Medium recycling for Nannochloropsis gaditana cultures for aquaculture

Bioresource Technology 129 (2013) 430–438

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Medium recycling for Nannochloropsis gaditana cultures for aquaculture C.V. González-López a, M.C. Cerón-García a,⇑, J.M. Fernández-Sevilla a, A.M. González-Céspedes b, J. Camacho-Rodríguez a, E. Molina-Grima a a b

Department of Chemical Engineering, University of Almería, E04120 Almería, Spain Department of Greenhouse Technology, Estación Experimental Fundación Cajamar, Almería E04710, Spain

h i g h l i g h t s " A methodology to recirculate medium in microalgae cultures is developed. " The most adequate sterilization method is established. " The recirculation reduces the demand of nutrients cutting costs in the process. " N. gaditana continuous cultures were maintained using the recirculated medium. " The biomass biochemical composition resulted of high interest for aquaculture.

a r t i c l e

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Article history: Received 1 August 2012 Received in revised form 12 November 2012 Accepted 16 November 2012 Available online 28 November 2012 Keywords: Medium recycling Sterilization Microalgae Aquaculture

a b s t r a c t Nannochloropsis gaditana is a good producer of proteins and valuable fatty acids for aquaculture. Recycling of culture medium is interesting for microalgae commercial production as it cuts costs and prevents environmental contamination. The recycled medium must be sterilized to prevent the buildup of unwanted metabolites and microorganisms. We tested several sterilization methods: filtration, ozonation, chlorination, addition of hydrogen peroxide and heating. Results showed that the most successful method is ozonation lowering the bacterial load to 1.9 103 CFUs/mL, which is 1000-fold and 10-fold lower than the supernatant obtained after harvesting and the initial filtered medium, respectively. Continuous cultures of N. gaditana were grown using this recirculated supernatant. A maximum biomass productivity of 0.8 g/L/d composed of 50% proteins and 40% lipids with more than 3% d.w. EPA was obtained making this biomass very interesting for aquaculture. Ó 2012 Elsevier Ltd. All rights reserved.

1. Introduction The production of microalgae is essential for the commercial cultivation of larvae of mollusks, crustaceans and fish. Microalgae can be used directly as live food (for mollusks and crustaceans) or indirectly as food for zooplankton species, such as the rotifer Brachiomus plicatilis and nauplii of Artemia, used as prey for fish larvae (Benemann, 1992). The microalgae used as food in the early larval stages of fish should ensure the nutritional requirements of the species, particularly in essential compounds like amino acids or polyunsaturated fatty acids. The microalgal diet must contain high levels of polyunsaturated fatty acids as docosahexanoic acid (DHA 22:6n3), eicosapentanoicacid (EPA 20:5n3) and araquidonic acid (AA 20:4n6) to ensure a good larvae growth and high levels of survival (Navarro and Villanueva, 2003; Morais et al., 2005).

⇑ Corresponding author. Tel.: +34 950 015981; fax: +34 950 015484. E-mail address: [email protected] (M.C. Cerón-García). 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.11.061

The maintenance of axenic mass cultures of microalgae requires the sterilization of large volumes of seawater and thus of a method that can be implemented in the large scale. Kawachi and Nöel (2005) reviewed several techniques to sterilize seawater. Among the methods discussed by these authors are filtration, application of ultraviolet radiation (UV), addition of sodium hypochlorite and pasteurization. Other techniques such as membrane-based separation methods have been suggested (Rathore and Shirke, 2011) but these are more difficult to implement in large-scale culture systems and especially in seawater. Many aquaculture facilities use filtration, UV irradiation or a combination of both methods to sterilize seawater. Ozonation and ultraviolet irradiation (UV) are the most frequently used methods for viral control in land based aquacultural systems. These two methods can be used to eliminate pathogens in the inlet seawater culture medium, in the supernatant and in the recirculated water (the supernatant obtained after harvesting). Disinfection by ozonation and UV irradiation are also used in other aquacultural applications, e. g., to reduce or eliminate the presence of potential

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pathogens associated with live prey, such as rotifers, in marine larval production systems or to disinfect the surface of fish eggs (Grotmol and Totland, 2000). The filtration only eliminates particles of the size of the smallest filter and does not eliminate viruses. A chemical sterilization method is the chlorination of water with bleach (sodium hypochlorite) for several hours, followed by the removal of the excess of chlorine by the addition of sodium thiosulphate and aeration (Kawachi and Nöel, 2005). Regarding thermal disinfection, pasteurization has proved to be an effective method for two species of algae: Rhodomonas lens (CCMP 739) and R. salina (CCMP 1319) (Rhodes et al., 2008). The use of ozone began in the 1990s, with the purification of drinking water and ever since it has become widespread. Ozone is used for disinfection of bottled water and in the oxidative decomposition of harmful impurities in industrial and domestic sewage (Martínez et al., 2011). It is a strong disinfectant with a high oxidation potential and it is one of the most effective and frequently used for deactivating pathogens in drinking water treatments (Langlais et al., 1991). Ozone also acts against various cellular substances including lipids, peptidoglycans, enzymes and viruses. It is a powerful disinfectant against bacteria, fungi, etc. and it can therefore be used to satisfy the highest disinfection standards (Tyrrell et al., 1995). The production of microalgae at large-scale with medium recycling has not yet been commercialized. The life-cycle impacts of large-scale microalgae productions are being debated, specially the impact on water usage, i. e., the water consumption per hectare of land used for algal feedstock production. As far as we know, the life-cycle of water when using seawater and recycling the harvested water to produce microalgae in aquaculture has not been established yet. The nearer attempts are those of Yang et al. (2011) and Jung and Lovitt (2010) who recycled the harvested water for microalgae production with the objective to obtain biodiesel and PUFAs, respectively. Therefore, if the supernatant obtained from the harvest of the biomass is not recirculated to the system, high amounts of nutrients are thrown out. It is possible to reuse those nutrients by using closed systems to recirculate the water and take the most of the nutrients not consumed. The biggest challenge of this issue is to develop a method to sterilize the supernatant obtained from the harvested biomass of microalgae cultures and to recirculate it to the system as to work in a closed circuit. The harvesting process involves the presence of suspended solids in the supernatant. These solids are organic matter that accumulates over time. So it is necessary to study the influence of this organic matter on the microalga growth. For example, Chlorella sp. uses organic matter to promote its growth and the subsequent reduction of chemical oxygen demand (COD) (Li et al. 2011). COD is an important and easily reproducible parameter in the analysis of domestic and industrial wastewaters in terms of the evaluation of their pollution levels. Nevertheless, the methodology for its determination is not well adapted to seawater so the results are not reliable. Saral and Goncalog (2008) developed a method to determine COD of saline wastewater (up to 14 gNaCl/L) but it involved the use of toxic chemicals. On the other hand, Aziz and Tebbut (1980) assessed that the total organic carbon (TOC) is an alternative to COD measurements. The TOC measurement is not dependent on the salt content of the sample. So to account the organic matter, the TOC content would be used as the preferable parameter. The aim of this work is to develop a methodology at laboratory scale that allows recirculating the supernatant obtained after harvesting microalgae cultures. For that, it is necessary to establish the best sterilization method to be used in order to avoid any influence on the microalgae production and/or on their biochemical composition. The experimentation has been designed bearing in mind

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that the technique selected will be later implemented in a large closed photobioreactor of commercial scale. In this way, the supernatant could be used as a nutrient source for the production of high value biomass reducing the demand of nutrients and emitting cleaner effluents. Once the optimal technique for recirculation is established, it will be possible in future contribution to study long-term recycling and its economic impact in large-scale systems. 2. Methods 2.1. Sterilization methods The artificial seawater culture medium used was provided by the microalgal production facility ‘‘Estación Experimental Fundación Cajamar’’ property of Cajamar (Paraje Las Palmerillas, Almería, Spain). The composition of this medium is the same as the commercial Algal medium (Bionova, Santiago de Compostela, Spain) prepared at 8 mM KNO3 concentration but using agricultural fertilizers instead of pure grade chemicals. The tests were performed at laboratory scale in 100 mL Erlenmeyer flasks containing 50 mL of the seawater culture medium. They were incubated on an orbital shaker (Orbital midi, Ovan, Barcelona, Spain) at 100 rpm without aeration under 25 °C and aseptic conditions during 7 days. Six different techniques were used to sterilize the medium: heating and addition of four different chemicals. The seawater culture medium was first sterilized in autoclave (Presoclave-II, J.P. Selecta, Barcelona, Spain) at 126 °C for 20 min in order to guarantee the sterilization used as the control test. The chemical sterilization was performed by the addition of four compounds: sodium hypochlorite (PANREAC Química S.L.U., Barcelona, Spain), dichloroisocyanurate (Suavinex C.N. 178665, Alicante, Spain), ozone (Ozone generator 3060, ETRON Ecology S.L., Spain) and hydrogen peroxide (PANREAC Química S.L.U., Barcelona, Spain). The chlorination was carried out with different doses of sodium hypochlorite: 0.44, 0.72, 0.86, 1.00, 1.50 and 2.00 mg/L. Sodium dichloroisocyanurate was added at concentrations of 200 and 400 mg/L. Ozonation was applied at 16, 95, 127, 174, 237, 300, 348, 396 and 459 mg/L with an ozone generator that produces 950 mgO3/h. The hydrogen peroxide dosages were: 2.6%, 3.8%, 5.0% and 10.0% v/v. Finally, the sterilization by pasteurization was carried out at temperatures of 40, 60, 70 and 90 °C for 5 min. 2.2. Evaluation of sterilization methods The effectiveness of each sterilization method was checked by following the growth of bacteria in the 100 mL Erlenmeyer flasks each day (during 7 days). The initial pH of each sample was adjusted to pH 7.8 to allow a comparison among flasks starting at the same conditions. As estimation, the bacterial contamination was measured spectrophotometrically by the absorbance at 680 nm (Helios Omega UV–VIS Spectrophotometer, Thermo Scientific, Horsham, England) as described by Brown (1980). The flasks which did not seem to present any contamination were checked by a bacterial count test as described by Eaton et al. (1995). This method was also used for the supernatant obtained after harvesting microalgae cultures and for seawater culture medium in order to check the effectiveness of the sterilization. It is also included water obtained from a filtration unit installed in the ‘‘Estación Experimental Fundación Cajamar’’ which consists of several filters of different pore size lasting in a filter of 1.2 lm (Merck Millipore, Germany). To carry out the bacterial count 0.1 mL aliquots of the samples were placed in Petri dishes (each sample in triplicate). Each sample was previously diluted (1:10, 1:100 or 1:1000 or 1:10,000) because of the different bacteria load in each one. The Petri dishes

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contained a culture medium composed of tryptic soy agar 1% w/v in seawater (TSA; Hardy, Santa Maria, CA, USA) as it is widely recommended for water and wastewater applications (Eaton et al., 1995). The samples were incubated for 48 h at 25 °C. After that, the grown bacteria colonies were counted to determine the concentration in CFU/mL. 2.3. Microorganism and culture conditions The strain Nannochloropsis gaditana B-3 was obtained from the Marine Culture Collection of the Institute of Marine Sciences of Andalucía (CSIC, Cádiz, Spain). The stock cultures were maintained photoautotrophically in 100 mL Erlenmeyer flasks containing 50 mL of culture. They were aseptically grown on an orbital shaker at 100 rpm without aeration under a continuous illumination of 100 lE/m2/s and a temperature of 25 °C. The seawater culture medium used was the commercial medium Algal (Bionova, Santiago de Compostela, Spain) prepared at 8 mM concentration of potassium nitrate. The inorganic components were added to natural seawater. The macronutrients were sterilized in autoclave (Presoclave-II, J.P. Selecta, Barcelona, Spain) at 126 °C for 20 min and the micronutrients were filtered (Whatman GF/F 47 mm, nominal pore size 0.22 lm, Maidstone, United Kingdom). The recirculated supernatant used for the tests was obtained from outdoor cultures in tubular photobioreactors (Estación Experimental Fundación Cajamar). These reactors were operated in continuous mode as described by Acién et al. (2012) harvesting the microalgae culture with a disc centrifuge (Westfalia OTC3, Holland at 268 L/h). 2.4. Effect of the sterilization methods in N. gaditana growth N. gaditana cultures were tested in order to check if the microalga is able to grow correctly using a recirculated supernatant (obtained after centrifugation of the cultures of the ‘‘Estación Experimental’’) or if it is necessary to mix this supernatant prior to the recirculation with new culture medium (Algal) and in what extent. For this, experiments in (i) batch mode and (ii) continuous modes were carried out. 2.4.1. Batch mode With regard to the batch mode 100 mL Erlenmeyer flasks were used in order to check the effect on N. gaditana growth under the temperature and irradiance previously described. The reported data are mean values of nine experimental measurements. In this sense, a second series of batch experiments in 100 mL Erlenmeyer flasks were carried out to select the proportion of the supernatant that is possible to recirculate. Therefore, the medium supplied to the microalgae cultures consisted of mixtures of supernatant and new seawater medium in the following proportions: 0– 100, 25–75, 50–50, 75–25 and 100–0 (supernatant-Algal medium). An additional test consisted of 100% recirculated water but replenishing the nutrients consumed during the culture (100⁄–0). The tests were carried out in triplicate. 2.4.2. Continuous mode After the batch tests, experiments at larger scale (1.8 L bubble column photobioreactors) were carried out indoors in continuous mode in order to verify the tests previously performed at smaller scale. The proportions of recirculated supernatant to Algal medium were the same than the described for the batch tests: 0–100, 50– 50, 75–25,100–0 and 100⁄–0. The culture system consisted of two bubble column photobioreactors (1.8 L capacity, 0.07 m diameter, 0.50 m height) jacketed for temperature control at 18 °C. They were bubbled with air at 0.5 v/v/min. The pH was controlled at 8.0

by on demand injection of carbon dioxide. The reactors were artificially illuminated simulating a solar cycle by six Phillips PL32 W/840/4p white-light lamps. The irradiance on the reactor surface was controlled by an automated system to provide a maximum irradiance of 1000 lE/m2/s at noon. The dilution rate was fixed at 0.3 1/d. The steady states attained were maintained at least for three days to completely ensure steady conditions regarding the biochemical composition of the biomass. Three samples were measured each day on the last three days of every steady state; therefore, the reported values are the means of nine measurements. For both operation modes the new seawater culture medium (Algal) and the recirculated supernatant were sterilized using the most successful method studied in this work, taking into account the obtained results in Section 2.2 and Section 2.4. The cultures were harvested by centrifugation (9100g, 5 min), washed with a 0.5 M aqueous ammonium bicarbonate solution, centrifuged again and freeze-dried. The dry biomass was analyzed immediately or stored at 22 °C for up to 10 days prior to analysis. 2.5. Analytical procedures The microalgal biomass concentration was estimated spectrophotometrically by measuring the absorbance at 540 nm (Helios Omega UV–VIS Spectrophotometer, Thermo Scientific, Horsham, England) and it was verified by dry weight measurements. The physiological status of the cells was determined by the fluorescence of chlorophylls (Fv/Fm) using a fluorimeter (AquaPen-C APC 100, Photon Systems Instruments, Czech Republic). The presence of organic matter in the recirculated supernatant was measured by the total organic carbon concentration (TOC analyzer, Shimadzu VCPH, Japan). The consumption of nutrients by the microalgae was analyzed after filtration to eliminate particles (0.22 lm) following the Official methods of analysis of AOAC (2002). Nitrates: by absorbance at 220 and 275 nm in an acid environment. Phosphorous: by a colorimetric method with ammonium molybdate at 430 nm. Sulfates: by turbidimetry. Iron, copper, manganese and zinc: by atomic absorption (Perkin-Elmer Analyst100, EE UU). The fatty acid content of the biomass was analyzed by gas chromatography as described by Rodríguez-Ruiz et al. (1998) using freeze-dried biomass. The total lipid content was determined as described by Kochert (1978). Ash and protein contents were analyzed by the methods described by Brown et al. (1989) and López et al. (2010), respectively. Finally, carbohydrates content was determined by difference between 100 and the percentage of the rest of the fractions (ashes, proteins and total lipids). The elemental analysis of the biomass was performed by means of a LECO CHNS932 analyzer (St. Joseph, MI, USA). Finally, the statistical analysis of the data was carried out using the software Statgraphics version 7.0. 2.6. Kinetic parameters The nutrient requirements of the biomass were determined using the growth stoichiometry of the microalga by the following equation:

aCO2 þ bKNO3 þ cH2 O ! Cw Hx Oy Nz þ dO2

ð1Þ

CwHxOyNz represents the dry microalgal biomass. This equation is presented from a macroscopic point of view and it only takes into account the nutrients that could be limiting for the growth of the microalga. To solve the equation one mol of biomass was fixed as the reference (w = 1). The elemental composition of the biomass was determined in order to calculate the stoichiometric

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where S and S0 are the final and initial substrate concentrations (g/ L), respectively and X is the average biomass concentration (g/L) at the stationary state. The substrate requirement (YS/X, g substrate consumed/g biomass) is easily obtained as the inverse of substrate yield. The kinetic parameter used to control the uptake of the substrate was the biomass specific uptake rate (g substrate/day/g biomass), which was calculated with the following equation where D is the dilution rate (0.3 1/d).

coefficients. The biomass is composed of C, H, O and N in 90– 95% d.w., so the atomic balances of these four elements (referred to one atom of carbon) would allow determining the coefficients. After this, this formula permitted to estimate the nutrient requirements of the microalgae. Then, the yield of consumption of a nutrient was obtained from the Eq. (2) and the substrate yield (g biomass/g substrate consumed) was determined by Eq. (3):

Y X=S ¼

Molecular mass of the biomass b  ðMolecular mass of substrateÞ

ð2Þ

Y S=X ¼

ðS  S0 Þ X

ð3Þ

ð4Þ

0.5

0.5 control 2 mg/L 1.5 mg/L 1 mg/L 0.86 mg/L 0.72 mg/L 0.44 mg/L

2

0.4

3

0.4

0.3

0.2

0.3

0.2

0.1

0.1

0.0

0.0 0

1

2

3

4

5

0

6

2

4

6

8

10

12

Time, d

Time, d 0.5

0.5 control 459 mg/L 396 mg/L 348 mg/L 300 mg/L 237 mg/L 174 mg/L 127 mg/L 95 mg/L 16 mg/L

4

0.4

0.3

control 10 %v/v 5 %v/v 3.8 %v/v 2.6 %v/v

Hydrogen peroxide

5

0.4

Absorbance, 680 nm

Ozone

Absorbance, 680 nm

control 200 mg/L 400 mg/L

Sodium Dichloroisocyanurate Absorbance, 680 nm

Sodium hypochlorite Absorbance, 680 nm

D  ðS  S0 Þ X

qs ¼

0.2

0.3

0.2

0.1

0.1

0.0

0.0 0

1

2

3

4

5

0

6

1

2

3

4

5

6

Time, d

Time, d 0.5 control 90 ºC 70 ºC 60 ºC 40 ºC

Pasteurization

6

Absorbance, 680 nm

0.4

0.3

0.2

0.1

0.0 0

2

4

6

8

10

Time, d Fig. 1. Influence of the sterilization method at different dosages and conditions in seawater culture medium (Algal medium). The first number in abscises axis denotes sterilization methods. 1: autoclave (control), 2: sodium hypochlorite, 3: sodium dichloroisocyanurate, 4: ozone, 5: hydrogen peroxide, 6: pasteurization.

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3. Results and discussion Regarding the tests about the sterilization methods (Section 2.1), the culture medium was sterilized by different techniques in order to determine the optimal conditions. The effectiveness of each method was first estimated by the absorbance of the sample at 680 nm during 7 days (Fig. 1). The control consisted of a sample that was autoclaved at 126 °C for 20 min to ensure a complete sterilization. So the culture mediums presenting absorbance values similar the control are considered appropriate for sterilizing. Chlorination seems to be effective at sodium hypochlorite concentrations over 0.88 mg/L. The procedure based on adding sodium dichloroisocyanurate sterilizes the medium at a concentration of 200 mg/L as recommended the ‘‘Milton method’’, used to sterilize materials for babies. Ozonation eliminates contaminants even at the lowest tested concentration, 16 mg/L. The addition of hydrogen peroxide sterilizes the medium at dosages above 5% v/v. Finally, pasteurization is effective at temperatures over 60 °C (5 min treatment). However, the absorbance measurement is not enough as a method to assess the quality of the sterilization. Because of that, the bacterial count has been used. The analysis made to the seawater culture medium showed a low bacterial load of 3.7104 CFUs/

mL (Fig. 2a). If this medium is filtered at 1.2 lm the bacterial load is reduced 33%, up to 2.5 104 CFUs/mL. Lastly, if the culture medium is ozonizated at 95 mg/L it does not show significant levels of contamination (2.0102 CFUs/mL) as the bacterial load is reduced in two orders of magnitude. The same analysis was applied to the recirculated supernatant obtained after harvesting the microalgal cultures and similar results were obtained (Fig. 2b). Thus, the initial bacterial load of the supernatant is high (1.8106 CFUs/mL) due to the recirculation of bacteria and organic matter that promote their development. If the supernatant is filtered the bacterial load is reduced only one order of magnitude (1.9105 CFUs/mL). However, when using ozonation it is possible to reach a reduction of three orders of magnitude (1.9103 CFUs/mL). It ozonation decreases the bacterial load 1000-fold and 10-fold with regard to the recirculated supernatant and the initial seawater medium, respectively. A statistic study was performed to show the mean values of contamination reached for each tested sterilization method and the intervals around each mean value. The method used to discriminate among the mean values was the Fisher’s Least Significant Difference (LSD) procedure. The confidence bars of these methods overlap, which means that all could be used without significative differences (Fig. 2a, similar methods noted with the same letter). This allows observing that there are significant differences

2.0e+6

2.0e+6

(a)

Algal medium

1.0e+6 5.0e+5

4.0e+4 3.0e+4

1.5e+6

Bacteria load, CFUs/mL

a

2.0e+4

1.0e+6 5.0e+5

4.0e+4 3.0e+4 2.0e+4 1.0e+4

1.0e+4

b

c

b

b

b

c

%

6. 90 ºC

5. 5

m

m

g/ L

g/ L 4. 95

g/ L m

%

4. 95

6. 90 ºC

m

5. 5

g/ L

g/ L m

m

g/ L 3. 20 0

2. 1

tio n Fi ltr a 1.

In itia l

0.0

0.0 3. 20 0

b

2. 1

b

1. F

b

In iti al

b

Sterilization method

Sterilization method 2.0e+6

2.0e+6

(c)

Pasteurization temperature, ºC

bc

0.0

b

b 4. 19 0

6. 60

6. 40

In itia l

b

6. 90

b

6. 70

a 0.0

a

5.0e+5

4. 95

5.0e+5

1.0e+6

4. 47

1.0e+6

1.5e+6

In iti al

1.5e+6

(d)

Recirculated supernatant Bacteria load, CFUs/mL

Recirculated supernatant Bacteria load, CFUs/mL

(b)

Recirculated supernatant

iltr at io n

Bacteria load, CFUs/mL

1.5e+6

Ozone concentration, mg/L

Fig. 2. Variation of bacterial load with the sterilization method used in Algal medium and recirculated supernatant (a, b). Influence of different temperatures: 40, 60, 70 and 90 °C for 5 min used in pasteurization (c). Ozonation: 2.5, 5, 10 min of exposition time (d). The first number in abscises axis denotes sterilization methods. 1: filtration (control), 2: sodium hypochlorite, 3: sodium dichloroisocyanurate, 4: ozone, 5: hydrogen peroxide, 6: pasteurization. Note, for example, that 5 min of ozone exposition corresponds to 95 mg O3/L. All the following figures result from a Multifactorial ANOVA Statistical Study (mean values with different small letters denote significant difference, P < 0.05).

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Biomass concentration, g/L

a

aa

a a

0.8

d d 0.6

e

ee

0.4

b b bb 0.2

c

c

cc

c

c c

c

c

c

1. f

ilt ra t 2. ion 2 m 2. g/ 1. L 5 m 2. g/L 1 m 2. g 0. 86 /L m 3. 20 g/L 0 m g 4. 96 /L m 4. g /L 16 0 m 4. 32 g/L 0 m 4. 48 g/L 0 m g 5. 10 /L % 5. v/v 5 % v 6. /v 60 º 6. C 90 ºC

0.0

Used Method Fig. 3. Influence of the sterilization method on the biomass concentration of N. gaditana. Error bars correspond to mean values of biomass concentration in stationary state for each method. 1: filtration (control), 2: sodium hypochlorite, 3: sodium dichloroisocyanurate, 4: ozone, 5: hydrogen peroxide, 6: pasteurization. All the following figures result from a Multifactorial ANOVA Statistical Study (mean values with different small letters denote significant difference, P < 0.05).

between the group of methods 2, 3, 4, 5 and 6 (sodium hypochlorite, dichloroisocyanurate, ozone, hydrogen peroxide and pasteurization) and the filtration (1). Similarly Fig. 2b shows that all methods of sterilization are effective for sterilizing the recirculated supernatant with the exception of filtration. It was expected that filtration would not cause a sharp decline in bacterial load because the pore size employed was 1.2 lm, which is high. The results show that with pasteurization (Fig. 2c) the best results were achieved at 90 °C (5 min). However, the most remarkable fact is the high effectiveness of the ozonation (Fig. 2d) at a concentration of 95 mg/L obtained with a short contact time (5 min) resulting in a complete reduction of the bacterial load. Other chemical methods such as the addition of hydrogen peroxide or sodium dichloroisocyanurate resulted in chemical alteration of the water, observed as changes in color, especially when using dichloroisocyanurate. To select the best method to reduce the bacterial load in the recirculated supernatant it is also important to pay attention to its influence on the microalga growth. Therefore, laboratory scale batch cultures of N. gaditana using the supernatant obtained after harvesting the microalgae have been performed (see Section 2.4). The aim is to check if the use of these sterilization methods involves a harmful effect on the growth of the microalga. For that, the growth of Nannochloropsis is followed as shown in Fig. 3. It is observed that, although several methods are able to sterilize in any extent the supernatant (2, 3, 4, 5 and 6), only filtration and ozonation are tolerated by the microalgae. It can be seen how only

the flasks grown with the ozonized supernatant at low concentration (95 mg/L and 160 mg/L) and the filtered one are not affected by the sterilization method attaining similar biomass concentration values (0.8 g/L). The statistical study reveals that these two methods do not show significant differences. Nevertheless, Fig. 2a and b shows that filtration did not reduce the bacteria load to suitable values. Kawachi and Noël (2005) reported that this procedure is inefficient killing the most resistant spores, although it does kill most life in the water. Regarding the other sterilizing methods it was observed that all of them affected negatively the growth of the N. gaditana. The chemical methods probably alter the composition of the medium and the pasteurization could oxidize compounds due to the high reached temperatures (usually 95 °C). These changes in the medium composition could influence the microalga metabolism. In aquaculture it is usual to have large tanks containing seawater. In that case, flash sterilization is commonly carried out using a titanium plate heat exchanger that could be quite expensive. On the other hand, an ozone generator can also be costly but the ozone is able to achieve complete bacteria elimination in contrast to filtration. Accordingly, ozonation seems to be the most adequate method to be used for the sterilization of mediums to grow microalgae. As ozonation seems to be the best sterilizing method, the total organic carbon concentration (TOC) of the samples treated by ozonation and filtration was analyzed in order to compare them (Table 1). The results obtained show that the organic matter in the recirculated supernatant reaches a value of TOC of 73.7 mg/L. This organic matter is mostly composed of cellular fragments and to a lesser extent of other dissolved organic compounds. The dissolved organic compounds are those that are potentially first oxidized and degraded in the presence of strong oxidizing agents such as ozone. By this way, the application of 95 mg/L of ozone to the sample reduces the TOC in 33.5%. Applying higher concentrations of ozone it is possible to achieve higher TOC depurations, but it would affect the microalga growth, as previously described. When the sample is just filtered at 1.2 lm the TOC depuration reaches 50.4%, which is quite higher. If the filtered sample is afterwards ozonized it is possible to increase the TOC depuration up to 72.1% but this requires a very high ozone dosage (3133 mg/L), so this process would show low effectiveness. It is possible to assess that filtration is not enough to ensure a properly sterilization of the medium in spite of these tests and taking into account the bacteria load measured in the previous experiments carried out. Once the best sterilization method has been chosen it is necessary to evaluate the behavior of the cultures of N. gaditana growing in the ozonized medium. They were first grown in batch mode in 100 mL flasks using the ozonized recirculated supernatant as the culture medium. No negative effect on the growth was observed (Fig. 4a) as the obtained biomass productivities were over the one attained in the control flask, which used Algal medium (0– 100). The higher the proportion of supernatant used, the higher the biomass concentration, reaching values from 25% to 31% higher than in the control flask in the best case. That demonstrated that

Table 1 Results obtained from the recirculated supernatant treated with ozone and filtration to reduce the dissolved organic carbon. Treatment

Initial without treatment

Ozone 16 mg/L

Ozone 95 mg/l

Ozone 197 mg/L

Filtration 1.2 lm

Filtration 1.2 lm + Ozone 16 mg/L

Filtration 1.2 lm + Ozone 95 mg/L

Filtration 1.2 lm + Ozone 197 mg/L

Filtration 1.2 lm + Ozone 3133 mg/L

TOC; mg/L Depuration, %

73.7 ± 2.2 0.0

71.0 ± 2.1 3.8

49.0 ± 1.5 33.5

43.9 ± 1.3 40.5

36.6 ± 1.1 50.4

28.0 ± 0.8 62.0

27.4 ± 0.8 62.8

27.3 ± 0.8 62.9

20.6 ± 0.6 72.1

C.V. González-López et al. / Bioresource Technology 129 (2013) 430–438 0.12

(a)

Batch cultivation

YX/S

3.0

1.5

YS/X

(a)

YX/S, g biomass/g nitrates

Biomass Productivity, g/L/d

0.10

0.08

0.06

0.04

2.5

2.0

1.0

1.5

1.0

0.5

0.5

0.02

0.0

0.00

0.0 0-100

0-100

25-75

50-50

75-25

100-0

50-50

75-25

100-0

100*-0

100*-0

Recirculated supernatant-algal medium

0-100 50-50 75-25 100-0 100*-0

0.06

1.0

(b)

Continuous cultivation 0.3 1/d

(b) b)

0.05

0.8 0.04

qS, g/d/g

Biomass Productivity, g/L/d

YS/X, g nitrates/g biomass

436

0.6

0.03

0.02

0.4

0.01 0.2 0.00 Nitrates

Phosphates

K+

Mg+2

Sulphates

0.0

0-100

50-50

75-25

100-0

100*-0

the recirculated supernatant was still rich in nutrients that could be reused by N. gaditana or even it could consume organic matter dissolved in the supernatant. However, if the lacking nutrients of the recirculated supernatant are replenished (100⁄–0) the biomass concentration is 15% higher than if it is not replenished (100–0). The bacteria load was measured in all the cultures obtaining very low values, below 1000 CFUs/mL. Microalgae production plants usually operate in continuous mode. During the culture large amounts of water and nutrients are discarded. Therefore, it is necessary to design a closed system to recirculate the medium making possible to use the nutrients not ingested by the microalgae, as a strategy for water and nutrients saving which would involve a reduction in costs. Accordingly, after the batch cultures in flasks, 1.8 L bubble column photobioreactors were inoculated with N. gaditana and batch cultures were performed to have a concentration of biomass of 1 g/L. Then, the continuous culture was carried out at a dilution rate of 0.3 1/d.

Biochemical composition, % d.w.

Fig. 4. Mean values of biomass productivity (g/L d) of cultures tested at different percentages of recirculated supernatant in (a) batch culture (0–100, 25–75, 50–50, 75–25, 100–0, 100⁄–0 and (b) continuous culture at a dilution rate of 0.3 1/d (0– 100, 50–50, 75–25, 100–0, 100⁄–0).

Ash Carbohidrate Protein Lipid

60

Recirculated supernatant-algal medium

50

(c) c)

40

30

20

10

0 0-100

50-50

75-25

100-0

100*-0

Fig. 5. Nutrient yield (g biomass/g nitrates) and nutrient requirements (g nitrates/g biomass) for all tests carried out in continuous mode (0–100, 50–50, 75–25, 100–0, 100⁄–0) (a). Biomass-specific uptake rate (g/d/g) of key inorganic compounds (NO 3, 2 + +2 PO3 4 , K , Mg , SO4 ) for all tests (b). Biochemical composition for all tests (c).

using mixtures of new seawater culture medium (Algal medium) and recirculated supernatant at different proportions (see Section 2.4.2). The obtained results (Fig. 4b) show how N. gaditana can properly grow using a mixture 50–50 of supernatant and Algal

Table 2 Empirical formula of the biomass obtained in the steady state of the continuous cultures of N. gaditana at 0.3 1/d using different mixtures of recirculated supernatant and Algal medium. Test

0–100

50–50

75–25

100–0

100⁄–0

Empirical formula

C1H1.36N0.14O0.51

C1H1.05N0.14O0.51

C1H1.19N0.11O0.50

C1H1.22N0.09O0.42

C1H1.39N0.10O0.68

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Table 3 Fatty acids profile expressed as percentage on a biomass dry weight basis for the continuous cultures of N. gaditana at 0.3 1/d dilution rate using different mixtures of recirculated supernatant and Algal medium. Note: N.I.: not identified. Fatty acid

14:0 16:0 16:1n7 18:0 18:1n9 18:2n6 20:4n6 20:5n3 N.I. Total

Test 0–100

50–50

75–25

100–0

100⁄-0

1.231±0.047 3.902±0.150 3.442±0.132 0.140±0.005 0.861±0.033 0.411±0.016 0.792±0.030 2.702±0.104 2.333±0.089 15.865±0.608

0.665±0.033 4.712±0.236 4.816±0.241 0.105±0.005 0.645±0.032 0.228±0.011 0.744±0.037 3.140±0.157 1.759±0.088 18.189±0.909

1.097±0.055 6.922±0.346 6.772±0.309 0.106±0.005 0.878±0.044 0.544±0.027 1.122±0.056 3.373±0.199 2.331±0.117 23.145±1.157

1.061±0.053 8.051±0.403 6.960±0.348 0.211±0.011 1.328±0.066 0.378±0.019 0.956±0.048 3.440±0.172 2.018±0.101 25.622±1.281

1.543±0.027 5.755±0.102 4.430±0.078 0.195±0.003 1.219±0.022 0.462±0.008 0.929±0.016 2.707±0.048 2.466±0.044 19.707±0.348

medium reaching a biomass productivity near 0.8 g/L/d. Nevertheless, when the ratio of supernatant recirculation is increased (75– 25 and 100–0) the biomass productivity decreases to values of 0.4 g/L/d mainly because of the lack of nutrients in the medium. There are hardly any difference between the tests 75–25 and 100–0 probably because of the lack of any essential nutrient to the growth in both cases. On the other hand, when using only the recirculated supernatant after the replenishment of the consumed nutrients (100⁄–0) it was possible to achieve a biomass productivity near 0.8 g/L/d, as the one obtained with Algal medium (0–100) with no statistically significant differences between them. The use of mixtures of recirculated supernatant and Algal medium 50:50 v/v or the use of only recirculated supernatant after the replenishment of the lacking nutrients lead to a reduction of 50% or 100%, respectively, in water usage (water evaporation and water loss in harvesting are not considered). It also leads to a reduction in the addition of nutrients up to 50% or 15%, respectively. The elemental composition of the microalgal biomass harvested in the steady state of the continuous cultures was analyzed in order to establish the empirical formula (Table 2). This formula is quite similar whatever the composition of the culture medium used. When using pure Algal medium the obtained formula is C1H1.36N0.14O0.51. From the elemental analysis reported by Pan et al. (2010) it is possible to determine an empirical formula for Nannochloropsis sp. of C1H1.63N0.08O0.40, which is analogous to the one obtained in this work. Having established the empirical formula it is possible to calculate the nitrate yield (Y X=S ) and the nitrate requirement (Y S=X ). It was calculated for the different ratios of recirculated supernatant and Algal medium (0–100, 50–50, 75–25, 100–0 and 100⁄–0). It was shown (Fig. 5a) that the higher the ratio of recirculated supernatant to Algal medium the higher the yield coefficient for nitrate (YX/S) and the lower the requirement of nitrate (YS/X). This would lead to an increase in the content in lipids and a decrease in proteins. However, with the recirculated supernatant after replenishing the nutrients (100⁄–0) is more than 30% lower than the obtained when using Algal medium (0–100). The biomass-specific uptake rates of key inorganic nutrients, 3 2 + +2 such as NO 3 , PO4 , K , Mg , and SO4 , was determined in order to compare them (Fig. 5b). As shown, the recirculation of the supernatant can significantly reduce the uptake rate of nutrients. 3 + The major usages correspond to NO 3 , PO4 and K with values of 0.049, 0.017 and 0.014 g/d/g, respectively. In the test with the replenished supernatant (100⁄–0) the uptake rate of nitrate decreases in 50% with regard to the test with Algal medium. Nevertheless, the rate for the uptake of the other inorganic nutrients remains constant. The reduction of nutrients consumption (up to 50%) is not noteworthy in the context of aquafeeds as biomass production for this purpose is not large. Otherwise, this reduction would have an important impact with regard to the sustainability of the processes related to the production of commodities.

The biochemical composition of the biomass was analyzed to check if the use of the recirculated supernatant had a negative or a positive effect on the biomass quality (Fig. 5c). Total lipids, proteins, carbohydrates and ashes were measured for the biomass of the steady state of the continuous cultures. Moreover, the fatty acids profile was determined (Table 3). The biomass was composed of a higher amount of total fatty acids when employing the mediums 50–50 and 100⁄–0 compared to the one of the control (Algal medium, 0–100). In particular, it had 96% and 26% more fatty acids for the tests at 50–50 and 100⁄–0, respectively, than for the test at 0–100, attaining a maximum value of 30.2% d.w. for the 50–50 test. That was accompanied by an increase in the amount of reserve lipids such as 14:0, 16:0, 16:1n7 and 18:1n9. The content in structural lipids (as 20:4n6 and 20:5n3) did not change. The content in proteins of the biomass was similar for the test at 0–100 and 100⁄–0, but it was lower for the one at 50–50 because the metabolism of the microalgae was more displaced to the production of lipids due to the lack of nutrients. The same tendency was observed regarding the carbohydrates content (Fig. 5c). Therefore, the lack of nutrients (test 50–50 or 100–0) leads to an increase of 50% or 100%, respectively in total lipids and a decrease in the amount of ashes. That is in accordance with the accumulation of fatty acids by carbon availability in excess (Roessler, 1990; Tan and Johns, 1996; Wen and Chen, 2000). Rodolfi et al. (2003) assess that the recirculation of the medium reduces the biomass productivity in 50% and favors its contamination. On the other hand, the biomass productivity obtained after the recirculation of the medium in this work is similar to the achieved using Algal medium. Moreover, the biomass is of a high quality for aquaculture because of its high content in proteins and lipids (particularly, EPA).

4. Conclusions If mass production of microalgae is ever to come about it is necessary that the culture medium be reused. This work has shown that the supernatant obtained from harvested N. gaditana cultures can be reused after ozonation at 95 mgO3/L as this ensures a proper sterilization. It will allow reducing the nutrient costs, in a extent we expect to be able to quantify in future contributions, once the method is scaled-up. It has also been shown that N. gaditana tolerates the recirculation of the medium replenishing the depleted nutrients, and the obtained microalgal biomass is of high quality, suitable for aquaculture because of its high content in proteins (50%) and lipids (40%) with more than 3% EPA (d.w.).

Acknowledgements This research was supported by the General Secretariat of Universities, Research and Technology of Andalusian Government

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(AGR-5334) and was co-financed by FEDER funds. We would also like to thank the support of Fundación CAJAMAR. References AOAC International, 2002. Official Methods of Analysis of AOAC International, seventeenth ed. Association of Analytical Communities, Gaithersburg, MD, USA, 1st revision, pp. 770. Acién, F.G., Fernández, J.M., Magán, J.J., Molina, E., 2012. Production cost of a real microalgae production plant and strategies to reduce it. Biotechnol. Adv. 30, 1344–1353. Aziz, J.A., Tebbut, T.H.Y., 1980. Significance of COD, BOD and TOC correlations in kinetic models of biological oxidation. Water Res. 14, 319–324. Benemann, J., 1992. Microalgae aquaculture feeds. J. Appl. Phycol. 4, 233–245. Brown, K.J., 1980. A method for turbidimetric measurement of bacterial growth in liquids cultures and agar plug diffusion cultures, using standard test tubes. Eur. J. Appl. Microbiol. Biotechnol. 9, 59–62. Eaton, A.D., Clesceri, L.S., Greenberg, A.E., 1995. Standard Methods for the Examination of Water and Wastewater, nineteenth ed. American Public Health Association, Washington D.C. Grotmol, S., Totland, G.K., 2000. Surface disinfection of Atlantic halibut (Hippoglossus hippoglossus) eggs with ozonated sea-water inactivates nodavirus and increases survival of the larvae. Dis. Aquat. Org. 39, 89–96. Jung, I.S., Lovitt, R.W., 2010. Integrated production of long chain polyunsaturated fatty acids (PUFA) rich Schizochytrium biomass using a nutrient supplemented marine aquaculture wastewater. Aquacult. Eng. 43, 51–61. Kawachi, M., Nöel, M.H., 2005. Sterilization and Sterile Techniques. In: Hellebust, J., Cragie, S. (Eds.), Algal Culturing Techniques Book. 5, 65–75. Cambridge University Press, London, United Kingdom, p. 189. Kochert, G., 1978. Quantitation of the macromolecular components of microalgae. Handbook of Phycological methods. Physiological and Biochemical methods. Ozone in water treatment application and engineering. Co-operative research report. Langlais, B., Reckhow, D.A., Brink, D.R. (Eds.), 1991. Ozone in Water Treatment Application and Engineering. Co-operative Research Report. CRC Press LLC, Florida, USA. Li, Y., Chen, Y.-F., Chen, P., Min, M., Zhou, W., Martínez, B., Zhu, J., Ruan, R., 2011. Characterizacion of a microalga Chlorella sp. Well adapted to highly concentrated municipal wastewater for nutrient removal and biodiesel production. Bioresour. Technol. 102, 5138–5144.

López, C.V., Cerón, M.C., Acién, F.G., Segovia, C., Chisti, Y., Fernández, J.M., 2010. Protein measurements of microalgal and cyanobacterial biomass. Bioresour. Technol. 101, 7587–7591. Martínez, S.B., Pérez-Parra, J., Suay, R., 2011. Use of ozone in wastewater treatment to produce water suitable for irrigation. Water Resour. Manage. 25, 2109–2124. Morais, S., Koven, W., Ronnertad, I., Conceicao, L.E.C., 2005. Lipid enrichment for Senegal sole (Soleasenegalensis) larvae: effect on growth survival and fatty acid profile. Aquac. Int. 12, 281–298. Navarro, J.C., Villanueva, R., 2003. The fatty acid composition of Octopus vulgaris par larvae reared with live and inert food: deviation from their natural fatty acid profile. Aquaculture 1–4, 613–631. Pan, P., Hu, C., Yang, W., Li, Y., Dong, L., Zhu, L., Tong, D., Qing, R., Fan, Y., 2010. The direct pyrolysis and catalytic pyrolysis of Nannochloropsis sp. Residue for renewable bio-oils. Bioresour. Technol. 101, 4593–4599. Rathore, A.S., Shirke, A., 2011. Recent developments in membrane-based separations in biotechnology processes: review. Prep. Biochem. Biotechnol. 41, 398–421. Rhodes, M.A., Sedlacek, C., Marcus, N.H., 2008. Large-scale batch algae cultivation: comparison of two seawater sterilization techniques. Aquac. Res. 39, 1128– 1130. Rodolfi, L., Chini Zitelli, G., Barsanti, L., Rosati, G., Tredici, M.R., 2003. Growth medium recycling in Nannochloropsis sp. Mass cultivation. Biomolec. Eng. 20, 243–248. Rodríguez-Ruiz, J., Belarbi, E., Sánchez, J.L.G., Alonso, D.L., 1998. Rapid simultaneous lipid extraction and transesterification for fatty acid analyses. Biotechnol. Tech. 12, 689–691. Roessler, P.G., 1990. Environmental control of glycolipid metabolism in microalgae: commercial implications and future research directions. J. Phycol. 26, 393– 399. Saral, A.B., Goncalog, B.I., 2008. Determination of real COD in highly chlorinated wastewaters. Clean 36, 996–1000. Tan, C.K., Johns, M.R., 1996. Screening of diatoms for heterotrophic eicosapentaenoic acid production. J. Appl. Phycol. 8, 59–64. Tyrrell, S.A., Ryppey, S.R., Watkins, W.D., 1995. Inactivation of bacterial and viral indicators in secondary sewage effluents using chlorine and ozone. Water Res. 29, 2483–2490. Wen, Z.Y., Chen, F., 2000. Production potential of eicosapentaenoic acid by the diatom Nitzschia. Biotechnol. Lett. 22, 727–733. Yang, J., Xu, M., Zhang, X., 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.