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Simultaneous wastewater treatment and lipid production by Scenedesmus sp. HXY2
Bioresource Technology 302 (2020) 122903
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Simultaneous wastewater treatment and lipid production by Scenedesmus sp. HXY2 Sisi Yea, Li Gaob, Jing Zhaoa, Mei Ana, Haiming Wua, Ming Lia,c,
T
⁎
a
College of Resources and Environment, Northwest A & F University, Yangling 712100, PR China SouthEast Water, 101 Wells Street, Frankston, VIC 3199, Australia c Scientific Laboratory of Heyang Agricultural Environment and Farmland Cultivation, Ministry of Agriculture and Rural Affairs, Heyang 715300, PR China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Scenedesmus Lipid production Wastewater treatment 18S rRNA gene
Screening for highly efficient microalgae is an important technique for improving treatment efficiency. In this study, eight species of microalgae (five Scenedesmus and three Desmodesmus) were isolated from water and soil in the Hexi Corridor region, China, and identified by 18S rRNA gene sequence analysis. Scenedesmus sp. HXY2 grew well under high total organic carbon and ammonia conditions and had the highest nutrient removal efficiency (> 95%). On day 12, the biomass of Scenedesmus sp. HXY2 was 7.2 × 106 cells mL−1. The lipid content and productivity of this species were 15.56% and 5.67 mg L-1 day−1, respectively. The proportion of unsaturated fatty acids (60.07%) indicated that the lipids of Scenedesmus sp. HXY2 were suitable for biodiesel production. Scenedesmus sp. HXY2 showed great potential for growth in wastewater with high ammonia and organic contents to simultaneously purify wastewater and produce lipids.
1. Introduction Biodiesel production can play an important role in countering the global energy crisis and climate change (Chisti, 2007; Jebali et al., 2015). Microalgae biomass is one of the most promising raw materials for biodiesel production because of its rapid growth rate and high lipid content (Abou-shanab et al., 2011a, b; Brennan and Owende, 2010; Fernando et al., 2006). However, the high production cost of microalgal biodiesel restricts its industrial development (Abou-shanab et al., 2011b). Nevertheless, using wastewater to culture microalgae is an important strategy for reducing the production costs of microalgae lipids for biodiesel (Clarens et al., 2010) because the process of nutrient removal can be accompanied by microalgal biomass production (Li et al., 2019). The isolation and identification of microalgae species that can produce high amounts of lipids and treat wastewater efficiently is important for this technique. Significant effort has been expended to screen and isolate microalgae species with efficient lipid-production characteristics. Several studies have reported that the lipid contents of microalgae isolated from natural conditions ranged from 47.9% to 72% when cultured in standard culture media (Yang et al., 2010; Abou-shanab et al., 2011b; Zhang et al., 2014). Thangavel et al. (2018) isolated Chlamydomonadales sp. from the Nilgiri Biosphere Reserve, India, and found that its lipid content was 29% when cultured in BG-11 medium. Even though ⁎
this lipid content was very high, their growth in wastewater was not assessed. Some microalgae species can survive and grow in wastewater, simultaneously treating the wastewater and producing biomass for biodiesel production (Jebali et al., 2015; Álvarez-Díaz et al., 2017). Inoculating microalgae in wastewater to produce microalgal biomass and purify wastewater can greatly reduce the cost of microalgae lipid production. Yang et al. (2011) found that the use of wastewater could reduce the nutrient demand by approximately 55%. Jebali et al. (2015) isolated Scenedesmus sp. from northeast Tunisia and cultured this alga in wastewater. Its lipid content was 14.8%, and the removal efficiencies of chemical oxygen demand (COD), ammonium-nitrogen (NH4+-N) and total phosphorus (TP) were 92–94%, 61–99% and 93–99%, respectively. Franchino et al. (2013) found that the removal rate of NH4+-N by Scenedesmus obliquus reached 83% when cultured in diluted livestock wastewater for 21 days. Álvarez-Díaz et al. (2017) reported the highest nitrogen removal rate for Chlorella sorokiniana reached 6.6 mg N L-1 d-1 when cultured in wastewater for 25 days, and the lipid content was 36.75%. However, further improvements in lipid production efficiency and decontamination abilities of microalgae are still required and will rely on isolating and screening of novel microalgae species. Besides biodiesel production (Tiwari et al., 2018; Sivaramakrishnan and Incharoensakdi, 2018), byproducts of lipid producing microalgae, including pigments, antioxidants, fatty acids and enzymes, are also of high commercial value (Hu et al., 2018). These high-value byproducts
Corresponding author. E-mail address: [email protected] (M. Li).
https://doi.org/10.1016/j.biortech.2020.122903 Received 27 November 2019; Received in revised form 23 January 2020; Accepted 24 January 2020 Available online 27 January 2020 0960-8524/ © 2020 Elsevier Ltd. All rights reserved.
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light:dark photoperiod. The isolates were cultured continuously for 12 days, and the water quality was measured every 4 days. The biomass, lipid productivity, fatty acid composition and pigment and protein contents were analyzed on day 12. All experiments were carried out in triplicate.
can effectively reduce the cost of microalgae biodiesel production, which is an important basis for screening microalgae species. Harsh environments, such as hot, cold, dry, high salinity, high osmotic pressure and strong ultraviolet radiation, are considered suitable for the isolation of microalgae species (Zhang et al., 2014). The purpose of this study was to identify and isolate microalgae species from the Hexi Corridor that could treat wastewater and produce high lipid contents using wastewater as the culture medium. The Hexi Corridor was selected because this area has a dry climate, low annual precipitation, large temperature difference between day and night, strong ultraviolet radiation and harsh natural conditions.
2.4. Biomass and water quality determination Cell density was measured by microscopic counting using a hemocytometer under a high magnification (400 × ) optical microscope (CX31, Tokyo, Japan). The dry weight of the microalgae was measured by the differential method. A 0.45 µm filter (Whatman #1) was dried to constant weight in the oven (m1). Ten mL of microalgae solution (recorded as V) was filtered through the filter with a vacuum pump (SHBIII, Zhengzhou, China). The filter membrane with microalgae was dried in an oven to a constant weight (m2). The dry weight of the microalgae was calculated according to Eq. (1).
2. Materials and methods 2.1. Microalgae isolation and purification Eight different strains were isolated and purified from the Hexi Corridor. Three strains were isolated from freshwater (Hexi College lake and fountain and Gaotai agricultural area pool) and named HXY2, HXY3 and HXY8; five strains were isolated from soils (Zhangye National Wetland Park, Zhangye Runquan Lake Park, Hexi College garden and Hexi College family building garden) and named HXY1, HXY4, HXY5, HXY6 and HXY7. In the current study, microalgae were isolated using BG-11 medium containing 1.5% agar. Spread plates were inoculated with 0.1–0.2 mL of water sample or soil suspension solution and incubated for 5–7 days at 25 °C, 60 µmol photons m−2 s−1 and a 14:10 h light:dark photoperiod. Single colonies were picked from the spread plates and inoculated into wells of a 12-well microtiter plate with liquid medium, where green colony was clearly observed under the microscope. Culture purity was confirmed by repeated plating and by regular observation under a microscope.
Microalgae dry weight(mg L−1) = (m2 − m1)/V
(1)
The filtrate (filtered through 0.45 μm filter (Whatman # 1) with a vacuum pump) was analyzed for water quality. The total organic carbon (TOC) concentration was determined using a total organic carbon analyzer (TOC-L CPN, Shimadzu Instruments Co., Ltd., Suzhou, China). Ammonia nitrogen (NH4+-N) was determined by Nessler's reagent spectrophotometry (APHA, 2005). Total dissolved nitrogen (TDN) was determined by potassium persulfate oxidation-ultraviolet spectrophotometry (Ebina et al., 1983). The total dissolved phosphorus (TDP) was determined by molybdenum antimony anti-spectrophotometry (Ebina et al., 1983). The removal efficiency of nutrients was calculated by Eq. (2) where ηi (i was TOC, NH4+-N, TDN or TDP) was the removal efficiency, Ci0 was the initial water concentration and Cia was the measured water concentration.
2.2. Identification of microalgae
ηi (%) = (Ci0 − Cia)/Ci0 × 100
Microalgae strains were preliminarily identified by optical microscope analysis and then were further confirmed by 18S rRNA gene sequence analysis. An aliquot of cultured cells (30–60 mL) was harvested during the exponential growth phase and centrifuged (10,000 rpm for 10 min) to concentrate. The microalgae pellet was then freeze-dried (LGJ-10C, China), and the DNA was extracted using a Plant Genomic DNA extraction kit. Eukaryotic primers P73F (AATCAGTTATAGTTTATTTGRTGGTACC) and P47R (TCTCAGGCTCCCTCTCCGGA) were used for PCR amplification. PCR products were analyzed by electrophoresis on a 0.8% (w/v) agarose gel to confirm the presence and approximate concentration of the products, which were then sequenced (Hongzhong Baide, Jiangsu, China). The 18S rRNA gene sequences of the isolates were searched against GenBank using BLAST and were manually aligned with representative sequences from microalgal strains and related taxa, according to similarities determined by the Clustal W program (Larkin et al., 2007). The phylogenetic tree was constructed using MEGA4 (Tamura and Dudley, 2007) software.
(2)
2.5. Lipid extraction and fatty acid analysis Microalgae solution (30 mL) was centrifuged in a high-speed centrifuge (10,000 rpm) for 6 min. The microalgal pellet was then placed in an oven (60 °C) and dried to constant weight. The lipids were extracted using the methanol-chloroform method (Dyer and Bligh, 1959). Approximately 100 mg of dry microalgae was weighed (Wa) into a 50 mL centrifuge tube, and 12 mL of 1:2 chloroform–methanol solution was added. The centrifuge tube was sonicated for 1 h at 25 °C and then centrifuged for 6 min (10,000 rpm); the sonicated suspension was filtered through a 0.45 µm filter membrane (Whatman #1) and transferred to a new centrifuge tube. The extraction was repeated for the remaining microalgal residue and both filtrates were combined. The combined filtrate was thoroughly mixed with 16 mL of 5% NaCl. The lower layer solution was separated into a round bottom flask and concentrated by a rotary evaporator (RE-2000A, Yarong Biochemical Instrument Company, Shanghai, China). The remaining solution was transferred to a 4 mL vial (vial dry weight recorded as Wi). The vial was dried using a nitrogen purging instrument (ND200-1, Hang Zhou Rui Cheng Instrument Co., Ltd., Hangzhou, China) at 45 °C until the sample reached a constant mass (recorded as Wf). The lipid content, production and productivity were calculated as in Eqs. (3)–(5), respectively.
2.3. Experimental culture conditions Simulated wastewater was used as the growth medium in the current study and was composed of (in mg L-1): CH3COONa 3000, NH4Cl 200, KH2PO4 60, FeSO4·7H2O 0.55, MgSO4·7H2O 66, CaCl2 6, CuSO4·5H2O 0.08, Co(NO3)2·6H2O 0.05, H3BO3 2.86, MnCl2·4H2O 1.86, ZnSO4·7H2O and Na2MoO4·2H2O 0.39. The experimental device was a glass tube with an inner diameter of 8 cm and a volume of 1.5 L. Aeration was applied at 0.16 L min−1; daytime aeration was intermittent (aeration:no aeration = 2:1h), whereas at night there was no aeration. The initial inoculation density was 1 × 105 cells mL−1. The culture temperature and illumination were 25 °C and 60 µmol photons m−2 s−1, respectively, and the cultures were incubated with a 14:10 h
Lipid content (%) = (Wf − W)/Wa × 100 i
(3)
Lipid production(mg L−1) = lipid content × microalgaedry weight
(4)
Lipid productivity(mg L−1day −1) = lipid production/day
(5)
Fatty acid methyl esters (FAMEs) were analyzed using the method of Lepage and Roy (1985). The extracted lipids were completely dissolved in 4 mL of 14% BF3-methanol solution and then boiled in a water bath 2
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wastewater containing complex pollutants and continuously purify the wastewater. Álvarez-Díaz et al. (2017) found that Scenedesmus obliquus could survive in municipal wastewater with TOC, TP and total nitrogen (TN) contents of 10.67, 1.55 and 20.09 mg L-1, respectively, with biomass productivity of 81.0 mg L-1 day−1. Komolafe et al. (2014), observed biomass productivity of Desmodesmus sp. of 13.0 mg L-1 day−1 using sewage treatment plant influent as the culture medium (with TP and TN of 35.4 and 65.6 mg L-1, respectively). In the current study, Desmodesmus sp. HXY7 had the highest biomass productivity of 57.9 mg L-1 day−1, which was much higher than that of Komolafe et al. (2014) and was close to the value shown by Álvarez-Díaz et al. (2017).
for 15 min. After cooling to room temperature, 2 mL of n-heptane and 4 mL of saturated NaCl solution were added, and the mixture was shaken well. After stratification, the supernatant was filtered using a 0.22-µm organic phase needle filter (TPFM012, Nanjing Tai Pu Rui Instrument Equipment Co., Ltd., China). FAMEs were analyzed using a gas chromatograph (GC-2014C, Shimadzu, Japan) equipped with flame ionization detector (FID) and a DB-5 ms (60 m) capillary column. The carrier gas was N2, and the injection and detector temperatures were 250 °C and 290 °C, respectively. The initial column temperature was 120 °C held for 3 min; it was then raised to 220 °C at 4 °C min−1 and held for 5 min; finally, the temperature was increased to 280 °C at 3 °C min−1 and held for 20 min. The injection volume was 1 μL.
3.3. Nutrient removal 2.6. Pigment analysis TOC, TP and TN concentrations in the simulated wastewater in this study were 878, 13.67 and 52.34 mg L−1, respectively. After 8 days of cultivation, the removal efficiency of NH4+-N by all eight microalgae species was > 98% (Fig. 3A). The removal efficiencies of TDN and TOC were > 90% and > 85%, respectively (Fig. 3B, C). > 70% of TDP was removed (Fig. 3D). Among the eight microalgae species, Scenedesmus sp. HXY2 showed the best nutrient removal efficiency, with removal efficiencies for TOC, NH4+-N, TDN and TDP of 96.07%, 99.09%, 96.62% and 94.52%, respectively. By day 12, the removal efficiencies for TOC, NH4+-N and TDN by the eight microalgae species were close to 100%. The removal efficiency for TDP by most species was > 95%, except for Scenedesmus sp. HXY4 and Desmodesmus sp. HXY8, which had removal efficiencies of ~72%. Kothari et al. (2013) cultured Chlamydomonas polypyrenoideum in dairy wastewater for 10 days and showed removal efficiencies for NO3–N, NH4+-N, TP and COD of 90%, 90%, 70% and 60%, respectively. Zhang et al. (2011) reported removal efficiencies for TOC, TN, and TP of 99.6%, 87.6%, and 69.8%, respectively, when Chlorella vulgaris was cultured in photomicrobial fuel cell wastewater for six days. Iasimone et al. (2018) showed that the removal rate of NH4+-N by mixed microalgae (60% Cyanobacteria and 40% Chlorophyta) in urban wastewater was > 74.8%. In this study, the NH4+-N removal efficiencies by the eight microalgae were higher than the values mentioned above. Additionally, all eight microalgae species could survive at high TOC concentrations (878 mg L-1). Furthermore, the isolated microalgae showed excellent abilities to degrade TOC, with removal rates of approximately 100 mg TOC L-1 day−1.
The microalgae solution (10 mL) was filtered through a 47-mm GF/ F filter (Whatman, Nanjing, China). The filter was placed in a 25-mL stoppered tube, and 5 mL of acetone solution was injected to permeate the filter; then it was treated with ultrasound for 10 min at 4 °C. The solution was incubated in the dark at −20 °C for 24 h, and then the filtrate was extracted with a polytetrafluoroethylene syringe filter and loaded into a 1.5 mL brown chromatography bottle for high performance liquid chromatography (HPLC) (Agilent-1100 series, USA) detection. Two mobile phases were used: A (2:1 = methanol: 0.5 mol L-1 ammonium acetate) and B (3:7 = methanol: acetonitrile). The linear gradients used were: (min, mobile phase A%, mobile phase B%) (0, 100, 0), (10, 40, 60), (20, 20, 80), (30, 0, 100), (40, 0, 100), (50, 0, 100), (52, 25, 75), (60, 100, 0), and (65, 100, 0). The injection volume was 100 μL, and the wavelengths for detection were 440, 266 and 300 nm. 2.7. Statistics All data were expressed as mean ± standard deviation. Differences in data between treatments were determined by analysis of variance (ANOVA) using SPSS 19.0 software (p < 0.05) (IBM Corp., Armonk, NY, USA). 3. Results and discussion 3.1. Identification of microalgae species Eight microalgae species were isolated in the current study. HXY1 existed as 2- or 4-cell populations. The cells were spindle-shaped with lengths of 7.81–10.53 µm. The other seven microalgae species were mostly unicellular. HXY2 and HXY3 cells were similar in morphology, with a spindle shapes and lengths of 7.00–12.09 µm. HXY5, HXY6 and HXY7 were larger, with lengths of 7.22–12.50 µm. HXY4 and HXY8 were much smaller, with spherical cell morphology having a diameter of 5.57–7.51 µm. The eight microalgae species were all in the phylum Chlorophyta and were identified as Scenedesmus sp. by morphological observation. The phylogenetic tree of the eight microalgae species obtained by 18S rRNA gene sequencing analysis is shown in Fig. 1. HXY1, HXY2, HXY4, HXY5 and HXY6 were classified as Scenedesmus (Scenedesmus sp. HXY1, HXY2, HXY4, HXY5 and HXY6, respectively). HXY3, HXY7 and HXY8 were classified as Desmodesmus (Desmodesmus sp. HXY3, HXY7 and HXY8, respectively).
3.4. Lipid productivity and fatty acid composition Desmodesmus sp. HXY3 had the highest lipid productivity of 7.50 mg L-1 day−1 (Fig. 4C) because of its high biomass productivity (Fig. 2A). Generally, higher lipid productivity can be obtained in standard media because of the balanced nutrient and mineral compositions. For example, Zhang et al. (2014) isolated Chlorella vulgaris CJ15 in the coastal areas of Qingdao using F/2 medium and demonstrated lipid productivity of 11.7 mg L-1 day−1. Chen et al. (2012) found that Scenedesmus sp. NJ-1, isolated from Antarctica, had a lipid productivity of 36.8 mg L-1 day−1 in BG-11 medium. Therefore, Desmodesmus sp. HXY3, isolated in this study, is expected to achieve higher lipid productivity in standard media. The medium nutrient and mineral compositions had significant effects on the lipid content of microalgae. The lipid content of the eight microalgae species ranged from 10% to 16% (Fig. 4A). Scenedesmus sp. HXY2 had the highest lipid content (15.56%). Yang et al. (2011) reported that Chlorella ellipsoidea YJ1, isolated from the secondary effluent of the Beijing Wastewater Treatment Plant, had lipid content and lipid productivity of 43% and 11.4 mg L-1 day−1, respectively, when it was cultured in secondary effluent (COD, TN and TP of 24, 7 and 0.46 mg L-1, respectively). Xin et al. (2010) found that the lipid content and lipid productivity of Scenedesmus sp. LX1 were 31–33% and 8 mg L1 day−1, respectively, when cultured in secondary effluent (COD, TN
3.2. Biomass production The biomass of the eight strains after 12 days of culture in simulated wastewater is shown in Fig. 2A. Desmodesmus sp. HXY7 had the highest cell density and dry weight of 1.72 × 107 cells mL−1 and 695 mg L-1, respectively. Microalgae, especially Chlorophyta, have high tolerance to pollutants, including ammonia, organic matter and heavy metals (Cai et al., 2013; Komolafe et al., 2014). Thus, they can survive in 3
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Fig. 1. Phylogenetic tree showing the relationships among the 18S rRNA gene sequences of the eight microalgae isolates and the most similar sequences retrieved from the NCBI nucleotide database.
and TP content of 24, 15.5 and 0.5 mg L-1, respectively) from the Beijing Domestic Sewage Treatment Plant. In the current study, the TOC, TN and TP contents in simulated wastewater were much higher than those of Yang et al. (2011) and Li et al. (2010), but the lipid productivity was much lower than in those studies. This was because
the high N concentration prevented lipid accumulation (Yeh and Chang, 2011; Yeh and Chang, 2012; Pan et al., 2011). The results of this study indicated that Scenedesmus sp. HXY2 had sufficient lipid content for microalgal biodiesel production in high ammonia and organic content wastewater.
Fig. 2. (A) The cell density and the dry weight and (B) the polysaccharides and protein contents of the eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05). 4
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Fig. 3. (A) The removal efficiency for total organic carbon (TOC), (B) ammonium nitrogen (NH4+-N), (C) total dissolved nitrogen (TDN) and (D) total dissolved phosphorus (TDP) by eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05).
Fig. 4. (A) The lipid content, (B) lipid production and (C) lipid productivity of eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05). 5
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Table 1 Fatty acid compositions of eight microalgae species (%).
C14:0 C15:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C22:0 C22:2 C24:0 Unsaturated C16-C18
Scenedesmus sp. HXY1
Scenedesmus sp. HXY2
Desmodesmus sp. HXY3
Scenedesmus sp. HXY4
Scenedesmus sp. HXY5
Scenedesmus sp. HXY6
Desmodesmus sp. HXY7
Desmodesmus sp. HXY8
1.64 ± 0.16 ND 43.68 ± 11.83 3.08 ± 0.9 4.85 ± 1.49 14.48 ± 2.59 32.27 ± 14.96 ND ND ND ND 49.83 98.36
1.26 ± 0.31 0.88 ± 0.09 33.63 ± 1.9 2.66 ± 0.1 3.49 ± 0.42 29.19 ± 14.92 25.57 ± 13.84 2.65 ± 0.23 0.95 ± 0.23 ND ND 60.07 97.19
1.43 ± 0.19 ND 45.91 ± 6.94 2.17 ± 0.15 5.31 ± 1.12 18.21 ± 10.51 23.52 ± 2.14 2.05 ± 0.29 1.41 ± 0.27 ND ND 45.95 97.17
1.94 ± 0.17 ND 41.23 ± 1.78 2.16 ± 0.2 5.91 ± 0.19 26.98 ± 1.54 17.08 ± 0.62 2.99 ± 1 ND ND 1.69 ± 0.23 49.21 96.35
ND ND 34.5 ± 0.01 1.9 ± 0.48 7.49 ± 0.44 17.46 ± 0.23 32.69 ± 0.17 5.96 ± 0.01 ND ND ND 58.01 100
ND ND 39.07 ± 2.87 1.89 ± 0.23 9.66 ± 0.48 22.6 ± 1.63 26.78 ± 0.99 ND ND ND ND 51.27 100
1.37 ± 0.69 ND 49.97 ± 4.64 1.18 ± 0.5 7.85 ± 1.19 7.96 ± 2.2 28.1 ± 6.44 1.67 ± 0.43 ND 1.9 ± 0.02 ND 40.81 96.73
1.89 ± 0.67 ND 45.78 ± 2.87 2.06 ± 0.26 8.22 ± 1.51 15.49 ± 1.59 15.72 ± 4.76 7.84 ± 5.32 ND 1.99 ± 0.06 1.51 ± 0.45 43.1 95.11
ND: not detected. Table 2 Pigment compositions and contents of eight microalgae species.
Scenedesmus sp. HXY1 Scenedesmus sp. HXY2 Desmodesmus sp. HXY3 Scenedesmus sp. HXY4 Scenedesmus sp. HXY5 Scenedesmus sp. HXY6 Desmodesmus sp. HXY7 Desmodesmus sp. HXY8
Viola (μg g−1)
Lut (μg g−1)
Chl-b (μg g−1)
Chl-a (μg g−1)
192.32 ± 14.51 ND 65.38 ± 12.96 59.38 ± 17.04 37.17 ± 32.33 ND 110.28 ± 12.15 ND
1241.21 ± 311.12 1476.15 ± 452.36 1097.87 ± 386.61 1635.37 ± 568.86 1040.15 ± 80.73 774.64 ± 454.17 3956.63 ± 226.15 1086.55 ± 216.56
92.94 ± 7.23 90.46 ± 22.21 159.53 ± 50.68 196.68 ± 75.66 172.35 ± 47.92 131.5 ± 64.11 499.57 ± 94.45 144.01 ± 66.44
318.31 ± 34.58 449.96 ± 120.54 511.67 ± 154.69 477.29 ± 162.24 570.11 ± 103.66 421.65 ± 184.39 1592.53 ± 362.23 404.35 ± 166.41
Viola: Violaxanthin. Lut: Lutein. Chl-b: Chlorophyll-b. Chl-a: Chlorophyll-a. ND: not detected.
cholesterol (Cristiano et al., 2018). For this study, the polysaccharide and protein contents are shown in Fig. 2B. Scenedesmus sp. HXY5 had the highest polysaccharide content (24.34 pg cell−1) and the highest protein content (46.39 pg cell−1). Lutein can play a positive role in preventing blindness and decreased vision caused by macular degeneration (Shi et al., 1997; Zhang et al., 1999; Wu et al., 2007). The pigment composition of the eight microalgae species is shown in Table 2. Desmodesmus sp. HXY7 had the highest lutein content (3.96 mg g−1) among all eight microalgae. The highest chlorophyll-a and chlorophyll-b contents were obtained for Desmodesmus sp. HXY7, 1.59 and 0.50 mg g−1, respectively. Minhas et al. (2016) summarized the lutein content of 22 microalgae, and the average value was 1.4 mg g−1. The lutein content of Desmodesmus sp. HXY7 in the current study (3.96 mg g−1) was much higher than the average value and higher than 91% of microalgae in the summary by Minhas et al. (2016). Ho et al. (2014) isolated six Scenedesmus species and reported that the average lutein content was 2.40 mg g−1, which was also lower than the value of Desmodesmus sp. HXY7. Therefore, the newly isolated Desmodesmus sp. HXY7 could be a potential source for lutein production.
The most common FAMEs present in biodiesel are palmitic acid (16:0), stearic acid (18:0), oleic acid (18:1), linoleic acid (18:2), and linolenic acid (18:3) (Knothe, 2013). The saturated fatty acids (16:0) and monounsaturated fatty acids (18:1) play an important role in determining biodiesel performance, such as oxidative stability and cold flow properties of biodiesel (Kaur et al., 2012). The fatty acid compositions of eight microalgae species cultured in simulated wastewater medium are shown in Table 1. The main fatty acids of the eight species of microalgae were palmitic acid (16:0), oleic acid (18:1) and linoleic acid (18:2), accounting for > 80% of the total fatty acid composition. Scenedesmus sp. HXY2 had the highest unsaturated fatty acid composition of 60.07%, followed by Scenedesmus sp. HXY5 (58.01%). The proportion of C16–C18 fatty acids for all eight microalgae species was > 95%. Thao et al. (2017) isolated lipid-producing microalgae in Vietnam, and, of these, Mychonastes sp. N16 had a maximum C16–C18 fatty acid content of 61.62%. Yeh and Chang, 2012 reported that the proportion of C16–C18 fatty acids of Chlorella vulgaris ESP-31 in standard medium was 66.99–89.59%. Abou-Shanab et al. (2011b) isolated five lipid-producing microalgae in South Korea, and their proportion of C16–C18 fatty acids ranged from 56% to 97%. In this study, the proportions of C16–C18 fatty acids for the eight microalgae species were similar to the values mentioned above, indicating that the isolated microalgae have good lipid production potential.
4. Conclusion Eight microalgae species were isolated from the Hexi Corridor. They were cultured in simulated wastewater for 12 days. Scenedesmus sp. HXY2 had the highest nutrient removal efficiency and lipid content. On day 8, the removal efficiencies of TOC, NH4+-N, TDN and TDP by Scenedesmus sp. HXY2 were 96.07%, 99.09%, 96.62% and 94.52%, respectively. The lipid content and lipid productivity of this alga was 15.56% and 5.67 mg L-1 day−1, respectively. Our results indicated that Scenedesmus sp. HXY2 has great potential for culturing in wastewater with high ammonia nitrogen and organic concentrations for simultaneous wastewater purification and lipid production.
3.5. Byproducts Polysaccharides produced by microalgae are usually very complex, and the complex structure of some polysaccharides can result in anticancer activity (Spolaore et al., 2006; Chaiklahan et al., 2013). Mišurcová et al. (2014) found that Chlorella sp. and Spirulina sp. had high protein contents (> 50% of dry weight) and could be used as sources of essential amino acids. Additionally, microalgae proteins have shown a significant effect in reducing blood pressure and lowering 6
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Manage. 223. Jebali, A., Acién, F.G., Gómez, C., Fernández-Sevilla, J.M., Mhiri, N., Karray, F., Dhouib, A., Molina-Grima, E., Sayadi, S., 2015. Selection of native Tunisian microalgae for simultaneous wastewater treatment and biofuel production. J. Bioresour Technol. 198. Kaur, S., Sarkar, M., Srivastava, R.B., Gogoi, H.K., Kalita, M.C., 2012. Fatty acid profiling and molecular characterization of some freshwater microalgae from India with potential for biodiesel production. New Biotech. 29 (3), 332–344. Knothe, G., 2013. “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energy Fuel 22 (2). Komolafe, O., Velasquez Orta, S.B., Monje-Ramirez, I., Noguez, I.Y., Harvey, A.P., Orta, L., María, T., 2014. Biodiesel production from indigenous microalgae grown in wastewater. Bioresour. Technol. 154, 297–304. Kothari, R., Prasad, R., Kumar, V., Singh, D.P., 2013. Production of biodiesel from microalgae Chlamydomonas polypyrenoideum grown on dairy industry wastewater. Bioresour. Technol. 144, 499–503. Larkin, M.A., Blackshields, G., Brown, N.P., Chenna, R., Mcgettigan, P.A., Mcwilliam, H., Valentin, F., Wallace, I.M., Wilm, A., Lopez, R., 2007. Clustal w and clustal x version 2.0. Bioinformatics 23 (21), 2947–2948. Lepage, G., Roy, C.C., 1985. Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J. Lipid Res. 25 (12), 1391–1396. Li, K., Liu, Q., Fang, F., Luo, R.H., Lu, Q., Zhou, W.G., Huo, S.H., Cheng, P.F., Liu, J.Z., Addy, M., Chen, P., Chen, D.J., Ruan, R., 2019. Microalgae-based wastewater treatment for nutrients recovery: a review. J. Bioresour. Technol. 291. Minhas, A.K., Hodgson, P., Barrow, C.J., Sashidhar, B., Adholeya, A., 2016. The isolation and identification of new microalgal strains producing oil and carotenoid simultaneously with biofuel potential. Bioresour. Technol S0960852416304242. Mišurcová, L., Buňka, F., Vávra, A.J., Machů, L., Samek, D., 2014. Amino acid composition of algal products and its contribution to RDI. Food Chem. 151, 120–125. Pan, Y.Y., Wang, S.T., Chuang, L.T., Chang, Y.W., Chen, C.N.N., 2011. Isolation of thermo-tolerant and high lipid content green microalgae: oil accumulation is predominantly controlled by photosystem efficiency during stress treatments in Desmodesmus. Bioresour. Technol. 102 (22), 10510–10517. Shi, X.M., Chen, F., Yuan, J.P., Chen, H., 1997. Heterotrophic production of lutein by selected chlorella strains. J. Appl. Phycol. 9 (5), 445–450. Sivaramakrishnan, R., Incharoensakdi, A., 2018. Microalgae as feedstock for biodiesel production under ultrasound treatment – a review. Bioresour. Technol. Spolaore, P., Joannis-Cassan, C., Duran, E., Isambert, A., 2006. Commercial applications of microalgae. J. Biosci. Bioeng. 101 (2), 87–96. Tamura, K., Dudley, J., 2007. MECA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596–1599. Thangavel, K., Krishnan, P.R., Nagaiah, S., Kuppusamy, S., Dananjeyan, B., 2018. Growth and metabolic characteristics of oleaginous microalgal isolates from Nilgiri biosphere reserve of India. BMC Microbiol. 18 (1), 1. Thao, T.Y., Linh, D.T.N., Si, V.C., Carter, T.W., Hill, R.T., 2017. Isolation and selection of microalgal strains from natural water sources in Viet Nam with potential for edible oil production. Mar. Drugs 15 (7), 194. Tiwari, A., Rajesh, V.M., Yadav, S., 2018. Biodiesel production in micro-reactors: a review. Energy. Sustain. Dev. 43, 143–161. Wu, Z.Y., Shi, C.L., Shi, X.M., 2007. Modeling of lutein production by heterotrophic Chlorellain batch and fed-batch cultures. World J. Microb. Biotechnol. 23 (9), 1233–1238. Xin, L., Hong-Ying, H., Jia, Y., 2010. Lipid accumulation and nutrient removal properties of a newly isolated freshwater microalga, Scenedesmus sp. LX1, growing in secondary effluent. New. Biotech. 27 (1), 59–63. Yang, H.L., Lu, C.K., Chen, S.F., Chen, Y.M., Chen, Y.M., 2010. Isolation and characterization of taiwanese heterotrophic microalgae: screening of strains for docosahexaenoic acid (dha) production. Mar. Biotechnol. 12 (2), 173–185. Yang, J., Xu, M., Hu, Q., Sommerfeld, M., Chen, Y., 2011. Life-cycle analysis on biodiesel production from microalgae: water footprint and nutrients balance. Bioresour. Technol. 102, 159–165. Yeh, K.L., Chang, J.S., 2011. Nitrogen starvation strategies and photobioreactor design for enhancing lipid content and lipid production of a newly isolated microalga chlorella vulgaris esp-31: implications for biofuels. Biotech. J. 6 (11), 1358–1366. Yeh, K.L., Chang, J.S., 2012. Effects of cultivation conditions and media composition on cell growth and lipid productivity of indigenous microalga chlorella vulgaris esp-31. Bioresour. Technol. 105, 120–127. Zhang, B., Wang, L., Hasan, R., Shahbazi, A., 2014. Characterization of a native algae species chlamydomonas debaryana: strain selection, bioremediation ability, and lipid characterization. BioResources. 9 (4). Zhang, X.W., Shi, X.M., Chen, F., 1999. A kinetic model for lutein production by the green microalga Chlorella protothecoidesin heterotrophic culture. J. Ind. Microbiol. Biotech. 23 (6), 503–507. Zhang, Y., Noori, J.S., Angelidaki, I., 2011. Simultaneous organic carbon, nutrients removal and energy production in a photomicrobial fuel cell (PFC). Energy Environ. Sci. 4 (10), 4340.
Sisi Ye: Data curation, Writing - original draft, Writing - review & editing. Li Gao: Writing - original draft, Writing - review & editing. Jing Zhao: Formal analysis, Investigation. Mei An: Methodology, Software. Haiming Wu: Writing - original draft, Validation. Ming Li: Conceptualization, Resources, Supervision, Writing - original draft, Writing - review & editing. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This work was supported by the Fundamental Research Funds for Central Universities (Northwest A&F University, Grant No. 2452018142); Dr. Ming Li is funded as Tang Scholar by Cyrus Tang Foundation and Northwest A&F University. References Abou-Shanab, R.A.I., Hwang, J.H., Cho, Y., Min, B., Jeon, B.H., 2011a. a. Characterization of microalgal species isolated from fresh water bodies as a potential source for biodiesel production. Appl. Energy 88 (10), 3300–3306. Abou-Shanab, R.A.I., Matter, I.A., Kim, S.N., Oh, Y.K., Choi, J., Jeon, B.H., 2011b. b. Characterization and identification of lipid-producing microalgae species isolated from a freshwater lake. Biomass. Bioenerg. 35 (7), 3079–3085. Álvarez-Díaz, P.D., Ruiz, J., Arbib, Z., Barragán, J., Garrido-Pérez, M.C., Perales, J. A., 2017. Freshwater microalgae selection for simultaneous wastewater nutrient removal and lipid production. Algal Res. APHA, 2005. Standard methods for the examination of water and wastewater, 21st ed.; APHA and AWWA and WEF DC, Washington, DC. Brennan, L., Owende, P., 2010. Biofuels from microalgae-a review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sust. Energy Rev. 14 (2), 557–577. Cai, T., Park, S.Y., Li, Y., 2013. Nutrient recovery from wastewater streams by microalgae: status and prospects. Renew. Sust. Energy Rev. 19, 360–369. Chaiklahan, R., Chirasuwan, N., Triratana, P., Loha, V., Tia, S., 2013. Polysaccharide extraction from Spirulina sp. And its antioxidant capacity. Int. J. Biol. Macromol. 58, 73–78. Chen, Z., Gong, Y., Fang, X.T., 2012. Scenedesmus. NJ-1 isolated from Antarctica: a suitable renewable lipid source for biodiesel production. World J. Microb Biotech. 28 (11), 3219–3225. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25 (3), 294–306. Clarens, A.F., Resurreccion, E.P., White, M.A., Colosi, L.M., 2010. Environmental life cycle comparison of algae to other bioenergy feedstocks. Environ. Sci. Technol. 44 (5), 1813–1819. Cristiano, J.A., Lidiane, M.A., Meriellen, D., Claudio, A.N., Maria, A.M., 2018. Chlorella and spirulina microalgae as sources of functional foods, nutraceuticals, and food supplements; an overview. MOJ Food Process Technol. 6 (2), 00144. Dyer, E.G., Bligh, W.J., 1959. A rapid method of total lipid extraction and purification. J. Biochem Physiol. 37. Ebina, J., Tsutsui, T., Shirai, T., 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res. 17 (12). Fernando, S., Hall, C., Jha, S., 2006. NOx reduction from biodiesel fuels. Energy Fuel. 20 (1), 376–382. Franchino, M., Comino, E., Bona, F., Riggio, V.A., 2013. Growth of three microalgae strains and nutrient removal from an agro-zootechnical digestate. Chemosphere 92 (6), 738–744. Ho, S.H., Chan, M.C., Liu, C.C., Chen, C.Y., Lee, W.L., Lee, D.J., Chang, J.S., 2014. Enhancing lutein productivity of an indigenous microalga Scenedesmus obliquus FSP3 using light-related strategies. Bioresour. Technol. 152, 275–282. Hu, J.J., Nagarajan, D., Zhang, Q.G., Chang, J.S., Lee, D.J., 2018. Heterotrophic cultivation of microalgae for pigment production: a review. Biotechnol. Adv. 36, 54–67. Iasimone, F., Panico, A., De Felice, V., Fantasma, F., Iorizzi, M., Pirozzi, F., 2018. Effect of light intensity and nutrients supply on microalgae cultivated in urban wastewater: Biomass production, lipids accumulation and settleability characteristics. J. Environ.
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Simultaneous wastewater treatment and lipid production by Scenedesmus sp. HXY2 Sisi Yea, Li Gaob, Jing Zhaoa, Mei Ana, Haiming Wua, Ming Lia,c,
T
⁎
a
College of Resources and Environment, Northwest A & F University, Yangling 712100, PR China SouthEast Water, 101 Wells Street, Frankston, VIC 3199, Australia c Scientific Laboratory of Heyang Agricultural Environment and Farmland Cultivation, Ministry of Agriculture and Rural Affairs, Heyang 715300, PR China b
A R T I C LE I N FO
A B S T R A C T
Keywords: Scenedesmus Lipid production Wastewater treatment 18S rRNA gene
Screening for highly efficient microalgae is an important technique for improving treatment efficiency. In this study, eight species of microalgae (five Scenedesmus and three Desmodesmus) were isolated from water and soil in the Hexi Corridor region, China, and identified by 18S rRNA gene sequence analysis. Scenedesmus sp. HXY2 grew well under high total organic carbon and ammonia conditions and had the highest nutrient removal efficiency (> 95%). On day 12, the biomass of Scenedesmus sp. HXY2 was 7.2 × 106 cells mL−1. The lipid content and productivity of this species were 15.56% and 5.67 mg L-1 day−1, respectively. The proportion of unsaturated fatty acids (60.07%) indicated that the lipids of Scenedesmus sp. HXY2 were suitable for biodiesel production. Scenedesmus sp. HXY2 showed great potential for growth in wastewater with high ammonia and organic contents to simultaneously purify wastewater and produce lipids.
1. Introduction Biodiesel production can play an important role in countering the global energy crisis and climate change (Chisti, 2007; Jebali et al., 2015). Microalgae biomass is one of the most promising raw materials for biodiesel production because of its rapid growth rate and high lipid content (Abou-shanab et al., 2011a, b; Brennan and Owende, 2010; Fernando et al., 2006). However, the high production cost of microalgal biodiesel restricts its industrial development (Abou-shanab et al., 2011b). Nevertheless, using wastewater to culture microalgae is an important strategy for reducing the production costs of microalgae lipids for biodiesel (Clarens et al., 2010) because the process of nutrient removal can be accompanied by microalgal biomass production (Li et al., 2019). The isolation and identification of microalgae species that can produce high amounts of lipids and treat wastewater efficiently is important for this technique. Significant effort has been expended to screen and isolate microalgae species with efficient lipid-production characteristics. Several studies have reported that the lipid contents of microalgae isolated from natural conditions ranged from 47.9% to 72% when cultured in standard culture media (Yang et al., 2010; Abou-shanab et al., 2011b; Zhang et al., 2014). Thangavel et al. (2018) isolated Chlamydomonadales sp. from the Nilgiri Biosphere Reserve, India, and found that its lipid content was 29% when cultured in BG-11 medium. Even though ⁎
this lipid content was very high, their growth in wastewater was not assessed. Some microalgae species can survive and grow in wastewater, simultaneously treating the wastewater and producing biomass for biodiesel production (Jebali et al., 2015; Álvarez-Díaz et al., 2017). Inoculating microalgae in wastewater to produce microalgal biomass and purify wastewater can greatly reduce the cost of microalgae lipid production. Yang et al. (2011) found that the use of wastewater could reduce the nutrient demand by approximately 55%. Jebali et al. (2015) isolated Scenedesmus sp. from northeast Tunisia and cultured this alga in wastewater. Its lipid content was 14.8%, and the removal efficiencies of chemical oxygen demand (COD), ammonium-nitrogen (NH4+-N) and total phosphorus (TP) were 92–94%, 61–99% and 93–99%, respectively. Franchino et al. (2013) found that the removal rate of NH4+-N by Scenedesmus obliquus reached 83% when cultured in diluted livestock wastewater for 21 days. Álvarez-Díaz et al. (2017) reported the highest nitrogen removal rate for Chlorella sorokiniana reached 6.6 mg N L-1 d-1 when cultured in wastewater for 25 days, and the lipid content was 36.75%. However, further improvements in lipid production efficiency and decontamination abilities of microalgae are still required and will rely on isolating and screening of novel microalgae species. Besides biodiesel production (Tiwari et al., 2018; Sivaramakrishnan and Incharoensakdi, 2018), byproducts of lipid producing microalgae, including pigments, antioxidants, fatty acids and enzymes, are also of high commercial value (Hu et al., 2018). These high-value byproducts
Corresponding author. E-mail address: [email protected] (M. Li).
https://doi.org/10.1016/j.biortech.2020.122903 Received 27 November 2019; Received in revised form 23 January 2020; Accepted 24 January 2020 Available online 27 January 2020 0960-8524/ © 2020 Elsevier Ltd. All rights reserved.
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light:dark photoperiod. The isolates were cultured continuously for 12 days, and the water quality was measured every 4 days. The biomass, lipid productivity, fatty acid composition and pigment and protein contents were analyzed on day 12. All experiments were carried out in triplicate.
can effectively reduce the cost of microalgae biodiesel production, which is an important basis for screening microalgae species. Harsh environments, such as hot, cold, dry, high salinity, high osmotic pressure and strong ultraviolet radiation, are considered suitable for the isolation of microalgae species (Zhang et al., 2014). The purpose of this study was to identify and isolate microalgae species from the Hexi Corridor that could treat wastewater and produce high lipid contents using wastewater as the culture medium. The Hexi Corridor was selected because this area has a dry climate, low annual precipitation, large temperature difference between day and night, strong ultraviolet radiation and harsh natural conditions.
2.4. Biomass and water quality determination Cell density was measured by microscopic counting using a hemocytometer under a high magnification (400 × ) optical microscope (CX31, Tokyo, Japan). The dry weight of the microalgae was measured by the differential method. A 0.45 µm filter (Whatman #1) was dried to constant weight in the oven (m1). Ten mL of microalgae solution (recorded as V) was filtered through the filter with a vacuum pump (SHBIII, Zhengzhou, China). The filter membrane with microalgae was dried in an oven to a constant weight (m2). The dry weight of the microalgae was calculated according to Eq. (1).
2. Materials and methods 2.1. Microalgae isolation and purification Eight different strains were isolated and purified from the Hexi Corridor. Three strains were isolated from freshwater (Hexi College lake and fountain and Gaotai agricultural area pool) and named HXY2, HXY3 and HXY8; five strains were isolated from soils (Zhangye National Wetland Park, Zhangye Runquan Lake Park, Hexi College garden and Hexi College family building garden) and named HXY1, HXY4, HXY5, HXY6 and HXY7. In the current study, microalgae were isolated using BG-11 medium containing 1.5% agar. Spread plates were inoculated with 0.1–0.2 mL of water sample or soil suspension solution and incubated for 5–7 days at 25 °C, 60 µmol photons m−2 s−1 and a 14:10 h light:dark photoperiod. Single colonies were picked from the spread plates and inoculated into wells of a 12-well microtiter plate with liquid medium, where green colony was clearly observed under the microscope. Culture purity was confirmed by repeated plating and by regular observation under a microscope.
Microalgae dry weight(mg L−1) = (m2 − m1)/V
(1)
The filtrate (filtered through 0.45 μm filter (Whatman # 1) with a vacuum pump) was analyzed for water quality. The total organic carbon (TOC) concentration was determined using a total organic carbon analyzer (TOC-L CPN, Shimadzu Instruments Co., Ltd., Suzhou, China). Ammonia nitrogen (NH4+-N) was determined by Nessler's reagent spectrophotometry (APHA, 2005). Total dissolved nitrogen (TDN) was determined by potassium persulfate oxidation-ultraviolet spectrophotometry (Ebina et al., 1983). The total dissolved phosphorus (TDP) was determined by molybdenum antimony anti-spectrophotometry (Ebina et al., 1983). The removal efficiency of nutrients was calculated by Eq. (2) where ηi (i was TOC, NH4+-N, TDN or TDP) was the removal efficiency, Ci0 was the initial water concentration and Cia was the measured water concentration.
2.2. Identification of microalgae
ηi (%) = (Ci0 − Cia)/Ci0 × 100
Microalgae strains were preliminarily identified by optical microscope analysis and then were further confirmed by 18S rRNA gene sequence analysis. An aliquot of cultured cells (30–60 mL) was harvested during the exponential growth phase and centrifuged (10,000 rpm for 10 min) to concentrate. The microalgae pellet was then freeze-dried (LGJ-10C, China), and the DNA was extracted using a Plant Genomic DNA extraction kit. Eukaryotic primers P73F (AATCAGTTATAGTTTATTTGRTGGTACC) and P47R (TCTCAGGCTCCCTCTCCGGA) were used for PCR amplification. PCR products were analyzed by electrophoresis on a 0.8% (w/v) agarose gel to confirm the presence and approximate concentration of the products, which were then sequenced (Hongzhong Baide, Jiangsu, China). The 18S rRNA gene sequences of the isolates were searched against GenBank using BLAST and were manually aligned with representative sequences from microalgal strains and related taxa, according to similarities determined by the Clustal W program (Larkin et al., 2007). The phylogenetic tree was constructed using MEGA4 (Tamura and Dudley, 2007) software.
(2)
2.5. Lipid extraction and fatty acid analysis Microalgae solution (30 mL) was centrifuged in a high-speed centrifuge (10,000 rpm) for 6 min. The microalgal pellet was then placed in an oven (60 °C) and dried to constant weight. The lipids were extracted using the methanol-chloroform method (Dyer and Bligh, 1959). Approximately 100 mg of dry microalgae was weighed (Wa) into a 50 mL centrifuge tube, and 12 mL of 1:2 chloroform–methanol solution was added. The centrifuge tube was sonicated for 1 h at 25 °C and then centrifuged for 6 min (10,000 rpm); the sonicated suspension was filtered through a 0.45 µm filter membrane (Whatman #1) and transferred to a new centrifuge tube. The extraction was repeated for the remaining microalgal residue and both filtrates were combined. The combined filtrate was thoroughly mixed with 16 mL of 5% NaCl. The lower layer solution was separated into a round bottom flask and concentrated by a rotary evaporator (RE-2000A, Yarong Biochemical Instrument Company, Shanghai, China). The remaining solution was transferred to a 4 mL vial (vial dry weight recorded as Wi). The vial was dried using a nitrogen purging instrument (ND200-1, Hang Zhou Rui Cheng Instrument Co., Ltd., Hangzhou, China) at 45 °C until the sample reached a constant mass (recorded as Wf). The lipid content, production and productivity were calculated as in Eqs. (3)–(5), respectively.
2.3. Experimental culture conditions Simulated wastewater was used as the growth medium in the current study and was composed of (in mg L-1): CH3COONa 3000, NH4Cl 200, KH2PO4 60, FeSO4·7H2O 0.55, MgSO4·7H2O 66, CaCl2 6, CuSO4·5H2O 0.08, Co(NO3)2·6H2O 0.05, H3BO3 2.86, MnCl2·4H2O 1.86, ZnSO4·7H2O and Na2MoO4·2H2O 0.39. The experimental device was a glass tube with an inner diameter of 8 cm and a volume of 1.5 L. Aeration was applied at 0.16 L min−1; daytime aeration was intermittent (aeration:no aeration = 2:1h), whereas at night there was no aeration. The initial inoculation density was 1 × 105 cells mL−1. The culture temperature and illumination were 25 °C and 60 µmol photons m−2 s−1, respectively, and the cultures were incubated with a 14:10 h
Lipid content (%) = (Wf − W)/Wa × 100 i
(3)
Lipid production(mg L−1) = lipid content × microalgaedry weight
(4)
Lipid productivity(mg L−1day −1) = lipid production/day
(5)
Fatty acid methyl esters (FAMEs) were analyzed using the method of Lepage and Roy (1985). The extracted lipids were completely dissolved in 4 mL of 14% BF3-methanol solution and then boiled in a water bath 2
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wastewater containing complex pollutants and continuously purify the wastewater. Álvarez-Díaz et al. (2017) found that Scenedesmus obliquus could survive in municipal wastewater with TOC, TP and total nitrogen (TN) contents of 10.67, 1.55 and 20.09 mg L-1, respectively, with biomass productivity of 81.0 mg L-1 day−1. Komolafe et al. (2014), observed biomass productivity of Desmodesmus sp. of 13.0 mg L-1 day−1 using sewage treatment plant influent as the culture medium (with TP and TN of 35.4 and 65.6 mg L-1, respectively). In the current study, Desmodesmus sp. HXY7 had the highest biomass productivity of 57.9 mg L-1 day−1, which was much higher than that of Komolafe et al. (2014) and was close to the value shown by Álvarez-Díaz et al. (2017).
for 15 min. After cooling to room temperature, 2 mL of n-heptane and 4 mL of saturated NaCl solution were added, and the mixture was shaken well. After stratification, the supernatant was filtered using a 0.22-µm organic phase needle filter (TPFM012, Nanjing Tai Pu Rui Instrument Equipment Co., Ltd., China). FAMEs were analyzed using a gas chromatograph (GC-2014C, Shimadzu, Japan) equipped with flame ionization detector (FID) and a DB-5 ms (60 m) capillary column. The carrier gas was N2, and the injection and detector temperatures were 250 °C and 290 °C, respectively. The initial column temperature was 120 °C held for 3 min; it was then raised to 220 °C at 4 °C min−1 and held for 5 min; finally, the temperature was increased to 280 °C at 3 °C min−1 and held for 20 min. The injection volume was 1 μL.
3.3. Nutrient removal 2.6. Pigment analysis TOC, TP and TN concentrations in the simulated wastewater in this study were 878, 13.67 and 52.34 mg L−1, respectively. After 8 days of cultivation, the removal efficiency of NH4+-N by all eight microalgae species was > 98% (Fig. 3A). The removal efficiencies of TDN and TOC were > 90% and > 85%, respectively (Fig. 3B, C). > 70% of TDP was removed (Fig. 3D). Among the eight microalgae species, Scenedesmus sp. HXY2 showed the best nutrient removal efficiency, with removal efficiencies for TOC, NH4+-N, TDN and TDP of 96.07%, 99.09%, 96.62% and 94.52%, respectively. By day 12, the removal efficiencies for TOC, NH4+-N and TDN by the eight microalgae species were close to 100%. The removal efficiency for TDP by most species was > 95%, except for Scenedesmus sp. HXY4 and Desmodesmus sp. HXY8, which had removal efficiencies of ~72%. Kothari et al. (2013) cultured Chlamydomonas polypyrenoideum in dairy wastewater for 10 days and showed removal efficiencies for NO3–N, NH4+-N, TP and COD of 90%, 90%, 70% and 60%, respectively. Zhang et al. (2011) reported removal efficiencies for TOC, TN, and TP of 99.6%, 87.6%, and 69.8%, respectively, when Chlorella vulgaris was cultured in photomicrobial fuel cell wastewater for six days. Iasimone et al. (2018) showed that the removal rate of NH4+-N by mixed microalgae (60% Cyanobacteria and 40% Chlorophyta) in urban wastewater was > 74.8%. In this study, the NH4+-N removal efficiencies by the eight microalgae were higher than the values mentioned above. Additionally, all eight microalgae species could survive at high TOC concentrations (878 mg L-1). Furthermore, the isolated microalgae showed excellent abilities to degrade TOC, with removal rates of approximately 100 mg TOC L-1 day−1.
The microalgae solution (10 mL) was filtered through a 47-mm GF/ F filter (Whatman, Nanjing, China). The filter was placed in a 25-mL stoppered tube, and 5 mL of acetone solution was injected to permeate the filter; then it was treated with ultrasound for 10 min at 4 °C. The solution was incubated in the dark at −20 °C for 24 h, and then the filtrate was extracted with a polytetrafluoroethylene syringe filter and loaded into a 1.5 mL brown chromatography bottle for high performance liquid chromatography (HPLC) (Agilent-1100 series, USA) detection. Two mobile phases were used: A (2:1 = methanol: 0.5 mol L-1 ammonium acetate) and B (3:7 = methanol: acetonitrile). The linear gradients used were: (min, mobile phase A%, mobile phase B%) (0, 100, 0), (10, 40, 60), (20, 20, 80), (30, 0, 100), (40, 0, 100), (50, 0, 100), (52, 25, 75), (60, 100, 0), and (65, 100, 0). The injection volume was 100 μL, and the wavelengths for detection were 440, 266 and 300 nm. 2.7. Statistics All data were expressed as mean ± standard deviation. Differences in data between treatments were determined by analysis of variance (ANOVA) using SPSS 19.0 software (p < 0.05) (IBM Corp., Armonk, NY, USA). 3. Results and discussion 3.1. Identification of microalgae species Eight microalgae species were isolated in the current study. HXY1 existed as 2- or 4-cell populations. The cells were spindle-shaped with lengths of 7.81–10.53 µm. The other seven microalgae species were mostly unicellular. HXY2 and HXY3 cells were similar in morphology, with a spindle shapes and lengths of 7.00–12.09 µm. HXY5, HXY6 and HXY7 were larger, with lengths of 7.22–12.50 µm. HXY4 and HXY8 were much smaller, with spherical cell morphology having a diameter of 5.57–7.51 µm. The eight microalgae species were all in the phylum Chlorophyta and were identified as Scenedesmus sp. by morphological observation. The phylogenetic tree of the eight microalgae species obtained by 18S rRNA gene sequencing analysis is shown in Fig. 1. HXY1, HXY2, HXY4, HXY5 and HXY6 were classified as Scenedesmus (Scenedesmus sp. HXY1, HXY2, HXY4, HXY5 and HXY6, respectively). HXY3, HXY7 and HXY8 were classified as Desmodesmus (Desmodesmus sp. HXY3, HXY7 and HXY8, respectively).
3.4. Lipid productivity and fatty acid composition Desmodesmus sp. HXY3 had the highest lipid productivity of 7.50 mg L-1 day−1 (Fig. 4C) because of its high biomass productivity (Fig. 2A). Generally, higher lipid productivity can be obtained in standard media because of the balanced nutrient and mineral compositions. For example, Zhang et al. (2014) isolated Chlorella vulgaris CJ15 in the coastal areas of Qingdao using F/2 medium and demonstrated lipid productivity of 11.7 mg L-1 day−1. Chen et al. (2012) found that Scenedesmus sp. NJ-1, isolated from Antarctica, had a lipid productivity of 36.8 mg L-1 day−1 in BG-11 medium. Therefore, Desmodesmus sp. HXY3, isolated in this study, is expected to achieve higher lipid productivity in standard media. The medium nutrient and mineral compositions had significant effects on the lipid content of microalgae. The lipid content of the eight microalgae species ranged from 10% to 16% (Fig. 4A). Scenedesmus sp. HXY2 had the highest lipid content (15.56%). Yang et al. (2011) reported that Chlorella ellipsoidea YJ1, isolated from the secondary effluent of the Beijing Wastewater Treatment Plant, had lipid content and lipid productivity of 43% and 11.4 mg L-1 day−1, respectively, when it was cultured in secondary effluent (COD, TN and TP of 24, 7 and 0.46 mg L-1, respectively). Xin et al. (2010) found that the lipid content and lipid productivity of Scenedesmus sp. LX1 were 31–33% and 8 mg L1 day−1, respectively, when cultured in secondary effluent (COD, TN
3.2. Biomass production The biomass of the eight strains after 12 days of culture in simulated wastewater is shown in Fig. 2A. Desmodesmus sp. HXY7 had the highest cell density and dry weight of 1.72 × 107 cells mL−1 and 695 mg L-1, respectively. Microalgae, especially Chlorophyta, have high tolerance to pollutants, including ammonia, organic matter and heavy metals (Cai et al., 2013; Komolafe et al., 2014). Thus, they can survive in 3
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Fig. 1. Phylogenetic tree showing the relationships among the 18S rRNA gene sequences of the eight microalgae isolates and the most similar sequences retrieved from the NCBI nucleotide database.
and TP content of 24, 15.5 and 0.5 mg L-1, respectively) from the Beijing Domestic Sewage Treatment Plant. In the current study, the TOC, TN and TP contents in simulated wastewater were much higher than those of Yang et al. (2011) and Li et al. (2010), but the lipid productivity was much lower than in those studies. This was because
the high N concentration prevented lipid accumulation (Yeh and Chang, 2011; Yeh and Chang, 2012; Pan et al., 2011). The results of this study indicated that Scenedesmus sp. HXY2 had sufficient lipid content for microalgal biodiesel production in high ammonia and organic content wastewater.
Fig. 2. (A) The cell density and the dry weight and (B) the polysaccharides and protein contents of the eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05). 4
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Fig. 3. (A) The removal efficiency for total organic carbon (TOC), (B) ammonium nitrogen (NH4+-N), (C) total dissolved nitrogen (TDN) and (D) total dissolved phosphorus (TDP) by eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05).
Fig. 4. (A) The lipid content, (B) lipid production and (C) lipid productivity of eight microalgae species cultured in simulated wastewater for 12 days. Different lowercase letters indicate significant differences between treatments (p < 0.05). 5
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Table 1 Fatty acid compositions of eight microalgae species (%).
C14:0 C15:0 C16:0 C16:1 C18:0 C18:1 C18:2 C18:3 C22:0 C22:2 C24:0 Unsaturated C16-C18
Scenedesmus sp. HXY1
Scenedesmus sp. HXY2
Desmodesmus sp. HXY3
Scenedesmus sp. HXY4
Scenedesmus sp. HXY5
Scenedesmus sp. HXY6
Desmodesmus sp. HXY7
Desmodesmus sp. HXY8
1.64 ± 0.16 ND 43.68 ± 11.83 3.08 ± 0.9 4.85 ± 1.49 14.48 ± 2.59 32.27 ± 14.96 ND ND ND ND 49.83 98.36
1.26 ± 0.31 0.88 ± 0.09 33.63 ± 1.9 2.66 ± 0.1 3.49 ± 0.42 29.19 ± 14.92 25.57 ± 13.84 2.65 ± 0.23 0.95 ± 0.23 ND ND 60.07 97.19
1.43 ± 0.19 ND 45.91 ± 6.94 2.17 ± 0.15 5.31 ± 1.12 18.21 ± 10.51 23.52 ± 2.14 2.05 ± 0.29 1.41 ± 0.27 ND ND 45.95 97.17
1.94 ± 0.17 ND 41.23 ± 1.78 2.16 ± 0.2 5.91 ± 0.19 26.98 ± 1.54 17.08 ± 0.62 2.99 ± 1 ND ND 1.69 ± 0.23 49.21 96.35
ND ND 34.5 ± 0.01 1.9 ± 0.48 7.49 ± 0.44 17.46 ± 0.23 32.69 ± 0.17 5.96 ± 0.01 ND ND ND 58.01 100
ND ND 39.07 ± 2.87 1.89 ± 0.23 9.66 ± 0.48 22.6 ± 1.63 26.78 ± 0.99 ND ND ND ND 51.27 100
1.37 ± 0.69 ND 49.97 ± 4.64 1.18 ± 0.5 7.85 ± 1.19 7.96 ± 2.2 28.1 ± 6.44 1.67 ± 0.43 ND 1.9 ± 0.02 ND 40.81 96.73
1.89 ± 0.67 ND 45.78 ± 2.87 2.06 ± 0.26 8.22 ± 1.51 15.49 ± 1.59 15.72 ± 4.76 7.84 ± 5.32 ND 1.99 ± 0.06 1.51 ± 0.45 43.1 95.11
ND: not detected. Table 2 Pigment compositions and contents of eight microalgae species.
Scenedesmus sp. HXY1 Scenedesmus sp. HXY2 Desmodesmus sp. HXY3 Scenedesmus sp. HXY4 Scenedesmus sp. HXY5 Scenedesmus sp. HXY6 Desmodesmus sp. HXY7 Desmodesmus sp. HXY8
Viola (μg g−1)
Lut (μg g−1)
Chl-b (μg g−1)
Chl-a (μg g−1)
192.32 ± 14.51 ND 65.38 ± 12.96 59.38 ± 17.04 37.17 ± 32.33 ND 110.28 ± 12.15 ND
1241.21 ± 311.12 1476.15 ± 452.36 1097.87 ± 386.61 1635.37 ± 568.86 1040.15 ± 80.73 774.64 ± 454.17 3956.63 ± 226.15 1086.55 ± 216.56
92.94 ± 7.23 90.46 ± 22.21 159.53 ± 50.68 196.68 ± 75.66 172.35 ± 47.92 131.5 ± 64.11 499.57 ± 94.45 144.01 ± 66.44
318.31 ± 34.58 449.96 ± 120.54 511.67 ± 154.69 477.29 ± 162.24 570.11 ± 103.66 421.65 ± 184.39 1592.53 ± 362.23 404.35 ± 166.41
Viola: Violaxanthin. Lut: Lutein. Chl-b: Chlorophyll-b. Chl-a: Chlorophyll-a. ND: not detected.
cholesterol (Cristiano et al., 2018). For this study, the polysaccharide and protein contents are shown in Fig. 2B. Scenedesmus sp. HXY5 had the highest polysaccharide content (24.34 pg cell−1) and the highest protein content (46.39 pg cell−1). Lutein can play a positive role in preventing blindness and decreased vision caused by macular degeneration (Shi et al., 1997; Zhang et al., 1999; Wu et al., 2007). The pigment composition of the eight microalgae species is shown in Table 2. Desmodesmus sp. HXY7 had the highest lutein content (3.96 mg g−1) among all eight microalgae. The highest chlorophyll-a and chlorophyll-b contents were obtained for Desmodesmus sp. HXY7, 1.59 and 0.50 mg g−1, respectively. Minhas et al. (2016) summarized the lutein content of 22 microalgae, and the average value was 1.4 mg g−1. The lutein content of Desmodesmus sp. HXY7 in the current study (3.96 mg g−1) was much higher than the average value and higher than 91% of microalgae in the summary by Minhas et al. (2016). Ho et al. (2014) isolated six Scenedesmus species and reported that the average lutein content was 2.40 mg g−1, which was also lower than the value of Desmodesmus sp. HXY7. Therefore, the newly isolated Desmodesmus sp. HXY7 could be a potential source for lutein production.
The most common FAMEs present in biodiesel are palmitic acid (16:0), stearic acid (18:0), oleic acid (18:1), linoleic acid (18:2), and linolenic acid (18:3) (Knothe, 2013). The saturated fatty acids (16:0) and monounsaturated fatty acids (18:1) play an important role in determining biodiesel performance, such as oxidative stability and cold flow properties of biodiesel (Kaur et al., 2012). The fatty acid compositions of eight microalgae species cultured in simulated wastewater medium are shown in Table 1. The main fatty acids of the eight species of microalgae were palmitic acid (16:0), oleic acid (18:1) and linoleic acid (18:2), accounting for > 80% of the total fatty acid composition. Scenedesmus sp. HXY2 had the highest unsaturated fatty acid composition of 60.07%, followed by Scenedesmus sp. HXY5 (58.01%). The proportion of C16–C18 fatty acids for all eight microalgae species was > 95%. Thao et al. (2017) isolated lipid-producing microalgae in Vietnam, and, of these, Mychonastes sp. N16 had a maximum C16–C18 fatty acid content of 61.62%. Yeh and Chang, 2012 reported that the proportion of C16–C18 fatty acids of Chlorella vulgaris ESP-31 in standard medium was 66.99–89.59%. Abou-Shanab et al. (2011b) isolated five lipid-producing microalgae in South Korea, and their proportion of C16–C18 fatty acids ranged from 56% to 97%. In this study, the proportions of C16–C18 fatty acids for the eight microalgae species were similar to the values mentioned above, indicating that the isolated microalgae have good lipid production potential.
4. Conclusion Eight microalgae species were isolated from the Hexi Corridor. They were cultured in simulated wastewater for 12 days. Scenedesmus sp. HXY2 had the highest nutrient removal efficiency and lipid content. On day 8, the removal efficiencies of TOC, NH4+-N, TDN and TDP by Scenedesmus sp. HXY2 were 96.07%, 99.09%, 96.62% and 94.52%, respectively. The lipid content and lipid productivity of this alga was 15.56% and 5.67 mg L-1 day−1, respectively. Our results indicated that Scenedesmus sp. HXY2 has great potential for culturing in wastewater with high ammonia nitrogen and organic concentrations for simultaneous wastewater purification and lipid production.
3.5. Byproducts Polysaccharides produced by microalgae are usually very complex, and the complex structure of some polysaccharides can result in anticancer activity (Spolaore et al., 2006; Chaiklahan et al., 2013). Mišurcová et al. (2014) found that Chlorella sp. and Spirulina sp. had high protein contents (> 50% of dry weight) and could be used as sources of essential amino acids. Additionally, microalgae proteins have shown a significant effect in reducing blood pressure and lowering 6
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Manage. 223. Jebali, A., Acién, F.G., Gómez, C., Fernández-Sevilla, J.M., Mhiri, N., Karray, F., Dhouib, A., Molina-Grima, E., Sayadi, S., 2015. Selection of native Tunisian microalgae for simultaneous wastewater treatment and biofuel production. J. Bioresour Technol. 198. Kaur, S., Sarkar, M., Srivastava, R.B., Gogoi, H.K., Kalita, M.C., 2012. Fatty acid profiling and molecular characterization of some freshwater microalgae from India with potential for biodiesel production. New Biotech. 29 (3), 332–344. Knothe, G., 2013. “Designer” biodiesel: optimizing fatty ester composition to improve fuel properties. Energy Fuel 22 (2). Komolafe, O., Velasquez Orta, S.B., Monje-Ramirez, I., Noguez, I.Y., Harvey, A.P., Orta, L., María, T., 2014. Biodiesel production from indigenous microalgae grown in wastewater. Bioresour. Technol. 154, 297–304. Kothari, R., Prasad, R., Kumar, V., Singh, D.P., 2013. Production of biodiesel from microalgae Chlamydomonas polypyrenoideum grown on dairy industry wastewater. Bioresour. Technol. 144, 499–503. Larkin, M.A., Blackshields, G., Brown, N.P., Chenna, R., Mcgettigan, P.A., Mcwilliam, H., Valentin, F., Wallace, I.M., Wilm, A., Lopez, R., 2007. Clustal w and clustal x version 2.0. Bioinformatics 23 (21), 2947–2948. Lepage, G., Roy, C.C., 1985. Improved recovery of fatty acid through direct transesterification without prior extraction or purification. J. Lipid Res. 25 (12), 1391–1396. Li, K., Liu, Q., Fang, F., Luo, R.H., Lu, Q., Zhou, W.G., Huo, S.H., Cheng, P.F., Liu, J.Z., Addy, M., Chen, P., Chen, D.J., Ruan, R., 2019. Microalgae-based wastewater treatment for nutrients recovery: a review. J. Bioresour. Technol. 291. Minhas, A.K., Hodgson, P., Barrow, C.J., Sashidhar, B., Adholeya, A., 2016. The isolation and identification of new microalgal strains producing oil and carotenoid simultaneously with biofuel potential. Bioresour. Technol S0960852416304242. Mišurcová, L., Buňka, F., Vávra, A.J., Machů, L., Samek, D., 2014. Amino acid composition of algal products and its contribution to RDI. Food Chem. 151, 120–125. Pan, Y.Y., Wang, S.T., Chuang, L.T., Chang, Y.W., Chen, C.N.N., 2011. Isolation of thermo-tolerant and high lipid content green microalgae: oil accumulation is predominantly controlled by photosystem efficiency during stress treatments in Desmodesmus. Bioresour. Technol. 102 (22), 10510–10517. Shi, X.M., Chen, F., Yuan, J.P., Chen, H., 1997. Heterotrophic production of lutein by selected chlorella strains. J. Appl. Phycol. 9 (5), 445–450. Sivaramakrishnan, R., Incharoensakdi, A., 2018. Microalgae as feedstock for biodiesel production under ultrasound treatment – a review. Bioresour. Technol. Spolaore, P., Joannis-Cassan, C., Duran, E., Isambert, A., 2006. Commercial applications of microalgae. J. Biosci. Bioeng. 101 (2), 87–96. Tamura, K., Dudley, J., 2007. MECA4: Molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596–1599. Thangavel, K., Krishnan, P.R., Nagaiah, S., Kuppusamy, S., Dananjeyan, B., 2018. Growth and metabolic characteristics of oleaginous microalgal isolates from Nilgiri biosphere reserve of India. BMC Microbiol. 18 (1), 1. Thao, T.Y., Linh, D.T.N., Si, V.C., Carter, T.W., Hill, R.T., 2017. Isolation and selection of microalgal strains from natural water sources in Viet Nam with potential for edible oil production. Mar. Drugs 15 (7), 194. Tiwari, A., Rajesh, V.M., Yadav, S., 2018. Biodiesel production in micro-reactors: a review. Energy. Sustain. Dev. 43, 143–161. Wu, Z.Y., Shi, C.L., Shi, X.M., 2007. Modeling of lutein production by heterotrophic Chlorellain batch and fed-batch cultures. World J. Microb. Biotechnol. 23 (9), 1233–1238. Xin, L., Hong-Ying, H., Jia, Y., 2010. Lipid accumulation and nutrient removal properties of a newly isolated freshwater microalga, Scenedesmus sp. LX1, growing in secondary effluent. New. Biotech. 27 (1), 59–63. Yang, H.L., Lu, C.K., Chen, S.F., Chen, Y.M., Chen, Y.M., 2010. Isolation and characterization of taiwanese heterotrophic microalgae: screening of strains for docosahexaenoic acid (dha) production. Mar. Biotechnol. 12 (2), 173–185. Yang, J., Xu, M., Hu, Q., Sommerfeld, M., Chen, Y., 2011. Life-cycle analysis on biodiesel production from microalgae: water footprint and nutrients balance. Bioresour. Technol. 102, 159–165. Yeh, K.L., Chang, J.S., 2011. Nitrogen starvation strategies and photobioreactor design for enhancing lipid content and lipid production of a newly isolated microalga chlorella vulgaris esp-31: implications for biofuels. Biotech. J. 6 (11), 1358–1366. Yeh, K.L., Chang, J.S., 2012. Effects of cultivation conditions and media composition on cell growth and lipid productivity of indigenous microalga chlorella vulgaris esp-31. Bioresour. Technol. 105, 120–127. Zhang, B., Wang, L., Hasan, R., Shahbazi, A., 2014. Characterization of a native algae species chlamydomonas debaryana: strain selection, bioremediation ability, and lipid characterization. BioResources. 9 (4). Zhang, X.W., Shi, X.M., Chen, F., 1999. A kinetic model for lutein production by the green microalga Chlorella protothecoidesin heterotrophic culture. J. Ind. Microbiol. Biotech. 23 (6), 503–507. Zhang, Y., Noori, J.S., Angelidaki, I., 2011. Simultaneous organic carbon, nutrients removal and energy production in a photomicrobial fuel cell (PFC). Energy Environ. Sci. 4 (10), 4340.
Sisi Ye: Data curation, Writing - original draft, Writing - review & editing. Li Gao: Writing - original draft, Writing - review & editing. Jing Zhao: Formal analysis, Investigation. Mei An: Methodology, Software. Haiming Wu: Writing - original draft, Validation. Ming Li: Conceptualization, Resources, Supervision, Writing - original draft, Writing - review & editing. Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This work was supported by the Fundamental Research Funds for Central Universities (Northwest A&F University, Grant No. 2452018142); Dr. Ming Li is funded as Tang Scholar by Cyrus Tang Foundation and Northwest A&F University. References Abou-Shanab, R.A.I., Hwang, J.H., Cho, Y., Min, B., Jeon, B.H., 2011a. a. Characterization of microalgal species isolated from fresh water bodies as a potential source for biodiesel production. Appl. Energy 88 (10), 3300–3306. Abou-Shanab, R.A.I., Matter, I.A., Kim, S.N., Oh, Y.K., Choi, J., Jeon, B.H., 2011b. b. Characterization and identification of lipid-producing microalgae species isolated from a freshwater lake. Biomass. Bioenerg. 35 (7), 3079–3085. Álvarez-Díaz, P.D., Ruiz, J., Arbib, Z., Barragán, J., Garrido-Pérez, M.C., Perales, J. A., 2017. Freshwater microalgae selection for simultaneous wastewater nutrient removal and lipid production. Algal Res. APHA, 2005. Standard methods for the examination of water and wastewater, 21st ed.; APHA and AWWA and WEF DC, Washington, DC. Brennan, L., Owende, P., 2010. Biofuels from microalgae-a review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sust. Energy Rev. 14 (2), 557–577. Cai, T., Park, S.Y., Li, Y., 2013. Nutrient recovery from wastewater streams by microalgae: status and prospects. Renew. Sust. Energy Rev. 19, 360–369. Chaiklahan, R., Chirasuwan, N., Triratana, P., Loha, V., Tia, S., 2013. Polysaccharide extraction from Spirulina sp. And its antioxidant capacity. Int. J. Biol. Macromol. 58, 73–78. Chen, Z., Gong, Y., Fang, X.T., 2012. Scenedesmus. NJ-1 isolated from Antarctica: a suitable renewable lipid source for biodiesel production. World J. Microb Biotech. 28 (11), 3219–3225. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25 (3), 294–306. Clarens, A.F., Resurreccion, E.P., White, M.A., Colosi, L.M., 2010. Environmental life cycle comparison of algae to other bioenergy feedstocks. Environ. Sci. Technol. 44 (5), 1813–1819. Cristiano, J.A., Lidiane, M.A., Meriellen, D., Claudio, A.N., Maria, A.M., 2018. Chlorella and spirulina microalgae as sources of functional foods, nutraceuticals, and food supplements; an overview. MOJ Food Process Technol. 6 (2), 00144. Dyer, E.G., Bligh, W.J., 1959. A rapid method of total lipid extraction and purification. J. Biochem Physiol. 37. Ebina, J., Tsutsui, T., Shirai, T., 1983. Simultaneous determination of total nitrogen and total phosphorus in water using peroxodisulfate oxidation. Water Res. 17 (12). Fernando, S., Hall, C., Jha, S., 2006. NOx reduction from biodiesel fuels. Energy Fuel. 20 (1), 376–382. Franchino, M., Comino, E., Bona, F., Riggio, V.A., 2013. Growth of three microalgae strains and nutrient removal from an agro-zootechnical digestate. Chemosphere 92 (6), 738–744. Ho, S.H., Chan, M.C., Liu, C.C., Chen, C.Y., Lee, W.L., Lee, D.J., Chang, J.S., 2014. Enhancing lutein productivity of an indigenous microalga Scenedesmus obliquus FSP3 using light-related strategies. Bioresour. Technol. 152, 275–282. Hu, J.J., Nagarajan, D., Zhang, Q.G., Chang, J.S., Lee, D.J., 2018. Heterotrophic cultivation of microalgae for pigment production: a review. Biotechnol. Adv. 36, 54–67. Iasimone, F., Panico, A., De Felice, V., Fantasma, F., Iorizzi, M., Pirozzi, F., 2018. Effect of light intensity and nutrients supply on microalgae cultivated in urban wastewater: Biomass production, lipids accumulation and settleability characteristics. J. Environ.
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