Nannochloropsis and Nannochloris Isolates with Potential for Biofuel Production
Alan Rodrigo López-Rosales1, Katia Ancona-Canché1, Juan Carlos Chavarria-Hernandez1, Felipe Barahona-Pérez1, Tanit Toledano-Thompson1, Gloria Garduño-Solórzano2,
Silvia López-Adrian3, Blondy Canto-Canché4, Erik Polanco-Lugo5and Ruby Valdez-Ojeda1,*
1 Unidad de Energía Renovable, Centro de Investigación Científica de Yucatán A.C., Parque Científico y Tecnológico del Estado de Yucatán, Carretera Sierra Papacal-ChuburnáPuerto, Yucatán 97302, Mexico;
[email protected] (A.R.L.-R.); [email protected] (K.A.-C.); [email protected] (J.C.C.-H.);
[email protected] (F.B.-P.); [email protected] (T.T.-T.)
2 Facultad de Estudios Superiores Iztacala, UNAM. Tlalnepantla, Edo. de México 54090, Mexico;
3 Facultad de Veterinaria y Zootecnia, Universidad Autónoma de Yucatán. Mérida, Yucatán 97288, Mexico;
4 Unidad de Biotecnología, Centro de Investigación Científica de Yucatán A.C., Parque Científico y Tecnológico del Estado de Yucatán, Carretera Sierra Papacal-ChuburnáPuerto, Yucatán 97302, Mexico;
5 Centro de Investigación y Asistencia en Tecnología y Diseño del Estado de Jalisco, Unidad Sureste, Parque Científico y Tecnológico del Estado de Yucatán, Carretera Sierra Papacal-ChuburnáPuerto, Yucatán 97302, Mexico; [email protected]
* Correspondence: [email protected]; Tel.: +52-999-9428-330; Fax: +52-999-9813-900 Received: 29 October 2018; Accepted: 25 December 2018; Published: 31 December 2018
Abstract: Marine microalgae are a promising feedstock for biofuel production given their high growth rates and biomass production together with cost reductions due to the use of seawater for culture preparation. However, different microalgae species produce different families of compounds.
Some compounds could be used directly as fuels, while others require thermochemical processing to obtain quality biofuels. This work focuses on the characterization of three marine microalgae strains native in Mexico and reported for the first time. Ultrastructure and phylogenetic analysis, suggested that they belong toNannochloropsissp. (NSRE-1 and NSRE-2) andNannochlorissp. (NRRE-1).
The composition of their lipid fractions included hydrocarbons, triacylglycerides (TAGs), free fatty acids (FFAs) and terpenes. Based on theoretical estimations from TAG and FFA composition, the potential biodiesels were found to comply with six of the seven estimated properties (ASTM D6751 and EN 14214). On the other hand, hydrocarbons and terpenes synthesized by the strains have outstanding potential as precursors for the production of other renewable fuels, mainly green diesel and bio-jet fuel, which are “drop-in” fuels with quality properties similar to fossil fuels.
The validity of this theoretical analysis was demonstrated for the oxygenates of strain NSRE-2, which were experimentally hydrodeoxygenated, obtaining a high-quality renewable diesel as the reaction product.
Keywords:biodiesel; bio-jet fuel; triacylglycerides; Fatty Acid Methyl Ester; lipids; hydrodeoxygenation;
drop-in fuel
1. Introduction
Microalgae offer an attractive way of generating renewable and sustainable biofuels [1] capable of helping to meet the global demand for transport fuels. However, a screening procedure to assess
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microalgae potential is a highly recommended prerequisite for biofuel feedstock production. Marine microalgae, which are abundant in seawater, are a particularly promising raw material due to the cost reductions resulting from using seawater for cultivation. The additional salinity level is also one of the methods of reducing contamination issues [2] in open-pond cultivation. For this reason, a number of companies currently operate commercial-scale mass cultivation of marine microalgae [3].
Proper characterization and adaptation to culture conditions of isolated native strains are the first steps in establishing any biotechnological process, including biofuel production. Strategies for improving microalgae growth include the variation of inoculum size, pH and culture medium composition. Inoculum size significantly affects microalgae cell growth characteristics, such as lag phase duration, maximal specific growth rate, biomass accumulation and metabolites production at the end of culturing [4]. An appropriate inoculum size can reduce cell mortality and increase biomass production and nutrient recovery [5]. With a proper culture pH, high cell densities and lipids accumulation can be obtained [6]. Culture medium composition affects the specific growth rate, biomass production and the biochemical composition of the resulting biomass and lipids [7].
Characterization of marine microalgae must include ultrastructure analysis of organelles and their disposition inside the cell, as well as light microscopy to elucidate reproduction mode, determination of the phylogenetic position of the strains under study, and analysis of their carotenoid and chlorophyll content in order to determine the taxonomic identity of each strain [8,9]. All of these analyses are particularly important in taxa such asNannochloropsissp. (Eustigmatophyceae) andNannochloris sp. (Chlorophyceae), both extremely small unicellular marine eukaryotic algae with easily confused characteristics, which can even be impossible to distinguish due to their tiny cell size and simple and smooth cell wall morphology [10]. Among the most important marine microalgae for obtaining biofuels, preciselyNannochloropsisspp. stand out due to their high growth rates and lipid contents in the form of TAGs [11].Nannochloris, meanwhile, is a species that has received less study for biofuel purposes, but has high lipid productivity [12]. To our knowledge, this is the first report on the isolation and characterization ofNannochloropsisandNannochlorisspecies in Mexico.
The lipidic fraction of the microalgal biomass is used for biofuel production. Triacylglycerides are converted to biodiesel, and this procedure has been extensively studied. However, other compounds produced by microalgae such as hydrocarbons and terpenes can also be used for the production of bio-jet fuel and green diesel. For example, phytol stands out as a natural diterpene alcohol that can be used as biofuel in diesel engines [13] or converted to gasoline by catalytic cracking [14]. Another terpene, neophytadiene, is one of the dominant ones in green microalgae [15] and it could also be converted to biofuel. Terpenes are hydrocarbons that are already important as bioactive compounds, with applications as functional compounds due to their broad spectrum of biological activity, such as antitumor and antiviral activity [16], among others. However, from more than 40,000 reported structures, specific terpenes (cyclic and acyclic forms) are recognized as special biofuels that can be used directly or blended with a range of fossil fuels, such as jet fuel, missile propellant, gasoline, or diesel fuels [17]. On the other hand, hydrocarbons produced from microalgae can be directly used in mixtures with the diesel fraction (C12–C22), or they can be subjected to a catalytic improvement stage (hydroisomerization or hydroisomerization plus hydrocracking process), through which paraffins with better fuel properties can be obtained [18]. The selection of the process will depend on the composition of the raw material and the type of fuel that is to be produced. For example, if the hydrocarbon fraction has a high content of paraffins with more than 21 carbon atoms, a hydroisomerization process with simultaneous hydrocracking would be recommended. This process would permit the formation of branched hydrocarbons with shorter carbon chains and products in the range of gasoline (bio-gasoline), jet fuel (bio-jet fuel) and diesel (green diesel).
The vast biodiversity available in Yucatan, Mexico, provides a valuable resource for the development of indigenous microalgal strains as biofuel feedstock. The objective of this research was to collect marine strains from coastal waters and analyze morphology, reproduction mode and organelle disposition by light microscopy and ultrastructure analysis to support phylogenetic studies
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and determine taxonomic identity. Additionally, the fatty acids and organic compounds synthesized by the strains under study were analyzed to determine their potential for biofuel production, either through catalytic processes for obtaining drop-in fuels (bio-jet fuel and renewable diesel) or biodiesel, or even using them without chemical modification in mixtures with fossil fuels. Finally, a catalytic hydrodeoxigenation reaction was carried out to experimentally demonstrate the potential of the extracted lipids as raw material to obtain high-quality renewable fuels.
2. Materials and Methods
2.1. Isolation and Cultivation of Microalgae
Marine microalgae were collected from the water column at three different sites in Progreso, Yucatan, located at 21◦1901.33N, 89◦4050.05W; 21◦1709.38N, 89◦4150.05W and 21◦1709.36 N, 89◦4150.07W. Samples were collected at three different depths (0.0, 0.5 and 1 m) from water bodies during the mornings of May 2014 using a Van Dorn bottle (1120-G45, Wildco, Yulee, FL, USA).
The pH, temperature and salinity values (Extech Stick EC500, Extech instruments, Boston, MA, USA) were registered [19]. The collected samples were allowed to settle for 3 to 4 h at 4◦C. The sedimented biomass was analyzed to proceed with the isolation of the microalgae species.
2.2. Obtaining Axenic Cultures
Microalgae cells were observed by inverted microscope (Axio Observer-5 ZEISS, Baden-Württemberg, Oberkoche, Germany) to select individual cells and inoculate in Guillard’s f/2 (G9903, Sigma-Aldrich, San Luis, MO, USA) and saline BG-11 medium (2% NaCl) (C3061, Sigma). Then microalgae colonies (500μL plated in solid Bold’s basal medium (BBM) were transferred to both media for decontamination:
Centrifugation. Microalgae cultures were centrifuged, pellets were washed in sterile seawater and centrifuged again, and were then inoculated [19]. Serial dilution. Dilutions of microalgae culture (1:10, 1:100 and 1:1000 (v/v)) were employed.Antibiotic treatment.After proliferation of bacteria in Müeller-Hinton media, antibiotic susceptibility testing disks (50295; Bio-Rad, Hercules, CA, USA) were used, giving maximum susceptibility to chloramphenicol, amikacin and streptomycin (100 mg/L) [20].
Acid shock method. Cultures in exponential phase were centrifuged at 2291×gfor 7 min at 10◦C (Digicen 21R, OrtoAlresa, Madrid, Spain). The pellet was resuspended three times in 1 mL HCl (1N) by manual agitation [21] and re-suspended in both media and incubated. The axenic microalgal strains were cultivated (10 mL) in Guillard’s f/2 (G9903, Sigma) and BG-11 (2% NaCl) (C3061, Sigma) under a 12:12 photoperiod at 25±2◦C, 110μmol m−2s−1light intensity and 120 rpm agitation. To improve cell densities, nutrient content (Guillard’s f and f/2 medium) was evaluated at different pH (6, 7 and 8) and inoculum size (10, 20 and 30%). Cell concentration was determined every 48 h using a Neubauer hemocytometer [22].
2.3. Morphological Analysis. Light Microscopy and Transmission Electron Microscopy (TEM)
Morphological features (cell shape and reproductive features) observed through optical microscopy (Eclipse E200, Nikon instruments, Melville, NY, USA) were recorded. For TEM, the strains were preserved in 2% glutaraldehyde for 4 h and then post-fixed in 1% OsO4for 30 min, followed by progressive dehydration in 30 to 100% ethanol at 4◦C. The samples were then transferred to 100%
propylene oxide and infiltrated into epon-propylene oxide (1-1) for 48 h, followed by embedding in epon and polymerization at 60◦C for 48 h [23]. Cross sections were performed with a Leica ultramicrotome (Leica, Wetzlar, Germany) and observed with the 1200-EX II TEM (JEOL, Peabody, MA, USA) located at the Institute of Cellular Physiology, UNAM.
2.4. DNA Extraction and 18S rRNA Gene Amplification
The microalgae cells were harvested at the stationary growth phase by centrifugation (30 mL) and washed three times in bidistilled sterile water. The cell (c.a. 200 mg) was stored at−20◦C until use.
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For genomic DNA extraction, the Youssef protocol was used with minor modifications [24]. 18S rRNA gene amplification was implemented using forward primer A (5-AACCTGGTTGATCCTGCCAG-3) and reverse primer SSU-inR1 (5-CACCAGACTTGCCCTCCA-3) based on the conserved domain region of 18S rDNA [25].
18S rRNA region amplification was carried out with a 10μL reaction mixture containing 1×
PCR Buffer, 2 mM MgCl2, 0.2 mM dNTPs, 1 mM primer, 1 UTaqDNA polymerase (Invitrogen, Waltman, MA, USA) and 10 ng of DNA. PCR conditions included 1 cycle of 1 min at 94◦C for initial denaturation, followed by 40 cycles of 30 s at 94◦C, 30 s at 57.5◦C, 1 min at 72◦C, and finally 7 min at 72◦C. Amplicons were checked by electrophoresis on 1% agarose gel in 1×TAE buffer.
The amplification products of approximately 500 pb were purified using NucleoSpin®Gel and PCR Clean-up commercial kit (Macherey-Nagel, Düren, Germany) to remove primer dimers according to the manufacturer’s instructions. Direct sequencing of the amplification products was carried out by a two-directional (reverse and forward) procedure (Clemson Sequencing Service, Clemson, South Caroline, USA).
The 18S rRNA region sequences from DNA of NSRE-1, NRRE-1 and NSRE-2 of 540 pb (approx.) were used as “queries” in nucleotide Blast tools (NCBI website) to recover the ten best matches for each input sequence, based on percentage of sequence coverage, E-value and percentage of identity of 100%. After manual adjustment with MEGA version 2.1, sequences with their homologies in GenBank were deleted to a final sequence of 536 pb and aligned using CLUSTALX (EMBL, Heidelberg, Germany). Molecular phylogenetic analyses by the maximum likelihood method based on the Kimura 2-parameter model were conducted using MEGA version 2.1 [26]. One thousand replica samplings were analyzed for percent bootstrap values in a neighbor-joining tree. The percentage of trees in which the associated taxa clustered together is shown next to the branches. Evolutionary analyses were conducted in MEGA 7 [26]. Reference sequences retrieved from GenBank of 18S rRNA region sequences were: NSRE-1, MH921192; NRRE-1, MH921193; and NSRE-2, MH921194.
2.5. Biomass Production
Algal biomass was produced in a culture system consisting of three serially connected 250 mL Erlenmeyer flasks with 200 mL of culture medium and a 250 mL Erlenmeyer flask with sterile distilled water (with the purpose of acting as an air humidifier). The culture system was maintained under the following conditions: an aeration rate of 3.6 vvm, temperature of 24±2◦C, and light intensity of 81μmol m−2s−1with a 16:8 (light:dark) photoperiod [27]. The biomass was recovered at the beginning of the stationary phase (15 days of cultivation) by centrifugation at 5057×gfor 7 min at 25◦C [28]. The pellet was washed with bidistilled sterile water to eliminate minerals and salts from the culture media. The total biomass was maintained at−80◦C for 24 h and freeze dried at 0.1 bar and
−53◦C for 72 h [29].
2.6. Total Lipid Extraction
One g of dried biomass was subjected to solvent extraction with 15 mL of a 1:2 (v/v) chloroform-methanol mixture [30–33] for 3 h at 40◦C and 150 rpm (3×). The solvents were then eliminated by vacuum evaporation.
2.7. Thin Layer Chromatography
It was carried out on TLC Silica gel 60 F254plates (Merck KGaA, New York, NY, USA). A 9:1:0.1 (v/v) mixture of hexane, ethyl acetate and acetic acid was used as the mobile phase. The plates were developed using a 10% (w/v) solution of phosphomolybdic acid in sulfuric acid [34].
2.8. Column Chromatography
The lipid extract was fractionated by column chromatography using silica gel (Kieselgel, 70–230 mesh) as the stationary phase. A solvent gradient elution was used to obtain five fractions from
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the lipid extract. The solvent system was hexane, then 9:1, 8:2 and 7:3 mixtures of hexane-ethyl acetate, and finally methanol [29,34,35].
2.9. Transesterification
The TAG fraction was transesterified with sodium metoxide (6%v/w) in a 25 mL vial with moderate agitation at 60◦C for 90 min. The reaction mixture was passed through silica gel to eliminate excess catalyst and glycerol. The solvent was then evaporated in a fume hood.
2.10. Lipid Profile Analysis (GC-MS)
The fractions were analyzed by gas chromatography–mass spectrometry (7890B and 5977A series GC/MSD, Agilent Technologies, Santa Clara, CA, USA), using a DB5-HT column (30 m×250μm ID×0.25μm film width) under the following conditions: 3 mL/min of helium flux as carrier gas.
The detector and injector temperatures were set at 300◦C. The GC temperature program started at 70◦C/5 min, and was then raised to 200◦C using a rate of 10◦C/min with a 5 min hold at 200◦C and 20◦C/min until 290◦C, which was held for 22.5 min. The injection volume was 1μL. The NIST 05 (National Institute of Standards and Technology, Gaithersburg, MD, USA) library was used for identification of the main peaks [36]. Six main groups of compounds were identified: hydrocarbons, FAMEs, organic acids, terpenes, sterols and amines.
2.11. Catalytic Hydrodeoxygenation of Oxygenates from Strain NSRE-2
The fraction of oxygenated compounds from strain NSRE-2 was subjected to a catalytic hydrodeoxygenation treatment (HDO) in a stainless steel (SS-316) fixed bed reactor (Microactivity Reference, Madrid, Sapin), with a length of 30.5 cm and a 0.9 cm internal diameter. The reaction mixture (31 mg lipid extract dissolved in 120 mL nonane) was fed using an HPLC pump (model 307 V 3.00, Gilson, Villiers-le-Bel, France) at a constant flow rate of 0.061 mL min-1 to attain a WHSV of 0.1 h−1. Pt/HZSM-22-γ-Al2O3synthesized in-house was used as catalyst (0.046 g) and activated in-situ under H2flow (50 mL min−1) at 400◦C for 2 h with a heating rate of 5◦C min−1and 20 bar with a pressure rate of 5 bar min−1. The reaction conditions were 300◦C at a heating rate of 5◦C min−1 and a H2pressure of 40 bar. H2was fed at a gas flow rate of 55 mL min−1and the total reaction time was 4 h. Samples of the reaction product were collected every hour and analyzed by GC-MS.
2.12. Statistical Analysis
Results of biomass and lipid production are presented as means±standard deviations (SD) from three replicates. The statistical analysis was carried out using OriginPro (v 9.0, OriginLab, Northampton, MA, USA) to determine the degree of significance using one-way analysis of variance (ANOVA) at a probability level of P < 0.05.
3. Results and Discussion
3.1. Obtaining Axenic Cultures
The three isolated microalgae were named NSRE-1, NRRE-1 and NSRE-2, and placed under the conditions mentioned. Application of HCl proved to be an effective decontamination technique.
This pH-shock treatment has been applied before for effective control ofV. chlorellavorus, a predatory bacterium that can destroy aChlorellaculture in just a few days [37]. This effective treatment with high selectivity and low cost can be considered throughout the scale-up and production process [37].
The pH-shock treatment applied in this study is simpler than the one executed by Ganuza et al. [37]
and patented by Ganuza and Tonkovich. Application of antibiotics has been used by other authors to obtain axenic cultures from freshwater bodies [38]. However, bacterial contamination persisted in all cultures when amikacin, active against aerobic gram-negative bacilli [19], was applied in this work.
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The culturing of NSRE-1, NRRE-1 and NSRE-2 in Guillard’s f/2 resulted in improved growth compared to BG-11 medium, as indicated by the cell densities (cell mL−1) of NSRE-1, NRRE-1 and NSRE-2. In Guillard’s f/2 they were 9.03×106, 5.46×106and 6.28×106, while in BG-11 they were 1.48×106, 2.56×105and 4.50×105respectively. This result could be explained by the fact that Guillard’s f/2 medium has a ratio of 16N:1P, which promotes higher cell density in shorter cultivation times [39]. Additionally, vitamins in Guillard’s f/2 medium (cyanocobalamin, thiamine and biotin), absent in BG-11, contribute to the initial growth of microalgae species [40].
Improving microalgae culture parameters to increase biomass and growth is considered the most important factor in sustainable product development [41]. This includes the improvement of the medium composition and physical parameters [42]. In this study, modifications to pH were carried out to increase cell density (Figure1). Inoculum size of microalgae is another important factor that significantly affects microalgae growth, specifically, the lag phase, maximum specific growth rate, biomass accumulation and the metabolite production at the end of culturing (Richmond 2004). In this study, an inoculum size of 10%v/v, yielded a higher cell density than 20% or 30%, probably because a greater availability of nutrients occurs at lower cell densities [43]. In this regard, inoculum size had an effect on final cell density. Selection of the appropriate inoculum size can help reduce the cell mortality ratio in microalgae and increase the biomass production and nutrient recovery [5].
The nutrient concentration of Guillard’s medium employed in this study were half strength and full strength, denoted as f/2 and f respectively. The cell density double increased in all strains under study by using f Guillard´s f media (Figure1). Notably in NRRE-1, cell density was ten times greater than NSRE-1 and NSRE-2. These results are consistent with those obtained by Malakootian et al.
(2015), inNannochloropsis oculatawith Walne medium, the composition of which is similar to Guillard´s f medium, resulting in a highest cell density than with Guillard’s f/2, Sato and TMRL media.
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Figure 1.Mean cell density±SE of NSRE-1, NRRE-1 and NSRE-2 strains at three different pH levels and inoculum sizes and two nutrient concentrations.
Based on the results obtained, stabilized microalgae strain culture conditions were as follows:
pH 8, inoculum size 10%v/vand Guillard’s f medium.