Bioresource Technology 121 (2012) 298–303
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Bioresource Technology
journal homepage: www.elsevier.com/locate/biortech
Using ammonia for algae harvesting and as nutrient in subsequent cultures
Fangjian Chen, Zhiyong Liu, Demao Li, Chenfeng Liu, Ping Zheng, Shulin Chen ⇑
Tianjin Key Laboratory for Industrial Biological Systems and Bioprocessing Engineering, Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences, Tianjin, China
h i g h l i g h t s
" We present a novel ﬂocculation process using ammonia as coagulant.
" The novel process is effective for the tested different algae strains.
" Ammonia is converted into ammonium using greenhouse gas during the novel process.
" The ammonia added will be reused as fertilizer in the subsequent cultures.
" Ammonia does not change the metabolic components without metal ions inputting.
a r t i c l e
i n f o
Article history:
Received 7 May 2012
Received in revised form 26 June 2012
Accepted 27 June 2012
Available online 3 July 2012
Keywords:
Aqueous ammonia
Algae
Harvest
Reuse
a b s t r a c t
Microalgae have been considered as a promising feedstock for biofuels and greenhouse gas reduction. A
low-cost harvesting technology without secondary contamination for down-stream extraction is a key
requirement to make algal biofuel commercially viable. A novel harvesting method using ammonia as
a ﬂocculant to make the algal biomass settable was devised and studied. Another major advantage of this
approach is that the ammonia added will be reused as fertilizer in the subsequent cultures. The results
indicated that ammonia-induced ﬂocculation led to more than 99% removal of algae at 12 h. The OD600
of algae growing in the ammonia-enriched ﬂocculation medium treated with heating and CO2 was 2
times than that of initial after 6 days. These results suggested that this ﬂocculation method was efﬁcient,
convenient and allowed the reuse of the ﬂocculated medium, therefore providing an option for economic
harvesting and cultivation of microalgae.
Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction
With the continuous depletion of petroleum and the rising CO2
concentration in the atmosphere from the combustion of fossil fuels,
increasing attention has been given to the development of the alternative fuels, such as lipid-derive biodiesel (Iglesias-Rodriguez,
2008) and renewable hydrocarbons (Graves et al., 2011). Biodiesel
has traditionally been produced with a variety of feedstocks such
as soybeans, canola oil, animal fat, palm oil, corn oil, waste cooking
oil (Felizardo et al., 2006), jatropha oil (Barnwal and Sharma, 2005).
These conventional lipid sources not only are subjected to quantity
limitation, but also cause concerns in competition with the need as
food for human. Algae have attracted a great interest in recent years
because of their various desirable characteristics such as high
potential productivity (Amaro et al., 2011; Brennan and Owende,
2010; Chisti, 2007). Additionally, algae can be potentially produced
with resources such as salt water or wastewater that are not used for
food production. Moreover, biofuel production from large-scale
⇑ Corresponding author.
E-mail address: [email protected] (S. Chen).
0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved.
http://dx.doi.org/10.1016/j.biortech.2012.06.076
cultivation of microalgae has been widely regarded as one of the
most promising alternatives that have potential to deal with the
problem of global warming (Benemann and Oswald, 1996; Chisti,
2007).
Many technical barriers, however, exist to the commercialization of algal biofuel systems among which concentrating the algal
biomass is a major challenge (Wang et al., 2008). It was estimated
that the cost of harvesting algal biomass was 20–30% of the total
costs of production (Gudin and Thepenier, 1986) due to small cell
size (typically in the range of 1–30 lm), low concentration (typically in the range of 0.3–5 g/l), unfavourable density (only slightly
greater than media) and negative charge (contributing to their stability in a dispersed state) (Brennan and Owende, 2010).
Major techniques available for harvesting microalgae include
ﬁltration, centrifugation, gravity sedimentation, ﬂocculation
(Benemann and Oswald, 1996; Molina et al., 2003) and electrophoresis (Amaro et al., 2011; Danquah et al., 2009). However, rapid
clogging, continuous backwashing and the costs arising from
pumping and membrane replacement are the major problems in
ﬁltration techniques (Becker, 1994; Molina et al., 2003). Centrifugation requires high energy input, complicated processing and
F. Chen et al. / Bioresource Technology 121 (2012) 298–303
299
large capital investment (Benemann and Oswald, 1996; Molina
et al., 2003). Gravity settling is suitable only to harvest large-sized
microalgal cells, e.g. Spirulina spp.(Xiong et al., 2008). Electrolytic
processes with increasing system temperature and cathode fouling
lead to high power consumption (Amaro et al., 2011).
The harvesting of microalgal cells by ﬂocculation is seen to be a
superior method to other aforementioned harvesting methods because of its effectiveness with given cost (Pushparaj et al., 1993).
Chemicals called ﬂocculants are usually added to induce ﬂocculation
followed by gravity separation. Multivalent metal salts like ferric
chloride (FeCl3), aluminium sulphate (Al2(SO4)3) and ferric sulphate
(Fe2(SO4)3) (Shelef et al., 1984) and certain cationic polymers such
as, chitosan, cationic polyacrlyamides, and cellulose, surfactants,
and other man-made ﬁbers (Bilanovic et al., 1988; Oh et al., 2001;
Pushparaj et al., 1993) have been tested effective. Although ﬂocculation has proven to be successful for concentrating microalgae, a large
amount of ﬂocculant is needed to cause solid–liquid separation of
the microalgae. The algal biomass as the end product is contaminated by the added ﬂocculant, thus algae harvesting with chemical
ﬂocculant is not suitable for biofuel application due the added cost
and residual effects on algal biomass and the culture water.
An ideal ﬂocculant for microalgae harvesting must meet the following criteria: (i) resulting in no residual in biomass, (ii) leading
to high efﬁcient subsequent settling of aglae, (iii) allowing reusing
the algal culture medium as growth supporting nutrients for subsequent algal cultivation, (iv) considering the environmental impact, reducing the greenhouse gas effect. A ﬂocculation process
using ammonia as coagulant was developed to meet these criteria.
In this process, aqueous ammonia is used to alter the pH of the culture that leads to the ﬂocculation and settling of the algae. Upon
the removal of the algal biomass, ﬂue gas containing CO2 is introduced to lower the pH to convert the un-ionized ammonia to its ionic form to reduce its potential toxicity when the ammonia laden
water is resued for algae culture (Fig. 1). This paper presents the
test results of this process, including ﬂocculation efﬁciency, ammonia conversion using CO2, analysis of cell metabolite and morphology, and reuse of the ﬂocculated culture water. Upon further
reﬁnement, this process has a potential to provide a viable option
for bio-friendly and economic mass cultivation and harvesting of
microalgae to make algal biofuel production more competitive.
(fresh water), Nannochlropsis oculata (marine), and native algal
species (marine, named HTBS, classiﬁed as Dunaliella). C. sorokiniana was cultivated using a Bold’s Basal Medium without glucose
(Bischoff and Bold, 1963), the pH of the medium was adjusted to
6.1 before sterilization. The marine algae were cultivated using a
f/2 medium (Guillard and Ryther, 1962) with ﬁltered sea water,
the composition brieﬂy was: 1.5 g NaNO3, 0.04 g K2HPO4, 0.006 g
ferric ammonium citrate, and trace metal mix A5 in 1 L of distilled
water without adjusting pH (pH was 8.0). The strains were incubated in plat bioreactor that contained 15 L of the f/2 medium. The
algal culture was continuously bubbling sterilized air with 3% CO2
under continuous illumination at 150 lmol m2 s1. The culture
temperature was 25 ± 1 °C and the strain was cultivated for
14 days.
2. Methods
2.3. Scanning electron microscopy (SEM) morphology analysis
2.1. Strains and culture medium
Scanning electron microscopy (JEOL-2100F, Japan) was used to
observe the morphology of HTBS algae cells, and the ﬂocs after
the ﬂocculation experiment. HTBS algae cells were placed on glass
slides and dried in air. Instead of ﬁxation as traditionally used to
The ammonia-based ﬂocculation process was evaluated with
both freshwater and marine algae, including Chlorella sorokiniana
2.2. Experimental design and analysis of ﬂocculation efﬁciency
In order to assess the ﬂocculation effect of aqueous ammonia,
the experiment of HTBS comparing with HTBS-treated algae with
aqueous ammonia was ﬁrst carried out. To further analyze the ﬂocculation effect quantitatively, the aforementioned three strains
were tested. The cultures were stirred using magnetic stirrer and
added different doses of commercial aqueous ammonia by titrimetry. For HTBS the concentrations were 0.09, 0.36, 1.09, 3.12, 12.90,
38.37 mmol L1, For N. oculata the concentrations were 0.74, 2.22,
4.44, 10.72, 26.62, 57.31, 118.7 mmol L1, and for C. sorokiniana
the concentrations were 0.52, 4.95, 9.54, 24.26, 48.14, 113.3,
240.6 mmol L1. After the ammonia addition, the removal efﬁciency
of ﬂocculated algal cultures was measured at 0.5 h, 1 h, 3 h, 6 h,
12 h. An aliquot was taken at a height of two-thirds from the bottom, and the pH was measured. The optical density (OD) of the aliquot was measured at 600 nm to evaluate the ﬂocculation efﬁciency
of the aqueous ammonia (Kim et al., 2011). The ﬂocculating efﬁciency was calculated using the following equation:
Flocculating efficiency ð%Þ ¼ ð1 B=AÞ 100
where, A is the optical density of the algal culture before the ﬂocculation measured at 600 nm and B is the optical density of the sample at 600 nm. The supernatant of culture medium was used to
carry out reuse experiment.
Fig. 1. New harvesting strategy using aqueous ammonia, ﬁrst algae is ﬂocculated and removed after settling, then the supernatant culture medium containing high
concentration of NH3H2O, ammonium, hydroxyl ion react with ﬂue gas (10–15% CO2), in which the CO2 and ammonia are converted into bicarbonate and ammonium ion
which can be used as the carbon and nitrogen sources, the remains of hydroxyl ion and NH3H2O are greatly reduced, the pH also can be return to normal levels, ﬁnally the
culture medium is simply treated and reused to culture the algae.
300
F. Chen et al. / Bioresource Technology 121 (2012) 298–303
Fig. 2. Flocculation effect of HTBS (A), N. oculata (B), C. sorokiniana (C) under different dose ammonia (/L) and time. Removal efﬁciency of HTBS (D), N. oculata (E), C.
sorokiniana (F) after different combination treatment of ammonia adjusting pH and medium, the pH of D–F was 10.8, 10.7, 10.0, respectively. +/pH normal (C): the normal
algae medium with/without adjusting corresponding pH as control group;+pH/dH2O (f/2, BBM) (S): the normal algae medium was centrifuged then adjusting corresponding
pH after resuspending with dH2O, f/2 or BBM.
prepare for SEM, small amounts of ﬂocs from ﬂocculation of algae
cells were simply placed on glass slides and air dried to avoid damaging the ﬂocs structure. Dried samples were mounted on copper
stubs and sputter coated with gold-palladium. The specimens were
observed at 5 kV(Yan et al., 2009).
and the bubbling was stopped when the pH was reduced to 6.5.
After that the medium was placed on magnetic stirrer and stirred
intensely, and the stirring stopped when the pH became stable.
2.4. pH kinetics changes of culture medium and ammonia conversion
using CO2
To further analyze the three algae metabolites such as protein
and lipid to evaluate whether they were affected by the ammonia
induced ﬂocculation, comparisons between ﬂocculation and centrifugation were made. The algae biomasses were harvested using
ammonia ﬂocculation and centrifugation, and the contents of neutral lipid, total protein, and pigments were measured.
The supernatant of ﬂocculated culture medium was bubbled
with pure CO2 using airstones, the velocity of CO2 was controlled
at 60 ml/min. The pH was monitored and recorded over time,
2.5. Measurement of protein, pigment and lipid content
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F. Chen et al. / Bioresource Technology 121 (2012) 298–303
Algal total lipid was extracted according to the procedure reported (Bligh and Dyer, 1959). For each 40 mg of sample, 6 ml of
CHCl3: methanol (2:1, v/v) was added and vortexed well, then
2 ml of methanol was added, the supernatant was mixed with
3.6 ml 5% NaCl, then centrifuged at 3000 g for 10 min, ﬁnally
the organic phase (bottom) was collected and dried at 60 °C using
pressured gas blowing concentrators, the crude oil was weighted.
Proteins were extracted using 0.5 M sodium hydroxide followed
by boiling 10 min, and then centrifuged at 8000 g for 5 min, the
supernatant was measured using Bradford assay (Bradford, 1976).
The samples were both centrifuged at 5000 g for 10 min, the
pellets were homogenized in an ice-cold 80% acetone (v/v) with a
chilled mortar and pestle. The chlorophyll content was determined
spectrophotometrically as described in (Arnon, 1949).
2.6. Reuse of ﬂocculated algal culture medium for subsequent
cultivation of algae
The algae HTBS was cultivated for 16 days at 26 °C using f/2 as
the growth medium. The algae were then ﬂocculated using the
ammonia-based process, ﬂocs from the culture medium were separated using gravity and centrifuged. The supernatant culture medium was treated with boiling, bubbling with 60 ml/min pure CO2
until pH recovery to about 8.0, combining of bubbling and heat,
respectively. After that trace metal and A5 were added before inoculation with a 10% (v/v) HTBS. Sampling was conducted every
2 days. The cultivation and harvest cycles were repeated three
times to investigate the growth supportability of the ﬂocculated
medium, which was measured based on the spectrophotometer
method. In a control experiment, fresh and sterilized f/2 with
1 mM ammonium bicarbonate (NH4HCO3) was used as extra nitrogen and carbon source.
3. Results and discussion
3.1. Analysis of ﬂocculation efﬁciency
The effectiveness of the ﬂocculation process appeared very
obvious based on the observation of the color change. Supplementa1 Fig. 1A and 1B show the color difference between the untreated and treated algae HTBS. After the addition of aqueous
ammonia to the medium and a short-time settling, the upper portion of the medium was highly transparent. The bottom portion of
the medium, on the other hand, existed a lot of distinct ﬂocs which
were deep green, compact and irregular surface shape. The similar
results were also observed with N. oculata and C. sorokiniana.
The above observation can be further conﬁrmed with the calculated ﬂocculation efﬁciency. The removal efﬁciency of marine algae
HTBS was 66.9% at 38.37 mmol L1 of ammonia within 0.5 h, to
91.2% at 3 h, and to more than 95% at 12 h (Fig. 2 A). The similar results were also obtained when using other two algae strains (Fig. 2 B
and C), the settling of N. oculata became substantial when the
ammonia concentration was at more than 57.31 mmol L1, the corresponding removal efﬁciency reached to 70.7% within 0.5 h, to 93%
at 3 h, and to more than 99% at 12 h. For C. sorokiniana, the trends of
the ﬂocculation effect were similar to the marine algae, but the removal efﬁciency was less distinct than marine algae. It was only
16.4% at 113.3 mmol L1 within half an hour, even at 12 h, it just
reached to 49.9%. However, the pH of HTBS adding 38.37 mmol
ammonia perlitre medium was 10.8; the pH of N. oculata adding
57.31 mmol was 10.7; whereas the pH of C. sorokiniana adding
113.3 mmol was 10.0. Therefore, it suggests that this ﬂocculation
process has a certain pH threshold above which it becomes more
effective. This is consistent with Vandamme’s study (Vandamme
et al., 2012). The pH effect is more sensitive for marine algae than
Table 1
Protein contents of three algae strains under centrifugation and aqueous ammonia
harvest.
Proteins (lg/ml)
Aqueous ammonia
Centrifugation
HTBS
C. sorokiniana
N. oculata
628 ± 149
286.4 ± 169
295.6 ± 105
626 ± 105
294.8 ± 99
305.6 ± 109
Table 2
Chlorophyll contents of three algae strains under centrifugation and aqueous
ammonia harvest.
Pigment (lg/ml)
HTBS
C. sorokiniana
N. oculata
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
Chlorophyll
a
b
(a + b)
a
b
(a + b)
a
b
(a + b)
Aqueous ammonia
Centrifugation
19.24 ± 1.562
3.24 ± 1.092
22.48 ± 2.655
10.39 ± 0.171
2.97 ± 0.115
13.36 ± 0.286
8.54 ± 0.314
1.65 ± 0.108
10.19 ± 0.422
16.59 ± 1.243
5.25 ± 0.163
21.84 ± 1.080
11.01 ± 0.275
3.69 ± 0.115
14.70 ± 0.390
8.96 ± 0.105
2.46 ± 0.196
11.42 ± 0.301
Table 3
Lipid contents of three algae strains under centrifugation and aqueous ammonia
harvest.
Lipid (w/dry)
Aqueous ammonia (%)
Centrifugation (%)
HTBS
C. sorokiniana
N. oculata
20.6 ± 0.549
23.6 ± 0.054
67.2 ± 1.151
21.1 ± 0.348
23.7 ± 0.026
68.6 ± 1.09
for fresh water algae. It was inferred that the compositions of sea
water are complex and some ions react with each other under high
pH condition to cause chemical formations that subsequently lead
the algal cell surface change to ﬂocculate, which accelerated the settling of the algae. The function of ions as co-factors was further veriﬁed by the results of the experiments testing the ﬂocculation effect
(Fig. 2D–F). Regardless the types of culture medium used, the effect
of adjusting pH on ﬂocculation performance was remarkable
(>99%). However, the removal efﬁciency difference caused by
adjusting pH in distilled water (dH2O) was not at all signiﬁcant
(<5%). This demonstrates that dramatically ﬂocculation effects
cannot be obtained by only increasing pH. These results are in
agreement with previous studies (Sukenik and Shelef, 1983; Vandamme et al., 2012). The pH-related coagulation and ﬂocculation
mechanisms were believed to be affected by the balance between
the electrostatic repulsion and the Van der Waals attraction through
increasing ionic strength in medium, reducing the microalgae surface charge or being charge neutralization (Vandamme et al.,
2012). In contrast, the results of this study suggest that ﬂocculation
of algae is caused by another mechanism: the rising pH generates
magnesium hydroxide, calcium hydroxide (Vandamme et al.,
2012), calcium phosphate (Sukenik and Shelef, 1983), and other
insoluble compound particles which covered the algae cell surface
(Supplemental Fig. 1D), which in combination with charge neutralization or adhesive attraction, forming heavier coagulation ﬂocs and
settling. Therefore, using the aqueous ammonia ﬂocculation process
to harvest algae was more suitable for marine algae applications.
3.2. SEM analysis
The results of SEM analysis showed that cell morphology was
fuzzy, ﬂagella shedded, surface damaged. Cells adhesion occurred
with a large number of coat (Supplemental Fig. 1D). The normal
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F. Chen et al. / Bioresource Technology 121 (2012) 298–303
were also observed (Tables 2 and 3). These results therefore suggest that ﬂocculation harvesting using aqueous ammonia rarely
inﬂuenced the changes of algae metabolite contents. The similar
results were also reported by other researchers (Knuckey et al.,
2006; Vandamme et al., 2012).
3.4. Reuse of ﬂocculated algal culture medium for cultivation of algae
Fig. 3. Reuse of ﬂocculated supernatant using aqueous ammonia. The medium was
treated with CO2 (square), heat (circle), combination of heat and CO2 (up
triangular), respectively, NH4HCO3 (down triangular) as control.
algae shape however, was clear, no damaged, recognizable ﬂagella,
no excess ﬂoc cover, with dispersed cell particles (Supplemental
Fig. 1C). This explains the reason that allows the cells to settle. Microalgae are generally small (1–30 lm) with low density (i.e. similar
to culture medium) and the surface is with negative charge. These
factors make the microalgae not ﬂocculate under normal growth
conditions. However, after adding aqueous ammonia, the surface
structure reacted with ammonia and was damaged, ﬁnally formed
lumps by incorporating other ions. That was equivalent to increase
the volume and cell density, which was higher than the density of
medium, causing sedimentation to occur. The speciﬁc components
of these coverings or adhesion complexes, however, are still unclear. More research is required to investigate the components of
ﬂocs after precipitation of algae occurs.
3.3. Metabolite inﬂuence of algae after ﬂocculation using aqueous
ammonia
The results showed that the total protein content of the HTBS,
C.sorokiniana, N. oculata harvested using centrifugation was
0.626, 0.294, 0.305 mg/ml, respectively, whereas that using aqueous ammonia was 0.628, 0.286, 0.295 mg/ml, respectively (Table
1). These results indicate that the protein content difference between ammonia ﬂocculation and centrifugation is not signiﬁcant
(p < 0.05). The similar results on chlorophyll and lipid contents
The HTBS algae strains were used to evaluate the suitability of
the recycled culture medium for new culture. The supernatant of
growing medium was simply treated by method described earlier
(2.6). The results showed that algae HTBS could normally grow in
the presence of 1 mM NH4HCO3, the OD600 increased 3.4 times after
6 days (Fig. 3), indicating that algae can utilize not only NH4+, but
also HCO3-. But algae were not good growth only treated by bubbling CO2 or heating. Although Conover (1975) observed that algae
could assimilate not only nitrate but also urea and ammonium,
ammonia inhibited photosynthesis and growth of Scenedesmus obliquus at concentrations over 2.0 mM and at pH values over 8.0, other
algae such as Chlorella pyrenoidosa, Anacystis nidulans, and Plectonema boryanum were also susceptible to ammonia inhibition
(Aharon and Yosef, 1976). However, the OD600 of algae under combination of heating and bubbling CO2 grew two times after 6 days.
Combining heating and bubbling CO2, on the one hand, might have
stripped out part of the ammonia, which was similar with Roney’s
results. The concentration of NH3 was mostly dependent on temperature, the solubility of NH3 in water could decrease to 18% (w/
w) at 50 °C from 47% (w/w) at 0 °C (Roney et al., 2004). At the same
time, CO2 addition helped to convert the residual ammonia into
ammonium. The amount of CO2 required to bring back the pH to
8.0 (proper for marine algae) was also estimated (Fig. 4) according
to the equilibrium NH3 + H2O ¡ NH3H2O ¡ NH4+ + OH (Nickolette et al., 2004), and the rate of CO2 outgassing through an air–
water surface to the atmosphere (Chi et al., 2011; Zeebe and
Wolf-Gladrow, 2001). With assumed conditions of 20 mM inorganic carbon, the transfer coefﬁcient which was 1 106 m/s, the
amount of CO2 required was approximately 38.37 mM to lower
the pH from 10.8 to 8.0 for HTBS culture. Improving the efﬁciency
of conversion of ammonia and carbon dioxide, however, needs further to be researched.
4. Conclusions
The novel process using ammonia to ﬂocculate algae is effective
for the tested different algae strains. The culture medium after the
treatment can be reused for algae culture. Ammonia as a ﬂocculant
without metal ions contamination also does not change the algae
Fig. 4. pH kinetic curve of ﬂocculated supernate using aqueous ammonia with CO2 (A) and stirred after stopping bubbling CO2 (B).
F. Chen et al. / Bioresource Technology 121 (2012) 298–303
cell metabolic components. Further research needs to be conducted reveal the speciﬁc mechanism and to optimize the process.
Acknowledgements
This work was supported partially by the National Program on
Key Basic Research Project (2011CB200905 and 2011CB200906)
and by Tianjin Municipal Science and Technology special project
(No. 10ZCKFSY05400), the Hi-Tech Research and Development Program (863) of China (2012AA052103), science and technology support key project plan of Tianjin (12ZCZDSFO2000).
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.biortech.2012.
06.076.
References
Aharon, A., Yosef, A., 1976. Toxicity of ammonia to algae in sewage oxidation ponds.
Appl. Environ. Microbiol. 31, 801–806.
Amaro, H.M., Guedes, A.C., Malcata, F.X., 2011. Advances and perspectives in using
microalgae to produce biodiesel. Appl. Energy. 88, 3402–3410.
Arnon, D.I., 1949. Copper enzymes in isolated chloroplasts. Polyphenoloxidase in
beta vulgaris. Plant Physiol. 24, 1–15.
Barnwal, B., Sharma, M., 2005. Prospects of biodiesel production from vegetables
oils in India. Renew. Sustain. Energy Rev. 9, 363–378.
Becker, E.W., 1994. Microalgae Biotechnology and Microbiology. Press Syndicate of
the University of Cambridge, Cambridge.
Benemann, J., Oswald, W.J., 1996. Systems and economic analysis of microalgae
ponds for conversion of CO2 to biomass. U.S. Department of, Energy, pp. 105–
109.
Bilanovic, D., Shelef, G., Sukenik, A., 1988. Flocculation of microalgae with cationic
polymers- Effects of medium salinity. Biomass 17, 65–76.
Bischoff, H.W., Bold, H.C., 1963. Some soil algae from enchanted rock and related
algal species. Psychol. Stud. 6318, 1–95.
Bligh, E.G., Dyer, W.J., 1959. A rapid method for total lipid extraction and
puriﬁcation. Can. J. Biochem. Physiol. 37, 911–917.
Bradford, M., 1976. A rapid and sensitive for the quantitation of microgram
quantitites of protein utilizing the principle of protein-dye binding. Anal.
Biochem. 72, 248–254.
Brennan, L., Owende, P., 2010. Biofuels from microalgae – a review of technologies
for production, processing, and extractions of biofuels and co-products. Renew.
Sustain. Energy Rev. 14, 557–577.
Chi, Z., Fallon, J.V.O., Chen, S., 2011. Bicarbonate produced from carbon capture for
algae culture. Trends Biotechnol. 11, 537–541.
Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306.
303
Danquah, M., Ang, L., Uduman, N., Moheimani, N., Fordea, G., 2009. Dewatering of
microalgal culture for biodiesel production: exploring polymer ﬂocculation and
tangential ﬂow ﬁltration. J. Chem. Technol. Biotechnol. 84, 1078–1083.
Felizardo, P., Correia, M., Raposo, I., Mendes, J., Berkemeier, R., Bordado, J., 2006.
Production of biodiesel from waste frying oil. Waste Manag. 26, 487–494.
Graves, C., Ebbesen, S.D., Mogensen, M., Lackner, K.S., 2011. Sustainable
hydrocarbon fuels by recycling CO2 and H2O with renewable or nuclear
energy. Renew. Sustain. Energy Rev. 15, 1–23.
Gudin, C., Thepenier, C., 1986. Bioconversion of solar energy into organic chemicals
by microalgae. Advances in Biotechnological Processes (USA).
Guillard, R.R.L., Ryther, J.H., 1962. Studies of marine planktonic diatoms: I.
Cyclotella nana hustedt, and detonula confervacea (cleve) gran. Can. J.
Microbiol. 8, 229–239.
Iglesias-Rodriguez, M.D., Halloran, P.R., Rickaby, R.E.M., Hall, I.R., ColmeneroHidalgo, E., Gittins, J.R., Green, D.R.H., Tyrrell, T., Gibbs, S.J., Dassow, P., Rehm, E.,
Armbrust, E.V., Boessenkool, K.P., 2008. Phytoplankton calciﬁcation in a highCO2 world. Science 320, 336–340.
Kim, D.G., La, H.J., Ahn, C.Y., Park, Y.H., Oh, H.M., 2011. Harvest of Scenedesmus sp.
with bioﬂocculant and reuse of culture medium for subsequent high-density
cultures. Bioresour. Technol. 102, 3163–3168.
Knuckey, R.M., Brown, M.R., Robert, R., Frampton, D.M.F., 2006. Production of
microalgal concentrates by ﬂocculation and their assessment as aquaculture
feeds. Aquacult. Eng. 35, 300–313.
Molina, E.G., Belarbi, E.-H., Acién Fernández, F.G., Medina, A.R., Chisti, Y., 2003.
Recovery of microalgal biomass and metabolites: process options and
economics. Biotechnol. Adv. 20, 491–515.
Nickolette, R., Fernando, L., 2004. Ammonia, agency for toxic substances and disease
registry. U.S. Department of Health and Human Services, Public Health Service,
pp. 113–116.
Oh, H.M., Lee, S.J., Park, M.H., Kim, H.S., Kim, H.C., Yoon, J.H., Kwon, G.S., Yoon, B.D.,
2001. Harvesting of Chlorella vulgaris using a bioﬂocculant from Paenibacillus sp.
AM49. Biotechnol. Lett. 23, 1229–1234.
Pushparaj, B., Pelosi, E., Torzillo, G., Materassi, R., 1993. Microbial biomass recovery
using a synthetic cationic polymer. Bioresour. Technol. 43, 59–62.
Roney, N., Llados, F., Little, S.S., 2004. Ammonia, agency for toxic substances and
disease registry. U.S. Department of Hhealth and Human Services, Public Health
Service, pp. 113–116.
Shelef, G., Sukenik, A., Green, M., 1984. Microalgae harvesting and processing: a
literature review. A subcontract, report. SERI/STR-231-2396.
Sukenik, A., Shelef, G., 1983. Algal autoﬂocculation-verif ication and proposed
mechanism. Biotechnol. Bioeng. 26, 142–147.
Vandamme, D., Foubert, I., Fraeye, I., Meesschaert, B., Muylaert, K., 2012.
Flocculation of Chlorella vulgaris induced by high pH: role of magnesium and
calcium and practical implications. Bioresour. Technol. 105, 114–119.
Wang, B., Li, Y., Wu, N., Lan, C.Q., 2008. CO2 bio-mitigation using microalgae. Appl.
Microbiol. Biotechnol. 79, 707–718.
Xiong, W., Li, X., Xiang, J., Wu, Q.Y., 2008. High-density fermentation of microalga
Chlorella protothecoides in bioreactor for microbio-diesel production. Appl.
Microbiol. Biotechnol. 78, 29–36.
Yan, D., Bai, Z., Mike, R., Gu, L., Ren, S., Yang, P., 2009. Bioﬁlm structure and its
inﬂuence on clogging in drip irrigation emitters distributing reclaimed
wastewater. J. Environ. Sci. (China) 21, 834–841.
Zeebe, R.E., Wolf-Gladrow, 2001. CO2 in Seawater: Equilibrium, Kinetics, Isotopes.
In: Oceanography Series. Elsevier, Amsterdam.