Accepted Manuscript Production of biodiesel from carbon sources of macroalgae, Laminaria japonica Xu Xu, Ji Young Kim, Yu Ri Oh, Jong Moon Park PII: DOI: Reference:
S0960-8524(14)00980-8 http://dx.doi.org/10.1016/j.biortech.2014.07.015 BITE 13660
To appear in:
Bioresource Technology
Received Date: Revised Date: Accepted Date:
5 May 2014 2 July 2014 3 July 2014
Please cite this article as: Xu, X., Kim, J.Y., Oh, Y.R., Park, J.M., Production of biodiesel from carbon sources of macroalgae, Laminaria japonica, Bioresource Technology (2014), doi: http://dx.doi.org/10.1016/j.biortech. 2014.07.015
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Production of biodiesel from carbon sources of macroalgae, Laminaria
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japonica
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Xu Xu a, Ji Young Kim b, Yu Ri Oh b, Jong Moon Park a,b,c *
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b
Department of Chemical Engineering,
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School of Environmental Science and Engineering ,
c
Division of Advanced Nuclear Engineering, Pohang University of Science and Technology, San 31, Hyoja-dong, Pohang 790-784, South Korea
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Corresponding author.
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*
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Tel: +82-54-279-2275; Fax: +82-54-279-8299; E-mail: [email protected] (J.M. Park)
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Abstract
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As aquatic biomass which is called “the third generation biomass”, Laminaria japonica (also
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known as Saccharina japonica) consists of mannitol and alginate which are the main
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polysaccharides of algal carbohydrates. In this study, oleaginous yeast (Cryptococcus
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curvatus) was used to produce lipid from carbon sources derived from Laminaria japonica.
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Volatile fatty acids (VFAs) were produced by fermentation of alginate extracted from L.
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japonica. Thereafter, mannitol was mixed with VFAs to culture the oleaginous yeast. The
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highest lipid content was 48.30 %. The composition of the fatty acids was similar to
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vegetable oils. This is the first confirmation of the feasibility of using macroalgae as a carbon
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source for biodiesel production.
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Keywords: Laminaria japonica, Cryptococcus curvatus, Manntiol, Alginate, Volatile fatty
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acids (VFAs), Fatty acid methyl esters (FAMEs), Biodiesel, Macroalgae
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1. Introduction Energy problem has become one of the most serious issues all over the world since energy
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consumption is inevitable for human existence. At this point, a large amount of alternative
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energy appears. Among them, microbial lipids can be used as a renewable alternative
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(biodiesel) to traditional fossil diesel fuel. Biodiesel, a mixture of fatty acid alkyl esters, can
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be obtained from various renewable lipid resources such as vegetable oils, fats and wastes of
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cooking oils (Liu et al., 2008). However, use of edible oils for biodiesel production competes
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with food production and requires land and irrigation water; these requirements and the high
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lignin content of terrestrial plants reduce the economic viability of this production mode.
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Especially in the countries like South Korea which has no vast territory to cultivate a large
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amount of high oil content plants, it is meaningless to use the edible oils for biodiesel
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production.
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On the basis of above mentioned issues, macroalgae which is called as “the 3rd generation
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biomass” is gaining increasingly more attentions as alternative renewable sources of inputs
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for biofuel production since it can deal with these drawbacks of terrestrial biomass and
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produce sustainable bioenergy and materials. Macroalgae (seaweed) have several
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advantageous characteristics such as high yield, low energy cost of production, low cost, low
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contaminant levels and low nutrient requirements, which can be confirmed from the results
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shown in Table 1 (Park, 2012). Macroalgae do not need land and freshwater for cultivation
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which is advantageous to the countries lack of land. In addition, macroalgae have a lower
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cost of the production of food and energy than other energy crops like corn and wheat.
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Because seaweed markets are mainly existed in a few East Asian countries where seaweed is
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utilized as food, hydrocolloids, fertilizer and animal feed (Jung et al., 2013). Macroalgae can
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convert solar energy into chemical energy with higher photosynthetic efficiency (6-8%) than
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terrestrial biomass (1.8-2.2%) (Jung et al., 2013). Moreover, the process of macroalgae 3
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production introduces no contaminants, such as soil which can subsequently lead to operation
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problems (McKendry, 2002). Comparing to the terrestrial plants, no utilization of chemical
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fertilizers and pesticides also reduce the contaminants produced during the biomass
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cultivation. Therefore, macroalgae have high potential as the feedstock of energy production,
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and may be the best choice as a feedstock for biodiesel production.
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Seaweed grows very quickly and can be harvested more than four crops per year compared
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to the crop and forest-derived biomass which are harvested less than two crops and one crop
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per year respectively (Table 1). Globally, about 7 million tons (wet weight) of macroalgae is
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harvested in 2008, and the most harvested species are Laminaria japonica (sea tangle),
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Undaria pinnatifiida (sea mustard), and Porphyra tenera (sea weed laver) (FAO, 2011).
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According to the “Food and Agricultural Organization stats”, the highest production of
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cultured seaweed throughout the world in 2008 was the brown seaweed, especially L.
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japonica which was 4.8 million tons per year. Among them, L. japonica and U. pinnatifiida
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were brown algae and L. japonica was accounted for about 65% of global production. In the
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case of South Korea, the two most cultured species occupying around 70 % of total
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aquaculture production are L. japonica and U. pinnatifiida. In our previous research, the
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composition of L. japonica had been analyzed. As the data shown in Table 2, the amount of
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carbohydrate of L. japonica is higher than that of U. pinnatifiida, which contains 60 – 67 %
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(W/W dry). However, U. pinnatifiida has more protein (Cho, 1995).
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In Table 3, it shows carbohydrate profile in brown algae. The two species of brown algae
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mainly consists of alginate, laminaran, fucoidan (Kim et al., 1995, Pyeun et al., 1977).
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Alginate and mannitol are the main structure and storage compound which account for about
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50% (W/W) of total carbohydrates respectively (Kloareg and Quatrano, 1988).
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Mannitol, a sugar alcohol equivalent to mannose, is one of the major carbon sources of brown algae. It is different from terrestrial biomass storing carbon source as cellulose, 4
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hemicellulose and lignin. Since cellulose is very difficult to be hydrolyzed and lignin cannot
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be used in biorefinery, it is required high costs of pretreatment for these compounds, which is
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the obstacle for commercialization of energy production from the terrestrial biomass.
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However, as mannitol is soluble and available carbohydrate, it can be directly utilized by
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microorganisms as carbon source. Alginate, the most abundant polysaccharide in L. japonica,
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is one of brown algae’s cell wall compounds with cellulose, fucoidan, and protein (Kloareg et
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al., 1986). The cells are strengthened to the algal tissue by both mechanical strength and
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flexibility of alginate (Andresen et al., 1977). In the L. japonica, there are two kinds of uronic
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acids linking each other which are polymannuronate and polyguluronate. The polymers
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accumulate in blocks and bind divalent metal ions through forming gels which are structural
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parts of algae. Therefore, it is less available to contact with alginate lyase. Since it is a
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polysaccharide, the microbial oil production using alginate would require extra
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saccharification or anaerobic fermentation process (Wang et al., 2013). Alginate has been
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used as a food additive for a long time. Its uses are based on the properties of thickening,
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gelling, film formation, stabilizing and general colloidal properties (Aliste et al., 2000). These
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properties are useful in pie, cake, ice cream, and canned food productions, which can reduce
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moisture retention, thicken batter and extend the shelf life.
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It is well known that the first step of anaerobic digestion is hydrolysis which breaks down
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biopolymers to monomers. In this step, carbohydrates, proteins and lipids are respectively
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transformed into sugars, amino acids and fatty acids. These small molecules are fermented to
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carbon dioxide and hydrogen along with several organic acids in the next step called
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acidogenesis (Angenent et al., 2008). The organic acids produced during the acidogenesis can
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be used by some microorganisms as carbon sources. Therefore, based on the processes
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introduced above, L. japonica was chosen as the most appropriate substrate in this study
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which has more carbohydrates than U. pinnatifida. 5
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As one of the most important renewable energy sources, biodiesel is produced through
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transesterification of vegetable oils or animal fats with short chain alcohols. However, high
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cost of raw materials for biodiesel production has become one of the major obstacles for wide
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application. Recently, it is gaining more attention to search for new oil sources for biodiesel
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production. Among them, microbial oils, also known as single cell oils (SCOs), can
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overcome these problems. In fact, all microorganisms can synthesize lipids, but only those
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which can produce more than 20% lipids of their dry biomass are called “oleaginous”
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including bacteria, fungi, and yeasts (Li et al., 2006, Meng et al., 2009). The oleaginous
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yeasts possess higher specific growth rate than molds and algae, which can be considered as
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favorable microorganisms for microbial lipids production (Li et al., 2007). The most
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productive oleaginous yeasts are Yarrowia lipolytica, Cryptococcus curvatus, and
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Rhodosporidium toruloides. They could produce from 40 % to 70 % of biomass (Ratledge
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and Cohen, 2008). The microbial lipids mainly consist of triacylglycerols (TAGs), and other
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components are free fatty acids, other neutral lipids, sterols and polar fractions (Fakas et al.,
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2006). The TAGs produced by oleaginous yeasts are mostly formed of C14 to C18 fatty acids
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which can be used as biofuel because they are similar to vegetable oils (Li et al., 2007, Meng
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et al., 2009).
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C. curvatus, an oleaginous yeast, is able to accumulate up to 60% oils by dry cell weight
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(DCW) (Ratledge, 1991) using economical carbon source . In this study, we assess whether C.
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curvatus can be used to produce significant quantities of lipids from mannitol and alginate
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extracted from L. japonica.
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Because mannitol is water-soluble whereas alginate is not, a stepwise process was used
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(Fig. 1): first, alginate was fermented under anaerobic condition to obtain VFAs which can be
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used by C. curvatus, then the VFAs were mixed with mannitol and then added to the lipid-
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producing fermentor. It was the first time to produce lipid by oleaginous yeast only using the 6
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L. japonica as the sole carbon source without adding any other nutrients.
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2. Materials and Methods
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2.1. Feedstock
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Dried kelp, Laminaria japocia, was purchased from Geumil-do Fisheries Cooperation
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Association (Wando, South Korea), milled to fine powder, and kept in a sealed container at
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room temperature until it was used. For the trials, 90 g of the powder was mixed in 3 L tap
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water to make the concentration 30 g/L and stirred for 6 h at room temperature to maximize
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mannitol recovery rate in the supernatant (based on results of preliminary work), then left
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unstirred to allow solids to precipitate. After 24 h, the mixture had separated into two layers.
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The supernatant (~2 L) was mannitol solution; the bottom layer (~1 L) was mainly alginate
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suspension. The supernatant was aspirated off and steam autoclaved at 121 °C for 20 min.
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The alginate suspension was used for VFA production.
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In the case of feedstock for lipid production, the effluent from the alginate fermentation
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was recovered, centrifuged for 10 min at 400 g-force, and then passed through a 1.20-µm
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filter (GF/C, Whatman, UK) to separate the liquid and solid components. The liquid part was
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then steam autoclaved at 121 °C for 20 min.
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2.2. Inoculum Anaerobic digester sludge from a wastewater treatment plant (Daegu, Korea), screened
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through a sieve (No. 10, diameter: 2 mm), then pretreated by heat at 90 °C for 20 min to
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inactive methanogens (Lee et al., 2008). First, populations of VFA-producing bacteria were
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increased and adapted to conditions in a 7 L fermentor (Biotron LiFlus GR Fernmentor)
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under anaerobic condition. Heat-treated sludge (1.5 L) and 2.5 L of ground kelp (4 L working
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volume) were added to the fermentor. It was operated at 35 °C and stirring rate was 200 ppm. 7
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After enrichment and adaptation, the 4 L content of the fermentor was poured into a 7 L
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continuous stirred tank reactor (CSTR) (Biotron LiFlus GR Fernmentor). Ground kelp mixed
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with tap water (30 g/L) was supplied to the reactor under anaerobic condition using peristaltic
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pumps (model No. 7523-30, 7521-57, Masterflex®, USA). The continuous process was
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operated at 4 d hydraulic retention time with 2.78 mL/min flow rate. After inoculation, the
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fermentor was purged using N2 gas for 10 min, and all instruments were maintained in strictly
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anaerobic condition. The pH was controlled at 7 by an automatic pH controller (Model KB-
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250, K&B) using 5 M NaOH (Samchun) and 1 M HCl (Samchun) solution because this pH
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yielded the highest VFA production rate in a preliminary study. Temperature and stirring rate
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were maintained at 35 °C and 200 rpm respectively. Ultimately, the effluent from the CSTR
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was directly used as the inoculum for alginate fermentation.
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C. curvatus (ATCC 20509) was obtained from the Korean Collection for Type Culture and
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pre-cultured in a medium composed of 1 g/L peptone, 1g/L yeast extract, and 10 g/L
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dextrose. C. curvatus was grown in 250 mL Erlenmeyer flasks containing 50 mL of medium
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and incubated at 28 °C in a rotary shaker which was set to 150 rpm. Before inoculation, seed
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cells were sub-cultured for 24 h in a medium that contained 10 g/L mannitol, 1 g/L peptone,
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and 1g/L yeast extract. The initial pH was 7.2 and the inoculum rate was 10% (v/v).
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2.3. Culture conditions
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Alginate fermentation and lipid production were conducted sequentially (Fig. 1). All
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instruments and reactors for lipid production were steam autoclaved before use. All batch
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tests were conducted in triplicate.
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For VFA production from alginate, a 1.5-L fermentor (Biotron LiFlus GR) containing 1 L
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alginate feedstock (Section 2.1.1) was seeded with 10 % (v/v) inoculum derived from the
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CSTR effluents, then purged using N2 gas for 10 min. pH was controlled at 7 by an automatic 8
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pH controller (Model KB-250, K&B). Temperature was maintained at 35 °C and the
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fermentor was stirred continuously at 400 rpm.
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For lipid production by C. curvatus, a 5-L fermentor (Biotron LiFlus GR) was utilized
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(Fig. 1). It was loaded with 2 L of mannitol supernatant and 1 L VFAs. The pH was
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controlled at 5.5 by an automatic pH controller (Model KB-250, K&B). Lipid production was
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aerobic, so air was supplied at 0.5 VVM using an aerator. Temperature was maintained at
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30 °C and the fermentor was stirred continuously at 400 rpm.
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2.4. Analytical method
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2.4.1. Physico-chemical analytical method
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Total solid (TS), volatile solid (VS), total suspended solid (TSS), volatile suspended solid
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(VSS) and total nitrogen (TN) were measured according to the standard method (APHA-
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AWWA-WEF, 1998). Every soluble sample was passed through a 0.45-µm pore
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polyethersulfone syringe filter (Millipore, USA) to remove insoluble particles.
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Mannitol, VFAs and ethanol concentrations were analyzed using a high performance liquid
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chromatography (HPLC, Agilent Technology 1100 series) equipped with a carbohydrate-
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analysis column (Aminex HPX-87H, BIORAD INC., USA), refractive index detector (RID)
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and diode array detector (DAD). The eluent was 0.004-M H2SO4 and the flow rate was
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0.6 mL/min. The column temperature was maintained at 50 °C. All liquid samples were
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diluted (1:5 or 1:10) with water and passed through a 0.2 µm syringe filter (Millipore, USA).
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2.4.2. Determination of dry cell weight For the dry cell weight (DCW) of C. curvatus was measured by forcing a known volume of
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cell suspension sample through a pre-dried, pre-weighed 0.45-µm nitrocellulose filter
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(Millipore, USA) using a vacuum pump. The samples were dried to constant weight at 9
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110 °C in an oven, then weighed.
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2.4.3. Lipid extraction The extraction of total cellular lipid was performed according to Bligh and Dyer (Bligh and
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Dyer, 1959) with modifications (Bourque and Titorenko, 2009). Lipid was extracted from
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lyophilized biomass using a mixture of chloroform and methanol (2:1 v/v). The mixture was
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centrifuged to partition the lipids into the solvent, then the solvent was evaporated using a
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Nitrogen Evaporation System (Organomation Associates Inc., USA). Lipid content was
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represented as a percentage of dry cell weight (%, W/W). Lipid concentration was defined as
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the amount of extracted cellular lipid in per liter working volume (g/L).
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2.4.4. Fatty acid methyl esters The fatty acid compositions of the lipid produced by C. curvatus were determined by
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analysis of fatty acid methyl esters (FAMEs). The FAMEs were produced by
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transesterification: methanol was added to the extracted lipid with sulfuric acid (2.5 % V/V
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H2SO4/CH3OH) as a catalyst; the reaction was allowed to proceed for 45 min at 90 °C. Then
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1 ml H2O and 2 ml n-hexane were added. The FAMEs dissolved into the n-hexane. The
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solution was centrifuged at 2000 rpm for 15 min, to separate the water from the
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FAME/hexane phase which was then transferred into glass vials using Pasteur pipettes.
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The FAMEs in n-hexane were analyzed using a gas chromatography (6890N, Agilent,
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USA) equipped with a flame ionized detector (FID) and an INNOWAX capillary column
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(Agilent, USA, 30 m × 0.32 mm × 0.5 µm). The column temperature was programmed as
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follows: (1) initial column temperature 100 °C, hold for 5 min, (2) increase to 250 °C at
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10 °C/min, hold for 30 min. The split ratio was 1:10 (v/v). The injector and the detector
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temperatures were both set at 250 °C. FAME components were identified and quantified by 10
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comparing the retention times and peak areas with those of standard solutions.
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3. Results and discussion
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3.1. VFA production by alginate fermentation using mixed cultivation system
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As mentioned in the materials and methods, the inoculum of alginate fermentation was the
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effluent from a CSTR. In fact, the CSTR was a VFA-producing system which can
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continuously produce acetate, propionate and butyrate using L. japonica as sole carbon
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source without adding any other nutrients. This system has been operated for over 1000 d in
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our previous research and stabilized more than 600 d. Therefore, the microbial community in
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the effluent from the CSTR was quite stable.
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To evaluate the profile changes of acids during alginate fermentation, mannitol
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consumption along with the accumulation and consumption of acidogenic products in batch
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reactor were demonstrated (Fig. 2). Although the mannitol and alginate were separated by
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natural sedimentation, the liquid (mannitol solution) could not be completely separated from
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the solid (alginate suspension). Therefore, liquid samples for HPLC analysis after
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centrifugation retained some mannitol (5.8 g/L initially). The products of alginate
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fermentation were acetate, succinate, lactate, formate, acetate, propionate, butyrate and
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ethanol. Ethanol concentration reached 1.0 g/L within 24 h, but gradually decreased to 0 by
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144 h, probably because ethanol initially produced was converted into acetate by
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acidogenesis (Angenent et al., 2008). The depletion of ethanol and production of acetate are
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thermodynamically favorable (Oh et al., 2003). Likewise, succinate, lactate and formate also
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increased to a certain amount and disappeared later. During the period of acidogenesis,
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acetate was produced as a major alginate fermentation product, followed by propionate and
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butyrate. The final concentration of acetate was more than 9.0 g/L at 6.5 d. Propionate
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concentration slowly increased to 3.5 g/L, and remained at that level until 156 h. Butyrate 11
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was produced until 24 h and its concentration began to increase from 36 h, ultimately to a
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concentration which was more than 0.5 g/L. At the end of digestion, the VFAs consisted of
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69.2 % acetate, 25.7 % propionate and 5.1 % butyrate.
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3.2. Cell growth and lipid accumulation by C.curvatus using carbon source derived from
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macroalgae by single batch cultivation system.
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Most of the microorganisms can synthesize lipids, but only the oleaginous strains may
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accumulate more than 20 % W/W lipids on dry cell basis (Papanikolaou and Aggelis, 2011a).
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Microorganisms synthesize lipids by either de novo processes that utilizes hydrophilic
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substances as substrates, or by ex novo processes that utilize hydrophobic substances as
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substrates. In this research, all lipid synthesis occurred by de novo processes. In de novo
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processes, lipid accumulation always happens after depletion of nitrogen (or to a lesser extent
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of other essential nutrient like phosphorus or sulfate) from the medium (Papanikolaou and
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Aggelis, 2011b). In addition, to ensure that VFAs are mostly converted into lipids rather than
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to biomass, these carbon sources should be added to a medium containing enough cells
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(Fontanille et al., 2012).
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Total nitrogen (TN) was 134 mg/L in the mannitol supernatant and 48 mg/L in the VFA
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effluent. Initially 1 L of VFA effluent was added to the 2 L of mannitol substrate; this
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addition diluted manntiol and VFA concentration in the substrate to 2/3 and 1/3 of pre-
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addition levels, respectively. During the first 12 h, VFAs were not dramatically consumed by
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C. curvatus. Lipid concentration and DCW increased slightly at the same time. However,
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lipid content increased rapidly from less than10 % to 30 %. During the next 36 h, lipid
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content, lipid concentration, and DCW increased to a much higher level until 48 h when all
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VFAs were used up by yeasts. As for lipid content, it achieved its highest point (48.30%). It
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is obvious that the concentration of mannitol was almost constant until 48 h, which could 12
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demonstrate that in lipid-producing metabolic pathway of C. curvatus, VFAs were more
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preferable than mannitol as a carbon sources under pH of 5.5. After the exhaustion of VFAs,
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mannitol began to gradually decrease and was consumed up within 84 h. In the meantime,
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DCW increased to 3.60 g/L until 72 h and then remained stable; whereas lipid content
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suddenly reduced which resulted in the decrease of lipid concentration. This phenomenon
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which is called as “lipid turnover” is related with storage lipid degradation (Chen et al., 2012,
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Peng et al., 2013). Microbial lipid turnover has been extensively studied in Y. lipolytic and C.
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echinulata (Papanikolaou and Aggelis, 2011a). Oleaginous yeasts consume their own
308
intracellular lipids to maintain lipid-free biomass when carbon source is exhausted or carbon
309
uptake rate decreases. Because there was still some mannitol left in the substrate, the lipid
310
turnover could result from the reduced carbon source uptake rate. When VFAs were
311
exhausted, C. curvatus had to consume mannitol as substitution, which could make the
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carbon source uptake rate decreased during this process. From 72 h, lipid content increased
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again with more mannitol assimilated by the yeasts.
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Several research groups have used VFAs to produce lipids by culturing oleaginous yeasts
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including Yarrowia lipolytica and Cryptococcus curvatus (Christophe et al., 2012, Fontanille
316
et al., 2012). The most significant difference was that in our study the microorganisms
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utilized mannitol and VFAs which were all derived from seaweed without addition of any
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other nutrients and reagents, whereas in other research, oleaginous yeasts used a synthetic
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medium containing glucose and acetate as substrates. Moreover, in other research the ratio of
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carbon to nitrogen (C/N) was artificially controlled by continuously adding carbon and
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nitrogen sources to achieve high lipid concentration and content. But in our work, we just
322
utilized the original mannitol supernatant and alginate fermentation effluents to culture the
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oleaginous yeasts.
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We compared our results to other researches that also used VFAs or acetate as carbon 13
325
sources (Table 4). In fact, little research has examined the feasibility of using mannitol as a
326
carbon source for oleaginous yeasts. In addition, the strains and concentration of carbon
327
source used were all different with each research. Therefore, the only relevant comparisons
328
was of the value of lipid content (% W/W). Among these results, apparently the lipid content
329
in this work (48.30%) was higher than that of any other researches. It indicated that the
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mannitol and VFAs derived from alginate anaerobic fermentation were favorable for
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microbial lipid production even without using synthetic medium or artificially adding any
332
other nutrients.
333 334
3.3. Fatty acid composition analysis
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The fatty acids produced in our study consisted mainly of palmitic acid (C16:0), stearic
336
acid (C18:0), oleic acid (C18:1), and linoleic acid (C18:2) (Table 5). Others indicates C8:0,
337
C10:0, C12:0, C14:0, C16:1, C18:3, C20:0, C20:1, C20:2, and C24:0. This composition is
338
similar to those of vegetable oils which have been utilized for industrial production of
339
biodiesel (Adamska ET, 2004). Over time, some changes in lipid composition occurred.
340
Obviously, there were three different trends shown in each fatty acid composition. As for
341
C18:0 and C18:1, they both increased during the period of VFAs consumption until 48 h,
342
while the percentages decreased when C. curvatus started to assimilate mannitol as the sole
343
carbon source. In contrast, in the case of palmitic acid, it was gradually reduced during the
344
first 48 h, then accumulated again until the end. However, the profile of linoleic acid
345
followed a totally different pattern. The percentage increased throughout the period from
346
8.95 % to 17.15 %, irrespective of which carbon source yeasts used. This composition was
347
comparable to vegetable fatty acids indicating that the fatty acids accumulated by C. curvatus
348
are appropriate for use in biodiesel production (Li et al., 2007, Meng et al., 2009).
349 14
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4. Conclusions
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We designed an efficient batch system for lipid production by the oleaginous yeast
352
Cryptococcus curvatus using two different carbon sources derived from a brown macroalga,
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Laminaria japonica: mannitol and VFAs obtained by anaerobic fermentation of alginate with
354
a mixed culture. This is the first demonstration of microbial oil production by bioconversion
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of macroalgae without addition of any other carbon sources or nutrients. This study has
356
shown that mannitol and VFAs derived from alginate fermentation are favorable feedstock
357
for lipid production. Future work can focus on optimizing and scaling up this process.
358 359
Acknowledgements
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This research was supported by Basic Science Research Program through the National
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Research Foundation of South Korea (NRF) funded by the Ministry of Education, Science
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and Technology (Grant number 2011-0001108), the Advanced Biomass R&D Center (ABC)
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of South Korea Grant funded by the Ministry of Education, Science and Technology (ABC-
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2011-0028387), Marine Biotechnology Program Funded by Ministry of Land, Transport and
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Maritime Affairs of South Korean Government, South Korea and the Manpower
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Development Program for Marine Energy funded by Ministry of Land, Transportation and
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Maritime Affairs (MLTM) of South Korean government and by the World Class University
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(WCU) program through the National Research Foundation of South Korea funded by the
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Ministry of Education, Science and Technology (R31-30005).
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15
372
Reference
373 374 375 376 377 378 379 380 381 382 383 384 385 386 387 388 389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406 407 408 409 410 411 412 413 414 415 416 417 418 419
[1]Adamska ET, C.T., Kaczmarek Z, Szala L. 2004. Multivariate approach to evaluating the fatty acid composition of seed oil in a doubled haploid population of winter oilseed rape (Brassica napus L.,). J. Appl. Genet 45, 6. [2]Aliste, A.J., Vieira, F.F., Del Mastro, N.L., 2000. Radiation effects on agar, alginates and carrageenan to be used as food additives. Radiat. Phys. Chem. 57, 305-308. [3]Andresen, I.L., Skipnes, O., Smidsrød, O., Østgaard, K., Hemmer, P. 1977. Some biological functions of matrix components in benthic algae in relation to their chemistry and the composition of seawater. ACS Symp. Ser. ACS Publications. pp. 361-381. [4]Angenent, L.T., Wrenn, B.A., Wall, J., Harwood, C., Demain, A. 2008. Optimizing mixedculture bioprocessing to convert wastes into bionergy. Bioenergy. ASM Press. pp. 179-194. [5]APHA-AWWA-WEF. 1998. Standard Methods for the Examination of Water and Wastewater, 20th ed., American Public Health Association. Washington, D.C. [6]Bligh, E., Dyer, W.J., 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911-917. [7]Bourque, S.D., Titorenko, V.I., 2009. A quantitative assessment of the yeast lipidome using electrospray ionization mass spectrometry. Journal of visualized experiments: JoVE. [8]Chen, X.F., Huang, C., Xiong, L., Chen, X.D., Chen, Y., Ma, L.L., 2012. Oil production on wastewaters after butanol fermentation by oleaginous yeast Trichosporon coremiiforme. Bioresour. Technol. 118, 594-7. [9]Cho, D.M., Kim, D.S., Lee, D.S., Kim, H.R., Pyeun, J.H., 1995. Trace Components and Functional Saccharides in Seaweed. 1: Changes in Proximate Composition and Trace Elements According to the Harvest Season and Places. Korean J. Fish. Aquat. Sci. 28, 49-59. [10]Christophe, G., Deo, J.L., Kumar, V., Nouaille, R., Fontanille, P., Larroche, C., 2012. Production of oils from acetic acid by the oleaginous yeast Cryptococcus curvatus. Appl. Biochem. Biotechnol. 167, 1270-9. [11]Fakas, S., Papanikolaou, S., Galiotou-Panayotou, M., Komaitis, M., Aggelis, G., 2006. Lipids of Cunninghamella echinulata with emphasis to gamma-linolenic acid distribution among lipid classes. Appl. Microbiol. Biotechnol. 73, 676-83. [12]FAO. 2011. 2008 Fishery and Aquaculture Statistics. Available from: . [13]Fei, Q., Chang, H.N., Shang, L., Choi, J.D.R., Kim, N., Kang, J., 2011a. The effect of volatile fatty acids as a sole carbon source on lipid accumulation by Cryptococcus albidus for biodiesel production. Bioresour. Technol. 102, 2695-2701. [14]Fei, Q., Chang, H.N., Shang, L.A., Choi, J.D.R., 2011b. Exploring low-cost carbon sources for microbial lipids production by fed-batch cultivation of Cryptococcus albidus. Biotechnol. Bioproc. E 16, 482-487. [15]Fontanille, P., Kumar, V., Christophe, G., Nouaille, R., Larroche, C., 2012. Bioconversion of volatile fatty acids into lipids by the oleaginous yeast Yarrowia lipolytica. Bioresour. Technol. 114, 443-9. [16]Jung, K.A., Lim, S.R., Kim, Y., Park, J.M., 2013. Potentials of macroalgae as feedstocks for biorefinery. Bioresour. Technol. 135, 182-90. [17]Kim, D.S., Cho, D.M., Lee, D.S., Kim, H.R., Pyeun, J.H., 1995. Trace components and functional saccharides in marine algae. J. Korean Fish. Soc. 28, 270-278. [18]Kloareg, B., Demarty, M., Mabeau, S., 1986. Polyanionic characteristics of purified 16
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sulphated homofucans from brown algae. Int. J. Biol. Macromol. 8, 380-386. [19]Kloareg, B., Quatrano, R., 1988. Structure of the cell walls of marine algae and ecophysiological functions of the matrix polysaccharides. Oceanogr. Mar. Biol. Annu. Rev. 26, 259-315. [20]Lee, M., Hidaka, T., Tsuno, H., 2008. Effect of temperature on performance and microbial diversity in hyperthermophilic digester system fed with kitchen garbage. Bioresour. Technol. 99, 6852-60. [21]Li, Y., Zhao, Z., Bai, F., 2007. High-density cultivation of oleaginous yeast Rhodosporidium toruloides Y4 in fed-batch culture. Enzyme Microb. Technol. 41, 312-317. [22]Li, Y.H., Liu, B., Zhao, Z.B., Bai, F.W., 2006. Optimization of Culture Conditions for Lipid Production by Rhodosporidium toruloides. Chin. J. Biotechnol. 22, 650-656. [23]Liu, Z.Y., Wang, G.C., Zhou, B.C., 2008. Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresour. Technol. 99, 4717-22. [24]McKendry, P., 2002. Energy production from biomass (part 1): overview of biomass. Bioresour. Technol. 83, 37-46. [25]Meng, X., Yang, J., Xu, X., Zhang, L., Nie, Q., Xian, M., 2009. Biodiesel production from oleaginous microorganisms. Renew. Energ 34, 1-5. [26]Oh, S.E., Van Ginkel, S., Logan, B.E., 2003. The relative effectiveness of pH control and heat treatment for enhancing biohydrogen gas production. Environ. Sci. Technol. 37, 5186-5190. [27]Papanikolaou, S., Aggelis, G., 2011a. Lipids of oleaginous yeasts. Part I: Biochemistry of single cell oil production. Eur. J. Lipid Sci. Technol. 113, 1031-1051. [28]Papanikolaou, S., Aggelis, G., 2011b. Lipids of oleaginous yeasts. Part II: Technology and potential applications. Eur. J. Lipid Sci. Technol. 113, 1052-1073. [29]Park, J.M. 2012. PETROFED (Petroleum Federation of India). Trends and Prospects of Biorefinery Research in Korea. Available from: . [30]Peng, W.F., Huang, C., Chen, X.F., Xiong, L., Chen, X.D., Chen, Y., Ma, L.L., 2013. Microbial conversion of wastewater from butanol fermentation to microbial oil by oleaginous yeast Trichosporon dermatis. Renew. Energ 55, 31-34. [31]Pyeun, J.H., Park, Y.H., Lee, K.H., 1977. Factors involved in the quality retention of cultured Undaria pinnatifida. Bull. Korean Fish. Soc 10, 125-135. [32]Ratledge, C., 1991. Microorganisms for lipids. Acta Biotechnol. 11, 429-438. [33]Ratledge, C., Cohen, Z., 2008. Microbial and algal oils: Do they have a future for biodiesel or as commodity oils. Lipid Technol. 20, 155-160. [34]Wang, J., Kim, Y.M., Rhee, H.S., Lee, M.W., Park, J.M., 2013. Bioethanol production from mannitol by a newly isolated bacterium, Enterobacter sp. JMP3. Bioresour. Technol. 135, 199-206.
459 460 461
17
462 463
Figure legends:
464
Fig. 1. Schematic diagram of alginate fermentation and lipid production processes.
465 466
Fig. 2. Profile of various carbon sources and volatile suspended solid (VSS) variations
467
during the alginate fermentation. (▲) VSS; (◇) mannitol; (○) acetic acid; (▼)
468
propionic acid; (□) butyric acid; (◆) ethanol; (ⅹ) formic acid; (▽) lactic acid; (△)
469
succinic acid.
470 471
Fig. 3. Profile of mannitol (◇), acetic acid (○), propionic acid (▼), butyric acid (□),
472
dry cell weight (DCW)(●), lipid concentration (▲), lipid content (■) during a two-step
473
batch cultivation of Cryptococcus curvatus in a 5 L fermentor.
18
st
nd
rd
Table 1. Comparisons between 1 , 2 , and 3 generation biomass Resourcea
Crop
Forest-derived
Seaweed
Harvesting
1 – 2 /yr
1 / 8yr
4 – 6 /yr
Production (ton/ha)
180
9
565
CO2 uptake (ton/ha)
5 – 10
4.6
36.7
Energy yield (%)
30 - 35
20 – 25
> 45
Cost ($/L)
0.2 – 0.3
0.4
0.2
a
(Park, 2012)
19
Table 2. Composition of Laminaria japonica and Undaria pinnadifida
(% w/w, dry base)
Protein
Lipid
Ash
Carbohydrate
Laminaria japonica
6.8 – 10.3
7.2 – 11.5
13.8 – 21.1
60.9 – 67.0
Undaria pinnadifidaa
13.7 – 28.7
3.6 – 6.2
29.4 – 46.5
26.5 – 42.8
a
(Cho, 1995)
20
Table 3. Characteristic algal polysaccharides in Laminaria japonica and Undaria pinnadifida Algal carbohydrate (% w/w, dry base) Alginate
Fucoidan
Laminaran
Mannitol
Total
Laminaria japonica
14.6 – 29.5
0 – 0.1
3.7 – 4.2
22.5 – 33.2
40.8 – 67.0
Undaria pinnadifidaa
22.3 – 32.9
1.4
3.6
4.1 – 7.3
31.4 – 45.2
a
(Kim et al., 1995, Pyeun et al., 1977)
21
Table 4. Lipid production by oleaginous yeasts using different substrate in two-step system Strains
Carbon source (g/L)
DCW (g/L)
Lipid conc. (g/L)
Lipid content (% W/W)
Reference
C. albidus
VFAs (2.00)
1.16
0.31
27.00
(Fei et al., 2011a)
C. albidus
Acetate (6.00)
2.85
0.74
25.80
(Fei et al., 2011b)
Y. lipolytica
Acetate (12.00)
5.98
1.84
30.76
(Fontanille et al., 2012)
C. curvatus
Mannitol + VFAs (9.30)
3.60
1.30
48.30
This study
a
Calculated based on references.
22
Table 5. Fatty acid methyl ester (FAME) profile of C. curvatus Fatty acid methyl ester (FAME) profile of C. curvatus (%) Fermentation time (h)
Palmitic AME (C16:0)
Stearic AME (C18:0)
Oleic AME (C18:1)
Linoleic AME (C18:2)
Others
0
25.88
12.83
48.16
8.95
4.18
12
25.62
11.64
48.36
10.65
3.73
24
21.16
15.45
49.38
11.41
2.60
36
14.72
22.41
50.34
11.49
1.05
48
14.70
19.46
52.26
12.78
0.80
60
15.33
18.20
52.13
13.29
1.06
72
16.75
15.76
49.31
16.32
1.86
84
18.18
14.03
48.71
17.15
1.93
a
Others: C8:0, C10:0, C12:0, C14:0, C16:1, C18:3, C20:0, C20:1, C20:2, and C24:0.
23
a
Aerator
Flow Meter
pH Controller
Mannitol Supernatant
Lipid Fermentor
VFAs
Substrate (30 g/L Laminaria japonica)
pH Controller Alginate Slurry
Alginate Fermentor
Fig. 1. Schematic diagram of alginate fermentation and lipid production processes. 24
12
40
30 8
6
20
VSS (g/L)
Concentration (g/L)
10
4 10 2
0
0 0
24
48
72
96
120
144
Time (h) Fig. 2. Profile of various carbon sources and volatile suspended solid (VSS) variations during alginate fermentation. (▲) VSS; (◇) mannitol; (○) acetic acid; (▼) propionic acid; (□) butyric acid; (◆) ethanol; (ⅹ) formic acid; (▽) lactic acid; (△) succinic acid.
25
2.0
50
40 1.5
6 1.0 4
30
20
Lipid content (%)
8
Lipid concn. (g/L)
DCW, Mannitol and VFAs (g/L)
10
0.5 2
10
0
0.0 0
12
24
36
48
60
72
0
84
Time (h) Fig. 3. Profile of mannitol (◇), acetic acid (○), propionic acid (▼), butyric acid (□), dry cell weight (DCW)(●), lipid concentration (▲), lipid content (■) during a batch cultivation of Cryptococcus curvatus in a 5 L fermentor.
26
H I G H LI G H TS ▶This
is the first demonstration of microbial oil production by using macroalgae.
▶Oleaginous ▶Carbon
sources were all derived from Laminaria japonica.
▶No addition ▶The
yeast Cryptococcus curvatus was used for microbial oil production.
of any other nutrients or synthetic medium was used.
composition of the fatty acids was found similar to vegetable oils.
27
S0960-8524(14)00980-8 http://dx.doi.org/10.1016/j.biortech.2014.07.015 BITE 13660
To appear in:
Bioresource Technology
Received Date: Revised Date: Accepted Date:
5 May 2014 2 July 2014 3 July 2014
Please cite this article as: Xu, X., Kim, J.Y., Oh, Y.R., Park, J.M., Production of biodiesel from carbon sources of macroalgae, Laminaria japonica, Bioresource Technology (2014), doi: http://dx.doi.org/10.1016/j.biortech. 2014.07.015
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1 2
Production of biodiesel from carbon sources of macroalgae, Laminaria
3
japonica
4
Xu Xu a, Ji Young Kim b, Yu Ri Oh b, Jong Moon Park a,b,c *
5 6 a
7
b
Department of Chemical Engineering,
8 9
School of Environmental Science and Engineering ,
c
Division of Advanced Nuclear Engineering, Pohang University of Science and Technology, San 31, Hyoja-dong, Pohang 790-784, South Korea
10 11 12 13 14 15 16 17 18 19
Corresponding author.
20
*
21
Tel: +82-54-279-2275; Fax: +82-54-279-8299; E-mail: [email protected] (J.M. Park)
22 23
1
24
Abstract
25
As aquatic biomass which is called “the third generation biomass”, Laminaria japonica (also
26
known as Saccharina japonica) consists of mannitol and alginate which are the main
27
polysaccharides of algal carbohydrates. In this study, oleaginous yeast (Cryptococcus
28
curvatus) was used to produce lipid from carbon sources derived from Laminaria japonica.
29
Volatile fatty acids (VFAs) were produced by fermentation of alginate extracted from L.
30
japonica. Thereafter, mannitol was mixed with VFAs to culture the oleaginous yeast. The
31
highest lipid content was 48.30 %. The composition of the fatty acids was similar to
32
vegetable oils. This is the first confirmation of the feasibility of using macroalgae as a carbon
33 34
source for biodiesel production.
35 36
Keywords: Laminaria japonica, Cryptococcus curvatus, Manntiol, Alginate, Volatile fatty
37
acids (VFAs), Fatty acid methyl esters (FAMEs), Biodiesel, Macroalgae
38 39 40 41 42 43 44 45 46 47 48 49
2
50 51
1. Introduction Energy problem has become one of the most serious issues all over the world since energy
52
consumption is inevitable for human existence. At this point, a large amount of alternative
53
energy appears. Among them, microbial lipids can be used as a renewable alternative
54
(biodiesel) to traditional fossil diesel fuel. Biodiesel, a mixture of fatty acid alkyl esters, can
55
be obtained from various renewable lipid resources such as vegetable oils, fats and wastes of
56
cooking oils (Liu et al., 2008). However, use of edible oils for biodiesel production competes
57
with food production and requires land and irrigation water; these requirements and the high
58
lignin content of terrestrial plants reduce the economic viability of this production mode.
59
Especially in the countries like South Korea which has no vast territory to cultivate a large
60
amount of high oil content plants, it is meaningless to use the edible oils for biodiesel
61
production.
62
On the basis of above mentioned issues, macroalgae which is called as “the 3rd generation
63
biomass” is gaining increasingly more attentions as alternative renewable sources of inputs
64
for biofuel production since it can deal with these drawbacks of terrestrial biomass and
65
produce sustainable bioenergy and materials. Macroalgae (seaweed) have several
66
advantageous characteristics such as high yield, low energy cost of production, low cost, low
67
contaminant levels and low nutrient requirements, which can be confirmed from the results
68
shown in Table 1 (Park, 2012). Macroalgae do not need land and freshwater for cultivation
69
which is advantageous to the countries lack of land. In addition, macroalgae have a lower
70
cost of the production of food and energy than other energy crops like corn and wheat.
71
Because seaweed markets are mainly existed in a few East Asian countries where seaweed is
72
utilized as food, hydrocolloids, fertilizer and animal feed (Jung et al., 2013). Macroalgae can
73
convert solar energy into chemical energy with higher photosynthetic efficiency (6-8%) than
74
terrestrial biomass (1.8-2.2%) (Jung et al., 2013). Moreover, the process of macroalgae 3
75
production introduces no contaminants, such as soil which can subsequently lead to operation
76
problems (McKendry, 2002). Comparing to the terrestrial plants, no utilization of chemical
77
fertilizers and pesticides also reduce the contaminants produced during the biomass
78
cultivation. Therefore, macroalgae have high potential as the feedstock of energy production,
79
and may be the best choice as a feedstock for biodiesel production.
80
Seaweed grows very quickly and can be harvested more than four crops per year compared
81
to the crop and forest-derived biomass which are harvested less than two crops and one crop
82
per year respectively (Table 1). Globally, about 7 million tons (wet weight) of macroalgae is
83
harvested in 2008, and the most harvested species are Laminaria japonica (sea tangle),
84
Undaria pinnatifiida (sea mustard), and Porphyra tenera (sea weed laver) (FAO, 2011).
85
According to the “Food and Agricultural Organization stats”, the highest production of
86
cultured seaweed throughout the world in 2008 was the brown seaweed, especially L.
87
japonica which was 4.8 million tons per year. Among them, L. japonica and U. pinnatifiida
88
were brown algae and L. japonica was accounted for about 65% of global production. In the
89
case of South Korea, the two most cultured species occupying around 70 % of total
90
aquaculture production are L. japonica and U. pinnatifiida. In our previous research, the
91
composition of L. japonica had been analyzed. As the data shown in Table 2, the amount of
92
carbohydrate of L. japonica is higher than that of U. pinnatifiida, which contains 60 – 67 %
93
(W/W dry). However, U. pinnatifiida has more protein (Cho, 1995).
94
In Table 3, it shows carbohydrate profile in brown algae. The two species of brown algae
95
mainly consists of alginate, laminaran, fucoidan (Kim et al., 1995, Pyeun et al., 1977).
96
Alginate and mannitol are the main structure and storage compound which account for about
97
50% (W/W) of total carbohydrates respectively (Kloareg and Quatrano, 1988).
98 99
Mannitol, a sugar alcohol equivalent to mannose, is one of the major carbon sources of brown algae. It is different from terrestrial biomass storing carbon source as cellulose, 4
100
hemicellulose and lignin. Since cellulose is very difficult to be hydrolyzed and lignin cannot
101
be used in biorefinery, it is required high costs of pretreatment for these compounds, which is
102
the obstacle for commercialization of energy production from the terrestrial biomass.
103
However, as mannitol is soluble and available carbohydrate, it can be directly utilized by
104
microorganisms as carbon source. Alginate, the most abundant polysaccharide in L. japonica,
105
is one of brown algae’s cell wall compounds with cellulose, fucoidan, and protein (Kloareg et
106
al., 1986). The cells are strengthened to the algal tissue by both mechanical strength and
107
flexibility of alginate (Andresen et al., 1977). In the L. japonica, there are two kinds of uronic
108
acids linking each other which are polymannuronate and polyguluronate. The polymers
109
accumulate in blocks and bind divalent metal ions through forming gels which are structural
110
parts of algae. Therefore, it is less available to contact with alginate lyase. Since it is a
111
polysaccharide, the microbial oil production using alginate would require extra
112
saccharification or anaerobic fermentation process (Wang et al., 2013). Alginate has been
113
used as a food additive for a long time. Its uses are based on the properties of thickening,
114
gelling, film formation, stabilizing and general colloidal properties (Aliste et al., 2000). These
115
properties are useful in pie, cake, ice cream, and canned food productions, which can reduce
116
moisture retention, thicken batter and extend the shelf life.
117
It is well known that the first step of anaerobic digestion is hydrolysis which breaks down
118
biopolymers to monomers. In this step, carbohydrates, proteins and lipids are respectively
119
transformed into sugars, amino acids and fatty acids. These small molecules are fermented to
120
carbon dioxide and hydrogen along with several organic acids in the next step called
121
acidogenesis (Angenent et al., 2008). The organic acids produced during the acidogenesis can
122
be used by some microorganisms as carbon sources. Therefore, based on the processes
123
introduced above, L. japonica was chosen as the most appropriate substrate in this study
124
which has more carbohydrates than U. pinnatifida. 5
125
As one of the most important renewable energy sources, biodiesel is produced through
126
transesterification of vegetable oils or animal fats with short chain alcohols. However, high
127
cost of raw materials for biodiesel production has become one of the major obstacles for wide
128
application. Recently, it is gaining more attention to search for new oil sources for biodiesel
129
production. Among them, microbial oils, also known as single cell oils (SCOs), can
130
overcome these problems. In fact, all microorganisms can synthesize lipids, but only those
131
which can produce more than 20% lipids of their dry biomass are called “oleaginous”
132
including bacteria, fungi, and yeasts (Li et al., 2006, Meng et al., 2009). The oleaginous
133
yeasts possess higher specific growth rate than molds and algae, which can be considered as
134
favorable microorganisms for microbial lipids production (Li et al., 2007). The most
135
productive oleaginous yeasts are Yarrowia lipolytica, Cryptococcus curvatus, and
136
Rhodosporidium toruloides. They could produce from 40 % to 70 % of biomass (Ratledge
137
and Cohen, 2008). The microbial lipids mainly consist of triacylglycerols (TAGs), and other
138
components are free fatty acids, other neutral lipids, sterols and polar fractions (Fakas et al.,
139
2006). The TAGs produced by oleaginous yeasts are mostly formed of C14 to C18 fatty acids
140
which can be used as biofuel because they are similar to vegetable oils (Li et al., 2007, Meng
141
et al., 2009).
142
C. curvatus, an oleaginous yeast, is able to accumulate up to 60% oils by dry cell weight
143
(DCW) (Ratledge, 1991) using economical carbon source . In this study, we assess whether C.
144
curvatus can be used to produce significant quantities of lipids from mannitol and alginate
145
extracted from L. japonica.
146
Because mannitol is water-soluble whereas alginate is not, a stepwise process was used
147
(Fig. 1): first, alginate was fermented under anaerobic condition to obtain VFAs which can be
148
used by C. curvatus, then the VFAs were mixed with mannitol and then added to the lipid-
149
producing fermentor. It was the first time to produce lipid by oleaginous yeast only using the 6
150
L. japonica as the sole carbon source without adding any other nutrients.
151 152
2. Materials and Methods
153
2.1. Feedstock
154
Dried kelp, Laminaria japocia, was purchased from Geumil-do Fisheries Cooperation
155
Association (Wando, South Korea), milled to fine powder, and kept in a sealed container at
156
room temperature until it was used. For the trials, 90 g of the powder was mixed in 3 L tap
157
water to make the concentration 30 g/L and stirred for 6 h at room temperature to maximize
158
mannitol recovery rate in the supernatant (based on results of preliminary work), then left
159
unstirred to allow solids to precipitate. After 24 h, the mixture had separated into two layers.
160
The supernatant (~2 L) was mannitol solution; the bottom layer (~1 L) was mainly alginate
161
suspension. The supernatant was aspirated off and steam autoclaved at 121 °C for 20 min.
162
The alginate suspension was used for VFA production.
163
In the case of feedstock for lipid production, the effluent from the alginate fermentation
164
was recovered, centrifuged for 10 min at 400 g-force, and then passed through a 1.20-µm
165
filter (GF/C, Whatman, UK) to separate the liquid and solid components. The liquid part was
166
then steam autoclaved at 121 °C for 20 min.
167 168 169
2.2. Inoculum Anaerobic digester sludge from a wastewater treatment plant (Daegu, Korea), screened
170
through a sieve (No. 10, diameter: 2 mm), then pretreated by heat at 90 °C for 20 min to
171
inactive methanogens (Lee et al., 2008). First, populations of VFA-producing bacteria were
172
increased and adapted to conditions in a 7 L fermentor (Biotron LiFlus GR Fernmentor)
173
under anaerobic condition. Heat-treated sludge (1.5 L) and 2.5 L of ground kelp (4 L working
174
volume) were added to the fermentor. It was operated at 35 °C and stirring rate was 200 ppm. 7
175
After enrichment and adaptation, the 4 L content of the fermentor was poured into a 7 L
176
continuous stirred tank reactor (CSTR) (Biotron LiFlus GR Fernmentor). Ground kelp mixed
177
with tap water (30 g/L) was supplied to the reactor under anaerobic condition using peristaltic
178
pumps (model No. 7523-30, 7521-57, Masterflex®, USA). The continuous process was
179
operated at 4 d hydraulic retention time with 2.78 mL/min flow rate. After inoculation, the
180
fermentor was purged using N2 gas for 10 min, and all instruments were maintained in strictly
181
anaerobic condition. The pH was controlled at 7 by an automatic pH controller (Model KB-
182
250, K&B) using 5 M NaOH (Samchun) and 1 M HCl (Samchun) solution because this pH
183
yielded the highest VFA production rate in a preliminary study. Temperature and stirring rate
184
were maintained at 35 °C and 200 rpm respectively. Ultimately, the effluent from the CSTR
185
was directly used as the inoculum for alginate fermentation.
186
C. curvatus (ATCC 20509) was obtained from the Korean Collection for Type Culture and
187
pre-cultured in a medium composed of 1 g/L peptone, 1g/L yeast extract, and 10 g/L
188
dextrose. C. curvatus was grown in 250 mL Erlenmeyer flasks containing 50 mL of medium
189
and incubated at 28 °C in a rotary shaker which was set to 150 rpm. Before inoculation, seed
190
cells were sub-cultured for 24 h in a medium that contained 10 g/L mannitol, 1 g/L peptone,
191
and 1g/L yeast extract. The initial pH was 7.2 and the inoculum rate was 10% (v/v).
192 193
2.3. Culture conditions
194
Alginate fermentation and lipid production were conducted sequentially (Fig. 1). All
195
instruments and reactors for lipid production were steam autoclaved before use. All batch
196
tests were conducted in triplicate.
197
For VFA production from alginate, a 1.5-L fermentor (Biotron LiFlus GR) containing 1 L
198
alginate feedstock (Section 2.1.1) was seeded with 10 % (v/v) inoculum derived from the
199
CSTR effluents, then purged using N2 gas for 10 min. pH was controlled at 7 by an automatic 8
200
pH controller (Model KB-250, K&B). Temperature was maintained at 35 °C and the
201
fermentor was stirred continuously at 400 rpm.
202
For lipid production by C. curvatus, a 5-L fermentor (Biotron LiFlus GR) was utilized
203
(Fig. 1). It was loaded with 2 L of mannitol supernatant and 1 L VFAs. The pH was
204
controlled at 5.5 by an automatic pH controller (Model KB-250, K&B). Lipid production was
205
aerobic, so air was supplied at 0.5 VVM using an aerator. Temperature was maintained at
206
30 °C and the fermentor was stirred continuously at 400 rpm.
207 208
2.4. Analytical method
209
2.4.1. Physico-chemical analytical method
210
Total solid (TS), volatile solid (VS), total suspended solid (TSS), volatile suspended solid
211
(VSS) and total nitrogen (TN) were measured according to the standard method (APHA-
212
AWWA-WEF, 1998). Every soluble sample was passed through a 0.45-µm pore
213
polyethersulfone syringe filter (Millipore, USA) to remove insoluble particles.
214
Mannitol, VFAs and ethanol concentrations were analyzed using a high performance liquid
215
chromatography (HPLC, Agilent Technology 1100 series) equipped with a carbohydrate-
216
analysis column (Aminex HPX-87H, BIORAD INC., USA), refractive index detector (RID)
217
and diode array detector (DAD). The eluent was 0.004-M H2SO4 and the flow rate was
218
0.6 mL/min. The column temperature was maintained at 50 °C. All liquid samples were
219
diluted (1:5 or 1:10) with water and passed through a 0.2 µm syringe filter (Millipore, USA).
220 221 222
2.4.2. Determination of dry cell weight For the dry cell weight (DCW) of C. curvatus was measured by forcing a known volume of
223
cell suspension sample through a pre-dried, pre-weighed 0.45-µm nitrocellulose filter
224
(Millipore, USA) using a vacuum pump. The samples were dried to constant weight at 9
225
110 °C in an oven, then weighed.
226 227 228
2.4.3. Lipid extraction The extraction of total cellular lipid was performed according to Bligh and Dyer (Bligh and
229
Dyer, 1959) with modifications (Bourque and Titorenko, 2009). Lipid was extracted from
230
lyophilized biomass using a mixture of chloroform and methanol (2:1 v/v). The mixture was
231
centrifuged to partition the lipids into the solvent, then the solvent was evaporated using a
232
Nitrogen Evaporation System (Organomation Associates Inc., USA). Lipid content was
233
represented as a percentage of dry cell weight (%, W/W). Lipid concentration was defined as
234
the amount of extracted cellular lipid in per liter working volume (g/L).
235 236 237
2.4.4. Fatty acid methyl esters The fatty acid compositions of the lipid produced by C. curvatus were determined by
238
analysis of fatty acid methyl esters (FAMEs). The FAMEs were produced by
239
transesterification: methanol was added to the extracted lipid with sulfuric acid (2.5 % V/V
240
H2SO4/CH3OH) as a catalyst; the reaction was allowed to proceed for 45 min at 90 °C. Then
241
1 ml H2O and 2 ml n-hexane were added. The FAMEs dissolved into the n-hexane. The
242
solution was centrifuged at 2000 rpm for 15 min, to separate the water from the
243
FAME/hexane phase which was then transferred into glass vials using Pasteur pipettes.
244
The FAMEs in n-hexane were analyzed using a gas chromatography (6890N, Agilent,
245
USA) equipped with a flame ionized detector (FID) and an INNOWAX capillary column
246
(Agilent, USA, 30 m × 0.32 mm × 0.5 µm). The column temperature was programmed as
247
follows: (1) initial column temperature 100 °C, hold for 5 min, (2) increase to 250 °C at
248
10 °C/min, hold for 30 min. The split ratio was 1:10 (v/v). The injector and the detector
249
temperatures were both set at 250 °C. FAME components were identified and quantified by 10
250
comparing the retention times and peak areas with those of standard solutions.
251 252
3. Results and discussion
253
3.1. VFA production by alginate fermentation using mixed cultivation system
254
As mentioned in the materials and methods, the inoculum of alginate fermentation was the
255
effluent from a CSTR. In fact, the CSTR was a VFA-producing system which can
256
continuously produce acetate, propionate and butyrate using L. japonica as sole carbon
257
source without adding any other nutrients. This system has been operated for over 1000 d in
258
our previous research and stabilized more than 600 d. Therefore, the microbial community in
259
the effluent from the CSTR was quite stable.
260
To evaluate the profile changes of acids during alginate fermentation, mannitol
261
consumption along with the accumulation and consumption of acidogenic products in batch
262
reactor were demonstrated (Fig. 2). Although the mannitol and alginate were separated by
263
natural sedimentation, the liquid (mannitol solution) could not be completely separated from
264
the solid (alginate suspension). Therefore, liquid samples for HPLC analysis after
265
centrifugation retained some mannitol (5.8 g/L initially). The products of alginate
266
fermentation were acetate, succinate, lactate, formate, acetate, propionate, butyrate and
267
ethanol. Ethanol concentration reached 1.0 g/L within 24 h, but gradually decreased to 0 by
268
144 h, probably because ethanol initially produced was converted into acetate by
269
acidogenesis (Angenent et al., 2008). The depletion of ethanol and production of acetate are
270
thermodynamically favorable (Oh et al., 2003). Likewise, succinate, lactate and formate also
271
increased to a certain amount and disappeared later. During the period of acidogenesis,
272
acetate was produced as a major alginate fermentation product, followed by propionate and
273
butyrate. The final concentration of acetate was more than 9.0 g/L at 6.5 d. Propionate
274
concentration slowly increased to 3.5 g/L, and remained at that level until 156 h. Butyrate 11
275
was produced until 24 h and its concentration began to increase from 36 h, ultimately to a
276
concentration which was more than 0.5 g/L. At the end of digestion, the VFAs consisted of
277
69.2 % acetate, 25.7 % propionate and 5.1 % butyrate.
278 279
3.2. Cell growth and lipid accumulation by C.curvatus using carbon source derived from
280
macroalgae by single batch cultivation system.
281
Most of the microorganisms can synthesize lipids, but only the oleaginous strains may
282
accumulate more than 20 % W/W lipids on dry cell basis (Papanikolaou and Aggelis, 2011a).
283
Microorganisms synthesize lipids by either de novo processes that utilizes hydrophilic
284
substances as substrates, or by ex novo processes that utilize hydrophobic substances as
285
substrates. In this research, all lipid synthesis occurred by de novo processes. In de novo
286
processes, lipid accumulation always happens after depletion of nitrogen (or to a lesser extent
287
of other essential nutrient like phosphorus or sulfate) from the medium (Papanikolaou and
288
Aggelis, 2011b). In addition, to ensure that VFAs are mostly converted into lipids rather than
289
to biomass, these carbon sources should be added to a medium containing enough cells
290
(Fontanille et al., 2012).
291
Total nitrogen (TN) was 134 mg/L in the mannitol supernatant and 48 mg/L in the VFA
292
effluent. Initially 1 L of VFA effluent was added to the 2 L of mannitol substrate; this
293
addition diluted manntiol and VFA concentration in the substrate to 2/3 and 1/3 of pre-
294
addition levels, respectively. During the first 12 h, VFAs were not dramatically consumed by
295
C. curvatus. Lipid concentration and DCW increased slightly at the same time. However,
296
lipid content increased rapidly from less than10 % to 30 %. During the next 36 h, lipid
297
content, lipid concentration, and DCW increased to a much higher level until 48 h when all
298
VFAs were used up by yeasts. As for lipid content, it achieved its highest point (48.30%). It
299
is obvious that the concentration of mannitol was almost constant until 48 h, which could 12
300
demonstrate that in lipid-producing metabolic pathway of C. curvatus, VFAs were more
301
preferable than mannitol as a carbon sources under pH of 5.5. After the exhaustion of VFAs,
302
mannitol began to gradually decrease and was consumed up within 84 h. In the meantime,
303
DCW increased to 3.60 g/L until 72 h and then remained stable; whereas lipid content
304
suddenly reduced which resulted in the decrease of lipid concentration. This phenomenon
305
which is called as “lipid turnover” is related with storage lipid degradation (Chen et al., 2012,
306
Peng et al., 2013). Microbial lipid turnover has been extensively studied in Y. lipolytic and C.
307
echinulata (Papanikolaou and Aggelis, 2011a). Oleaginous yeasts consume their own
308
intracellular lipids to maintain lipid-free biomass when carbon source is exhausted or carbon
309
uptake rate decreases. Because there was still some mannitol left in the substrate, the lipid
310
turnover could result from the reduced carbon source uptake rate. When VFAs were
311
exhausted, C. curvatus had to consume mannitol as substitution, which could make the
312
carbon source uptake rate decreased during this process. From 72 h, lipid content increased
313
again with more mannitol assimilated by the yeasts.
314
Several research groups have used VFAs to produce lipids by culturing oleaginous yeasts
315
including Yarrowia lipolytica and Cryptococcus curvatus (Christophe et al., 2012, Fontanille
316
et al., 2012). The most significant difference was that in our study the microorganisms
317
utilized mannitol and VFAs which were all derived from seaweed without addition of any
318
other nutrients and reagents, whereas in other research, oleaginous yeasts used a synthetic
319
medium containing glucose and acetate as substrates. Moreover, in other research the ratio of
320
carbon to nitrogen (C/N) was artificially controlled by continuously adding carbon and
321
nitrogen sources to achieve high lipid concentration and content. But in our work, we just
322
utilized the original mannitol supernatant and alginate fermentation effluents to culture the
323
oleaginous yeasts.
324
We compared our results to other researches that also used VFAs or acetate as carbon 13
325
sources (Table 4). In fact, little research has examined the feasibility of using mannitol as a
326
carbon source for oleaginous yeasts. In addition, the strains and concentration of carbon
327
source used were all different with each research. Therefore, the only relevant comparisons
328
was of the value of lipid content (% W/W). Among these results, apparently the lipid content
329
in this work (48.30%) was higher than that of any other researches. It indicated that the
330
mannitol and VFAs derived from alginate anaerobic fermentation were favorable for
331
microbial lipid production even without using synthetic medium or artificially adding any
332
other nutrients.
333 334
3.3. Fatty acid composition analysis
335
The fatty acids produced in our study consisted mainly of palmitic acid (C16:0), stearic
336
acid (C18:0), oleic acid (C18:1), and linoleic acid (C18:2) (Table 5). Others indicates C8:0,
337
C10:0, C12:0, C14:0, C16:1, C18:3, C20:0, C20:1, C20:2, and C24:0. This composition is
338
similar to those of vegetable oils which have been utilized for industrial production of
339
biodiesel (Adamska ET, 2004). Over time, some changes in lipid composition occurred.
340
Obviously, there were three different trends shown in each fatty acid composition. As for
341
C18:0 and C18:1, they both increased during the period of VFAs consumption until 48 h,
342
while the percentages decreased when C. curvatus started to assimilate mannitol as the sole
343
carbon source. In contrast, in the case of palmitic acid, it was gradually reduced during the
344
first 48 h, then accumulated again until the end. However, the profile of linoleic acid
345
followed a totally different pattern. The percentage increased throughout the period from
346
8.95 % to 17.15 %, irrespective of which carbon source yeasts used. This composition was
347
comparable to vegetable fatty acids indicating that the fatty acids accumulated by C. curvatus
348
are appropriate for use in biodiesel production (Li et al., 2007, Meng et al., 2009).
349 14
350
4. Conclusions
351
We designed an efficient batch system for lipid production by the oleaginous yeast
352
Cryptococcus curvatus using two different carbon sources derived from a brown macroalga,
353
Laminaria japonica: mannitol and VFAs obtained by anaerobic fermentation of alginate with
354
a mixed culture. This is the first demonstration of microbial oil production by bioconversion
355
of macroalgae without addition of any other carbon sources or nutrients. This study has
356
shown that mannitol and VFAs derived from alginate fermentation are favorable feedstock
357
for lipid production. Future work can focus on optimizing and scaling up this process.
358 359
Acknowledgements
360
This research was supported by Basic Science Research Program through the National
361
Research Foundation of South Korea (NRF) funded by the Ministry of Education, Science
362
and Technology (Grant number 2011-0001108), the Advanced Biomass R&D Center (ABC)
363
of South Korea Grant funded by the Ministry of Education, Science and Technology (ABC-
364
2011-0028387), Marine Biotechnology Program Funded by Ministry of Land, Transport and
365
Maritime Affairs of South Korean Government, South Korea and the Manpower
366
Development Program for Marine Energy funded by Ministry of Land, Transportation and
367
Maritime Affairs (MLTM) of South Korean government and by the World Class University
368
(WCU) program through the National Research Foundation of South Korea funded by the
369
Ministry of Education, Science and Technology (R31-30005).
370 371
15
372
Reference
373 374 375 376 377 378 379 380 381 382 383 384 385 386 387 388 389 390 391 392 393 394 395 396 397 398 399 400 401 402 403 404 405 406 407 408 409 410 411 412 413 414 415 416 417 418 419
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sulphated homofucans from brown algae. Int. J. Biol. Macromol. 8, 380-386. [19]Kloareg, B., Quatrano, R., 1988. Structure of the cell walls of marine algae and ecophysiological functions of the matrix polysaccharides. Oceanogr. Mar. Biol. Annu. Rev. 26, 259-315. [20]Lee, M., Hidaka, T., Tsuno, H., 2008. Effect of temperature on performance and microbial diversity in hyperthermophilic digester system fed with kitchen garbage. Bioresour. Technol. 99, 6852-60. [21]Li, Y., Zhao, Z., Bai, F., 2007. High-density cultivation of oleaginous yeast Rhodosporidium toruloides Y4 in fed-batch culture. Enzyme Microb. Technol. 41, 312-317. [22]Li, Y.H., Liu, B., Zhao, Z.B., Bai, F.W., 2006. Optimization of Culture Conditions for Lipid Production by Rhodosporidium toruloides. Chin. J. Biotechnol. 22, 650-656. [23]Liu, Z.Y., Wang, G.C., Zhou, B.C., 2008. Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresour. Technol. 99, 4717-22. [24]McKendry, P., 2002. Energy production from biomass (part 1): overview of biomass. Bioresour. Technol. 83, 37-46. [25]Meng, X., Yang, J., Xu, X., Zhang, L., Nie, Q., Xian, M., 2009. Biodiesel production from oleaginous microorganisms. Renew. Energ 34, 1-5. [26]Oh, S.E., Van Ginkel, S., Logan, B.E., 2003. The relative effectiveness of pH control and heat treatment for enhancing biohydrogen gas production. Environ. Sci. Technol. 37, 5186-5190. [27]Papanikolaou, S., Aggelis, G., 2011a. Lipids of oleaginous yeasts. Part I: Biochemistry of single cell oil production. Eur. J. Lipid Sci. Technol. 113, 1031-1051. [28]Papanikolaou, S., Aggelis, G., 2011b. Lipids of oleaginous yeasts. Part II: Technology and potential applications. Eur. J. Lipid Sci. Technol. 113, 1052-1073. [29]Park, J.M. 2012. PETROFED (Petroleum Federation of India). Trends and Prospects of Biorefinery Research in Korea. Available from: . [30]Peng, W.F., Huang, C., Chen, X.F., Xiong, L., Chen, X.D., Chen, Y., Ma, L.L., 2013. Microbial conversion of wastewater from butanol fermentation to microbial oil by oleaginous yeast Trichosporon dermatis. Renew. Energ 55, 31-34. [31]Pyeun, J.H., Park, Y.H., Lee, K.H., 1977. Factors involved in the quality retention of cultured Undaria pinnatifida. Bull. Korean Fish. Soc 10, 125-135. [32]Ratledge, C., 1991. Microorganisms for lipids. Acta Biotechnol. 11, 429-438. [33]Ratledge, C., Cohen, Z., 2008. Microbial and algal oils: Do they have a future for biodiesel or as commodity oils. Lipid Technol. 20, 155-160. [34]Wang, J., Kim, Y.M., Rhee, H.S., Lee, M.W., Park, J.M., 2013. Bioethanol production from mannitol by a newly isolated bacterium, Enterobacter sp. JMP3. Bioresour. Technol. 135, 199-206.
459 460 461
17
462 463
Figure legends:
464
Fig. 1. Schematic diagram of alginate fermentation and lipid production processes.
465 466
Fig. 2. Profile of various carbon sources and volatile suspended solid (VSS) variations
467
during the alginate fermentation. (▲) VSS; (◇) mannitol; (○) acetic acid; (▼)
468
propionic acid; (□) butyric acid; (◆) ethanol; (ⅹ) formic acid; (▽) lactic acid; (△)
469
succinic acid.
470 471
Fig. 3. Profile of mannitol (◇), acetic acid (○), propionic acid (▼), butyric acid (□),
472
dry cell weight (DCW)(●), lipid concentration (▲), lipid content (■) during a two-step
473
batch cultivation of Cryptococcus curvatus in a 5 L fermentor.
18
st
nd
rd
Table 1. Comparisons between 1 , 2 , and 3 generation biomass Resourcea
Crop
Forest-derived
Seaweed
Harvesting
1 – 2 /yr
1 / 8yr
4 – 6 /yr
Production (ton/ha)
180
9
565
CO2 uptake (ton/ha)
5 – 10
4.6
36.7
Energy yield (%)
30 - 35
20 – 25
> 45
Cost ($/L)
0.2 – 0.3
0.4
0.2
a
(Park, 2012)
19
Table 2. Composition of Laminaria japonica and Undaria pinnadifida
(% w/w, dry base)
Protein
Lipid
Ash
Carbohydrate
Laminaria japonica
6.8 – 10.3
7.2 – 11.5
13.8 – 21.1
60.9 – 67.0
Undaria pinnadifidaa
13.7 – 28.7
3.6 – 6.2
29.4 – 46.5
26.5 – 42.8
a
(Cho, 1995)
20
Table 3. Characteristic algal polysaccharides in Laminaria japonica and Undaria pinnadifida Algal carbohydrate (% w/w, dry base) Alginate
Fucoidan
Laminaran
Mannitol
Total
Laminaria japonica
14.6 – 29.5
0 – 0.1
3.7 – 4.2
22.5 – 33.2
40.8 – 67.0
Undaria pinnadifidaa
22.3 – 32.9
1.4
3.6
4.1 – 7.3
31.4 – 45.2
a
(Kim et al., 1995, Pyeun et al., 1977)
21
Table 4. Lipid production by oleaginous yeasts using different substrate in two-step system Strains
Carbon source (g/L)
DCW (g/L)
Lipid conc. (g/L)
Lipid content (% W/W)
Reference
C. albidus
VFAs (2.00)
1.16
0.31
27.00
(Fei et al., 2011a)
C. albidus
Acetate (6.00)
2.85
0.74
25.80
(Fei et al., 2011b)
Y. lipolytica
Acetate (12.00)
5.98
1.84
30.76
(Fontanille et al., 2012)
C. curvatus
Mannitol + VFAs (9.30)
3.60
1.30
48.30
This study
a
Calculated based on references.
22
Table 5. Fatty acid methyl ester (FAME) profile of C. curvatus Fatty acid methyl ester (FAME) profile of C. curvatus (%) Fermentation time (h)
Palmitic AME (C16:0)
Stearic AME (C18:0)
Oleic AME (C18:1)
Linoleic AME (C18:2)
Others
0
25.88
12.83
48.16
8.95
4.18
12
25.62
11.64
48.36
10.65
3.73
24
21.16
15.45
49.38
11.41
2.60
36
14.72
22.41
50.34
11.49
1.05
48
14.70
19.46
52.26
12.78
0.80
60
15.33
18.20
52.13
13.29
1.06
72
16.75
15.76
49.31
16.32
1.86
84
18.18
14.03
48.71
17.15
1.93
a
Others: C8:0, C10:0, C12:0, C14:0, C16:1, C18:3, C20:0, C20:1, C20:2, and C24:0.
23
a
Aerator
Flow Meter
pH Controller
Mannitol Supernatant
Lipid Fermentor
VFAs
Substrate (30 g/L Laminaria japonica)
pH Controller Alginate Slurry
Alginate Fermentor
Fig. 1. Schematic diagram of alginate fermentation and lipid production processes. 24
12
40
30 8
6
20
VSS (g/L)
Concentration (g/L)
10
4 10 2
0
0 0
24
48
72
96
120
144
Time (h) Fig. 2. Profile of various carbon sources and volatile suspended solid (VSS) variations during alginate fermentation. (▲) VSS; (◇) mannitol; (○) acetic acid; (▼) propionic acid; (□) butyric acid; (◆) ethanol; (ⅹ) formic acid; (▽) lactic acid; (△) succinic acid.
25
2.0
50
40 1.5
6 1.0 4
30
20
Lipid content (%)
8
Lipid concn. (g/L)
DCW, Mannitol and VFAs (g/L)
10
0.5 2
10
0
0.0 0
12
24
36
48
60
72
0
84
Time (h) Fig. 3. Profile of mannitol (◇), acetic acid (○), propionic acid (▼), butyric acid (□), dry cell weight (DCW)(●), lipid concentration (▲), lipid content (■) during a batch cultivation of Cryptococcus curvatus in a 5 L fermentor.
26
H I G H LI G H TS ▶This
is the first demonstration of microbial oil production by using macroalgae.
▶Oleaginous ▶Carbon
sources were all derived from Laminaria japonica.
▶No addition ▶The
yeast Cryptococcus curvatus was used for microbial oil production.
of any other nutrients or synthetic medium was used.
composition of the fatty acids was found similar to vegetable oils.
27
