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doi.org/10.26434/chemrxiv.12479195.v1
Sustainable Harvesting of Microalgae by Coupling Chitosan Flocculationand Electro-FloatationLin Zhu, Hui Xu, Weijie Guo, Jianghua Yu, Wenqing Shi
Submitted date: 14/06/2020 • Posted date: 16/06/2020Licence: CC BY-NC 4.0Citation information: Zhu, Lin; Xu, Hui; Guo, Weijie; Yu, Jianghua; Shi, Wenqing (2020): SustainableHarvesting of Microalgae by Coupling Chitosan Flocculation and Electro-Floatation. ChemRxiv. Preprint.https://doi.org/10.26434/chemrxiv.12479195.v1
In this study, we proposed a new method for edible microalgae harvesting by coupling chitosan flocculationand electro-floatation using carbon electrodes, and tested it at different operation conditions. The surfacecharge of microalgae cells was measured to explore the underlying mechanisms. The responses of growthmedium to microalgae harvesting were also investigated to evaluate the feasibility of sustainable utilization ofculture medium. (1) Microalgae harvesting efficiencyThe microalgae harvesting efficiency of the proposedmethod were tested using Chlorella vulgaris (C. vulgaris). C. vulgariscells (FACHB-24) were purchased fromthe Institute of Hydrobiology, Chinese Academy of Sciences, and cultured in the BG11 medium, whichconsists of 500 mg L-1 Bicin, 100 mg L-1 KNO3, 100 mg L-1 b-C3H7O6PNa2, 50 mg L-1 NaNO3, 50 mgL-1Ca(NO3)2•4H2O, 50 mg L-1 MgCl2•6H2O, 40 mg L-1 Na2SO4, 20 mg L-1 H3BO3, 5 mg L-1 Na2EDTA, 5mg L-1 MnCl2•4H2O, 5 mg L-1 CoCl2•6H2O and 0.8 mg L-1 Na2MoO4•2H2O, 0.5 mg L-1 FeCl3•6H2O and 0.5mg L-1 ZnCl2. The batch cultures were conducted in an illuminating incubator (LRH-250-G, GuangdongMedical Apparatus Co., Ltd., China) with continuous cool white fluorescent light of 2500 ± 500 lux on a 12 hlight and 12 h darkness regime at the temperature of 30 ± 1°C.The microalgae harvesting efficiency systemconsists of a flat stir paddle (Zhongrun Water Industry Technology Development Co., Ltd., China) for mixingduring chitosan flocculation and two round carbon electrode plates (Jinjia Metal Co., Ltd., China) forelectro-floatation. The carbon electrode plate has a surface area of 55.4 cm2 and a thickness of 0.2 cm, whichwas horizontally installed at the bottom with a gap of 2 cm between the two plates. There are 85 small roundholes on each carbon electrode plate to allow gas bubbles freely pass it during electrolysis, such that theeffective surface area was 38.7 cm2. The electric current was supplied by a direct current power supply(DF1730SL5A, Ningbo Zhongce Dftek Electronics Co., Ltd., China). C. vulgaris culture at the exponentialgrowth phase was used in the microalgae harvesting test. The initial cell concentration was set to 3.63 × 1010
cells L-1. 0.5 L of readily prepared C. vulgaris solution was transferred to the harvesting cell. Water-solublechitosan was purchased from Qingdao Yunzhou Bioengineering Co. Ltd., China. Prior to the test, a chitosanstock solution (2 g L-1) was prepared as follows: 1 g chitosan was added to 0.5 L distilled water andcompletely diluted by stirring. After chitosan was added, the microalgae solution was stirred at 200 rpm for 2min and 40 rpm for another 10 min; electro-floatation was started in the last 5 min during chitosan flocculation.
The microalgae solution was allowed to stand for 10 min, and then water samples were carefully collectedfrom an outlet 2 cm above the carbon electrode plate to enumerate the cell number using an Axioskop 2 motplus microscope (Carl ZEISS, Germany). The microalgae harvesting efficiency was calculated as (initial cellconcentration-sample cell concentration)/initial cell concentration × 100%. In the test, the chitosan dosagewas set to 0, 2, 4, 6, 8, 10, 12 and 15 mg L-1, and the current density was set to 0, 0.2, 0.4 and 0.6 A. All thetests were conducted in triplicate at the raw microalgae solution pH of 8.6.(2) Surface charges of chitosan,microalgae cells and microalgae flocsThe surface charges of chitosan, microalgae cells and microalgae flocswere characterized using a Zetasizer 2000 (Malvern Co. United Kingdom). (3) Sustainable utilization ofmicroalgae culture mediumBefore and after microalgae harvesting, medium nutrients (phosphate, ammoniumand nitrate) were measured according to Chinese Monitoring Analysis Method of Water and Wastewater;medium pH and temperature were measured using a Yellow Springs Instruments (Yellow Springs, Ohio,USA)(4) Cost evaluationThe cost of microalgae harvesting was estimated by summing flocculants and energycosts per unit of harvested microalgae biomass. The chitosan and electric power are 0.03 USD g-1 and 0.08USD (kWh)-1, respectively.
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Sustainable harvesting of microalgae by coupling chitosan flocculation and electro-
floatation
Lin Zhu†, Hui Xu‡, Weijie Guo§, Jianghua Yu†, Wenqing Shi†,*
†Jiangsu Collaborative Innovation Center of Atmospheric Environment and Equipment
Technologies, Jiangsu Key Laboratory of Atmospheric Environmental Monitoring &
Pollution Control, School of Environmental Science & Engineering, Nanjing University of
Information Science & Technology, Nanjing 210044, China‡Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences, Beijing
100085, China§Basin Water Environmental Research Department, Changjiang River Scientific Research
Institute, Wuhan 430010, China*Corresponding author: Wenqing Shi ([email protected])
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Abstract. Microalgae have been widely used as animal feeds, healthcare products and food
additives; however, it still remains a challenge to harvest them safely and effectively from
growth medium. In this study, we tested a green method for microalgae harvesting by
coupling chitosan flocculation and electro-floatation. Results demonstrated that microalgae
can be preliminarily flocculated into unstable flocs by chitosan, and then floated by electro-
floatation, during which chitosan was charged by electrolysis and activated to flocculate
microalgae, producing a positive priming effect. It is possibly amino groups of chitosan were
positively charged by electrolysis, increasing the charge neutralization ability for microalgae
flocculation. Thus, the use of stronger current yielded a higher harvesting efficiency at a
lower chitosan dosage. At the current intensity of 0.2, 0.4 and 0.6 A, microalgae harvesting
efficiency reached a maximum of 33.2%, 59.6% and 63.5%, and the optimal chitosan dosage
was 6.0, 4.0 and 2.0 mg L-1, respectively. The use of edible chitosan and inert carbon
electrodes makes it possible to harvest microalgae biomass safely for food or healthcare use
and achieve sustainable utilization of culture medium, which will be beneficial to microalgae-
based engineering.
Keywords: microalgae harvesting, chitosan flocculation, electro-floatation, sustainable
utilization, microalgae-based engineering
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IntroductionIn recent years, microalgae have been widely used as animal feeds, healthcare products and
food additives.1,2 Microalgae are more nutritious than traditional animal and aquatic feed like
millet, grams and other small fishes in terms of protein, omega 3 fatty acids and carotenoids
content.3,4 Several kinds of antioxidant compounds have been extracted from microalgae to
protect against oxidative stress, a cause of many diseases and aging.1,5 However, it still
remains a challenge to find an effective and especially safe method for microalgae harvesting.Many physical and chemical methods have been tested to harvest microalgae, including
sedimentation, centrifugation, filtration, chemical flocculation and electro-flocculation.
Sedimentation is simple but time-consuming and only suitable for harvesting large-size
microalgae, since most of microalgae have similar density to water, negative surface charge,
and thereby low settling velocities.6 Centrifugation and filtration are rapid but less cost-
effective, limiting their applications at large scale.7,8 Mata et al. (2010) considered that
filtration consumes large amounts of membranes and pumping energy, especially for
harvesting unicellular small microalgae cells.8 Chemical flocculation and electro-flocculation
are rapid and effective, but residual chemical flocculants in the harvested biomass pose
potential health risks and limit their utilizations as food and healthcare products.9 So far, there
are few safe and effective technologies for microalgae harvesting.Chitosan is the second-most abundant natural biopolymer, which is mainly derived from the
shells of shrimp and other crustaceans.10,11 It has received widespread applications in food and
pharmaceutical industries due to its biological compatibility, biodegradability, and nontoxic
properties.12,13 Chitosan has positively charged amine groups and thereby exhibits flocculation
potentials for negatively charged microalgae.14 Pan et al. (2011) used chitosan to flocculate
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microalgae cells and then sink them with the aid of soil particles, achieving effective harmful
microalgae removal in Lake Taihu, China.15 Electro-floatation is a separation process in
which solids are separated from liquid by micro-bubbles produced from electrode surfaces.16
It has been widely used to remove floatable materials including activated sludge, oil,
surfactants, and flocculants etc.17,18 The use of non-sacrificial electrodes in electro-floatation
will not introduce chemical flocculants.19,20 Hence, if microalgae are preliminarily flocculated
into unstable flocs by chitosan, and then floated by electro-floatation using non-sacrificial
electrodes, it is possibly to harvest microalgae biomass safely and effectively. In this study, we proposed a new method for edible microalgae harvesting by coupling
chitosan flocculation and electro-floatation using carbon electrodes, and tested it at different
operation conditions. The surface charge of microalgae cells was measured to explore the
underlying mechanisms. The responses of growth medium to microalgae harvesting were also
investigated to evaluate the feasibility of sustainable utilization of culture medium. The
objective of this study is to develop a green method for the harvesting of edible microalgae.
Materials and methods
Microalgae species and culture
In this study, Chlorella vulgaris (C. vulgaris), an edible green microalgae species, was chosen
to test the new harvesting method.21,22 Here, C. vulgaris cells (FACHB-24) were purchased
from the Institute of Hydrobiology, Chinese Academy of Sciences, and cultured in the BG11
medium, which consists of 500 mg L-1 Bicin, 100 mg L-1 KNO3, 100 mg L-1 b-C3H7O6PNa2,
50 mg L-1 NaNO3, 50 mg L-1 Ca(NO3)2•4H2O, 50 mg L-1 MgCl2•6H2O, 40 mg L-1 Na2SO4, 20
mg L-1 H3BO3, 5 mg L-1 Na2EDTA, 5 mg L-1 MnCl2•4H2O, 5 mg L-1 CoCl2•6H2O and 0.8 mg
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L-1 Na2MoO4•2H2O, 0.5 mg L-1 FeCl3•6H2O and 0.5 mg L-1 ZnCl2. The batch cultures were
conducted in an illuminating incubator (LRH-250-G, Guangdong Medical Apparatus Co.,
Ltd., China) with continuous cool white fluorescent light of 2500 ± 500 lux on a 12 h light
and 12 h darkness regime at the temperature of 30 ± 1°C.
Microalgae harvesting system
The microalgae harvesting system consists of a flat stir paddle (Zhongrun Water Industry
Technology Development Co., Ltd., China) for mixing during chitosan flocculation and two
round carbon electrode plates (Jinjia Metal Co., Ltd., China) for electro-floatation (Fig. 1).
The carbon electrode plate has a surface area of 55.4 cm2 and a thickness of 0.2 cm, which
was horizontally installed at the bottom with a gap of 2 cm between the two plates. There are
85 small round holes on each carbon electrode plate to allow gas bubbles freely pass it during
electrolysis, such that the effective surface area was 38.7 cm2. The electric current was
supplied by a direct current power supply (DF1730SL5A, Ningbo Zhongce Dftek Electronics
Co., Ltd., China).
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Fig. 1. The schematic diagram of microalgae harvesting system.Microalgae harvesting testC. vulgaris culture at the exponential growth phase was used in the microalgae harvesting
test. The initial cell concentration was set to 3.63 × 1010 cells L-1. 0.5 L of readily prepared C.
vulgaris solution was transferred to the harvesting cell. Water-soluble chitosan was purchased
from Qingdao Yunzhou Bioengineering Co. Ltd., China. Prior to the test, a chitosan stock
solution (2 g L-1) was prepared as follows: 1 g chitosan was added to 0.5 L distilled water and
completely diluted by stirring. After chitosan was added, the microalgae solution was stirred
at 200 rpm for 2 min and 40 rpm for another 10 min; electro-floatation was started in the last
5 min during chitosan flocculation. The microalgae solution was allowed to stand for 10 min,
and then water samples were carefully collected from an outlet 2 cm above the carbon
electrode plate to enumerate the cell number using an Axioskop 2 mot plus microscope (Carl
ZEISS, Germany). The microalgae harvesting efficiency was calculated as (initial cell
concentration-sample cell concentration)/initial cell concentration × 100%. In the test, the
chitosan dosage was set to 0, 2, 4, 6, 8, 10, 12 and 15 mg L -1, and the current density was set
to 0, 0.2, 0.4 and 0.6 A. All the tests were conducted in triplicate at the raw microalgae
solution pH of 8.6.
The surface charge of microalgae cells was characterized using a Zetasizer 2000 (Malvern
Co. United Kingdom). To study the responses of culture medium to microalgae harvesting,
medium nutrients (phosphate, ammonium and nitrate) were measured according to Chinese
Monitoring Analysis Method of Water and Wastewater;23 medium pH and temperature were
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measured using a Yellow Springs Instruments (Yellow Springs, Ohio, USA) before and after
microalgae harvesting.
Cost evaluation
The cost of microalgae harvesting was estimated by summing flocculants and energy costs
per unit of harvested microalgae biomass as follows:
Cost=(Cost floccluation+Costeletrofloatation)/W biomass (1)
Cost floccluation=Costchitosan+Costmixting energy (2)
Costeletrofloatation=Costeletrolysis energy (3)
Costchitosan=C∗v∗a (4)
Costmixing energy=P∗T∗b (5)
Costeletrolysis energy=U∗I∗t∗b (6)
W biomass=v∗β∗θ∗σ (7)
where C is the chitosan dosage, mg L-1; v is the volume of microalgae solution, L; a is the
chitosan price, which is 0.03 USD g-1 ; P is the stirrer power, which is 40 W; T is the stirring
time, h; b is the electric power price, which is 0.08 USD (kWh)-1; U is the electrolysis
voltage, V; I is the current intensity, A; t is the electrolysis time, h; β is the initial microalgae
concentration, cell L-1; θ is the microalgae harvesting efficiency, %; σ is the weight per
microalgae cell, 32 × 10-12 g cell-1.
Data analysis
One-way analysis of variance (ANOVA) was employed to test the statistical significance of
differences between treatments. Post-hoc multiple comparisons of treatment means were
performed using the Tukey’s least significant difference procedure. All statistical calculations
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were performed using the SPSS (v22.0) statistical package for personal computers. The level
of significance was P < 0.05 for all tests.
Results
Surface charge properties of chitosan and C. vulgaris cells
The isoelectric point of chitosan was pH 9.7, making it possess net positive charges under
most microalgae culture conditions. The zeta potential of chitosan kept above +1.5 mV in the
wide pH range of < 9.0, and then decreased to nearly zero at pH 9.7. In contrast, the zeta
potential of C. vulgaris cells kept below zero in the pH range of > 2.5 (Fig. 2).
Fig. 2. The surface charge properties of chitosan and C. vulgaris cells. Error bars indicate
standard deviations.
Microalgae harvesting efficiency
The use of chitosan flocculation alone only achieved limited microalgae harvesting no matter
what the chitosan dosage was. The microalgae harvesting efficiency reached 16.9% at the
chitosan dosage of 2 mg L-1 and maintained stably at this value as the chitosan dosage further
increased. Similarly, the use of electro-floatation alone could not effectively harvest
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microalgae either and the harvesting efficiency was only 4.5%, 13.4% and 22.9% at the
current intensity of 0.2, 0.4 and 0.6 A, respectively. When chitosan flocculation and electro-
floatation were used together, the microalgae harvesting was greatly improved by increasing
harvesting efficiency and simultaneously decreasing the optimal chitosan dosage, and this
effect was enhanced as the current density increased. When the current intensity of 0.2, 0.4
and 0.6 A was applied, microalgae harvesting efficiency reached a maximum of 33.2%,
59.6% and 63.5%, and the optimal chitosan dosage was 6.0, 4.0 and 2.0 mg L-1, respectively.
However, a remarkable decrease in microalgae harvesting efficiency was observed as
chitosan was overdosed at the current intensity of 0.4 and 0.6 A, and the harvesting efficiency
decreased to 35.2% at the chitosan dosage of 15 mg L-1 (Fig. 3).
Fig. 3. Effects of chitosan dosage and current density on microalgae harvesting efficiency.
Error bars indicate standard deviations.
Microalgae surface charge
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After chitosan was added, the zeta potential of microalgae cells showed a remarkable
increase. The zeta potential of microalgae cells was gradually increased from –9.2 to –0.9 mV
as the chitosan dosage increased from 0 to 15 mg L-1. The use of electro-floatation further
increased the zeta potential of microalgae cells, indicating a charging effect, and this effect
was enhanced as the current intensity increased (Fig. 4A). When the current density of 0.2,
0.4 and 0.6 A was applied, the zeta potential of microalgae cells was finally increased to +7.3,
+8.4 and +10.6 mV at the chitosan dosage of 15 mg L-1, respectively. The charging effect was
further evaluated by subtracting the zeta potential of microalgae cells in presence of electro-
floatation from the value in the absence of electro-floatation. It exhibited chitosan limitation
at low chitosan dosages and current limitation at high chitosan dosages (Fig. 4B). As the
chitosan dosage increased, the increase amplitude of zeta potential gradually increased and
then reached an equilibrium; the use of higher intensity currents yielded a higher equilibrium
value at a lower chitosan dosage. At the current intensity of 0.2, 0.4 and 0.6 A, the increase
amplitude of zeta potential of microalgae cells reached a maximum of 9.3, 11.0 and 11.5 mV,
and the optimal chitosan dosage was 2, 4 and 8 mg L-1, respectively. In contrast, in the
absence of chitosan, there were no significant differences in zeta potential of microalgae cells
among different intensity currents (P < 0.05), and the zeta potential values kept stably at –9.4
mV at the chitosan dosage of 0 mg L-1 (Fig. 4A).
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Fig. 4. Changes of microalgae cell surface change during microalgae harvesting. (A) Zeta
potential of microalgae cells; (B) The charging effect of electro-floatation. Error bars indicate
standard deviations.
Microalgae culture medium
Compared with chitosan flocculation alone, ammonium in culture medium was significantly
increased after electro-floatation was introduced (P < 0.05), and the use of higher intensity
currents yielded higher ammonium concentrations. After microalgae harvesting, ammonium
concentration maintained stably below 0.2 mg L-1 in chitosan flocculation alone, and was
increased to 1.0, 1.6 and 2.2 mg L-1 when electro-floatation was applied at the current
intensity of 0.2, 0.4 and 0.6 A, respectively (Fig. 5A). In contrast, there were no significant
changes in other parameters observed after microalgae harvesting (P > 0.05). In all the tests,
medium nitrate and phosphate maintained stably at 250.9 and 4.66 mg L-1 (Fig. 5B); medium
pH, conductivity, and temperature maintained stably at 7.0, 2.4 mS cm-1 and 19.6°C,
respectively (Table S2).
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Fig. 5. Changes of nutrients in culture medium after microalgae harvesting. (A) Ammonium;
(B) Phosphate and nitrate. Error bars indicate standard deviations.
Microalgae harvesting cost
Compared with chitosan flocculation (0.08 × 10-3 USD g-1 biomass), it costed much more to
harvest microalgae using electro-floatation, reaching 2.83 × 10-3, 2.76 × 10-3 and 3.30 × 10-3
USD g-1 biomass at the 0.2, 0.4 and 0.6 A, respectively. When chitosan flocculation and
electro-floatation were used together, the cost could be greatly reduced, but exhibited a
potential increase as the current intensity increased. The cost reached 0.41 × 10-3, 0.64 × 10-3
and 1.18 × 10-3 USD g-1 biomass at 0.2, 0.4 and 0.6 A, respectively.
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Fig. 6. The cost evaluation for microalgae harvesting using the coupled chitosan flocculation
and electro-flocculation. The single red and blue circles indicated chitosan flocculation and
electro-floatation were used alone, respectively. The combined red and blue circles indicated
that chitosan flocculation and electro-floatation were used together.
Discussion
The synergistic effect of chitosan flocculation and electro-floatation
Microalgae particles have negative surface charge and often stably suspend in the solution
with electrostatic repulsion.24,25 Charge neutralization is an essential step in microalgae
flocculation, which eliminates energy barrier for microalgae aggregation.26-28 Chitosan is
positively charged over a wide range of pH < 9.7, which made it obtain flocculation potential
for negatively charged microalgae (Fig. 2). However, most of the formed microalgae flocs
still suspended with the aid of buoyancy. The zeta potential of microalgae cells showed a
remarkable increase as the chitosan dosage increase (Fig. 4). As a result, the use of chitosan
flocculation alone yielded limited microalgae harvesting efficiency whatever the chitosan
dosage was (Fig. 3). After electro-floatation was introduced, large amounts of tiny gas
bubbles were produced, carrying the flocs to water surface where they can be easily collected
(Fig. S1). Microalgae harvesting efficiency were therefore remarkably increased when
chitosan flocculation and electro-floatation were used together (Fig. 3). However, the use of
electro-floatation alone could not effectively harvest microalgae either. This is because
microalgae cells stably suspend with electrostatic repulsion, and it is difficult to float them by
bubbles without preliminary flocculation.29
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The positive priming effect of electro-floatation on chitosan flocculation
Chitosan was appointed to neutralize microalgae surface charge for microalgae flocculation
(Fig.2 and Fig. 4). Surprisingly, after electro-floatation was introduced, the zeta potential of
microalgae cells differed at the same chitosan dosage, depending on current intensity applied.
The zeta potential of microalgae cells generally exhibited an increase as the current intensity
increased, but this effect disappeared in the absence of chitosan (Fig. 4A). It indicated that
electrolysis charged chitosan and increased its charge neutralization ability for microalgae
flocculation, producing a positive priming effect. This explained that the use of higher current
density yielded higher microalgae harvesting efficiency at a lower optimal chitosan dosage.
There are a lot of amino groups on the chain of chitosan, which were possibly charged by
electrolysis during electro-floatation.30 At the low chitosan dosage, the receptor capacity was
limited and the charging effect of electrolysis exhibited a chitosan limitation. As the chitosan
dosage increased, chitosan limitation was transferred to current limitation (Fig. 4B), and the
stronger current has a higher charging effect. However, excess loads of positive charges can
cause microalgae cells positively charged and re-establish electrostatic repulsion.31 Thus, a
remarkable decrease in microalgae harvesting efficiency was observed as chitosan was
overdosed at the high current intensity (0.4 and 0.6 A, Fig. 3).
Recommendations for future applications
In the microalgae-based engineering, culture medium reuse can offer a promising strategy for
saving water and nutrients.32,33 Because of the electrolysis, the physico-chemical properties of
culture medium may change after microalgae harvesting.34,35 For instance, water temperature
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may increase as waste heat releases. However, there were no significant changes in medium
pH (P > 0.05), temperature (P > 0.05) and conductivity (P > 0.05) after microalgae
harvesting in this study (Table S1), which is possibly due to weak electrolysis (low current
intensity and short electrolysis time). As for main nutrients, there was a significant increase in
ammonium (P < 0.05) (Fig. 5A). We attributed it to the transformation from nitrate to
ammonium under electrolysis according to our additional experiments (Fig. S2). During
electrolysis, nitrate reduction (NO3- + 10H+ + 8e- = NH4
+ + 3H2O) can occur at the cathode,
releasing ammonium to the medium.36,37 Thus, the stronger current had a higher ammonium
yield (Fig.5A and Fig. S2). However, we did not detect the significant decrease in nitrate (P >
0.05, Fig. 5B) because of the extremely high levels of nitrate in the BG11 medium (> 250 mg
L-1). The shift of nitrate to ammonium can increase nitrogen bioavailability since ammonium
is generally favored by microalgae relative to nitrate.38-40 In contrast, phosphate did not show
significant changes after microalgae harvesting (P > 0.05, Fig. 5B) due to the use of non-
sacrificial electrodes, which differs with the decrease of phosphate concentration by
electrolysis using Fe/Al electrodes.41,42 Hence, nutrient bioavailability increase and other
parameter stability (Fig. 5B, Table S2) make medium reusable for microalgae continuous
culture.
In practical applications, cost is often another major concern for a technology. For the
microalgae harvesting technology in this study, there was a trade-off between the harvesting
efficiency and the cost, since the cost exhibited a potential increase as the current intensity
increased (Fig. 6). It will be cost-effective to apply the low current intensity to harvest
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microalgae in the continuous system, and the remaining cells can benefit microalgae
recovery. Despite the microalgae harvesting efficiency was greatly increased by coupling
chitosan flocculation and electro-floatation (Fig. 3), it may be further increased by optimizing
operation condition or screening other edible flocculants, such as moringa oleifera and
tannin.43,44
Conclusions
Microalgae harvesting is a crucial step but still remains a challenge for microalgae-based
engineering. This study proposed a green way to harvest microalgae by coupling chitosan
flocculation and electro-floatation. Microalgae can be preliminarily flocculated into unstable
flocs by chitosan, and then floated by electro-floatation; electro-floatation charged chitosan
and activated it to flocculate microalgae, producing a positive priming effect. The use of
edible chitosan and inert carbon electrodes makes it possible to harvest microalgae biomass
safely and effectively for food or healthcare use and achieve the sustainable utilization of
culture medium. Further studies are needed to optimize operation conditions to increase
harvesting efficiency and reduce the cost.AcknowledgmentsThis study was supported by the National Natural Science Foundation of China (No.
41701112, 51709181, 51979171), Starting Research Fund of Nanjing University of
Information Engineering (No. 20181015), Major Science and Technology Program for Water
Pollution Control and Treatment (No. 2017ZX07501-002, 2017ZX07108-002).Supporting InformationThe Supporting Information is available free of charge on the web.The microalgae floc image; the effects of electrolysis using carbon electrodes on BG11
medium; changes of physical properties of growth medium after microalgae harvesting.
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Notes The authors declare no competing financial interest.
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For Table of Contents Use Only
Synopsis: A green method for harvesting microalgae biomass will be beneficial to
microalgae-based engineering for healthcare and food use.
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download fileview on ChemRxivACS-MS-20200614.docx (896.06 KiB)
1
Sustainable harvesting of microalgae by coupling chitosan flocculation and electro-1
floatation 2
3
Lin Zhu†, Hui Xu‡, Weijie Guo§, Jianghua Yu†, Wenqing Shi†,* 4
5
6
†Jiangsu Collaborative Innovation Center of Atmospheric Environment and Equipment 7
Technologies, Jiangsu Key Laboratory of Atmospheric Environmental Monitoring & Pollution 8
Control, School of Environmental Science & Engineering, Nanjing University of Information 9
Science & Technology, Nanjing 210044, China 10
‡Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences, Beijing 11
100085, China 12
§Basin Water Environmental Research Department, Changjiang River Scientific Research 13
Institute, Wuhan 430010, China 14
*Corresponding author: Wenqing Shi ([email protected]) 15
16
2
Abstract. Microalgae have been widely used as animal feeds, healthcare products and food 17
additives; however, it still remains a challenge to harvest them safely and effectively from 18
growth medium. In this study, we tested a green method for microalgae harvesting by coupling 19
chitosan flocculation and electro-floatation. Results demonstrated that microalgae can be 20
preliminarily flocculated into unstable flocs by chitosan, and then floated by electro-floatation, 21
during which chitosan was charged by electrolysis and activated to flocculate microalgae, 22
producing a positive priming effect. It is possibly amino groups of chitosan were positively 23
charged by electrolysis, increasing the charge neutralization ability for microalgae flocculation. 24
Thus, the use of stronger current yielded a higher harvesting efficiency at a lower chitosan 25
dosage. At the current intensity of 0.2, 0.4 and 0.6 A, microalgae harvesting efficiency reached 26
a maximum of 33.2%, 59.6% and 63.5%, and the optimal chitosan dosage was 6.0, 4.0 and 2.0 27
mg L-1, respectively. The use of edible chitosan and inert carbon electrodes makes it possible 28
to harvest microalgae biomass safely for food or healthcare use and achieve sustainable 29
utilization of culture medium, which will be beneficial to microalgae-based engineering. 30
31
Keywords: microalgae harvesting, chitosan flocculation, electro-floatation, sustainable 32
utilization, microalgae-based engineering 33
34
3
Introduction 35
In recent years, microalgae have been widely used as animal feeds, healthcare products and 36
food additives.1,2 Microalgae are more nutritious than traditional animal and aquatic feed like 37
millet, grams and other small fishes in terms of protein, omega 3 fatty acids and carotenoids 38
content.3,4 Several kinds of antioxidant compounds have been extracted from microalgae to 39
protect against oxidative stress, a cause of many diseases and aging.1,5 However, it still remains 40
a challenge to find an effective and especially safe method for microalgae harvesting. 41
Many physical and chemical methods have been tested to harvest microalgae, including 42
sedimentation, centrifugation, filtration, chemical flocculation and electro-flocculation. 43
Sedimentation is simple but time-consuming and only suitable for harvesting large-size 44
microalgae, since most of microalgae have similar density to water, negative surface charge, 45
and thereby low settling velocities.6 Centrifugation and filtration are rapid but less cost-46
effective, limiting their applications at large scale.7,8 Mata et al. (2010) considered that 47
filtration consumes large amounts of membranes and pumping energy, especially for 48
harvesting unicellular small microalgae cells.8 Chemical flocculation and electro-flocculation 49
are rapid and effective, but residual chemical flocculants in the harvested biomass pose 50
potential health risks and limit their utilizations as food and healthcare products.9 So far, there 51
are few safe and effective technologies for microalgae harvesting. 52
Chitosan is the second-most abundant natural biopolymer, which is mainly derived from the 53
shells of shrimp and other crustaceans.10,11 It has received widespread applications in food and 54
pharmaceutical industries due to its biological compatibility, biodegradability, and nontoxic 55
4
properties.12,13 Chitosan has positively charged amine groups and thereby exhibits flocculation 56
potentials for negatively charged microalgae.14 Pan et al. (2011) used chitosan to flocculate 57
microalgae cells and then sink them with the aid of soil particles, achieving effective harmful 58
microalgae removal in Lake Taihu, China.15 Electro-floatation is a separation process in which 59
solids are separated from liquid by micro-bubbles produced from electrode surfaces.16 It has 60
been widely used to remove floatable materials including activated sludge, oil, surfactants, and 61
flocculants etc.17,18 The use of non-sacrificial electrodes in electro-floatation will not introduce 62
chemical flocculants.19,20 Hence, if microalgae are preliminarily flocculated into unstable flocs 63
by chitosan, and then floated by electro-floatation using non-sacrificial electrodes, it is possibly 64
to harvest microalgae biomass safely and effectively. 65
In this study, we proposed a new method for edible microalgae harvesting by coupling chitosan 66
flocculation and electro-floatation using carbon electrodes, and tested it at different operation 67
conditions. The surface charge of microalgae cells was measured to explore the underlying 68
mechanisms. The responses of growth medium to microalgae harvesting were also investigated 69
to evaluate the feasibility of sustainable utilization of culture medium. The objective of this 70
study is to develop a green method for the harvesting of edible microalgae. 71
Materials and methods 72
Microalgae species and culture 73
In this study, Chlorella vulgaris (C. vulgaris), an edible green microalgae species, was chosen 74
to test the new harvesting method.21,22 Here, C. vulgaris cells (FACHB-24) were purchased 75
from the Institute of Hydrobiology, Chinese Academy of Sciences, and cultured in the BG11 76
5
medium, which consists of 500 mg L-1 Bicin, 100 mg L-1 KNO3, 100 mg L-1 b-C3H7O6PNa2, 77
50 mg L-1 NaNO3, 50 mg L-1 Ca(NO3)2•4H2O, 50 mg L-1 MgCl2•6H2O, 40 mg L-1 Na2SO4, 20 78
mg L-1 H3BO3, 5 mg L-1 Na2EDTA, 5 mg L-1 MnCl2•4H2O, 5 mg L-1 CoCl2•6H2O and 0.8 mg 79
L-1 Na2MoO4•2H2O, 0.5 mg L-1 FeCl3•6H2O and 0.5 mg L-1 ZnCl2. The batch cultures were 80
conducted in an illuminating incubator (LRH-250-G, Guangdong Medical Apparatus Co., Ltd., 81
China) with continuous cool white fluorescent light of 2500 ± 500 lux on a 12 h light and 12 h 82
darkness regime at the temperature of 30 ± 1°C. 83
Microalgae harvesting system 84
The microalgae harvesting system consists of a flat stir paddle (Zhongrun Water Industry 85
Technology Development Co., Ltd., China) for mixing during chitosan flocculation and two 86
round carbon electrode plates (Jinjia Metal Co., Ltd., China) for electro-floatation (Fig. 1). The 87
carbon electrode plate has a surface area of 55.4 cm2 and a thickness of 0.2 cm, which was 88
horizontally installed at the bottom with a gap of 2 cm between the two plates. There are 85 89
small round holes on each carbon electrode plate to allow gas bubbles freely pass it during 90
electrolysis, such that the effective surface area was 38.7 cm2. The electric current was supplied 91
by a direct current power supply (DF1730SL5A, Ningbo Zhongce Dftek Electronics Co., Ltd., 92
China). 93
6
94
Fig. 1. The schematic diagram of microalgae harvesting system. 95
Microalgae harvesting test 96
C. vulgaris culture at the exponential growth phase was used in the microalgae harvesting test. 97
The initial cell concentration was set to 3.63 × 1010 cells L-1. 0.5 L of readily prepared C. 98
vulgaris solution was transferred to the harvesting cell. Water-soluble chitosan was purchased 99
from Qingdao Yunzhou Bioengineering Co. Ltd., China. Prior to the test, a chitosan stock 100
solution (2 g L-1) was prepared as follows: 1 g chitosan was added to 0.5 L distilled water and 101
completely diluted by stirring. After chitosan was added, the microalgae solution was stirred 102
at 200 rpm for 2 min and 40 rpm for another 10 min; electro-floatation was started in the last 5 103
min during chitosan flocculation. The microalgae solution was allowed to stand for 10 min, 104
and then water samples were carefully collected from an outlet 2 cm above the carbon electrode 105
plate to enumerate the cell number using an Axioskop 2 mot plus microscope (Carl ZEISS, 106
Germany). The microalgae harvesting efficiency was calculated as (initial cell concentration-107
sample cell concentration)/initial cell concentration × 100%. In the test, the chitosan dosage 108
7
was set to 0, 2, 4, 6, 8, 10, 12 and 15 mg L-1, and the current density was set to 0, 0.2, 0.4 and 109
0.6 A. All the tests were conducted in triplicate at the raw microalgae solution pH of 8.6. 110
The surface charge of microalgae cells was characterized using a Zetasizer 2000 (Malvern Co. 111
United Kingdom). To study the responses of culture medium to microalgae harvesting, medium 112
nutrients (phosphate, ammonium and nitrate) were measured according to Chinese Monitoring 113
Analysis Method of Water and Wastewater;23 medium pH and temperature were measured 114
using a Yellow Springs Instruments (Yellow Springs, Ohio, USA) before and after microalgae 115
harvesting. 116
Cost evaluation 117
The cost of microalgae harvesting was estimated by summing flocculants and energy costs per 118
unit of harvested microalgae biomass as follows: 119
𝐶𝑜𝑠𝑡 = (𝐶𝑜𝑠𝑡𝑓𝑙𝑜𝑐𝑐𝑙𝑢𝑎𝑡𝑖𝑜𝑛 + 𝐶𝑜𝑠𝑡𝑒𝑙𝑒𝑡𝑟𝑜𝑓𝑙𝑜𝑎𝑡𝑎𝑡𝑖𝑜𝑛)/𝑊𝑏𝑖𝑜𝑚𝑎𝑠𝑠 (1) 120
𝐶𝑜𝑠𝑡𝑓𝑙𝑜𝑐𝑐𝑙𝑢𝑎𝑡𝑖𝑜𝑛 = 𝐶𝑜𝑠𝑡𝑐ℎ𝑖𝑡𝑜𝑠𝑎𝑛 + 𝐶𝑜𝑠𝑡𝑚𝑖𝑥𝑡𝑖𝑛𝑔 𝑒𝑛𝑒𝑟𝑔𝑦 (2) 121
𝐶𝑜𝑠𝑡𝑒𝑙𝑒𝑡𝑟𝑜𝑓𝑙𝑜𝑎𝑡𝑎𝑡𝑖𝑜𝑛 = 𝐶𝑜𝑠𝑡𝑒𝑙𝑒𝑡𝑟𝑜𝑙𝑦𝑠𝑖𝑠 𝑒𝑛𝑒𝑟𝑔𝑦 (3) 122
𝐶𝑜𝑠𝑡𝑐ℎ𝑖𝑡𝑜𝑠𝑎𝑛 = 𝐶 ∗ 𝑣 ∗ 𝑎 (4) 123
𝐶𝑜𝑠𝑡𝑚𝑖𝑥𝑖𝑛𝑔 𝑒𝑛𝑒𝑟𝑔𝑦 = 𝑃 ∗ 𝑇 ∗ 𝑏 (5) 124
𝐶𝑜𝑠𝑡𝑒𝑙𝑒𝑡𝑟𝑜𝑙𝑦𝑠𝑖𝑠 𝑒𝑛𝑒𝑟𝑔𝑦 = 𝑈 ∗ 𝐼 ∗ 𝑡 ∗ 𝑏 (6) 125
𝑊𝑏𝑖𝑜𝑚𝑎𝑠𝑠 = 𝑣 ∗ 𝛽 ∗ 𝜃 ∗ 𝜎 (7) 126
where C is the chitosan dosage, mg L-1; v is the volume of microalgae solution, L; a is the 127
chitosan price, which is 0.03 USD g-1 ; P is the stirrer power, which is 40 W; T is the stirring 128
time, h; b is the electric power price, which is 0.08 USD (kWh)-1; U is the electrolysis voltage, 129
8
V; I is the current intensity, A; t is the electrolysis time, h; β is the initial microalgae 130
concentration, cell L-1; θ is the microalgae harvesting efficiency, %; σ is the weight per 131
microalgae cell, 32 × 10-12 g cell-1. 132
Data analysis 133
One-way analysis of variance (ANOVA) was employed to test the statistical significance of 134
differences between treatments. Post-hoc multiple comparisons of treatment means were 135
performed using the Tukey’s least significant difference procedure. All statistical calculations 136
were performed using the SPSS (v22.0) statistical package for personal computers. The level 137
of significance was P < 0.05 for all tests. 138
Results 139
Surface charge properties of chitosan and C. vulgaris cells 140
The isoelectric point of chitosan was pH 9.7, making it possess net positive charges under most 141
microalgae culture conditions. The zeta potential of chitosan kept above +1.5 mV in the wide 142
pH range of < 9.0, and then decreased to nearly zero at pH 9.7. In contrast, the zeta potential 143
of C. vulgaris cells kept below zero in the pH range of > 2.5 (Fig. 2). 144
145
9
Fig. 2. The surface charge properties of chitosan and C. vulgaris cells. Error bars indicate 146
standard deviations. 147
Microalgae harvesting efficiency 148
The use of chitosan flocculation alone only achieved limited microalgae harvesting no matter 149
what the chitosan dosage was. The microalgae harvesting efficiency reached 16.9% at the 150
chitosan dosage of 2 mg L-1 and maintained stably at this value as the chitosan dosage further 151
increased. Similarly, the use of electro-floatation alone could not effectively harvest microalgae 152
either and the harvesting efficiency was only 4.5%, 13.4% and 22.9% at the current intensity 153
of 0.2, 0.4 and 0.6 A, respectively. When chitosan flocculation and electro-floatation were used 154
together, the microalgae harvesting was greatly improved by increasing harvesting efficiency 155
and simultaneously decreasing the optimal chitosan dosage, and this effect was enhanced as 156
the current density increased. When the current intensity of 0.2, 0.4 and 0.6 A was applied, 157
microalgae harvesting efficiency reached a maximum of 33.2%, 59.6% and 63.5%, and the 158
optimal chitosan dosage was 6.0, 4.0 and 2.0 mg L-1, respectively. However, a remarkable 159
decrease in microalgae harvesting efficiency was observed as chitosan was overdosed at the 160
current intensity of 0.4 and 0.6 A, and the harvesting efficiency decreased to 35.2% at the 161
chitosan dosage of 15 mg L-1 (Fig. 3). 162
10
163
Fig. 3. Effects of chitosan dosage and current density on microalgae harvesting efficiency. 164
Error bars indicate standard deviations. 165
Microalgae surface charge 166
After chitosan was added, the zeta potential of microalgae cells showed a remarkable increase. 167
The zeta potential of microalgae cells was gradually increased from –9.2 to –0.9 mV as the 168
chitosan dosage increased from 0 to 15 mg L-1. The use of electro-floatation further increased 169
the zeta potential of microalgae cells, indicating a charging effect, and this effect was enhanced 170
as the current intensity increased (Fig. 4A). When the current density of 0.2, 0.4 and 0.6 A was 171
applied, the zeta potential of microalgae cells was finally increased to +7.3, +8.4 and +10.6 172
mV at the chitosan dosage of 15 mg L-1, respectively. The charging effect was further evaluated 173
by subtracting the zeta potential of microalgae cells in presence of electro-floatation from the 174
value in the absence of electro-floatation. It exhibited chitosan limitation at low chitosan 175
dosages and current limitation at high chitosan dosages (Fig. 4B). As the chitosan dosage 176
increased, the increase amplitude of zeta potential gradually increased and then reached an 177
equilibrium; the use of higher intensity currents yielded a higher equilibrium value at a lower 178
11
chitosan dosage. At the current intensity of 0.2, 0.4 and 0.6 A, the increase amplitude of zeta 179
potential of microalgae cells reached a maximum of 9.3, 11.0 and 11.5 mV, and the optimal 180
chitosan dosage was 2, 4 and 8 mg L-1, respectively. In contrast, in the absence of chitosan, 181
there were no significant differences in zeta potential of microalgae cells among different 182
intensity currents (P < 0.05), and the zeta potential values kept stably at –9.4 mV at the chitosan 183
dosage of 0 mg L-1 (Fig. 4A). 184
185
Fig. 4. Changes of microalgae cell surface change during microalgae harvesting. (A) Zeta 186
potential of microalgae cells; (B) The charging effect of electro-floatation. Error bars indicate 187
standard deviations. 188
Microalgae culture medium 189
Compared with chitosan flocculation alone, ammonium in culture medium was significantly 190
increased after electro-floatation was introduced (P < 0.05), and the use of higher intensity 191
currents yielded higher ammonium concentrations. After microalgae harvesting, ammonium 192
concentration maintained stably below 0.2 mg L-1 in chitosan flocculation alone, and was 193
increased to 1.0, 1.6 and 2.2 mg L-1 when electro-floatation was applied at the current intensity 194
of 0.2, 0.4 and 0.6 A, respectively (Fig. 5A). In contrast, there were no significant changes in 195
12
other parameters observed after microalgae harvesting (P > 0.05). In all the tests, medium 196
nitrate and phosphate maintained stably at 250.9 and 4.66 mg L-1 (Fig. 5B); medium pH, 197
conductivity, and temperature maintained stably at 7.0, 2.4 mS cm-1 and 19.6°C, respectively 198
(Table S2). 199
200
Fig. 5. Changes of nutrients in culture medium after microalgae harvesting. (A) Ammonium; 201
(B) Phosphate and nitrate. Error bars indicate standard deviations. 202
Microalgae harvesting cost 203
Compared with chitosan flocculation (0.08 × 10-3 USD g-1 biomass), it costed much more to 204
harvest microalgae using electro-floatation, reaching 2.83 × 10-3, 2.76 × 10-3 and 3.30 × 10-3 205
USD g-1 biomass at the 0.2, 0.4 and 0.6 A, respectively. When chitosan flocculation and electro-206
floatation were used together, the cost could be greatly reduced, but exhibited a potential 207
increase as the current intensity increased. The cost reached 0.41 × 10-3, 0.64 × 10-3 and 1.18 208
× 10-3 USD g-1 biomass at 0.2, 0.4 and 0.6 A, respectively. 209
13
210
Fig. 6. The cost evaluation for microalgae harvesting using the coupled chitosan flocculation 211
and electro-flocculation. The single red and blue circles indicated chitosan flocculation and 212
electro-floatation were used alone, respectively. The combined red and blue circles indicated 213
that chitosan flocculation and electro-floatation were used together. 214
Discussion 215
The synergistic effect of chitosan flocculation and electro-floatation 216
Microalgae particles have negative surface charge and often stably suspend in the solution with 217
electrostatic repulsion.24,25 Charge neutralization is an essential step in microalgae flocculation, 218
which eliminates energy barrier for microalgae aggregation.26-28 Chitosan is positively charged 219
over a wide range of pH < 9.7, which made it obtain flocculation potential for negatively 220
charged microalgae (Fig. 2). However, most of the formed microalgae flocs still suspended 221
with the aid of buoyancy. The zeta potential of microalgae cells showed a remarkable increase 222
as the chitosan dosage increase (Fig. 4). As a result, the use of chitosan flocculation alone 223
yielded limited microalgae harvesting efficiency whatever the chitosan dosage was (Fig. 3). 224
After electro-floatation was introduced, large amounts of tiny gas bubbles were produced, 225
14
carrying the flocs to water surface where they can be easily collected (Fig. S1). Microalgae 226
harvesting efficiency were therefore remarkably increased when chitosan flocculation and 227
electro-floatation were used together (Fig. 3). However, the use of electro-floatation alone 228
could not effectively harvest microalgae either. This is because microalgae cells stably suspend 229
with electrostatic repulsion, and it is difficult to float them by bubbles without preliminary 230
flocculation.29 231
The positive priming effect of electro-floatation on chitosan flocculation 232
Chitosan was appointed to neutralize microalgae surface charge for microalgae flocculation 233
(Fig.2 and Fig. 4). Surprisingly, after electro-floatation was introduced, the zeta potential of 234
microalgae cells differed at the same chitosan dosage, depending on current intensity applied. 235
The zeta potential of microalgae cells generally exhibited an increase as the current intensity 236
increased, but this effect disappeared in the absence of chitosan (Fig. 4A). It indicated that 237
electrolysis charged chitosan and increased its charge neutralization ability for microalgae 238
flocculation, producing a positive priming effect. This explained that the use of higher current 239
density yielded higher microalgae harvesting efficiency at a lower optimal chitosan dosage. 240
There are a lot of amino groups on the chain of chitosan, which were possibly charged by 241
electrolysis during electro-floatation.30 At the low chitosan dosage, the receptor capacity was 242
limited and the charging effect of electrolysis exhibited a chitosan limitation. As the chitosan 243
dosage increased, chitosan limitation was transferred to current limitation (Fig. 4B), and the 244
stronger current has a higher charging effect. However, excess loads of positive charges can 245
cause microalgae cells positively charged and re-establish electrostatic repulsion.31 Thus, a 246
15
remarkable decrease in microalgae harvesting efficiency was observed as chitosan was 247
overdosed at the high current intensity (0.4 and 0.6 A, Fig. 3). 248
Recommendations for future applications 249
In the microalgae-based engineering, culture medium reuse can offer a promising strategy for 250
saving water and nutrients.32,33 Because of the electrolysis, the physico-chemical properties of 251
culture medium may change after microalgae harvesting.34,35 For instance, water temperature 252
may increase as waste heat releases. However, there were no significant changes in medium 253
pH (P > 0.05), temperature (P > 0.05) and conductivity (P > 0.05) after microalgae harvesting 254
in this study (Table S1), which is possibly due to weak electrolysis (low current intensity and 255
short electrolysis time). As for main nutrients, there was a significant increase in ammonium 256
(P < 0.05) (Fig. 5A). We attributed it to the transformation from nitrate to ammonium under 257
electrolysis according to our additional experiments (Fig. S2). During electrolysis, nitrate 258
reduction (NO3- + 10H+ + 8e- = NH4
+ + 3H2O) can occur at the cathode, releasing ammonium 259
to the medium.36,37 Thus, the stronger current had a higher ammonium yield (Fig.5A and Fig. 260
S2). However, we did not detect the significant decrease in nitrate (P > 0.05, Fig. 5B) because 261
of the extremely high levels of nitrate in the BG11 medium (> 250 mg L-1). The shift of nitrate 262
to ammonium can increase nitrogen bioavailability since ammonium is generally favored by 263
microalgae relative to nitrate.38-40 In contrast, phosphate did not show significant changes after 264
microalgae harvesting (P > 0.05, Fig. 5B) due to the use of non-sacrificial electrodes, which 265
differs with the decrease of phosphate concentration by electrolysis using Fe/Al electrodes.41,42 266
16
Hence, nutrient bioavailability increase and other parameter stability (Fig. 5B, Table S2) make 267
medium reusable for microalgae continuous culture. 268
In practical applications, cost is often another major concern for a technology. For the 269
microalgae harvesting technology in this study, there was a trade-off between the harvesting 270
efficiency and the cost, since the cost exhibited a potential increase as the current intensity 271
increased (Fig. 6). It will be cost-effective to apply the low current intensity to harvest 272
microalgae in the continuous system, and the remaining cells can benefit microalgae recovery. 273
Despite the microalgae harvesting efficiency was greatly increased by coupling chitosan 274
flocculation and electro-floatation (Fig. 3), it may be further increased by optimizing operation 275
condition or screening other edible flocculants, such as moringa oleifera and tannin.43,44 276
Conclusions 277
Microalgae harvesting is a crucial step but still remains a challenge for microalgae-based 278
engineering. This study proposed a green way to harvest microalgae by coupling chitosan 279
flocculation and electro-floatation. Microalgae can be preliminarily flocculated into unstable 280
flocs by chitosan, and then floated by electro-floatation; electro-floatation charged chitosan and 281
activated it to flocculate microalgae, producing a positive priming effect. The use of edible 282
chitosan and inert carbon electrodes makes it possible to harvest microalgae biomass safely 283
and effectively for food or healthcare use and achieve the sustainable utilization of culture 284
medium. Further studies are needed to optimize operation conditions to increase harvesting 285
efficiency and reduce the cost. 286
Acknowledgments 287
17
This study was supported by the National Natural Science Foundation of China (No. 41701112, 288
51709181, 51979171), Starting Research Fund of Nanjing University of Information 289
Engineering (No. 20181015), Major Science and Technology Program for Water Pollution 290
Control and Treatment (No. 2017ZX07501-002, 2017ZX07108-002). 291
Supporting Information 292
The Supporting Information is available free of charge on the web. 293
The microalgae floc image; the effects of electrolysis using carbon electrodes on BG11 medium; 294
changes of physical properties of growth medium after microalgae harvesting. 295
Notes 296
The authors declare no competing financial interest. 297
298
18
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For Table of Contents Use Only 424
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Synopsis: A green method for harvesting microalgae biomass will be beneficial to microalgae-426
based engineering for healthcare and food use. 427
download fileview on ChemRxivACS-MS-20200614.pdf (0.95 MiB)
Supporting Information
Sustainable harvesting of microalgae by coupling chitosan flocculation and electro-floatation
Lin Zhu†, Hui Xu‡, Weijie Guo§, Jianghua Yu†, Wenqing Shi†,*
†Jiangsu Collaborative Innovation Center of Atmospheric Environment and Equipment
Technologies, Jiangsu Key Laboratory of Atmospheric Environmental Monitoring & Pollution
Control, School of Environmental Science & Engineering, Nanjing University of Information
Science & Technology, Nanjing 210044, China
‡Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences, Beijing 100085,
China
§Basin Water Environmental Research Department, Changjiang River Scientific Research Institute,
Wuhan 430010, China
*Corresponding author: Wenqing Shi ([email protected])
Fig. S1. The microalgae floc image. For the floc image study, the flocs were carefully transferred
onto a glass slide and then photographed by an electromotive microscope (ST-CV320, Chongqing
UOP Photoelectric Technology Co., Ltd., China).
Fig. S2. The effects of electrolysis using carbon electrodes on BG11 medium. Three potential
precursors of ammonium in the BG11 medium were tested using the electro-floatation cell,
including sodium nitrate (NaNO3), ammonium ferric citrate (EDTA-2Na) and
ethylenediaminetetraacetic acid disodium salt (C6H8FeNO7). Sodium chloride (NaCl) was used as
electrolyte for EDTA-2Na and C6H8FeNO7 electrolysis. The concentrations of NaNO3, EDTA-2Na
and C6H8FeNO7 were the same as those in the BG11 medium. The molar concentration of NaCl
was the same as that of NaNO3. Error bars indicate standard deviations.
Table S1. Changes of physical properties of growth medium after microalgae harvesting
Current
(A)
Chitosan dosage (mg L-1)
0 2 4 6 8 10 12 15
pH
0 6.93 6.92 7.12 7.26 7.17 7.19 7.16 7.10
0.2 7.25 7.32 7.29 7.13 7.06 6.77 6.92 6.89
0.4 7.02 6.88 6.94 7.02 6.88 6.98 7.02 6.86
0.6 6.84 6.92 6.92 6.82 6.77 6.61 7.36 6.67
Conductivity
（mS cm-1）
0 2.46 2.42 2.43 2.43 2.41 2.46 2.45 2.44
0.2 2.43 2.44 2.44 2.48 2.44 2.43 2.45 2.43
0.4 2.45 2.47 2.44 2.44 2.42 2.46 2.45 2.44
0.6 2.48 2.49 2.47 2.49 2.47 2.45 2.46 2.45
Temperature
(°C)
0 19.0 19.4 19.0 18.9 18.8 18.4 18.5 19.3
0.2 19.2 19.7 19.4 19.9 19.7 20.0 19.5 19.9
0.4 19.7 20.2 19.6 19.7 19.9 19.5 19.5 19.9
0.6 19.4 19.4 20.0 20.2 20.2 20.1 20.4 20.5
download fileview on ChemRxivACS-SI-20200614.pdf (249.37 KiB)
Supporting Information
Sustainable harvesting of microalgae by coupling chitosan flocculation and electro-
floatation
Lin Zhu†, Hui Xu‡, Weijie Guo§, Jianghua Yu†, Wenqing Shi†,*
†Jiangsu Collaborative Innovation Center of Atmospheric Environment and Equipment
Technologies, Jiangsu Key Laboratory of Atmospheric Environmental Monitoring & Pollution
Control, School of Environmental Science & Engineering, Nanjing University of Information
Science & Technology, Nanjing 210044, China‡Research Center for Eco-Environmental Sciences, Chinese Academy of Sciences, Beijing
100085, China§Basin Water Environmental Research Department, Changjiang River Scientific Research
Institute, Wuhan 430010, China
*Corresponding author: Wenqing Shi ([email protected])
Fig. S1. The microalgae floc image. For the floc image study, the flocs were carefully transferred
onto a glass slide and then photographed by an electromotive microscope (ST-CV320,
Chongqing UOP Photoelectric Technology Co., Ltd., China).
Fig. S2. The effects of electrolysis using carbon electrodes on BG11 medium. Three potential
precursors of ammonium in the BG11 medium were tested using the electro-floatation cell,
including sodium nitrate (NaNO3), ammonium ferric citrate (EDTA-2Na) and
ethylenediaminetetraacetic acid disodium salt (C6H8FeNO7). Sodium chloride (NaCl) was used
as electrolyte for EDTA-2Na and C6H8FeNO7 electrolysis. The concentrations of NaNO3, EDTA-
2Na and C6H8FeNO7 were the same as those in the BG11 medium. The molar concentration of
NaCl was the same as that of NaNO3. Error bars indicate standard deviations.
Table S1. Changes of physical properties of growth medium after microalgae harvesting
Current
(A)
Chitosan dosage (mg L-1)
0 2 4 6 8 10 12 15
pH
0 6.93 6.92 7.12 7.26 7.17 7.19 7.16 7.10
0.2 7.25 7.32 7.29 7.13 7.06 6.77 6.92 6.89
0.4 7.02 6.88 6.94 7.02 6.88 6.98 7.02 6.86
0.6 6.84 6.92 6.92 6.82 6.77 6.61 7.36 6.67
Conductivity（mS cm-1）
0 2.46 2.42 2.43 2.43 2.41 2.46 2.45 2.44
0.2 2.43 2.44 2.44 2.48 2.44 2.43 2.45 2.43
0.4 2.45 2.47 2.44 2.44 2.42 2.46 2.45 2.44
0.6 2.48 2.49 2.47 2.49 2.47 2.45 2.46 2.45
Temperature(°C)
0 19.0 19.4 19.0 18.9 18.8 18.4 18.5 19.3
0.2 19.2 19.7 19.4 19.9 19.7 20.0 19.5 19.9
0.4 19.7 20.2 19.6 19.7 19.9 19.5 19.5 19.9
0.6 19.4 19.4 20.0 20.2 20.2 20.1 20.4 20.5
download fileview on ChemRxivACS-SI-20200614.docx (235.67 KiB)
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