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Lipid production ofChlorella vulgariscultured in artificial wastewater medium
Yujie Feng a,*, Chao Li a, Dawei Zhang b
a State Key Laboratory of Urban Water Resource and Environment, Harbin Institute of Technology, No 73 Huanghe Road, Nangang District, Harbin 150090, PR Chinab Department of Environmental Science & Engineering, Harbin Institute of Technology (Weihai), Weihai 264200, China
a r t i c l e i n f o
Article history:
Received 31 March 2010
Received in revised form 1 June 2010
Accepted 2 June 2010
Keywords:
Chlorella vulgaris
Lipid content
Artificial wastewater
Energy and cost analyses
a b s t r a c t
Chlorella vulgariswas used to study algal lipid production with wastewater treatment. Artificial wastewa-
ter was used to cultivate C. vulgaris in a column aeration photobioreactor (CAP) under batch and semi-continuous cultivation with various daily culture replacements (0.5 l1.5 l per 2 l reactor). The cell den-
sity was decreased from 0.89 g/l with the daily replacement of 0.5 l to 0.28 g/l with 1.5 l replacement.
However, C. vulgaris culture achieved the highest lipid content (42%, average value of the phase) and
the lipid productivity (147 mg/l d1) with daily replacement of 1.0 l. And then the nutrient removal effi-
ciency were 86% (COD), 97% (NH4 ) and 96% (TP), respectively. Analyses of energy efficiency showed that
the net energy ratio (NER) for lipid production with daily replacement of 1.0 l (1.25) was higher than the
other volume replacement protocols. And cost analyses showed that the algal biomass can be competitive
with petroleum at US$ 63.97 per barrel with the potential credit for wastewater treatment. According to
the above results, it is concluded that the present research will lead to an economical technology of algal
lipid production.
2010 Elsevier Ltd. All rights reserved.
1. Introduction
Global demand for food is expected to double within 50 years,
and the demand for transportation fuels is expected to increase
even more rapidly (Hill et al., 2006). Diversion of food crops to bio-
fuels would not be right approach to solve the problems because
they compete with food production for high-grade arable land
(Rittmann, 2008). There is a great need for renewable energy sup-
plies that do not cause significant environmental harm and not
competed with food supply. Because of their higher photosynthetic
efficiency, higher biomass production and faster growth compared
with other energy crops, microalgae have been receiving attentions
as candidates for fuel production (Minowa et al., 1995). Microalgae
can be used to produce various forms of biofuel including biodiesel
(Converti et al., 2009; Gao et al., 2010), ethanol (Shirai et al., 1998),
bioelectricity (Powell et al., 2009), hydrogen (Ghirardi, 2006;
Hemschemeier et al., 2009), and methane (Stucki et al., 2009).
Biodiesel is produced from plant oils or animal fats, and biodie-
sel industries are expanding rapidly both in the United States and
in Europe with soybean or rapeseed oils as the feedstock. However,
the potential market for biodiesel far surpasses the availability of
plant oils, waste cooking oil and animal fats. Therefore, microalgae
have been studied as alternative feedstock for biodiesel production
recently. Use of microalgae to produce biodiesel would not com-
promise production of food, fodder and other products derived
from crops (Chisti, 2007). Many microalgae accumulate lipids as
storage materials and their accumulation is stimulated under envi-
ronment stress, such as nutrient deficiency (Dunahay et al., 1996)
or salt stress (Takagi et al., 2006).Widjaja et al. (2009) reported
that maximum lipid content ofChlorella vulgariswas only 26% un-
der normal nutrition medium with nitrogen (NaNO3) content of
70.02 mg/l. However, after normal nutrition cultivation, the med-
ium was changed into nitrogen depletion(0.02 mg/l) continued
for 7 d and 17 d, and the lipid contents were 36% and 43%, respec-
tively. Furthermore, according to the results obtained by Converti
et al. (2009), a threefold increase (from 5.9% to 15.3%) in lipid con-
tent took place with NaNO3 concentration decrease from 1.5 to
0.375 g/l.Hsieh et al. (2009)used urea as the nitrogen source at
concentrations of 0.025, 0.050, 0.100, 0.150, and 0.200 g/l. After
6 days of cultivation, the lipid contents of Chlorella sp. were 66%,
60%, 52%, 37%, and 33% respectively.
Microalgal biomass can be produced through autotrophic culti-
vation in open ponds or photobioreactors by using solar energy
and fixing carbon dioxide. Alternatively they are cultivated hetero-
trophically or mixotrophically using organic compounds as energy
and carbon sources. Due to the reduction in light penetration
(Chaumont, 1993) in autotrophic culture, the cell density is usually
less than 1 g/l (Borowitzka, 1994). So far as we know, there is
no effective cultivated method to increase cell density in the
autotrophic cultivation processes. Therefore, the downstream
0960-8524/$ - see front matter 2010 Elsevier Ltd. All rights reserved.doi:10.1016/j.biortech.2010.06.016
* Corresponding author. Tel.: +86 451 86283068; mobile: +13069891017; fax:
+86 451 87162150.
E-mail address:[email protected](Y. Feng).
Bioresource Technology 102 (2011) 101105
Contents lists available at ScienceDirect
Bioresource Technology
j o u r n a l h o m e p a g e : w w w . e l s e v i e r . c o m / l o c a t e / b i o r t e c h
http://dx.doi.org/10.1016/j.biortech.2010.06.016mailto:[email protected]://dx.doi.org/10.1016/j.biortech.2010.06.016http://www.sciencedirect.com/science/journal/09608524http://www.elsevier.com/locate/biortechhttp://www.elsevier.com/locate/biortechhttp://www.sciencedirect.com/science/journal/09608524http://dx.doi.org/10.1016/j.biortech.2010.06.016mailto:[email protected]://dx.doi.org/10.1016/j.biortech.2010.06.0168/12/2019 1-s2.0-S096085241000996X-main
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processing costs are relatively high (Shi et al., 1997). For the het-
erotrophic or mixotrophic cultivation, organic carbon compounds
such as glucose are responsible for higher production costs.
Glucose used in this process comprises about 80% of the total costs
(Li et al., 2007).
In this work C. vulgaris was used to produce algal lipid using
wastewater as medium. In order to simply investigate the factors
related with algal lipid production during wastewater treatment,
preliminary results were obtained here using synthetic wastewater
instead of real wastewater. The use of wastewater as feedstock for
algal lipid is economically attractive since the production costs can
be reduced with credits for wastewater treatment as well as with
reduction in the greenhouse gas emission.
2. Methods
2.1. Algal strain and culture medium
C. vulgaris (FACHB1068) was purchased from Freshwater Algae
Culture Collection, Institute of Hydrobiology, Chinese Academy of
Sciences (Wuhan, China). The strain was preserved in the BG11
medium containing following chemicals: NaNO3 (1.5 g/l),K2HPO43H2O (0.04 g/l), MgSO47H2O (0.075g/l), CaCl22H2O
(0.036 g/l), Na2CO3 (0.02 g/l), citric acid (0.006 g/l), Ferric ammo-
nium citrate (0.006 g/l), EDTA (0.001 g/l), and A5 + Co solution
(1 ml/l) that consists of H3BO3 (2.86 g/l), MnCl2H2O (1.81 g/l),
ZnSO47H2O (0.222 g/l), CuSO45H2O (0.079 g/l), Na2MoO42H2O
(0.390 g/l) and Co(NO3)26H2O (0.049 g/l). C. vulgaris was inocu-
lated at 20% (v/v) in 250 ml Erlenmeyer flasks containing 100 ml
BG11 medium. The flasks were incubated under stationary condi-
tion at 30 C with 3000 lx continuous cool-white fluorescent light
illumination, and were hand shaken three to five times daily to
avoid sticking. The algal cells which just reached the stationary
phase were used to inoculate the column aeration photobioreactor
(CAP).
2.2. Artificial wastewater
The artificial wastewater was prepared dissolving following
chemicals; glucose (0.4125 g/l), NH4Cl (0.078 g/l), KH2PO4(0.018 g/l), MgSO47H2O (0.013 g/l), CaCl22H2O (0.043 g/l), FeS-
O7H2O (0.005 g/l), and A5 + Co solution (1 ml/l). The initial pH
was adjusted to 7.08.0 and sterilized at 121 C for 20 min before
inoculation. The initial N-NH4+, total phosphate (TP), and COD con-
centration were 20, 4, and 400 mg/l, respectively.
2.3. Reactor design and its operation
Four 2.2 l CAPs were consructed with 2 l effective volume
(10 cm diameter and 25 cm height) using polymethyl methacrylate(PMMA). The CAPs containing 1.5 l sterilized artificial wastewater
were inoculated with 0.5 l flask culture ofC. vulgaris. The culture
pH decreased due to NH4+ assimilation, which was observed in
our previous study (data not shown). Therefore, the pH of culture
was maintained between 8 and 10 during Day 2 to 14. All the
experiments were carried out at 30 C and 3000 lx continuous
cool-white fluorescent light illumination. The reactors were aer-
ated with sterilized air at 0.5 vvm (volumes of air per total volume
of bioreactor per minute) to provide mixing and CO2, as well as O2to the algae.
Before the cultures reach stationary phase various volume was
replaced daily with fresh medium to operate the reactors in the
semi-continuous mode. When the cell density reached about
0.8 g/l on Day 4 after the inoculation, the culture was operated inthe first phase of semi-continuous cultivation for 3 d by replacing
0.5 l of the culture with fresh artificial wastewater everyday. After
that, the culture was operated in the second and third phases of
semi-continuous mode by replacing 1.0 and 1.5 l of the culture
with fresh medium everyday, respectively. Both these phases were
maintained for 4 days each. The batch culture under the same con-
dition except for medium replacement was used as positive con-
trol. It was operated for 14 days. All the experiments were
carried out in duplicate and average values are reported.
2.4. Lipid extraction
Algal cells were harvested by centrifugation at 10,000 rpm, 4 C
for 10 min. Supernatant was decanted and cell pellets were washed
with distilled water and then freeze-dried under 80 C. Thereaf-
ter, the total lipids were extracted from microalgal biomass using
a modified method ofBligh and Dyer (1959). Fifty mg of lyophi-
lized microalgal biomass was placed into a 15 ml test tube and
1.6 ml water, 4.0 ml methanol and 2.0 ml chloroform were added.
The solution was mixed for 30 s. Thereafter, an additional 2.0 ml of
chloroform and 2.0 ml water were added and the content of the
test tube was mixed for 30 s. The test tubes were centrifuged at
5000 rpm for 10 min. The upper layer was withdrawn by using apipette and the lower chloroform phase containing the extracted
lipids was transferred into a 30-ml culture tube. The solid material
left at the bottom of extraction tube was extracted with the same
procedure two more times and the chloroform phases were mixed
together and then evaporated in a nitrogen evaporator until
obtaining dry lipid. Thereafter, the total lipids were measured
gravimetrically, and then lipid content and lipid yields were
calculated.
2.5. Analyses
Samples were taken from CAPs each day for analyses. Optical
density (OD) of the algae culture at 658 nm was measured daily
as the cell density indicator using a spectrophotometer (752 Grat-
ing Spectrophotometer, Shandong Gaomi Caihong Analytical
Instrument Factory, China). A linear relationship between OD658and dry weight (DW, g/L) of algal biomass was determined previ-
ously for this strain:
Dry weight g=l 0:4818 OD658; R2 0:9962 1
Samples were centrifuged at 10,000 rpm for 10 min to determine
NH4 , TP and COD concentrations in the supernatants. COD was
determined by a Multi-Function Reactor (ET3150B Multi-Function
Reactor, Euro Tech, China), Nash reagent photometry was used for
measuring NH4 concentration. TP was determined by molybde-
numantimony anti-spectrophotometric method.
For the energy analysis, the values of lipid content, cell den-sity, and hydraulic retention time were obtained from the results
of the semi-continuous cultivation. According to Jorqueras
(2010) research, a production scale of 100 ton of algal biomass
per year was set as the basis to calculate energy balance. And
the energy consumption term included only the energy required
for air pumping that was used to maintain appropriate culture
mixing and liquid/gas mass transfer. Thereafter, volumetric pro-
ductivity, reactor volume required for a biomass production of
100 ton/year, net lipid yield, energy consumption required for a
biomass production of 100 ton/year, total energy consumption,
energy produced as lipid, and NER for lipid production were cal-
culated. Because the structure and the operational mechanism of
CAP were similar to flat-plate photobioreactor, the energy con-
sumption of CAP was assumed equal to flat-plate photobioreactor(53 W/m3).
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3. Results and discussion
3.1. Algal growth and lipid content
CAPs inoculated by the test organism were operated for 14 days
in a batch mode monitoring algal growth and lipid content (Fig. 1).
As shown inFig. 1, the alga grew up to 7 days of cultivation to the
cell density of 1.581.72 g/l with a lag phase of one day. On theother hand the lipid content increased to 37% from 15% in Day 2,
and decreased to 7.8% during the growth phase. However, the lipid
content increased to 34.1% in day 10 but did not increase further at
the stationary phase.
The high lipid content at Day 2 is probably because the
culture was under heterotrophic/mixotrophic conditions. Organic
carbon in the culture was quickly consumed and was exhausted
on Day 2 (see Section 3.2), and then C. vulgaris switched from
mixotrophic metabolism to autotrophic metabolism. It had been
reported (Miao and Wu, 2004) that the autotrophic microalgae
had low lipid content in comparison with those under heterotro-
phic and mixotrophic conditions. Therefore, the switch of metab-
olism on Day 2 resulted in the significantly decrease of lipid
content.
The lipid content in the stationary phase was up to 37%.Li et al.
(2008)reported that the nitrogen deficiency would result in more
metabolic flux generated from photosynthesis to be turned to lipid
accumulation in Neochloris oleoabundans. The reason may be that
under nitrogen deficiency or limitations the synthetic rate of
essential cell structures including proteins and nucleic acids be-
comes low. Therefore, the major part of carbon fixed is converted
into carbohydrate or lipid (Richardson et al., 1969). The artificial
wastewater used in these experiments contains only 20 mg/l nitro-
gen in form of N NH4 . Although N NH4 was depleted on Day 3
(seeFig. 2c), the growth rate of algal cell was not limited signifi-
cantly. The reason may be that nitrate has been introduced to
the culture at inoculation (The initial N NO3 concentration in
BG11 medium is 247 mg/l). Only ammonium is utilized when cul-
tures containing both nitrate and ammonium (Ahmad and Helle-bust, 1990). Therefore, ammonium was depleted first in the
culture, and then algal cell grew with nitrate as nitrogen source un-
til nitrate was depleted. Thus, the lipid accumulation was not en-
hanced up due to most metabolic flux generated from
photosynthesis was still used for cell synthesis from Day 3 to 8.
After Day 8, the growth of algae was nearly ceased, thus resulting
in the increased lipid synthesis. The relatively high lipid content at
Day 1 is believed due to the fact that cells in this phase are similar
to those in the stationary phase culture in BG11 medium which
was used as the inoculums.
CAPs was operated in semi-continuous mode at Day 4 by
replacing different volume of the culture with fresh medium every
day; 0.5 l (first phase), 1.0 l (second phase) and 1.5 l (third phase).
The cell density and lipid content were determined, and the lipid
productivity was calculated based on the results. As shown in
Table 1, cell density decreased from 0.89 g/l in the first phase to
0.28 g/l in the third phase with the increase in daily changed cul-
ture volume during the semi-continuous cultivation. As for the li-
pid content, it increased significantly from 20% in the first phase
to 42% in the second phase, and then decreased slightly to 38% in
the third phase. According to the results of cell density and lipid
content, lipid productivity was calculated. The highest lipid pro-
ductivity (147 mg/l d1) was achieved during the second phase
compared with the first (44 mg/l d1) and the third (79 mg/l d1)
phases.
The main reason for the reduction of cell density is due to the
reduced algal cell retention time in the reactor with the increaseddaily changed volume. As discussed earlier, the lipid content is
dependent on the nitrogen limitations and on the trophic condi-
tions. Higher lipid content is expected in the cells facing nitrogen
limitation under mixotrophic conditions. In a semi-continuous
0
0.4
0.8
1.2
1.6
2
0 2 4 6 8 10 12 14 16
Days of cultivation
Celldensity(g/l)
0
10
20
30
40
Lipidcontent(%)
cell density
lipid content
Fig. 1. Cell density and lipid content ofC. vulgaris culture in the batch cultivation.
0
100
200
300
400
0 2 4 6 8 10 12 14 16Days of cultivation
C
ODconcentrations
(mg/l)
0
20
40
60
80
100
Removalefficiency(%)
semi-continuous
batch
removal efficiency in semi-continuous
0
1
2
3
4
0 2 4 6 8 10 12 14 16
Days of cultivation
TPconcentrations
(mgP/l)
0
20
40
60
80
100
Removalefficiency(%)
semi-continuous
batch
removal efficiency
in semi-continuous
0
5
10
15
20
25
0 2 4 6 8 10 12 14 16Days of cultivation
NH4+concentrations
(mgN/l)
0
20
40
60
80
100
Removalefficiency(%)
semi-continuous
batch
removal efficiency
in semi-continuous
A
B
C
Fig. 2. Removal efficiency and mean concentration of nutrients for C. vulgaris
growing in the batch and semi-continuous cultivation. (A) COD; (B) TP; (C) NH4 .
Table 1
Cell density and lipid content ofC. vulgarisin different phases of the semi-continuous
cultivation.
First Second Third
Daily change medium (l/2 l reactor) 0.5 1.0 1.5
Cell density (g/l) 0.89 0.69 0.28
Lipid content (%) 20 42 38
Lipid productivity (mg/l d1) 44 147 79
*Cell density, lipid content and productivity were average value in the phase.
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culture system the nitrogen availability and trophic conditions are
determined by the volume of daily changed medium. The lipid
content was low in the first phase with low volume change. This
is believed due to the fact that organic carbon was not enough to
supportC. vulgarisgrown in mixotrophic metabolism for long time
to maintain a high lipid synthesis rate. And nitrate may have been
introduced to the culture at inoculation, thus resulting in abundant
nitrogen in culture in the first phase. On the other hand the highest
lipid content of 42% was achieved during the second phase of the
semi-continuous cultivation. This result suggests that the culture
was supplied with enough organic carbon to maintain mixotrophic
metabolism with nitrogen limitation. However, further increase in
daily changed volume during the third phase caused a slight
decrease in lipid content (38%). These results suggest that the
culture had abundant nutrient during the third phase with high
changed volume, therefore, the algae grew vigorously and more
assimilated organic carbon was used for cell growth. Thus, the lipid
content obtained a slight decrease.
3.2. Nutrients removal efficiency
The culture supernatant was analyzed for COD, TP, and NH4+ to
determine the process performance of the system (Fig. 2). As ex-
pected the nutrients removal efficiency was poor at the beginning
of the reactor operation (Day 01) due to lowcell density (0.05 g/l).
It is expected that the higher the cell density, the better the nutri-
ent removal efficiency (Lau et al., 1995). Thereafter, the removal
efficiency of nutrient achieved higher level during the growth
phase, due to the higher cell density and vigorous growth. On
Day 2 the removal efficiencies of COD, TP, and NH4+ were 87%,
94%, and 90%, respectively. It was interesting to note that COD in
the supernatant prepared from the batch cultivation was higher
than those from the semi-continuous cultivation during Days 2
to 14, although the higher cell density and longer HRT in batch
cultivation. The possible reason for this result is thatC. vulgaris
secretes extra cellular substances during the growth process (Babel
et al., 2002; Paralkar and Edzwald, 1996), which was hardlydegradable by the alga. In the semi-continuous cultivation, the
extra cellular substances were removed by medium replacement.
Therefore, the removal efficiency of COD in semi-continuous
cultivation increased from 85% to 88% along with the increasing
daily changed medium. As for TP and NH4+, the batch cultivation
had good removal efficiency which was 96% and 97%, respectively.
The removal efficiency of TP in the third phase of semi-continuous
cultivation was 92% that was lower than that of batch cultivation.
This might be due to low cell growth. The removal efficiency of
NH4 was high (97%) in the whole semi-continuous processes.
3.3. Energy and cost analysis
The net energy ratio (NER) for lipid production was defined asthe ratio of the energy produced as lipid over the total energy con-
sumption. As shown in Table 2, the second phase not only achieved
the highest value of energy produced as lipid, but also the lowest
total energy consumption among the three phases. Therefore, the
NER for lipid production in the second phase (1.25) was higher
than the others. It suggests that the semi-continuous in the second
phase was the most efficiency energy production system among
others.
A large part of the production cost of algal lipid is downstream
processing costs including cell harvest and lipid extraction costs
that are dependent considerably on the cell density and lipid con-
tent (Li et al., 2008). High cell density reduces the cell harvesting
cost, so does high lipid content to lipid extraction cost. Therefore,
the downstream processing costs of algal lipid can be calculatedbased on the following equation:
z xDy=DL 2
where z, downstream processing cost of algal lipid ($/g); x, algae
harvesting cost ($/l), y, lipid extraction cost ($/g); D, cell density
(g/l);L , lipid content (%). It was assumed that the harvesting cost
is dependent on the volume of cultures, and that the extraction cost
dependent on the weight of algae.
Cell density and lipid content of each phase were used to calcu-
late the downstream processing cost according to Equation (2). The
downstream processing costs of algal lipid were 5.6x+ 5y,
3.4x+ 2.4y, and 9.5x+ 2.6y for the first, second, and third phases,
respectively. Hence, the downstream processing of the secondphase was the lowest among three phases.
According to Chistis (2008) research, the algal biomass (lipid
content of 42%) with the production costs of US$ 217.22/ton be-
comes competitive with petroleum at US$ 60.00 per barrel. Using
a value of US$ 0.22/kWh for the energy consumption, the produc-
tion cost of 1 ton algal biomass in the second phase was estimated
to be US$ 808.79. To produce 1 ton algal biomass, 1443 m3 of
wastewater is treated. If the credit for wastewater treatment at
US$ 0.4/m3 is counted, the price of 1 ton of biomass would be re-
duced to US$ 231.59. This figure shows that algal biomass can be
competitive when supposing the price of petroleum is US$ 63.97
per barrel. However, since the energy and cost analyses were car-
ried out based on synthetic wastewater, the results could be differ-
ent when using different real wastewater.
4. Conclusions
The results in this study showed that microalgae cultivation
with wastewater as medium is a promising method to produce al-
gal lipid. The highest lipid content (42%) and productivity (147 mg/
l d1) were achieved in the semi-continuous cultivation with daily
replacement of 1.0 l of the 2.0 l culture. And then the nutrient re-
moval efficiencies were 86% (COD), 97% NH4 and 96% (TP),
respectively. These results were used to analyze the energy effi-
ciency. The NER for lipid production (1.25) was greater than unity.
And cost analysis exhibited that the algal biomass can be compet-
itive with petroleum at US$ 63.97 per barrel with the potential
credit for wastewater treatment. Furthermore, this process also re-duces the greenhouse gas emission in wastewater treatment.
Table 2
Comparative analyses of algal biomass and lipid production in different phases of the
semi-continuous cultivation.
Variable First Second Third
Cell density (g/l) 0.89 0.69 0.28
Hydraulic retention time (d) 4 2 1.3
Volumetric productivity (g/l d1) 0.223 0.346 0.21
Reactor volume required for a biomass
production of 100 ton/yeara (m3)
1246 803 1323
Lipid content (%) 20 42 38
Net lipid yieldb (m3/year) 22 47 42
Energy consumption (W/m3) 53 53 53
Energy consumption required for a biomass
production of 100 ton/yearc (W)
66,038 42,550 70,119
Total energy consumptiond (GJ/year) 2054.05 1323.48 2180.99
Energy produced as lipide (GJ/year) 772.93 1651.27 1475.60
NER for lipid production 0.38 1.25 0.68
a Determined by dividing the annual biomass production by the volumetric
productivity.b Determined by dividing the product of annual biomass and lipid content by the
density of lipid (assumed to be 0.9 kg/l).c Determined by multiplying the energy consumption by the reactor volume
required.d Determined by multiplying the energy consumption by the number of hours of
air pumping (it was 24 h of one day).e Determined by multiplying the net lipid yield by energy content of lipid
(assumed value of 35, 133.33 kJ/l).
104 Y. Feng et al. / Bioresource Technology 102 (2011) 101105
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Acknowledgements
The research is supported by the Scientific Research Foundation
for the Returned Overseas Chinese Scholars, State Education Minis-
try, China. The authors also acknowledge the support of the Na-
tional Creative Research Groups of China (50821002) and the
technical and financial support of the State Key Laboratory of Ur-
ban Water Resource and Environment (2010TS08), HIT, China. Prof.Byung Hong Kim (Water Environment and Remediation Research
Center, Korea Institute of Science and Technology) is gratefully
thanked for his efforts on this paper.
References
Ahmad, I., Hellebust, J., 1990. Regulation of chloroplast development by nitrogen
source and growth conditions in a Chlorella protothecoides strain. PlantPhysiology 94 (3), 944949.
Babel, S., Takizawa, S., Ozaki, H., 2002. Factors affecting seasonal variation of
membrane filtration resistance caused by Chlorella algae. Water Research 36
(5), 11931202.
Bligh, E.G., Dyer, W.J., 1959. A rapid method of total lipid extraction and
purification. Canadian Journal of Biochemistry and Physiology 37 (8), 911917.
Borowitzka, M.A., 1994. Large-scale algal culture systems: the next generation.
Australasian Biotechnology 4 (4), 212215.
Chaumont, D., 1993. Biotechnology of algal biomass production a review ofsystems for outdoor mass-culture. Journal of Applied Phycology 5 (6), 593604.
Chisti, Y., 2007. Biodiesel from microalgae. Biotechnology Advances 25 (3), 294
306.
Chisti, Y., 2008. Biodiesel from microalgae beats bioethanol. Trends in
Biotechnology 26 (3), 126131.
Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., Del Borghi, M., 2009. Effect of
temperature and nitrogen concentration on the growth and lipid content of
Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. ChemicalEngineering and Processing 48 (6), 11461151.
Dunahay, T.G., Jarvis, E.E., Dais, S.S., Roessler, P.G., 1996. Manipulation of microalgal
lipid production using genetic engineering. Applied Biochemistry and
Biotechnology 5758, 223231.
Gao, C.F., Zhai, Y., Ding, Y., Wu, Q.Y., 2010. Application of sweet sorghum for
biodiesel production by heterotrophic microalga Chlorella protothecoides.Applied Energy 87 (3), 756761.
Ghirardi, M.L., 2006. Hydrogen production by photosynthetic green algae. Indian
Journal of Biochemistry and Biophysics 43 (4), 201210.
Hemschemeier, A., Melis, A., Happe, T., 2009. Analytical approaches to
photobiological hydrogen production in unicellular green algae.Photosynthesis Research 102 (23), 523540.
Hill, J., Nelson, E., Tilman, D., Polasky, S., Tiffany, D., 2006. Environmental, economic,
and energetic costs and benefits of biodiesel and ethanol biofuels. Proceedings
of the National Academy of Sciences of the United States of America 103 (30),
1120611210.
Hsieh, C.H., Wu, W.T., 2009. Cultivation of microalgae for oil production with a
cultivation strategy of urea limitation. Bioresource Technology 100 (17), 3921
3926.
Jorquera, O., Kiperstok, A., Sales, E.A., Embirucu, M., Ghirardi, M.L., 2010.
Comparative energy life-cycle analyses of microalgal biomass production in
open ponds and photobioreactors. Bioresource Technology 101 (4), 14061413.
Lau, P.S., Tam, N.F.Y., Wong, Y.S., 1995. Effect of algal density on nutrient removalfrom primary settled waste-water. Environmental Pollution 89 (1), 5966.
Li, X.F., Xu, H., Wu, Q.Y., 2007. Large-scale biodiesel production from microalga
Chlorella protothecoides through heterotropic cultivation in bioreactors.Biotechnology and Bioengineering 98 (4), 764771.
Li, Y.Q., Horsman, M., Wang, B., Wu, N., Lan, C.Q., 2008. Effects of nitrogen sources
on cell growth and lipid accumulation of green alga Neochloris oleoabundans.Applied Microbiology and Biotechnology 81 (4), 629636.
Miao, X.L., Wu, Q.Y., 2004. High yield bio-oil production from fast pyrolysis by
metabolic controlling ofChlorella protothecoides. Journal of Biotechnology 110(1), 8593.
Minowa, T., Yokoyama, S., Kishimoto, M., Okakura, T., 1995. Oil production from
algal cells ofDunaliella tertiolecta by direct thermochemical liquefaction. Fuel74 (12), 17351738.
Paralkar, A., Edzwald, J.K., 1996. Effect of ozone on EOM and coagulation. Journal
American Water Works Association 88 (4), 143154.
Powell, E.E., Mapiour, M.L., Evitts, R.W., Hill, G.A., 2009. Growth kinetics ofChlorellavulgarisandits use as a cathodic half cell. Bioresource Technology 100(1), 269274.
Richardson, B., Orcutt, D.M., Schwertner, H.A., Martinez, C.L., Wickline, H.E., 1969.
Effects of nitrogen limitation on the growth and composition of unicellular
algae in continuous culture. Applied Microbiology 18 (2), 245250.
Rittmann, B.E., 2008. Opportunities for renewable bioenergy using microorganisms.
Biotechnology and Bioengineering 100 (2), 203212.
Shi, X.M., Chen, F., Yuan, J.P., Chen, H., 1997. Heterotrophic production of lutein by
selected Chlorella strains. Journal of Applied Phycology 9 (5), 445450.
Shirai, F., Kunii, K., Sato, C., Teramoto, Y., Mizuki, E., Murao, S., Nakayama, S., 1998.
Cultivation of microalgae in thesolution fromthe desaltingprocess of soysauce
waste treatment and utilization of the algal biomass for ethanol fermentation.
World Journal of Microbiology and Biotechnology 14 (6), 839842.
Stucki, S., Vogel, F., Ludwig, C., Haiduc, A.G., Brandenberger, M., 2009. Catalytic
gasification of algae in supercritical water for biofuel production and carbon
capture. Energy and Environmental Science 2 (5), 535541.
Takagi, M., Karseno, Yoshida, T., 2006. Effect of salt concentration on intracellular
accumulation of lipids and triacylglyceride in marine microalgae Dunaliella
cells. Journal of Bioscience and Bioengineering 101 (3), 223226.
Widjaja, A., Chien, C.C., Ju, Y.H., 2009. Study of increasing lipid production from
fresh water microalgae Chlorella vulgaris. Journal of the Taiwan Institute ofChemical Engineers 40 (1), 1320.
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