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Use of the Dinoflagellate Karlodinium veneficum as a
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Sustainable Source of Biodiesel Production
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C. Fuentes-Grünewald a,b* , E. Garcés a, S. Rossi b , J. Camp a
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a
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b Universidad
Institut de Ciències del Mar, ICM-CSIC, Passeig Marítim de la Barceloneta, 37-49., E-08003, Barcelona, Spain.
Autónoma de Barcelona, Instituto de Ciencias y Tecnología Ambiental, E-08193, Bellaterra, Spain.
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Abstract
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Microalgae are microscopic heterotrophic-autotrophic photosynthesizing organisms
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with enormous potential as a source of biofuel. Dinoflagellates, a class of microalgae,
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contain large amounts of high-quality lipids, the principal component of fatty acid
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methyl esters (FAMEs). The properties of the dinoflagellate species Karlodinium
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veneficum include a growth rate of 0.14 day-1, a wet biomass of 16.4 g/L, a growth
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period ≈ 30 days, and a ~97% increase in fatty-acid content during the transition from
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exponential phase to stationary phase. These parameters make K. veneficum a suitable
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choice as a bioresource for biodiesel production. Similarly, two other species were also
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determined to be appropriate for biodiesel production: the Dinophyceae Alexandrium
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andersoni and the raphidophyte Heterosigma akashiwo.
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Keywords: Biodiesel; microalgae; dinoflagellates; lipids; Karlodinium veneficum
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1. Introduction
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Biodiesel is a biofuel that, through transesterification, can be produced from different
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feedstocks, including grease, vegetable oils, waste oils, animal fats, and microalgae. In
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this reaction, triglycerides are converted into fatty acid methyl esters (FAMEs) in the
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presence of an alcohol, such as methanol or ethanol, and either an alkaline or acidic
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catalyst. The reaction produces two immiscible layers, biodiesel and, as a by product,
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glycerol (Palligarnai and Briggs, 2008).
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The unstable price of fossil fuel, worldwide interest in reducing the amount of CO2
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emitted into the atmosphere, and the attempts of petroleum-dependent countries to
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enlarge their energy matrix have led to increasing interest in biofuel production. Until
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recently, the synthesis of biodiesel derived mostly from terrestrial plants. This strategy
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has become controversial because of the lack of sustainability of plant-based biofuel,
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specifically, the resulting deforestation of extensive land otherwise devoted to the
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cultivation of soybean, palm, sugarcane, rapeseed, and other food plants (Lian, 2007),
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the consumption of scarce water resources, the degradation of arable land, and the
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reduced amount of CO2 fixation. Moreover, the transformation of primary food
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resources into biofuels has led to a clash of interests, as plant-derived biodiesel has
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deprived poor countries of food and increased its cost (Puppán, 2002). This has
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stimulated the search for other sources of biodiesel production, ones that are both
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sustainable and economical (Chisti, 2007).
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Microalgae are microscopic heterotrophic-autotrophic photosynthesizing organisms that
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inhabit many different types of environments, including freshwater, brackish water, and
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seawater. More than 40,000 different species of microalgae are known, most of which
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have a high content of lipids, accounting for between 20 and 50% of their total biomass
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(Chisti, 2007). Accordingly, microalgae have the potential to synthesize 30 times more
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oil per hectare than terrestrial plants (Sheehan et al., 2006). They are widely used in
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industry in the synthesis of pigments and additives, as a source of protein, and in biofuel
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production.
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Marine microalgae shows several advantages compared to other sources of biodiesel
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production: Their high growth rate has the potential to satisfy the enormous demand for
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biofuels but they can be cultured on non-agricultural land or even in coastal areas and
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without the need of freshwater. In addition, the tolerance of microalgae to a high CO2
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content in gas streams allows high-efficiency CO2 mitigation (Chang et al., 2003; Hsueh
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et al., 2007). Biodiesel from microalgae does not contain sulfurs, is highly
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biodegradable, and is associated with minimal nitrous oxide release. Microalgal farming
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is also potentially more cost-effective than conventional farming (Chisti, 2008; Yanqun
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et al., 2008).
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In the 1970s, an important effort to identify the optimal type of algae for biodiesel
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production was made by the US government, in response to the petroleum crisis
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(Sheehan et al., 1998). As a result, more than 3,000 different microalgae species,
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including those belonging to the Bacillariophyceae, Chlorophyceae, Cyanophyceae,
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Prymnesiophyceae, Eustigmatophyceae, and Prasinophyceae, with natural habitats in
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different parts of the country, were examined. A few of these algae were shown to be
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cultivable on a large scale such as in a photobioreactor or in ponds.
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Nonetheless, the production of biodiesel from microalgae has thus far been restricted to
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a few species, i.e., those for which the culture conditions in high biomass systems are
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known: the cyanobacteria Spirulina platensis (protein production), the Chlorophyceae
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Chlorella protothecoids (heterotrophic cultivation in photobioreactors for biomass),
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Tetraselmis suecica (food source in aquaculture hatcheries), and Haematoccocus
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pluvialis (pigment production). Despite their relatively widespread use in these and
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other applications, these species are controversial sources of biodiesel because their
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cultivation requires the input of large amounts of freshwater (Spirulina, Chlorella,
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Haematoccocus) or because their oil content is too low to be of economic interest
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(Tetraselmis,).
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Microalgae with a high content of fatty acids, neutral lipids, and polar lipids as well as a
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high growth rate in the natural environment have yet to be exploited for biodiesel, and
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the isolation and characterization of microalgae with the potential for more efficient
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lipid/oil production remain subjects of research (Qiang et al., 2008).
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A high content of fatty acids, as a neutral lipids or triacylglycerols (TAG), is found
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naturally in a group of microalgae, the dinoflagellates. Additionally, these organisms
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occasionally form explosive and extensive proliferations (blooms) in coastal waters all
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over the world. These episodic blooms extend for hundreds of kilometers and their cell
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concentrations are in the millions per liter (Clement, et al., 2002; Basterretxea, et al.,
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2005). These properties make dinoflagellates of potential interest as a source of biofuel.
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Therefore, in this study, three genus of dinoflagellate and one raphidophyte species
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(Alexandrium, Karlodinium, Scripsiella, and Heterosigma) were investigated as
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alternatives for biodiesel production and as sources of biomass in biofuel production.
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Specifically, the properties of these microalgae were compared with those of microalgae
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traditionally used in biodiesel production, in terms of growth rate, cell yield, and
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quantity and quality of their oil content.
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2. Materials and Methods
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2.1 Strain cultures
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All species were cultivated in multiple and in batch cultures at the Marine Science
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Institute (ICM, CSIC). The characteristics of the species are presented in Table 1. The
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strains were grown in L1 medium under the same conditions (Guillard and Hargraves
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1993). The ten strains, belonging to six genera, were inoculated at an average
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concentration of 4200 cells/mL seawater (salinity = 36, neutral pH) into 2-L Nalgene
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flasks and incubated at 21°C ± 1°C in prefiltered air (Iwaki filter, 0.2-µm pore size).
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Flasks were maintained using a 12:12 h light:dark cycle, with illumination provided by
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fluorescence tubes (Gyrolux, Sylvania, Germany) emitting a photon irradiance of 110
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μmol photons m-2 s-1 (measured with a Licor sensor).
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Table 1
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2.2 Growth rates
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Subsamples (10 mL) of each culture were fixed in Lugol’s iodine. During the lag phase,
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the cultures were sampled every day and during stationary phase every 4 days. Samples
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were counted in a Sedgewick-rafter chamber under an inverted optical microscope
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(Leica-Leitz DM-II, Leica Microsystems GMbH, Wetzlar, Germany) at 200–400×
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magnification. Cell abundance data were used to calculate the exponential growth rate
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of the cultures. Species-specific net growth rates were estimated from μ = ln(N0/Nt)/t,
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where N0 and Nt are the initial and final cell densities, and t the time interval in days
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(Guillard, 1973).
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2.3 Wet weight (biomass)
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Wet weight (WW) was determined by filtering duplicate subsamples (10 mL) through
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pre-weighed glass-fiber filters (Whatman GF/F 25 mm, nominal pore size 0.7 µm) and
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then weighing the filters on a Sartorious balance (precision of 0.001).
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2.4 Lipid extraction and fatty-acid analyses
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Primary lipid analyses were carried out for all strains at the Institute of Science and
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Environmental Technology (ICTA, Autonomous University of Barcelona, Spain). The
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lipids of one control strain (Tetraselmis suecica; number 10, see Table 1) and of five
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different strains (number 1, 2, 3, 5, 7, see Table 1) were analyzed at three different time
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in the growth curve: lag phase (day 6), exponential phase (day 21), and stationary phase
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(day 35).
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Triplicates of a 50-mL subsample were filtered on previously combusted (450ºC 4 h)
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GF/F Whatman glass-fiber filters, immediately frozen in liquid N2, freeze-dried for 12 h
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and then stored at -20ºC until analysis. The filters were placed in a tube with 3:1
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dichloromethane-methanol (DCM:MeOH), spiked with an internal standard (2-
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octyldodecanoic acid and the 5β-cholanic acid), and the lipids extracted using a
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microwave-assisted technique (5 min. at 70ºC). After centrifugation, the extract was
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taken to near dryness in a centrifugal vacuum concentrator maintained at constant
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temperature and then fractionated by solid-phase extraction according to a previously
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published method (Ruiz, et al., 2004) The sample was subsequently re-dissolved in 0.5
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mL of chloroform and eluted through a 500-mg aminopropyl mini-column (Waters Sep-
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Pak® Cartridges) previously activated with 4 mL of n-hexane. The first fraction was
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eluted with 3 mL chloroform:2-propanol (2:1) and the fatty acids recovered with 8.5 mL
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of diethyl ether:acetic acid (98:2). Despite reported concerns on the background free
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fatty acids (FFAs) levels of aminopropyl columns (Russel, and Werne, 2007), the
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concentrations of target FFAs in the SPE cartridges used were below the detection limit.
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The FFAs fraction was methylated using a 20% solution of MeOH/BF3 heated at 90ºC
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for 1 hr. The reaction was quenched with 4 mL of NaCl-saturated water. FAMEs were
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recovered by extracting the samples twice with 3 mL of n-hexane. The combined
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extracts were taken to near dryness, re-dissolved with 1.5 mL of chloroform, eluted
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through a glass column filled with Na2SO4 to remove residual water, and, after removal
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of the chloroform, with nitrogen evaporation. The extracted sample was stored at -20ºC
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until gas chromatography analysis.
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Gas chromatography analysis of extracts re-dissolved in 30 µL of iso-octane was carried
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out in a Thermo Finnigan Trace GC Ultra instrument equipped with a flame ionization
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detector and a splitless injector, and fitted with a DB-5 Agilent column (30-m length,
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0.25-mm internal diameter, 0.25-µm phase thickness). Helium was used as the carrier
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gas, delivered at a rate of 33 cm s-1. The oven temperature was programmed to increase
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from 50 to 320ºC at 10ºC min-1. Injector and detector temperatures were 300ºC and
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320ºC, respectively. FAMEs were identified by comparison of their retention times with
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those of standard fatty acids (37 FAME compounds, Supelco® Mix C4-C24) and
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quantified by integrating the areas under the curves in the gas chromatograph traces
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(Chromquest 4.1 software), using calibrations derived from internal standards.
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2.5 Lipids fluorescence in microalgae
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The intracellular neutral lipid distribution in microalgal cells was examined by staining
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a 3.mL suspension of the algae with 10 µL (7.8 × 10-4 M) of Nile Red fluorescent dye
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(Sigma-Aldrich) dissolved in acetone (final concentration 0.26 µM). The samples were
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examined by epifluorescent microscopy (Leica-Leitz DM-II, Leica Microsystems
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GMbH, Wetzlar, Germany) with an excitation wavelength of 486 nm and emission
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measured at 570 nm, following the method of Cooney et al. (2007). Photographs were
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taken with a Sigmapro software image analyzer, which was also used to calculate the
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percentage of positive stained cells.
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3. Results
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3.1 Strains growth rate
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The net growth rate differed among the examined species (Fig. 1) and was highest for
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Tetraselmis suecica (Prasinophyceae), which grew at a rate of 0.23 day-1 (1 division
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every 2.8 days) during the exponential phase of growth. Accordingly, at 24 days of
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culture, the cell abundance was nearly 85 × 106 cells/L, after which the culture entered
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stationary phase and decayed. Among the three dinoflagellates, the growth rate of
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Karlodinium veneficum was the highest, 0.14 day-1 in the exponential phase,
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corresponding to an abundance of 44 × 106 cells/L at day 30 of culture. For
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Heterosigma akashiwo, maximum abundance was approximately 26 × 106 cells/L at day
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35 of culture, reflecting a growth rate of 0.10 day-1 (1 division every 10 days). Among
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the six species examined in the study, this raphidophyte was unique in that cell
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abundance was maintained for more than 6 months (data not shown). Moreover, the
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cells remained healthy without the addition of fresh medium. This was in contrast to the
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other cultures, which gradually decayed such that total cell lysis has occurred ~2 months
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after inoculation.
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The growth rate of dinoflagellates belonging to the genus Alexandrium differed
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depending on the species. The highest growth rate was that of A. andersoni, 0.10 day-1
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similar to that of H akashiwo, but the cell abundance of the former (maximum of 9 ×
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106 cells/L) was lower than that of the raphidophyte. The growth rates of A. minutum
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and A. catenella were two orders of magnitude slower (0.04 day-1 and 0.03 day-1,
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respectively) than those of faster-growing microalgae. In terms of abundance, A.
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minutum reached a maximum of 2.6 × 106 cells/L and A catenella a maximum of 9.4 ×
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105 cells/L at culture day 36 and 35, respectively.
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Figure 1
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3.2 Wet Biomass
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In the lag phase, the maximum biomass was achieved by H. akashiwo (WW of 14.3 g/l),
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and the minimum by A. andersoni (9.4 g/l). In the exponential phase, the maximum wet
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weight was that of A. catenella (20.2 g/l), and the lowest that of K. veneficum (average
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of 16.1 g/l). The increase in wet weight during exponential phase was more evident in
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the genus Alexandrium than in the other microalgae (Fig. 2). The different microalgal
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strains evidenced similar results, with maximum biomass reached in the late-
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exponential phase. At stationary phase, the wet weight of all the cultures diminished.
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The only exception was Tetraselmis suecica, which retained the weight it had obtained
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in exponential phase (18.5 g/l).
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Figure 2
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3.3 Total lipid content
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The lipid concentration as a percentage of total fatty acid was determined at the
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different phases of culture in the strains grown under equivalent culture conditions. The
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lipid content of the control strain, Tetraselmis suecica, was maintained at almost the
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same level throughout the experiment, with only a slight decrease (7.8%) in the
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stationary phase (data not shown). The behaviour of the genus Alexandrium differed
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depending on the species. The lipid content of A. catenella diminished by 7.0% from lag
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phase to exponential phase, and then increased by nearly 48% from exponential phase to
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stationary phase. In A. minutum, the lipid content increased by approximately 97% from
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lag phase to exponential phase whereas that of A. andersoni diminished by about 27%
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from exponential phase to stationary phase. The best performance in terms of lipid
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accumulation was that of Karlodinium veneficum. The lipid content of this
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dinoflagellate increased throughout the different growth phases: by 40% from lag phase
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to exponential phase followed a large and intense accumulation, ~97%, from
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exponential phase to stationary phase. The raphidophyte H. akashiwo maintained its
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total lipid content from lag phase to exponential phase, but there was an extreme
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reduction (~43%) from exponential phase to stationary phase.
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The highest total lipid content (Fig. 3) was achieved at the stationary phase of culture,
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by the dinoflagellate K. veneficum, whereas during this growth phase, among the
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microalgal strains analyzed, the lipid content of Tetraselmis suecica was the lowest.
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Figure 3
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3.4 Fatty acid composition
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The most abundant fatty acids expressed by the different classes of microalgae during
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stationary phase were those of the 18:0, 16:0, 20:3, and 17:1 type (Table 2). In general,
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the highest levels of saturated lipids were found in A. catenella (42.3%), A. minutum
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(40.6%), and K. veneficum (39.7%). In diatoms, saturated fatty acid content ranged from
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22.3% (the minimum of the six strains tested) in P. delicatissima to 32.3% in C. affinis.
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In the dinoflagellate S. trochoidea saturated fatty acids accounted for 29.8% of the lipid
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content, compared to 39.7% in the control algae T. suecica. Monounsaturated fatty acids
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varied from 5.8 to 9.8% of the total fatty acids. Polyunsaturated fatty acids (PUFA)
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consisted mainly of 18:5(n3) (0.8–5.0%), 20:3(n3) (2.8–6.4%), and, in lesser amounts,
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20:4(n3) (0.4–2.5%).
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Table 2
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3.5 Change in fatty acid composition at different growth phases
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The fatty acid composition in members of the Dinophyceae changed depending on the
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culture phase. In most cases, the lipid concentration, especially of PUFAs such as
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C18:5(n3) and C20:3(n3), increased during the transition from exponential to stationary
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phase. In A. minutum and A. catenella, these polyunsaturated acids increased by
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approximately 5% whereas in A. andersoni their amounts did not change. Among the
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Dinophyceae, Karlodinium veneficum had the greatest increase in lipid content, 45%,
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from exponential to stationary phase. This increase was seen not only in the main
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compounds C16:0 and C18:0, but also in all fatty acids measured. The lipid content of
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the raphidophyte H. akashiwo also increased, by about 20%, between lag phase and
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stationary phase (data not shown). In the control algae T. suecica, a smooth increase of
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only ~3%, in the main fatty acids C16:0 and C18:0, was observed (data not shown).
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Figure 4
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3.6 Fluorescence of neutral lipids
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The liposoluble fluorescence probe Nile Red was used to visualize neutral lipids in the
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cells. This method has several advantages over in situ screening. The dye is relatively
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photostable, intensely fluorescent when dissolved in organic solvent and in a
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hydrophobic environment, and it is sensitive to non-polar lipids in living cells. In this
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study, microphotographs (Fig. 5) were obtained from stationary-phase cultures.
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The highest lipid content, as determined by Nile Red staining, was observed in K.
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veneficum, in which 81% of the cells were lipid-positive. In this species, small drops of
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neutral lipids were seen dispersed throughout the cytoplasm. While some A. minutum
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cells in the sample stained highly positively for lipid, others did not such that, overall,
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the percentage of stained cells was low (21.3%). T. suecica was also comparatively poor
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in neutral lipid content during stationary phase. By contrast, many of the cells of A.
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andersoni showed massive concentrations of neutral lipids, usually located in the
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hypotheca of the cell.
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Figure 5
B
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D
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4. Discussion
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Characteristic growth curves for marine microalgae obtained for the 6 microalgal strains
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showed that for most of them the lag phase occurred from day 0 to day 6; the exception
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was T. suecica, consistent with previously reported data for this species (Fábregas, et
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al., 2001; Fábregas, eta al 1991). The biomass of T. suecica doubles every 24 hours
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during the exponential growth phase, thus reaching high densities. For this reason, it has
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become one the microalgal strain most frequently used in industrial aquaculture.
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Among the Dinophyceae, Karlodinium veneficum showed the best performance in terms
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of growth, although the rate measured in this study was much lower than that of wild
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populations (Stolte, and Garcés, 2006). Nonetheless, it was high enough to yield a large
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biomass in culture within a reasonable period of time. Further studies will be needed to
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determine whether the growth rate in culture can be improved, e.g., by isolating new
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strains and/or inoculating the cells in exponential phase, before the maximum growth
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rate is established.
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In the genus Alexandrium, different strategies of growth and abundance were observed:
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1) The best growth results were obtained with A. andersoni, with a rate similar to that of
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wild populations in the Mediterranean Sea. For this species, maximum cell abundance
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occurred at day 35 and the growth curve was longer than that of either the control algae
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or the best Dinophyceae. 2) A. minutum and A. catenella had low growth rates.
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Moreover, the cell abundance of A. catenella was the lowest of the strains studied,
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specifically, almost two orders of magnitude less than the best microalgae in this study
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(T. suecica). The growth rate of the raphidophyte Heterosigma akashiwo was similar to
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that of A. andersoni, and the maximum density was measured at day 36. Furthermore,
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the species was able to remain in stationary phase for more than 6 months. This feature
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could be taken advantage of to maintain this organism as a constant inoculum in a high-
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biomass culture strategy. The different growth rates and cell abundances suggest that
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the biovolume of the cells greatly influences the carrying capacity of the population
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when the microalgae are cultured in flasks or tanks such as those used in this study.
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The biomasses of the strains were similar during the different culture phase. For
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example, the biomass of all strains was lower during lag phase than during exponential
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phase, with the maximum biomass being that of H. akashiwo and the minimum that of
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A. andersoni. In the transition from lag phase to exponential phase, the biomass of all
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strains increased, most importantly in A. catenella and A. andersoni, as both strains
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almost doubled their biomass weight within 10 days. By contrast, the biomass of H.
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akashiwo in the two growth phases differed by only a few grams. In stationary phase,
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culture biomass decreased in all cases; the only exception was Tetraselmis, which
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maintained its biomass between exponential phase and lag phase.
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For commercial or industrial applications, it is important to determine the optimal phase
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to harvest the microalgae. Our results suggest that, in terms of growth phase and
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biomass in this type of culture (batch cultures), harvest during late exponential phase
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results in the highest yields. For example, the average wet weight of K. veneficum, A.
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andersoni, and H. akashiwo during late exponential phase was ~15g/L. This value is
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low compared to the 50 g/L reported for Spirulina platensis, a typical cyanobacteria
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used in protein production and cultured in open ponds, but is similar to the weights
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obtained following heterotrophic cultivation of Chlorella protothecoides, which is used
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for biodiesel production. The wet weight of this green algae when cultured in a
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bioreactor was reported to be about 15.5 g/L in 5 L, 12.8 g/L in 750 L, and 14.2 g/L in
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11,000 L vessels (Li et al., 2007).
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The total lipid content in our strains, especially those belonging to the dinophyceae and
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the raphidophytes, increased from the lag and exponential phases to stationary phase.
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These results are consistent with those of previous studies on other dinoflagellates
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species. Mansour et al. (2003) found that in Gymnodinium sp. the proportion of
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triacylglycerols increased almost four-fold during stationary phase compared to the
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level measured during the exponential phase. Triacyglycerols function as storage lipids
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and thus in most microalgae are usually at their lowest levels during exponential growth
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(Volkman et al., 1989) but increase during stationary phase, as nitrogen or phosphorus
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is depleted (Volkman et al., 1993; Olsen et al., 1994; Brown et al., 1996). This
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observation can be practically applied in strategies aimed at exploiting the high biomass
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reached by cultures of dinoflagellates such as Karlodinium veneficum. Accordingly, it is
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important to evaluate growth phase, biomass, and storage lipids with respect to
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achieving the highest production under different limiting conditions. Based on our
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results on biomass and lipid content, late stationary phase is the optimal time to harvest
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microalgae in order to obtain the highest concentrations of oil.
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The characterization of marine microalgae lipid composition has been suggested as a
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chemotaxonomic tool to distinguish between orders and classes of these organisms
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(Hallegraef et al., 1999; Mooney et al., 2007). However, the lipid composition within
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the same species can vary in response to growth conditions and other, related factors. In
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the microalgae studied in the present work, the characteristic fatty acid composition is
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C16:0 and C18:0 (Mathews and Van Holde, 2000), and these lipids were present in high
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concentrations in our strains. Additionally, the PUFAs octadecapentaenoic acid (OPA
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18:5n3), and 20:5n3 were presents in all dinoflagellate species in varying amounts. A
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requirement of the raw material used for biodiesel production is that it contain high
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amounts of saturated fatty acids. Our results show that the oil extracted from
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dinoflagellates is highly similar to palm oil (Fig 4). With respect to biodiesel
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production, the fatty acid profile of dinoflagellate suggest that the obtained product
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offers several advantages in terms of quality because it results in a high-cetane fuel and
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thus a high quality of ignition
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Staining cells with Nile Red confirmed the presence of oils drops distributed throughout
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the cytoplasm. Within the same culture of A. minutum, some cells had large amounts of
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neutral lipids whereas others had none. There were indeed differences in the per-cell oil
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content in this species. In Karlodinium, small oil drops were heterogeneously
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distributed in the cell body while in almost all cells of A. andersoni they were
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concentrated in the hypotheca. Oil drops were difficult to observe in T. suecica, most
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likely due to the low lipid content of the cells. Taken together, our staining results
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demonstrate that Nile Red can be used prior to the rapid quantification of neutral lipids
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by spectrophotometry, as described by other groups (Cooney et al. 2007; Qingyu et al.
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2008).
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To validate our findings, that dinoflagellates offer a sustainable approach to biodiesel
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production, pilot studies using large-scale cultures are needed. In these studies, a natural
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source of light should be used and culture strategies, such as photobioreactor and open-
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pond cultures, should be developed with the aim of enhancing the quantity and the
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quality of the microalgal lipids.
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5. Conclusion
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Dinoflagellates are widely distributed and readily isolated in many different countries.
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As shown here, they comprise several strategic species that can be used as a source of
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raw material for biofuels. An analysis of the biotic characteristics (growth rate, biomass,
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cell yield, lipid content) of several species of microalgae supports their use as
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feedstocks for biodiesel production. Two species of Dinophyceae, Karlodinium
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veneficum and Alexandrium andersoni, and one species of raphidophyte, Heterosigma
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akashiwo, were found to be of particular interest as a bioresource for biodiesel
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production, based on: 1) their high lipid content; 2) their moderate net growth rate; 3)
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their high average wet biomass; and 4) their short period of growth (28–35 days)
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compared with terrestrial plants;
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Future work will need to focus on improving the biotic features of microalgal cultures
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relevant to biodiesel production, mainly by changing environmental parameters such as
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salinity, nutrient, temperature, and light conditions.
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Acknowledgements
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We thank the members of the L´Esfera Ambiental laboratory, Universitat Autònoma de
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Barcelona, for their help in gas chromatography analyses. This study was funded by the
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Departament de Medi Ambient, CSIC, Generalitat de Catalunya, through the contract
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“Plà de vigilància nociu i toxic a la costa Catalana”. We also thank the Comisión
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Nacional de Investigación Ciencia y Tecnología (CONICYT) Chile, for its support of
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the scholarship “Beca de Gestión Propia,” which finances the PhD studies of CFG.
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References
412
413
Palligarnai, T. Vasudevan., Briggs, M. Biodiesel production-current state of the art and
414
challenges. J. Ind. Microbiol. Biotechnol. 35:421–430. 2008
415
416
Chisti, Y. Biodiesel from microalgae. Biotechnol. Adv. 25, 294-306. 2007
417
418
Lian, P. H., Potential habitat and biodiversity losses from intensified Biodiesel
419
feedstock production. Ecology & Conservation Biology. pp. 1373-1375. 2007
420
421
Puppán, D. Environmental evaluation of biofuels. Periodica polytechnica Ser. Soc.
422
Man. Sci. Vol 10, No. 1, pp. 95-116. 2002
423
424
Sheehan, J. et al. Are a biofuel sustainable?. National Renewable Energy Laboratory.
425
2006
426
427
Chang, E. H. Yang, S. S. Some characteristics of microalgae isolated in Taiwan for
428
biofixation of carbon dioxide. Bot. Bull. Acad. Sin. 44 (1), 43-52 2003
429
430
Hsueh, H. T. Chu, H. Yu, S. T. A batch study on the bio-fixation of carbon dioxide in
431
the absorbed solution from a chemical wet scrubber by a hot spring and marine algae.
432
Chemosphere. 66 (5), 878 – 886. 2007
433
434
Chisti, Y. Biodiesel from microalgae beats bioethanol. Trends in Biotechnology. 26, n°
435
3: 126-131. 2008
436
437
Yanqun, L. Horsman, M. Nan W. Christopher Q.L. Dubois-Calero, N. Biofuels from
438
microalgae. American Chemical Society. 24: 815-820. 2008
439
440
Sheehan, J. Dunahay, T. Benemann, J. Roesler, P. A look back at the U.S. Department
441
of Energy´s Aquatic Species Program: Biodiesel from Algae. National Renewable
442
Energy Laboratory. 1998
443
444
Qiang, H. Sommerfeld, M. Jarvis, E, Ghirardi, M. Posewitz, M. Seibert, M. Darzins, A.
445
Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and
446
advances. The Plant Journal. 54, pp. 621-639, 2008
447
448
Clement, A., Aguilera, A., Fuentes-Grünewald, C. Análisis de marea roja en
449
Archipiélago de Chiloe , contingencia verano 2002. In: XXII Congreso de Ciencias del
450
Mar, Valdivia, Chile, 2002.
451
452
Basterretxea G, Garcés E, A. Jordi A, M. Masó M, Tintoré J. Breeze conditions as a
453
favoring mechanism of Alexandrium taylori blooms at a Mediterranean beach.
454
Estuarine, Coastal and Shelf Science, 32 (1-2):1-12. 2005
455
456
Guillard, R. R. L. & Hargraves, P. E. Stichocrysis immobilis is a diatom, not a
457
chrysophyte. Phycologia 32:234-6. 1993
458
Guillard, R. R. L. Division rates. In: Stein, J.R. (ed), Handbook of phycological
459
methods. Cambridge University Press, Cambridge, pp. 289-312. 1973
460
461
Zhu C.J. & Lee Y.K. Determination of biomass dry weight of marine microalgae.
462
Journal of applied phycology 9: 189-194, 1997.
463
464
Ruiz J, Antequera T, Andres AI, Petron MJ, Muriel E. Improvement of a solid phase
465
extraction method for analysis of lipid fractions in muscle foods. Anal Chim Acta 520:
466
201-205. 2004
467
468
Russell JM, Werne JP. The use of solid phase extraction columns in fatty acid
469
purification. Org Geochem 38: 48-51 2007
470
471
Cooney, M.J, Elsey, D. Jameson, D. Raleigh, B. Fluorescent measurement of
472
microalgal neutral lipids. Note in Journal of Microbiological Methods 68: 639-642.
473
2007
474
475
Ramos, M.J. et al.,. Influence of fatty acid composition of raw materials on biodiesel
476
properties, Bioresources.Technology. 100: 261-268. 2008
477
478
Fábregas, J. Otero, A. Dominguez, A. Patiño, M. Growth rate of the microalgae
479
Tetraselmis suecica changes the biochemical composition of Artemia species. Mar.
480
Biotechnol. 3, 256-263, 2001
481
482
Fábregas, J. Abalde, J. Herrero, C. Cid, A. Yields in biomass and chemical constituents
483
of four commercial important marine microalgae with different culture media.
484
Aquacultural Engineering 10, 99-110, 1991
485
486
Stolte, W. Garcés, E. Ecological aspects of harmful algal in situ population growth
487
rates. Ecological Studies, vol. 189, 2006
488
489
Li, X. Xu, H. Wu, Q. Large-scale biodiesel production from microalgae Chlorella
490
protothecoides through heterotrophic cultivation in bioreactors. Biotechnol Bioeng. 1;
491
98(4):764-71 2007
492
493
Mansour, P. Volkman, J. Blackburn, S. The effect of growth phase on the lipid class,
494
fatty acid and sterol composition in the marine dinoflagellate, Gymnodinium sp. in batch
495
culture. Phytochemistry 63, 145-153. 2003
496
497
Volkman, J.K. Fatty acids of microalgae used as feedstocks in aquaculture. In: Cambie,
498
R.C. (ed) Fats for the future. Ellis Horwood, Chichester, pp. 286-283. 1989
499
500
Dunstan, G.A., Volkman, J.K., Barret, S.M., Garland, C.D. Changes in the lipid
501
composition and maximization of the polyunsaturated fatty acid content of three
502
microalgae grown in mass culture. J. Appl. Phycol. 34, 71-83. 1993
503
504
Reitan, K.L., Rainuzzo, J.R., Olsen, Y. Effect of nutrient limitation on fatty acid and
505
lipid content of marine microalgae. J. Phycol. 30, 972-979. 1994
506
507
Brown, M.R., Dunstan, G.A., Norwood, S.J., Miller, K.A. Effects of harvest stage and
508
light in biochemical composition of the diatom Thalassiosira pseudonana. J. Phycol.
509
34, 712 – 721. 1996
510
511
Hallegraeff, G.M. Nichols, P.D. Volkman, J.K. Blackburn, S. & Everit, D. Pigments,
512
fatty acids, and sterols of the toxic dinoflagellate Gymnodinium catenatum. J. Phycol.
513
27, 591-599. 1999
514
515
Mooney, D.B. Nichols, P.D. De Salas, M.F. & Hallegraeff, G.M. Lipid, fatty acid, and
516
sterols composition of eight species of kareniaceae (Dinophyta): Chemotaxonomy and
517
putative lipid phycotoxins. J. Phycol. 43, 101-111. 2007
518
519
Mathews & Van Holde. Bioquimica. Seg. ed. MacGraw-Hill, Interamericana. 2000
520
521
Qingyu, Gao, Chungfang, Xiong, Wei, Zhang, Yiliang, Yuang, Wenqiao, Wu. Rapid
522
quantification of lipids in microalgae by time domain nuclear resonance. J Microbiol
523
Methods. 75: 437-440. 2008
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Fig. 1 Growth curve of obtained from the populations of different marine microalgae
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incubated at 21°C, at neutral pH, in a salinity of 36, and in prefiltered air. The flasks
526
were maintained using a 12:12 h light:dark cycle at a photon irradiance of 110 μmol
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photons m-2 s-1. Errors bars denote standard deviations among replicates.
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Fig. 2 Total wet weight of six marine microalgae cultures at different phases of the
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growth cycle (lag, exponential, and stationary phases). Errors bars denote standard
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deviations between two replicates. Significances differences are marked *p < 0.05
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Fig. 3 Total lipid content as measured by gas chromatography, with respect to oil
534
concentration (g/l), during the stationary phase of culture. Errors bars denote standard
535
deviations between three replicates.
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Fig. 4 Different types of oil (from Ramos, M.J. et al. 2008) compared with the fatty
538
acids profile obtained from dinoflagellates in this study.
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Fig. 5. Epifluorescent microphotographs (60× magnification) of microalgae stained with
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the fluorochrome Nile red. Neutral lipids are seen as yellows drops. (A) Alexandrium
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andersoni, (B) Karlodinium veneficum, (C) Alexandrium minutum, and (D) Tetraselmis
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suecica
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Table 1. Algal strains analyzed in this study. Strain origin, year of isolation, and
546
identification code are indicated.
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Table 2. Relative abundance (%) of fatty acid composition in different marine
549
microalgae at the stationary phase of culture. SFA, saturated fatty acid; MUFA,
550
monounsaturated fatty acid; PUFA, polyunsaturated fatty acid.
551
(Standard deviation n = 3)
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