species was mainly C12:0, C16:0, C18:0, C18:1n9c, C18:3n6, C16:1, and C14:0. The results of this study indicate that the naturally isolated microalgal strain Chla. pitschmannii YSL03 may be a valuable candidate for biodiesel production.
Acknowledgements This work was supported by the Students’ Association of the Graduate School of Yonsei University and was funded by the Graduate School of Yonsei University, and Yonsei University research fund of 2009, 21st Frontier research project (Sustainable Water Resources Research Center 3-4-3), Global Research Laboratory project (Korea Institute of Geosciences and Mineral Resources NP2008-019) and the Brain Korea-21 (BK-21) program of the Ministry of Education, Korea.
references
[1] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25: 294e306. [2] Widjaja A, Chao-Chang Chien, Yi-Hsu Ju. Study of increasing lipid production from fresh water microalgae Chlorella vulgaris. J Taiwan Ins Chem Engin 2009;40:13e20. [3] Lang X, Dalai AK, Bakhshi NN, Reaney MJ, Hertz PB. Preparation and characterization of biodiesels from various Bio-Oils. Biores Technol 2002;80:53e62.
[4] Puppan D. Environmental evaluation of biofuels. Periodica Polytechnic Ser Soc Man Sci 2002;10:95e116. [5] Richmond A. Handbook of microalgal culture: Biotechnology and applied phycology. Blackwell Science Ltd; 2004. [6] Becker K. Measurement of algal growth. In: Microalgae biotechnology and microbiology. Cambridge University Press; 1994. p. 56e62. [7] Chisti Y. Biodiesel from microalgae beats bioethanol. Trends Biotechnol 2008;26:126e31. [8] Sheehan J, Dunahay T, Benemann J, Roessler P. A look back at the U.S. Department of Energy’s aquatic species program: biodiesel from algae. Close-Out report. Golden, Colorado, U.S. A: National Renewable Energy Lab, Department of Energy; 1998. Report number NREL/TP-580e24190. [9] Los DA, Murata N. Membrane fluidity and its roles in the perception of environmental signals. Biochim Biophys ActaBiomembranes 2004;1666:142e57. [10] Qiang H, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, et al. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J 2008; 54:621e39. [11] Kanz T, Bold HC. Publication No. In: Physiological studies, morphological and taxonomical investigation of nostoc and anabaena in culture. Austin, TX: University of Texas; 1969. p. 6924. [12] John DM, Whitton BA, Brook AJ. The freshwater algal flora of the British Isles an identification guide to freshwater and terrestrial algae. Cambridge: Cambridge University press; 2003. 39e43. [13] Sonnenberg R, Nolte AW, Tautz D. An evaluation of LSU rDNA D1-D2 sequences for their use in species identification. Front Zool 2007;4:1e12. [14] Altschul SF, Thomas LM, Alejandro AS, Jinghui Z, Zheng Z, Webb M, et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 1997;25:3389e402. [15] Nicholas KB, Nicholas HB. Alignment editor and shading utility. 2.6.001 Ed, http://www.PSC.edu/biomed/genedoc; 1997. [16] Tamura K, Dudley J. MECA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 2007;24:1596e9. [17] American Public Health Association. Methods for biomass production. In: Standard methods for the examination of water and wastewater. Baltimore, MD, USA: American Public Health Association; 1998. [18] Takagi M, Karseno S, Yoshida T. Effect of salt concentration on intracellular accumulation of lipids and triacylglyceride in marine microalgae Dunaliella cells. J Biosci Bioeng 2006;101:223e6. [19] Bligh EG, Dyer WJ. A rapid method of total lipid extraction and purification. Can J Biochem Physiol 1959;37:911e7. [20] Lepage G, Roy CC. Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J Lipid Res 1984;25:1391e6. [21] Xu N, Zhang X, Fan X, Han L, Zeng C. Effects of nitrogen source and concentration on growth rate and fatty acid composition of Ellipsoidion sp. (Eustigmatophyta). J Appl Phycol 2001;13:463e9. [22] Komarek J, Marvan P. Morphological differences in natural populations of the genus Botryococcus (chlorophyceae). Archiv Fur Protistenkunde 1992;141:65e100. [23] Metzger P, Largeau C. Botryococcus braunii: a rich source for hydrocarbons and related ether lipids. Appl Microbiol Biotechnol 2005;66:486e96. [24] Wang L, Yecong L, Chen P, Min M, Chen Y, Zhu J, et al. Anaerobic digested dairy manure as a nutrient supplement for cultivation of oil-rich green microalgae Chlorella sp. Biores Technol 2010;101:2623e8. [25] Lau PS, Tam NFY, Wong YS. Effect of algal density on nutrient removal from primary settled wastewater. Environ Pollut 1995;89:59e66.
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[26] Griffiths MJ, Harrison STL. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. J Appl Phycol 2009;21:493e507. [27] Spolaore P, Joannis-Cassan C, Duran E, Isambert A. Commercial applications of microalgae. J Biosci Bioeng 2006;101:87e96. [28] Li Y, Horsman M, Wu N, Lan CQ, Dubois-Calero N. Biofuels from microalgae. Biotechnol Prog 2008;24:815e20. [29] Illman AM, Scragg AH, Shales SW. Increase in Chlorella strains calorific values when grown in low nitrogen medium. Enz Microb Technol 2000;27:631e5. [30] Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonin G, et al. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol Bioeng 2009;102:100e12.
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[31] Wackett LP. Biomass to fuels via microbial transformations. Curr Opin Chem Biol 2008;12:187e93. [32] Knothe G. “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energy Fuels 2008; 22:1358e64. [33] Ranga Rao A, Sarada TR, Ravishankar GA. Influence of CO2 on growth and hydrocarbon production in Botryococcus braunii. J Microbiol Biotechnol 2007;17: 414e9. [34] Petkov G, Garcia G. Which are fatty acids of the green algae Chlorella. Biochem Syst Ecol 2007;35:281e5. [35] Stournas S, Lois E, Serdari A. Effects of fatty acid derivatives on the ignition quality and cold flow of diesel fuel. J Am Oil Chem Soc 1995;72:433e7.
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Characterization and identification of lipid-producing microalgae species isolated from a freshwater lake Reda A.I. Abou-Shanab a,b, Ibrahim A. Matter a, Su-Nam Kim c, You-Kwan Oh d, Jaeyoung Choi c, Byong-Hun Jeon a,* a
Department of Environmental Engineering, Yonsei University, Wonju, Gangwon-do 220-710, South Korea Department of Environmental Biotechnology, Mubarak City for Scientific Research and Technology Applications, New Borg El Arab City, Alexandria 21934, Egypt c KIST Gangneung Institute, Gangneung 210-340, South Korea d Bioenergy Center, Korea Institute of Energy Research, Daejeon 305-343, South Korea b
article info
abstract
Article history:
Microalgal lipids are the oils of the future for sustainable biodiesel production. One of the
Received 31 March 2010
most important decisions in obtaining oil from microalgae is the choice of species. A total
Received in revised form
of 45 algal cultures were isolated from a freshwater lake at Wonju, South Korea. Five
10 April 2011
microalgal isolates were selected based on their morphology and ease of cultivation
Accepted 15 April 2011
under our test conditions. These cultures were identified as strains of Scenedesmus obli-
Available online 5 May 2011
quus YSL02, Chlamydomonas pitschmannii YSL03, Chlorella vulgaris YSL04, S. obliquus YSL05, and Chlamydomonas mexicana YSL07 based on microscopic examination and LSU rDNA
Keywords:
(D1-D2) sequence analysis. S. obliquus YSL02 reached a higher biomass concentration
Biodiesel
(1.84 0.30 g L1) with a lower lipid content (29% w/w), than did Chla. pitschmannii YSL03
Freshwater
(maximum biomass concentration of 1.04 0.09 with a 51% lipid content). Our results
Isolation
suggest that Chla. pitschmannii YSL03 is appropriate for producing biodiesel based on its
LSU rDNA (D1-D2)
high lipid content and oleic acid proportion. ª 2011 Elsevier Ltd. All rights reserved.
Microalgae
1.
Introduction
The basic resources currently exploited to obtain energy are petroleum, natural gas, coal, hydropower, and nuclear power. Continued use of petroleum-based fuels is now widely recognized as unsustainable because of limited supplies and the contribution of these fuels to atmospheric pollution. Fossil fuel combustion is also a major source of greenhouse gases responsible for global warming. Renewable, carbon-neutral, economically viable alternatives to fossil fuels are urgently needed to avert the impending oil crisis and the dramatic consequences of climate change [1].
Biomass is one of the better sources of energy to mitigate greenhouse gas emissions and to function as a substitute for fossil fuels [2]. Large-scale introduction of biomass energy could contribute to sustainable development on environmental, social, and economic fronts. Biodiesel (monoalkyl esters) is one such alternative fuel, obtained by the transesterification of triglyceride oil with monohydric alcohols. Commercial biodiesel has been obtained successfully from rapeseed, soybean, sunflower, corn, palm, and waste cooking oil, as well as from animal fat [3]. However, large-scale production of biodiesel from those resources cannot realistically satisfy the existing demand for transport fuels [1]. Biodiesel has received considerable
* Corresponding author. Tel.: þ82 33 760 2446; fax: þ82 33 760 2571. E-mail address: [email protected] (B.-H. Jeon). 0961-9534/$ e see front matter ª 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.biombioe.2011.04.021
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attention in recent years, as it is a biodegradable, renewable, and non-toxic fuel. It contributes no net carbon dioxide or sulfur to the atmosphere and emits less gaseous pollutants than does normal diesel [3]. While biodiesel is a desirable product, the significant economic and environmental impacts of using agricultural crops, especially food crops, as a feedstock for biofuels has raised crucial sustainability issues. For example, the transformation of primary food resources into bio-fuels has led to a clash of interests, as using food crops to derived biodiesel has resulted in a reduced supply to poor countries and an increase in food costs [4]. All of these factors have stimulated the search for other sources of biodiesel production that are both sustainable and economical. Microalgae are microscopic heterotrophiceautotrophic photosynthesizing organisms that are able to use solar energy to combine water with carbon dioxide to create biomass. Microalgae are present in all existing earth ecosystems, both aquatic and terrestrial, and can flourish under a wide range of environmental conditions, including freshwater, brackish water, seawater, and even wastewater [5]. Microalgae have been suggested as good candidates for fuel production because of their higher photosynthetic efficiency, higher biomass production and faster growth compared to those of other energy crops [6]. Microalgae systems also use far less water than do traditional oilseed crops. For these reasons, microalgae are capable of producing more oil per unit area of land compared to terrestrial oilseed crops [7]. According to some estimates, the yield (per acre) of oil from algae is over 200 times the yield from the bestperforming plant/vegetable oils [8]. Hundreds of microalgal strains capable of producing large quantities of lipids have been screened, and their lipid production metabolisms characterized and reported. Most of these organisms are marine microalgae [8]. Some species of algae produce large quantities of oil as a storage product, regularly achieving 50%e60% of their dry weights as lipid [7]. Biosynthesis of fatty acids can vary significantly according to the external environmental conditions [9] with temperature, composition in the culture medium and bubbling gas concentration being important in microalgal growth. Microalgae with high contents of fatty acids, neutral lipids, and polar lipids as well as a high growth rate in the natural environment have yet to be exploited for biodiesel production, and the isolation and characterization of microalgae with the potential for more efficient oil production remain the focus of continuing research [10]. In this study, the growth rate, algal biomass, and lipid content of some environmental microalgal isolates were determined. Furthermore, these naturally isolated microalgal species were subjected to fatty acid profile analysis.Moreover, the large subunit (LSU) ribosomal DNA (D1-D2)-encoding gene of the isolates was sequenced to confirm the identities of the microalgal species.
2.
Materials and methods
2.1. Isolation, purification and identification of microalgae Water samples used to isolate microalgae were collected aseptically from sites that appeared to contain algal growth in
a freshwater lake at Yonsei University, Wonju, South Korea. Bold basal medium (BBM) was used in this study [11]. The media was autoclaved at 1.2 atm for 15 min before use. Ten milliliters of water sample was inoculated into 200 mL media in a 500 mL conical flask, and then incubated on a rotary shaker at 27 C and 150 rpm under continuous illumination with white fluorescent light for three weeks. Every two days, the flasks were examined for algal growth using an optical microscope, and serial dilutions were prepared in BBM from flasks showing growth. Subcultures were made by inoculating 50 mL onto petri plates containing BBM solidified with 1.5% (w/v) of bacteriological agar. Further, 50 mL aliquots of the same dilution were placed into wells of a 96-well microtiter plate containing 200 mL BBM. These procedures were repeated for each of the original flasks. Both the Petri and microtiter plates were incubated at 27 C under continuous illumination using white fluorescent light for two weeks. The purities of the culture were ensured by repeated plating and regular observation under a microscope. Microscopic identification was performed [12] and was confirmed using molecular markers.
2.2. DNA extraction, PCR amplification, sequencing, and phylogenetic analysis An aliquot of cultured cells (1 mL) was harvested in the mid- to late exponential growth phase (10e14 days) by centrifugation (13,000 g for 3 min at 4 C) in a sterile microcentrifuge tube. Genomic DNA was extracted using a Plant Genomic DNA extraction kit (SolGent, Daejeon, S. Korea) according to the manufacturer’s instructions and protocols. The DNA concentration of the extracted DNA was measured at 260 nm using a spectrophotometer (HACH, DR/4000v, USA). To amplify the D1-D2 (LSU) coding region of the rDNA, amplification reactions were performed on a T-Gradient thermocycler (Biometra GmbH, Gottingen, Germany) using the universal eukaryotic primers 50 -AGCGGAGGAAAAGAAACTA’3 as forward and 50 -TACTAGAAGGTTCGATTAGTC-’3 as reverse, according to the PCR protocol described by Sonnenberg et al. [13]. Aliquots (10 mL) of the reaction mixtures were analyzed by 1% horizontal agarose gel electrophoresis to confirm the presence of product. The PCR products were purified using the Gel PCR Clean-Up System (Applied Biosystems, Foster, CA). Sequencing reactions were performed using a Dye Deoxy Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems, Foster City, CA), and sequencing fragments were analyzed on a ABI Prism 377 DNA Sequencer. Ribosomal RNA gene sequences from the isolates were searched against GenBank using BLAST [14]. Aligned sequences were checked manually and were edited with Genedoc [15]. Sequences containing fewer than 200 nucleotides or in excess of 1000 nucleotides were removed, and sequences not belonging to green microalgal species were also discarded from our study. A phylogenetic tree was constructed using the neighbor-joining (NJ) algorithm using Kimura’s two-parameter model of sequence evolution, as implemented in the MEGA4 program package [16].
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2.3.
Microalgae cultivation and biomass
One hundred milliliters BBM in a 250 mL Erlenmeyer flask was inoculated with the cells (OD680 0.05) and incubated at 27 C with shaking at 150 rpm under continuous illumination for three weeks. Algal growth was monitored by measuring daily changes in optical density at 680 nm with a spectrophotometer. If the optical density of the undiluted sample was greater than 1.0, the sample was diluted to give an absorbance in the range of 0.1e1.0. Microalgae dry weight per liter (g L1) was measured according to a method previously reported [17]. Microalgal cells were harvested by centrifugation and washed twice with deionized water. Microalgal pellets were dried overnight at 105 C for dry weight measurement [18]. Experiments were carried out in triplicate, and data are expressed as mean SD.
2.4.
Extraction of total lipids
Total lipids were extracted from fresh microalgal biomass using a slightly modified method of Bligh and Dyer [19]. The lipids were extracted with chloroform-methanol (2:1, v/v) and then separated into chloroform and aqueous methanol layers by the addition of methanol and water to give a final solvent ratio of chloroform: methanol: water of 1:1:0.9. The chloroform layer was washed with 20 mL of a 5% NaCl solution, and evaporated to dryness. Thereafter, the weight of the crude lipid obtained from each sample was measured gravimetrically. Experiments were carried out in triplicate, and data are expressed as mean SD.
2.5.
Fatty acid composition analysis
The fatty acids were analyzed using the modified method of Lepage and Roy [20]. The crude lipid (w10 mg) was dissolved using 2 mL of a freshly prepared chloroform-methanol mixture (2: 1, v/v) and transferred into capped test tube. One mL of chloroform containing nonadecanoic acid (500 mg L1) as internal standard, 1 mL methanol, and 300 mL of sulfuric acid as transmethylation reagents were added to the tube, mixed for 5 min and then incubated at 100 C for 10 min. The fatty acid-containing phase was separated by adding 1 ml distilled water and was then recovered. The organic phase was filtered using a hypodermic 0.22 mm PVDF syringe filter (Millex-GV, Millipore, USA). Methyl esters of fatty acids were analyzed
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using a gas chromatograph (GC-7890, Agilent, USA) equipped with a flame ionization detector anda HP-INNO wax capillary column (Agilent Technologies, USA). The temperatures of injector and detector were set at 250 C and 275 C, respectively. Oven temperature conditions were maintained at 50 C for 1 min, 200 C for 12 min, and 250 C for 2 min. Mix RM3, Mix RM5, GLC50, and GLC70 (Supelco Co., USA), and a-linolenic acid (Sigma Chemical Co. USA) were used as standard materials. All reagents were of analytical grade. The components were identified by comparing their retention times and fragmentation patterns with those of the standards [21].
3.
Results and discussion
3.1.
Isolation and identification of microalgae
A total of 45 algal cultures were isolated from a freshwater lake at Yonsei University, Wonju, S. Korea. Out of 45 cultures, five green microalgal isolates (YSL02, YSL03, YSL05, YSL04 and YSL07) were selected based on their morphologies (i.e., cell shape and size) and because they could be successfully cultivated in pure form under our test conditions. Light microscopic images of the new species isolated in this study are shown in Fig. 1. Microscopic observation of algal isolates revealed their colonial existences and purities. Microscopic analysis of the samples allowed preliminary identification of isolates YSL02, YSL03, YSL04, YSL05, and YSL07 as genus Scenedesmus, Chlamydomonas, Chlorella, Scenedesmus, and Chlamydomonas, respectively. Komarek and Marvan [22] proposed the existence of at least 13 species of Botryococcus on the basis of morphological differences by omitting the chemical analyses. Metzger and Largeau [23] reported that for algae, within each chemical race and for the same strain, morphology could vary in relation to age and culture conditions. The morphological heterogeneity of algae makes microscopic identification difficult. Therefore, we isolated total DNA and PCR-amplified rRNA (LSU) to confirm our morphology-based species identifications.
3.1.1. LSU-rRNA (D1-D2) coding region amplification and sequencing PCR amplification of the genomic DNA of the algal isolates with the universal forward and reverse primers revealed efficient amplification. A single band of amplified LSU rDNA
Fig. 1 e Light microscope (40x) pictures of the tested microalgal isolates.
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(D1-D2) product with a size of w 850 bp was recorded for all isolates. The LSU rRNA gene has a higher evolutionary rate than does the SSU rRNA gene [13] and should therefore allow for better discrimination between closely-related species using short diagnostic sequences. Based on the LSU rDNA (D1D2) sequences, we concluded that microalgal isolates YSL02, YSL03, YSL04, YSL05 and YSL07 were closely related to Scenedesmus obliquus, Chlamydomonas pitschmannii, Chlorella vulgaris, S. obliquus and Chlamydomonas mexicana, based on 97%, 99%, 97%, 97% and 99% sequence similarities, respectively. The DNA sequences were published in the NCBI databases (accession numbers are provided in Table 1). The lengths of the LSU rDNA (D1-D2) regions of the five species of microalgae, their specific accession numbers and the nearest identifiable match present in the GenBank nucleotide database are shown in Table 1. Identification of the five microalgal strains was also supported by the results from the phylogenetic analysis of the LSU rDNA D1-D2 sequence. In the phylogram (Fig. 2), YSL02 and YSL05 which were identified as S. obliquus clearly group with the microalgal strain S. obliquus AF183482 (Table 1). The LSU rDNA sequences of isolates YSL03, YSL04, and YSL07 confirmed their identification as Chla. pitschmannii, C. vulgaris, and Chla. mexicana; they had sequence similarities of 99%, 97%, and 99% to Chla. pitschmannii AF183462, C. vulgaris AB237642, and Chla. mexicana AF395501, respectively (Fig. 2 and Table 1).
3.2.
Fig. 2 e Phylogenetic tree showing the relationships among LSU rDNA D1-D2 sequences of isolates YSL02, YSL03, YSL04, YSL05, and YSL07 and the most similar sequences retrieved from the NCBI nucleotide database.
Growth rates of the microalgal strains
Under suitable conditions and sufficient nutrients, microalgae can grow profusely. Their biomass usually doubles within 3.5e24 h during the exponential growth phase [1]. The net growth rates differed among the examined species under similar environmental conditions (Fig. 3). The average growth rates of C. vulgaris YSL04, Chla. mexicana YSL07, S. obliquus YSL05, S. obliquus YSL02, and Chla. pitschmannii YSL03, were 1.83 0.07, 1.65 0.04, 1.24 0.07, 1.07 0.17, and 0.61 0.11 days1, respectively. The growth rate of C. vulgaris YSL04 after 20 days of incubation was 3.29 0.20 compared with an initial reading of 0.69 0.19 at OD 680 nm. This result indicates that C. vulgaris YSL04 is suitable for high-density culture. Algal growth is directly affected by the availability of nutrients, light, the stability of pH, temperature and the initial inoculum density [24]. An increase in the initial inoculum density leads to better algal growth and increases the nutrient removal efficiency [25].
3.3.
Biomass, lipid content, and lipid productivity
The five microalgal species were tested for lipid production by evaluating biomass productivity and lipid content in 250 mL flask laboratory cultures under the same conditions after 21 days of incubation (Table 2). Biomass productivities (g dwt L1) of 1.84 0.30; 1.71 0.53; 1.65 0.07, and 1.53 0.30 were found for S. obliquus YSL02; S. obliquus YSL05; C. vulgaris YSL04 and Chla. mexicana YSL07, respectively. S. obliquus YSL02 showed the highest biomass productivity at 1.84 0.30 g dwt L1, while Chla. pitschmannii YSL03 had the lowest biomass productivity at 1.04 0.09 g dwt L1. Under our experimental growth conditions, the total lipid contents of the microalgae cultured in this study ranged from
Table 1 e The accession numbers, lengths in base pairs, similarities between amplified sequences, and the closest relative sequences for five strains of environmentally isolated microalgae. Microalgal strain S. obliquus YSL02 Chla. pitschmannii YSL03 C. vulgarisYSL04 S. obliquus YSL05 Chla. mexicana YSL07
Accession number
Length (nt)
GU732415 GU732416 GU732417 GU732418 GU732420
864 874 881 864 868
S ¼ Scenedesmus, Chla ¼ Chlamydomonas, C ¼ Chlorella and (nt) ¼ nucleotides.
Closest relative and GenBank accession number S. obliquus AF183482 Chla. pitschmannii AF183462 C. vulgarisAB237642 S. obliquus AF183482 Chla. mexicana AF395501
% Similarity 97 99 97 97 99
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Fig. 3 e Growth curves of five microalgal species cultivated in a batch experiment on a rotary shaker at 27 C and 150 rpm under continuous illumination for 21 days. Experiments were carried out in triplicate.
26% to 51% of their dry weight. The lipid content of Chla. pitschmannii YSL03 was 51% of its dry weight, which was about two-fold higher than that from C. vulgaris YSL04. S. obliquus YSL02, S. obliquus YSL05, and Chla. mexicana YSL07 had largely similar lipid contents (Table 2). The lipid productivity of Chla. pitschmannii YSL03 was the highest at 0.54 0.9 g L1 (Table 2). Lipid content data for different algal species are readily available in the literature [26]. Many microalgae species can be induced to accumulate substantial quantities of lipids [8], thus contributing to a high oil yield. In previous studies [27,28], a total lipid content of 20e50% of the dry biomass weight was quite common, and some microalgae had a total lipid content of 90% of the dry biomass in response to different culture conditions. However, previous studies have demonstrated that some Chlorella and Scenedesmus species can produce more lipids under certain conditions [29,30].
3.4.
Fatty acids in the five strains of microalgae (S. obliquus YSL02, Chla. pitschmannii YSL03, C. vulgaris YSL04, S. obliquus YSL05,
Table 2 e Biomass productivities, lipid contents and lipid productivities of five microalgal strains.
S. obliquus YSL02 Chla. pitschmannii YSL03 C. vulgaris YSL04 S. obliquus YSL05 Chla. mexicana YSL07
Table 3 e Fatty acid composition of the different microalgal species. Fatty acid
Fatty acid composition
Microalgal strain
and Chla. mexicana YSL07) were primarily esterified and the major fatty acid composition of each isolate was determined using GC analysis (Table 3). Biodiesel consists largely of fatty acid methyl esters, which are produced by the transesterification of biologically-derived lipids [31], and the quality of biodiesel is considerably affected by the composition of the fatty acids in the biodiesel. In a previous report [32], palmitic, stearic, oleic, linoleic and linolenic acid were recognized as the most common fatty acids in biodiesel. The fatty acids profiles of the isolates (Table 3), indicated the presences of lauric (C12:0), myristic (C14:0), palmitic (C16:0), heptadecanoic (C17:0), stearic (C18:0), palmitoleic (C16:1), oleic (C18:1n9c), alinolenic (C18:3n3), and g-linolenic acid (C18:3n6). The major fatty acids in the five isolates were palmitic, lauric, and stearic acids comprising 22%e50%, 5%e34% and 5%e23% of the total fatty acids, respectively, whereas palmitoleic, heptadecanoic, and a-linolenic acid existed as minor fatty acids. Oleic and palmitic acid comprised 53% and 50% of total fatty acids in C. vulgaris YSL04 and Chal. mexicana YSL07, respectively. Oleic acid, which is an ideal component of biodiesel, occupied up to 53% of the total fatty acids in C. vulgaris YSL04, which is a much higher proportion than the 24.8% reported by Ranga Rao et al. [33]. For the green alga Chlorella, the fatty acid compositions of 14:0, 16:0, 16:1, 16:2, 16:3, 18:0, 18:1, 18:2, a-183 have been confirmed under many conditions including photoautotrophic and heterotrophic cultivation, nitrogen starvation, and outdoors in a photobioreactor [34]. In particular, oils with high oleic acid content have been reported to have reasonable ignition quality, combustion heat, cold filter plugging point (CFPP), oxidative stability, viscosity, and lubricity. Biodiesel fuels enriched in methyl oleate are desirable, relatively small percentages of saturated fatty esters can wreck the cold flow properties of biodiesel. Our finding shows
Lipid Lipid content Biomass productivity productivity (% biomass) (g L1) (g dwt L1) 1.84 0.30 1.04 0.09
0.53 0.04 0.54 0.13
29 51
1.65 0.07 1.71 0.53 1.53 0.30
0.44 0.05 0.48 0.01 0.45 0.05
26 28 29
Experiments were carried out in triplicate and data are expressed as mean SD.
Fatty acid composition (wt %) S. Chla. C. S. Chla. obliquus pitschmannii vulgaris obliquus mexicana YSL02 YSL03 YSL04 YSL05 YSL07
Lauric acid (C12:0) Myristic acid (C14:0) Palmitic acid (C16:0) Palmitoleic acid (C16:1) Heptadecanoic acid (C17:0) Stearic acid (C18:0) Oleic acid (C18:1n9c) a-Linolenic acid (C18:3n3) g-Linolenic acid (C18:3n6) Total
11
10
5
7
34
0
8
2
2
10
29
26
22
25
50
0
0
5
4
0
0
0
0
2
0
17
20
5
23
6
20
13
53
8
0
0
0
0
4
0
23
23
8
25
0
100
100
100
100
100
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that most of these strains contain w 20% of saturated fatty acids (C16 and C18). Certainly this amount would negatively impact cold flow properties such as CFPP. Higher oleic acid content increases the oxidative stability of fuel, enabling longer storage [32], and decreases the CFPP of the fuel, allowing it to be used in cold regions [35]. Among the tested microalgal species, C. vulgaris RAI04 showed the highest oleic acid content, making it the most suitable isolate for the production of good quality biodiesel.
4.
Conclusions
Biomass productivity, lipid cell content, and overall lipid productivity are some of the key parameters that determine the economic feasibility of algal oil for biodiesel production. To select microalgae with a high biomass and lipid productivity, five microalgal cultures were selected based on their ease of cultivation and were identified as S. obliquus YSL02, Chla. pitschmannii YSL03, C. vulgaris YSL04, S. obliquus YSL05, and Chla. mexicana YSL07. Under similar environmental conditions, the average growth rates (OD680 nm) of C. vulgaris YSL04, Chla. mexicana YSL07, S. obliquus YSL05, S. obliquus YSL02, and Chla. pitschmannii YSL03 were 1.83 0.071, 1.65 0.04, 1.24 0.07, 1.07 0.17 and 0.61 0.11 days1, respectively. These results indicated that the C. vulgaris YSL04 strain was the most suitable of the five strains for high-density culture. The total lipid contents of the algae were 51%, 29%, 29%, 28%, and 26% for Chla. pitschmannii YSL03, Chla. mexicana YSL07, S. obliquus YSL02, S. obliquus YSL05, and C. vulgaris YSL04, respectively. The composition of fatty acids in the studied species was mainly C12:0, C16:0, C18:0, C18:1n9c, C18:3n6, C16:1, and C14:0. The results of this study indicate that the naturally isolated microalgal strain Chla. pitschmannii YSL03 may be a valuable candidate for biodiesel production.
Acknowledgements This work was supported by the Students’ Association of the Graduate School of Yonsei University and was funded by the Graduate School of Yonsei University, and Yonsei University research fund of 2009, 21st Frontier research project (Sustainable Water Resources Research Center 3-4-3), Global Research Laboratory project (Korea Institute of Geosciences and Mineral Resources NP2008-019) and the Brain Korea-21 (BK-21) program of the Ministry of Education, Korea.
references
[1] Chisti Y. Biodiesel from microalgae. Biotechnol Adv 2007;25: 294e306. [2] Widjaja A, Chao-Chang Chien, Yi-Hsu Ju. Study of increasing lipid production from fresh water microalgae Chlorella vulgaris. J Taiwan Ins Chem Engin 2009;40:13e20. [3] Lang X, Dalai AK, Bakhshi NN, Reaney MJ, Hertz PB. Preparation and characterization of biodiesels from various Bio-Oils. Biores Technol 2002;80:53e62.
[4] Puppan D. Environmental evaluation of biofuels. Periodica Polytechnic Ser Soc Man Sci 2002;10:95e116. [5] Richmond A. Handbook of microalgal culture: Biotechnology and applied phycology. Blackwell Science Ltd; 2004. [6] Becker K. Measurement of algal growth. In: Microalgae biotechnology and microbiology. Cambridge University Press; 1994. p. 56e62. [7] Chisti Y. Biodiesel from microalgae beats bioethanol. Trends Biotechnol 2008;26:126e31. [8] Sheehan J, Dunahay T, Benemann J, Roessler P. A look back at the U.S. Department of Energy’s aquatic species program: biodiesel from algae. Close-Out report. Golden, Colorado, U.S. A: National Renewable Energy Lab, Department of Energy; 1998. Report number NREL/TP-580e24190. [9] Los DA, Murata N. Membrane fluidity and its roles in the perception of environmental signals. Biochim Biophys ActaBiomembranes 2004;1666:142e57. [10] Qiang H, Sommerfeld M, Jarvis E, Ghirardi M, Posewitz M, Seibert M, et al. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J 2008; 54:621e39. [11] Kanz T, Bold HC. Publication No. In: Physiological studies, morphological and taxonomical investigation of nostoc and anabaena in culture. Austin, TX: University of Texas; 1969. p. 6924. [12] John DM, Whitton BA, Brook AJ. The freshwater algal flora of the British Isles an identification guide to freshwater and terrestrial algae. Cambridge: Cambridge University press; 2003. 39e43. [13] Sonnenberg R, Nolte AW, Tautz D. An evaluation of LSU rDNA D1-D2 sequences for their use in species identification. Front Zool 2007;4:1e12. [14] Altschul SF, Thomas LM, Alejandro AS, Jinghui Z, Zheng Z, Webb M, et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 1997;25:3389e402. [15] Nicholas KB, Nicholas HB. Alignment editor and shading utility. 2.6.001 Ed, http://www.PSC.edu/biomed/genedoc; 1997. [16] Tamura K, Dudley J. MECA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol Biol Evol 2007;24:1596e9. [17] American Public Health Association. Methods for biomass production. In: Standard methods for the examination of water and wastewater. Baltimore, MD, USA: American Public Health Association; 1998. [18] Takagi M, Karseno S, Yoshida T. Effect of salt concentration on intracellular accumulation of lipids and triacylglyceride in marine microalgae Dunaliella cells. J Biosci Bioeng 2006;101:223e6. [19] Bligh EG, Dyer WJ. A rapid method of total lipid extraction and purification. Can J Biochem Physiol 1959;37:911e7. [20] Lepage G, Roy CC. Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J Lipid Res 1984;25:1391e6. [21] Xu N, Zhang X, Fan X, Han L, Zeng C. Effects of nitrogen source and concentration on growth rate and fatty acid composition of Ellipsoidion sp. (Eustigmatophyta). J Appl Phycol 2001;13:463e9. [22] Komarek J, Marvan P. Morphological differences in natural populations of the genus Botryococcus (chlorophyceae). Archiv Fur Protistenkunde 1992;141:65e100. [23] Metzger P, Largeau C. Botryococcus braunii: a rich source for hydrocarbons and related ether lipids. Appl Microbiol Biotechnol 2005;66:486e96. [24] Wang L, Yecong L, Chen P, Min M, Chen Y, Zhu J, et al. Anaerobic digested dairy manure as a nutrient supplement for cultivation of oil-rich green microalgae Chlorella sp. Biores Technol 2010;101:2623e8. [25] Lau PS, Tam NFY, Wong YS. Effect of algal density on nutrient removal from primary settled wastewater. Environ Pollut 1995;89:59e66.
b i o m a s s a n d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9 e3 0 8 5
[26] Griffiths MJ, Harrison STL. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. J Appl Phycol 2009;21:493e507. [27] Spolaore P, Joannis-Cassan C, Duran E, Isambert A. Commercial applications of microalgae. J Biosci Bioeng 2006;101:87e96. [28] Li Y, Horsman M, Wu N, Lan CQ, Dubois-Calero N. Biofuels from microalgae. Biotechnol Prog 2008;24:815e20. [29] Illman AM, Scragg AH, Shales SW. Increase in Chlorella strains calorific values when grown in low nitrogen medium. Enz Microb Technol 2000;27:631e5. [30] Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonin G, et al. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol Bioeng 2009;102:100e12.
3085
[31] Wackett LP. Biomass to fuels via microbial transformations. Curr Opin Chem Biol 2008;12:187e93. [32] Knothe G. “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energy Fuels 2008; 22:1358e64. [33] Ranga Rao A, Sarada TR, Ravishankar GA. Influence of CO2 on growth and hydrocarbon production in Botryococcus braunii. J Microbiol Biotechnol 2007;17: 414e9. [34] Petkov G, Garcia G. Which are fatty acids of the green algae Chlorella. Biochem Syst Ecol 2007;35:281e5. [35] Stournas S, Lois E, Serdari A. Effects of fatty acid derivatives on the ignition quality and cold flow of diesel fuel. J Am Oil Chem Soc 1995;72:433e7.