- Open Access
Cultivation of lipid-producing microalgae in struvite-precipitated liquid digestate for biodiesel production
© The Author(s) 2018
Received: 27 December 2017
Accepted: 31 March 2018
Published: 7 April 2018
Using liquid digestate from the biogas industry as a medium to culture lipid-producing microalgae is considered mutually beneficial for digestate valorization and for reducing the cost of microalgal cultivation. However, the low transmittance and high ammonium (NH4+-N) levels in liquid digestate negatively influence microalgae growth.
Struvite precipitation was used to pretreat liquid digestate. To obtain struvite-precipitated supernatant with an ideal transmittance, NH4+-N concentration, salinity, and N:P ratio for microalgal growth, there should be a 1:1.2:1.2 NH4+:Mg2+:PO43− molar ratio in the liquid digestate, with KH2PO4 and MgCl2 added through continuous stirring. The addition and stirring was subsequently stopped when the pH reached 8.5. Of the nine tested microalgae species, Dictyosphaerium ehrenbergianum exhibited the best growth in the supernatant. The biomass productivity and lipid content of D. ehrenbergianum cultured in the struvite-precipitated supernatant were 161.06 mg/l/days and 34.33%, respectively, which was higher than when cultured in the standard BG-11 medium. Moreover, the struvite-precipitated supernatant improved the accumulation of monounsaturated fatty acids and saturated fatty acids.
Microalgae are considered one of the most promising biofuel feedstocks [1, 2]. However, the high cultivation cost is still a limiting factor for its further commercialization . Using wastewater for microalgal cultures is considered mutually beneficial, because the wastewater provides nutrients for microalgal growth, while the microalgae remove pollutants from the wastewater [4, 5].
Digestate is an important byproduct of the anaerobic digestion of organic waste [5, 6]. Traditionally, land application is the primary method for managing digestate from livestock farms . However, with the rapid development of the biogas industry, the volume of digestate has increased substantially in recent years. Land application and other digestate processing techniques require large energy inputs and increase environmental risk, especially with respect to liquid digestate from large scale biogas plants in large livestock or poultry farms . Digestate management has become a major bottleneck in the development of the biogas industry, as well as for the livestock and poultry breeding industry . On the other hand, the liquid digestate is rich in nitrogen (N), phosphorous (P), potassium (K), and other nutrients essential for microalgal growth ; thus, its use as a microalgal culture medium is considered a new opportunity for digestate valorization [6, 9].
Culturing microalgae in liquid digestate can reduce the cost of nutrients for microalgal cultivation, while simultaneously reusing liquid digestate ; however, there are some limiting and inhibitory factors in liquid digestate-based microalgal cultivation. First, the high turbidity of liquid digestate, caused by suspended materials, can lead to low transmittance. This reduces the efficiency of photosynthesis and the growth of microalgae . Moreover, the ammonium-nitrogen (NH4+-N) levels in liquid digestate are usually high and can potentially inhibit microalgal growth . Pretreatments, such as separation and dilution, are often used to reduce the negative effects associated with high NH4+-N and turbidity; however, these processes consume a large quantity of energy and fresh water .
Struvite (magnesium ammonium phosphate; MAP) precipitation can reduce NH4+ and suspended solids (SS) in wastewater under alkaline conditions, while simultaneously generating slow-release fertilizer [12, 13]. This technique has been also used to recover N from digestate . However, the pH of liquid digestate is usually neutral or weak alkaline, and it has less PO43− and Mg2+ compared to NH4+. As such, chemicals containing PO43−, Mg2+, or OH− must be added to facilitate effective NH4+ removal [15, 16]. As a result, the candidate supernatant for microalgal culture following precipitation usually has a high pH and salinity, as well as an unsatisfactory N:P ratio for microalgal growth. The suggested optimal N:P mass ratio for microalgal growth is approximately 7:1, based on the composition of microalgae ; however, the N:P molar ratio in liquid digestate is usually adjusted to approximately 1:1 (mass ratio around 0.45) at the beginning of the reaction to achieve a high precipitation efficiency [18, 19]. In fact, the remaining N and P levels, as well as the N:P ratio in the supernatant, are under the control of the reaction conditions, such as pH value and PO43−:Mg2+:NH4+ molar ratios . Thus, one goal of this study was to determine the optimal combination of chemical additives and reaction conditions of struvite precipitation in liquid digestate, to obtain an optimal supernatant for microalgal growth.
Most microalgae prefer neutral environments; as such, the high pH and salinity in the struvite-precipitated supernatant may negatively affect algal growth. However, reducing the pH and salinity in the supernatant is usually not economical. Microalgal species have a range of optimal pH and salinities, with some species tolerating high pH and salt conditions . Selecting high pH and salt tolerant species is beneficial for culturing microalgae in liquid digestate after struvite precipitation. Therefore, in this study, struvite precipitation was selected as a digestate pretreatment technique to determine the optimal combination of chemical additives and reaction conditions, and to investigate suitable microalgal species. This provides a new way for combining liquid digestate treatment and microalgal cultivation .
Liquid digestate collection and characterization
The liquid digestate used in this study was obtained from a pig farm located in Jianyang, Sichuan province in China. The samples were collected from a storage pond after treatment in an anaerobic continuous stirred tank reactor (CSTR) with the raw materials of swine wastewater after separation. They were immediately transported to the laboratory and stored at 4 °C until use.
Microalgal strains and growth medium
Nine microalgal strains were collected from the Freshwater Algae Culture Collection at the Institute of Hydrobiology (FACHB-Collection, Wuhan, China), including Chlorella regularis FACHB-1068, Chlorella pyrenoidosa FACHB-9, Botryococcus braunii FACHB-357, Scenedesmus obliquus FACHB-417, Dictyosphaerium ehrenbergianum FACHB-1223, Haematococcus pluvialis FACHB-712, Spirulina subsalsa FACHB-351, Spirulina platensis FACHB-900, and Spirulina maxima FACHB-438. The three Spirulina strains were cultured in the Spirulina medium  and the other strains were cultured in BG-11 medium .
Effect of KH2PO4 and NaH2PO4 on transmittance, NH4+-N removal rate, and salinity
NH4+-N removal rate (%)
KH2PO4 + MgCl2
80.83 ± 0.43a
0.67 ± 0.03a
93.18 ± 0.74a
NaH2PO4 + MgCl2
74.76 ± 0.11b
0.77 ± 0.05b
89.18 ± 0.02b
To evaluate the effect of the Mg2+ and PO43− sources and stirring time on struvite reaction, 300 ml of liquid digestate were fed into a 500-ml beaker. After this, different combinations of Mg2+ and PO43− sources were added and mixed by continuous stirring with a magnetic stirrer at a constant rate (150 rpm) to make the PO43−:Mg2+:NH4+ molar ratios of each solution at 1:1.2:1.2. When the exogenous compounds were dissolved, the pH of the mixture was adjusted to 9.0 using 0.1-M NaOH . The stirring continued for 0, 1, 2, 5, 10, and 20 min after the pH reached 9.0; stirring then stopped to allow sampling. A 10-ml sample of each mixture was placed into its own 15-ml centrifuge tube. After settling for 30 min, the supernatant from each centrifuge tube was used for analysis.
To select the best precipitation conditions for optimal microalgal growth in terms of residual NH4+-N concentration and N:P mass ratio, further experiments were done at different pH values (8.0, 8.5, and 9.0) and at different P:Mg:N molar ratios (1:1:0.95, 1:1:1, and 1:1.2:1.2) with the ideal Mg2+ and PO43− sources and stirring pattern identified above.
The struvite-precipitated supernatant obtained above acted as a nutrient source to cultivate nine microalgal strains. Each strain was cultivated in an Erlenmeyer flask (250 ml) as a single batch (100-ml working volume) at a constant temperature (25 °C). A 12-h light/12-h dark cycle was provided using daylight fluorescent tubes with a photon flux density of 40–50 μmol/m2/s. The cultures were manually shaken 2–3 times per day to prevent biomass sedimentation as described previously . The biomass in each Erlenmeyer flask was after 7-day cultivation.
Once a suitable microalgal strain was chosen, laboratory-scale cultivations were conducted in 1.2 l reactors (1 l working volume) for 10-day cultivation. Either struvite-precipitated supernatant or BG-11 was used as the culture medium. An ambient air flow of 0.2 l/min was provided to each reactor in a 12 h/12 h cycle, consistent with the photoperiod described above.
Water quality analysis
The pH value and salinity of the solution was measured using a pH meter and conductivity meter, respectively. Transmittance was measured using spectrophotometry at 680 nm . Chemical oxygen demand (COD) was determined according to standard methods described by APHA . After filtration through a 0.45 μm membrane, the concentration of NH4+-N and NO3−-N was analyzed using an AA-3 autoanalyzer (Bran + Luebbe, Germany). PO43−-P was measured by the molybdate-ascorbic acid colorimetric method described by APHA . Scanning electron microscopy and X-ray diffraction analysis were used to analyze the struvite precipitate.
The struvite precipitate collected from the bottom of the beakers was dried at room temperature (25 ± 2 °C) and ground using a mortar. The powder (with particles that passed through a 200 mesh) was imaged using scanning electron microscopy (Hitachi SU1510, Japan) with an energy-dispersive spectrometer system (SEM–EDS) (Horiba EX-250, Japan). The crystal structures of struvite precipitation were measured using X-ray powder diffractometer (XRD) (Bruker D8 ADVANCE, Germany). The scattering was operated at a power level of 60 kV and at 80 mA. The data were recorded at a speed of 4°/min over the angular range of 10°–60°.
Microalgal growth and total lipid analysis
In this expression, Xfinal and X0 are DCW (mg/l) at the first (t0) and last time point (tfinal), respectively.
This calculation applied two time intervals of DCW (mg/l).
Fatty acid profile analysis
Fatty acid content and composition analysis were determined in two consecutive steps, including the preparation of fatty acid methyl ester (FAME) and the analysis using Gas Chromatography–Mass Spectrometry (GC–MS) (Agilent, USA). FAME was prepared using a one-step extraction–transesterification method, as described by Indarti et al. , with a minor modification. Dried microalgal samples (approximately 500 mg) were weighed into clean, 50-ml screw-top glass bottles, to which a 20 ml mixture of methanol, concentrated sulfuric acid, and chloroform (4.25:0.75:5) were added. Transesterification was carried out in a 90 °C water bath for 90 min. Once the reaction was completed, the chloroform layer containing the FAME was carefully collected for GC–MS analysis. The oven temperature was set at 80 °C, held steady for 5 min, was then raised to 290 °C at a rate of 4 °C/min, and was then held at 290 °C for 5 min. The resulting compounds were identified in the NIST Mass Spectral Database and quantified by the area normalization method.
Each experiment was performed in triplicate and was repeated at least three times. The experimental results were reported as the mean value of each parameter with standard deviation. Statistical analysis was performed using a one-way analysis of variance (ANOVA) followed by a Tukey pairwise comparison, using Origin 10.0. A p value < 0.05 was considered statistically significant.
Results and discussion
Optimization of struvite precipitation
The concentrations of COD, NH4+-N, NO3−-N, and PO43−-P in the digestate used in this study were 629.05, 591.2, 0.07, and 9.87 mg/l, respectively. The pH value was 8.2; the transmittance was 0.11%; and the salinity was 0.26%.
Additional Mg2+ and PO43− ions were required to achieve high struvite precipitation efficiency in the liquid digestate. The sources of Mg2+ and PO43− significantly affected the quality of the struvite-precipitated supernatant . In this study, six combinations were first tested at a NH4+:Mg2+:PO43− molar ratio of 1:1.2:1.2 and a pH of 9.0, as described by Perera et al. . Figure 1 shows that the combination of KH2PO4 + MgCl2 resulted in the highest transmittance at the first test time point (0 min), reaching 80%; this combination was followed by KH2PO4 + MgSO4, K2HPO4 + MgCl2, and K2HPO4 + MgSO4, which resulted in transmittances of more than 60%. The two MgO combinations exhibited low transmittance due to the low solubility, while the K2HPO4 + MgO combination resulted in the lowest transmittance of approximately 10%. This indicated that KH2PO4 and MgCl2 were the better choices for obtaining supernatant with high transmittance after struvite precipitation.
In addition, the maximum transmittance of each combination occurred when the pH reached 9.0, and it gradually decreased with continued stirring (Fig. 1). The struvite formed flocs and rapid precipitation reduced the suspended solids, thereby improving the transmittance of the liquid digestate. Continuous stirring could destroy the flocs, causing suspended fine particles that could decrease the transmittance. Therefore, in subsequent experiments, the stirring was stopped as soon as the pH reached the predetermined value.
In nature, microalgae have a large species diversity, allowing them to adapt to salinity changes. In the open ocean, salinity varies between 3.3 and 3.7% , which is significantly higher than the salinity obtained in this study. Moreover, salt stress could increase the lipid content of some microalgal species . Of the six combinations, we selected KH2PO4 + MgCl2 for further investigation. Other studies have confirmed that the dominant form of P in MAP formation reaction is H2PO4− or HPO42− . In this study, the H2PO4− achieved better results with higher transmittance, NH4+-N removal rate, and lower salinity. Yetilmezsoy and Sapci-Zengin  found similar results when recovering NH4+-N from the effluent of a UASB treating poultry manure using MAP precipitation.
NaH2PO4 is often used as a PO43− source for struvite precipitation . As such, we further analyzed the differences between NaH2PO4 and KH2PO4. The results indicate that the KH2PO4 + MgCl2 combination achieves higher transmittance, higher NH4+ removal rate, and lower salinity (Table 1). The MgKPO4·6H2O (MKP) is one of the struvite analogs, but it is more likely to form if NH4+ concentrations are low [31, 33]. The liquid digestate used in this study was high in NH4+; the presence of K+ did not interfere with removing NH4+. Otherwise, potassium is an essential macronutrient and is most abundantly absorbed cation playing an important role in algae growth.
Characteristics of struvite-precipitated supernatant under different NH4+:Mg2+:PO43− molar ratio and pH
NH4+:Mg2+:PO43− molar ratio
N:P mass ratio
NH4+-N removal rate (%)
142.52 ± 2.10a
0.692 ± 0.021a
35.47 ± 0.48a
4.12 ± 0.68a
70.00 ± 0.76a
0.72 ± 0.21a
81.18 ± 0.37a
118.71 ± 0.48b
0.771 ± 0.049a
29.87 ± 0.32b
4.05 ± 0.22a
71.12 ± 0.66a
0.69 ± 0.11b
83.33 ± 0.08b
78.06 ± 0.25c
0.755 ± 0.074a
26.24 ± 0.70c
3.06 ± 0.42b
71.78 ± 0.74a
0.70 ± 0.01ab
86.41 ± 0.04c
90.07 ± 0.08d
0.030 ± 0.019b
34.12 ± 0.39d
2.70 ± 0.73b
74.65 ± 1.33b
0.67 ± 0.07b
85.32 ± 0.01d
83.78 ± 0.82e
0.092 ± 0.076b
22.78 ± 0.51e
3.75 ± 0.04b
77.36 ± 1.34b
0.61 ± 0.08b
86.41 ± 0.14c
66.56 ± 0.52f
0.063 ± 0.048b
16.36 ± 0.07f
4.16 ± 0.23a
74.82 ± 0.86b
0.62 ± 0.08b
88.41 ± 0.09e
71.08 ± 1.38 g
0.053 ± 0.032b
18.92 ± 1.52 g
3.86 ± 0.57b
72.19 ± 1.53a
0.64 ± 0.03b
89.62 ± 0.24f
57.85 ± 0.66 h
0.046 ± 0.042b
8.51 ± 0.84 h
7.04 ± 0.72c
78.28 ± 0.38bc
0.66 ± 0.03b
91.63 ± 0.11 g
38.95 ± 0.74i
0.071 ± 0.016b
7.02 ± 1.21 h
5.65 ± 0.43d
79.60 ± 0.01d
0.67 ± 0.02b
93.22 ± 0.13 h
The initial NH4+-N concentration of the liquid digestate used in this study was 591.2 mg/l. As more PO43− and Mg2+ was added, less NH4+-N remained. In addition, at the same NH4+:Mg2+:PO43− molar ratio, the remaining NH4+-N concentration decreased with an increasing pH value. At a NH4+:Mg2+:PO43− ratio of 1:1.2:1.2 and pH of 9.0, the NH4+-N concentration in supernatant was as low as 38.95 mg/l, indicating a NH4+ removal rate of more than 90% (Table 2). This led to the hypothesis that this method could be used to treat liquid digestates containing up to 1000 mg/l of NH4+-N, to meet the 100-mg/l NH4+-N concentration requirements for microalgal growth.
The N:P ratio is another important factor affecting microalgae growth. According to the typical microalgae composition formula (C106H181O45N16P), optimal microalgae growth occurs when the mass ratio of N to P that can be absorbed by microalgae is approximately 7:1 . NH4+-N, NO3−-N, and PO43−-P are the main forms of N and P absorbed by microalgae ; as such, the ratio of the sum of NH4+-N and NO3−-N to PO43−-P was used to evaluate the N:P ratio in this study. The N:P mass ratio in the supernatants ranged from 2.70 to 7.04 (Table 2). At a NH4+:Mg2+:PO43− molar ratio of 1:1.2:1.2 and a pH of 8.5, the N:P mass ratio in the supernatant was 7.04, approaching the ideal value. Moreover, at this condition, the supernatant had the second highest transmittance and NH4+ removal rate.
In summary, to achieve optimal transmittance, NH4+-N levels and N:P mass ratio for microalgal growth, the NH4+:Mg2+:PO43− ratio of the liquid digestate should first be adjusted to 1:1.2:1.2 using KH2PO4 and MgCl2 with continuous stirring. The pH should then be adjusted to 8.5 by adding NaOH. At that point, the stirring should be stopped. The supernatant obtained under this condition was used for the subsequent microalgae culture.
Screening of suitable algal species
Specific growth rate (μ) of different microalgae strains cultured in struvite-precipitated supernatant
Specific growth rate
Chlorella regularis FACHB-1068
0.097 ± 0.016a
C. pyrenoidosa FACHB-9
Botryococcus braunii FACHB-357
Scenedesmus obliquus FACHB-417
0.089 ± 0.026a
Dictyosphaerium ehrenbergianum FACHB-1223
0.144 ± 0.025b
Haematococcus pluvialis FACHB-712
Spirulina subsalsa FACHB-351
0.069 ± 0.031c
S. platensis FACHB-900
S. maxima FACHB-438
0.084 ± 0.015a
The genus Dictyosphaerium is found in both marine and fresh water environments  and some species in this genus have a strong ability to adapt to extreme environments . For example, D. chlorelloides can survive in alkaline and moderately acidified aquatic environments containing hexavalent chromium  and Dictyosphaerium sp. has been found in high rate algal ponds (HRAPs) used for wastewater treatment . D. ehrenbergianum was designated as a type species of the genus of Dictyosphaerium . To date, research on this species has mainly focused on its taxonomy [40, 41]; therefore, this study provides a novel application for this alga.
Growth of D. ehrenbergianum in struvite-precipitated supernatant
Biomass productivity, lipid content, lipid productivity, and fatty acid profiles of Dictyosphaerium ehrenbergianum in struvite-precipitated supernatant and BG-11
In struvite-precipitated supernatant
Biomass productivity (mg/l/days)
161.06 ± 3.71
84.87 ± 6.24
Lipid content (% DW)
34.33 ± 1.52
23.67 ± 1.15
Lipid productivity (mg/l/days)
55.29 ± 2.37
20.09 ± 1.83
Fatty acids (% of total fatty acids)
21.09 ± 1.04
17.46 ± 0.62
3.27 ± 0.16
1.32 ± 0.33
7.43 ± 0.13
2.41 ± 0.47
13.48 ± 0.63
11.05 ± 0.59
7.05 ± 0.75
8.60 ± 0.50
9.46 ± 0.60
15.42 ± 1.21
17.76 ± 0.73
20.23 ± 1.21
19.37 ± 0.73
22.24 ± 1.09
In this study, the lipid level of D. ehrenbergianum cultured in BG-11 was 23.67%, which is close to the level found for Dictyosphaerium CFR 5-01/FW by Vidyashankar et al. . However, the lipid content of D. ehrenbergianum cultured in the struvite-precipitated supernatant was as high as 34.33%, resulting in lipid productivity of 55.29 mg/l/days. This indicates that the supernatant was also beneficial for D. ehrenbergianum lipid accumulation. Other research has shown that lipid accumulation can be promoted when microalgal cells are cultivated in stressed conditions, such as high salinity . By adding KH2PO4 and MgCl2, the presence of K+, and other ions, increased the salinity (from 0.26 to 0.66%) of the supernatant following struvite precipitation; therefore, the increased salinity in the supernatant did not inhibit microalgae growth, but actually contributed to greater lipid accumulation.
The predominant fatty acids of D. ehrenbergianum cultured in both the struvite-precipitated supernatant and BG-11 were C16 and C18 (Table 4), which are the main components of biodiesels . This indicates that the oil from D. ehrenbergianum is an ideal biodiesel alternative. The microalgae biodiesels usually have poor oxidative stability due to the low monounsaturated fatty acid (MUFA) and saturated fatty acid (SaFA) content . The proportion of total MUFAs and SaFAs of D. ehrenbergianum increased from 32.24% in BG-11 to 45.27% in the supernatant, whereas the proportion of polyunsaturated fatty acid (PUFA) was decreased. This indicated that the lipid profiles of D. ehrenbergianum could change with growth conditions, the supernatant generated through in this study was more favorable for culturing D. ehrenbergianum for ideal biodiesel production. Similar results were also found in the previous study .
This study constructed a new way to combine liquid digestate treatment and microalgal cultivation. The study also determined the optimal struvite precipitation conditions for pretreating liquid digestate intended for use as a microalgal culture medium. KH2PO4 and MgCl2 were the optimum source of Mg2+ and PO43− to adjust the NH4+:Mg2+:PO43− molar ratio to 1:1.2:1.2. Continued stirring should be stopped when the pH reaches 8.5. D. ehrenbergianum grew best in the struvite-precipitated supernatant, which enhanced D. ehrenbergianum biomass productivity and lipid content, and also improved the accumulation of MUFAs and SaFAs. The struvite-precipitated liquid digestate can be used to culture D. ehrenbergianum for biodiesel production.
YJ and WW prepared the manuscript. WW and LD designed the research. YJ, XP, DZ, TZ, and SW carried out the experimental works. All authors read and approved the final manuscript.
The study was supported by the China Agriculture Research System [Grant Number CARS-35], Fundamental Research Funds for Central Non-profit Scientific Institution [Grant Numbers Y2016CG34, 12017206030202203, and 12017206030202205], and Sichuan Science and Technology Project [Grant Number 2016JY0154].
The authors declare that there are no competing interests.
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- Wijffels RH, Barbosa MJ. An outlook on microalgal biofuels. Science. 2010;329:796–9.View ArticleGoogle Scholar
- Williams PJL, Laurens LML. Microalgae as biodiesel & biomass feedstocks: review & analysis of the biochemistry, energetics & economics. Energ Environ Sci. 2010;3:554–90.View ArticleGoogle Scholar
- Zhu L. Microalgal culture strategies for biofuel production: a review. Biofuels Bioprod Bioref. 2015;9:801–14.View ArticleGoogle Scholar
- Delrue F, Alvarez-Diaz PD, Fon-Sing S, Fleury G, Sassi JF. The environmental biorefinery: using microalgae to remediate wastewater, a win–win paradigm. Energies. 2016;9:3.View ArticleGoogle Scholar
- Chiu SY, Kao CY, Chen TY, Chang YB, Kuo CM, Lin CS. Cultivation of microalgal Chlorella for biomass and lipid production using wastewater as nutrient resource. Bioresour Technol. 2014;184:179–89.View ArticleGoogle Scholar
- Monlau F, Sambusiti C, Ficara E, Aboulkas A, Barakat A, Carrere H. New opportunities for agricultural digestate valorization: current situation and perspectives. Energ Environ Sci. 2015;8:2600–21.View ArticleGoogle Scholar
- Nkoa R. Agricultural benefits and environmental risks of soil fertilization with anaerobic digestates: a review. Agron Sustain Dev. 2013;34:473–92.View ArticleGoogle Scholar
- Fuchs W, Drosg B. Assessment of the state of the art of technologies for the processing of digestate residue from anaerobic digesters. Water Sci Technol. 2013;67:1984–93.View ArticleGoogle Scholar
- Xia A, Murphy JD. Microalgal cultivation in treating liquid digestate from biogas systems. Trends Biotechnol. 2016;34:264–75.View ArticleGoogle Scholar
- Wang L, Li Y, Chen P, Min M, Chen Y, Zhu J, et al. Anaerobic digested dairy manure as a nutrient supplement for cultivation of oil-rich green microalgae Chlorella sp. Bioresour Technol. 2010;101:2623–8.View ArticleGoogle Scholar
- Uggetti E, Sialve B, Latrille E, Steyer JP. Anaerobic digestate as substrate for microalgae culture: the role of ammonium concentration on the microalgae productivity. Bioresour Technol. 2014;152:437–43.View ArticleGoogle Scholar
- Darwish M, Aris A, Puteh MH, Abideen MZ, Othman MN. Ammonium–nitrogen recovery from wastewater by struvite crystallization technology. Sep Purif Rev. 2015;45:261–74.View ArticleGoogle Scholar
- Talboys PJ, Heppell J, Roose T, Healey JR, Jones DL, Withers PJ. Struvite: a slow-release fertiliser for sustainable phosphorus management. Plant Soil. 2016;401:109–23.View ArticleGoogle Scholar
- Estevez MM, Linjordet R, Horn SJ, Morken J. Improving nutrient fixation and dry matter content of an ammonium-rich anaerobic digestion effluent by struvite formation and clay adsorption. Water Sci Technol. 2014;70:337–44.View ArticleGoogle Scholar
- Cheng J, Xu J, Huang Y, Li Y, Zhou J, Cen K. Growth optimisation of microalga mutant at high CO(2) concentration to purify undiluted anaerobic digestion effluent of swine manure. Bioresour Technol. 2015;177:240–6.View ArticleGoogle Scholar
- Nelson NO, Mikkelsen RL, Hesterberg DL. Struvite precipitation in anaerobic swine lagoon liquid: effect of pH and Mg:P ratio and determination of rate constant. Bioresour Technol. 2003;89:229–36.View ArticleGoogle Scholar
- Karapinar Kapdan I, Aslan S. Application of the Stover–Kincannon kinetic model to nitrogen removal by Chlorella vulgaris in a continuously operated immobilized photobioreactor system. J Chem Technol Biotechnol. 2008;83:998–1005.View ArticleGoogle Scholar
- Cerrillo M, Palatsi J, Comas J, Vicens J, Bonmatí A. Struvite precipitation as a technology to be integrated in a manure anaerobic digestion treatment plant—removal efficiency, crystal characterization and agricultural assessment. J Chem Technol Biotechnol. 2015;90:1135–43.View ArticleGoogle Scholar
- Perera PWA, Han ZY, Chen YX, Wu WX. Recovery of nitrogen and phosphorous as struvite from swine waste biogas digester effluent. Biomed Environ Sci. 2007;20:343–50.Google Scholar
- Khatoon H, Abdu Rahman N, Banerjee S, Harun N, Suleiman SS, Zakaria NH, et al. Effects of different salinities and pH on the growth and proximate composition of Nannochloropsis sp. and Tetraselmis sp. isolated from South China Sea cultured under control and natural condition. Int Biodeter BiodegrI. 2014;95:11–8.View ArticleGoogle Scholar
- Zhang DM, Chen YX, Jilani G, Wu WX, Liu WL, Han ZY. Optimization of struvite crystallization protocol for pretreating the swine wastewater and its impact on subsequent anaerobic biodegradation of pollutants. Bioresour Technol. 2012;116(7):386–95.View ArticleGoogle Scholar
- Aiba S, Ogawa T. Assessment of growth yield of a blue–green alga: spirulina platensis, in axenic and continuous culture. J General Microbiol. 1977;102:179–82.View ArticleGoogle Scholar
- Stanier RY, Kunisawa R, Mandel M, Cohen-Bazire G. Purification and properties of unicellular blue–green algae (Order Chroococcales). Bacteriol Rev. 1971;35:171–205.Google Scholar
- Song M, Pei H, Hu W, Ma G. Evaluation of the potential of 10 microalgal strains for biodiesel production. Bioresour Technol. 2013;141(4):245–51.View ArticleGoogle Scholar
- Santana H, Cereijo CR, Teles VC, Nascimento RC, Fernandes MS, Brunale P, et al. Microalgae cultivation in sugarcane vinasse: selection, growth and biochemical characterization. Bioresour Technol. 2017;228:133–40.View ArticleGoogle Scholar
- Association APHA. Standard methods for the examination of water and wastewater. 21st ed. Washington: American Public Health Association; 2005.Google Scholar
- Bligh EGDWJ. A rapid method of total lipid extraction and purification. Can J Chem. 1959;37:911–7.Google Scholar
- Indarti E, Majid MIA, Hashim R, Chong A. Direct FAME synthesis for rapid total lipid analysis from fish oil and cod liver oil. J Food Compos Anal. 2005;18:161–70.View ArticleGoogle Scholar
- Kirst GO. Salinity tolerance of eukaryotic marine algae. Annu Rev Plant Physiol Plant Mol Biol. 1989;40:21–53.Google Scholar
- Hwang JH, Church J, Lee SJ, Park J, Lee WH. Use of microalgae for advanced wastewater treatment and sustainable bioenergy generation. Environ Eng Sci. 2016;33:882–97.View ArticleGoogle Scholar
- Wilsenach JA, Schuurbiers CAH, van Loosdrecht MCM. Phosphate and potassium recovery from source separated urine through struvite precipitation. Water Res. 2007;41:458–66.View ArticleGoogle Scholar
- Yetilmezsoy K, Sapci-Zengin Z. Recovery of ammonium nitrogen from the effluent of uasb treating poultry manure wastewater by map precipitation as a slow release fertilizer. J Hazard Mater. 2009;166(1):260–9.View ArticleGoogle Scholar
- Martí N, Pastor L, Bouzas A, et al. Phosphorus recovery by struvite crystallization in WWTPs: influence of the sludge treatment line operation. Water Res. 2010;44(7):2371–9.View ArticleGoogle Scholar
- Park J, Jin HF, Lim BR, Park KY, Lee K. Ammonia removal from anaerobic digestion effluent of livestock waste using green alga Scenedesmus sp. Bioresour Technol. 2010;101:8649–57.View ArticleGoogle Scholar
- Richmond A, Hu Q. Handbook of microalgal culture: biotechnology and applied phycology. West Sussex: Wiley; 2013.View ArticleGoogle Scholar
- Guiry MD, Guiry GM. AlgaeBase. World-wide electronic publication. Galway: National University of Ireland; 2016. http://www.algaebase.org. Accessed 27 Dec 2016.
- Lopez-Rodas V, Marva F, Rouco M, Costas E, Flores-Moya A. Adaptation of the chlorophycean Dictyosphaerium chlorelloides to stressful acidic, mine metal-rich waters as result of pre-selective mutations. Chemosphere. 2008;72:703–7.View ArticleGoogle Scholar
- Pereira M, Bartolome MC, Sanchez-Fortun S. Influence of pH on the survival of Dictyosphaerium chlorelloides populations living in aquatic environments highly contaminated with chromium. Ecotoxicol Environ Saf. 2013;98:82–7.View ArticleGoogle Scholar
- Park JB, Craggs RJ, Shilton AN. Enhancing biomass energy yield from pilot-scale high rate algal ponds with recycling. Water Res. 2013;47:4422–32.View ArticleGoogle Scholar
- Bock C, Pröschold T, Krienitz L. Updating the genus Dictyosphaerium and description of Mucidosphaerium gen. nov (Trebouxiophyceae) based on morphological and molecular data. J Phycol. 2011;47:638–52.View ArticleGoogle Scholar
- Zou S, Fei C, Song J, Bao Y, He M, Wang C. Combining and comparing coalescent, distance and character-based approaches for barcoding microalgaes: a test with chlorella-like species (Chlorophyta). PLoS ONE. 2016;11:e0153833.View ArticleGoogle Scholar
- Cho DH, Ramanan R, Heo J, Shin DS, Oh HM, Kim HS. Influence of limiting factors on biomass and lipid productivities of axenic Chlorella vulgaris in photobioreactor under chemostat cultivation. Bioresour Technol. 2016;211:367–73.View ArticleGoogle Scholar
- Zhou X, Ge H, Xia L, Zhang D, Hu C. Evaluation of oil-producing algae as potential biodiesel feedstock. Bioresour Technol. 2013;134:24–9.View ArticleGoogle Scholar
- Vidyashankar S, VenuGopal KS, Swarnalatha GV, Kavitha MD, Chauhan VS, Ravi R, et al. Characterization of fatty acids and hydrocarbons of chlorophycean microalgae towards their use as biofuel source. Biomass Bioenerg. 2015;77:75–91.View ArticleGoogle Scholar
- Markou G, Nerantzis E. Microalgae for high-value compounds and biofuels production: a review with focus on cultivation under stress conditions. Biotechnol Adv. 2013;31:1532–42.View ArticleGoogle Scholar
- Stansell GR, Gray VM, Sym SD. Microalgal fatty acid composition: implications for biodiesel quality. J Appl Phycol. 2011;24:791–801.View ArticleGoogle Scholar
- Olmstead ILD, Hill DRA, Dias DA, et al. A quantitative analysis of microalgal lipids for optimization of biodiesel and omega-3 production. Biotechnol Bioeng. 2013;110:2096–104.View ArticleGoogle Scholar