- Email: [email protected]
Carbon, nitrogen, and phosphorus removal, and lipid production by three saline microalgae grown in synthetic wastewater irradiated with different photon fluxes
Algal Research 34 (2018) 97–103
Contents lists available at ScienceDirect
Algal Research journal homepage: www.elsevier.com/locate/algal
Carbon, nitrogen, and phosphorus removal, and lipid production by three saline microalgae grown in synthetic wastewater irradiated with different photon fluxes
T
⁎
Manuel Sacristán de Alvaa, Víctor Manuel Luna Pabelloa, , María Teresa Orta Ledesmab, Modesto Javier Cruz Gómezc a Laboratorio de Microbiología Experimental, Departamento de Biología, Facultad de Química, UNAM, Av. Universidad No. 3000, Col. Universidad, Nacional Autónoma de México C.U., Delegación Coyoacán, C.P. 04510 Ciudad de México, México, Mexico b Laboratorio de Ingeniería Ambiental, Edificio 5, Instituto de Ingeniería, UNAM, Mexico c Laboratorio 212, Departamento de Ingeniería Química, Conjunto E. Facultad de Química, UNAM, Mexico
A R T I C LE I N FO
A B S T R A C T
Keywords: Mariculture Lipids Biodiesel Nutrient stress Marine microalgae
Mariculture production has increased in the last decades, with untreated wastewater discharged directly into the sea, impacting coastal ecosystems. There is a need for mariculture wastewater treatment systems that are costeffective. This can be met by the implementation of wastewater treatment systems that in addition to removing pollutants are capable of producing valuable by-products such as biomass for the biofuel industry. In this study, Dunaliella sp., Nannochloropsis sp. and Tetraselmis sp. microalgae were cultivated in controlled environments simulating mariculture wastewaters. Single stage culture systems were used to grow these microalgae, the growing conditions included inducing stress with different photon flux densities (900, 1500 and 2000 μmol m−2 s−1), and low carbon, nitrogen and phosphorus concentrations obtained at the stationary phase, in order to force these microalgae to increase their lipid content. The best results were obtained with Tetraselmis sp., which achieved 132.8 mg L−1 day−1 of biomass productivity at 900 μmol m−2 s−1. Nevertheless the best lipid productivity was reached at 1500 μmol m−2 s−1, also by Tetraselmis sp., being 29.5 mg L−1 day−1, where biomass productivity was of 124.5 mg L−1 day−1. All three microalgae species were able to remove > 90% of nitrogen and orthophosphates, and 80% of carbon, which makes them suitable for treating mariculture wastewater, and in addition, represent a valuable high lipid content biomass byproduct usable as raw material for biodiesel synthesis.
1. Introduction Aquaculture production has increased substantially for the past 40 years [1,2]. This industry is responsible of producing wastewaters with high contents of suspended solids and dissolved nutrients, which are discharged untreated into water bodies, causing eutrophication [3], generating damages to coastal ecosystems, and eventually the loss of biodiversity [4]. Conventional biological and physicochemical wastewater treatments can remove nutrients contained in this kind of wastewaters, and produce a good quality effluent. However, these methods involve the production of sludges which in turn must be treated prior to discharge; generating additional costs. In this regard, microalgae culture in aquaculture wastewaters offers the advantages of removing nutrients
while simultaneously producing biomass that may be used as raw material for valuable products, such as biofuels [5]. The economic feasibility of microalgae biofuels faces various challenging limiting factors, such as the need for cultures with high lipid productivity in the shortest time period [6,7]. It is well-known that cell growth and lipid accumulation do not take place simultaneously during culture, so the techniques for increasing lipid accumulation often render low biomass quantities, and consequently, low lipid productivity [8]. To address this issue, the microalgae culture is carried out in two stages. In the first stage, microalgae are cultured in optimum growth conditions; and in the second stage, the cultures are subject to stress conditions in order to enhance the accumulation of energy in the form of neutral lipids (triacylglycerols) by altering their biosynthetic pathways. The result is no increase in microalgae density, but an important
⁎
Corresponding author. E-mail addresses: [email protected] (M. Sacristán de Alva), [email protected] (V.M. Luna Pabello), [email protected] (M.T. Orta Ledesma), [email protected] (M.J. Cruz Gómez). https://doi.org/10.1016/j.algal.2018.07.006 Received 22 February 2018; Received in revised form 26 June 2018; Accepted 8 July 2018 2211-9264/ © 2018 Elsevier B.V. All rights reserved.
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
Fig. 1. Conceptual model of lipid synthesis during nitrogen limitation and light excess in a microalgae.
increase in lipid content [9,10]. There are several physicochemical factors that affect microalgae metabolism, for enhancing lipid production, such as light intensity, photoperiod, temperature, salinity, nutrient concentration in the growth medium (carbon, phosphates, nitrogen), and nitrogen (ammonium or nitrate) and carbon (glucose or glycerol, among others) sources. By exposing microalgae to environmental stress in laboratory scale cultures, through the modification of said factors, high lipid productivity has been achieved [11]. Nitrogen limitation causes three main changes: decrease in the thylakoid membrane; stimulation of phospholipid hydrolysis; and fatty acid synthesis enzyme activation. These changes cause an increase of the intracellular fatty acids content [12]. Generally, when microalgae are grown under low light intensities, the assimilated carbon is used for the synthesis of amino acids and other essential components, but under light saturation conditions sugars, lipids, and starch are formed [13]. An adequate light intensity contributes to lipid overproduction, which may be the result of the overgeneration of photo-assimilated compounds that are converted into chemical energy [14]. Fig. 1 shows a conceptual model of lipid accumulation in microalgae. During the nitrogen limitation stage, the pathways marked with dashed arrows are activated for lipid production; other pathways are the membrane lipid recycling, and the protein degradation. Pathways indicated with X are inhibited mostly because there is a reduction in the photosynthetic rate, and carbon fixation through photosynthesis is reduced, as well as protein synthesis. When there is light excess the
Table 1 Doubling times of the 3 microalgae species (Data shown is the mean, n = 3). Microalgae
Tetraselmis sp. Dunaliella sp. Nannochloropsis sp.
Photosynthetically active photon flux density (PAPFD) (μmol m−2 s−1) 900
1500
2000
3.21 4.13 5.83
3.56 4.13 5.27
4.23 4.80 6.67
Table 2 Specific growth rate and biomass productivity of the 3 microalgae species (Data shown is the mean, n = 3). Microalgae
PAPFD (μmol m−2 s−1)
900
1500
2000
Dunaliella sp.
Specific growth rate (day−1) Biomass productivity (mg L−1 day−1) Specific growth rate (day−1) Biomass productivity (mg L−1 day−1) Specific growth rate (day−1) Biomass productivity (mg L−1 day−1)
0.19 68.9
0.20 69.3
0.17 52.1
0.15 88.0
0.18 99.5
0.17 84.9
0.29 132.8
0.28 124.5
0.23 101.2
Nannochloropsis sp.
Tetraselmis sp.
Table 3 Nutrient removal kinetics and removal percentages of the 3 microalgae species (Data shown is the mean, n = 3). PAPFD (μmol m−2 s−1)
900
1500
2000
a
Total nitrogena Orthophosphates COD Total nitrogena Orthophosphates COD Total nitrogena Orthophosphates COD
Dunaliella sp.
Nannochloropsis sp.
Tetraselmis sp.
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
96.91 94.02 81.85 97.26 94.25 82.38 97.53 93.27 81.29
3.50 0.87 17.35 3.57 0.94 17.99 3.51 0.89 16.71
94.16 97.12 81.45 90.07 97.16 81.02 94.24 97.20 81.85
3.85 0.78 10.45 3.68 0.81 10.17 4.02 0.83 10.74
97.99 95.62 81.56 98.03 95.94 80.97 98.06 95.67 75.60
7.07 1.49 20.11 7.25 1.48 19.34 7.34 1.48 14.08
Total nitrogen: is the sum of the concentrations of nitrates, nitrites and ammonia. 98
Algal Research 34 (2018) 97–103
900 mol m s
1.6
1500 mol m s
-2 -1
-2 -1
2000 mol m s
1.4
250 -1
1.2 1.0 0.8 0.6 0.4
20
Nitrogen 900 µmol m-2s-1 COD 900 µmol m-2s-1 Orthophosphates 900 µmol m-2s-1 Nitrogen 1500 µmol m-2s-1 COD 1500 µmol m-2s-1 Orthophosphates 1500 µmol m-2s-1 Orthophosphates 2000 µmol m-2s-1 Nitrogen 2000 µmol m-2s-1 COD 2000 µmol m-2s-1
300
-2 -1
Total nitrogen (mg L ) COD (mg O2 L-1)
Dry biomass (g L-1)
1.8
Dunaliella sp.
350
200 150
18 16 14 12 10 8
100
6 50
0.2
4
0
0.0 0
5
10
15
-1
a) Tetraselmis sp.
2.0
Orthophosphates (mg L )
M. Sacristán de Alva et al.
2
20
0 0
Time (days)
2
4
6
8
10
12
14
16
18
20
Time (days) -1
Growth kinetics: k 900 =0.22 day , k 1500 =0.19 day , k 2000 =0.16 day 2.0 1.8
300
-2 -1
900 mol m s
250
-2 -1
1500 mol m s
1.6
-2 -1
2000 mol m s
Total nitrogen (mg L ) COD (mg O2 L-1)
1.4
-1
1.2 1.0 0.8 0.6
200
20 18 16 14 12
150
10 8
100
6 50
4
0.4
2
0
0.2
0
0.0 5
10
15
-50
20
0
5
10
Time (days)
Growth kinetics: k 900 =0.17 day-1, k 1500 =0.17 day-1, k 2000 =0.14 day-1
-2 -1
Dry biomass (g L-1)
Total nitrogen (mg L ) COD (mg O2 L-1)
1500 mol m s
1.6
-1
-2 -1
2000 mol m s
1.4 1.2 1.0 0.8 0.6 0.4
20
Nitrogen 900 µ mol m-2s-1 COD 900 µ mol m-2s-1 Orthophosphates 900 µ mol m-2s-1 -2 -1 Nitrogen 1500 µ mol m s
250
-2 -1
900 mol m s
1.8
20
Tetraselmis sp.
300
c) Nannochloropsis sp.
2.0
15
Time (days)
200
18 16 14
-2 -1
COD 1500 µ mol m s
12
Orthophosphates 1500 µ mol m-2s-1 Orthophosphates 2000 µ mol m-2s-1 -2 -1 Nitrogen 2000 µ mol m s
150 100
10 8
-2 -1
COD 2000 µ mol m s
6 50
4 2
0
0
0.2 0.0 0
5
10
15
0
20
-1
0
Orthophosphates (mg L )
Dry biomass (g L-1)
Nitrogen 900 µmol m-2s-1 COD 900 µmol m-2s-1 Orthophosphates 900 µmol m-2s-1 Orthophosphates 1500 µmol m-2s-1 Nitrogen 1500 µmol m-2s-1 COD 1500 µmol m-2s-1 Nitrogen 2000 µmol m-2s-1 COD 2000 µmol m-2s-1 Orthophosphates 2000 µmol m-2s-1
Nannochloropsis sp.
b) Dunaliella sp.
-1
-1
Orthophosphates (mg L )
-1
5
10
15
20
Time (days)
Time (days)
Growth kinetics: k 900 =0.12 day-1, k 1500 =0.13 day-1, k 2000 =0.10 day-1
Fig. 3. Nutrient concentration during the culture of the 3 microalgae species (Data shown is the mean +/−SD, n = 3).
Fig. 2. Growth curves of: a) Tetraselmis sp., b) Dunaliella sp. and c) Nannochloropsis sp. (Data shown is the mean +/−SD, n = 3).
is required [27]. In view of the foregoing, the purpose of this study is separately growing three marine microalgae in a single stage culture to overcome the difficulties of a two stage culture. Firstly, we seek to obtain the maximum microalgal biomass productivity during the logarithmic growing phase where the nutrients are present. Secondly obtaining high lipid accumulation as a result of the stress produced by the lack of nutrients when reaching the stationary phase. In order to determine the effect on the productivity of lipids in microalgae, three levels of photosynthetically active photon flux density were evaluated.
pathways marked with grey arrows get activated for lipid production, carotenoid synthesis pathways are also activated since carotenoids act as a sunblock for the cells. Adapted from [14–26]. Two-stage culture has the disadvantage of complicating the culture process by requiring additional infrastructure for transferring the culture from the first stage to the second, adding costs to the process. Furthermore, even if lipid accumulation increases, the growth rate is reduced, affecting the net lipid productivity. Consequently, a strategy that stimulates lipid accumulation without affecting microalgae growth
99
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
a) Tetraselmis sp.
120
2. Materials and methods 2.1. Microalgae and growth medium
900 µmol m s
-2 -1
1500 µmol m s
-2 -1
100
The microalgae used in this study were Dunaliella sp., Nannochloropsis, and Tetraselmis sp., because they are used as live food for mariculture organisms, and because of their lipid content assessed in other studies [28–30]. The synthetic wastewater (SWW) used in this study was prepared based in the characterization made of actual wastewater (AWW) effluents from growing commercial marine species, such as octopus, shrimp, seabass, corvina, snapper, and ornamental species, collected at the facilities of the Yucatan Academic Unit of the UNAM located in Sisal, Yucatan, Mexico. The values obtained for the AWW were the following: nitrate, 45 mg L−1; nitrite, 10 mg L−1; ammonium, 30 mg L−1; orthophosphate, 17 mg L−1; and dissolved organic matter (as chemical oxygen demand, COD), 270 mg O2 L−1. SWW was prepared from distilled water added with a sufficient amount of Instant Ocean sea salt to obtain the desired salinity (31 psu for Nannochloropsis sp. and Tetraselmis sp., and 41 psu for Dunaliella sp.; these salinities provide optimum growth for each species. In order to match the nitrogen, phosphorus, and carbon concentrations, of the AWW, KNO3, NaNO2, (NH4)2SO4, K3PO4, and sodium acetate were added in sufficient quantities to mimic AWW concentrations (sodium acetate, 370 mg L−1, K3PO4, 15.6 mg L−1, (NH4)2SO4, 110.1 mg L−1, NaNO2, 7.5 mg L−1, KNO3, 73.4 mg L−1).
2000 µmol m s
-2 -1
-1
Lipids (mg L )
80
60
40
20
0 0
5
10
15
20
Time (days)
b) Dunaliella sp.
120 900 µmol m s
-2 -1
100
1500 µmol m s
-2 -1
2000 µmol m s
Lipids (mg L-1)
-2 -1
80
2.2. Microalgae culture 60
The experiments were carried out in 2-L Erlenmeyer flasks with 1.2 L of SWW, and 100 mL of inoculum of 2.3 g L−1 of pre-conditioned microalgae culture. Cultures were exposed to light with 27 W Grow Led PAR 38 lamps (Grupo Adapta) at 3 different Photosynthetically Active Photon Flux Densities (PAPFD): 900, 1500, and 2000 μmol m−2 s−1, measured at site (at morning, noon, and afternoon) with a photo radiometer Delta Ohm HD 2302.0, and a Delta Ohm Lp 471 PAR probe. 27 W Grow LED PAR38 lightbulbs were used. The light:dark photoperiod was set to 12 h:12 h. The cultures were continuously aerated with Elite P-799 air pumps to provide uniform mixing. Temperature was 30 °C ± 3 °C, measured with a mercury thermometer.
40
20
0 0
5
10
15
20
Time (days)
c) Nannochloropsis sp.
120 900 µmol m s
-2 -1
2.3. Biomass determination
1500 µmol m s
-2 -1
100
Lipids (mg L-1)
-2 -1 2000 µmol m s
Microalgae biomass was quantified daily by filtering 10 mL of the growth medium through a 0.45 μm filter (Millipore), which was preweighted, and dried at 50 °C for 24 h [31].
80
60
2.4. Biovolume determination 40
The biovolume of Tetraselmis sp. and Dunaliella sp. were estimated as 363.7 μm3 and 115.8 μm3, respectively, considering an ellipsoid shape (4/3 π xyz); for Nannochloropsis sp. the biovolume estimate was 10.3 μm3, considering a spherical shape (4/3 π r3). The microalgae linear dimensions were measured by a Nikkon Eclipse 80i phase-contrast microscope.
20
0 0
5
10
15
20
Time (days)
2.5. Determinations in SWW
Fig. 4. Lipid content in the cultures of: a) Tetraselmis sp., b) Dunaliella sp. and c) Nannochloropsis sp. (Data shown is the mean +/−SD, n = 3).
The filtrate obtained from the biomass determination was used for daily physicochemical determinations. Orthophosphate concentration was determined by a colorimetric method in which phosphomolybdate is formed and then reduced to a blue compound [32]. Nitrites were determined by a colorimetric chemical reaction forming an azo dye [32]. Ammonium was determined by oxidation to 100
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
Table 4 Lipid productivities of the 3 microalgae species (Data shown is the mean, n = 3). PAPFD (μmol m−2 s−1) 900 1500 2000
Dunaliella sp. −1
Nannochloropsis sp. −1
−1
16.43 mg L day 19.42 mg L−1 day−1 15.95 mg L−1 day−1
−1
7.34 mg L day 9.11 mg L−1 day−1 13.03 mg L−1 day−1
Tetraselmis sp. 21.90 mg L−1 day−1 29.48 mg L−1 day−1 20.41 mg L−1 day−1
during the log phase, according to [39]. For Nannochloropsis sp. the values were similar at 1500 and 2000 μmol m−2 s−1 PAPFDs (0.18 and 0.17 day−1, respectively), and the lowest value was at 900 μmol m−2 s−1 (0.15 day−1). Values were similar for Dunaliella sp. and Tetraselmis sp., at PAPFDs of 900 and 1500 μmol m−2 s−1 (0.19 and 0.20 day−1, and 0.28 and 0.29 day−1, respectively), and the lowest specific growth rate was obtained at 2000 μmol m−2 s−1 for both microalgae (0.17 and 0.23 day−1). Tetraselmis sp. showed the highest specific growth rates at 900 and 1500 μmol m−2 s−1, 0.28 and 0.29 day−1 respectively, being the species that grows faster, and reaches its death phase in 13 days, while the other two species reached their stationary phase at the same 13 days. Biomass productivity was calculated according to [31] (Table 2). The lowest biomass productivity for all 3 species was obtained at 2000 μmol m−2 s−1 (52.1 mg L−1 day−1 for Dunaliella sp., −1 −1 84.9 mg L day for Nannochloropsis sp., and 101.2 mg L−1 day−1 for Tetraselmis sp.). Dunaliella sp. biomass productivities were very similar at 900 and 1500 μmol m−2 s−1 PAPFDs (68.9 day 69.3 mg L−1 day−1, respectively). The biomass productivity of Nannochloropsis sp., at 900 μmol m−2 s−1 (88.0 mg L−1 day−1) was similar to that obtained at 2000 μmol m−2 s−1 (84.9 mg L−1 day−1), while the highest productivity was obtained at 1500 μmol m−2 s−1 (99.5 mg L−1 day−1). Once again, Tetraselmis sp. showed the highest biomass productivities because of its faster growth rate; the highest productivity was at a PAPFD of 900 μmol m−2 s−1 (132.8 mg L−1 day−1). PAPFD inhibits the microalgae growth as evidenced by the decrease in biomass concentration, as the PAPFD increases. This photoinhibition at high PAPFDs, is due to the reduction of photosynthetic capacity as the proteins required for photosynthesis are degraded [40]. The productivities obtained by us are much lower than those obtained by [41,42]. [41] worked with Nannochloropsis gaditana cultured in water obtained from dried activated sludge, and estimated a productivity of 400 mg L−1 day−1, and [42] worked with Tetraselmis suecica cultivated in wastewater from a fish farm, estimating a productivity of 350 mg L−1 day−1, both authors worked with continuous cultures unlike this study.
nitrites [33]. Nitrates were determined by reduction to nitrites [34]. pH and dissolved oxygen were determined by electrodes [35]. Salinity was determined by an InstantOcean hydrometer. Chemical oxygen demand (COD) was determined by the closed reflux colorimetric method described by [36]. 2.6. Determination of lipid content Biomass lipid content was determined by the colorimetric method of phosphovanillin with a spectrophotometer Pharmacia Biotech, Ultrospec 3000 [37]. 2.7. Statistical analysis All reported statistical analyses were carried out by the R software. An analysis of variance was made to determine if there was inter experimental significant difference, and said differences were determined by a Tukey's test. The confidence interval was of 5% (α = 0.05). 3. Results and analysis 3.1. Microalgae growth Fig. 1 shows the growing curve of Tetraselmis sp., Dunaliella sp., and Nannochloropsis sp., observing different growing rates. Tetraselmis sp. showed the fastest growing rate by reaching the stationary phase in 9 days of culture, in contrast with Dunaliella sp. and Nannochloropsis sp. that reached the stationary phase in 13 days. The stationary phase of Tetraselmis sp. lasted only 3 days, while Nannochloropsis sp. and Dunaliella sp. stationary phases lasted 5 days. Doubling times were calculated according to [38], and are shown in Table 1: It can be seen that for the three microalgae the greatest doubling time was achieved at the PAPFD of 2000 μmol m−2 s−1, and the doubling times at the PAPFDs of 900 and 1500 μmol m−2 s−1 were very similar. The reason for this might be that there is more photosynthetic activity at higher PAPFDs. The dimensions of the microalgae are: Tetraselmis sp. 4.9 ± 0.2 μm depth, 10.5 ± 0.6 μm width, and 13.5 ± 0.5 μm length; Dunaliella sp. 3.7 ± 0.3 μm depth, 6.7 ± 0.3 μm width, and 9.0 ± 0.4 μm length; Nannochloropsis sp. 2.7 ± 0.3 μm diameter. There might be a relationship between biovolume and growing rate since Tetraselmis sp. showed the highest biovolume and reached the stationary phase in the shortest time (8 days), while Nannochloropsis sp., lowest biovolume, took longer to reach the stationary phase (13 days). In terms of biomass production, the highest biomass quantity was obtained at 900 μmol m−2 s−1 PAPFD for the three microalgae. Nannochloropsis sp. showed the highest biomass quantity (1.81 g L−1, on day 15), while Dunaliella sp. and Tetraselmis sp. had lower results (1.37 g L−1 on day 14, and 1.33 L−1 on day 12, respectively). The lowest biomass quantities were obtained at the 2000 μmol m−2 s−1 PAPFD (1.29 g L−1 for Nannochloropsis sp., 1.01 g L−1 for Dunaliella sp., and 1.04 for Tetraselmis sp.). However, these values were obtained at different culture times. Nannochloropsis sp., with the lowest biovolume, showed the highest biomass concentrations, while Dunaliella sp. and Tetraselmis sp. showed similar biomass concentrations. With the obtained data, the specific growth rate was calculated,
3.2. Nutrient removal Fig. 3 shows a decrease in nutrient concentration in the growth medium for Dunaliella sp., Nannochloropsis sp., and Tetraselmis sp., during the whole culture period. Table 3 shows that, in general, a removal higher than 90% for nitrogen and orthophosphate was obtained for all 3 microalgae species. The removal of dissolved organic matter measured as COD was close to 80% for all 3 microalgae species, except for Tetraselmis sp. at the 2000 μmol m−2 s−1 PAPFD, where the removal was 75%. These results are similar to those reported by [43] where a consortia of microalgae and bacteria were cultured in treated wastewater from a fish farm, reaching a COD removal of 77%, with a much higher initial concentration (678 ± 249 mg O2 L−1) than the concentration used in the present study. Total nitrogen and orthophosphate removal were both higher than 90% for all three microalgae species. Removal values for total nitrogen in this study were higher than those obtained in other studies: [42] reported a nitrogen removal of 49.4% with Tetraselmis suecica cultivated in fish farm wastewater (initial concentration of 41.3 mg L−1); 101
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
smallest biovolume (Nannochloropsis sp.) also showed the lowest biomass productivities, while the microalga with the biggest biovolume (Tetraselmis sp.) showed the highest biomass productivities at all 3 PAPFDs. All lipid productivities were significantly different for all 3 microalgae species (p < 0.05). Nitrogen and phosphorus concentrations had a significant effect in the lipid production in all 3 microalgae species (p < 0.05), this is because at the logarithmic growth phase there is sufficient nitrogen and phosphorous in the culture for the microalgae to grow. In late phases of the culture concentrations of nitrogen and phosphorous decrease. Once at the stationary phase where there are low concentrations of both nutrients, the microalgae become stressed and start accumulating lipids as an energy reserve. However, PAPFD had no significant effect in the lipid productivity (p > 0.05), in any of the microalgae species. It is important to direct future research to the study of continuous cultures to determine if the microalgal biomass productivity and lipid content can be increased.
[43] reported 80% of nitrogen removal, with a total initial nitrogen concentration (31 ± 10 mg L−1) lower than the initial nitrogen concentration in this study; [44] reported a 78% removal with N. oculata cultivated in aquaculture wastewater (initial concentration of 40 mg L−1); and [41] reported only 40% of total nitrogen removal due to a nitrogen overconcentration in the tested wastewater (338 mg L−1). Orthophosphates removal values in our results were similar to [41,42], 90% and 99% removal, respectively (with initial concentrations of 25 mg L−1 and 4.96 mg L−1); while [43] reported an orthophosphates removal of 78% (initial concentration of 14 ± 7 mg L−1); and [44] reported 79% of phosphorus removal (initial concentration of 10 mg L−1) with T. chuii; being these removal percentages lower than those reported here. The nitrogen and phosphorus removal kinetics for Tetraselmis sp. were higher than for the other two microalgae species (Table 3), this is explained because a faster growing rate, resulting in a rapid uptake of nutrients. However, Dunaliella sp. and Tetraselmis sp. showed very similar dissolved organic matter removal kinetics, measured as COD, despite their different growing rates. The removal kinetics reported by [41] were much higher (35 mg L−1 day−1 for total nitrogen, and 5.7 mg L−1 day−1 for total phosphorus) than the values reported in this study, since the culture technique was a continuous culture where a volume of medium was extracted daily and replaced with wastewater.
4. Conclusions It has been demonstrated that microalgae are useful in treating mariculture wastewater in view of their high nutrient removal values (i.e., nitrogen, phosphorus, and carbon), especially because these nutrients are commonly found in this type of wastewater. It is possible to enhance the lipid productivities of the tested microalgae by keeping them in the culture wastewater during the stationary phase of the culture, when low-nutrient concentrations are present in the wastewater. This strategy induces stress conditions forcing the microalgae to accumulate lipids as an energy source. Also the photosynthetically active photon flux density has an influence over the biomass and lipid productivities, since at specific PAPFDs the highest biomass and lipid productivities were obtained. This would translate in reducing the wastewater treatment cost because the obtained microalgae biomass byproduct would render a higher biodiesel production.
3.3. Lipid content Fig. 4 shows the lipid production for all 3 microalgae species. In all 3 cases it is evident that once the stationary phase is reached (Fig. 2), the lipid content exponentially increases. At the stationary phase, nutrient concentrations are low, so the microalgae start producing lipids as an energy reserve [45]. Fig. 2 shows that Tetraselmis sp. reached the stationary phase on day 9 of culture. Fig. 4 shows that on day 8, the exponential lipid production starts, precisely at the time where the nutrient concentration is low (Fig. 3). In the case of Dunaliella sp., the stationary phase was reached on day 13 of culture (Fig. 2), when nutrient concentration is low (Fig. 3), and as a result the quantity of lipids increases (Fig. 4). Nannochloropsis sp. showed a similar pattern, reaching the stationary phase on day 14 of culture (Fig. 2) when the nutrient concentration is low (Fig. 3), however, the lipid production starts increasing around day 16 (Fig. 4). Tetraselmis sp. showed the highest lipid concentration, probably due to its bigger size it can store more lipids inside the cell. The microalgae lipid productivity was calculated according to [31] (Table 4), in this case Nannochloropsis sp. showed the lowest lipid productivities, due to its long culture time. Tetraselmis sp. and Dunaliella sp. showed their highest lipid productivities at 1500 μmol m−2 s−1 PAPFD (29.48 and 19.42 mg L−1 day−1, respectively), while Nannochloropsis sp. showed its highest lipid productivity at 2000 μmol m−2 s−1 PAPFD (13.03 mg L−1 day−1). For Dunaliella sp. and Tetraselmis sp. the lipid productivities at 900 and 2000 μmol m−2 s−1 PAPFDs were similar. Tetraselmis sp. showed the highest lipid productivity (29.48 mg L−1 day−1), at 1500 μmol m−2 s−1 PAPFD. The productivities observed for Nannochloropsis sp. are lower than those reported by [46], i.e. 12.9 mg L−1 day−1 for Nannochloropsis salina, and 18.0 mg L−1 day−1 for Nannochloropsis gaditana. The lipid productivities obtained for Tetraselmis sp. are higher than those reported by [47] for Tetraselmis suecica in a nitrogen-limited medium (18.7 mg L−1 day−1), but lower than the value reported for this microalga in a growth medium (34.6 mg L−1 day−1). Dunaliella sp. productivities were lower than the values reported by [48], which were 38.0 mg L−1 day−1 and 56.0 mg L−1 day−1 in a nitrogen-limited medium and a nutrient-sufficient medium, respectively. Tetraselmis sp. showed the best results with the highest biomass productivity at 900 μmol m−2 s−1 (132.8 mg L−1 day−1). However, the highest lipid productivity (29.5 mg L−1 day−1) was observed at 1500 μmol m−2 s−1. Again, it is noticeable that the microalga with the
Conflict of interest The authors declare that there is no conflict of interest. No conflicts, informed consent, human or animal rights are applicable for this work. Acknowledgements We thanks for the economic support provided by Project UNAM/ DGAPA PAPIIT IT202818 “Desarrollo de una estrategia para el tratamiento de aguas residuales de maricultura empleando microalgas y aprovechamiento de la biomasa producida” and PAIP (VMLP) 50009111 granted to VMLP by the Facultad de Química, UNAM. CONACYT (421332), Mexico grant received by Manuel Sacristán for doctoral studies (Ph.D. studies) on Engineering in the PMyD at UNAM. Iveth Gabriela Palomino Albarrán, MSc, is kindly acknowledged for the donation of the microalgae cultures. Korinthya López Aguiar, MSc, is kindly acknowledged for the technical assistance in the wastewater analysis. Luciano Hernández Gómez, MSc, for his technical support. References [1] FAO, The State of World Fisheries and Aquaculture. Contributing to Food Security and Nutrition for All, (2016). [2] N.M. Nasir, N.S.A. Bakar, F. Lananan, S.H. Abdul Hamid, S.S. Lam, A. Jusoh, Treatment of African catfish, Clarias gariepinus wastewater utilizing phytoremediation of microalgae, Chlorella sp. with Aspergillus niger bio-harvesting, Bioresour. Technol. (2015), https://doi.org/10.1016/j.biortech.2015.03.023 (in press). [3] S.A. Castine, A.D. McKinnon, N.A. Paul, L.A. Trott, R. de Nys, Wastewater treatment for land-based aquaculture: improvements and value-adding alternatives in model systems from Australia, Aquac. Environ. Interact. 4 (2013) 285–300, https://doi. org/10.3354/aei00088. [4] S. Dinesh Kumar, P. Santhanam, R. Nandakumar, S. Ananth, P. Nithya, B. Dhanalakshmi, et al., Bioremediation of shrimp (Litopenaeus vannamei) cultured effluent using copepod (Oithona rigida) and microalgae (Picochlorum maculatam &
102
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
[5]
[6] [7]
[8]
[9]
[10]
[11]
[12]
[13]
[14] [15]
[16]
[17]
[18]
[19]
[20]
[21]
[22]
[23]
[24]
[25]
10.1016/j.biotechadv.2016.06.004. [26] P. Singh, S. Kumari, A. Guldhe, R. Misra, I. Rawat, F. Bux, Trends and novel strategies for enhancing lipid accumulation and quality in microalgae, Renew. Sust. Energ. Rev. 55 (2016) 1–16, https://doi.org/10.1016/j.rser.2015.11.001. [27] S. Esakkimuthu, V. Krishnamurthy, R. Govindarajan, K. Swaminathan, Augmentation and starvation of calcium, magnesium, phosphate on lipid production of Scenedesmus obliquus, Biomass Bioenergy 88 (2016) 126–134, https://doi. org/10.1016/j.biombioe.2016.03.019. [28] Y. Gong, M. Jiang, Biodiesel production with microalgae as feedstock: from strains to biodiesel, Biotechnol. Lett. 33 (2011) 1269–1284, https://doi.org/10.1007/ s10529-011-0574-z. [29] A. Demirbas, M. Fatih Demirbas, Importance of algae oil as a source of biodiesel, Energy Convers. Manag. 52 (2011) 163–170, https://doi.org/10.1016/j.enconman. 2010.06.055. [30] R. Huerlimann, R. de Nys, K. Heimann, Growth, lipid content, productivity, and fatty acid composition of tropical microalgae for scale-up production, Biotechnol. Bioeng. 107 (2010) 245–257, https://doi.org/10.1002/bit.22809. [31] A. Toledo-Cervantes, M. Morales, E. Novelo, S. Revah, Carbon dioxide fixation and lipid storage by Scenedesmus obtusiusculus, Bioresour. Technol. 130 (2013) 652–658, https://doi.org/10.1016/j.biortech.2012.12.081. [32] K. Grasshoff, K. Kremling, M. Ehrhardt, Methods of Seawater Analysis, third edition, Wiley-VCH, Germany, 1999. [33] F.A. Richards, R.A. Kletsch, The Spectophotometric Determination of Ammonia and Labile Amino Compounds in Fresh and Seawater by Oxidation to Nitrite, (1964). [34] B. Schnetger, C. Lehners, Determination of nitrate plus nitrite in small volume marine water samples using vanadium(III)chloride as a reduction agent, Mar. Chem. 160 (2014) 91–98, https://doi.org/10.1016/j.marchem.2014.01.010. [35] APHA, AWWA, WEF, Standard Methods for the Examination of Water and Wastewater, 20th ed., American Public Health Association, American Water Works Association, Water Environment Federation, Washington D.C., 1998. [36] I. Vyrides, D.C. Stuckey, A modified method for the determination of chemical oxygen demand (COD) for samples with high salinity and low organics, Bioresour. Technol. 100 (2009) 979–982, https://doi.org/10.1016/j.biortech.2008.06.038. [37] S.K. Mishra, W.I. Suh, W. Farooq, M. Moon, A. Shrivastav, M.S. Park, et al., Rapid quantification of microalgal lipids in aqueous medium by a simple colorimetric method, Bioresour. Technol. 155 (2014) 330–333, https://doi.org/10.1016/j. biortech.2013.12.077. [38] R.S. Gour, A. Kant, R.S. Chauhan, Screening of micro algae for growth and lipid accumulation properties, J. Algal Biomass Util. 5 (2014) 38–46. [39] FAO, Manual on the Production and Use of Live Food for Aquaculture, (1996), https://doi.org/10.1007/s13398-014-0173-7.2. [40] D. Hanelt, Photoinhibition of photosynthesis in marine microalgae, Sci. Mar. 60 (1996) 243–248. [41] C. Sepúlveda, F.G. Acién, C. Gómez, N. Jiménez-Ruíz, C. Riquelme, E. MolinaGrima, Utilization of centrate for the production of the marine microalgae Nannochloropsis gaditana, Algal Res. 9 (2015) 107–116, https://doi.org/10.1016/j. algal.2015.03.004. [42] M.H.A. Michels, M. Vaskoska, M.H. Vermuë, R.H. Wiffels, Growth of Tetraselmis suecica in a tubular photobioreactor on wastewater from a fish farm, Water Res. 65 (2014) 290–296, https://doi.org/10.1016/j.watres.2014.07.017. [43] E. Posadas, A. Muñoz, M.-C. García-González, R. Muñoz, P.A. García-Encina, A case study of a pilot high rate algal pond for the treatment of fish farm and domestic wastewaters, J. Chem. Technol. Biotechnol. (2014), https://doi.org/10.1002/jctb. 4417 (n/a-n/a). [44] I.N. Sirakov, K.N. Velichkova, Bioremediation of wastewater originate from aquaculture and biomass production from microalgae species - Nannochloropsis oculata and Tetraselmis chuii, Bulgarian J. Agr. Sci. 20 (2014) 66–72. [45] I.A. Guschina, J.L. Harwood, Algal lipids and effect of the environment on their biochemistry in lipids in aquatics, Ecosystems (2009), https://doi.org/10.1017/ CBO9781107415324.004. [46] A. Taleb, J. Pruvost, J. Legrand, H. Marec, B. Le-Gouic, B. Mirabella, et al., Development and validation of a screening procedure of microalgae for biodiesel production: application to the genus of marine microalgae Nannochloropsis, Bioresour. Technol. 177 (2015) 224–232, https://doi.org/10.1016/j.biortech.2014. 11.068. [47] E. Del Río, A. Armendáriz, E. García-Gómez, M. García-González, M.G. Guerrero, Continuous culture methodology for the screening of microalgae for oil, J. Biotechnol. 195 (2015) 103–107, https://doi.org/10.1016/j.jbiotec.2014.12.024. [48] G. Breuer, P.P. Lamers, D.E. Martens, R.B. Draaisma, R.H. Wijffels, The impact of nitrogen starvation on the dynamics of triacylglycerol accumulation in nine microalgae strains, Bioresour. Technol. 124 (2012) 217–226, https://doi.org/10. 1016/j.biortech.2012.08.003.
Amphora sp.)—an integrated approach, Desalin. Water Treat. (2016) 1–10, https:// doi.org/10.1080/19443994.2016.1163509. F. Gao, C. Li, Z. Yang, G. Zeng, L. Feng, J. Liu, et al., Continuous microalgae cultivation in aquaculture wastewater by a membrane photobioreactor for biomass production and nutrients removal, Ecol. Eng. 92 (2016) 55–61, https://doi.org/10. 1016/j.ecoleng.2016.03.046. D.G. Kim, Y.-E. Choi, Microalgae cultivation using LED light, Korean Chem. Eng. Res. 52 (2014) 8–16, https://doi.org/10.9713/kcer.2014.52.1.8. F. Bumbak, S. Cook, V. Zachleder, S. Hauser, K. Kovar, Best practices in heterotrophic high-cell-density microalgal processes: achievements, potential and possible limitations, Appl. Microbiol. Biotechnol. 91 (2011) 31–46, https://doi.org/10. 1007/s00253-011-3311-6. J. Wang, H. Yang, F. Wang, Mixotrophic cultivation of microalgae for biodiesel production: status and prospects, Appl. Biochem. Biotechnol. 172 (2014) 3307–3329, https://doi.org/10.1007/s12010-014-0729-1. W.-W. Li, H.-Q. Yu, Z. He, Towards sustainable wastewater treatment by using microbial fuel cells-centered technologies, Energy Environ. Sci. 7 (2014) 911–924, https://doi.org/10.1039/c3ee43106a. K.K. Sharma, H. Schuhmann, P.M. Schenk, High lipid induction in microalgae for biodiesel production, Energies 5 (2012) 1532–1553, https://doi.org/10.3390/ en5051532. A.K. Minhas, P. Hodgson, C.J. Barrow, A. Adholeya, A review on the assessment of stress conditions for simultaneous production of microalgal lipids and carotenoids, Front. Microbiol. 7 (2016) 1–19, https://doi.org/10.3389/fmicb.2016.00546. L. Xin, H. Hu, G. Ke, Y. Sun, Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid accumulation of a freshwater microalga Scenedesmus sp. Bioresour. Technol. 101 (2010) 5494–5500, https://doi. org/10.1016/j.biortech.2010.02.016. J.-V. Pérez-Pazos, P. Fernández-Izquierdo, Synthesis of neutral lipids in Chlorella sp. under different light and carbonate conditions, Ciencia Tecnol. Futur. 4 (2011) 47–58. L.D. Zhu, Z.H. Li, E. Hiltunen, Strategies for lipid production improvement in microalgae as a biodiesel feedstock, Biomed. Res. Int. 2016 (2016) 1–8. Q. Hu, M. Sommerfeld, E. Jarvis, M. Ghirardi, M. Posewitz, M. Seibert, et al., Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances, Plant J. 54 (2008) 621–639, https://doi.org/10.1111/j.1365-313X.2008. 03492.x. O. Perez-Garcia, F.M.E. Escalante, L.E. De-Bashan, Y. Bashan, Heterotrophic cultures of microalgae: metabolism and potential products, Water Res. 45 (2011) 11–36, https://doi.org/10.1016/j.watres.2010.08.037. A.M.J. Kliphuis, A.J. Klok, D.E. Martens, P.P. Lamers, M. Janssen, R.H. Wijffels, Metabolic modeling of Chlamydomonas reinhardtii: energy requirements for photoautotrophic growth and maintenance, J. Appl. Phycol. 24 (2012) 253–266, https:// doi.org/10.1007/s10811-011-9674-3. Y. Li, X. Fei, X. Deng, Novel molecular insights into nitrogen starvation-induced triacylglycerols accumulation revealed by differential gene expression analysis in green algae Micractinium pusillum, Biomass Bioenergy 42 (2012) 199–211, https:// doi.org/10.1016/j.biombioe.2012.03.010. A. Patel, S. Barrington, M. Lefsrud, Microalgae for phosphorus removal and biomass production: a six species screen for dual-purpose organisms, GCB Bioenergy 4 (2012) 485–495, https://doi.org/10.1111/j.1757-1707.2012.01159.x. J. Msanne, D. Xu, A. Reddy, J.A. Casas-Mollano, T. Awada, E.B. Cahoon, et al., Metabolic and gene expression changes triggered by nitrogen deprivation in the photoautotrophically grown microalgae Chlamydomonas reinhardtii and Coccomyxa sp. C-169, Phytochemistry 75 (2012) 50–59, https://doi.org/10.1016/j.phytochem. 2011.12.007. L.A. Garay, K.L. Boundy-Mills, J.B. German, Accumulation of high-value lipids in single-cell microorganisms: a mechanistic approach and future perspectives, J. Agric. Food Chem. 62 (2014) 2709–2727, https://doi.org/10.1021/jf4042134. J.J. Mayers, K.J. Flynn, R.J. Shields, Influence of the N:P supply ratio on biomass productivity and time-resolved changes in elemental and bulk biochemical composition of Nannochloropsis sp. Bioresour. Technol. 169 (2014) 588–595, https:// doi.org/10.1016/j.biortech.2014.07.048. H.J. Choi, S.M. Lee, Effect of the N/P ratio on biomass productivity and nutrient removal from municipal wastewater, Bioprocess Biosyst. Eng. 38 (2015) 761–766, https://doi.org/10.1007/s00449-014-1317-z. N.W. Rasdi, J.G. Qin, Effect of N:P ratio on growth and chemical composition of Nannochloropsis oculata and Tisochrysis lutea, J. Appl. Phycol. 27 (2015) 2221–2230, https://doi.org/10.1007/s10811-014-0495-z. S.K. Lenka, N. Carbonaro, R. Park, S.M. Miller, I. Thorpe, Y. Li, Current advances in molecular, biochemical, and computational modeling analysis of microalgal triacylglycerol biosynthesis, Biotechnol. Adv. 34 (2016) 1046–1063, https://doi.org/
103
Contents lists available at ScienceDirect
Algal Research journal homepage: www.elsevier.com/locate/algal
Carbon, nitrogen, and phosphorus removal, and lipid production by three saline microalgae grown in synthetic wastewater irradiated with different photon fluxes
T
⁎
Manuel Sacristán de Alvaa, Víctor Manuel Luna Pabelloa, , María Teresa Orta Ledesmab, Modesto Javier Cruz Gómezc a Laboratorio de Microbiología Experimental, Departamento de Biología, Facultad de Química, UNAM, Av. Universidad No. 3000, Col. Universidad, Nacional Autónoma de México C.U., Delegación Coyoacán, C.P. 04510 Ciudad de México, México, Mexico b Laboratorio de Ingeniería Ambiental, Edificio 5, Instituto de Ingeniería, UNAM, Mexico c Laboratorio 212, Departamento de Ingeniería Química, Conjunto E. Facultad de Química, UNAM, Mexico
A R T I C LE I N FO
A B S T R A C T
Keywords: Mariculture Lipids Biodiesel Nutrient stress Marine microalgae
Mariculture production has increased in the last decades, with untreated wastewater discharged directly into the sea, impacting coastal ecosystems. There is a need for mariculture wastewater treatment systems that are costeffective. This can be met by the implementation of wastewater treatment systems that in addition to removing pollutants are capable of producing valuable by-products such as biomass for the biofuel industry. In this study, Dunaliella sp., Nannochloropsis sp. and Tetraselmis sp. microalgae were cultivated in controlled environments simulating mariculture wastewaters. Single stage culture systems were used to grow these microalgae, the growing conditions included inducing stress with different photon flux densities (900, 1500 and 2000 μmol m−2 s−1), and low carbon, nitrogen and phosphorus concentrations obtained at the stationary phase, in order to force these microalgae to increase their lipid content. The best results were obtained with Tetraselmis sp., which achieved 132.8 mg L−1 day−1 of biomass productivity at 900 μmol m−2 s−1. Nevertheless the best lipid productivity was reached at 1500 μmol m−2 s−1, also by Tetraselmis sp., being 29.5 mg L−1 day−1, where biomass productivity was of 124.5 mg L−1 day−1. All three microalgae species were able to remove > 90% of nitrogen and orthophosphates, and 80% of carbon, which makes them suitable for treating mariculture wastewater, and in addition, represent a valuable high lipid content biomass byproduct usable as raw material for biodiesel synthesis.
1. Introduction Aquaculture production has increased substantially for the past 40 years [1,2]. This industry is responsible of producing wastewaters with high contents of suspended solids and dissolved nutrients, which are discharged untreated into water bodies, causing eutrophication [3], generating damages to coastal ecosystems, and eventually the loss of biodiversity [4]. Conventional biological and physicochemical wastewater treatments can remove nutrients contained in this kind of wastewaters, and produce a good quality effluent. However, these methods involve the production of sludges which in turn must be treated prior to discharge; generating additional costs. In this regard, microalgae culture in aquaculture wastewaters offers the advantages of removing nutrients
while simultaneously producing biomass that may be used as raw material for valuable products, such as biofuels [5]. The economic feasibility of microalgae biofuels faces various challenging limiting factors, such as the need for cultures with high lipid productivity in the shortest time period [6,7]. It is well-known that cell growth and lipid accumulation do not take place simultaneously during culture, so the techniques for increasing lipid accumulation often render low biomass quantities, and consequently, low lipid productivity [8]. To address this issue, the microalgae culture is carried out in two stages. In the first stage, microalgae are cultured in optimum growth conditions; and in the second stage, the cultures are subject to stress conditions in order to enhance the accumulation of energy in the form of neutral lipids (triacylglycerols) by altering their biosynthetic pathways. The result is no increase in microalgae density, but an important
⁎
Corresponding author. E-mail addresses: [email protected] (M. Sacristán de Alva), [email protected] (V.M. Luna Pabello), [email protected] (M.T. Orta Ledesma), [email protected] (M.J. Cruz Gómez). https://doi.org/10.1016/j.algal.2018.07.006 Received 22 February 2018; Received in revised form 26 June 2018; Accepted 8 July 2018 2211-9264/ © 2018 Elsevier B.V. All rights reserved.
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
Fig. 1. Conceptual model of lipid synthesis during nitrogen limitation and light excess in a microalgae.
increase in lipid content [9,10]. There are several physicochemical factors that affect microalgae metabolism, for enhancing lipid production, such as light intensity, photoperiod, temperature, salinity, nutrient concentration in the growth medium (carbon, phosphates, nitrogen), and nitrogen (ammonium or nitrate) and carbon (glucose or glycerol, among others) sources. By exposing microalgae to environmental stress in laboratory scale cultures, through the modification of said factors, high lipid productivity has been achieved [11]. Nitrogen limitation causes three main changes: decrease in the thylakoid membrane; stimulation of phospholipid hydrolysis; and fatty acid synthesis enzyme activation. These changes cause an increase of the intracellular fatty acids content [12]. Generally, when microalgae are grown under low light intensities, the assimilated carbon is used for the synthesis of amino acids and other essential components, but under light saturation conditions sugars, lipids, and starch are formed [13]. An adequate light intensity contributes to lipid overproduction, which may be the result of the overgeneration of photo-assimilated compounds that are converted into chemical energy [14]. Fig. 1 shows a conceptual model of lipid accumulation in microalgae. During the nitrogen limitation stage, the pathways marked with dashed arrows are activated for lipid production; other pathways are the membrane lipid recycling, and the protein degradation. Pathways indicated with X are inhibited mostly because there is a reduction in the photosynthetic rate, and carbon fixation through photosynthesis is reduced, as well as protein synthesis. When there is light excess the
Table 1 Doubling times of the 3 microalgae species (Data shown is the mean, n = 3). Microalgae
Tetraselmis sp. Dunaliella sp. Nannochloropsis sp.
Photosynthetically active photon flux density (PAPFD) (μmol m−2 s−1) 900
1500
2000
3.21 4.13 5.83
3.56 4.13 5.27
4.23 4.80 6.67
Table 2 Specific growth rate and biomass productivity of the 3 microalgae species (Data shown is the mean, n = 3). Microalgae
PAPFD (μmol m−2 s−1)
900
1500
2000
Dunaliella sp.
Specific growth rate (day−1) Biomass productivity (mg L−1 day−1) Specific growth rate (day−1) Biomass productivity (mg L−1 day−1) Specific growth rate (day−1) Biomass productivity (mg L−1 day−1)
0.19 68.9
0.20 69.3
0.17 52.1
0.15 88.0
0.18 99.5
0.17 84.9
0.29 132.8
0.28 124.5
0.23 101.2
Nannochloropsis sp.
Tetraselmis sp.
Table 3 Nutrient removal kinetics and removal percentages of the 3 microalgae species (Data shown is the mean, n = 3). PAPFD (μmol m−2 s−1)
900
1500
2000
a
Total nitrogena Orthophosphates COD Total nitrogena Orthophosphates COD Total nitrogena Orthophosphates COD
Dunaliella sp.
Nannochloropsis sp.
Tetraselmis sp.
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
Removal percentage (%)
Removal kinetics (mg L−1 day−1)
96.91 94.02 81.85 97.26 94.25 82.38 97.53 93.27 81.29
3.50 0.87 17.35 3.57 0.94 17.99 3.51 0.89 16.71
94.16 97.12 81.45 90.07 97.16 81.02 94.24 97.20 81.85
3.85 0.78 10.45 3.68 0.81 10.17 4.02 0.83 10.74
97.99 95.62 81.56 98.03 95.94 80.97 98.06 95.67 75.60
7.07 1.49 20.11 7.25 1.48 19.34 7.34 1.48 14.08
Total nitrogen: is the sum of the concentrations of nitrates, nitrites and ammonia. 98
Algal Research 34 (2018) 97–103
900 mol m s
1.6
1500 mol m s
-2 -1
-2 -1
2000 mol m s
1.4
250 -1
1.2 1.0 0.8 0.6 0.4
20
Nitrogen 900 µmol m-2s-1 COD 900 µmol m-2s-1 Orthophosphates 900 µmol m-2s-1 Nitrogen 1500 µmol m-2s-1 COD 1500 µmol m-2s-1 Orthophosphates 1500 µmol m-2s-1 Orthophosphates 2000 µmol m-2s-1 Nitrogen 2000 µmol m-2s-1 COD 2000 µmol m-2s-1
300
-2 -1
Total nitrogen (mg L ) COD (mg O2 L-1)
Dry biomass (g L-1)
1.8
Dunaliella sp.
350
200 150
18 16 14 12 10 8
100
6 50
0.2
4
0
0.0 0
5
10
15
-1
a) Tetraselmis sp.
2.0
Orthophosphates (mg L )
M. Sacristán de Alva et al.
2
20
0 0
Time (days)
2
4
6
8
10
12
14
16
18
20
Time (days) -1
Growth kinetics: k 900 =0.22 day , k 1500 =0.19 day , k 2000 =0.16 day 2.0 1.8
300
-2 -1
900 mol m s
250
-2 -1
1500 mol m s
1.6
-2 -1
2000 mol m s
Total nitrogen (mg L ) COD (mg O2 L-1)
1.4
-1
1.2 1.0 0.8 0.6
200
20 18 16 14 12
150
10 8
100
6 50
4
0.4
2
0
0.2
0
0.0 5
10
15
-50
20
0
5
10
Time (days)
Growth kinetics: k 900 =0.17 day-1, k 1500 =0.17 day-1, k 2000 =0.14 day-1
-2 -1
Dry biomass (g L-1)
Total nitrogen (mg L ) COD (mg O2 L-1)
1500 mol m s
1.6
-1
-2 -1
2000 mol m s
1.4 1.2 1.0 0.8 0.6 0.4
20
Nitrogen 900 µ mol m-2s-1 COD 900 µ mol m-2s-1 Orthophosphates 900 µ mol m-2s-1 -2 -1 Nitrogen 1500 µ mol m s
250
-2 -1
900 mol m s
1.8
20
Tetraselmis sp.
300
c) Nannochloropsis sp.
2.0
15
Time (days)
200
18 16 14
-2 -1
COD 1500 µ mol m s
12
Orthophosphates 1500 µ mol m-2s-1 Orthophosphates 2000 µ mol m-2s-1 -2 -1 Nitrogen 2000 µ mol m s
150 100
10 8
-2 -1
COD 2000 µ mol m s
6 50
4 2
0
0
0.2 0.0 0
5
10
15
0
20
-1
0
Orthophosphates (mg L )
Dry biomass (g L-1)
Nitrogen 900 µmol m-2s-1 COD 900 µmol m-2s-1 Orthophosphates 900 µmol m-2s-1 Orthophosphates 1500 µmol m-2s-1 Nitrogen 1500 µmol m-2s-1 COD 1500 µmol m-2s-1 Nitrogen 2000 µmol m-2s-1 COD 2000 µmol m-2s-1 Orthophosphates 2000 µmol m-2s-1
Nannochloropsis sp.
b) Dunaliella sp.
-1
-1
Orthophosphates (mg L )
-1
5
10
15
20
Time (days)
Time (days)
Growth kinetics: k 900 =0.12 day-1, k 1500 =0.13 day-1, k 2000 =0.10 day-1
Fig. 3. Nutrient concentration during the culture of the 3 microalgae species (Data shown is the mean +/−SD, n = 3).
Fig. 2. Growth curves of: a) Tetraselmis sp., b) Dunaliella sp. and c) Nannochloropsis sp. (Data shown is the mean +/−SD, n = 3).
is required [27]. In view of the foregoing, the purpose of this study is separately growing three marine microalgae in a single stage culture to overcome the difficulties of a two stage culture. Firstly, we seek to obtain the maximum microalgal biomass productivity during the logarithmic growing phase where the nutrients are present. Secondly obtaining high lipid accumulation as a result of the stress produced by the lack of nutrients when reaching the stationary phase. In order to determine the effect on the productivity of lipids in microalgae, three levels of photosynthetically active photon flux density were evaluated.
pathways marked with grey arrows get activated for lipid production, carotenoid synthesis pathways are also activated since carotenoids act as a sunblock for the cells. Adapted from [14–26]. Two-stage culture has the disadvantage of complicating the culture process by requiring additional infrastructure for transferring the culture from the first stage to the second, adding costs to the process. Furthermore, even if lipid accumulation increases, the growth rate is reduced, affecting the net lipid productivity. Consequently, a strategy that stimulates lipid accumulation without affecting microalgae growth
99
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
a) Tetraselmis sp.
120
2. Materials and methods 2.1. Microalgae and growth medium
900 µmol m s
-2 -1
1500 µmol m s
-2 -1
100
The microalgae used in this study were Dunaliella sp., Nannochloropsis, and Tetraselmis sp., because they are used as live food for mariculture organisms, and because of their lipid content assessed in other studies [28–30]. The synthetic wastewater (SWW) used in this study was prepared based in the characterization made of actual wastewater (AWW) effluents from growing commercial marine species, such as octopus, shrimp, seabass, corvina, snapper, and ornamental species, collected at the facilities of the Yucatan Academic Unit of the UNAM located in Sisal, Yucatan, Mexico. The values obtained for the AWW were the following: nitrate, 45 mg L−1; nitrite, 10 mg L−1; ammonium, 30 mg L−1; orthophosphate, 17 mg L−1; and dissolved organic matter (as chemical oxygen demand, COD), 270 mg O2 L−1. SWW was prepared from distilled water added with a sufficient amount of Instant Ocean sea salt to obtain the desired salinity (31 psu for Nannochloropsis sp. and Tetraselmis sp., and 41 psu for Dunaliella sp.; these salinities provide optimum growth for each species. In order to match the nitrogen, phosphorus, and carbon concentrations, of the AWW, KNO3, NaNO2, (NH4)2SO4, K3PO4, and sodium acetate were added in sufficient quantities to mimic AWW concentrations (sodium acetate, 370 mg L−1, K3PO4, 15.6 mg L−1, (NH4)2SO4, 110.1 mg L−1, NaNO2, 7.5 mg L−1, KNO3, 73.4 mg L−1).
2000 µmol m s
-2 -1
-1
Lipids (mg L )
80
60
40
20
0 0
5
10
15
20
Time (days)
b) Dunaliella sp.
120 900 µmol m s
-2 -1
100
1500 µmol m s
-2 -1
2000 µmol m s
Lipids (mg L-1)
-2 -1
80
2.2. Microalgae culture 60
The experiments were carried out in 2-L Erlenmeyer flasks with 1.2 L of SWW, and 100 mL of inoculum of 2.3 g L−1 of pre-conditioned microalgae culture. Cultures were exposed to light with 27 W Grow Led PAR 38 lamps (Grupo Adapta) at 3 different Photosynthetically Active Photon Flux Densities (PAPFD): 900, 1500, and 2000 μmol m−2 s−1, measured at site (at morning, noon, and afternoon) with a photo radiometer Delta Ohm HD 2302.0, and a Delta Ohm Lp 471 PAR probe. 27 W Grow LED PAR38 lightbulbs were used. The light:dark photoperiod was set to 12 h:12 h. The cultures were continuously aerated with Elite P-799 air pumps to provide uniform mixing. Temperature was 30 °C ± 3 °C, measured with a mercury thermometer.
40
20
0 0
5
10
15
20
Time (days)
c) Nannochloropsis sp.
120 900 µmol m s
-2 -1
2.3. Biomass determination
1500 µmol m s
-2 -1
100
Lipids (mg L-1)
-2 -1 2000 µmol m s
Microalgae biomass was quantified daily by filtering 10 mL of the growth medium through a 0.45 μm filter (Millipore), which was preweighted, and dried at 50 °C for 24 h [31].
80
60
2.4. Biovolume determination 40
The biovolume of Tetraselmis sp. and Dunaliella sp. were estimated as 363.7 μm3 and 115.8 μm3, respectively, considering an ellipsoid shape (4/3 π xyz); for Nannochloropsis sp. the biovolume estimate was 10.3 μm3, considering a spherical shape (4/3 π r3). The microalgae linear dimensions were measured by a Nikkon Eclipse 80i phase-contrast microscope.
20
0 0
5
10
15
20
Time (days)
2.5. Determinations in SWW
Fig. 4. Lipid content in the cultures of: a) Tetraselmis sp., b) Dunaliella sp. and c) Nannochloropsis sp. (Data shown is the mean +/−SD, n = 3).
The filtrate obtained from the biomass determination was used for daily physicochemical determinations. Orthophosphate concentration was determined by a colorimetric method in which phosphomolybdate is formed and then reduced to a blue compound [32]. Nitrites were determined by a colorimetric chemical reaction forming an azo dye [32]. Ammonium was determined by oxidation to 100
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
Table 4 Lipid productivities of the 3 microalgae species (Data shown is the mean, n = 3). PAPFD (μmol m−2 s−1) 900 1500 2000
Dunaliella sp. −1
Nannochloropsis sp. −1
−1
16.43 mg L day 19.42 mg L−1 day−1 15.95 mg L−1 day−1
−1
7.34 mg L day 9.11 mg L−1 day−1 13.03 mg L−1 day−1
Tetraselmis sp. 21.90 mg L−1 day−1 29.48 mg L−1 day−1 20.41 mg L−1 day−1
during the log phase, according to [39]. For Nannochloropsis sp. the values were similar at 1500 and 2000 μmol m−2 s−1 PAPFDs (0.18 and 0.17 day−1, respectively), and the lowest value was at 900 μmol m−2 s−1 (0.15 day−1). Values were similar for Dunaliella sp. and Tetraselmis sp., at PAPFDs of 900 and 1500 μmol m−2 s−1 (0.19 and 0.20 day−1, and 0.28 and 0.29 day−1, respectively), and the lowest specific growth rate was obtained at 2000 μmol m−2 s−1 for both microalgae (0.17 and 0.23 day−1). Tetraselmis sp. showed the highest specific growth rates at 900 and 1500 μmol m−2 s−1, 0.28 and 0.29 day−1 respectively, being the species that grows faster, and reaches its death phase in 13 days, while the other two species reached their stationary phase at the same 13 days. Biomass productivity was calculated according to [31] (Table 2). The lowest biomass productivity for all 3 species was obtained at 2000 μmol m−2 s−1 (52.1 mg L−1 day−1 for Dunaliella sp., −1 −1 84.9 mg L day for Nannochloropsis sp., and 101.2 mg L−1 day−1 for Tetraselmis sp.). Dunaliella sp. biomass productivities were very similar at 900 and 1500 μmol m−2 s−1 PAPFDs (68.9 day 69.3 mg L−1 day−1, respectively). The biomass productivity of Nannochloropsis sp., at 900 μmol m−2 s−1 (88.0 mg L−1 day−1) was similar to that obtained at 2000 μmol m−2 s−1 (84.9 mg L−1 day−1), while the highest productivity was obtained at 1500 μmol m−2 s−1 (99.5 mg L−1 day−1). Once again, Tetraselmis sp. showed the highest biomass productivities because of its faster growth rate; the highest productivity was at a PAPFD of 900 μmol m−2 s−1 (132.8 mg L−1 day−1). PAPFD inhibits the microalgae growth as evidenced by the decrease in biomass concentration, as the PAPFD increases. This photoinhibition at high PAPFDs, is due to the reduction of photosynthetic capacity as the proteins required for photosynthesis are degraded [40]. The productivities obtained by us are much lower than those obtained by [41,42]. [41] worked with Nannochloropsis gaditana cultured in water obtained from dried activated sludge, and estimated a productivity of 400 mg L−1 day−1, and [42] worked with Tetraselmis suecica cultivated in wastewater from a fish farm, estimating a productivity of 350 mg L−1 day−1, both authors worked with continuous cultures unlike this study.
nitrites [33]. Nitrates were determined by reduction to nitrites [34]. pH and dissolved oxygen were determined by electrodes [35]. Salinity was determined by an InstantOcean hydrometer. Chemical oxygen demand (COD) was determined by the closed reflux colorimetric method described by [36]. 2.6. Determination of lipid content Biomass lipid content was determined by the colorimetric method of phosphovanillin with a spectrophotometer Pharmacia Biotech, Ultrospec 3000 [37]. 2.7. Statistical analysis All reported statistical analyses were carried out by the R software. An analysis of variance was made to determine if there was inter experimental significant difference, and said differences were determined by a Tukey's test. The confidence interval was of 5% (α = 0.05). 3. Results and analysis 3.1. Microalgae growth Fig. 1 shows the growing curve of Tetraselmis sp., Dunaliella sp., and Nannochloropsis sp., observing different growing rates. Tetraselmis sp. showed the fastest growing rate by reaching the stationary phase in 9 days of culture, in contrast with Dunaliella sp. and Nannochloropsis sp. that reached the stationary phase in 13 days. The stationary phase of Tetraselmis sp. lasted only 3 days, while Nannochloropsis sp. and Dunaliella sp. stationary phases lasted 5 days. Doubling times were calculated according to [38], and are shown in Table 1: It can be seen that for the three microalgae the greatest doubling time was achieved at the PAPFD of 2000 μmol m−2 s−1, and the doubling times at the PAPFDs of 900 and 1500 μmol m−2 s−1 were very similar. The reason for this might be that there is more photosynthetic activity at higher PAPFDs. The dimensions of the microalgae are: Tetraselmis sp. 4.9 ± 0.2 μm depth, 10.5 ± 0.6 μm width, and 13.5 ± 0.5 μm length; Dunaliella sp. 3.7 ± 0.3 μm depth, 6.7 ± 0.3 μm width, and 9.0 ± 0.4 μm length; Nannochloropsis sp. 2.7 ± 0.3 μm diameter. There might be a relationship between biovolume and growing rate since Tetraselmis sp. showed the highest biovolume and reached the stationary phase in the shortest time (8 days), while Nannochloropsis sp., lowest biovolume, took longer to reach the stationary phase (13 days). In terms of biomass production, the highest biomass quantity was obtained at 900 μmol m−2 s−1 PAPFD for the three microalgae. Nannochloropsis sp. showed the highest biomass quantity (1.81 g L−1, on day 15), while Dunaliella sp. and Tetraselmis sp. had lower results (1.37 g L−1 on day 14, and 1.33 L−1 on day 12, respectively). The lowest biomass quantities were obtained at the 2000 μmol m−2 s−1 PAPFD (1.29 g L−1 for Nannochloropsis sp., 1.01 g L−1 for Dunaliella sp., and 1.04 for Tetraselmis sp.). However, these values were obtained at different culture times. Nannochloropsis sp., with the lowest biovolume, showed the highest biomass concentrations, while Dunaliella sp. and Tetraselmis sp. showed similar biomass concentrations. With the obtained data, the specific growth rate was calculated,
3.2. Nutrient removal Fig. 3 shows a decrease in nutrient concentration in the growth medium for Dunaliella sp., Nannochloropsis sp., and Tetraselmis sp., during the whole culture period. Table 3 shows that, in general, a removal higher than 90% for nitrogen and orthophosphate was obtained for all 3 microalgae species. The removal of dissolved organic matter measured as COD was close to 80% for all 3 microalgae species, except for Tetraselmis sp. at the 2000 μmol m−2 s−1 PAPFD, where the removal was 75%. These results are similar to those reported by [43] where a consortia of microalgae and bacteria were cultured in treated wastewater from a fish farm, reaching a COD removal of 77%, with a much higher initial concentration (678 ± 249 mg O2 L−1) than the concentration used in the present study. Total nitrogen and orthophosphate removal were both higher than 90% for all three microalgae species. Removal values for total nitrogen in this study were higher than those obtained in other studies: [42] reported a nitrogen removal of 49.4% with Tetraselmis suecica cultivated in fish farm wastewater (initial concentration of 41.3 mg L−1); 101
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
smallest biovolume (Nannochloropsis sp.) also showed the lowest biomass productivities, while the microalga with the biggest biovolume (Tetraselmis sp.) showed the highest biomass productivities at all 3 PAPFDs. All lipid productivities were significantly different for all 3 microalgae species (p < 0.05). Nitrogen and phosphorus concentrations had a significant effect in the lipid production in all 3 microalgae species (p < 0.05), this is because at the logarithmic growth phase there is sufficient nitrogen and phosphorous in the culture for the microalgae to grow. In late phases of the culture concentrations of nitrogen and phosphorous decrease. Once at the stationary phase where there are low concentrations of both nutrients, the microalgae become stressed and start accumulating lipids as an energy reserve. However, PAPFD had no significant effect in the lipid productivity (p > 0.05), in any of the microalgae species. It is important to direct future research to the study of continuous cultures to determine if the microalgal biomass productivity and lipid content can be increased.
[43] reported 80% of nitrogen removal, with a total initial nitrogen concentration (31 ± 10 mg L−1) lower than the initial nitrogen concentration in this study; [44] reported a 78% removal with N. oculata cultivated in aquaculture wastewater (initial concentration of 40 mg L−1); and [41] reported only 40% of total nitrogen removal due to a nitrogen overconcentration in the tested wastewater (338 mg L−1). Orthophosphates removal values in our results were similar to [41,42], 90% and 99% removal, respectively (with initial concentrations of 25 mg L−1 and 4.96 mg L−1); while [43] reported an orthophosphates removal of 78% (initial concentration of 14 ± 7 mg L−1); and [44] reported 79% of phosphorus removal (initial concentration of 10 mg L−1) with T. chuii; being these removal percentages lower than those reported here. The nitrogen and phosphorus removal kinetics for Tetraselmis sp. were higher than for the other two microalgae species (Table 3), this is explained because a faster growing rate, resulting in a rapid uptake of nutrients. However, Dunaliella sp. and Tetraselmis sp. showed very similar dissolved organic matter removal kinetics, measured as COD, despite their different growing rates. The removal kinetics reported by [41] were much higher (35 mg L−1 day−1 for total nitrogen, and 5.7 mg L−1 day−1 for total phosphorus) than the values reported in this study, since the culture technique was a continuous culture where a volume of medium was extracted daily and replaced with wastewater.
4. Conclusions It has been demonstrated that microalgae are useful in treating mariculture wastewater in view of their high nutrient removal values (i.e., nitrogen, phosphorus, and carbon), especially because these nutrients are commonly found in this type of wastewater. It is possible to enhance the lipid productivities of the tested microalgae by keeping them in the culture wastewater during the stationary phase of the culture, when low-nutrient concentrations are present in the wastewater. This strategy induces stress conditions forcing the microalgae to accumulate lipids as an energy source. Also the photosynthetically active photon flux density has an influence over the biomass and lipid productivities, since at specific PAPFDs the highest biomass and lipid productivities were obtained. This would translate in reducing the wastewater treatment cost because the obtained microalgae biomass byproduct would render a higher biodiesel production.
3.3. Lipid content Fig. 4 shows the lipid production for all 3 microalgae species. In all 3 cases it is evident that once the stationary phase is reached (Fig. 2), the lipid content exponentially increases. At the stationary phase, nutrient concentrations are low, so the microalgae start producing lipids as an energy reserve [45]. Fig. 2 shows that Tetraselmis sp. reached the stationary phase on day 9 of culture. Fig. 4 shows that on day 8, the exponential lipid production starts, precisely at the time where the nutrient concentration is low (Fig. 3). In the case of Dunaliella sp., the stationary phase was reached on day 13 of culture (Fig. 2), when nutrient concentration is low (Fig. 3), and as a result the quantity of lipids increases (Fig. 4). Nannochloropsis sp. showed a similar pattern, reaching the stationary phase on day 14 of culture (Fig. 2) when the nutrient concentration is low (Fig. 3), however, the lipid production starts increasing around day 16 (Fig. 4). Tetraselmis sp. showed the highest lipid concentration, probably due to its bigger size it can store more lipids inside the cell. The microalgae lipid productivity was calculated according to [31] (Table 4), in this case Nannochloropsis sp. showed the lowest lipid productivities, due to its long culture time. Tetraselmis sp. and Dunaliella sp. showed their highest lipid productivities at 1500 μmol m−2 s−1 PAPFD (29.48 and 19.42 mg L−1 day−1, respectively), while Nannochloropsis sp. showed its highest lipid productivity at 2000 μmol m−2 s−1 PAPFD (13.03 mg L−1 day−1). For Dunaliella sp. and Tetraselmis sp. the lipid productivities at 900 and 2000 μmol m−2 s−1 PAPFDs were similar. Tetraselmis sp. showed the highest lipid productivity (29.48 mg L−1 day−1), at 1500 μmol m−2 s−1 PAPFD. The productivities observed for Nannochloropsis sp. are lower than those reported by [46], i.e. 12.9 mg L−1 day−1 for Nannochloropsis salina, and 18.0 mg L−1 day−1 for Nannochloropsis gaditana. The lipid productivities obtained for Tetraselmis sp. are higher than those reported by [47] for Tetraselmis suecica in a nitrogen-limited medium (18.7 mg L−1 day−1), but lower than the value reported for this microalga in a growth medium (34.6 mg L−1 day−1). Dunaliella sp. productivities were lower than the values reported by [48], which were 38.0 mg L−1 day−1 and 56.0 mg L−1 day−1 in a nitrogen-limited medium and a nutrient-sufficient medium, respectively. Tetraselmis sp. showed the best results with the highest biomass productivity at 900 μmol m−2 s−1 (132.8 mg L−1 day−1). However, the highest lipid productivity (29.5 mg L−1 day−1) was observed at 1500 μmol m−2 s−1. Again, it is noticeable that the microalga with the
Conflict of interest The authors declare that there is no conflict of interest. No conflicts, informed consent, human or animal rights are applicable for this work. Acknowledgements We thanks for the economic support provided by Project UNAM/ DGAPA PAPIIT IT202818 “Desarrollo de una estrategia para el tratamiento de aguas residuales de maricultura empleando microalgas y aprovechamiento de la biomasa producida” and PAIP (VMLP) 50009111 granted to VMLP by the Facultad de Química, UNAM. CONACYT (421332), Mexico grant received by Manuel Sacristán for doctoral studies (Ph.D. studies) on Engineering in the PMyD at UNAM. Iveth Gabriela Palomino Albarrán, MSc, is kindly acknowledged for the donation of the microalgae cultures. Korinthya López Aguiar, MSc, is kindly acknowledged for the technical assistance in the wastewater analysis. Luciano Hernández Gómez, MSc, for his technical support. References [1] FAO, The State of World Fisheries and Aquaculture. Contributing to Food Security and Nutrition for All, (2016). [2] N.M. Nasir, N.S.A. Bakar, F. Lananan, S.H. Abdul Hamid, S.S. Lam, A. Jusoh, Treatment of African catfish, Clarias gariepinus wastewater utilizing phytoremediation of microalgae, Chlorella sp. with Aspergillus niger bio-harvesting, Bioresour. Technol. (2015), https://doi.org/10.1016/j.biortech.2015.03.023 (in press). [3] S.A. Castine, A.D. McKinnon, N.A. Paul, L.A. Trott, R. de Nys, Wastewater treatment for land-based aquaculture: improvements and value-adding alternatives in model systems from Australia, Aquac. Environ. Interact. 4 (2013) 285–300, https://doi. org/10.3354/aei00088. [4] S. Dinesh Kumar, P. Santhanam, R. Nandakumar, S. Ananth, P. Nithya, B. Dhanalakshmi, et al., Bioremediation of shrimp (Litopenaeus vannamei) cultured effluent using copepod (Oithona rigida) and microalgae (Picochlorum maculatam &
102
Algal Research 34 (2018) 97–103
M. Sacristán de Alva et al.
[5]
[6] [7]
[8]
[9]
[10]
[11]
[12]
[13]
[14] [15]
[16]
[17]
[18]
[19]
[20]
[21]
[22]
[23]
[24]
[25]
10.1016/j.biotechadv.2016.06.004. [26] P. Singh, S. Kumari, A. Guldhe, R. Misra, I. Rawat, F. Bux, Trends and novel strategies for enhancing lipid accumulation and quality in microalgae, Renew. Sust. Energ. Rev. 55 (2016) 1–16, https://doi.org/10.1016/j.rser.2015.11.001. [27] S. Esakkimuthu, V. Krishnamurthy, R. Govindarajan, K. Swaminathan, Augmentation and starvation of calcium, magnesium, phosphate on lipid production of Scenedesmus obliquus, Biomass Bioenergy 88 (2016) 126–134, https://doi. org/10.1016/j.biombioe.2016.03.019. [28] Y. Gong, M. Jiang, Biodiesel production with microalgae as feedstock: from strains to biodiesel, Biotechnol. Lett. 33 (2011) 1269–1284, https://doi.org/10.1007/ s10529-011-0574-z. [29] A. Demirbas, M. Fatih Demirbas, Importance of algae oil as a source of biodiesel, Energy Convers. Manag. 52 (2011) 163–170, https://doi.org/10.1016/j.enconman. 2010.06.055. [30] R. Huerlimann, R. de Nys, K. Heimann, Growth, lipid content, productivity, and fatty acid composition of tropical microalgae for scale-up production, Biotechnol. Bioeng. 107 (2010) 245–257, https://doi.org/10.1002/bit.22809. [31] A. Toledo-Cervantes, M. Morales, E. Novelo, S. Revah, Carbon dioxide fixation and lipid storage by Scenedesmus obtusiusculus, Bioresour. Technol. 130 (2013) 652–658, https://doi.org/10.1016/j.biortech.2012.12.081. [32] K. Grasshoff, K. Kremling, M. Ehrhardt, Methods of Seawater Analysis, third edition, Wiley-VCH, Germany, 1999. [33] F.A. Richards, R.A. Kletsch, The Spectophotometric Determination of Ammonia and Labile Amino Compounds in Fresh and Seawater by Oxidation to Nitrite, (1964). [34] B. Schnetger, C. Lehners, Determination of nitrate plus nitrite in small volume marine water samples using vanadium(III)chloride as a reduction agent, Mar. Chem. 160 (2014) 91–98, https://doi.org/10.1016/j.marchem.2014.01.010. [35] APHA, AWWA, WEF, Standard Methods for the Examination of Water and Wastewater, 20th ed., American Public Health Association, American Water Works Association, Water Environment Federation, Washington D.C., 1998. [36] I. Vyrides, D.C. Stuckey, A modified method for the determination of chemical oxygen demand (COD) for samples with high salinity and low organics, Bioresour. Technol. 100 (2009) 979–982, https://doi.org/10.1016/j.biortech.2008.06.038. [37] S.K. Mishra, W.I. Suh, W. Farooq, M. Moon, A. Shrivastav, M.S. Park, et al., Rapid quantification of microalgal lipids in aqueous medium by a simple colorimetric method, Bioresour. Technol. 155 (2014) 330–333, https://doi.org/10.1016/j. biortech.2013.12.077. [38] R.S. Gour, A. Kant, R.S. Chauhan, Screening of micro algae for growth and lipid accumulation properties, J. Algal Biomass Util. 5 (2014) 38–46. [39] FAO, Manual on the Production and Use of Live Food for Aquaculture, (1996), https://doi.org/10.1007/s13398-014-0173-7.2. [40] D. Hanelt, Photoinhibition of photosynthesis in marine microalgae, Sci. Mar. 60 (1996) 243–248. [41] C. Sepúlveda, F.G. Acién, C. Gómez, N. Jiménez-Ruíz, C. Riquelme, E. MolinaGrima, Utilization of centrate for the production of the marine microalgae Nannochloropsis gaditana, Algal Res. 9 (2015) 107–116, https://doi.org/10.1016/j. algal.2015.03.004. [42] M.H.A. Michels, M. Vaskoska, M.H. Vermuë, R.H. Wiffels, Growth of Tetraselmis suecica in a tubular photobioreactor on wastewater from a fish farm, Water Res. 65 (2014) 290–296, https://doi.org/10.1016/j.watres.2014.07.017. [43] E. Posadas, A. Muñoz, M.-C. García-González, R. Muñoz, P.A. García-Encina, A case study of a pilot high rate algal pond for the treatment of fish farm and domestic wastewaters, J. Chem. Technol. Biotechnol. (2014), https://doi.org/10.1002/jctb. 4417 (n/a-n/a). [44] I.N. Sirakov, K.N. Velichkova, Bioremediation of wastewater originate from aquaculture and biomass production from microalgae species - Nannochloropsis oculata and Tetraselmis chuii, Bulgarian J. Agr. Sci. 20 (2014) 66–72. [45] I.A. Guschina, J.L. Harwood, Algal lipids and effect of the environment on their biochemistry in lipids in aquatics, Ecosystems (2009), https://doi.org/10.1017/ CBO9781107415324.004. [46] A. Taleb, J. Pruvost, J. Legrand, H. Marec, B. Le-Gouic, B. Mirabella, et al., Development and validation of a screening procedure of microalgae for biodiesel production: application to the genus of marine microalgae Nannochloropsis, Bioresour. Technol. 177 (2015) 224–232, https://doi.org/10.1016/j.biortech.2014. 11.068. [47] E. Del Río, A. Armendáriz, E. García-Gómez, M. García-González, M.G. Guerrero, Continuous culture methodology for the screening of microalgae for oil, J. Biotechnol. 195 (2015) 103–107, https://doi.org/10.1016/j.jbiotec.2014.12.024. [48] G. Breuer, P.P. Lamers, D.E. Martens, R.B. Draaisma, R.H. Wijffels, The impact of nitrogen starvation on the dynamics of triacylglycerol accumulation in nine microalgae strains, Bioresour. Technol. 124 (2012) 217–226, https://doi.org/10. 1016/j.biortech.2012.08.003.
Amphora sp.)—an integrated approach, Desalin. Water Treat. (2016) 1–10, https:// doi.org/10.1080/19443994.2016.1163509. F. Gao, C. Li, Z. Yang, G. Zeng, L. Feng, J. Liu, et al., Continuous microalgae cultivation in aquaculture wastewater by a membrane photobioreactor for biomass production and nutrients removal, Ecol. Eng. 92 (2016) 55–61, https://doi.org/10. 1016/j.ecoleng.2016.03.046. D.G. Kim, Y.-E. Choi, Microalgae cultivation using LED light, Korean Chem. Eng. Res. 52 (2014) 8–16, https://doi.org/10.9713/kcer.2014.52.1.8. F. Bumbak, S. Cook, V. Zachleder, S. Hauser, K. Kovar, Best practices in heterotrophic high-cell-density microalgal processes: achievements, potential and possible limitations, Appl. Microbiol. Biotechnol. 91 (2011) 31–46, https://doi.org/10. 1007/s00253-011-3311-6. J. Wang, H. Yang, F. Wang, Mixotrophic cultivation of microalgae for biodiesel production: status and prospects, Appl. Biochem. Biotechnol. 172 (2014) 3307–3329, https://doi.org/10.1007/s12010-014-0729-1. W.-W. Li, H.-Q. Yu, Z. He, Towards sustainable wastewater treatment by using microbial fuel cells-centered technologies, Energy Environ. Sci. 7 (2014) 911–924, https://doi.org/10.1039/c3ee43106a. K.K. Sharma, H. Schuhmann, P.M. Schenk, High lipid induction in microalgae for biodiesel production, Energies 5 (2012) 1532–1553, https://doi.org/10.3390/ en5051532. A.K. Minhas, P. Hodgson, C.J. Barrow, A. Adholeya, A review on the assessment of stress conditions for simultaneous production of microalgal lipids and carotenoids, Front. Microbiol. 7 (2016) 1–19, https://doi.org/10.3389/fmicb.2016.00546. L. Xin, H. Hu, G. Ke, Y. Sun, Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid accumulation of a freshwater microalga Scenedesmus sp. Bioresour. Technol. 101 (2010) 5494–5500, https://doi. org/10.1016/j.biortech.2010.02.016. J.-V. Pérez-Pazos, P. Fernández-Izquierdo, Synthesis of neutral lipids in Chlorella sp. under different light and carbonate conditions, Ciencia Tecnol. Futur. 4 (2011) 47–58. L.D. Zhu, Z.H. Li, E. Hiltunen, Strategies for lipid production improvement in microalgae as a biodiesel feedstock, Biomed. Res. Int. 2016 (2016) 1–8. Q. Hu, M. Sommerfeld, E. Jarvis, M. Ghirardi, M. Posewitz, M. Seibert, et al., Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances, Plant J. 54 (2008) 621–639, https://doi.org/10.1111/j.1365-313X.2008. 03492.x. O. Perez-Garcia, F.M.E. Escalante, L.E. De-Bashan, Y. Bashan, Heterotrophic cultures of microalgae: metabolism and potential products, Water Res. 45 (2011) 11–36, https://doi.org/10.1016/j.watres.2010.08.037. A.M.J. Kliphuis, A.J. Klok, D.E. Martens, P.P. Lamers, M. Janssen, R.H. Wijffels, Metabolic modeling of Chlamydomonas reinhardtii: energy requirements for photoautotrophic growth and maintenance, J. Appl. Phycol. 24 (2012) 253–266, https:// doi.org/10.1007/s10811-011-9674-3. Y. Li, X. Fei, X. Deng, Novel molecular insights into nitrogen starvation-induced triacylglycerols accumulation revealed by differential gene expression analysis in green algae Micractinium pusillum, Biomass Bioenergy 42 (2012) 199–211, https:// doi.org/10.1016/j.biombioe.2012.03.010. A. Patel, S. Barrington, M. Lefsrud, Microalgae for phosphorus removal and biomass production: a six species screen for dual-purpose organisms, GCB Bioenergy 4 (2012) 485–495, https://doi.org/10.1111/j.1757-1707.2012.01159.x. J. Msanne, D. Xu, A. Reddy, J.A. Casas-Mollano, T. Awada, E.B. Cahoon, et al., Metabolic and gene expression changes triggered by nitrogen deprivation in the photoautotrophically grown microalgae Chlamydomonas reinhardtii and Coccomyxa sp. C-169, Phytochemistry 75 (2012) 50–59, https://doi.org/10.1016/j.phytochem. 2011.12.007. L.A. Garay, K.L. Boundy-Mills, J.B. German, Accumulation of high-value lipids in single-cell microorganisms: a mechanistic approach and future perspectives, J. Agric. Food Chem. 62 (2014) 2709–2727, https://doi.org/10.1021/jf4042134. J.J. Mayers, K.J. Flynn, R.J. Shields, Influence of the N:P supply ratio on biomass productivity and time-resolved changes in elemental and bulk biochemical composition of Nannochloropsis sp. Bioresour. Technol. 169 (2014) 588–595, https:// doi.org/10.1016/j.biortech.2014.07.048. H.J. Choi, S.M. Lee, Effect of the N/P ratio on biomass productivity and nutrient removal from municipal wastewater, Bioprocess Biosyst. Eng. 38 (2015) 761–766, https://doi.org/10.1007/s00449-014-1317-z. N.W. Rasdi, J.G. Qin, Effect of N:P ratio on growth and chemical composition of Nannochloropsis oculata and Tisochrysis lutea, J. Appl. Phycol. 27 (2015) 2221–2230, https://doi.org/10.1007/s10811-014-0495-z. S.K. Lenka, N. Carbonaro, R. Park, S.M. Miller, I. Thorpe, Y. Li, Current advances in molecular, biochemical, and computational modeling analysis of microalgal triacylglycerol biosynthesis, Biotechnol. Adv. 34 (2016) 1046–1063, https://doi.org/
103