Chemical Engineering and Processing 48 (2009) 1146–1151
Contents lists available at ScienceDirect
Chemical Engineering and Processing:
Process Intensification
journal homepage: www.elsevier.com/locate/cep
Effect of temperature and nitrogen concentration on the growth
and lipid content of Nannochloropsis oculata and Chlorella vulgaris
for biodiesel production
Attilio Converti ∗ , Alessandro A. Casazza, Erika Y. Ortiz, Patrizia Perego, Marco Del Borghi
Department of Chemical and Process Engineering “G.B. Bonino”, University of Genoa, Via Opera Pia 15, 16145 Genoa, Italy
a r t i c l e
i n f o
Article history:
Received 1 February 2009
Received in revised form 23 March 2009
Accepted 24 March 2009
Available online 1 April 2009
Keywords:
Biodiesel
Microalgae
Lipids
Extraction
Nannochloropsis oculata
Chlorella vulgaris
Carbon dioxide
a b s t r a c t
A possible source of biological material for the production of biodiesel is represented by microalgae, in
particular by their lipid content. The aim of the present work was to study of the effects of temperature
and nitrogen concentration on the lipid content of Nannochloropsis oculata and Chlorella vulgaris in view of
their possible utilization as novel raw materials for biodiesel production. In addition, various lipid extraction methods were investigated. The extracted lipids were quantitatively and qualitatively analyzed by
gravimetric and gas chromatographic methods, respectively, in order to check their suitability according
to the European standards for biodiesel. The lipid content of microalgae was strongly influenced by the
variation of tested parameters; indeed, an increase in temperature from 20 to 25 ◦ C practically doubled
the lipid content of N. oculata (from 7.90 to 14.92%), while an increase from 25 to 30 ◦ C brought about a
decrease of the lipid content of C. vulgaris from 14.71 to 5.90%. On the other hand, a 75% decrease of the
nitrogen concentration in the medium, with respect to the optimal values for growth, increased the lipid
fractions of N. oculata from 7.90 to 15.31% and of C. vulgaris from 5.90 to 16.41%, respectively.
© 2009 Elsevier B.V. All rights reserved.
1. Introduction
The microalgae are unicellular photosynthetic organisms that
use light energy and carbon dioxide, with higher photosynthetic
efficiency than plants for the production of biomass [1,2]. They may
be destined to different applications, such as biofuel production,
purification of wastewater under either autotrophic or mixotrophic
conditions [3,4], extractions of high added value foods and pharmaceutical products, or as food for aquaculture [5].
The lipids from microalgae could be used in different processes
for energy exploitation, including the simple combustion in boiler
or in a diesel engine. However, the best possible use of this oil is
certainly its transformation to a biofuel, especially biodiesel [6].
Moreover, high added value compounds can be extracted from
microalgae, such as fatty acids (␥-linolenic, arachidonic, eicosapentaenoic, docosahexaenoic acids, etc.) [7,8], pigments (carotenoids
and ficobiliproteins), biochemically stable isotopes [6,9–11] and
vitamins such as biotin [12], vitamins C [13] and E [14,15]; also some
metabolites appear to have some pharmacological activities, among
others the anticholesterolemic, antitumoral, immunomodulatory,
antibacterial and antimycotic ones.
∗ Corresponding author. Tel.: +39 010 353 2593; fax: +39 010 353 2586.
E-mail address: [email protected] (A. Converti).
0255-2701/$ – see front matter © 2009 Elsevier B.V. All rights reserved.
doi:10.1016/j.cep.2009.03.006
Finally in aquaculture, the microalgae are used as main food
of rotifers, and as very important live food for larvae of marine
fish, filter-feeding invertebrates, etc. In 1999, the production of
microalgae for aquaculture was 1000 tons: 62% for mollusks, 21%
for shrimps and 16% for fish [16].
Microalgae biomass contains approximately 50% of carbon on a
dry weight basis [17], and all the carbon present in cell is usually
from carbon dioxide. With the production of 100 tons of microalgae
biomass, around 180 tons of CO2 can be disposed using natural or
artificial light.
The microalgae for biodiesel production can potentially use part
of carbon dioxide from industrial plants [18,19]; from this point of
view the microalgae can also be seen as simple CO2 sequestrants
to use in plants for greenhouse gas emissions control (ET—Kyoto
Protocol).
Given the high productivity, justified by the shorter generation
time and higher oil content compared to crops (up to 80% on dry
weight) [20], it is clear the potential contribution that microalgae
can give on large scale fuel oils production. A further benefit is that
microalgae may also be grown on arid lands unsuitable for conventional agriculture, such as desert areas, or in large reservoirs of
saline water.
Several studies have shown that the quantity and quality of lipids
within the cell can vary as a result of changes in growth conditions
(temperature and light intensity) or nutrient media characteristics
(concentration of nitrogen, phosphates, and iron) [21,22].
A. Converti et al. / Chemical Engineering and Processing 48 (2009) 1146–1151
This work is a first attempt to increase the lipid content in
microalgae as a result of variations in temperature and concentration of nitrogen in two different species of microalgae: Nannochloropsis oculata H. and Chlorella vulgaris. The lipids extracted
from the two microalgae grown in Erlenmeyer flasks under conditions favoring lipid content in the cell have been analyzed
quantitatively and qualitatively, by gravimetric and gas chromatographic methods, and different extraction methods have been
investigated to maximize the yield in biodiesel.
2. Materials and methods
2.1. Microalgae
Two microalgal species were used in this study, specifically N.
oculata H. (from aquaculture of Rosignano’s Society of Marine fish,
Italy) and C. vulgaris CCAP 211 (Culture Collection of Algae and Protozoa, Argyll, UK). Both microalgae are eukaryotic photosynthetic
microorganisms that grow rapidly due to their simple structure
[23]. Because of their small size ranging from 2 to 4 ␮m, they are
considered to be part of phytoplankton. C. vulgaris was grown in the
Bold’s Basal Medium and N. oculata on the Guillard f2 one [24,25]
using the CO2 contained in air (about 300 ppm) and NaNO3 as the
sole sources of carbon and nitrogen [26], respectively.
2.2. Culture system
Growth experiments were done at different temperatures and
concentrations of nitrogen in 2.0 L-Erlenmeyer flasks. The medium
and flasks were sterilized in an autoclave for 20 min at 121 ◦ C
in order to prevent any contamination during the early stages of
growth.
The growth was done using a refrigerated incubator series 6000
(Termaks, Milan, Italy) equipped with artificial lighting, and the air,
and consequently the CO2 contained in it, was injected continuously
through pumps M2K3 (Schego, Offenbach, Germany) after passing
through a cotton trap.
Each autotrophic batch cultivation was carried out in duplicate
for 14 days at a continuous photon flux density of 70.0 ␮E m−2 s−1 ,
which was measured by a Radiometer HD9021 (Delta OHM, Padua,
Italy). The duration of cultivations and light intensity were established on the basis of literature [25,27,28].
Temperature and nitrogen concentration in the medium were
selected as the independent variables. The central values of
temperature, 20 ◦ C for N. oculata and 30 ◦ C for C. vulgaris, were chosen on the basis of data reported in the literature [28,29]. Then,
additional experiments were done either increasing or reducing
the growth temperature by 5 ◦ C intervals with respect to the above
optima. Moreover, C. vulgaris was also grown at 38 ◦ C to evaluate
the effect of high temperature on the lipid content of the microalga.
Based on these considerations, the temperature values tested for N.
oculata were 15, 20 and 25 ◦ C and those for C. vulgaris 25, 30, 35 and
38 ◦ C.
Because the literature suggested that nitrogen limitation could
exalt the lipid content of many microalgae [23], the central concentrations of nitrogen in medium (0.300 g L−1 for N. oculata and
1.50 g L−1 for C. vulgaris) were selected according to Guillard [25],
and additional cultivations were run at 0.150 and 0.075 g L−1 with
N. oculata and at 0.750 and 0.375 g L−1 with C. vulgaris, respectively.
2.3. Microalgal biomass concentration
The microalgae concentration was determined daily by optical
density measurements at 625 nm by a UV–vis spectrophotometer,
model Lambda 25 (PerkinElmer, Milan, Italy). All measures were
1147
carried out in triplicate, and biomass concentration was related to
optical density by the equation y = 2.416 × (R2 = 0.994) for N. oculata
and y = 4.203 × (R2 = 0.990) for C. vulgaris, respectively.
After growth, biomass was separated from the medium by centrifugation at 5000 rpm for 15 min, using a centrifuge model 42426
(ALC, Milan, Italy). Biomass was dried at 105 ◦ C for 48 h, pulverized
in a mortar and stored at −20 ◦ C for later analysis.
2.4. Lipids extraction
The lipid fraction was extracted by
• Classic extraction, using petroleum ether (EtP) as a solvent and
an extraction time of 4 h;
• Soxhlet extraction [30] under the same conditions;
• Folch method [27] that makes use of a mixture of chloroform and
methanol (2:1, v/v) for 1.5 h;
• The methodology described by Krienitz and Wirth [31] that consists in the Folch method combined to the use of ultrasounds
(mod. UP100H, Hielscher, Teltow, Germany) and increasing the
extraction time from 1.5 h to 6.0 h;
• Ultrasonic extraction using petroleum ether as a solvent [32] and
the same instrumentation as in the preceding method.
All extractions were performed on dry biomass. A further test
was also carried out on wet biomass by the Folch method combined
to ultrasounds to assess the possible influence of water on the action
of ultrasounds.
2.5. Kinetic and yield parameters
• The specific growth rate was calculated by the equation:
=
1
ln
t
X m
X0
(1)
where Xm and X0 are the concentrations of biomass at the end and
at the beginning of a batch run, respectively, and t is the duration
of the run.
• The lipid productivity was calculated by the equation:
=
CL
t
(2)
where CL in the concentration of lipids at the end of the batch run
and t is the duration of the run.
• The yield of the microalgae lipids was calculated by the equation:
Y (%) =
WL
WDA
(3)
where WL and WDA are the weights of the extracted lipids and of
the dry algae biomass, respectively.
2.6. Lipids analysis
To remove residual microalgae, the extraction products were
filtered on membranes Type FH (Millipore, Bedford, MA) with
0.50 ␮m mean pore diameter. After washing twice with the extraction solvent and its complete evaporation, gravimetric analysis was
done. Part of the lipid fraction was transesterified through the
method described by Zunin et al. [33] and qualitatively characterized by a gas chromatograph, model Ultra Trace (Thermo Finnigan,
Milan, Italy) equipped with ZB Vax column and FID detector (Thermoscientific, Milan, Italy) using appropriate reference standards
(Sigma–Aldrich, Milan, Italy).
1148
A. Converti et al. / Chemical Engineering and Processing 48 (2009) 1146–1151
Fig. 1. Lipid yield (glipids /100 gmass ) obtained by different techniques of extraction
from N. oculata grown at 20 ◦ C, 70 ␮E m−2 s−1 and 0.3 g L−1 NaNO3 . ( ) Wet biomass;
( ) dry biomass. S (classic extraction), SL (Soxhlet), UAE (Ultrasound-assisted extraction), and F (Folch method).
3. Results and discussion
3.1. Selection of lipid extraction methodology
The first part of this study dealt with the extraction of lipids from
the cell, in order to determine the percentage of this component
in the microalgae dry mass. Different methods of extraction were
compared so as to select the one able to extract the highest oily
fraction and to increase the reliability of results on lipid content
variations due to changes in growth conditions. For those purposes,
N. oculata was initially grown under the conditions suggested by
the literature (20 ◦ C, 70 ␮E m−2 s−1 and 0.3 g L−1 of NaNO3 ) to get
a total amount of biomass (about 20 g) sufficient to perform all the
extraction tests.
Fig. 1 shows that the most effective extraction method, among
those taken into consideration, was the combination of ultrasound
with the Folch method, which led to a yield (>20%) that by far
exceeded those obtained by the traditional Soxhlet (7%) and Folch
(8%) methods. However, such a methodology did not show significant differences when applied to dried and wet biomass, the
maximum lipid extraction yields being 24.3 and 22.8%, respectively.
The kinetics of extraction using the Folch method combined
with ultrasounds is shown in Fig. 2. As can be seen, most of lipids
Fig. 2. Kinetics of lipids extraction from N. oculata using Folch method combined
with the use of ultrasounds. () Wet biomass; () dry biomass.
(80%) were transferred to the chloroform/methanol solvent within
the first 90 min of extraction, either using dry or wet biomass,
even if the complete extraction required a total time period of
6 h.
Based on these results, it was decided to use the combination of
ultrasounds and Folch method to extract the lipid fraction from dry
biomass grown at different temperatures and nitrogen concentrations in the medium, as described later.
3.2. Effect of temperature and nitrogen concentration on
microalgae growth
In the second part of this work, we studied the effects of
temperature and nitrate concentration on batch growth of the two
selected microalgae.
C. vulgaris growth appeared to be affected at temperatures
above 30 ◦ C (Table 1). At 35 ◦ C, this microalga did in fact exhibit a
17% decrease in its growth rate () when compared to 30 ◦ C. Further
increase in temperature (38 ◦ C) led to an abrupt interruption of
microalgal growth, and later the cells dead. This result was easily
visible because of color change of the cells from green to brown,
and consequently the microalgal growth rate was even negative.
As far as N. oculata is concerned, changes from the optimal
conditions of growth (20 ◦ C) also resulted in significant changes
in the microalgae growth rate (Table 2). For temperatures below
this threshold, growth rate more than halved, falling from 0.13 to
Table 1
Main parameters of growth and lipid production of C. vulgaris at different temperatures.
Temperature (◦ C)
a (days−1 )
25
30
35
38
0.14
0.14
0.12
−0.01
a
b
c
±
±
±
±
0.00
0.00
0.01
0.01
Yb (glipids (100 gdry algae )−1 )
14.71
5.90
6.60
11.32
±
±
±
±
0.30
0.42
0.59
0.20
c (mglipids L−1 days−1 )
20.22
8.16
8.21
−2.72
±
±
±
±
0.60
0.65
0.17
1.62
Specific growth rate.
Lipid yield using the Folch method combined with ultrasounds.
Lipid productivity.
Table 2
Main parameters of growth and lipid production of N. oculata at different temperatures.
Temperature (◦ C)
a (days−1 )
Yb (glipids (100 gdry algae )−1 )
c (mglipids L−1 days−1 )
15
20
25
0.06 ± 0.00
0.13 ± 0.00
0.07 ± 0.01
14.92 ± 0.82
7.90 ± 0.21
13.89 ± 0.61
9.11 ± 0.30
10.01 ± 0.22
10.10 ± 2.09
a
b
c
Specific growth rate.
Lipid yield using Folch method and ultrasound.
Lipid productivity.
A. Converti et al. / Chemical Engineering and Processing 48 (2009) 1146–1151
1149
Table 3
Main parameters of growth and lipid production of C. vulgaris at different NaNO3 concentration in the growth medium.
NaNO3 (g L−1 )
1.500
0.750
0.375
a
b
c
a (days−1 )
0.14 ± 0.00
0.14 ± 0.01
0.13 ± 0.00
Yb (glipids (100 gdry algae )−1 )
5.90 ± 0.42
14.37 ± 0.64
15.31 ± 0.51
c (mglipids L−1 days−1 )
8.16 ± 0.65
20.44 ± 0.75
20.30 ± 0.40
Specific growth rate.
Lipid yield using Folch method and ultrasound.
Lipid productivity.
Table 4
Main parameters of growth and lipid production of N. oculata H at different NaNO3 concentration in the growth medium.
NaNO3 (g L−1 )
0.300
0.150
0.075
a
b
c
a (days−1 )
0.13 ± 0.00
0.10 ± 0.00
0.10 ± 0.00
Yb (glipids (100 gdry algae )−1 )
c (mglipids L−1 days−1 )
7.88 ± 0.21
13.01 ± 0.39
15.86 ± 0.59
10.01 ± 0.16
13.61 ± 1.10
16.41 ± 0.11
Specific growth rate.
Lipid yield using Folch method and ultrasound.
Lipid productivity.
0.06 day−1 . A sharp drop in microalgae growth rate was also noticed
at higher temperature (25 ◦ C), as reported in the literature [28].
Using the same microalga, an additional investigation was
carried out on the effect resulting from a reduction of the nitrogen
concentration in the medium. Nitrogen limiting conditions were
in fact reported to significantly increase the lipid fraction of many
microalgae [21]. For this purpose, the concentration of nitrate in
both media for N. oculata and C. vulgaris batch growth was reduced
to half and quarter of the standard media described in Section 2
while the light intensity and the air flux were kept the same trough
the experiments.
The effect of a reduction of NaNO3 concentration on C. vulgaris
growth is summarized in Table 3. Whereas its specific growth rate
was not significantly affected, a threefold increase in lipid content
took place.
In contrast, N. oculata showed a gradual decrease in the growth
rate accompanied by almost a duplication of the lipid content
(Table 4).
These results as a whole suggest that, to perform an effective
biodiesel production from microalgae, the optimum compromise
between a slowdown in growth and the increase in the lipid fraction
should be achieved.
3.3. Effect of temperature and concentration of nitrogen source
on the quantity and composition of the lipid fraction
Variations of temperature and decrease in the concentration of
nitrate in the medium resulted in a significant change in cell composition, favoring the accumulation of lipid components in both
microalgae during the batch growths (Tables 1–4).
A decrease in the growth temperature from 30 to 25 ◦ C led to
an increase in the lipid content of C. vulgaris from 5.9 to 14.7%,
while the rate of growth remained unchanged and, as a result, lipid
productivity () increased from 8 to 20 mg L−1 day−1 (Table 1).
Even in N. oculata, there was an increase in the lipid component
passing from 25 ◦ C (13.9%) to suboptimal temperature conditions
Fig. 3. Percentages of individual fatty acid methyl ester (FAME) on the total FAMEs (g/100 gFAME ) in C. vulgaris at different temperatures and concentrations of NaNO3 in the
growth medium. () 16:0; ( ) 18:0; ( ) 18:1; () 18:2; () 18:3.
1150
A. Converti et al. / Chemical Engineering and Processing 48 (2009) 1146–1151
Fig. 4. Percentages of individual fatty acid methyl ester (FAME) on the total FAMEs (g/100 gFAME ) in N. oculata at different temperatures and concentrations of NaNO3 in the
growth medium. () 16:0; ( ) 18:0; ( ) 18:1; () 18:2; () 18:3.
(7.9% at 20 ◦ C and 14.9% at 15 ◦ C). However, also the growth rate
was significantly affected by changes in temperature, thus leading
a lipid productivity which was approximately the same for all three
growth runs (Table 2).
The reduction of the concentration of nitrate in the growth
medium increased the lipid fraction in both species, despite of
an almost constant growth rate, thereby doubling the productivity of the oil (Tables 3 and 4). The lack of NaNO3 limited the protein
biosynthesis [25] thus increasing the lipid/protein ratio.
The final phase of the research was the qualitative analysis of the
lipids contained in the oily fraction by gas chromatography, in order
to assess their quality for the production of biodiesel according to
the European standards (EN 14214 and EN 14213) [34].
Analysis of fatty acid methyl esters (FAME) produced by the reaction between fatty acids and methanol (transesterification), yielded
very high values of palmitic acid (16:0), which constituted about
60% (mol/mol) of the overall lipid fraction in C. vulgaris (Fig. 3) and
N. oculata (Fig. 4).
In accordance to literature [35] no significant change in the
amount of palmitic acid was observed varying the concentration of
NaNO3 (Figs. 3 and 4). However, the concentration of linolenic acid
(18:3) in C. vulgaris makes the biodiesel derived from this microalga
able to meet the level of European legislation for transportation use
(12%, mol/mol) [34]. On the other hand, that coming from N. oculata
requires additional treatment of catalytic hydrogenation or the use
in mixture with a biodiesel richer in saturated fatty acids.
Finally, it is noteworthy the fact that increasing the temperature over the optimum value lead to increased content of oleic acid,
which was particularly high for C. vulgaris (up to 34% at 38 ◦ C).
4. Conclusions
The method combining chloroform/methanol and ultrasounds
allowed the complete extraction of the microalgae fatty component.
The variation of parameters tested (temperature and concentration of nitrogen) strongly influenced the lipid content of microalgae.
Almost all the stress conditions investigated led not only to the
accumulation of lipids, but also to a reduction in microalgae growth,
thereby affecting the lipids productivity. In particular, whereas the
growth of C. vulgaris was not significantly influenced by temperature, a decrease from 30 to 25 ◦ C brought about lipid content 2.5
times higher. For N. oculata the lipid productivity seems not to
depend on temperature as a result of increased lipid content and
reduced growth.
Of particular interest was the influence of the concentration of
NaNO3 . A doubling of lipid content was in fact observed in both
microalgae when nitrate concentration in the medium was reduced
by 75%, although the growth practically kept unchanged.
The qualitative analysis of fatty acids showed very high values
of palmitic acid (around 60%, mol/mol of the overall lipid fraction)
in both microalgae. The concentration of linolenic acid in C. vulgaris
met the requirement of European legislation for biodiesel, whereas
the higher content of this acid in N. oculata requires an additional
treatment to this purpose.
As far as the possible developments of this research are concerned, the next effort will deal with the growth of both microalgae
in appropriate and efficient photobioreactors under the stress conditions identified in this study, and varying the light intensity.
Acknowledgements
The authors are grateful to the Italian Ministry of the Instruction,
University and Research (MIUR) (FIRB, Prot. N. RBIP06993E) for the
financial support of this work.
References
[1] J.R. Benemann, CO2 mitigation with microalgae systems, Energy Convers. Manage. 38 (1997) 475–479.
[2] X. Miao, Q. Wu, Biodiesel production from heterotrophic microalgal oil, Bioresour. Technol. 97 (2006) 841–846.
[3] M.I. Orús, E. Marco, F. Martínez, Suitability of Chlorella vulgaris UAM 101 for
heterotrophic biomass production, Bioresour. Technol. 38 (1991) 179–184.
[4] R. Muñoz, B. Guieysse, Algal–bacterial processes for the treatment of hazardous
contaminants: a review, Water Res. 40 (2006) 2799–2815.
[5] P. Spolaore, C. Joannis-Cassan, E. Duran, A. Isambert, Commercial applications
of microalgae, J. Biosci. Bioeng. 101 (2006) 87–96.
[6] Y. Chisti, Biodiesel from microalgae, Biotechnol. Adv. 25 (2007) 294–306.
[7] K.H.M. Cardozo, T. Guaratini, M.P. Barros, V.R. Falcão, A.P. Tonon, N.P. Lopes, S.
Campos, M.A. Torres, A.O. Souza, P. Colepicolo, E. Pinto, Metabolites from algae
with economical impact, Comp. Biochem. Physiol. C: Toxicol. Pharmacol. 146
(2007) 60–78.
[8] I. Valencia, D. Ansorena, I. Astiasarán, Development of dry fermented sausages
rich in docosahexaenoic acid with oil from the microalgae Schizochytrium sp.:
influence on nutritional properties, sensorial quality and oxidation stability,
Food Chem. 104 (2007) 1087–1096.
A. Converti et al. / Chemical Engineering and Processing 48 (2009) 1146–1151
[9] L.J. Borowitzka, Development of Western Biotechnology’s algal ␤-carotene
plant, Bioresour. Technol. 38 (1991) 251–252.
[10] T. Furuki, S. Maeda, S. Imajo, T. Hiroi, T. Amaya, T. Hirokawa, K. Ito,
H. Nozawa, Rapid and selective extraction of phycocyanin from Spirulina
platensis with ultrasonic cell disruption, J. Appl. Phycol. 15 (2003) 319–
324.
[11] M. Olaizola, Commercial development of microalgal biotechnology: from the
test tube to the marketplace, Biomol. Eng. 20 (2003) 459–466.
[12] E. Baker, J.J.A. McLaughlin, S.S.H. Hutner, B. DeAngelis, S. Feingold, O. Frank,
H. Baker, Water-soluble vitamins in cells and spent culture supernatants of
Poteriochromonas stipitata, Euglena gracilis, and Tetrahymena thermophila, Arch.
Microbiol. 129 (1981) 310–313.
[13] S.A. Survase, I.B. Bajaj, R.S. Singhal, Biotechnological production of vitamins,
Food Technol. Biotechnol. 44 (2006) 381–396.
[14] J.A. Running, D.K. Severson, K.J. Schneider, Extracellular production of l-ascorbic
acid by Chlorella protothecoides, Prototheca species, and mutants of P. moriformis
during aerobic culturing at low pH, J. Ind. Microbiol. Biotechnol. 29 (2002)
93–98.
[15] C. Bremus, U. Herrmann, S. Bringer Meyera, H. Sahm, The use of microorganisms
in l-ascorbic acid production, J. Biotechnol. 124 (2006) 196–205.
[16] F.A. Muller, The role of microalgae in aquaculture: situation and trends, J. Appl.
Phycol. 12 (2000) 527–534.
[17] A. Sánchez Mirón, M.C. Cerón García, A. Contreras Gómez, F. García Camacho,
E. Molina Grima, Y. Chisti, Shear stress tolerance and biochemical characterization of Phaeodactylum tricornutum in quasi steady-state continuous culture in
outdoor photobioreactors, Biochem. Eng. J. 16 (2003) 287–297.
[18] S. Sawayama, CO2 fixation and oil production through microalga, Fuel. Energ.
Abstr. 37 (1996) 217.
[19] Y.S. Yun, S.B. Lee, J.M. Park, C.I. Lee, J.W. Yang, Carbon dioxide fixation by algal
cultivation using wastewater nutrients, J. Chem. Technol. Biotechnol. 69 (1997)
451–455.
[20] F.B. Metting, Biodiversity and application of microalgae, J. Ind. Microbiol.
Biotechnol. 17 (1996) 477–489.
[21] A.M. Illman, A.H. Scragg, S.W. Shales, Increase in Chlorella strains calorific values when grown in low nitrogen medium, Enzyme Microb. Technol. 27 (2000)
631–635.
[22] Z.Y. Liu, G.C. Wang, B.C. Zhou, Effect of iron on growth and lipid accumulation
in Chlorella vulgaris, Bioresour. Technol. 99 (2008) 4717–4722.
1151
[23] Y. Li, M. Horsman, N. Wu, C.Q. Lan, N. Dubois Calero, Biofuels from microalgae,
Biotechnol. Progr. 24 (2008) 815–820.
[24] R.R.L. Guillard, J.H. Ryther, Studies of marine planktonic diatoms. I. Cyclotella
nana Hustedt and Detonula confervacea Cleve, Can. J. Microbiol. 8 (1962)
229–239.
[25] R.R.L. Guillard, Culture of phytoplankton for feeding marine invertebrates,
in: W.L. Smith, M.H. Chanley (Eds.), Culture of Marine Invertebrate Animals,
Plenum Press, New York, 1975, pp. 26–60.
[26] E. Huertas, O. Montero, L.M. Lubián, Effects of dissolved inorganic carbon availability on growth, nutrient uptake and chlorophyll fluorescence of two species
of marine microalgae, Aquacult. Eng. 22 (2000) 181–197.
[27] J. Folch, M. Lees, G.H. Sloane Stanley, A simple method for the isolation and
purification of total lipids from animal tissues, J. Biol. Chem. 226 (1957)
497–509.
[28] M.R. Brown, M.A. McCausland, K. Kowalski, The nutritional value of four Australian microalgal strains fed to Pacific oyster Crassostrea gigas spat, Aquaculture
165 (1998) 281–293.
[29] J.-P. Hernandez, L.E. de-Bashan, D.J. Rodriguez, Y. Rodriguez, Y. Bashan, Growth
promotion of the freshwater microalga Chlorella vulgaris by the nitrogen-fixing,
plant growth-promoting bacterium Bacillus pumilus from arid zone soils, Eur. J.
Soil Biol. 45 (2009) 88–93.
[30] R.J. Maxwell, Determination of total lipid and lipid subclasses in meat and meat
products, J. Assoc. Off. Anal. Chem. 70 (1987) 74–77.
[31] L. Krienitz, M. Wirth, The high content of polyunsaturated fatty acids in Nannochloropsis limnetica (Eustigmatophyceae) and its implication for food web
interactions, freshwater aquaculture and biotechnology, Limnologica 36 (2006)
204–210.
[32] G. Cravotto, L. Boffa, S. Mantegna, P. Perego, M. Avogadro, P. Cintas,
Improved extraction of vegetable oils under high-intensity ultrasound and/or
microwaves, Ultrason. Sonochem. 15 (2008) 898–902.
[33] P. Zunin, P. Salvadeo, R. Boggia, F. Evangelisti, Sterol oxidation in meat- and fishbased homogenized baby foods containing vegetable oils, J. AOAC Int. 89 (2006)
441–446.
[34] G. Knothe, Analyzing biodiesel: standards and other methods, J. Am. Oil Chem.
Soc. 83 (2006) 823–833.
[35] M. Piorreck, K.H. Baasch, P. Pohl, Biomass production, total protein, chlorophylls,
lipids and fatty acids of freshwater green and blue-green algae under different
nitrogen regimes, Phytochemistry 23 (1984) 217–223.