yield coefficient based on glucose, the perfusion culture system potentially may be used to grow algae for heterotrophic production of bio-oils.
A modified perfusion culture system that introduces cell bleeding during perfusion oper- ation was also developed for heterotrophic production of algae (Figure 6.5c;Wen and Chen, 2001b). This system could potentially improve the biomass productivity but at the same time lower the cell density, e.g., from 40 g L–1to less than 20 g L–1(Wen and Chen, 2001b; Wen and Chen, 2002a).
It is worth mentioning that different algal species/strains may favor different culture systems to achieve maximized cell density, biomass productivity, and oil productivity.
An experimental optimization is required for a selected algal strain to demonstrate which culture system is best for the heterotrophic production of oils. Regardless of the algal strain selected and culture system used, the key to optimizing a production system rests with the cost balance of output and input from a cost-effectiveness point of view.
6.6 CHLORELLA AS THE CELL FACTORY
the data obtained from thoseChlorellaspecies that may have been excluded from theChlorella genus by the studies mentioned.
6.6.1 Oil Production Potential
Chlorella has long been used as human health food. Under certain stress conditions, Chlorellaspecies are capable of accumulating as high as 60% (w/w, on dry-weight basis) oil within cells (Table 6.4). Together with the characteristics of high growth rate and ease of culture and scale-up in bioreactors, Chlorella has attracted unprecedented interest as a feedstock for biofuels, in particular biodiesel (Xu et al 2006; Li et al 2007a; Xiong et al 2008; Hsieh and Wu 2009; Gao et al 2010; Liu et al 2010, 2012a). The synthesized fatty acids inChlorellaare mainly of medium length, ranging from 16 to 18 carbons, despite the great variation in fatty acid composition (Table 6.5). Generally, saturated fatty esters possess high cetane numbers and superior oxidative stability, whereas unsaturated, especially poly- unsaturated, fatty esters have improved low-temperature properties (Knothe, 2008). It is suggested that the modification of fatty esters—for example, enhancing the proportion of oleic acid (C18:1) ester—can provide a compromise solution between oxidative stability and low-temperature properties and therefore promote the quality of biodiesel (Knothe, 2009). In this regard, C. protothecoides, with the highest proportion of oleic acid (71.6%), may be better than other Chlorellaspecies as biodiesel feedstock (Cheng et al., 2009). The properties ofC.protothecoides-derived biodiesel were assessed, and most of them proved to comply with the limits established by the American Society for Testing and Materials (Xu et al., 2006).
There are increasing reports of using heterotrophicC.protothecoidescultures for oil produc- tion, from laboratory scale to large scale of 11,000-L of culture volume (Table 6.4). The scale-up from 5 to 11,000 L just caused a slight decrease in productivities, suggesting theC.protothecoides may represent a potential producer of oils for commercially large-scale production (Li et al., 2007a). In a nonoptimized fed-batch culture ofC.protothecoides, the record cell density, biomass productivity, and oil productivity were achieved by Yan et al (2011), namely, 97.1 g L–1, 12.8 g L–1day–1, and 7.3 g L–1day–1, respectively. Later, using a nonlinear-mode-based optimi- zation approach,De la Hoz Siegler et al. (2012)maximized the cell density and oil productivity of fed-batch culture ofC.protothecoidesto 144 g L–1and 20.2 g L–1day–1.
6.6.2 Downstream Processes
Downstream processes ofC.protothecoidescultures include biomass harvest and drying, cell disruption, oil extraction, and transesterification for biodiesel. Various harvesting methods are applied toChlorellacultures, including flocculation, flotation, filtration, gravity sedimen- tation, and centrifugation (Lin, 2005; Wiley et al., 2009; Papazi et al., 2010; Lee et al., 2012).
The harvest efficiency rests not only with harvesting methods used but also algal species, cul- ture ages, and cell densities. Usually, a harvesting method is not used alone but is coupled with one or more other methods to achieve the highest harvesting efficiency, e.g., a preceding treatment of flocculation was used to improve the performance of flotation, filtration, sedi- mentation, or centrifugation (Sim et al., 1988; Liu et al., 1999; Wiley et al., 2009). A drying
130 6. HETEROTROPHIC PRODUCTION OF ALGAL OILS
process following biomass harvest may be needed, depending on whether drying or wet bio- mass is used for oil extraction. The harvest and drying processes may contribute 20–30% of the total cost of photoautotrophic algal biomass production (Molina Grima et al., 2003). Although the high cell density associated with heterotrophic algae can reduce the cost contribution, TABLE 6.4 Growth and Lipid Production ofC.protothecoidesFeeding on Various Organic Carbon Sources.
Cell Density (g L1)
Biomass Productivity (g L1day1)
Lipid Productivity
(g L1day1) Organic Carbons Culture
Conditionsa References
16.5 3.6 1.60 Hydrolysate of
Jerusalem artichoke tuber
B, flask, 1 L Cheng et al., 2009
10.8 1.7 0.95 Glucose B, flask, 1 L De la Hoz Siegler
et al., 2011
30 3.3 1.9 Glucose FB, bioreactor, 2 L
— — 12.3b Glucose C, bioreactor, 2 L
144 — 20.2 Glucose FB, bioreactor, 2 L De la Hoz Siegler
et al., 2012
6 1.2 0.59 Hydrolysate of
sweet sorghum juice
B, flask, 500 mL Gao et al., 2010
15.5 2.0 0.93 Glucose FB, bioreactor, 5 L Li et al., 2007a
12.8 1.7 0.81 Glucose FB, bioreactor,
750 L
14.2 1.7 0.73 Glucose FB, bioreactor,
11,000 L
14 3.2 1.85 Glycerol B, flask O’Grady et al., 2011
13.1 1.46 0.85 Glucose B, flask, 250 mL Shen et al., 2010
14.2 2.2 1.2 Glucose B, bioreactor, 5 L Xiong et al., 2010b
51.2 6.6 3.3 Glucose FB, bioreactor, 5 L Xiong et al., 2008
15.5 2.0 1.1 Glucose FB, bioreactor, 5 L Xu et al., 2006
3.7 0.7 0.36 Corn powder
hydrolysate
B, flask, 500 mL
17.9 3.6 1.45 Hydrolyzed
molasses
B, flask, 500 mL Yan et al., 2011
97.1 12.8 7.3 Hydrolyzed
molasses
FB, bioreactor, 5 L
46 6.28 2.06 Glucose FB, bioreactor, 7 L Chen and Walker,
2012
aB, batch; FB, fed-batch; C, continuous.
bPredicted value.
131
6.6 CHLORELLAAS THE CELL FACTORY FOR HETEROTROPHIC OILS
TABLE 6.5 Fatty Acid Profiles of SelectedChlorellaSpecies.
Chlorella
Species C14:0 C15:0 C16:0 C16:1 C16:2 C16:3 C17:0 C18:0 C18:1 C18:2 C18:3 C20 or
Above References
C. ellipsoidea 2 26 4 40 23 5 Abou-Shanab et al.,
2011
C. minutissima 2.8 13.5 1.1 3.4 46.1 26.7 3.3 Li et al., 2011
C.protothecoides 14.3 1 2.7 71.6 9.7 Cheng et al., 2009
C.protothecoides 1.1 11.7 0.3 0.4 5.6 59.4 19.1 2.1 0.5 Chen and Walker,
2012
C.protothecoides 2.3 26.2 0.8 17.6 47.6 0.8 0.1 4.5 De la Hoz Siegler
et al., 2012
C. pyrenoidosa 17.3 7 9.3 1.2 3.3 18.5 41.8 D’oca et al., 2011
C. saccharophila 2.7 17.6 4.9 32.2 31.1 9.8 Isleten-Hosoglu
et al., 2012
C. sorokiniana 25.4 3.1 10.7 4.1 1.4 12.4 34.4 7.1 Chen and Johns,
1991
C. sp 19.1 1 3.1 25.9 6.8 44.2 Matsumoto et al.,
2010
C. sp 20.6 6.6 10.4 6 3.4 2.4 12.5 27.2 10.2 Wang et al., 2010
C. sp 3.3 6.4 49.5 10.1 28.5 1.3 Yeesang and
Cheirsilp, 2011
C. vulgaris 19.2 4.2 14.6 12.7 3.8 21.1 13.8 Cleber Bertoldi
et al., 2006
C. vulgaris 63 9 3 11 13 Converti et al., 2009
C. vulgaris 24 2.1 1.3 24.8 47.8 Yoo et al., 2010
C. vulgaris 1 32 26 1 5 14 28 3 Heredia-Arroyo
et al., 2011
C. zofingiensis 22.6 2 7.4 2 2.1 35.7 18.5 7.8 Liu et al., 2010
C. zofingiensis 22.8 2.5 7.5 1.8 2.7 34.2 19.7 7.3 Liu et al., 2012a
1326.HETEROTROPHICPRODUCTIONOFALGALOILS
finding ways to improve the cost-effectiveness of the harvest and drying steps still represents a big challenge for theChlorellaindustry if it is to expand from the current high-value, low- quantity specialty products market to a low-value, high-volume commodity products market.
Chlorellahas a tough, rigid cell wall, and thus disruption of the cell wall is required for the facilitation of oil extraction. Various disruption methods, e.g., mechanical crushing, ultra- sonic treatment, and enzymatic degradation, can be employed for cell-wall disruption. The oils released after cell disruption are suitable for extraction using organic solvents. Supercrit- ical CO2is another way for efficiently extracting algal oils, but it is expensive and energy in- tensive, which restricts the commercialization of this technology (Herrero et al., 2010).
The extracted algal oils are suitable for biodiesel conversion through transesterification.
Transesterificationis a catalytic reaction of oils with a short-chain alcohol (typically methanol or ethanol) to form fatty acid esters. The reaction is reversible; as such, a large excess of alcohol is used in industrial processes to ensure the direction of fatty acid esters. Methanol is the pre- ferred alcohol for industrial use because of its low cost. Commonly, a catalyst is required to facilitate the transesterification, including acids, alkalis, and enzymes. Acid transesteri- fication is considered suitable for the conversion of oils with high free fatty acids but with low reaction rate (Gerpen, 2005). In contrast, alkali catalyzes a much higher transesterification rate, thought it is unfavorable for free fatty acids (Fukuda et al., 2001). As a result, alkalis are preferred catalysts for industrial production of biodiesel, and acid pretreatment is usually employed when the oils contain a high content of free fatty acids. The use of lipases for transesterification has also attracted much attention because it produces a high-purity prod- uct and enables easy separation of biodiesel from the byproduct glycerol (Ranganathan et al., 2008). However, the cost of the enzyme is still relatively high and remains a barrier to its industrial implementation.