- Email: [email protected]
A comparison of harvesting and drying methodologies on fatty acids composition of the green microalga Scenedesmus obliquus
Biomass and Bioenergy 132 (2020) 105437
Contents lists available at ScienceDirect
Biomass and Bioenergy journal homepage: http://www.elsevier.com/locate/biombioe
Research paper
A comparison of harvesting and drying methodologies on fatty acids composition of the green microalga Scenedesmus obliquus Carlos Yure Barbosa de Oliveira a, *, Thayna Lye Viegas a, Rafael Garcia Lopes a, Herculano Cella a, Rafael Silva Menezes b, Aline Terra Soares b, Nelson Roberto Antoniosi Filho b, Roberto Bianchini Derner a a b
Laboratory of Algae Cultivation, Aquaculture Department, Federal University of Santa Catarina (UFSC), 88061-600, Florian� opolis, Brazil Laboratory of Extraction and Separation Methods, Chemistry Institute, Federal University of Goias (UFG), 74690-900, Goi^ ania, Brazil
A R T I C L E I N F O
A B S T R A C T
Keywords: Flocculation Biomass recovery Biomass drying Saturated fatty acids Biodiesel
Harvesting and drying processes are still obstacles in the microalgae production chain. In this study, a combi nation of different harvesting and drying methods for Scenedesmus obliquus was proposed. S. obliquus was cultivated in a pond raceway for 15 days until reaching stationary phase. The culture was separated by different harvesting methods (centrifugation and flocculation) and drying methods (freezing, freeze-drying and oven drying), each combination in triplicate. Flocculation did not influence FAME yield of S. obliquus, except when the biomass was dried in an oven. The biomass that was only frozen had the lowest FAME content due to the presence of reactive water in the biomass. In contrast, frozen biomass had higher content of saturated and monosaturated fatty acids; freeze-drying or oven drying caused an enrichment of the polyunsaturated fatty acids content. In conclusion, if the biomass will be used to extract polyunsaturated fatty acids, freeze-drying would be more appropriate. On the other hand, if the purpose of the biomass is to produce biodiesel, the best process would be to just freeze the biomass.
1. Introduction
standards requirements [5]. Unfortunately, strains with high oil contents in their biomass often present low growth rates [6]. Therefore, developing large scale pro duction techniques for strains that have satisfactory growth rates and high lipid content is one of the challenges to the feasibility of industrial production of microalgae biodiesel. In this context, Scenedesmus obliquus is considered an organism with great potential [5] because it has fea tures that can facilitate massive outdoor production such as resistance to high temperatures and pH, and high growth rates [7,8]. The technical-economic feasibility of using microalgae biomass as raw material for biodiesel production presents other challenges, mainly in the stages of harvesting, drying and storage [9]. Microalgae cultures can be harvested by several individual methods, or by a combination of methods (e.g. flocculation plus centrifugation), and although they are important steps in the microalgae biomass production chain, clear har vesting and storage protocols have not yet been established [10]. The microalgae biomass harvesting and drying processes influence the costs and quality of the final products [11]. Generally, centrifugation is the most efficient harvesting method and it has been used in most pilot-scale
Microalgae oils have advantages over animal fats and higher plant oils for the production of biodiesel, since microalgae can be grown in areas unsuitable for agriculture [1], and liquid and gaseous effluents can be used for microalgae production [2]. In addition, microalgae biomass contains other high added-value bioproducts, such as vitamins, pig ments and amino acids, with potential use in several industrial segments and whose sale can help compensate for the production cost of biofuels made from microalgae [3]. The oil content of the most frequently studied microalgae species varies from 20 to 30% of dry weight [1]. Among the lipid classes, fatty acids are a group of major economic interest, and they are mainly classified according to its unsaturation levels. Different fatty acids classes might be used for different purposes, e.g., mono and saturated fatty acid esters are preferable to biodiesel production, while poly unsaturated fatty acids are considerate essential to animal diet and present bioactive properties [4]. So, besides showing a reasonable lipidic content, the fatty acid profile must also meet the international
* Corresponding author. E-mail addresses: [email protected], [email protected] (C.Y.B. Oliveira). https://doi.org/10.1016/j.biombioe.2019.105437 Received 25 May 2019; Received in revised form 23 November 2019; Accepted 27 November 2019 Available online 9 December 2019 0961-9534/© 2019 Elsevier Ltd. All rights reserved.
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
productions [12]. However, centrifuging is often economically unviable on an industrial scale due to high operating costs [13]. Flocculation has been a convenient separation method for concen trating large volumes of microalgae [14]. Although it is still necessary to filter or centrifuge the biomass after flocculation, the fact that this process concentrates the crop at a considerably lower volume can reduce the time and energy costs associated with separation [15]. The biomass harvesting and drying processes must be simple, fast, low cost and have high efficiency in the preservation of the biochemical quality of the biomass [16]. Although much has been studied about biomass harvesting; little is known of the impacts caused by the different drying methods, and less about the harvesting and drying interaction [12,16–18]. Determining the methodology that provides the best performance in harvesting and drying without qualitative damages to the algal biomass is important to establishing the production chain of any bioproduct derived from microalgae. For this reason, this study evaluated the influence of har vesting and drying methods on the yield and profile of fatty acid methyl esters (FAME) extracted from the green microalga Scenedesmus obliquus.
20 � C and stored until the moment of the next analysis. The frozen biomass remained at this temperature until the direct transesterification step. 2.2.4. Freeze-drying The samples subjected to the freeze-drying process were frozen to 20 � C, and then freeze-dried (L101 model, Liobras, Brazil), at 55 � C and 0.13 mbar, over 48 h. 2.2.5. Oven-drying The process in the oven-drying occurred with continuous forced air flow at 45 � C in 48 h. 2.3. Direct transesterification of microalgal biomass Approximately 200 mg of microalgal biomass was weighed. Next, 3.0 mL of 0.5 mol L 1 sodium hydroxide solution (MERCK ®) in dry methanol (TEDIA ®) was added, and the test tube was heated for 10 min in a 90 � C water bath. The test tube was cooled in an ice bath, and 9.0 mL of the mixture previously prepared for esterification was added ac cording to Soares et al. [3]. The test tube was heated again for 10 min in a 90 � C water bath. The test tube was cooled in an ice bath, and 5.0 mL of n-heptane (TEDIA ®) and 2.0 mL of distilled water were added and was put to rest by 2 h. The test tubes were shaken again for 1 min at 3,000 rpm and allowed to stand for 24 h until phase separation. The assays were performed in triplicate. The heptanic phases were collected with a Pasteur pipettes, inserted into 1.5 mL vials and analyzed by gas chro matography. The extraction procedure was performed in triplicate. To determine the FAME content from S. obliquus, a graph was created that correlated the sum of the peak area of the FAME (Y) of the soybean oil as a function of soybean oil mass (X) used in the FAME production through the transesterification of the oil, as the method described above. The correlation coefficient value was 0.999 for the calibration curve.
2. Material and methods 2.1. Microalga and growth conditions Scenedesmus obliquus strain was provided by the Culture Center of the Laboratory of Algae Cultivation, Brazil and maintained in LCA-AD (adapted from Bold’s Basal Medium) containing: 1.0 g L 1 NaNO3; 0.75 g L 1 MgSO4 7H2O; 0.6 g L 1 KH2PO4; 0.05 g L 1 EDTA-Na2; 0.03 g L 1 KOH; 0.25 g L 1 K2HPO4; 0.25 g L 1 NaCl; 0.25 g L 1 CaCl2 2H2O; 0.11 g L 1 H3BO3; 0.05 g L 1 FeSO4 7H2O; 0.014 g L 1 MnCl 4H2O; 0.016 g L 1 CuSO4 5H2O; 0.00071 g L 1 MoO3; 0.0004 g L 1 Co(NO3) 6H2O. S. obliquus was maintained in 2-L borosilicate glass flasks at 22 � 1 � C, 560 μmol photons m 2 s 1 irradiance with continuous lighting and stirred by bubbling air enriched with 0.5% CO 2. 200-L inoculum of S. obliquus was transferred to an outdoor raceway tank containing 2,000-L of culture medium. S. obliquus was cultured for 15 days, at outdoor temperatures (24 � 4 � C), with an average of 1,000 μmol photons m 2 s 1 of natural irradiance during the illuminated period, with a 12:12 photoperiod and agitated by a continuous flow of 0.5 m s 1. Cultivation was interrupted in the stationary phase when biomass reached 0.5 g L 1. The biomass obtained was fractionated in the different treatments according to the harvesting methodology: centrifugation; flocculation; and subsequent drying: frozen biomass only; freeze-dried and; oven dried, corresponding to a two-factorial design (2 � 3), in triplicate.
2.4. Gas chromatographic analysis An Agilent 7890 gas chromatograph equipped with a Flame Ioniza tion Detector (FID) and a split/splitless injector was used to analyze the FAME composition. The capillary column used was the DB-WAX (30 m � 0.25 mm � 0.25 μm). The initial oven temperature was 70 � C, and it was heated at a rate of 10 � C min 1 to 240 � C and maintained at this temperature for 13 min. It was heated again at a rate of 5 � C min 1 to 250 � C. The injector temperature was kept at 310 � C in split mode with a split ratio of 10:1 for an injection volume of 2 μL. The oven temperature was maintained at 310 � C. Hydrogen (5.0) was used as the carrier gas at a linear velocity of 42 cm s 1, and nitrogen was used as the auxiliary gas at a rate of 20 mL min 1. The FAMEs were identified by comparison with the retention times of samples of known composition such as soybean, peanut, and crambe (Crambe abyssinica) oils, by analyzing FAME reference standards (NuCheck-Prep ®) and by gas chromatography coupled with high resolution mass spectrometry (GC-HRMS) using a Shimadzu model 17A chro matograph coupled with a Shimadzu QP-5050 mass spectrometer. The carrier gas was helium with a linear velocity of 42 cm s 1. The GC-FID operating conditions (oven, injector, interface, and capillary column) were maintained for GC-HRMS [19].
2.2. Harvesting and drying methods 2.2.1. Centrifugation Three biomass samples, each in triplicate, were concentrated using an industrial centrifuge (SSD-06 model, GEA-Westfalia, Germany) in continuous processing (12,000 rpm). 2.2.2. Flocculation Since charge neutralization is the basis of flocculation and the elec trical charge of the microalgae cell wall is negative, cationic flocculants are generally more effective than anionic flocculants. For this reason, cationic polyacrylamide FLOPAM ® (courtesy of Professor Paulo Abreu, Federal University of Rio Grande) was used as the flocculating agent, with a medium charge density and high molecular weight, at 10 mg L 1 final concentration [9]. The biomass concentrated in this process was centrifuged using the same centrifuge and rotation used in the previous treatments.
2.5. Statistical analysis Two-way analysis of variance (ANOVA), followed by the Tukey’s post-hoc test was applied when necessary to compare the fatty acids profile among the harvesting and storage methodologies. Principal component analysis (PCA) was performed to determine possible patterns among the harvesting and drying combinations, using a correlation to produce a cross product matrix [20]. All statistical analyses were
2.2.3. Freezing Samples from both harvesting processes were immediately frozen at 2
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
Fig. 1. Fatty acid methyl esters content (mean � SD) of Scenedesmus obliquus harvested and dried by different methodologies. Different letters represent significant differences among the drying methods; * represent significant differences between the harvest methods, by Tukey’s post-hoc test.
performed at 5% significance level using RStudio® software version 3.5.1 (RStudio Inc, USA).
oven drying treatments was 3.2 � 0.2%; and for the biomass that was not dried (freezing), the moisture content was 64.5 � 12.37%. Thus, if the average moisture content contained in the frozen biomass is dis regarded, the FAME yield per gram of dry biomass would be approxi mately 119.41 mg g 1, similar to that found for the treatments where the biomass was dried. However, the removal of water from the reaction medium favors the transesterification reaction, since the acidic and basic catalysts will interact more effectively with the microalgae biomass. The presence of water may lead to a decrease in the base and acid concentrations present in the esterification mixture, since these solutions are in methanol, which interacts with water from the medium. Moreover, oils can be difficult to extract using solvent from wet biomass [21], especially when a cell wall disruption strategy is not adopted. These characteristics may be related to the different morphological types of microalgae cell walls. Indeed, for the diatom Isochrysis galbana, the lipids were also easily extracted when the biomass was freeze-dried [22]. Cationic polymers, polyacrylamide based like FLOPAM, which was used in the present study, act as coagulants, neutralizing part of the negative charges on microalgae cell walls at neutral pH and thus, it re duces the cellular repulsion. After the flocculation occurs, the resulting
3. Results and discussion 3.1. Direct transesterification of microalgal biomass Fig. 1 shows the direct transesterification of Scenedesmus obliquus biomass from the different combinations of harvesting and drying methods. The percentage of FAME extracted from the dry biomass of S. obliquus varied between 30 mg g (moist weight) 1, for the biomass that underwent flocculation and freezing, and 140 mg g (dry weight) 1, for the biomass that underwent centrifugation and freeze-drying, for the minimum and maximum, respectively. However, no significant differ ences were observed between the centrifugation plus freeze-drying, flocculation plus freeze-drying and centrifugation plus oven-drying combinations. The flocculation only had a negative influence (p < 0.05) when the biomass was dried in an oven. For the other treatments, flocculation using cationic polyacrylamide did not influence the direct transesterification of S. obliquus. The moisture content in the biomass that underwent freeze-drying or
Fig. 2. Fatty acids classes (in % FAME) of the Scenedesmus obliquus for the different combinations of biomass harvesting and drying methodologies. 3
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
flocs were also centrifuged (same conditions of the other treatments) and since the flocculation does not negatively impact the cell composi tion [23], it was expected that the FAME profile would not be affected. However, the combination of flocculation plus oven drying resulted in a low FAME yield, as it was observed that dry biomass had a thin layer of the flocculant. This might have acted as a physical barrier, not allowing an efficient extraction of the non-polar fraction of S. obliquus biomass with the use of solvents. The high temperatures of the oven may have favored the binding of the polymers with the cell walls, making it difficult to extract the lipids [11,24]. This is, supposedly, the reason to the differences found in FAME concentration between centrifugation and flocculation with posterior oven drying, since in the absence of the flocculant agent, FAME concentration was not impacted. For the higher FAME yield, the results found in this study were similar to those reported by El-Sheekh et al. [25], about 150 mg g 1. However, S. obliquus can reach higher FAME contents, of 340 mg g 1, when grown under controlled conditions [26]. Although the results re ported here were similar to other studies, the production of S. obliquus in an outdoor raceway tank possibly influenced the FAME concentration, mostly due to oscillations of environmental parameters that occur in open systems.
Table 1 Fatty acids profile of Scenedesmus obliquus for the different combinations of biomass harvesting and drying methodologies. Fatty acid
Centrifugation Freezing
Freezedrying
Ovendrying
Freezing
Freezedrying
Ovendrying
C14:0
1.13 � 0.06 24.97 � 0.55a 13.63 � 0.21a 2.07 � 0.11a 7.57 � 0.21a 0.37 � 0.11 n.d.b
1.07 � 0.38 12.5 � 0.79b 3.30 � 0.10b 1.40 � 0.00b 3.53 � 0.21b 0.20 � 0.01 0.13 � 0.06a 0.87 � 0.06a
1.57 � 0.50 29.5 � 1.39a 11.17 � 2.97a 2.93 � 0.21a 8.47 � 0.60a 0.27 � 0.06 n.d.b
0.50 � 0.00b*
0.8 � 0.01 11.67 � 0.25b 3.4 � 0.20b 1.23 � 0.06c 3.07 � 0.15b 0.13 � 0.06 0.13 � 0.06a 0.87 � 0.06a
1.03 � 0.11 14.07 � 1.27b 3.53 � 0.90b 1.77 � 0.11b 3.67 � 0.38b 0.30 � 0.26 0.20 � 0.01a 1.10 � 0.10a
1.00 � 0.00 13.77 � 0.38b 3.57 � 0.21b 1.90 � 0.01b 3.97 � 0.06b 0.67 � 0.06 0.17 � 0.06a 1.10 � 0.00a
5.87 � 0.67b
21.17 � 0.64a
20.43 � 1.01a
3.40 � 0.44b
17.97 � 2.58a
19.5 � 0.61a
1.10 � 0.00a 10.03 � 0.58a* 3.03 � 0.06a* 3.73 � 0.06 0.20 � 0.01b
0.40 � 0.01c 4.83 � 0.11b* 1.4 � 0.01b* 4.17 � 0.11 0.30 � 0.00a
0.67 � 0.06a 5.23 � 0.40b* 1.60 � 0.10b* 4.20 � 0.20 0.30 � 0.00a
1.07 � 0.15a 12.53 � 0.25a 4.10 � 0.14a 4.13 � 0.11 n.d.b
0.43 � 0.06c 6.43 � 0.58b 2.07 � 0.21b 4.83 � 0.23 0.30 � 0.00a
0.63 � 0.06b 6.17 � 0.15b 2.13 � 0.06b 4.50 � 0.17 0.30 � 0.00a
22.27 � 0.75b
41.17 � 0.15a
25.10 � 0.75b
17.57 � 0.57c
25.1 � 21.316b
36.13 � 0.50a
1.53 � 0.11b
4.37 � 0.11a*
4.13 � 0.25a
0.90 � 0.10b
3.70 � 0.36a*
3.60 � 0.10a
1.10 � 0.00a 0.90 � 0.00a 6.23 � 0.31a
0.43 � 0.06b 0.30 � 0.00b 3.97 � 0.21b
0.47 � 0.15b* 0.37 � 0.06b* 4.37 � 0.38b
1.43 � 0.64 0.90 � 0.00a 6.67 � 0.76
1.67 � 1.94 0.40 � 0.10b 6.69 � 3.14
1.03 � 0.06 n.d.b
C16:0 C16:1 cis 9 C16:1 C16:1 cis 11 C16:2 cis 7,10 C17:0 C16:3 cis 6, 9, 12 C16:4 cis 6, 9, 12, 15 C18:0
3.2. FAME profile of S. obliquus Fig. 2 shows the fatty acids classes for the different combinations of harvesting and drying. The biomass that was not dried (directly frozen after harvesting) showed the best profile for biodiesel production. The polyunsaturated fatty acids percentage nearly doubled when the biomass was dried in a freeze-dryer or oven. The content of diunsatu rated fatty acids remained close to 4% regardless of the combination used. The harvesting methodology did not influence significant modi fications in the fatty acid classes, since once these possible cell disrup tions occurred by the centrifugation forces, followed by possible intracellular material leakage, were present in all treatments. Drying (and storage) methodologies represent an obstacle in the microalgae biomass production chain. In addition to freeze-drying and oven drying, spray drying, sun drying, and drum drying have already been studied [17]. The correct selection of the drying method to produce biodiesel from microalgae is an important step still to be taken [18]. The use of freeze-dryers has already been ruled out for large-scale production because of the high operating costs. The findings reported in this study represent an important advance, especially considering that the highest saturated and monosaturated fatty acids levels were found when the biomass was not dried. This can represent great cost and time saving in the microalgae biodiesel production chain [27]. Drum-drying, spray-drying or freeze-drying of Dunaliella, β-carotenerich, produced satisfactory results in dry biomass uniformity and sta bility of β-carotene. The influence of the drying method on the compo sition and amount of the metabolite is probably linked to molecule stability [28], cell wall composition, resistance [29] and microalgal cell integrity [30]. It should also be emphasized that if the goal is the re covery of polyunsaturated fatty acids (those with nutraceutical proper ties), freeze-drying has considerably increased the levels of these fatty acids. In the extraction of bioactive compounds from S. subspicatus, the use of water was more efficient than that of potentially toxic solvents [31]. Possible interactions among saturated and monounsaturated fatty acids with polar solvents (such as water) have not yet been established. However, in this study, the extracellular water in the frozen biomass preserved a more saturated and monosaturated fatty acid profile. The use of oven drying resulting from harvesting processes can cause pyrolysis of intracellular organic compounds [32]. Although the decomposition of volatiles, such as carbohydrates and lipids, usually occurs at higher temperatures (around 120–500 � C) in microalgae, the exposure of S. obliquus to 45 � C for 48 h may have oxidized some of the saturated and monosaturated fraction. The fatty acid profiles (Table 1) of S. obliquus had higher levels of
C18:1 cis 9 C18:1 cis 11 C18:2 cis 9, 12 C18:3 cis 6, 9, 12 C18:3 cis 9, 12, 15 C18:4 cis 6, 9, 12, 15 C22:0 C24:0 Others
Floculation
n.d.b
5.67 � 0.11
Different letters represent significant differences among the drying methods; * represent significant differences between the harvest methods, by Tukey’s posthoc test. n.d. not detected.
Linolenic (C18:3 cis 9, 12, 15), Palmitic (C16:0) and Hexadecatetraenoic (C16:4 cis 6, 9, 12, 15) acids. Again, no significant differences were observed between the biomass harvesting methods. However, individual FAME differed significantly between the drying methods. The highest Linolenic percentage (the most abundant fatty acid) was reported for the centrifugation plus freeze-drying and centrifugation plus oven-drying combinations. The Palmitic acid concentrations were significantly higher in the treatments in which the biomass was frozen soon after being concentrated (centrifuged or flocculated) (Table 1). However, the drying of the biomass, by both freeze- and oven-drying, may have caused the biomass to oxidize due to the loss of free water, which could lead to an increase in saturated and monounsaturated fractions of FAME. This could explain why higher content of poly unsaturated fatty acids was observed in the treatments of biomass with drying than in those without drying. PCA ordination (F ¼ 41.76; p ¼ 0.002) for 6 combinations of har vesting and drying methodologies, explained 99.34% of data variability in the first two components (Fig. 3). Polyunsaturated (r ¼ -0.552), 4
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
Fig. 3. Principal components analysis (PCA) ordination diagram for the combinations of harvesting (CEN centrifugation, FLO flocculation) and drying (FZ freezing, FD freeze-drying and OD oven-drying) methodologies.
Monounsaturated (r ¼ 0.553) and Saturated (r ¼ 0.549) were the principal fatty acids in the composition of component 1 (79.60%), while Diunsaturated (r ¼ 0.95) was the most important fatty acid for compo nent 2 (19.74%). The diagram for PCA indicates a clear distinction be tween the treatments that were only frozen, independent of having been harvested by centrifugation or flocculation, in relation to mono- and saturated fatty acids. Even with the low polyunsaturated fatty acids content, the oil of S. obliquus presents restrictions to the production of biodiesel that at tends European regulation EN 14214 (which allows only 1% of fatty acids with more than three double bonds). Choricystis minor var. minor was considered a promising species for the production of biodiesel because of its fatty acid profile [33]. However, the linolenic acid content reported by Menezes et al. [19] was greater than 12% (the maximum limit acceptable to EN 14214) using centrifugation plus freeze-drying as biomass harvesting and drying methods, respectively. In this study, this combination showed a close to 100% increase in the linolenic acid content when compared to biomass that was not dried (only frozen). However, it is worth noting that the combination of harvesting and drying methodologies may present different performances in other species, due to the specific characteristics already mentioned.
flocculation and oven-drying reduced the FAME yield. Human and animal rights The study did not involve human subjects or animal models. Author contributions CYBO and TLV analyzed the data. RGL, HC and ATS conducted the experiments. All the authors read and edited the manuscript and pro vided valuable inputs to improve the quality of the article. Declaration of Competing interest The authors declare that they have no competing interests. Acknowledgment The authors would like to thank the Ministry of Science, Technology and Innovation (MCTI) for financial support provided by FINEP (Agreement No. 01.10.0457.00), Conselho Nacional de Desenvolvi �gico (Case No. 574796/2008-8), Coor mento Científico e Tecnolo ~o de Aperfeiçoamento de Pessoal de Nível Superior Brazil denaça (Finance Code 001).
4. Conclusions The two harvesting methodologies that were analyzed in this study did not affect the fatty acid profile of Scenedesmus obliquus. In contrast, the fatty acid content did change when they were dried using different methodologies. If the goal is to increase the polyunsaturated fatty acids content, the biomass should be freeze-dried or oven-dried. On other hand, if the biomass will be used to produce biodiesel, the biomass should be frozen after harvesting. Finally, the combination of
References [1] Y. Chisti, Biodiesel from microalgae, Biotechnol. Adv. 25 (2007) 294–306, https:// doi.org/10.1016/j.biotechadv.2007.02.001. [2] C.Y.B. Oliveira, J.L. Abreu, C.D.L. Oliveira, P.C. Lima, A.O. G� alvez, D.M.M. Dantas, Growth of Chlorella vulgaris using wastewater from Nile tilapia (Oreochromis niloticus) farming in a low-salinity biofloc system, Acta Sci. Technol. 42 (2020), e46232, https://doi.org/10.4025/actascitechnol.v42i1.46232.
5
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
[3] A.T. Soares, D.C. da Costa, B.F. Silva, R.G. Lopes, R.B. Derner, N.R. Antoniosi Filho, Comparative analysis of the fatty acid composition of microalgae obtained by different oil extraction methods and direct biomass transesterification, Bioenergy Res. 7 (2014) 1035–1044, https://doi.org/10.1007/s12155-014-9446-4. [4] B. Ramesh Kumar, G. Deviram, T. Mathimani, P.A. Duc, A. Pugazhendhi, Microalgae as rich source of polyunsaturated fatty acids, Biocatal. Agric. Biotechnol. 17 (2019) 583–588, https://doi.org/10.1016/j.bcab.2019.01.017. [5] S. Mandal, N. Mallick, Microalga Scenedesmus obliquus as a potential source for biodiesel production, Appl. Microbiol. Biotechnol. 84 (2009) 281–291, https://doi. org/10.1007/s00253-009-1935-6. [6] C. Dayananda, R. Sarada, M. Usha Rani, T.R. Shamala, G.A. Ravishankar, Autotrophic cultivation of Botryococcus braunii for the production of hydrocarbons and exopolysaccharides in various media, Biomass Bioenergy 31 (2007) 87–93, https://doi.org/10.1016/j.biombioe.2006.05.001. [7] M.G. de Morais, J.A.V. Costa, Biofixation of carbon dioxide by Spirulina sp. and Scenedesmus obliquus cultivated in a three-stage serial tubular photobioreactor, J. Biotechnol. 129 (2007) 439–445, https://doi.org/10.1016/j. jbiotec.2007.01.009. [8] T. de Marchin, M. Erpicum, F. Franck, Photosynthesis of Scenedesmus obliquus in outdoor open thin-layer cascade system in high and low CO2 in Belgium, J. Biotechnol. 215 (2015) 2–12, https://doi.org/10.1016/j.jbiotec.2015.06.429. [9] R.B. K€ onig, R. Sales, F. Roselet, P.C. Abreu, Harvesting of the marine microalga Conticribra weissflogii (Bacillariophyceae) by cationic polymeric flocculants, Biomass Bioenergy 68 (2014) 1–6, https://doi.org/10.1016/j. biombioe.2014.06.001. [10] T.M. Mata, A.A. Martins, N.S. Caetano, Microalgae for biodiesel production and other applications: a review, Renew. Sustain. Energy Rev. 14 (2010) 217–232, https://doi.org/10.1016/j.rser.2009.07.020. [11] L. Borges, J.A. Mor� on-Villarreyes, M.G.M. D’Oca, P.C. Abreu, Effects of flocculants on lipid extraction and fatty acid composition of the microalgae Nannochloropsis oculata and Thalassiosira weissflogii, Biomass Bioenergy 35 (2011) 4449–4454, https://doi.org/10.1016/j.biombioe.2011.09.003. [12] A.J. Dassey, C.S. Theegala, Harvesting economics and strategies using centrifugation for cost effective separation of microalgae cells for biodiesel applications, Bioresour. Technol. 128 (2013) 241–245, https://doi.org/10.1016/j. biortech.2012.10.061. [13] J.J. Milledge, S. Heaven, A review of the harvesting of micro-algae for biofuel production, Rev. Environ. Sci. Biotechnol. 12 (2013) 165–178, https://doi.org/ 10.1007/s11157-012-9301-z. [14] R.M. Knuckey, M.R. Brown, R. Robert, D.M.F. Frampton, Production of microalgal concentrates by flocculation and their assessment as aquaculture feeds, Aquacult. Eng. 35 (2006) 300–313, https://doi.org/10.1016/j.aquaeng.2006.04.001. [15] R. Sales, P.C. Abreu, Use of natural pH variation to increase the flocculation of the marine microalgae Nannochloropsis oculata, Appl. Biochem. Biotechnol. 175 (2014) 2012–2019, https://doi.org/10.1007/s12010-014-1412-2. [16] M.K. Danquah, L. Ang, N. Uduman, N. Moheimani, G.M. Forde, Dewatering of microalgal culture for biodiesel production: exploring polymer flocculation and tangential flow filtration, J. Chem. Technol. Biotechnol. 84 (2009) 1078–1083, https://doi.org/10.1002/jctb.2137. [17] E. Molina Grima, E.H. Belarbi, F.G. Aci�en Fern� andez, A. Robles Medina, Y. Chisti, Recovery of microalgal biomass and metabolites: process options and economics, Biotechnol. Adv. 20 (2003) 491–515, https://doi.org/10.1016/S0734-9750(02) 00050-2. [18] K.Y. Show, D.-J. Lee, A.S. Mujumdar, Advances and challenges on algae harvesting and drying, Dry. Technol. 33 (2015) 386–394, https://doi.org/10.1080/ 07373937.2014.948554.
[19] R.S. Menezes, A.T. Soares, J.G. Marques Júnior, R.G. Lopes, R.F. da Arantes, R. B. Derner, N.R.A. Filho, Culture medium influence on growth, fatty acid, and pigment composition of Choricystis minor var. minor: a suitable microalga for biodiesel production, J. Appl. Phycol. 28 (2016) 2679–2686, https://doi.org/ 10.1007/s10811-016-0828-1. [20] C.Y.B. Oliveira, C.D.L. Oliveira, A.J.G. Almeida, A.O. G� alvez, D.M. Dantas, Phytoplankton responses to an extreme drought season: a case study at two reservoirs from a semiarid region, Northeastern Brazil, J. Limnol. 78 (2) (2019) 176–184, https://doi.org/10.4081/jlimnol.2019.1869. [21] E.H. Belarbi, E. Molina, Y. Chisti, A process for high yield and scaleable recovery of high purity eicosapentaenoic acid esters from microalgae and fish oil, Enzym. Microb. Technol. 26 (7) (2000) 516–529. [22] E.M. Grima, A.R. Medina, A.G. Gim� enez, J.A. S� anchez P�erez, F.G. Camacho, J. L. García S� anchez, Comparison between extraction of lipids and fatty acids from microalgal biomass, J. Am. Oil Chem. Soc. 71 (1994) 955–959, https://doi.org/ 10.1007/BF02542261. [23] R. Sales, R.B. Derner, M.Y. Tsuzuki, Effects of different harvesting and processing methods on Nannochloropsis oculata concentrates and their application on rotifer Brachionus sp. cultures, J. Appl. Phycol. (2019), https://doi.org/10.1007/s10811019-01877-8. [24] N. Uduman, Y. Qi, M.K. Danquah, G.M. Forde, A. Hoadley, Dewatering of microalgal cultures: a major bottleneck to algae-based fuels, J. Renew. Sustain. Energy 2 (2010), https://doi.org/10.1063/1.3294480. [25] M. El-Sheekh, A.E.F. Abomohra, D. Hanelt, Optimization of biomass and fatty acid productivity of Scenedesmus obliquus as a promising microalga for biodiesel production, World J. Microbiol. Biotechnol. 29 (2013) 915–922, https://doi.org/ 10.1007/s11274-012-1248-2. [26] E.S. Salama, H.C. Kim, R.A.I. Abou-Shanab, M.K. Ji, Y.K. Oh, S.H. Kim, B.H. Jeon, Biomass, lipid content, and fatty acid composition of freshwater Chlamydomonas mexicana and Scenedesmus obliquus grown under salt stress, Bioproc. Biosyst. Eng. 36 (2013) 827–833, https://doi.org/10.1007/s00449-013-0919-1. [27] F. Fasaei, J.H. Bitter, P.M. Slegers, A.J.B. van Boxtel, Techno-economic evaluation of microalgae harvesting and dewatering systems, Algal Res. 31 (2018) 347–362, https://doi.org/10.1016/j.algal.2017.11.038. [28] P.W. Behrens, V.J. Sicotte, J. Delente, Microalgae as a source of stable isotopically labeled compounds, J. Appl. Phycol. 6 (1994) 113–121, https://doi.org/10.1007/ BF02186065. [29] H.G. Gerken, B. Donohoe, E.P. Knoshaug, Enzymatic cell wall degradation of Chlorella vulgaris and other microalgae for biofuels production, Planta 237 (2013) 239–253, https://doi.org/10.1007/s00425-012-1765-0. [30] U.D. Keris-Sen, U. Sen, G. Soydemir, M.D. Gurol, An investigation of ultrasound effect on microalgal cell integrity and lipid extraction efficiency, Bioresour. Technol. 152 (2014) 407–413, https://doi.org/10.1016/j.biortech.2013.11.018. [31] D.M. de M. Dantas, C.Y.B. de Oliveira, R.M.P.B. Costa, M. das G. Carneiro-daCunha, A.O. G� alvez, R. de S. Bezerra, Evaluation of antioxidant and antibacterial capacity of green microalgae Scenedesmus subspicatus, Food Sci. Technol. Int. 25 (2019) 318–326, https://doi.org/10.1177/1082013218825024. [32] C. Chen, S. Yang, X. Bu, Microwave drying effect on pyrolysis characteristics and kinetics of microalgae, Bioenergy Res. (2019), https://doi.org/10.1007/s12155019-09970-z. [33] T. Mazzuca Sobczuk, Y. Chisti, Potential fuel oils from the microalga Choricystis minor, J. Chem. Technol. Biotechnol. 85 (2010) 100–108, https://doi.org/ 10.1002/jctb.2272.
6
Contents lists available at ScienceDirect
Biomass and Bioenergy journal homepage: http://www.elsevier.com/locate/biombioe
Research paper
A comparison of harvesting and drying methodologies on fatty acids composition of the green microalga Scenedesmus obliquus Carlos Yure Barbosa de Oliveira a, *, Thayna Lye Viegas a, Rafael Garcia Lopes a, Herculano Cella a, Rafael Silva Menezes b, Aline Terra Soares b, Nelson Roberto Antoniosi Filho b, Roberto Bianchini Derner a a b
Laboratory of Algae Cultivation, Aquaculture Department, Federal University of Santa Catarina (UFSC), 88061-600, Florian� opolis, Brazil Laboratory of Extraction and Separation Methods, Chemistry Institute, Federal University of Goias (UFG), 74690-900, Goi^ ania, Brazil
A R T I C L E I N F O
A B S T R A C T
Keywords: Flocculation Biomass recovery Biomass drying Saturated fatty acids Biodiesel
Harvesting and drying processes are still obstacles in the microalgae production chain. In this study, a combi nation of different harvesting and drying methods for Scenedesmus obliquus was proposed. S. obliquus was cultivated in a pond raceway for 15 days until reaching stationary phase. The culture was separated by different harvesting methods (centrifugation and flocculation) and drying methods (freezing, freeze-drying and oven drying), each combination in triplicate. Flocculation did not influence FAME yield of S. obliquus, except when the biomass was dried in an oven. The biomass that was only frozen had the lowest FAME content due to the presence of reactive water in the biomass. In contrast, frozen biomass had higher content of saturated and monosaturated fatty acids; freeze-drying or oven drying caused an enrichment of the polyunsaturated fatty acids content. In conclusion, if the biomass will be used to extract polyunsaturated fatty acids, freeze-drying would be more appropriate. On the other hand, if the purpose of the biomass is to produce biodiesel, the best process would be to just freeze the biomass.
1. Introduction
standards requirements [5]. Unfortunately, strains with high oil contents in their biomass often present low growth rates [6]. Therefore, developing large scale pro duction techniques for strains that have satisfactory growth rates and high lipid content is one of the challenges to the feasibility of industrial production of microalgae biodiesel. In this context, Scenedesmus obliquus is considered an organism with great potential [5] because it has fea tures that can facilitate massive outdoor production such as resistance to high temperatures and pH, and high growth rates [7,8]. The technical-economic feasibility of using microalgae biomass as raw material for biodiesel production presents other challenges, mainly in the stages of harvesting, drying and storage [9]. Microalgae cultures can be harvested by several individual methods, or by a combination of methods (e.g. flocculation plus centrifugation), and although they are important steps in the microalgae biomass production chain, clear har vesting and storage protocols have not yet been established [10]. The microalgae biomass harvesting and drying processes influence the costs and quality of the final products [11]. Generally, centrifugation is the most efficient harvesting method and it has been used in most pilot-scale
Microalgae oils have advantages over animal fats and higher plant oils for the production of biodiesel, since microalgae can be grown in areas unsuitable for agriculture [1], and liquid and gaseous effluents can be used for microalgae production [2]. In addition, microalgae biomass contains other high added-value bioproducts, such as vitamins, pig ments and amino acids, with potential use in several industrial segments and whose sale can help compensate for the production cost of biofuels made from microalgae [3]. The oil content of the most frequently studied microalgae species varies from 20 to 30% of dry weight [1]. Among the lipid classes, fatty acids are a group of major economic interest, and they are mainly classified according to its unsaturation levels. Different fatty acids classes might be used for different purposes, e.g., mono and saturated fatty acid esters are preferable to biodiesel production, while poly unsaturated fatty acids are considerate essential to animal diet and present bioactive properties [4]. So, besides showing a reasonable lipidic content, the fatty acid profile must also meet the international
* Corresponding author. E-mail addresses: [email protected], [email protected] (C.Y.B. Oliveira). https://doi.org/10.1016/j.biombioe.2019.105437 Received 25 May 2019; Received in revised form 23 November 2019; Accepted 27 November 2019 Available online 9 December 2019 0961-9534/© 2019 Elsevier Ltd. All rights reserved.
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
productions [12]. However, centrifuging is often economically unviable on an industrial scale due to high operating costs [13]. Flocculation has been a convenient separation method for concen trating large volumes of microalgae [14]. Although it is still necessary to filter or centrifuge the biomass after flocculation, the fact that this process concentrates the crop at a considerably lower volume can reduce the time and energy costs associated with separation [15]. The biomass harvesting and drying processes must be simple, fast, low cost and have high efficiency in the preservation of the biochemical quality of the biomass [16]. Although much has been studied about biomass harvesting; little is known of the impacts caused by the different drying methods, and less about the harvesting and drying interaction [12,16–18]. Determining the methodology that provides the best performance in harvesting and drying without qualitative damages to the algal biomass is important to establishing the production chain of any bioproduct derived from microalgae. For this reason, this study evaluated the influence of har vesting and drying methods on the yield and profile of fatty acid methyl esters (FAME) extracted from the green microalga Scenedesmus obliquus.
20 � C and stored until the moment of the next analysis. The frozen biomass remained at this temperature until the direct transesterification step. 2.2.4. Freeze-drying The samples subjected to the freeze-drying process were frozen to 20 � C, and then freeze-dried (L101 model, Liobras, Brazil), at 55 � C and 0.13 mbar, over 48 h. 2.2.5. Oven-drying The process in the oven-drying occurred with continuous forced air flow at 45 � C in 48 h. 2.3. Direct transesterification of microalgal biomass Approximately 200 mg of microalgal biomass was weighed. Next, 3.0 mL of 0.5 mol L 1 sodium hydroxide solution (MERCK ®) in dry methanol (TEDIA ®) was added, and the test tube was heated for 10 min in a 90 � C water bath. The test tube was cooled in an ice bath, and 9.0 mL of the mixture previously prepared for esterification was added ac cording to Soares et al. [3]. The test tube was heated again for 10 min in a 90 � C water bath. The test tube was cooled in an ice bath, and 5.0 mL of n-heptane (TEDIA ®) and 2.0 mL of distilled water were added and was put to rest by 2 h. The test tubes were shaken again for 1 min at 3,000 rpm and allowed to stand for 24 h until phase separation. The assays were performed in triplicate. The heptanic phases were collected with a Pasteur pipettes, inserted into 1.5 mL vials and analyzed by gas chro matography. The extraction procedure was performed in triplicate. To determine the FAME content from S. obliquus, a graph was created that correlated the sum of the peak area of the FAME (Y) of the soybean oil as a function of soybean oil mass (X) used in the FAME production through the transesterification of the oil, as the method described above. The correlation coefficient value was 0.999 for the calibration curve.
2. Material and methods 2.1. Microalga and growth conditions Scenedesmus obliquus strain was provided by the Culture Center of the Laboratory of Algae Cultivation, Brazil and maintained in LCA-AD (adapted from Bold’s Basal Medium) containing: 1.0 g L 1 NaNO3; 0.75 g L 1 MgSO4 7H2O; 0.6 g L 1 KH2PO4; 0.05 g L 1 EDTA-Na2; 0.03 g L 1 KOH; 0.25 g L 1 K2HPO4; 0.25 g L 1 NaCl; 0.25 g L 1 CaCl2 2H2O; 0.11 g L 1 H3BO3; 0.05 g L 1 FeSO4 7H2O; 0.014 g L 1 MnCl 4H2O; 0.016 g L 1 CuSO4 5H2O; 0.00071 g L 1 MoO3; 0.0004 g L 1 Co(NO3) 6H2O. S. obliquus was maintained in 2-L borosilicate glass flasks at 22 � 1 � C, 560 μmol photons m 2 s 1 irradiance with continuous lighting and stirred by bubbling air enriched with 0.5% CO 2. 200-L inoculum of S. obliquus was transferred to an outdoor raceway tank containing 2,000-L of culture medium. S. obliquus was cultured for 15 days, at outdoor temperatures (24 � 4 � C), with an average of 1,000 μmol photons m 2 s 1 of natural irradiance during the illuminated period, with a 12:12 photoperiod and agitated by a continuous flow of 0.5 m s 1. Cultivation was interrupted in the stationary phase when biomass reached 0.5 g L 1. The biomass obtained was fractionated in the different treatments according to the harvesting methodology: centrifugation; flocculation; and subsequent drying: frozen biomass only; freeze-dried and; oven dried, corresponding to a two-factorial design (2 � 3), in triplicate.
2.4. Gas chromatographic analysis An Agilent 7890 gas chromatograph equipped with a Flame Ioniza tion Detector (FID) and a split/splitless injector was used to analyze the FAME composition. The capillary column used was the DB-WAX (30 m � 0.25 mm � 0.25 μm). The initial oven temperature was 70 � C, and it was heated at a rate of 10 � C min 1 to 240 � C and maintained at this temperature for 13 min. It was heated again at a rate of 5 � C min 1 to 250 � C. The injector temperature was kept at 310 � C in split mode with a split ratio of 10:1 for an injection volume of 2 μL. The oven temperature was maintained at 310 � C. Hydrogen (5.0) was used as the carrier gas at a linear velocity of 42 cm s 1, and nitrogen was used as the auxiliary gas at a rate of 20 mL min 1. The FAMEs were identified by comparison with the retention times of samples of known composition such as soybean, peanut, and crambe (Crambe abyssinica) oils, by analyzing FAME reference standards (NuCheck-Prep ®) and by gas chromatography coupled with high resolution mass spectrometry (GC-HRMS) using a Shimadzu model 17A chro matograph coupled with a Shimadzu QP-5050 mass spectrometer. The carrier gas was helium with a linear velocity of 42 cm s 1. The GC-FID operating conditions (oven, injector, interface, and capillary column) were maintained for GC-HRMS [19].
2.2. Harvesting and drying methods 2.2.1. Centrifugation Three biomass samples, each in triplicate, were concentrated using an industrial centrifuge (SSD-06 model, GEA-Westfalia, Germany) in continuous processing (12,000 rpm). 2.2.2. Flocculation Since charge neutralization is the basis of flocculation and the elec trical charge of the microalgae cell wall is negative, cationic flocculants are generally more effective than anionic flocculants. For this reason, cationic polyacrylamide FLOPAM ® (courtesy of Professor Paulo Abreu, Federal University of Rio Grande) was used as the flocculating agent, with a medium charge density and high molecular weight, at 10 mg L 1 final concentration [9]. The biomass concentrated in this process was centrifuged using the same centrifuge and rotation used in the previous treatments.
2.5. Statistical analysis Two-way analysis of variance (ANOVA), followed by the Tukey’s post-hoc test was applied when necessary to compare the fatty acids profile among the harvesting and storage methodologies. Principal component analysis (PCA) was performed to determine possible patterns among the harvesting and drying combinations, using a correlation to produce a cross product matrix [20]. All statistical analyses were
2.2.3. Freezing Samples from both harvesting processes were immediately frozen at 2
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
Fig. 1. Fatty acid methyl esters content (mean � SD) of Scenedesmus obliquus harvested and dried by different methodologies. Different letters represent significant differences among the drying methods; * represent significant differences between the harvest methods, by Tukey’s post-hoc test.
performed at 5% significance level using RStudio® software version 3.5.1 (RStudio Inc, USA).
oven drying treatments was 3.2 � 0.2%; and for the biomass that was not dried (freezing), the moisture content was 64.5 � 12.37%. Thus, if the average moisture content contained in the frozen biomass is dis regarded, the FAME yield per gram of dry biomass would be approxi mately 119.41 mg g 1, similar to that found for the treatments where the biomass was dried. However, the removal of water from the reaction medium favors the transesterification reaction, since the acidic and basic catalysts will interact more effectively with the microalgae biomass. The presence of water may lead to a decrease in the base and acid concentrations present in the esterification mixture, since these solutions are in methanol, which interacts with water from the medium. Moreover, oils can be difficult to extract using solvent from wet biomass [21], especially when a cell wall disruption strategy is not adopted. These characteristics may be related to the different morphological types of microalgae cell walls. Indeed, for the diatom Isochrysis galbana, the lipids were also easily extracted when the biomass was freeze-dried [22]. Cationic polymers, polyacrylamide based like FLOPAM, which was used in the present study, act as coagulants, neutralizing part of the negative charges on microalgae cell walls at neutral pH and thus, it re duces the cellular repulsion. After the flocculation occurs, the resulting
3. Results and discussion 3.1. Direct transesterification of microalgal biomass Fig. 1 shows the direct transesterification of Scenedesmus obliquus biomass from the different combinations of harvesting and drying methods. The percentage of FAME extracted from the dry biomass of S. obliquus varied between 30 mg g (moist weight) 1, for the biomass that underwent flocculation and freezing, and 140 mg g (dry weight) 1, for the biomass that underwent centrifugation and freeze-drying, for the minimum and maximum, respectively. However, no significant differ ences were observed between the centrifugation plus freeze-drying, flocculation plus freeze-drying and centrifugation plus oven-drying combinations. The flocculation only had a negative influence (p < 0.05) when the biomass was dried in an oven. For the other treatments, flocculation using cationic polyacrylamide did not influence the direct transesterification of S. obliquus. The moisture content in the biomass that underwent freeze-drying or
Fig. 2. Fatty acids classes (in % FAME) of the Scenedesmus obliquus for the different combinations of biomass harvesting and drying methodologies. 3
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
flocs were also centrifuged (same conditions of the other treatments) and since the flocculation does not negatively impact the cell composi tion [23], it was expected that the FAME profile would not be affected. However, the combination of flocculation plus oven drying resulted in a low FAME yield, as it was observed that dry biomass had a thin layer of the flocculant. This might have acted as a physical barrier, not allowing an efficient extraction of the non-polar fraction of S. obliquus biomass with the use of solvents. The high temperatures of the oven may have favored the binding of the polymers with the cell walls, making it difficult to extract the lipids [11,24]. This is, supposedly, the reason to the differences found in FAME concentration between centrifugation and flocculation with posterior oven drying, since in the absence of the flocculant agent, FAME concentration was not impacted. For the higher FAME yield, the results found in this study were similar to those reported by El-Sheekh et al. [25], about 150 mg g 1. However, S. obliquus can reach higher FAME contents, of 340 mg g 1, when grown under controlled conditions [26]. Although the results re ported here were similar to other studies, the production of S. obliquus in an outdoor raceway tank possibly influenced the FAME concentration, mostly due to oscillations of environmental parameters that occur in open systems.
Table 1 Fatty acids profile of Scenedesmus obliquus for the different combinations of biomass harvesting and drying methodologies. Fatty acid
Centrifugation Freezing
Freezedrying
Ovendrying
Freezing
Freezedrying
Ovendrying
C14:0
1.13 � 0.06 24.97 � 0.55a 13.63 � 0.21a 2.07 � 0.11a 7.57 � 0.21a 0.37 � 0.11 n.d.b
1.07 � 0.38 12.5 � 0.79b 3.30 � 0.10b 1.40 � 0.00b 3.53 � 0.21b 0.20 � 0.01 0.13 � 0.06a 0.87 � 0.06a
1.57 � 0.50 29.5 � 1.39a 11.17 � 2.97a 2.93 � 0.21a 8.47 � 0.60a 0.27 � 0.06 n.d.b
0.50 � 0.00b*
0.8 � 0.01 11.67 � 0.25b 3.4 � 0.20b 1.23 � 0.06c 3.07 � 0.15b 0.13 � 0.06 0.13 � 0.06a 0.87 � 0.06a
1.03 � 0.11 14.07 � 1.27b 3.53 � 0.90b 1.77 � 0.11b 3.67 � 0.38b 0.30 � 0.26 0.20 � 0.01a 1.10 � 0.10a
1.00 � 0.00 13.77 � 0.38b 3.57 � 0.21b 1.90 � 0.01b 3.97 � 0.06b 0.67 � 0.06 0.17 � 0.06a 1.10 � 0.00a
5.87 � 0.67b
21.17 � 0.64a
20.43 � 1.01a
3.40 � 0.44b
17.97 � 2.58a
19.5 � 0.61a
1.10 � 0.00a 10.03 � 0.58a* 3.03 � 0.06a* 3.73 � 0.06 0.20 � 0.01b
0.40 � 0.01c 4.83 � 0.11b* 1.4 � 0.01b* 4.17 � 0.11 0.30 � 0.00a
0.67 � 0.06a 5.23 � 0.40b* 1.60 � 0.10b* 4.20 � 0.20 0.30 � 0.00a
1.07 � 0.15a 12.53 � 0.25a 4.10 � 0.14a 4.13 � 0.11 n.d.b
0.43 � 0.06c 6.43 � 0.58b 2.07 � 0.21b 4.83 � 0.23 0.30 � 0.00a
0.63 � 0.06b 6.17 � 0.15b 2.13 � 0.06b 4.50 � 0.17 0.30 � 0.00a
22.27 � 0.75b
41.17 � 0.15a
25.10 � 0.75b
17.57 � 0.57c
25.1 � 21.316b
36.13 � 0.50a
1.53 � 0.11b
4.37 � 0.11a*
4.13 � 0.25a
0.90 � 0.10b
3.70 � 0.36a*
3.60 � 0.10a
1.10 � 0.00a 0.90 � 0.00a 6.23 � 0.31a
0.43 � 0.06b 0.30 � 0.00b 3.97 � 0.21b
0.47 � 0.15b* 0.37 � 0.06b* 4.37 � 0.38b
1.43 � 0.64 0.90 � 0.00a 6.67 � 0.76
1.67 � 1.94 0.40 � 0.10b 6.69 � 3.14
1.03 � 0.06 n.d.b
C16:0 C16:1 cis 9 C16:1 C16:1 cis 11 C16:2 cis 7,10 C17:0 C16:3 cis 6, 9, 12 C16:4 cis 6, 9, 12, 15 C18:0
3.2. FAME profile of S. obliquus Fig. 2 shows the fatty acids classes for the different combinations of harvesting and drying. The biomass that was not dried (directly frozen after harvesting) showed the best profile for biodiesel production. The polyunsaturated fatty acids percentage nearly doubled when the biomass was dried in a freeze-dryer or oven. The content of diunsatu rated fatty acids remained close to 4% regardless of the combination used. The harvesting methodology did not influence significant modi fications in the fatty acid classes, since once these possible cell disrup tions occurred by the centrifugation forces, followed by possible intracellular material leakage, were present in all treatments. Drying (and storage) methodologies represent an obstacle in the microalgae biomass production chain. In addition to freeze-drying and oven drying, spray drying, sun drying, and drum drying have already been studied [17]. The correct selection of the drying method to produce biodiesel from microalgae is an important step still to be taken [18]. The use of freeze-dryers has already been ruled out for large-scale production because of the high operating costs. The findings reported in this study represent an important advance, especially considering that the highest saturated and monosaturated fatty acids levels were found when the biomass was not dried. This can represent great cost and time saving in the microalgae biodiesel production chain [27]. Drum-drying, spray-drying or freeze-drying of Dunaliella, β-carotenerich, produced satisfactory results in dry biomass uniformity and sta bility of β-carotene. The influence of the drying method on the compo sition and amount of the metabolite is probably linked to molecule stability [28], cell wall composition, resistance [29] and microalgal cell integrity [30]. It should also be emphasized that if the goal is the re covery of polyunsaturated fatty acids (those with nutraceutical proper ties), freeze-drying has considerably increased the levels of these fatty acids. In the extraction of bioactive compounds from S. subspicatus, the use of water was more efficient than that of potentially toxic solvents [31]. Possible interactions among saturated and monounsaturated fatty acids with polar solvents (such as water) have not yet been established. However, in this study, the extracellular water in the frozen biomass preserved a more saturated and monosaturated fatty acid profile. The use of oven drying resulting from harvesting processes can cause pyrolysis of intracellular organic compounds [32]. Although the decomposition of volatiles, such as carbohydrates and lipids, usually occurs at higher temperatures (around 120–500 � C) in microalgae, the exposure of S. obliquus to 45 � C for 48 h may have oxidized some of the saturated and monosaturated fraction. The fatty acid profiles (Table 1) of S. obliquus had higher levels of
C18:1 cis 9 C18:1 cis 11 C18:2 cis 9, 12 C18:3 cis 6, 9, 12 C18:3 cis 9, 12, 15 C18:4 cis 6, 9, 12, 15 C22:0 C24:0 Others
Floculation
n.d.b
5.67 � 0.11
Different letters represent significant differences among the drying methods; * represent significant differences between the harvest methods, by Tukey’s posthoc test. n.d. not detected.
Linolenic (C18:3 cis 9, 12, 15), Palmitic (C16:0) and Hexadecatetraenoic (C16:4 cis 6, 9, 12, 15) acids. Again, no significant differences were observed between the biomass harvesting methods. However, individual FAME differed significantly between the drying methods. The highest Linolenic percentage (the most abundant fatty acid) was reported for the centrifugation plus freeze-drying and centrifugation plus oven-drying combinations. The Palmitic acid concentrations were significantly higher in the treatments in which the biomass was frozen soon after being concentrated (centrifuged or flocculated) (Table 1). However, the drying of the biomass, by both freeze- and oven-drying, may have caused the biomass to oxidize due to the loss of free water, which could lead to an increase in saturated and monounsaturated fractions of FAME. This could explain why higher content of poly unsaturated fatty acids was observed in the treatments of biomass with drying than in those without drying. PCA ordination (F ¼ 41.76; p ¼ 0.002) for 6 combinations of har vesting and drying methodologies, explained 99.34% of data variability in the first two components (Fig. 3). Polyunsaturated (r ¼ -0.552), 4
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
Fig. 3. Principal components analysis (PCA) ordination diagram for the combinations of harvesting (CEN centrifugation, FLO flocculation) and drying (FZ freezing, FD freeze-drying and OD oven-drying) methodologies.
Monounsaturated (r ¼ 0.553) and Saturated (r ¼ 0.549) were the principal fatty acids in the composition of component 1 (79.60%), while Diunsaturated (r ¼ 0.95) was the most important fatty acid for compo nent 2 (19.74%). The diagram for PCA indicates a clear distinction be tween the treatments that were only frozen, independent of having been harvested by centrifugation or flocculation, in relation to mono- and saturated fatty acids. Even with the low polyunsaturated fatty acids content, the oil of S. obliquus presents restrictions to the production of biodiesel that at tends European regulation EN 14214 (which allows only 1% of fatty acids with more than three double bonds). Choricystis minor var. minor was considered a promising species for the production of biodiesel because of its fatty acid profile [33]. However, the linolenic acid content reported by Menezes et al. [19] was greater than 12% (the maximum limit acceptable to EN 14214) using centrifugation plus freeze-drying as biomass harvesting and drying methods, respectively. In this study, this combination showed a close to 100% increase in the linolenic acid content when compared to biomass that was not dried (only frozen). However, it is worth noting that the combination of harvesting and drying methodologies may present different performances in other species, due to the specific characteristics already mentioned.
flocculation and oven-drying reduced the FAME yield. Human and animal rights The study did not involve human subjects or animal models. Author contributions CYBO and TLV analyzed the data. RGL, HC and ATS conducted the experiments. All the authors read and edited the manuscript and pro vided valuable inputs to improve the quality of the article. Declaration of Competing interest The authors declare that they have no competing interests. Acknowledgment The authors would like to thank the Ministry of Science, Technology and Innovation (MCTI) for financial support provided by FINEP (Agreement No. 01.10.0457.00), Conselho Nacional de Desenvolvi �gico (Case No. 574796/2008-8), Coor mento Científico e Tecnolo ~o de Aperfeiçoamento de Pessoal de Nível Superior Brazil denaça (Finance Code 001).
4. Conclusions The two harvesting methodologies that were analyzed in this study did not affect the fatty acid profile of Scenedesmus obliquus. In contrast, the fatty acid content did change when they were dried using different methodologies. If the goal is to increase the polyunsaturated fatty acids content, the biomass should be freeze-dried or oven-dried. On other hand, if the biomass will be used to produce biodiesel, the biomass should be frozen after harvesting. Finally, the combination of
References [1] Y. Chisti, Biodiesel from microalgae, Biotechnol. Adv. 25 (2007) 294–306, https:// doi.org/10.1016/j.biotechadv.2007.02.001. [2] C.Y.B. Oliveira, J.L. Abreu, C.D.L. Oliveira, P.C. Lima, A.O. G� alvez, D.M.M. Dantas, Growth of Chlorella vulgaris using wastewater from Nile tilapia (Oreochromis niloticus) farming in a low-salinity biofloc system, Acta Sci. Technol. 42 (2020), e46232, https://doi.org/10.4025/actascitechnol.v42i1.46232.
5
C.Y.B. Oliveira et al.
Biomass and Bioenergy 132 (2020) 105437
[3] A.T. Soares, D.C. da Costa, B.F. Silva, R.G. Lopes, R.B. Derner, N.R. Antoniosi Filho, Comparative analysis of the fatty acid composition of microalgae obtained by different oil extraction methods and direct biomass transesterification, Bioenergy Res. 7 (2014) 1035–1044, https://doi.org/10.1007/s12155-014-9446-4. [4] B. Ramesh Kumar, G. Deviram, T. Mathimani, P.A. Duc, A. Pugazhendhi, Microalgae as rich source of polyunsaturated fatty acids, Biocatal. Agric. Biotechnol. 17 (2019) 583–588, https://doi.org/10.1016/j.bcab.2019.01.017. [5] S. Mandal, N. Mallick, Microalga Scenedesmus obliquus as a potential source for biodiesel production, Appl. Microbiol. Biotechnol. 84 (2009) 281–291, https://doi. org/10.1007/s00253-009-1935-6. [6] C. Dayananda, R. Sarada, M. Usha Rani, T.R. Shamala, G.A. Ravishankar, Autotrophic cultivation of Botryococcus braunii for the production of hydrocarbons and exopolysaccharides in various media, Biomass Bioenergy 31 (2007) 87–93, https://doi.org/10.1016/j.biombioe.2006.05.001. [7] M.G. de Morais, J.A.V. Costa, Biofixation of carbon dioxide by Spirulina sp. and Scenedesmus obliquus cultivated in a three-stage serial tubular photobioreactor, J. Biotechnol. 129 (2007) 439–445, https://doi.org/10.1016/j. jbiotec.2007.01.009. [8] T. de Marchin, M. Erpicum, F. Franck, Photosynthesis of Scenedesmus obliquus in outdoor open thin-layer cascade system in high and low CO2 in Belgium, J. Biotechnol. 215 (2015) 2–12, https://doi.org/10.1016/j.jbiotec.2015.06.429. [9] R.B. K€ onig, R. Sales, F. Roselet, P.C. Abreu, Harvesting of the marine microalga Conticribra weissflogii (Bacillariophyceae) by cationic polymeric flocculants, Biomass Bioenergy 68 (2014) 1–6, https://doi.org/10.1016/j. biombioe.2014.06.001. [10] T.M. Mata, A.A. Martins, N.S. Caetano, Microalgae for biodiesel production and other applications: a review, Renew. Sustain. Energy Rev. 14 (2010) 217–232, https://doi.org/10.1016/j.rser.2009.07.020. [11] L. Borges, J.A. Mor� on-Villarreyes, M.G.M. D’Oca, P.C. Abreu, Effects of flocculants on lipid extraction and fatty acid composition of the microalgae Nannochloropsis oculata and Thalassiosira weissflogii, Biomass Bioenergy 35 (2011) 4449–4454, https://doi.org/10.1016/j.biombioe.2011.09.003. [12] A.J. Dassey, C.S. Theegala, Harvesting economics and strategies using centrifugation for cost effective separation of microalgae cells for biodiesel applications, Bioresour. Technol. 128 (2013) 241–245, https://doi.org/10.1016/j. biortech.2012.10.061. [13] J.J. Milledge, S. Heaven, A review of the harvesting of micro-algae for biofuel production, Rev. Environ. Sci. Biotechnol. 12 (2013) 165–178, https://doi.org/ 10.1007/s11157-012-9301-z. [14] R.M. Knuckey, M.R. Brown, R. Robert, D.M.F. Frampton, Production of microalgal concentrates by flocculation and their assessment as aquaculture feeds, Aquacult. Eng. 35 (2006) 300–313, https://doi.org/10.1016/j.aquaeng.2006.04.001. [15] R. Sales, P.C. Abreu, Use of natural pH variation to increase the flocculation of the marine microalgae Nannochloropsis oculata, Appl. Biochem. Biotechnol. 175 (2014) 2012–2019, https://doi.org/10.1007/s12010-014-1412-2. [16] M.K. Danquah, L. Ang, N. Uduman, N. Moheimani, G.M. Forde, Dewatering of microalgal culture for biodiesel production: exploring polymer flocculation and tangential flow filtration, J. Chem. Technol. Biotechnol. 84 (2009) 1078–1083, https://doi.org/10.1002/jctb.2137. [17] E. Molina Grima, E.H. Belarbi, F.G. Aci�en Fern� andez, A. Robles Medina, Y. Chisti, Recovery of microalgal biomass and metabolites: process options and economics, Biotechnol. Adv. 20 (2003) 491–515, https://doi.org/10.1016/S0734-9750(02) 00050-2. [18] K.Y. Show, D.-J. Lee, A.S. Mujumdar, Advances and challenges on algae harvesting and drying, Dry. Technol. 33 (2015) 386–394, https://doi.org/10.1080/ 07373937.2014.948554.
[19] R.S. Menezes, A.T. Soares, J.G. Marques Júnior, R.G. Lopes, R.F. da Arantes, R. B. Derner, N.R.A. Filho, Culture medium influence on growth, fatty acid, and pigment composition of Choricystis minor var. minor: a suitable microalga for biodiesel production, J. Appl. Phycol. 28 (2016) 2679–2686, https://doi.org/ 10.1007/s10811-016-0828-1. [20] C.Y.B. Oliveira, C.D.L. Oliveira, A.J.G. Almeida, A.O. G� alvez, D.M. Dantas, Phytoplankton responses to an extreme drought season: a case study at two reservoirs from a semiarid region, Northeastern Brazil, J. Limnol. 78 (2) (2019) 176–184, https://doi.org/10.4081/jlimnol.2019.1869. [21] E.H. Belarbi, E. Molina, Y. Chisti, A process for high yield and scaleable recovery of high purity eicosapentaenoic acid esters from microalgae and fish oil, Enzym. Microb. Technol. 26 (7) (2000) 516–529. [22] E.M. Grima, A.R. Medina, A.G. Gim� enez, J.A. S� anchez P�erez, F.G. Camacho, J. L. García S� anchez, Comparison between extraction of lipids and fatty acids from microalgal biomass, J. Am. Oil Chem. Soc. 71 (1994) 955–959, https://doi.org/ 10.1007/BF02542261. [23] R. Sales, R.B. Derner, M.Y. Tsuzuki, Effects of different harvesting and processing methods on Nannochloropsis oculata concentrates and their application on rotifer Brachionus sp. cultures, J. Appl. Phycol. (2019), https://doi.org/10.1007/s10811019-01877-8. [24] N. Uduman, Y. Qi, M.K. Danquah, G.M. Forde, A. Hoadley, Dewatering of microalgal cultures: a major bottleneck to algae-based fuels, J. Renew. Sustain. Energy 2 (2010), https://doi.org/10.1063/1.3294480. [25] M. El-Sheekh, A.E.F. Abomohra, D. Hanelt, Optimization of biomass and fatty acid productivity of Scenedesmus obliquus as a promising microalga for biodiesel production, World J. Microbiol. Biotechnol. 29 (2013) 915–922, https://doi.org/ 10.1007/s11274-012-1248-2. [26] E.S. Salama, H.C. Kim, R.A.I. Abou-Shanab, M.K. Ji, Y.K. Oh, S.H. Kim, B.H. Jeon, Biomass, lipid content, and fatty acid composition of freshwater Chlamydomonas mexicana and Scenedesmus obliquus grown under salt stress, Bioproc. Biosyst. Eng. 36 (2013) 827–833, https://doi.org/10.1007/s00449-013-0919-1. [27] F. Fasaei, J.H. Bitter, P.M. Slegers, A.J.B. van Boxtel, Techno-economic evaluation of microalgae harvesting and dewatering systems, Algal Res. 31 (2018) 347–362, https://doi.org/10.1016/j.algal.2017.11.038. [28] P.W. Behrens, V.J. Sicotte, J. Delente, Microalgae as a source of stable isotopically labeled compounds, J. Appl. Phycol. 6 (1994) 113–121, https://doi.org/10.1007/ BF02186065. [29] H.G. Gerken, B. Donohoe, E.P. Knoshaug, Enzymatic cell wall degradation of Chlorella vulgaris and other microalgae for biofuels production, Planta 237 (2013) 239–253, https://doi.org/10.1007/s00425-012-1765-0. [30] U.D. Keris-Sen, U. Sen, G. Soydemir, M.D. Gurol, An investigation of ultrasound effect on microalgal cell integrity and lipid extraction efficiency, Bioresour. Technol. 152 (2014) 407–413, https://doi.org/10.1016/j.biortech.2013.11.018. [31] D.M. de M. Dantas, C.Y.B. de Oliveira, R.M.P.B. Costa, M. das G. Carneiro-daCunha, A.O. G� alvez, R. de S. Bezerra, Evaluation of antioxidant and antibacterial capacity of green microalgae Scenedesmus subspicatus, Food Sci. Technol. Int. 25 (2019) 318–326, https://doi.org/10.1177/1082013218825024. [32] C. Chen, S. Yang, X. Bu, Microwave drying effect on pyrolysis characteristics and kinetics of microalgae, Bioenergy Res. (2019), https://doi.org/10.1007/s12155019-09970-z. [33] T. Mazzuca Sobczuk, Y. Chisti, Potential fuel oils from the microalga Choricystis minor, J. Chem. Technol. Biotechnol. 85 (2010) 100–108, https://doi.org/ 10.1002/jctb.2272.
6