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Environmental manipulation of select algae strains for maximaloil productionAuthor(s): Travis A. Lyon , Joel G. Primm , Michael S. Wojdan , Kristopher D.Morehouse , Melanie L. Grogger , Simina Vintila , and Donald V. VeverkaSource: BIOS, 84(1):21-29. 2013.Published By: Beta Beta Beta Biological SocietyDOI: http://dx.doi.org/10.1893/0005-3155-84.1.21URL: http://www.bioone.org/doi/full/10.1893/0005-3155-84.1.21

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Research Article

Environmental manipulation of select algae strainsfor maximal oil production

Travis A. Lyon, Joel G. Primm, Michael S. Wojdan, Kristopher D. Morehouse,

Melanie L. Grogger, Simina Vintila, and Donald V. Veverka

Department of Biology, United States Air Force Academy, Colorado Springs, CO 80840

Abstract. Consideration for the use of microalgae as a feedstock for commercial biofuels

production began in the 1970s. While high production costs and inefficiencies impeded private

industry scale up to meet national energy needs, recent energy supply concerns and market

instability has revitalized this alternative energy process. A key factor in using microalgae comes

from efficiently harvesting algae oils which are chemically similar to fossil fuel oils. Technological

advances have been made to genetically engineer microalgae to accumulate oils but these

procedures are time consuming and expensive. Others have made progress in using natural

processes in the environment to cause algae to produce oils. Considerable research has shown

determining the appropriate mix of light, temperature and CO2 can foster oil accumulation in select

microalgae strains. Using an experimental design, the freshwater algae, Chlorella pyrenoidosa and

Ettlia sp. were subjected to various environmental conditions to investigate effects on oil

production. When exposed to a higher irradiance level (400 lE), C. pyrenoidosa achieved a

significantly higher cell count leading to an increase in lipid productivity. Under media

manipulation effects, the cell count for nitrogen-limited cultures was significantly lower than that of

cultures grown on replete medium. Nitrogen limitation seems to negatively impact lipid

productivity under our experimental conditions. Supplementation with 2% CO2, led to an increase

in growth after seven days of incubation; however, growth decreased subsequently to a slower rate

as compared to growth with air sparging after 12 days of incubation. There appeared to be a degree

of variability with regards to lipid ratios, with increases in neutral lipids in cultures exposed to

higher levels of irradiance, nitrogen limitation and additional CO2. Although analysis showed that

environmental manipulation is feasible in terms of oil accumulation for these select strains, further

study is warranted in investigating other oil-accumulating microalgae strains for commercial

production.

Introduction

Investigation involving the use of micro-

algae species for the purposes of produc-

ing biofuels began in earnest at the onset

of this country’s energy crisis in the 1970s (Hill

et al., 2006). The term microalga describes a

very diverse group of thousands of organisms

with different types of cells displaying a wide

spectrum in their potential to produce biofuels

or precursor molecules to biofuels. For example,

certain algae accumulate lipids which can be

chemically modified into hydrocarbon fuels or

produce hydrogen, which can be subsequently

used for power generation in vehicles, planes

and other fuel-consuming devices/equipment.Correspondence to: simina.vintila@gmail.com

21BIOS 84(1) 21–29, 2013

Copyright Beta Beta Beta Biological Society

These photosynthetic microorganisms have the

potential to compete favorably with other oil

crops/feedstocks such as soybeans and oil palm.

A major advantage microalgae have over other

types of oil crops is their potential to produce

higher yields of oil per hectare of land. Many

have reported that microalgae have the capabil-

ity to produce a several-fold increase in oil per

acre over that seen in soybean production

(Chisti, 2008; Hu et al., 2008; Sheehan et al.,

1998). Some estimate that even with palm oil,

considered one of the most productive of the oil

crops, production would fall significantly short

of current U.S. biodiesel demands (Chisti,

2007).

The problem largely resides with available

land to grow the requisite amounts of crop oil

needed for conversion to biofuels. Considerable

agricultural land (used for food production)

would be needed to support oil crop require-

ments, severely impacting U.S. capability in

terms of providing cereal grains and livestock.

In contrast, depending on the cultivation

method, microalgae could be grown on margin-

al lands (non-arable, arid, etc.) not suitable for

conventional agricultural practices (Chisti,

2007). Further, many species of algae flourish

in saline/brackish water or coastal seawaters and

can utilize growth nutrients from wastewater

such as nitrogen and phosphorus. Even when

grown under environmentally controlled condi-

tions (photo-bioreactors), microalgae outper-

form land- raised crops by several fold.

Although the literature (NREL, 2006; Sheehan

et al., 1998; Shelef et al., 1984) provides a

myriad of questions that should be addressed to

feasibly use microalgae for renewable and

dependable energy sources, more recent inves-

tigation (Durrett et al., 2008; Hu et al., 2008)

has focused on adequately building sufficient

algal stocks through strain selection (identifica-

tion/screening), biomass accumulation (ponds

vs. photo-bioreactors), production methods

(lipid yield) and harvesting (extraction meth-

ods).

While tens of thousands of species of algae

have been identified, those belonging within the

green algae taxa (Chlorophyceae) appear to hold

the greatest promise (Hu et al., 2008). Green

algae have a tendency to produce more starch

than lipid but have high growth rates at higher

temperatures and lighting conditions. Keeping

in mind the goal to cultivate high quantities of

select microalgae feedstocks to produce oils,

there are some green algae (Botryococcus

braunii and Nannochloropsis oculata) that have

been cited in the literature as having higher than

usual lipid yields (B. braunii reported yields of

up to 80% dry cell weight and Nannochloropsis

species have provided yields nearing 70%)

(Chisti, 2007; Hu et al., 2008). Lipids produced

by these strains contain very long carbon chains

(C23-40), similar to those in petroleum, and

make attractive feedstocks for investigation and

development (Banerjee et al., 2002, Metzger

and Largeau, 2004). Diatoms (Bacillariophyta)

have also been investigated as potential feed-

stocks; however, their nutrient needs are much

different from other algae in that silicate is an

important nutrient for growth.

Determining optimal strains requires in-

depth understanding of the microalgae lipid

production process. Given the pressing eco-

nomic realities associated with biofuels produc-

tion, work that uncovers key environmental

factors which promote cellular lipid develop-

ment would be of great value. While the

literature varies as to what causes the cells of

algae to produce high quantities of cellular

lipids, most agree that stress (nutrient depriva-

tion, varying temperatures, or high light condi-

tions) predisposes the microalgal cell to move

toward the triacylglycerol (TAG) pathway (Hu

et al., 2008). Li et al. (2011) found that

Scenedesmus sp. could grow under a wide

range in temperatures but reported 208 C as the

optimal temperature for greatest level of bio-

mass and lipid production. Additional work by

Lin and Lin (2010) revealed that certain

nitrogen sources can alter fatty acid profiles in

another freshwater algae strain similar to

Scenedesmus rubesens. Weldy and Huesemann

(2007) investigated the results of nitrogen

limitation and light intensity on Dunaliella

salina cultures cultivated within a closed photo-

bioreactor system. Manipulating light and

nutrient levels between comparable systems,

authors found that while nutrient deprivation

22 BIOS

Volume 84, Number 1, 2013

and increased light intensity can increase lipid

content as a percentage of dry weight, total lipid

productivity is much greater under conditions

where Dunaliella salina is nutrient-replete

under high light intensity. While percent lipid

content is an important factor, Weldy and

Huesemann (2007) argue that greater amounts

of oils can be accumulated by healthier

Dunaliella salina cultures via increased cell

numbers (maximizing biomass). Total lipid

productivity should thus be of great importance

to private industry when depending on econo-

mies of scale to ensure profitability and

efficiency of operations.

A number of studies have shown that in

many microalgae cellular lipid accumulation,

generally in the form of triacylglycerides

(neutral lipids), is positively affected by stress

conditions such as nutrient deficiency or high

irradiance (Hu et al., 2008; Roessler, 1990). It

has also been shown that changes in growth

conditions (light intensity, temperature) and/or

nutrient composition can not only affect growth

and lipid quantity but also lipid composition

(Greenwell et al., 2010; Hu et al., 2008; Lin and

Lin, 2010). It should be noted that stressful

growth conditions may actually decrease total

lipid productivity through a negative effect on

growth. However, according to Rodolfi (2009),

few studies have investigated the effect of

different environmental parameters on lipid

productivity (Rodolfi et al., 2009). Nutrient

composition and ratios determine where the

cellular metabolism is directed. Key nutrients

that reportedly affect lipid yield include carbon

(in the form of carbon dioxide), nitrogen and

phosphorus. The carbon/nitrogen (C/N) ratio of

the cell generally determines if energy is to be

spent or stored (in the form of starch or

triacylglycerols). Changes in irradiance levels

primarily affect cells by increasing or decreas-

ing the rate of photosynthesis and subsequently

the rate of carbon fixation, although it is

important to keep in mind that cellular respons-

es to irradiance changes are largely species-

dependent (Guschina and Harwood, 2006).

The manner in which algae partition their

carbon sources towards various metabolic

processes, especially for lipid accumulation,

remains unclear. Developing a balanced set of

environmental variables that promote lipid

productivity, at least for a few specific strains,

would provide insight for an additional com-

mercial process in which to cost- effectively

produce algae-produced biofuels.

This study examines the effect of irradiance,

nitrogen limitation and carbon dioxide sparging

on growth and lipid productivity of the

chlorophyte algae Chlorella pyrenoidosa and

Ettlia sp. We also examined the effect of

osmotic stress on these strains as another

potential environmental factor to ascertain algal

strain robustness under varying conditions. The

genus Chlorella is generally a freshwater alga

but, interestingly, C. pyrenoidosa has been

reported to grow in saline medium (Poyton,

1970). Using marine medium also demonstrates

the flexibility of microalgae as a biofuels

feedstock to avoid competing for freshwater

resources. While the effects of irradiance,

nitrogen starvation and carbon dioxide (CO2)

supplementation on total lipid accumulation

have been investigated for a number of green

algae, no study has investigated the effects of

these parameters on lipid productivity during

osmotic stress.

Materials and Methods

Growth conditions

Chlorella pyrenoidosa (obtained from Air

Force Research Laboratories, Tyndall AFB, FL)

was grown in modified f/2 medium (Guillard

and Ryther, 1962) excluding vitamins. Cultures

of Ettlia sp. (obtained from Dr. Juergen Polle,

Brooklyn College) were grown in modified

Bold’s Basal medium with 1/3 of nitrogen and

without vitamins (Culture Collection of Algae

and Protozoa, http://www.ccap.ac.uk/media/

documents/3N_BBM_V_000.pdf). For the ni-

trogen limitation experiment, Chlorella cultures

were grown at 258C and 200 lE irradiance,

either in nitrogen-replete or nitrogen-limited

medium using batch cultures, and samples were

collected 6 and 12 days after the start of the

experiment. Nitrogen limitation was achieved

by decreasing the nitrate levels 20-fold com-

pared to the original concentration in the f/2

Manipulation of algae for maximal oil production 23

Volume 84, Number 1, 2013

medium (from 8.8 to 0.44 mM). For the

irradiance experiment, batch cultures were

grown at 200 lE and 400 lE in f/2 medium

on an orbital shaker. To investigate the effects of

CO2 on growth and lipid accumulation, semi-

continuous cultures of C. pyrenoidosa and Ettlia

sp. were grown in a 14 L BioFlo 115 photo-

bioreactor (New Brunswick Scientific, Edison,

NJ) in f/2 or BBM medium, respectively, with a

media flow rate of 1 L/day (0.6944 ml/min) for

C. pyrenoidosa and 6 L/day (4.2 ml/min) for

Ettlia sp. The cultures were sparged with 2%CO2 and 98% air at 0.5 L/min.

Growth was monitored during all experiments

by cell counts using either a hemocytometer or

flow cytometer (Accuri C6 Flow Cytometer, Ann

Arbor, MI). All experiments were run in

biological triplicates (except experiment on

Ettlia sp.). Samples were taken in duplicates.

Sampling and lipid extraction

For lipid extraction, 500 ml of cell culture

was centrifuged at 6000 x g for 10 min. Cell

pellets were freeze dried and stored at -208C

until subsequent lipid extraction. Lipid extrac-

tion was performed as previously reported with

modifications (Bigogno et al., 2002). Briefly,

freeze-dried cells were suspended in 4-5 ml of a

chloroform : methanol mixture (2:1, v/v) with

0.2 g quartz particles (size 1.2-20 lm, Sigma).

Each mixture was homogenized with a needle

sonicator at 18 W for 3 minutes. For maximum

extraction, the suspension was allowed to stand

for at least 6-8 h. Each vial was then centrifuged

at 7000 x g for 10 min and the organic phase

was transferred to a new vial. Chloroform:water

(1:1, v/v) was added to rinse non-lipid mole-

cules from the organic phase and vials were

centrifuged at 7000 x g for 10 min. The organic

phase was transferred to pre-weighed glass vials

and the solvent was evaporated under N2

atmosphere. The remaining algal oil was

weighed and normalized.

Lipid analysis

Lipid classes were analyzed using automated

thin layer chromatography coupled to a flame

ionization detector (TLC-FID) system (Iatro-

scan MK6, Iatron, Japan). The lipid samples

were dissolved in 200 ll of a chloroform:me-

thanol mixture (2:1, v/v), then 1 ll of each

sample was spotted onto silicon rods. The rods

were developed using a two-bath system. The

first bath contained a mixture of hexane:diethyl

ether:formic acid (98:2:0.1, v/v/v) and rods

were developed for 20 min. The second bath

contained a mixture of hexane:diethyl ether:

formic acid (80:20:0.1, v/v/v) and the rods were

developed for 10 min. The rods were then dried

at 1208C for 2 minutes. The samples were

placed in the Iatroscan, which was set at 2 L/

min air and, 160 ml/min H2, and set to scan for

30 seconds. Samples were run in technical

duplicates. Glycerol Trioleate, Palmitic Acid,

Cholesterol, Nonadecane and Palmityl Palmitate

(Sigma Aldrich, St Louis, MO) were used as

external standards.

Statistical analysis

ANOVA followed by Tukey HSD was

carried out using the R statistical software

program, version 2.3.1 for Windows XP

(http://www.r-project.org/).

Results

The effects of irradiance, nitrogen limitation

and CO2 supplementation on growth, lipid

composition and productivity of the green alga

Chlorella pyrenoidosa were examined. In addi-

tion, the effect of CO2 supplementation was

investigated for one other green alga, Ettlia sp.

The results indicate that all three of the above

mentioned parameters affect growth and lipid

productivity.

Effect of irradiance on growth and lipid

productivity

C. pyrenoidosa cultures grown under a

higher irradiance (400 lE) as compared to

control cultures (200 lE) achieved a signifi-

cantly higher (p < 0.001) cell count (Figure 1a)

during the incubation period. At the same time,

the proportion of neutral lipids was 25 % lower

24 BIOS

Volume 84, Number 1, 2013

(p = 0.02) at 400 lE than 200 lE (Figure 1b).

While the lipid amount per cell may be lower

after nine days of incubation than for cultures

grown under 200 lE, the higher cell count

under 400 lE actually led to higher lipid

accumulation per liter of culture. Taken togeth-

er, the results indicate that higher irradiance has

a positive effect on lipid productivity.

Effect of nitrogen limitation on growth and

lipid productivity

To achieve nitrogen-limiting conditions, the

nitrate amount in the medium was reduced from

8.8 mM to 0.44 mM (a 20-fold reduction). C.

pyrenoidosa cultures were incubated for a total

of 12 days. As shown in Figure 2a, no

difference in cell density could be seen between

nitrogen-limited cultures and control cultures

for the first 6 days of the experiment (p = 0.99).

After 12 days of incubation, the cell count for

nitrogen-limited cultures was significantly lower

(p < 0.001) than that of cultures grown on

replete medium (Figure 2a). Interestingly, the

proportion of neutral lipids decreased signifi-

cantly (p = 0.004) between day 6 and day 12 of

incubation (Figure 2 b) for the control culture

and while a slight decrease could also be

observed for the nitrogen-limited culture, the

decrease in neutral lipids was not statistically

significant (p = 0.144).The proportion of

neutral lipids was also somewhat higher under

nitrogen-limiting conditions (Figure 2c). While

the relative amount of lipid did not differ

between cultures grown on replete medium

and nitrogen-limited cultures (day 6 p = 0.2;

day 12 p = 0.4), the cell counts were

significantly different (p < 0.001). When

growth rate is taken into account, nitrogen

limitation, at the level tested, has a negative

effect on total lipid productivity (lipid accumu-

lation per liter of culture).

Effect of CO2 supplementation on growth

and lipid productivity

Cultures subjected to CO2 supplementation

were grown under semi-continuous conditions.

C. pyrenoidosa cultures supplemented with 2%

CO2 displayed an increase in growth after seven

Figure 1. Cell density and neutral:polar lipid ratio at varying irradiance a) Cell densities of C. pyrenoidosa culturesgrown at 200 lE and 400 lE. b) Relative proportions of neutral and polar lipids in C. pyrenoidosa cultures grown at 200 lEand 400 lE.

Manipulation of algae for maximal oil production 25

Volume 84, Number 1, 2013

days of incubation but growth decreased and

cell densities were lower than at the start of the

experiment after 12 days of incubation (p <0.001; Figure 3a). The proportion of neutral

lipids was negatively correlated with growth

(Figure 3b). During the increase in cell growth,

the proportion of neutral lipids decreases but

increases again after 12 days of incubation

(Figure 3 b) There was however some variabil-

ity between samples and therefore the observed

changes in neutral lipid proportion are not

statistically significant (p = 0.209).

The same experiment was performed on

another freshwater alga, Ettlia sp. As shown in

Figure 4a, the response to CO2 supplementation

of Ettlia sp. cultures was similar to that of C.

pyrenoidosa. During the first 6 days of incuba-

tion, the cultures displayed an increase in

growth accompanied by a decrease in the

proportion of neutral lipids (Figure 4b). After

8 days of incubation, the growth was markedly

reduced and the cells had an increased propor-

tion of neutral lipids accumulated. While total

lipid accumulation per cell was increased at the

end of each experiment for both species, the

overall growth decreased. Considering both

total lipid accumulation per cell and growth,

the highest lipid productivity can be found 6 to

7 days after CO2 supplementation, coinciding

with the increase in growth for both species.

Discussion

The main goal of this study was to

investigate the effect of three environmental

conditions on lipid productivity for species not

previously investigated. The results are in

agreement with previous studies showing that

stress conditions seem to positively affect the

cellular accumulation of lipids, in particular

Figure 2. Cell densities and neutral:polar lipid ratio on replete and N-limited media a) Cell densities of C. pyrenoidosacultures grown in full medium and nitrogen limited medium. b) Relative proportions of neutral and polar lipids in C.pyrenoidosa cultures grown in replete medium. c) Relative proportions of neutral and polar lipids in C. pyrenoidosa culturesgrown in nitrogen-limited medium

26 BIOS

Volume 84, Number 1, 2013

neutral lipids. However, this study also shows

that while certain stress conditions, such as

nitrogen limitation, may increase the lipid

accumulation or the percentage of neutral lipids,

the total lipid productivity may be lower due to

the negative effect on growth. This study finds

that irradiance levels of 400 lE have a positive

effect on lipid productivity for C. pyrenoidosa

compared to 200 lE. We did not, however,

investigate higher irradiances. While higher

irradiance levels may have had an even greater

effect on lipid productivity, high irradiance

levels may also reduce algal growth rate through

photoinhibition (Radakovits et al., 2010) and

thereby reduce lipid productivity overall. These

results are also in agreement with previous

studies showing an increase in the proportion of

polar lipids with an increase in irradiance levels

(Gordillo et al., 1998). This can be explained by

the positive effect of a higher irradiance level on

growth. During a fast growth rate cells have to

produce polar lipids, such as phospho- and

sphingolipids, for cellular membranes while

storage lipids are mobilized to sustain a fast

growth rate.

Many studies have investigated the effect of

nitrogen limitation on lipid accumulation and

shown that during nitrogen starvation cells

increase total lipid accumulation, mainly in the

form of storage lipids (neutral lipids) (Converti

et al., 2009; Gordillo et al., 1998; Guschina and

Harwood 2006; Illman et al., 2000). Some

studies (Converti et al., 2009; Zhila et al., 2005)

have mentioned the negative effect of nitrogen

starvation on cellular growth but did not

correlate the effect of nitrogen limitation on

lipid productivity. The results of this study

suggest that nitrogen limitation has a negative

effect on lipid productivity if cultures are

limited for more than a week for the species

and conditions tested. As we only investigated

two levels of nitrogen, however, future studies

are planned to investigate a range of nitrogen

levels. This will help determine the most

optimal nutrient composition(s) that induce the

highest level of lipid accumulation without

excessively inhibiting growth, thus yielding

the greatest total lipid productivity

Carbon dioxide supplementation to algal

cultures is known to increase the growth rate

since the CO2 levels in aqueous environments

are very low. Previous studies have reported that

CO2 supplementation leads to an increase in

growth rate but no apparent increase in the

Figure 3. Cell densities and neutral:polar lipid ratio with 2 % CO2 sparging (C. pyrenoidosa) a) Cell densities of C.pyrenoidosa cultures grown with 2 % CO2. b) Relative proportions of neutral and polar lipids in C. pyrenoidosa culturesgrown with 2 % CO2.

Manipulation of algae for maximal oil production 27

Volume 84, Number 1, 2013

proportion of neutral lipids (Gordillo et al.,

1998, Chiu et al., 2009). Our results show the

same trend although prolonged supplementation

of CO2 led to a decrease of pH in the media

which probably led to the observed decrease in

growth. Based on the acquired data, a few days’

exposure to 2% CO2 or longer exposure to

lower levels of CO2 will increase lipid produc-

tivity by increasing cell count.

While the results of this are generally in

agreement with previous studies showing that

lipid accumulation is positively affected by

environmental stress, this stress-induced lipid

accumulation correlates negatively with total

lipid productivity due to a negative impact of

stress on growth. Our results stress the need for

future algal biodiesel research to focus more on

parameters increasing lipid productivity, not

only lipid accumulation, if biodiesel is to

become a viable option to gasoline.

Acknowledgements: Funding support for this

research was made possible through the Air

Force Office of Scientific Research and Air

Force Research Laboratories. The authors

would also like to thank Dr. Juergen Polle,

Brooklyn College, for supplying promising

algae strains.

The views expressed in this article are those of

the authors and do not reflect the official policy

or position of the United States Air Force, the

Department of Defense, or the US Government.

References

Banerjee, A., Sharma, R., Chisti, Y. and Banerjee, U. (2002).

Botryococcus braunii: A Renewable Source of Hydro-

carbons and Other Chemicals. Crit Rev Biotechnol. 22

(3), 245–279.

Bigogno, C., Khozin-Goldberg, I., Boussiba, S., Vonshak,

A., and Cohen, Z. (2002). Lipid and fatty acid

composition of the green oleaginous alga Parietochloris

incisa, the richest plant source of arachidonic acid.

Phytochemistry 60, 497–503.

Chisti, Y. (2007). Biodiesel from microalgae. Biotechnol

Adv. May-June, 25 (3), 294–306.

Chisti, Y. (2008). Biodiesel from microalgae beats bio-

ethanol. Trends Biotechnol. March, 26 (3), 126–131.

Chiu, S-Y., Kao, C-Y., Tsai, M-T., Ong, S-C., Chen, C-H.,

and Lin, C-S. (2009). Lipid accumulation and CO2

utilization of Nannochloropsis oculata in response to

CO2 aeration. Biores Technol, 100, 833–838.

Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P. and del

Borghi, M. (2009). Effect of temperature and nitrogen

concentration on the growth and lipid content of

Nannochloropsis oculata and Chlorella vulgaris for

Figure 4. Cell densities and neutral:polar lipid ratio with 2% CO2 sparging (Ettlia sp.) a) Cell densities of Ettlia sp.cultures grown with 2 % CO2. b) Relative proportions of neutral and polar lipids in Ettlia sp. cultures grown with 2 % CO2.

28 BIOS

Volume 84, Number 1, 2013

biodiesel production. Chem Eng Process, 48, 1146–1151.

Durrett, T.P., Benning, C., and Ohlrogge, J. (2008). Planttriacylglycerols as feedstocks for the production ofbiofuels. Plant J, May, 54 (4), 593–607.

Gordillo, F.J.C., Goutx, M., Figueroa, F.L. and Niell, F.X.(1998). Effects of light intensity, CO2 and nitrogensupply on lipid class composition of Dunaliella viridis. JAppl Phycol, 10, 135–144.

Greenwell, H.C., Laurens, L.M.L., Shields, R.J., Lovitt,R.W., and Flynn, K.J. (2010). Placing microalgae on thebiofuels priority list: a review of the technologicalchallenges. J R Soc Interface. 7, 703–726.

Guillard, R.R.L. and Ryther, J.H. (1962). Studies of marineplanktonic diatoms. I. Cyclotella nana Hustedt andDetonula confervaceae (Cleve). Can. J. Microbiol. 8,229–239.

Guschina, I. A and Harwood, J. L. (2006). Lipids and lipidmetabolism in eukaryotic algae. Prog Lipid Res, 45,160–186.

Hill, J., Nelson, E., Tilman, D., Polasky, S., and Tiffany, D.(2006). Environmental, economic, and energetic costsand benefits of biodiesel and ethanol biofuels. Proc NatlAcad Sci USA. July 25, 103(30), 206–10.

Hu, Q., Sommerfeld, M., Jarvis, E., Ghirardi, M., Posewitz,M., Seibert, M., and Darzins, A. (2008). Microalgaltriacylglycerols as feedstocks for biofuel production:perspectives and advances. Plant J, May, 54(4), 621–39.

Illman, A.M., Scragg, A.H. and Shales, S.W. (2000).Increase in Chlorella strains calorific values when grownin low nitrogen medium. Enzyme Microb Technol, 27,631–635.

Li, X., Hu, H.Y., and Zhang, Y.P. (2011). Growth and lipidaccumulation properties of a freshwater microalgaScenedesmus sp. under different cultivation temper-ature.Bioresour Technol. Feb;102(3), 3098–102

Lin, Q. and Lin, J. (2010). Effects of nitrogen source andconcentration on biomass and oil production of aScenedesmus rubescens like microalga. Bioresour Tech-nol. 102, 1615–1621.

Metzger, P. and Largeau, C. (2004). Botryococcus braunii: a

rich source for hydrocarbons and related ether lipids.

Appl Microbiol Biotechnol. February, 66 (5), 486–496.

National Renewable Energy Laboratory (NREL). Jet Fuel

from Microalgal Lipids. (Fact Sheet). 2006. 2 pp.; NREL

Report No. FS-840–40352.

Poyton, R.O. (1970). The Characterization of Hyalochlor-

ella marina gen. et sp.nov. a New Colourless Counter-

part of Chlorella. J Gen Microbiol. 62, 171–188.

Radakovits, R., Jinkerson, R.E., Darzins, A. and Posewitz,

M.C. (2010). Genetic engineering of algae for enhanced

biofuel production. Eukaryot Cell, 9, 486–501.

Rodolfi, L., Chini Zittellini, G., Bassi, N., Padovani, G.,

Biondi, N., Bonini, G. and Tredici, M. R. (2009).

Microalgae for oil: strain selection, induction of lipid

synthesis and outdoor mass cultivation in a low cost

photobioreactor. Biotechnol Bioeng, 102, 100–112

Roessler, P.G. (1990). Environmental control of glycerolipid

metabolism in microalgae: commercial implications and

future research directions. J Phycol. 26, 393–399.

Sheehan, J., Dunahay, T.G., Benemann, J.R., Roessler, P.G.,

and Weissman, J.C. (1998). A Look Back at the US

Department of Energy’s Aquatic Species Program:

Biodiesel from Algae - Close Out Report. 328 pp.;

NREL Report No. TP-580-24190.

Shelef, G., Sukenik, A., and Green, M. Microalgae

harvesting and processing: a literature review. Solar

Energy Research Institute, U.S. Department of Energy

Technical Report (SERI/STR-231-2396), August 1984.

Weldy C.S., and Huesemann M. (2007). Lipid production by

Dunaliella salina in batch culture: effects of nitrogen

limitation and light intensity. U.S. Department of Energy

Journal of Undergraduate Research. 7, 115–122.

Zhila N.O., Kalacheva, G.S. and Volova, T.G. (2005). Effect

of nitrogen limitation on the growth and lipid compo-

sition of the green alga Botryococcus braunii KutzIPPAS H-252. J Plant Physiol. 52(3), 311–319.

Received 18 December 2011; accepted 19 July 2012.

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