Characterization of microalgal species isolated from fresh water bodies as a potential source for biodiesel production

b i om a s s a n d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 5

Avai lab le a t www.sc iencedi rec t .com

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Characterization and identification of lipid-producingmicroalgae 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,*aDepartment of Environmental Engineering, Yonsei University, Wonju, Gangwon-do 220-710, South KoreabDepartment of Environmental Biotechnology, Mubarak City for Scientific Research and Technology Applications, New Borg El Arab City,

Alexandria 21934, EgyptcKIST Gangneung Institute, Gangneung 210-340, South KoreadBioenergy Center, Korea Institute of Energy Research, Daejeon 305-343, South Korea

a r t i c l e i n f o

Article history:

Received 31 March 2010

Received in revised form

10 April 2011

Accepted 15 April 2011

Available online 5 May 2011

Keywords:

Biodiesel

Freshwater

Isolation

LSU rDNA (D1-D2)

Microalgae

* Corresponding author. Tel.: þ82 33 760 244E-mail address: [email protected] (B.-H

0961-9534/$ e see front matter ª 2011 Elsevdoi:10.1016/j.biombioe.2011.04.021

a b s t r a c t

Microalgal lipids are the oils of the future for sustainable biodiesel production. One of the

most important decisions in obtaining oil from microalgae is the choice of species. A total

of 45 algal cultures were isolated from a freshwater lake at Wonju, South Korea. Five

microalgal isolates were selected based on their morphology and ease of cultivation

under our test conditions. These cultures were identified as strains of Scenedesmus obli-

quus YSL02, Chlamydomonas pitschmannii YSL03, Chlorella vulgaris YSL04, S. obliquus YSL05,

and Chlamydomonas mexicana YSL07 based on microscopic examination and LSU rDNA

(D1-D2) sequence analysis. S. obliquus YSL02 reached a higher biomass concentration

(1.84 � 0.30 g L�1) with a lower lipid content (29% w/w), than did Chla. pitschmannii YSL03

(maximum biomass concentration of 1.04 � 0.09 with a 51% lipid content). Our results

suggest that Chla. pitschmannii YSL03 is appropriate for producing biodiesel based on its

high lipid content and oleic acid proportion.

ª 2011 Elsevier Ltd. All rights reserved.

1. Introduction Biomass is one of the better sources of energy to mitigate

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].

6; fax: þ82 33 760 2571.. Jeon).

ier Ltd. All rights reserved

greenhouse gas emissions and to function as a substitute for

fossil fuels [2]. Large-scale introductionof 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 bio-

diesel has been obtained successfully from rapeseed, soybean,

sunflower, corn, palm, and waste cooking oil, as well as from

animal fat [3].However, large-scaleproductionofbiodiesel from

those resources cannot realistically satisfy the existing demand

for transport fuels [1]. Biodiesel has received considerable

.

b i om a s s an d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 53080

attention in recent years, as it is a biodegradable, renewable,

andnon-toxic fuel. It contributesnonet carbondioxideor 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 bio-

fuels 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

photosynthesizingorganismsthatareabletousesolarenergyto

combine water with carbon dioxide to create biomass. Micro-

algae are present in all existing earth ecosystems, both aquatic

and terrestrial, and can flourish under a wide range of envi-

ronmental conditions, including freshwater, brackish water,

seawater, and even wastewater [5]. Microalgae have been sug-

gested 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

producingmore 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 best-

performing 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 concen-

tration 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 iso-

lated 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 ofmicroalgae

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 andmicrotiter plates were incubated at 27 �Cunder 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, andphylogenetic analysis

An aliquot of cultured cells (1mL) was harvested in themid- 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, amplifi-

cation reactions were performed on a T-Gradient thermo-

cycler (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 Sonnen-

berg 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 Bio-

systems, 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 nucleo-

tides or in excess of 1000 nucleotides were removed, and

sequences not belonging to greenmicroalgal specieswere also

discarded from our study. A phylogenetic tree was con-

structed using the neighbor-joining (NJ) algorithm using

Kimura’s two-parameter model of sequence evolution, as

implemented in the MEGA4 program package [16].

b i om a s s a n d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 5 3081

2.3. Microalgae cultivation and biomass

One hundredmilliliters BBM in a 250mL Erlenmeyer flask was

inoculated with the cells (OD680 0.05) and incubated at 27 �Cwith 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 spectrophotom-

eter. 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 L�1) 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 L�1) 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

Fig. 1 e Light microscope (40x) picture

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, respec-tively. Oven temperature conditions were maintained at 50 �Cfor 1min, 200 �C for 12min, and 250 �C for 2min. Mix RM3,Mix

RM5, GLC50, and GLC70 (Supelco Co., USA), and a-linolenic

acid (Sigma Chemical Co. USA) were used as standard mate-

rials. All reagents were of analytical grade. The components

were identified by comparing their retention times and frag-

mentation 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 exis-

tences and purities. Microscopic analysis of the samples

allowed preliminary identification of isolates YSL02, YSL03,

YSL04, YSL05, and YSL07 as genus Scenedesmus, Chlamydo-

monas, Chlorella, Scenedesmus, and Chlamydomonas, respec-

tively. Komarek and Marvan [22] proposed the existence of

at least 13 species of Botryococcus on the basis of morpho-

logical 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 andsequencingPCR 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

s of the tested microalgal isolates.

b i om a s s an d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 53082

(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 (D1-

D2) sequences, we concluded that microalgal isolates YSL02,

YSL03, YSL04, YSL05 and YSL07 were closely related to Scene-

desmus 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 ofmicroalgae,

their specific accession numbers and the nearest identifiable

match present in the GenBank nucleotide database are shown

in Table 1. Identification of the fivemicroalgal 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).

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.

3.2. 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

days�1, 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].

Table 1 e The accession numbers, lengths in base pairs, similasequences for five strains of environmentally isolated microal

Microalgal strain Accession number Length

S. obliquus YSL02 GU732415 864

Chla. pitschmannii YSL03 GU732416 874

C. vulgarisYSL04 GU732417 881

S. obliquus YSL05 GU732418 864

Chla. mexicana YSL07 GU732420 868

S ¼ Scenedesmus, Chla ¼ Chlamydomonas, C ¼ Chlorella and (nt) ¼ nucleotid

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 L�1)

of 1.84� 0.30; 1.71� 0.53; 1.65� 0.07, and 1.53� 0.30were found

for S. obliquus YSL02; S. obliquus YSL05; C. vulgaris YSL04 and

Chla.mexicanaYSL07, respectively. S. obliquusYSL02 showed the

highest biomassproductivity at 1.84� 0.30 gdwt L�1,whileChla.

pitschmannii YSL03 had the lowest biomass productivity at

1.04 � 0.09 g dwt L�1.

Under our experimental growth conditions, the total lipid

contents of the microalgae cultured in this study ranged from

rities between amplified sequences, and the closest relativegae.

(nt) Closest relative andGenBank accession number

% Similarity

S. obliquus AF183482 97

Chla. pitschmannii AF183462 99

C. vulgarisAB237642 97

S. obliquus AF183482 97

Chla. mexicana AF395501 99

es.

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.

Table 3 e Fatty acid composition of the differentmicroalgal species.

Fatty acid Fatty acid composition (wt %)

S.obliquusYSL02

Chla.pitschmannii

YSL03

C.vulgarisYSL04

S.obliquusYSL05

Chla.mexicanaYSL07

b i om a s s a n d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 5 3083

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 L�1 (Table 2).

Lipid content data for different algal species are readily

available in the literature [26]. Manymicroalgae 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 somemicroalgae 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 acid composition

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 lipidproductivities of five microalgal strains.

Microalgalstrain

Biomassproductivity(g dwt L�1)

Lipidproductivity

(g L�1)

Lipid content(% biomass)

S. obliquus YSL02 1.84 � 0.30 0.53 � 0.04 29

Chla. pitschmannii

YSL03

1.04 � 0.09 0.54 � 0.13 51

C. vulgaris YSL04 1.65 � 0.07 0.44 � 0.05 26

S. obliquus YSL05 1.71 � 0.53 0.48 � 0.01 28

Chla. mexicana

YSL07

1.53 � 0.30 0.45 � 0.05 29

Experiments were carried out in triplicate and data are expressed

as mean � SD.

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 trans-

esterification 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), a-

linolenic (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-18-

3 have been confirmed under many conditions including

photoautotrophic and heterotrophic cultivation, nitrogen

starvation, and outdoors in a photobioreactor [34]. In partic-

ular, 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 desir-

able, relatively small percentages of saturated fatty esters can

wreck the cold flow properties of biodiesel. Our finding shows

Lauric acid

(C12:0)

11 10 5 7 34

Myristic acid

(C14:0)

0 8 2 2 10

Palmitic acid

(C16:0)

29 26 22 25 50

Palmitoleic acid

(C16:1)

0 0 5 4 0

Heptadecanoic

acid (C17:0)

0 0 0 2 0

Stearic acid

(C18:0)

17 20 5 23 6

Oleic acid

(C18:1n9c)

20 13 53 8 0

a-Linolenic acid

(C18:3n3)

0 0 0 4 0

g-Linolenic acid

(C18:3n6)

23 23 8 25 0

Total 100 100 100 100 100

b i om a s s an d b i o e n e r g y 3 5 ( 2 0 1 1 ) 3 0 7 9e3 0 8 53084

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 produc-

tivity, 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 days�1,

respectively. These results indicated that the C. vulgaris YSL04

strainwas themost 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.

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