Extraction and characterization of the saponifiable lipid fraction
from microalgae Chlamydomonas sp. cultivated under stress
Anderson F. Gomes1•Tatiana de C. Bicudo2•Marta Costa1•Luiz Di Souza3•Luciene S. de Carvalho1
Received: 14 August 2018 / Accepted: 2 February 2019 / Published online: 15 February 2019 Akade´miai Kiado´, Budapest, Hungary 2019
Abstract
Microalgae when subjected to nutrient restriction conditions, inducing environmental stress, are prone to total lipid accumulation and changes in the fatty profile. In this context, the green microalgae Chlamydomonas sp. was submitted for 3 days to the absence of nutrients. Thus, the efficacy of the extraction of the crude hexane lipid fraction (CHF), using mechanical stirred associated with ultrasound technique, was evaluated in two different times (2 and 4 h), using scanning electron microscopy and thermogravimetric analysis (TG). Saponifiable fractions, CHF extracts and crude chloroform fraction (CCF) were esterified and characterized by TG, FTIR and GC/MS to identify their fatty profile. Residual biomass (RB) was analyzed by elemental analysis, and the resulting data were employed to estimate the higher heating value (HHV) and crude protein percentage. The micrographs of the biomass surfaces before and after the extraction process showed significant fragmentation only for the time of 4 h. A pronounced mass loss in the typical lipid range was evidenced by TG being consistent with the significant difference of material extracted in the time of 2 h (2.95 ± 0.28)% and 4 h (10.54 ± 0.46)%. The total lipid fraction (CHF and CCF) was approximately 29%. The RB revealed values of HHV and crude protein of 18 MJ kg-1and 56%, respectively. The TG data for the CCF extract revealed a material consisting mainly of saponifiable fractions, unlike CHF which is composed predominantly of unsaponifiables. Therefore, the TG curve ratified a better conversion rate to CCF (approximately 89%), consistent with the value obtained by GC/MS (about 86%). The fatty profile for the saponifiable fractions of both extracts showed that the major fatty acids are C16:0 and C18:3 (x-3 and x-6). The ester profile with elevated concentration of polyunsaturated fatty acids (C18:3) is unfeasible in their application for biodiesel production; nevertheless, the fatty profile of microalgae suggests pharmacological potential for diet or therapy, since some of the main components are reported as bioactive metabolites.
Keywords Microalgae Thermal analysis Cultivation Biomass Saponifiable
Introduction
The development of new processes for the functional production of biofuels, on a large scale, emanates as a viable and increasingly necessary proposal in the face of a
world scenario of energy insecurity and oil dependency. Thus, the search for clean resources to meet future energy needs is one of the greatest challenges today. The use of biomass, whether of animal or vegetable origin, for the production of biofuel is a valid alternative to minimize the environmental, social and economic impacts of society. Microalgal biomass has enormous potential as an energy feedstock, and its advantages are many when compared to traditional oleaginous plants (soybean, cotton, sunflower, etc.). Microalgae are autotrophic microorganisms that use light energy and inorganic nutrients (carbon dioxide, nitrogen, phosphorus, etc.) and synthesize compounds of high commercial value such as lipids, proteins, carbohy-drates, pigments and vitamins [1–3]. Among the advan-tages of microalgae are: exponential growth, need for less
& Anderson F. Gomes
gomesfisica007@hotmail.com
1 Institute of Chemistry, Federal University of Rio Grande do Norte, Natal, RN 59078-970, Brazil
2 Science and Technology School, Federal University of Rio Grande do Norte, Natal, RN 59078-970, Brazil
3 Catalysis Laboratory Environment and Materials-LACAM, Chemistry Department, FANAT, University of the State of Rio Grande do Norte, Mossoro´, RN 59620676, Brazil https://doi.org/10.1007/s10973-019-08071-5(0123456789().,-volV)(0123456789().,- volV)
area for cultivation and lipid percentage above 20% [4–6]. It is worth mentioning in relation to these advantages that the processing of any raw material should optimize the use of resources, aiming to maximize profitability and benefits and minimize waste and operational and environmental problems. The green microalgae Chlamydomonas sp. in optimized conditions stands out due to the high produc-tivity of biomass and lipid, requiring, on average, only 7 days for the renewal of its cultivation and presents sig-nificant tolerance to salinity [7,8]. Furthermore, it has been reported that when submitted to nitrogen deprivation con-ditions these microalgae are able to substantially accumu-late lipids in its cells [9].
The biorefinery is an industrial process, where biomass is converted into chemicals and energy, and the similar concept to that used to define the oil refinery. However, fractionation, an important step in the biorefinery process, is reported in the literature as an obstacle, since it is very difficult to separate the different classes of compounds [10–16]. Therefore, considering the concept of biorefinery, it is important to develop techniques that allow a wide withdrawal of metabolites, considering the difficulty of recovery, inaccessibility and/or their variety of polarities [17]. Solvent access to the metabolites of the intracellular region is a major obstacle in obtaining the lipid fraction, since the procedure should conserve the composition of the extract and dispense the use of too much solvent. In addition, it should efficiently induce the breakdown of the cell wall of the microalgae and allow the solvent to entrain the material of desirable polarity [18]. The use of the ultrasonic bath has been shown to be a very efficient technique in algae cell rupture [18, 19]. In addition, it consists of a cold process and requires a discrete amount of solvent without damaging the composition of the extract. The association of the ultrasonic method with mechanical agitation simultaneously was even more efficient, signifi-cantly increasing the lipid content extracted [20].
Microalgae may have their biosynthetic pathways manipulated and directed, through changes in cultivation conditions and environmental stress, to the production and substantial accumulation of compounds desirable for the production of biofuels [16]. The use of environmental stress in the cultivation of microalgae includes the absence or decrease in the concentration of some nutrients, control of the incidence of luminosity, increase in salinity and control of pH. Thus, the literature has reported that stress conditions help in the accumulation of specific secondary metabolites of lipids and carbohydrates (mainly pigments and vitamins). The absence of nutrients, mainly nitrogen, has provided several microalgae to accumulate lipids, predominantly saponifiable fractions (SF), including tria-cylglycerides (TAGs), fatty acids and phospholipids, and in some cases, these compounds account for more than 30%
of the dry biomass mass. In addition, they commonly present fatty acids with a diversified profile, which may include high concentration of unsaturated (x-3 and x-6) with high commercial value in the cosmetic and pharma-ceutical industry [21–23].
Residual biomass resulting from the fractionation of more refined chemical substances has been reported as a promising raw material for thermochemical conversion processes such as instantaneous pyrolysis (500C), con-sidered a technique with good viability for a future sub-stitution of fossil fuels due to the high conversion rate of biomass/liquid (95.5%) [24]. Generally, the microalgal biomass yields higher heating value (HHV) than that of herbaceous biomass, which is commonly employed for thermal energy obtainment. Moreover, it presents lower decomposition, ignition and burn temperatures due to the high level of lipids and carbohydrates, in addition to the reduced content of lignocellulose, thus enabling better efficiency in heat transfer and combustion [25, 26]. Aquaculture is another promising area for the use of algal biomass including applications such as fish feed, larval nutrition for mollusks or shrimp, and stabilization and improvement in the quality of the culture medium (‘‘green water’’ technique) [27,28].
The present study suggests a preliminary evaluation of the biorefinery potential of the microalgal biomass of the species Chlamydomonas sp. previously cultured under conditions optimized for the growth of their cells and subsequently subjected to complete nutrient deprivation. Thus, the efficacy of the extraction of its metabolites in particular the hexane lipid (CHF) and chloroform fraction (CCF) was evaluated through SEM and TG techniques. The residual biomass (RB) from the lipid fraction removal stage had its elemental composition elucidated, as well as its higher heating value (HHV) and estimated crude protein content. Finally, the CHF and CCF were hydroesterified for the thermal characterization of their SF and their fatty profile by gas chromatography coupled to mass spectrom-etry (GC/MS).
Materials and methods
Culture conditions of the microalgae
Chlamydomonas sp
The microalgae of the species Chlamydomonas sp. were provided by Petrobras/CENPES-UFRN partnership by means of the research project: ‘‘Implementation of a pilot plant for production of microalgal biomass, aiming bio-diesel production.’’ The microalgae were cultivated in the first week of May 2015 in the metropolitan region of Natal, state of Rio Grande do Norte, Brazil. Initially the culture
was performed in an open system (raceway) for 2 days, with a mean light intensity of 11 klx, salinity of 10%, pH = 6.7 and using commercial nutrients for microalgal biomass growth: 50 mg L-1urea, 20 mg L-1monoanionic phosphate (MAP) and 10 mg L-1 MgSO4. It should be noted that the nutrients were agitated by CO2-reinforced aeration 2. At the end of culture period, the microalgal cells were pumped to another raceway and then subjected to nutrient deprivation (total absence) for 3 days, maintaining the mentioned conditions. Finally, the biomass was pumped into a flocculation tank, maintained until its total decanting, and then collected and stored in a freezer at a temperature of - 20C. The control group (CTG) was cultivated for 5 days without interruption in nutrient supply and following the conditions already mentioned above.
Pretreatment of the microalgal biomass
and extraction of the crude hexane fraction
The microalgal biomass was initially dried in an Enterprise I series freeze-drier using N2gas under a reduced pressure of 7.10-3 bar and a temperature of - 5C. Then, the dry biomass was ground and its granulometry was controlled at 32 mesh. The extraction of HF (hexane fraction) was done using 1 g of biomass and 100 mL of n-hexane HPLC grade. The mixture under these conditions was subjected to extraction in a sealed glass vial with a small orifice for passing the propeller of the mechanical stirrer of the Fisatom-712 brand, employing rotation of 2000 rpm, and simultaneously, this system was coupled to an ultrasonic bath with frequency of 40 kHz and power of 90 W to aid in the process of cellular rupture for 2 and 4 h. It is worth mentioning that extractions of CHF were performed in triplicates. At the end of the extractive process, the resulting material was filtered to retain the residual biomass (RB), while the filtrate hexane fraction was concentrated in a rotary evaporator under reduced pressure under a ther-mostated bath at 50C. The crude hexane fraction (CHF) is calculated by Eq. (1):
CHF %ð Þ ¼ EHF 100 DBM
r ð1Þ
where EHF is the extracted hexane fraction (mg); DBM the dry microalgal biomass (mg); and r the standard deviation. Residual biomass after 2 h and 4 h of extraction (RBHF2h and RBHF4h, respectively) were transferred to laminated paper-covered flasks (with small holes) and allowed to stand for 24 h for residual solvent volatilization. After the end of the period, the material was weighed and residual biomass percentage (RBHF) was determined using the results and Eq. (2).
RBHF %ð Þ ¼ BBE 100 BAE
r ð2Þ
where BBE is the biomass before the extraction process (mg) and BAE the biomass after the extraction process.
Extractions of crude chloroform fractions
The CCF was extracted from the residual microalgal bio-mass (RBHF4h) using 100 mL of chloroform for each 1 g of biomass. After mixing, the system was sonicated for 1 h at a frequency of 20 kHz and a power of 50 W. At the end of the process, the solubilized material was transferred to test tubes which were subsequently centrifuged at 3000 rpm for 10 min. Then, the resulting mixture was fil-tered and the filtrate was concentrated in a rotary evapo-rator at 60C under reduced pressure. All extractions were performed in triplicates, and the residual biomass (RBCF) and percentage of crude chloroform fraction (CCF) are calculated with Eqs. (3) and (4), respectively. The total lipid fraction (TLF) was determined by the sum of the two fractions, according to Eq. (5):
RBCF %ð Þ ¼ BBE 100 DBM r ð3Þ CCF %ð Þ ¼ ECF 100 DBM r ð4Þ TLF %ð Þ ¼ CHF þ CCF ð5Þ
where CCF is the crude chloroform fraction; ECF the extracted chloroform fraction; BMS the dry biomass (mg); and TLF the total lipid fraction.
Hydroesterification of CHF and CCF
The CHF and CCF lipid fractions were initially hydrolyzed in a reflux system for 2 h under constant stirring using an alkoxide solution (methanol/NaOH) and a stoichiometric ratio of 5 mL of methanol to 100 mg of organic extract. The other reaction conditions included temperature of 60C and the use of 2% NaOH. Then, the reaction mixture was transferred to a separatory funnel, adding distilled water and the same volume of n-hexane. The resulting biphasic system contains the unsaponifiable material as the supernatant, while the denser phase is composed of the saponifiable substances (mainly fatty acids).
Fatty acids (FA) were obtained by separating the aque-ous phase and then acidifying with a 6 mol L-1 HCl solution until pH 2. After acidification, the mixture con-taining the FA was transferred to a separatory funnel, and for extraction, ethyl ether was added again forming a two-phase system. The upper two-phase (FA) was separated and concentrated in a rotaevaporator at 40 C under reduced
pressure. The FA were esterified at 60C for 3 h in a constant stirring reflux system using a stoichiometric ratio of 5 mL of methanol to 100 mg of FA using 1% catalyst (H2SO4ACS).
At the end of the reaction, the reaction mixture was transferred to a separatory funnel and washed two times with 20 mL of distilled water, and subsequently, for extraction of the MME dichloromethane was again added, resulting in a two-phase system again. Finally the denser phase composed of the MME was separated and concen-trated in a rotaevaporator at 60C under reduced pressure.
Scanning electron microscopy
The biomass samples before the extractive processes (BBE) and after the extraction processes RBHF2h and RBHF4h were placed on graphite carbon ribbons and made superficial images with a scanning electron microscope model Hitachi Tabletop Microscope TM-3000, with backscattered electron detector of semiconductor type, acceleration voltage of 5–15 kV and with magnification capacity of 15–30,000 times. The applied software to obtain the average diameter of the biomass fragments for SEM image analysis was the TopoMAPS Software from Thermo Fisher Scientific.
Elemental analysis of microalgal biomass
The microalgal biomass before (BBE) and after extraction with hexane (RBHF4h) and hexane followed by chloro-form (RBCF) were determined by an elemental analyzer (CHNS), brand vario MACRO cube. In the characteriza-tion, 50 mg of sample was used and the elements quantified by combustion at 1150C. The higher heating value (HHV) of the biomasses was estimated using the percent-age values of the elemental composition [29], according to Eq. (6):
HHV kJ kg1¼ 5:22C2319C1647H þ 38:66C H
þ 133N þ 21028
ð6Þ The crude protein values present in the samples were also estimated using the elemental analysis data according to Eq. (7) [30]:
Crude protein content %ð Þ ¼ N content %ð Þ 6:25 ð7Þ
Characterization by FTIR
The analysis were performed on a Shimadzu FTIR spec-trophotometer, model IRAffinity-1 using an attenuated total reflection (ATR) device with ZnSe crystal. The
readings included spectral range 800–4000 cm-1, 32 scans, 4 cm-1resolution, mode transmittance. In the spectra, the main stretches and angular deformations were identified.
Thermogravimetric characterization
Thermal analyzes were performed using a thermobalance SDT Q 600 from the manufacturer TA instruments, using approximately 10 mg sample and alumina crucible. The conditions used were inert atmosphere, via N2flow at the flow rate of 50 mL min-1, heating range of 30–800C and heating rate of 10C min-1.
Characterization by GC/MS of the saponifiable
fraction from CHF and CCF
Initially the samples were filtered using syringe filters with a porosity of 0.45 lm, the equipment used for the char-acterization was a chromatograph coupled to mass spec-trometer (GC/MS), brand Thermo Scientific model Focus GC and the column used was the NST-5MS with the dimensions of 30 m 9 0.25 mm 9 10 lm. The conditions of analysis include: 1.0 lL injection volume; drag gas flow of 1.5 mL min-1, injector temperature of 250C; interface temperature of 300C; ionization source of 250 C; ion-ization mode by electrons using energy of 70 eV for fragmentation; and scanning acquisition mode of com-pounds with m/z between 35 and 500. The data acquisition start time was 3.5 min. The temperature gradient started with maintaining the oven at 80C for 5 min. The tem-perature was raised at a rate of 5C min-1to 260C; then, the temperature was raised at a rate of 10C min-1 to 280 C when the temperature was held constant for a fur-ther 5 min. The identification of the compounds was per-formed by comparing the spectrum obtained with the databases of the NIST libraries versions 8.0.
Results and discussion
Extraction of lipid fractions characterization
of residual biomass by SEM
The values of the crude hexane fractions extracted using different times (CHF2h and CHF4h) as well as the chlo-roform fraction (CCF) obtained from the residual biomass are shown in Fig.1. The amount of CHF4h (10.54 ± 0.46)% is clearly superior to the content extrac-ted in CHF2h (2.95 ± 0.28)%. The performance employ-ing the system that couples two simultaneous effects, cavitation generated by ultrasonic bath sound waves and the shear provided by the mechanical stirrer blades, con-tributes synergistically to rupture of the cell wall of the
microalgae, consequently allowing the solvent to potentiate the entrainment of the intracellular materials in a shorter time, when compared to the use of the systems individually [31–34]. It is worth noting that the shearing action does not depend exclusively on the time, but also on the rotation used [20]. However, the use of high rotations may com-promise the microalgal biomass content to be recovered.
The lipid content extracted from BS and CTG groups was remarkably different. CHF4h and CCF samples had their obtained lipid fractions increased up to 47.62% and 65.40%, respectively. Therefore, there was a meaningful growth of approximately 58% in the extracted TLF* value (from 18.45% to 29.23%). Yang et al. [35] stated that the cultivation of Chlamydomonas reinhardtii sp. microalgae under nutrient reduction, specifically phosphorus and nitrogen, resulted in an increment of fatty acids content superior to 104% if compared to the control group. Ch-lamydomonas vulgaris cultivated under nitrogen restriction showed an increase of more than 150% in the lipid content [36,37].
After the biomass growth stage, the microalgae culti-vated under limitation or the absence of nutrients (espe-cially nitrogen) undergone processes such as nitrogen compounds degradation and substantial accumulation of lipids and carbohydrates [38]. Furthermore, Courchensne et al. [39] evidenced the correlation between lipid storage and nutrient restriction since it results in efficient accommodation of these molecules in small cel-lular compartments; in addition, it may also be used as energy reserve during adverse conditions for cell survival and proliferation. Chlamydomonas zofingiensis cultivated under conditions of nutritional stress showed that the nitrogen restriction is more effective to induce substantial accumulation of lipids when compared to phosphorus limitation [40].
The results reported for CHF2h and CHF4h are con-sistent with the images generated by the electron micro-scopy, where no differences between the morphological aspects presented by BBE and RBHF2h surfaces were observed (Fig.1a, b). However, there is significant
fragmentation of the biomass when comparing RBHF2h and RBHF4h (Fig.1c). This fact is evidenced by the sig-nificant reduction in the mean biomass fragment size (from 4.5 mm to 0.8 mm, respectively). Therefore, it is plausible to consider that the substantial difference between the lipid fractions extracted in the times of 2 and 4 h (more than doubling the value) is associated with a significant increase in the surface area in RBHF4h, which contributes to sol-vent diffusion and increased entrainment of intracellular material [41].
The images amplified at 250 and 2000 times of the surface of the BBE (Fig.2a, c) show a surface with rough texture and a cluster of cells, confirming that before the extractive process the cells have discrete damages [42]. However, the appearance of RBHF4h is considerably modified (Fig.2b, d), and the smooth surface supposedly indicates extravasation of intracellular material, justified by the expressive extraction of CHF4h.
The value of CCF extracted (18.69 ± 0.79)% was sig-nificantly higher, a fact that may be associated with the lipid composition of the fraction being composed mainly of more polar substances, including phospholipids and fatty acids, which favored the extraction with the most polar solvent [43]. In addition, the residual biomass also has a significant number of injured cells due to the first extrac-tion with the hexane, another factor that contributes to a greater drag of intracellular material (Table 1).
The microalgal biomass of the species Chlamydomonas sp. cultivated under conditions of nutrient limitation and open system showed a TLF (29.23 ± 0.62)% after 4 h of extraction with hexane followed by 1 h of extraction with chloroform, within the range of values reported in the lit-erature [44]. It is worth mentioning that the limitation of nutrients, especially nitrogen, has been shown to have the potential to substantially increase the lipid percentage in microalgae, since nitrogen deprivation induces the genes that encode enzymes directly involved in lipid biosynthesis [45]. The residual biomass at each fractionation stage was above 70%, and its reuse in sectors that add commercial value to its use is plausible.
Fig. 1 Micrographs obtained by SEM of the surface of the samples analyzed under a 50-fold increase. a BBE; bRBHF2h; c RBHF4h
Thermal characterization of the BBE microalgal
biomass, RBHF2h and RBHF4h
The TG curves show that the analyzed biomass presents three main thermal events (TEs) (see Table2). TE1 for BBE (in the range of 30.0–170C, maximum at 91.5 C) is simultaneous thermal events and ratifies the presence of moisture in the sample (up to approximately 120C) and the presence of low molecular mass carbohydrate frag-ments resulting from the processing of the biomass prior to extraction. TE2 (in the range of 170–380C, maximum 316C) refers to the volatilization of different microalgal constituents including cellulose and lignin, polymeric car-bohydrates that make up the cell wall of microalgae, pro-teins and, mainly, of lipid components such as fatty acids (up to 20 carbons) [46]. TE3 (380–600C) appears as a shoulder for BBE and RBHF2h, and as a well-defined peak for RBHF4h, events are reported as TAG and heavier phospholipids [47,48].
The biomass characterization by TG can make it pos-sible to estimate the average content (%) of TAG and other saponifiable lipid fractions (including fatty acids and phospholipids) by evaluating the results before and after the extraction process, providing a practical and fast way to size the extraction efficiency. The literature reports that the thermal events related to the lipid fraction (exclusively TAG) are between 350 and 450 C, events ratified as decomposition or volatilization of triacylglycerides [49,50].
The quantification of this region in terms of mass (%) provided a value of approximately 16.48%, higher than the maximum value of RBHF4h obtained during extractions that was 10.54 ± 0.46%, a result that shows the efficiency of the method considering the difficulty of extracting these materials. The DTG curve of RBHF4h shows a decrease in the fraction due to the presence of fatty acids (range from 170 to 380C, maximum at 316 C), compared to the BBE curve and an increase in the corresponding fraction of TAG
Fig. 2 Micrographies obtained by SEM of the surface of the samples analyzed under 250-fold increase: a BBE, bRBHF4h and 2000 times: cBBE, d RBHF4h
Table 1 Hexanic fractions extracted at different times, chloroform fraction obtained from residual biomass and total lipid fraction (TLF) Culture conditions CHF2h/% CHF4h/% CCF/% TLF*/% Control group (CTG) 2.59 ± 0.18 7.15 ± 0.14 11.30 ± 0.07 18.45 ± 0.10 Recovered biomass/% 91.04 ± 1.48 87.78 ± 2.34 80.67 ± 2.90 Under stress (BS) 2.95 ± 0.28 10.54 ± 0.46 18.69 ± 0.79 29.23 ± 0.29 Recovered biomass/% 90.34 ± 1.98 85.45 ± 2.58 75.85 ± 3.36
(380 at 450C). This probably occurs because after removal of the fatty acids, shown by the decrease in the RBHF4h curve compared to the BBE curve, the TAG fraction becomes proportionally larger in the amount of material analyzed.
Note also that there is a shift of the peaks of events I and II to lower values of temperature indicating an evolution over time in the preferential extraction of lower molecular mass materials because they are more easily dragged. It is possible to infer that the observed behavior is due to RBHF4h being more concentrated of less volatile compo-nents, as there is a notable reduction in mass loss in the TE3 of RBHF4h, a characteristic region of the saponifiable lipid fraction. It is important to note that several authors have been using the technique for quantitative purposes and obtaining relevant results [50]. The remaining residues of the pyrolysis of each biomass (Fig.3and Table2) show a consistent relationship between the above results with the content of CHF4h extracted from the microalgae. The increase in the time of extraction causes a significant increase in the lipid fraction obtained; associated with this fact, an increase in the content of generated residues is also observed, which is coherent and explainable by the breaking of the material seen in the SEM and that tends to increase residues.
Elementary composition of microalgal biomass
The elemental analysis (see Table3) of the biomass before the extractive processes (BBE) revealed a composition rich in carbon (47.11%), a value consistent with crops under nutrient deprivation [44], because the above-mentioned conditions reduce the rate of microalgal metabolism;consequently, the excess carbon generated is directed to energy storage, mainly in the form of lipids (TAG and phospholipids) and carbohydrates, substances mainly composed of carbon and hydrogen [44]. The elemental composition of BBE still counts on hydrogen (8.01%), nitrogen (9.01%), sulfur (0.45%) and oxygen (35.37%). The crude protein content estimated for BBE was around 56%, a relatively high value for crops under nutrient lim-itation, as nitrogen scarcity reduces protein synthesis in order to adapt the cells to lower availability of amino acids [51, 52]. However, microalgae of the genus Chlamy-domonas have been shown to be less sensitive to crops under nitrogen deprivation, so the behavior is subject to oscillations depending on the species analyzed [53].
The residual biomasses RBHF4h and RBCF (residual biomass after extraction with hexane and chloroform) revealed an elemental composition with a significant reduction in the percentages of carbon and hydrogen; the result indicates that the extractive processes removed the lipids for the most part. It is worth noting that RBHF and RBCF have significant nitrogen content indicating a residual biomass with good protein potential.
The remarkable value of crude protein, associated with a relatively high recoverable biomass content (above 70%) even after extracting apolar fractions, makes algal biomass a source with potential for animal supplementation (mainly
20 – 4.5 – 3.6 – 2.7 – 1.8 – 0.9 0 DTG/°C –1 120 220 320 420 520 620 720 820 DTG-BBE DTG-RBHF2h DTG-RBHF4h TG-BBE TG-RBHF2h TG-RBHF4h 20 20 Mass/% 120 (a) (b) 220 320 420 520 Temperature/°C Temperature/°C 620 720 820 0 40 60 80 100
Fig. 3 TG (a) and DTG (b) curves of the biomass BBE, RBHF2h and RBHF4h
Table 2 Thermal event records for the biomasses BBE, RBHF2h and RBHF4h
Samples Thermal events Temperature/C Mass/%
BBE 1 30–170 9.87 2 170–380 42.85 3 380–600 20.62 Residue/% [ 600 26.66 RBHF2h 1 30–164 11.91 2 170–380 41.75 3 380–600 18.58 Residue/% [ 600 27.76 RBHF4h 1 30–170 12.32 2 170–380 31.93 3 380–600 21.11 Residue/% [ 600 34.64
aquaculture) and cosmetic industries due to its nutritional value related to the presence of proteins, pigments and vitamins [10, 11]. However, microalgal cultivation still needs adaptations established by the Food and Drug Administration (FDA) agency, aiming to ensure the safety in the use of algae extracts in humans [54]. The use of algal biomass associated with biofuel and fine chemistry industry is an imminent alternative to make the production cost feasible since these microorganisms can be used to syn-thesize compounds of high commercial value, such as polyunsaturated fatty acids, natural antioxidants and sub-stances with bioactive potential [55,56]. For this purpose, extraction procedures that allow compounds recovery are required, considering energy consumption and viability of the processing.
The HHV for the dried biomasses BBE, RBHF4h and RBCF was 20.16, 19.36 and 18.48 MJ kg-1, respectively, values that are higher than the values of other microalgae reported in the literature as Scenedesmus obliquus (16.69 MJ kg-1) and Dunaliella tertiolecta (11.66 MJ kg-1) [57, 58]. The calorific value is an important parameter to measure the presence of metabo-lites such as lipids and carbohydrates, molecules respon-sible for storing much of the energy content present in the microalgal biomass.
Characterization of extracts (CHF and CCF)
and their saponifiable fractions
Spectroscopic for FTIR
The FTIR spectra (see Fig.4a, b) show similarities between the CHF and CCF crude extracts, as well as the presence of the strains referring to the C–H (sp3–s) axial deformations near 2922 cm-1, but also C–H near 1462 cm-1, characteristic of methylene groups (CH2-), expected absorptions for lipid fractions, mostly composed of carbon and hydrogen. In addition, there is a band with maximum absorption at approximately 3400 cm-1, which, plausibly, can be attributed to O–H or N–H stretches, which may be subject to signal coalescence due to heterogeneity of the extracts [59]. The stretches occurring in the range of 1600-1800 cm1are mainly associated with different types of C=O axial formations. The CHF extract
shows stretches at 1670 and 1710 cm-1 referring to C=O of amide I and fatty acids, respectively. CCF includes signals related to C=O deformations of dimerized fatty acids (1709 cm-1) and esters (1739 cm-1), most likely indicating the presence of TAG and phospholipids, mate-rials that can be saponified.
The FTIR spectra shown in Fig.4c, d confirm the SF conversion of both extracts to fatty acids, as the spectra show the presence of approximate stretches of 1710 cm-1relative to acid carbonyl, such as a widening in the band relative to the axial deformation OH (about 3200–2500 cm-1). The esterification of the saponifiable fractions is confirmed by the shift in absorption of the C=O draw from 1710 cm-1 to approximately 1743 cm-1(attributed to ester C=O) and by the appearance of signals at approximately 1095 cm-1 associated with the draw C–O–C, also, attached to the ester group, as indicated in Fig.4e, f [60].
Thermal analysis (TG/DTG)
The TG curves show significant differences in the com-position of the extracts CCF and CHF, Fig.5a, b, respec-tively. The CCF has three main TE, with TE1 (30–318C) referring to the vaporization of water molecules (at about 100 C), an event that also occurs for CHF. It is worth mentioning that TE2 (main event) for CHF (100–194C, maximum 117.7C) occurs simultaneously with TE1, mainly referring to low molecular mass residues, suppos-edly due to the degradation of pigments (mainly chloro-phyll and carotenoids), a fact common in cultures under stress conditions [61]. Still the TE1 from CCF in the temperature range of 100–318C (maximum at 262.6 C) and TE3 for CHF (194–260C with maximum at 242.4C) are mainly related to the volatilization of unsaturated fatty acids such as linolenic acid (C18:3) and palmitic acid (C16:1) and saturated carbon chain of less than C16[51].
The TE3 for CCF (318–600 C, two simultaneous events) includes up to 370C the volatilization of saturated fatty acids (C16–C18), including stearic and oleic acids, common in several species of microalgae grown under conditions of nutrient limitation [30, 44,45]. In addition, prolongation of the thermal event (above 370C) may be associated with volatilization and decomposition of heavier lipid fractions such as TAG and phospholipids [62]. The TE4 for CHF (260–600 C with maximum at 318 C) is associated with the substances mentioned above for the TE3 of the CCF.
The TG curves of the saponifiable fractions after ester-ification (Fig. 6a, b) show samples with a single major thermal event, as also indicated in Table4. TE2 for SF_CHF-Esterified (range 100–300 C, maximum at 213.4C) and SF_CCF-Esterified (range 100–300 C,
Table 3 Elemental compositions and higher heating value (HHV) of biomasses before extractive processes (BBE) and after extractions with hexane (RBHF4h) and hexane followed by chloroform (RBCF)
Samples C/% H/% N/% S/% O/% HHV/MJ Kg-1
BBE 47.11 8.01 9.01 0.45 35.37 20.16 RBHF4h 45.45 7.74 9.22 0.48 37.11 19.36 RBCF 43.24 7.47 9.97 0.52 38.81 18.48
maximum at 218.9C) refer to the main products obtained in the esterification and the small peaks above 300C which complete losses for CHF are due to the minority presence of compounds of high molecular mass.
The reduction in the thermal stability of the esterified samples, when compared to the respective crude extracts, is an evidence corroborating the formation of MME [50,63,64]. The saponifiable fraction includes mainly fatty acids, TAGs and phospholipids which are more thermally stable. It is worth mentioning that the presence of a
shoulder (region close to 90C) in the DTG of the ther-mogravimetric curve for SF_CHF-Esterified shows a more heterogeneous composition for the sample, probably linked to SF being from a less processed biomass. In addition, even with successive washing steps it is plausible that the drag of unsaponifiable material occurs. The SF_CCF-Es-terified TG curve aspect indicates a more homogeneous product (DTG curve less expanded) and with significant conversion of SF to MME (approximately 89%). The results confirm that the CCF concentrates most of the SF in
800 1300 1800 2300 2800 Wavenumber/cm–1 3300 Transmitance/% Transmitance/% Transmitance/% Transmitance/% Transmitance/% Transmitance/% 3800 800 1300 1800 2300 2800 Wavenumber/cm–1 3300 3800 800 1300 1800 1710 3380 1741 1710 1710 1710 1739 1740 1094 1097 3380 2300 2800 Wavenumber/cm–1 3300 3800 (a) (c) (d) (e) (f) (b) 800 1300 1800 2300 2800 Wavenumber/cm–1 3300 3800 800 1300 1800 2300 2800 Wavenumber/cm–1 3300 3800 800 1300 1800 2300 2800 Wavenumber/cm–1 3300 3800
Fig. 4 FTIR spectra of the samples of CHF (a), CCF (b), saponified CHF (c), saponified CCF (d), saponified CHF after esterification (e) and saponified CCF after esterification (f)
the lipid material extracted from the microalgae, and the data are consistent with the TG curves shown by the crude extracts.
Characterization of the saponifiable fraction by GC/MS
The results of GC/MS revealed a significant percentage of substances that differ from the MME, especially for sample SF_CHF-Esterified as indicated in Table5. The most rep-resentative fraction of the above compounds includes the phytol and phytol acetate with 34.64% and 18.48%, respectively; such substances are derived from the degra-dation process of chlorophyll a and b (nitrogenated pig-ments), common in microalgae of the genus Chlamydomonas [61]. In addition, microalgal biomass when grown under nitrogen constraints tends to increase the rate of enzymes responsible for the degradation of organic nitrogen in the cells [65]. Therefore, microalgal cells could plausibly degrade nitrogenous pigments to balance cellular metabolic functions [31]. However, the MME obtained (31.24%) consists mainly of methyl linoleate (14.58%) and methyl palmitate (15.21%).
The substances exhibited by GC/MS for sample SF_CCF-Esterified (Table6) are predominantly MME (86.31%), the value being consistent with thermogravi-metric conversion data (89.10%). The composition includes the MME methyl 4Z,7Z,10Z,13Z hexadecate-traenoate (10.87%), methyl palmitoleate (6.73%), methyl palmitate (23.02%) and methyl linolenate (45.68%). Therefore, it is mainly composed of unsaturated fatty acids (approximately 73%). The results of the microalgal fatty profile are similar to the data reported in the literature for species under nutrient limitation that exhibit C16:0 satu-rated fatty acid (palmitic acid) and C18:3 polyunsatusatu-rated fatty acid (linolenic acid, x-3 and x-6) as predominant, respectively [66–68]. 20 120 220 320 420 520 620 720 820 – 0.6 – 0.5 – 0.4 – 0.3 – 0.3 – 0.2 – 0.2 – 0.25 – 0.35 – 0.1 – 0.1 – 0.15 0 DTG/1/°C DTG/1/°C 0 – 0.05 Temperature/°C 20 120 220 320 420 520 620 720 820 TG_CHF TG_CCF DTG_CHF DTG_CCF Temperature/°C 20 Mass/% (a) (b) 0 40 60 80 100 20 Mass/% 0 40 60 80 100
Fig. 5 TG and DTG curves of crude extracts CCF (a) and CHF (b)
20 120 220 320 420 520 620 720 820 Temperature/°C 20 120 220 320 420 520 TG-SF_CCF-Esterified TG-SF_CHF-Esterified DTG-SF_CHF-Esterified DTG-SF_CCF-Esterified 620 720 820 Temperature/°C 20 Mass/% (b) 0 40 60 80 100 20 Mass/% (a) 0 40 60 80 100 – 0.3 – 0.4 – 0.45 – 0.2 – 0.25 – 0.35 – 0.1 – 0.15 DTG/1/°C 0 – 0.05 – 0.3 – 0.2 – 0.25 – 0.1 – 0.15 DTG/1/°C 0 – 0.05
Fig. 6 Esterified saponifiable fractions feed SF_CCF-Esterified (a) and SF_CHF-Esterified (b)
Table 4 Data extracted from the TG curves of crude hexane (CHF), esterified saponifiable (SF_CHF-Esterified) from CHF, crude chlo-roform (CCF) and esterified saponifiable fraction (SF_CCF-Esteri-fied) from CCF
Samples Thermal events Temperature/C Mass/%
CHF 1 30–194 62.33 2 194–260 15.55 3 260–600 20.36 Residue/% [ 600 1.76 SF_CHF-Esterified 1 30–100 5.64 2 100–600 90.63 Residue/% [ 600 3.73 CCF 1 30–318 40.87 2 and 3 318–600 46.23 Residue/% [ 600 12.90 SF_CCF-Esterified 1 30–100 1.36 2 100–600 89.1 Residue/% [ 600 9.54
The SF_CCF-Esterified from BS showed a slight growth on the conversion rate, a minor diversity of substances and 36.48% increment of linolenic acid in comparison with CTG. Yang et al. [62] verified that Chlamydomonas rein-hardtii cultivation under nitrogen and phosphorous restriction resulted in a notable rise of polyunsaturated fatty acids (specially c-linolenic acid); however, C18:0 e C18:2 acids were not detected in the BS sample. Moreover, there were no changes in the predominant fat profile for both conditions analyzed and the major fatty acids remained to be C16:0 e C18:3, which represent more than 70% of total ester composition.
The ester profile with relatively elevated concentration of polyunsaturated fatty acids (C18:3) is unfeasible in their application for biodiesel production, for which the content should be under 12% in order to avoid a low oxidative stability of the biofuel. Nevertheless, the fatty profile of microalgae under nutritional stress conditions shows
pharmacological potential for diet or therapy, since some of the main components such as arachidonic, linolenic and eicosapentaenoic acids are reported as bioactive metabo-lites [55,69].
Conclusions
The comparative micrographs of the surfaces BBE, RBHF2h and RBHF4h demonstrated that the extractive method, mechanical stirred coupled to the sonication sys-tem, promotes greater fragmentation of the biomass in the time of 4 h, allowing greater extraction of CCF. The TG curves for the biomasses showed compatibility with the aforementioned data, evidencing significant reductions in mass loss for RBHF4h, in the temperature range charac-teristic of lipids. The total lipid fraction value for biomass cultivated under stress had an increment of 65% when
Table 5 Substances identified in SF_CHF-Esterified by GC/ MS
Substances Retention time/min Peak area/%
2,6,11-Trimethyldodecane 29.951 6.10 3,7,11,15-Tetramethyl-2-hexadecene 32.743 5.90 Methyl palmitoleate (C16:1) 34,588 1.45 Methyl palmitate (C16:0) 34.700 15.21 Phytol acetate 35.487 18.48 Neophytadiene 36.813 3.64 Phytol 37.369 34.64 Methyl linolenate (C18:3) 38.091 14.58
Methyl monoesters (MME) Several 31.24
Other substances Several 68.76
Table 6 Substances identified in SF_CCF-Esterified (BS)1e SF_CCF-Esterified (CTG)2by GC/MS
Substances Retention time/min 1Peak area/% 2Peak area/%
Butylated hydroxytoluene 25.380 Absent 4.74
Hydroxylamine, O-decyl 29.947 Absent 1.66
Methyl 4Z,7Z,10Z,13Z hexadecatetraenoate 33.861 10.87 5.87 Methyl 12,13 tetradecadienoate 34.135 Absent 3.46
Methyl myristate (C14:0) 34.205 Absent 3.58
Methyl palmitoleate (C16:1) 34.588 6.73 0.51
Methyl palmitate (C16:0) 34.700 23.02 24.68
Phytol acetate 35.487 1.92 0.99
Neophytadiene 36.813 0.96 1.41
Phytol 37.369 10.80 8.62
Methyl stearate (C18:0) 37.762 Absent 1.17
Methyl linoleate (C18:2) 38.056 Absent 9.84
Methyl linolenate (C18:3) 38.091 45.68 33.47
Methyl monoesters (MME) Several 86.31 82.58
MME (saturated) Several 23.03 28.28
MME (unsaturated) Several 63.28 54.32
compared to the control group. The high recoverability of the residual biomass associated with the crude protein content allows several applications, such as nutritional supplementation in aquaculture, as well as the subsequent evaluation of more polar fractions for use in fine chemistry. The TG data for the CCF extract reveal a material con-stituted mainly by saponifiable fractions, unlike the behavior exhibited by CHF TG curves, predominantly composed of unsaponifiables. Consequently, the TG curve for SF_CCF-Esterified reveals a better conversion of the sample to MME (about 89%). The saponifiable fractions, practically, did not present significant changes in the MME obtained, being the predominant fatty acids—the C16:0 and C18:3 (x-3 and x-6); however, the SF_CCF-Esterified fraction exhibited a higher conversion rate (86.31%) when compared to the SF_CHF-Esterified sample (31.24%) and the CTG (82.58%). Finally, the ester profile with relatively elevated concentration of polyunsaturated fatty acids (C18:3) is unfeasible in their application for biodiesel production; nevertheless, the fatty profile of microalgae suggests pharmacological potential for diet or therapy, since some of the main components are reported as bioactive metabolites.
References
1. Morioca C. Floculac¸a˜o de Chlorella sp. produzida em concen-trado de dessalinizac¸a˜o e estudo de me´todo de extrac¸a˜o de lipı´-deos intracelulares. Quı´mi Nova. 2014. https://doi.org/10.1590/
s0100-40422014000100009.
2. Markou G, Nerantzis E. Microalgae for high-value compounds and biofuels production: a review with focus on cultivation under stress conditions. Biotechnol Adv. 2013.https://doi.org/10.1016/
j.biotechadv.2013.07.011.
3. Trivedi J, Aila M, Bangwal DP, Kaul S, Garg MO. Algae based biorefinery—how to make sense? Renew Sustain Energy Rev. 2015.https://doi.org/10.1016/j.rser.2015.03.052.
4. Ahmad AL, Yasin NHM, Derek JK, Lim K. Microalgae as a sustainable energy source for biodiesel production: a review. Renew Sustain Energy Rev. 2011.https://doi.org/10.1016/j.rser.
2010.09.018.
5. Pereira CMP, Hobuss CB, Maciel JV, Ferreira LR, Del Pino FB, Mesko MF, Jacob LE, Neto PC. Biodiesel renova´vel derivado de microalgas: avanc¸os e perspectivas tecnolo´gicas. Quı´mi Nova. 2012.https://doi.org/10.1590/s0100-40422012001000022. 6. Abdul RWAKG, Wan AYH, Taufiq Y, Michael KD, Razif H.
Thermochemical conversion of microalgal biomass for biofuel production. Renew Sustain Energy Rev. 2015.https://doi.org/10.
1016/j.rser.2015.04.186.
7. Nakanishi A, Aikawa S, Ho S, Chen C, Chang J, Hasunuma T, Kondo A. Development of lipid productivities under different CO2 conditions of marine microalgae Chlamydomonas sp. JSC4. Bioresour Technol. 2014.https://doi.org/10.1016/j.biortech.2013.
11.009.
8. Kato Y, Ho S, Vavricka CJ, Chang JS, Hasunuma T, Kondo A. Evolutionary engineering of salt-resistant Chlamydomonas sp. strains reveals salinity stress-activated starch-to-lipid
biosynthesis switching. Bioresour Technol. 2017.https://doi.org/
10.1016/j.biortech.2017.06.035.
9. Magali S, Ste´phan C, Caroline C, Boris F, Mai N, Patrick C, Audrey B, Fred B, Christian T, Yonghua L, Gilles P. Oil accu-mulation in the model green alga Chlamydomonas reinhardtii: characterization, variability between common laboratory strains and relationship with starch reserves. BMC Biotechnol. 2011.
https://doi.org/10.1186/1472-6750-11-7.
10. Zhu L. The combined production of ethanol and biogas from microalgal residuals to sustain microalgal biodiesel: a theoretical evaluation. Biofuels Bioprod Biorefining. 2014. https://doi.org/
10.1002/bbb.1442.
11. Zhu L. Biorefinery as a promising approach to promote microalgae industry: an innovative framework. Renew Sustain Energy Rev. 2015.https://doi.org/10.1016/j.rser.2014.09.040. 12. Chernova NI, Kiseleva SV. Microalgae biofuels: induction of
lipid synthesis for biodiesel production and biomass residues into hydrogen conversion. Int J Hydrogen Energy. 2017.https://doi.
org/10.1016/j.ijhydene.2016.05.302.
13. Demirbas A. Biomass resource facilities and biomass conversion processing for fuels and chemicals. Energy Convers Manag. 2001.https://doi.org/10.1016/s0196-8904(00)00137-0.
14. Nobre BP, Villalobos F, Barraga´n BE, Oliveira AC, Batista AP, Marques PASS, Mendes RL, Sovova´ H, Palavra AF, Gouveia L. A biorefinery from Nannochloropsis sp microalga—Extraction of oils and pigments Production of biohydrogen from the leftover biomass. Bioresour Technol. 2013.https://doi.org/10.1016/j.bior
tech.2012.11.084.
15. Subhadra B, Grinson G. Algal biorefinery-based industry: an approach to address fuel and food insecurity for a car-bon-smart world. J Sci Food Agric. 2011.https://doi.org/10.1002/jsfa.4207. 16. Vanthoor-Koopmans M, Wijffels RH, Barbosa MJ, Eppink MHM. Biorefinery of microalgae for food and fuel. Bioresour Technol. 2013.https://doi.org/10.1016/j.biortech.2012.10.135. 17. Service RF. Algae’s second try. Science. 2011.https://doi.org/10.
1126/science.333.6047.1238.
18. Araujo GS, Matos LJBL, Fernandes JO, Cartaxo SJM, Gonc¸alves LRB, Fernandes FAN, Farias WRL. Extraction of lipids from microalgae by ultrasound application: prospection of the optimal extraction method. Ultrason Sonochem. 2013.https://doi.org/10.
1016/j.ultsonch.2012.07.027.
19. Lee J, Yoo C, Jun SY, Ahn C, Oh H. Comparison of several methods for effective lipid extraction from microalgae. Bioresour Technol. 2010.https://doi.org/10.1016/j.biortech.2009.03.058. 20. Gomes AF. Extrac¸a˜o e ana´lise da frac¸a˜o lipı´dica da microalga
Monoraphidium sp., sı´ntese e caracterizac¸a˜o do seu biodiesel. Dissertac¸a˜o de Mestrado, Universidade Federal do Rio Grande do Norte, Brasil. 2013. https://repositorio.ufrn.br/jspui/bitstream/
123456789/17702/1/AndersonFG_DISSERT.pdf. Accessed 15
oct 2018.
21. Bigogno C, Khozin-Goldberg I, Boussiba S, Vonshak A, Cohen Z. Lipid and fatty acid composition of the green oleaginous alga Parietochloris incisa, the richest plant source of arachidonic acid. Phytochemistry. 2002.
https://doi.org/10.1016/s0031-9422(02)00100-0.
22. Ferruzzi MG, Blakeslee J. Digestion, absorption, and cancer preventative activity of dietary chlorophyll derivatives. Nutr Res. 2007.https://doi.org/10.1016/j.nutres.2006.12.003.
23. Chew KW, Yap JY, Show PL, Suan HN, Juan JC, Ling TC, Lee DJ, Chang JS. Microalgae biorefinery: high value products per-spectives. Bioresour Technol. 2017. https://doi.org/10.1016/j.
biortech.2017.01.006.
24. Demirbas A. Oily products from mosses and algae via pyrolysis. Energ Source Part A. 2007. https://doi.org/10.1080/
25. Zhao Z, Liu P, Wang S, Ma S, Cao J. Combustion characteristics and kinetics of five tropic oilgal strains using thermogravimetric analysis. J Therm Anal Calorim. 2018. https://doi.org/10.1007/
s10973-017-6646-0.
26. Paniagua S, Calvo LF, Escapa C, Coimbra RN, Otero M, Garcı´a AI. Chlorella sorokiniana thermogravimetric analysis and com-bustion characteristic indexes estimation. J Therm Anal Calorim. 2018.https://doi.org/10.1007/s10973-017-6734-1.
27. Brown MR, Jeffrey SW, Volkman JK, Dunstan GA. Nutritional properties of microalgae for mariculture. Aquaculture. 1997.
https://doi.org/10.1016/s0044-8486(96)01501-3.
28. Chuntapa B, Powtongsook S, Menasveta P. Water quality control using Spirulina platensis in shrimp culture tanks. Aquaculture. 2003.https://doi.org/10.1016/s0044-8486(02)00428-3.
29. Friedl A, Padouvas E, Rotter H, Varmuza K. Prediction of heating values of biomass fuel from elemental composition. Anal Chim Acta. 2005.https://doi.org/10.1016/j.aca.2005.01.041. 30. Becker EW. Microalgae: biotechnology and microbiology.
Cambridge: Cambridge University Press; 1994.
31. Toma M, Vinatoru M, Paniwnyk L, Mason TJ. Investigation of the effects of ultrasound on vegetal tissues during solvent extraction. Ultrason Sonochem. 2001. https://doi.org/10.1016/
s1350-4177(00)00033-x.
32. Jaouen P, Vandanjon L, Que´me´neur F. The shear stress of microalgal cell suspensions (Tetraselmis suecica) in tangential flow filtration systems: the role of pumps. Bioresour Technol. 1999.https://doi.org/10.1016/s0960-8524(98)00144-8.
33. Conney M, Young G, Nagle N. Extraction of bio-oils from microalgae. Sep Purif Rev. 2009. https://doi.org/10.1080/
15422110903327919.
34. Shirsath SR, Sonawane SH, Gogate PR. Intensification of extraction of natural products using ultrasonic irradiations—A review of current status. Chem Eng Process. 2012.https://doi.org/
10.1016/j.cep.2012.01.003.
35. Yang L, Chen J, Qin S, Zeng M, Jiang Y, Hu L, Wang J. Growth and lipid accumulation by different nutrients in the microalga Chlamydomonas reinhardtii. Biotechnol Biofuels. 2018.https://
doi.org/10.1186/s13068-018-1041-z.
36. Converti A, Casazza AA, Ortiz EY, Perego P, Borghi MD. Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem Eng Process. 2009.https://doi.org/10.
1016/j.cep.2009.03.006.
37. Mujtaba G, Choi W, Lee CG, Lee K. Lipid production by Chlorella vulgaris after a shift from nutrient-rich to nitrogen starvation conditions. Bioresour Technol. 2012. https://doi.org/
10.1016/j.biortech.2012.07.057.
38. Simionato D, Block MA, La Rocca N, Jouhet J, Mare´chal E, Finazzi G, Morosinotto T. The response of Nannochloropsis gaditana to nitrogen starvation includes de novo biosynthesis of triacylglycerols, a decrease of chloroplast galactolipids, and reorganization of the photosynthetic apparatus. Eukaryot Cell. 2013.https://doi.org/10.1128/ec.00363-12.
39. Courchesne NMD, Parisien A, Wang B, Lan CQ. Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. J Biotechnol. 2009. https://doi.
org/10.1016/j.jbiotec.2009.02.018.
40. Feng P, Deng Z, Fan L, Hu Z. Lipid accumulation and growth characteristics of Chlorella zofingiensis under different nitrate and phosphate concentrations. J Biosci Bioeng. 2012.https://doi.
org/10.1016/j.jbiosc.2012.05.007.
41. Ansari FA, Gupta SK, Nasr M, Rawat I, Bux F. Evaluation of various cell drying and disruption techniques for sustainable metabolite extractions from microalgae grown in wastewater: a multivariate approach. J Clean Prod. 2018. https://doi.org/10.
1016/j.jclepro.2018.02.098.
42. Pohndorf RS, Camara AS, Larrosa APQ, Pinheiro CP, Strieder MM, Pinto LAA. Production of lipids from microalgae Spirulina sp: influence of drying, cell disruption and extraction methods. Biomass Bioenergy. 2016. https://doi.org/10.1016/j.biombioe.
2016.06.020.
43. Luque R, Herrero-Davila L, Campelo JM, Clark JH, Hidalgo JM, Luna D, et al. Biofuels: a technological perspective. Energy Environ Sci. 2008.https://doi.org/10.1039/b807094f.
44. Shekh AY, Shrivastava P, Krishnamurthi K, Mudliar SN, Sara-vana Devi S, Kanade GS, Chakrabarti T. Stress enhances poly-unsaturation rich lipid accumulation in Chlorella sp. and Chlamydomonas sp. Biomass Bioenergy. 2016. https://doi.org/
10.1016/j.biombioe.2015.11.013.
45. Amoah J, Ho SH, Hama S, Yoshida A, Nakanishi A, Hasunuma T, Ogino C, Kondo A. Conversion of Chlamydomonas sp JSC4 lipids to biodiesel using Fusarium heterosporum lipase-express-ing Aspergillus oryzae whole-cell as biocatalyst. Algal Res. 2017.
https://doi.org/10.1016/j.algal.2017.10.003.
46. Fu L, Cui X, Li Y, Xu L, Zhang C, Xiong R, Zhou D, Crittenden JC. Excessive phosphorus enhances Chlorella regularis lipid production under nitrogen starvation stress during glucose het-erotrophic cultivation. Chem Eng J. 2017. https://doi.org/10.
1016/j.cej.2017.07.182.
47. Chen W, Huang M, Chang J, Chen C. Thermal decomposition dynamics and severity of microalgae residues in torrefaction. Bioresour Technol. 2014.https://doi.org/10.1016/j.biortech.2014.
06.086.
48. Campanella A, Muncrief R, Harold MP, Griffith DC, Whitton NM, Weber RS. Thermolysis of microalgae and duckweed in a CO2-swept fixed-bed reactor: bio-oil yield and compositional effects. Bioresour Technol. 2012. https://doi.org/10.1016/j.bior
tech.2011.12.115.
49. Garcia CC, Franco PIBM, Zuppa TO, Antoniosi Filho NR, Leles MIG. Thermal stability studies of some cerrado plant oils. J Therm Anal Calorim. 2007.
https://doi.org/10.1007/s10973-006-7769-x.
50. Gomes AF, Gomes MPS, Souza DL, Costa M. Thermal and chromatographic characterization of biomass, lipid material, and microalgal biodiesel from Monoraphidium sp. J Therm Anal Calorim. 2015.https://doi.org/10.1007/s10973-014-43475. 51. Chand P, Reddy V, Verkade JG, Wang T, Grewell D.
Thermo-gravimetric quantification of biodiesel produced via alkali cat-alyzed transesterification of soybean oil. Energy Fuels. 2009.
https://doi.org/10.1021/ef800668u.
52. Anand J, Arumugam M. Enhanced lipid accumulation and bio-mass yield of Scenedesmus quadricauda under nitrogen starved condit ion. Bioresour Technol. 2015. https://doi.org/10.1016/j.
biortech.2014.12.097.
53. Zhu S, Huang W, Xu J, Wang, Xu J, Yuan Z. Metabolic changes of starch and lipid triggered by nitrogen starvation in the microalga Chlorella zofingiensis. Bioresour Technol. 2014.
https://doi.org/10.1016/j.biortech.2013.10.092.
54. Chew KW, Yap JY, Show PL, Suan NH, Juan JC, Ling TC, Lee D, Chang J. Microalgae biorefinery: high value products per-spectives. Bioresour Techenol. 2017. https://doi.org/10.1016/j.
biortech.2017.01.006.
55. Banskota AH, Stefanova R, Sperker S, Lall S, Craigie JS, Hafting JT. Lipids isolated from the cultivated red alga Chondrus crispus inhibit nitric oxide production. J Appl Phycol. 2014.https://doi.
org/10.1007/s10811-013-0174-5.
56. Vaz BS, Moreira JB, Morais MG, Costa JAV. Microalgae as a new source of bioactive compounds in food supplements. Curr Opin Food Sci. 2016.https://doi.org/10.1016/j.cofs.2015.12.006. 57. Dean AP, Sigee DC, Estrada B, Pittman JK. Using FTIR spec-troscopy for rapid determination of lipid accumulation in response to nitrogen limitation in freshwater microalgae.
Bioresour Technol. 2010.https://doi.org/10.1016/j.biortech.2010.
01.065.
58. Kim SS, Ly HV, Kim J, Lee EY, Woo HC. Pyrolysis of microalgae residual biomass derived from Dunaliella tertiolecta after lipid extraction and carbohydrate saccharification. Chem Eng J. 2015.https://doi.org/10.1016/j.cej.2014.11.045.
59. Ido AL, Luna MDG, Capareda SC, Maglinao AL, Nam H. Application of central composite design in the optimization of lipid yield from Scenedesmus obliquus microalgae by ultrasound-assisted solvent extraction. Energy. 2018.https://doi.org/10.1016/
j.energy.2018.04.171.
60. Wang J, Ma X, Yu Z, Peng X, Lin Y. Studies on thermal decomposition behaviors of demineralized low-lipid microalgae by TG-FTIR. Thermochim Acta. 2018.https://doi.org/10.1016/j.
tca.2018.01.001.
61. Li F, Srivatsa SC, Batchelor W, Bhattachary S. A study on growth and pyrolysis characteristics of microalgae using Ther-mogravimetric Analysis-Infrared Spectroscopy and synchrotron Fourier Transform Infrared Spectroscopy. Bioresour Technol. 2017.https://doi.org/10.1016/j.biortech.2017.01.005.
62. Dweck J, Sampaio CMS. Analysis of thermal decomposition of comercial vegetable oils in air simultaneous TG/DTA. J Therm Anal Calorim. 2004. https://doi.org/10.1023/b:jtan.0000027124.
96546.0f.
63. Goodrum JW. Volatility and boiling points of biodiesel from vegetable oils and tallow. Biomass Bioenergy. 2002.https://doi.
org/10.1016/s0961-9534(01)00074-5.
64. Chen WT, Chen WC, Ma CM, Laiwang B, Shen SJ, You ML, Shu CM. Structural characteristics and decomposition analyses of four commercial essential oils by thermal approaches and GC/
MS. J Therm Anal Calorim. 2018. https://doi.org/10.1007/
s10973-017-6594-8.
65. Deschoenmaeker F, Bayon-Vicente G, Sachdeva N, Depraetere O, Pino JCC, Leroy B, Muylaert K, Wattiez R. Impact of dif-ferent nitrogen sources on the growth of Arthrospira sp. PCC 8005 under batch and continuous cultivation—A biochemical, transcriptomic and proteomic profile. Bioresour Technol. 2017.
https://doi.org/10.1016/j.biortech.2017.03.145.
66. Pinho DMM, Oliveira RS, Santos VML, Marques WF, Pinto AC, Rezende, Michelle JC, Paulo AZ. Evaluating the potential of biodiesel production through microalgae farming in photobiore-actor and high rate ponds from wastewater treatment. J Braz Chem Soc. 2017.https://doi.org/10.21577/0103-5053.20170097. 67. Chen M, Tang H, Ma H, Holland TC, Simon KY, Salley SO. Effect of nutrients on growth and lipid accumulation in the green algae Dunaliella tertiolecta. Bioresour Technol. 2011.https://doi.
org/10.1016/j.biortech.2010.09.062.
68. O¨ rdo¨g V, Stirk WA, Ba´lint P, Aremu AO, Okem A, Lova´sz C, Molna´r Z, Staden JV. Effect of temperature and nitrogen con-centration on lipid productivity and fatty acid composition in three Chlorella strains. Algal Res. 2017.https://doi.org/10.1016/j.
algal.2016.03.001.
69. Sathasivam R, Radhakrishnan R, Hashem A, Abd Allah EF. Microalgae metabolites: a rich source for food and medicine. Saudi J Biol Sci. 2017. https://doi.org/10.1016/j.sjbs.2017.11.
003.
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