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Production potential of Chlorella zofingienesis as a feedstock for biodiesel
Bioresource Technology 101 (2010) 8658–8663
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Production potential of Chlorella zofingienesis as a feedstock for biodiesel Jin Liu a, Junchao Huang a,*, King Wai Fan a, Yue Jiang b,**, Yujuan Zhong a, Zheng Sun a, Feng Chen a a b
School of Biological Science, The University of Hong Kong, Hong Kong, China Department of Biology, Hong Kong Baptist University, Kowloon Tong, Hong Kong, China
a r t i c l e
i n f o
Article history: Received 12 January 2010 Received in revised form 20 May 2010 Accepted 25 May 2010 Available online 7 July 2010 Keywords: Biodiesel Chlorella zofingiensis Fed-batch fermentation Lipid Microalgae
a b s t r a c t The lipid contents and fatty acid profiles of Chlorella zofingiensis cultured in the dark with various carbon sources were investigated. Glucose was found to be the best carbon source for the growth and lipid production. When cultivated with 50 g L1 glucose, C. zofingiensis accumulated lipids up to 52% of the dry biomass, with triacylglycerols (TAGs) accounting for 72.1% of the total lipids. Fatty acid profiles revealed that glucose contributed to the highest yield of total fatty acids (TFAs) and proportion of oleic acid (35.7% of TFAs), which corresponded to the strongest up-regulation of biotin carboxylase (BC) and stearoyl ACP desaturase (SAD) genes. In fed-batch cultivation based on glucose, the lipid yield and productivity of C. zofingiensis were further increased to 20.7 g L1 and 1.38 g d1 L1 respectively, representing 3.9-fold of those achieved in batch culture. We conclude that C. zofingiensis has great potential for biodiesel production. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction Petroleum-based fuels are recognized as unsustainable energy source due to their depleting supplies and contribution to global warming (Chisti, 2008). Renewable biofuels are promising alternatives to petroleum-based fuels, among which biodiesel has attracted the most attention in recent years (Knothe et al., 1997; Hu et al., 2008; Huang et al., 2010). Biodiesel is a diesel-equivalent fuel derived from biological feedstocks and is chemically referred to as a fatty acid methyl ester (FAME). Compared with traditional fuels, biodiesel is carbon neutral, contributes less emission of gaseous pollutants and hence is environmentally beneficial (Ma and Hanna, 1999). Currently biodiesel is mainly derived from vegetable oils, animal fats and waste cooking oils (Knothe et al., 1997; Lang et al., 2001; Zhang et al., 2003). However, those sources cannot even meet the growing need for transportation energy as large land areas have to be used for oil crops cultivation (Chisti, 2008). In addition, the relatively low productivity by oil plants also restricts the commercialization of biodiesel from plants (Lang et al., 2001). There is the need therefore, to search for a cost-effective source of biodiesel to fulfil industrial sustainability. Microalgae are currently being considered as feedstocks for biodiesel production because of their rapid growth rates and high yield of lipids (Chisti, 2008; Hu et al., 2008; Huang et al., 2009). Furthermore, unlike oil crops, microalgae can be easily cultured in outdoor ponds or indoor well-established bioreactors or fermentors (Chen,
* Corresponding author. Tel.: +852 22990309; fax: +852 22990311. ** Corresponding author. Tel.: +852 34117062; fax: +852 34115995. E-mail addresses: [email protected] (J. Huang), [email protected] (Y. Jiang). 0960-8524/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2010.05.082
1996; Del Campo et al., 2007; Pruvost et al., 2009). Thus microalgae are superior to oil crops for the sustainable production of biomass in an environmental-friendly operation mode. When a microalga is considered for mass biodiesel production, two key factors, namely cell biomass and lipid content are essential for the initial assessment. The green microalgae in the genus Chlorella consist of about 10 species that can grow photoautotrophically, mixotrophically and heterotrophically with high biomass concentration (Chen, 1996; Shi and Chen, 2002; Ip et al., 2004; Miao and Wu, 2004). Considering their lipid contents, C. vulgaris and C. protothecoides have been reported to be the candidates for biodiesel production under photoautotrophic or heterotrophic culture conditions (Miao and Wu, 2004; Liu et al., 2008; Hsieh and Wu, 2009). Certain Chlorella species was reported to attain very high biomass concentration in heterotrophic culture (Sun et al., 2008). In addition, lipids extracted from heterotrophically grown cells had similar properties to fuel oil in terms of oxygen content, heating value, density and viscosity (Miao and Wu, 2004). Moreover, cultivating Chlorella heterotrophically could eliminate the light requirement that could definitely reduce the cost of the final product (Chen, 1996). We reported previously that C. zofingiensis could grow rapidly in the dark with glucose as the sole carbon and energy source (Ip and Chen, 2005). Furthermore, by using fed-batch culture strategy, up to 53 g L1 biomass could be reached for this alga (Sun et al., 2008). In this study, we investigated the production potential of C. zofingiensis as a biodiesel feedstock, focusing on its growth and lipid contents under heterotrophic culture conditions with various organic carbon sources. The regulation of two key genes involved in fatty acid biosynthesis was also surveyed in order to deepen our fundamental understanding of biodiesel formation.
J. Liu et al. / Bioresource Technology 101 (2010) 8658–8663
2. Methods 2.1. Microalga strain C. zofingiensis (ATCC 30412) was obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). This alga was maintained at 4 °C on an agar slant of Kuhl medium (Kuhl and Lorenzen, 1964) consisting of (per liter) 1.01 g KNO3; 0.62 g NaH2PO4H2O; 0.089 g Na2HPO42H2O; 0.247 g MgSO47H2O; 14.7 mg CaCl22H2O; 6.95 mg FeSO47H2O; 0.061 mg H3BO3; 0.169 mg MnSO4H2O; 0.287 mg ZnSO47H2O; 0.0025 mg CuSO45H2O; and 0.01235 mg (NH4)6MO7O244H2O. The pH of the medium was adjusted to pH 6.5 prior to autoclaving. 2.2. Batch and fed-batch culture Inocula were prepared by growing the microalga in 250 mL Erlenmeyer flask containing 50 ml of the Kuhl medium at 25°C for 4 days with orbital shaking at 150 rpm and illuminated with continuous light of 30 lmol photon m2 s1. For batch culture, Erlenmeyer flasks, each containing 90 mL medium supplemented with various carbon sources (lactose, galactose, sucrose, fructose, mannose and glucose), were inoculated with 10% (v/v) of exponentially growing inoculum and then incubated at 25 °C in an orbital shaker at 150 rpm in the dark. The two-stage fed-batch strategy previously developed by us for high-biomass of C. zofingiensis (Sun et al., 2008) was used to investigate if the well-controlled fermentation system is superior to the classical batch culture system. The working volume of the fermentor (3.7-L, Bioengineering Ag, Wald, Switzerland) was 3.0 L. The cultivation conditions in the fermenter were controlled as follows: pH 6.5; temperature 25 °C; agitation 450 rpm; and dissolved oxygen concentration at 50% saturation. The pH value of medium was regulated by phosphate buffer which maintained the value near 6.5. During fed-batch cultivation, the sterilized stock nutrient solution was fed into the fermenter to maintain the glucose concentration at 5–20 g L1. 2.3. Determination of glucose concentration, nitrate concentration, dry cell weight and specific growth rate The cells were centrifuged at 3800g for 5 min. Glucose and nitrate concentrations in the supernatant were determined according to Miller (1959) and Elton-Bott (1979), respectively. The pellet was re-suspended in distilled water and filtered through a pre-dried Whatman GF/C filter paper (1.2 lm pore size). The algal cells on the filter paper discs were dried at 70 °C in a vacuum oven until constant weight and were cooled down to room temperature in a desiccator before weighting. The specific growth rate (l) at the exponential phase was calculated according to the equation, l = (ln X2 ln X1)/(t2 t1), where X2 and X1 are the dry cell weight concentration (g L1) at time t2 and t1, respectively.
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(GLs), and phospholipids (PLs) on silica cartridges (Waters, Milford, MA, USA) by sequential elution with chloroform, acetone, and methanol, as previously described (Christie, 2003). Neutral lipids were further resolved to subclasses by TLC (silica gel 60, 20 20 cm plates, 0.25 mm thickness; Merck, Whitehouse Station, NJ, USA) using a solvent system of petroleum ether:diethyl ether:acetic acid (70:30:1, v/v/v). Lipids were visualized by brief exposure to 2,7-dichlorofluorescein (Sigma, St. Louis, MO, USA) vapors and were identified by comparison with the standards (Sigma). Fatty acid methyl esters (FAMEs) were prepared by direct transmethylation of freeze-dried cells, lipid extracts, or individual lipids with sulphuric acid in methanol (Christie, 2003). The FAMEs were analyzed by using a HP 6890 capillary gas chromatograph (Hewlett–Packard, Palo Alto, CA) equipped with a flame ionization detector (FID) and a HP-INNOwax capillary column (30 m 0.32 mm) (Agilent Technologies, Inc., Wilmington, DE). Nitrogen was used as carrier gas. Initial column temperature was set at 170 °C, which was subsequently raised to 230 °C at 1 °C min1. The injector was kept as 250 °C with an injection volume of 2 lL under splitless mode. The FID temperature was set at 270 °C. FAMEs were identified by chromatographic comparison with authentic standards (Sigma). The quantities of individual FAMEs were estimated from the peak areas on the chromatogram using heptadecanoic acid as the internal standard. 2.5. RNA isolation and RT-PCR assay RNA was isolated from aliquots of about 108 cells using the TRI Reagent (Molecular Research Center, Cincinnati, OH, USA) according to the manufacturer’s instructions. The concentration of total RNA was determined spectrophotometrically at 260 nm. Total RNA (1 lg) extracted from different samples was reverse transcribed to cDNA by using a SuperScript III First-Strand Synthesis System (Invitrogen, Carlsbad, CA, USA) for reverse transcriptionPCR (RT-PCR) primed with oligo(dT) according to the manufacturer’s instructions. PCR amplification was carried out according to Li et al. (2008a) using specific primers of BC (forward, 50 GTGCGATTGGGTATGTGGGGGTG-30 , and reverse, 50 -CGACCAGGAC CAGGGCGGAAAT-30 ) and SAD (forward, 50 -TCCAGGAACGTGCCAC CAAG-30 , and reverse, 50 -GCGCCCTGTCTTGCCCTCATG-30 ). C. zofingiensis actin (ACT) primers (forward, 50 -TGCCGAGCGTGAAATTGT GAG-30 , and reverse, 50 -CGTGAATGCCAGCAGCCTCCA-30 ) were used to demonstrate equal amounts of templates and loading. The GenBank accession numbers for BC and SAD were GQ996717 and GQ996719, respectively. Amplification of the cDNA was done by conventional PCR [94 °C for 2 min followed by 24 cycles (for ACT gene) or 26 cycles (for BC and SAD genes) of 94 °C for 15 s, 58 °C for 20 s, 72 °C for 30 s]. PCR products were separated on a 2% agarose gel and stained with ethidium bromide for photography (Biorad, Hercules, CA, USA). 3. Results and discussion
2.4. Lipid extraction and fractionation and fatty acid analysis Cells were harvested and lyophilized for lipid extraction and fatty acids analysis. Lipids were extracted with chloroform–methanol (2:1, v/v), and then separated into chloroform and aqueous methanol layers by addition of methanol and water to give a final solvent ratio of chloroform:methanol:water of 1:1:0.9. The chloroform layer was washed with 20 mL of a 5% NaCl solution, and evaporated to dryness. Thereafter, the total lipids were measured gravimetrically and stored at 20 °C under nitrogen gas to prevent lipid oxidation or used directly for subsequent analysis. Total lipid extracts were fractionated into neutral lipids (NLs), glycolipids
3.1. Heterotrophic growth and lipid production of C. zofingiensis with various carbon sources We have previously reported that C. zofingiensis could grow well in the dark with various sugars as carbon sources (Sun et al. 2008). To investigate the influences of carbon sources on lipid production in C. zofingiensis cells, we cultured the algal cells in the dark with lactose, galactose, sucrose, fructose, mannose or glucose as a carbon source, respectively. Among the tested sugars, glucose gave the highest growth rate (0.03 h1), cell biomass (10.1 g L1), lipid content (0.52 g g1), and lipid yield (5.27 g L1). Algal cells cultured
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with mannose, fructose, or sucrose produced slightly lower amounts of cell biomass and lipids. In contrast, lactose and galactose were observed to be poor carbon sources for lipid production since low biomass and lipid contents were obtained (Fig. 1). These findings demonstrated a close relationship between lipid biosynthesis and cell growth for the heterotrophic alga. As shown in Fig. 2, based on lipid class analysis of C. zofingiensis cultured with glucose, neutral lipids (NLs) were found to be the major constituent that accounted for 80.9% of the total lipids, while phospholipids (PLs) and glycolipids (GLs) together accounted for 19.1%. In NLs, TAGs were the predominant component, accounting for 72.1% of the total lipids. In the batch culture, up to 0.375 g g1 TAGs were accumulated in heterotrophic C. zofingiensis cells. Unlike PLs and GLs that are membrane-bounded lipids, TAGs serve primarily as a storage form of carbon and energy (Hu et al., 2008). In addition, TAGs are superior to phospholipids (PLs) or glycolipids (GLs) for biodiesel due to their higher content of fatty acids (Pruvost et al., 2009). Hence C. zofingiensis may act as a promising host for biodiesel production due to its fast cell growth and high content of TAGs.
3.2. Fatty acid profiles of dark-grown C. zofingiensis cultures Fatty acid composition considerably influences the properties of biodiesel such as cetane number, heat of combustion, oxidative stability, cloud point, lubricity, which will finally influence the quality of biodiesel (Knothe, 2009). In this study, the fatty acid compositions of the heterotrophically cultured C. zofingiensis with various sugars were investigated (Table 1). It was found that C16:0, C16:2, C18:1, C18:2 and C18:3 (n-3) were the major fatty acids, which accounted for more than 84% of the total fatty acids (TFAs). The highest amounts of TFAs (45.4% of cell biomass) and C18:1 (35.7% of TFAs) were achieved in C. zofingiensis cultured with glucose as the carbon source, which were about five and three times, respectively of those given by lactose. Furthermore, the fatty acid profiles of NLs, GLs and PLs from algal cells cultivated with glucose were also investigated. Compared with GLs and PLs, NLs had a lower proportion of C16:0 but a much higher percentage of oleic acid (Table 2). As ideal biodiesel, the fatty acids should be oxidative and low-temperature stable. Generally, saturated fatty acids are oxidative stable, while unsaturated fatty acids give low-temperature stability (Knothe, 2008). The enhanced proportion of oleic acid (C18:1) in the total fatty acids has been considered as a feasible ap-
Fig. 2. Distribution of NLs, GLs and PLs in total lipids extracted from heterotrophic C. zofingiensis cells in batch culture with 50 g L1 of glucose for 14 days. The horizontal line inside the NLs bar marks the portion of TAGs in this fraction.
proach to balance the oxidative and low-temperature stability with retaining the cetane number at an acceptable level (Knothe, 2009). The high content of oleic acid in the heterotrophic cells further supports that C. zofingiensis is a favorable host for producing high quality biodiesel. 3.3. Sugars up-regulate the transcription of BC and SAD genes of C. zofingiensis The de novo biosynthesis of fatty acid initially catalyzed in chloroplast by the key enzyme acetyl-CoA carboxylase (ACCase); while the introduction of the first double bond to acyl chain is carried out by stearoyl ACP desaturae (SAD), an enzyme playing an important role in determining the ratio of unsaturated and saturated fatty acids (Hu et al., 2008). To reveal the relationship between the fatty acid profiles and the regulation of ACCase and SAD, we investigated the transcript levels of SAD and biotin carboxylase (BC, a subunit of ACCase) in dark-grown C. zofingiensis cells cultured with various sugars using the RT-PCR approach. The transcription of the two genes was shown to be differentially regulated by various carbon sources (Fig. 3). In coincidence with the higher contents of TFAs in cells cultured with sucrose, fructose, mannose or glucose, the transcript levels of BC in the cells were strongly up-regulated. In contrast, galactose moderately enhanced the transcription of BC whereas lactose, the poor carbon source for the cell growth and fatty acid accumulation, caused a slight up-regulation of BC. These results are consistent with the findings of Ohlrogge and Jaworski (1997) that the first reaction step of fatty acid biosynthesis catalyzed by ACCase is a major point of flux control for this pathway. A similar pattern induced by sugars was also found for SAD at mRNA levels, although much stronger up-regulation was observed when compared with the expression of BC (Fig. 3). Similarly, lactose induced low-level expression of SAD while glucose triggered high amounts of SAD transcripts, which is consistent with the unsaturation values of fatty acids in the algal cells cultured with the tested sugars (Table 1). 3.4. Fed-batch fermentation enhances lipid production by C. zofingiensis
Fig. 1. Cell biomass (black column), lipid content (gray column) and yield (white column) of C. zofingiensis in batch culture with 50 g L1 of various sugars. Lac, lactose; Gal, galactose; Suc, sucrose; Fru, fructose; Man, mannose; Glu, glucose.
As shown from the above results, the productivity of lipid is closely related to the growth rate, cell density and cellular lipid content. Compared with batch culture, fed-batch cultivation mode can extend the exponential growth phase of the cell and further
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J. Liu et al. / Bioresource Technology 101 (2010) 8658–8663 Table 1 Fatty acid profiles of dark-grown C. zofingiensis cultured with 50 g L1 of various sugars for 14 days. Fatty acids
Sugars
C16:0 C16:1 C16:2 C16:3 C16:4 C18:0 C18:1 C18:2 C18:3 (n-6) C18:3 (n-3) C18:4 Others Mounsatda Pounsatdb Unsatdc DUS (r/mol)d TFAe a b c d e
Lactose
Galactose
Sucrose
Fructose
Mannose
Glucose
28.73 ± 1.22 3.42 ± 0.12 9.02 ± 0.35 5.28 ± 0.27 0.92 ± 0.05 0.95 ± 0.06 12.59 ± 0.45 20.87 ± 0.87 1.39 ± 0.03 12.86 ± 0.39 1.24 ± 0.04 2.72 ± 0.09 16.01 ± 0.55 51.59 ± 2.11 67.59 ± 2.95 1.43 ± 0.05 8.48 ± 0.33
27.81 ± 0.99 2.52 ± 0.13 11.08 ± 0.61 2.52 ± 0.11 0.65 ± 0.03 1.61 ± 0.09 25.15 ± 0.89 16.48 ± 0.83 1.17 ± 0.05 7.74 ± 0.31 1.10 ± 0.06 2.17 ± 0.11 27.67 ± 0.82 40.74 ± 2.05 68.42 ± 2.31 1.24 ± 0.06 20.30 ± 0.71
23.16 ± 0.83 1.49 ± 0.06 8.48 ± 0.39 1.96 ± 0.07 0.23 ± 0.02 2.80 ± 0.09 31.99 ± 1.21 19.51 ± 0.75 0.55 ± 0.01 7.36 ± 0.35 0.46 ± 0.01 2.00 ± 0.07 33.48 ± 1.12 38.57 ± 1.28 72.05 ± 3.72 1.22 ± 0.06 40.48 ± 1.29
23.62 ± 1.12 1.71 ± 0.08 8.31 ± 0.44 2.12 ± 0.09 0.17 ± 0.01 2.34 ± 0.12 32.06 ± 0.93 19.27 ± 0.49 0.52 ± 0.03 7.48 ± 0.18 0.45 ± 0.01 1.94 ± 0.04 33.78 ± 1.03 38.33 ± 1.49 72.10 ± 3.11 1.22 ± 0.03 43.01 ± 1.11
23.38 ± 1.03 1.49 ± 0.05 7.94 ± 0.29 1.83 ± 0.06 0.19 ± 0.01 2.44 ± 0.10 33.30 ± 1.45 19.36 ± 0.98 0.52 ± 0.03 7.31 ± 0.23 0.45 ± 0.02 1.78 ± 0.08 34.79 ± 1.37 37.61 ± 1.44 72.40 ± 3.13 1.21 ± 0.03 43.10 ± 1.79
22.62 ± 0.77 1.97 ± 0.08 7.38 ± 0.33 1.94 ± 0.04 0.22 ± 0.02 2.09 ± 0.08 35.68 ± 1.23 18.46 ± 0.66 0.51 ± 0.02 7.24 ± 0.30 0.49 ± 0.02 1.40 ± 0.08 37.64 ± 1.26 36.24 ± 1.63 73.89 ± 2.99 1.21 ± 0.05 45.38 ± 1.83
Mounsatd: percentage of monounsaturated fatty acids (% of total fatty acids). Pounsatd: percentage of polyunsaturated fatty acids (% of total fatty acids). Unsatd: percentage of unsaturated fatty acids (% of total fatty acids). DUS (r/mol): degree of fatty acid unsaturation = [1.0 (% monoenes) + 2.0 (% dienes) + 3.0 (% trienes) + 4.0 (% tetraenes)]/100. TFA: total fatty acids (g)/cell dry weight (g) 100%.
Table 2 Fatty acid profiles of neutral lipids, glycolipids and phospholipids from heterotrophic C. zofingiensis cells cultivated with 50 g L1 glucose for 14 days. Fatty acids
NLs
GLs
PLs
C16:0 C16:1 C16:2 C16:3 C16:4 C18:0 C18:1 C18:2 C18:3 (n-6) C18:3 (n-3) C18:4 Others Mounsatd Pounsatd Unsatd DUS (r/mol)
20.10 ± 0.83 1.65 ± 0.06 6.38 ± 0.35 1.80 ± 0.07 0.25 ± 0.02 2.76 ± 0.11 38.41 ± 2.15 19.20 ± 1.1 0.45 ± 0.02 6.86 ± 0.28 0.42 ± 0.02 1.71 ± 0.05 40.06 ± 2.13 35.37 ± 1.57 75.42 ± 4.19 1.21 ± 0.05
26.45 ± 1.28 2.18 ± 0.12 18.42 ± 1.06 3.23 ± 0.19 0.76 ± 0.04 8.03 ± 0.51 14.29 ± 0.63 5.07 ± 0.19 1.26 ± 0.01 16.42 ± 0.91 0.36 ± 0.01 3.53 ± 0.12 16.47 ± 0.66 45.52 ± 2.45 61.99 ± 2.88 1.31 ± 0.04
42.40 ± 1.87 1.53 ± 0.05 6.93 ± 0.26 0.99 ± 0.03 0.35 ± 0.01 1.82 ± 0.06 16.51 ± 0.71 17.50 ± 0.96 0.96 ± 0.04 8.97 ± 0.64 0.49 ± 0.02 1.55 ± 0.04 18.04 ± 1.12 36.19 ± 2.24 54.23 ± 2.07 1.03 ± 0.06
on cell growth and lipid accumulation in a fed-batch culture system. The fed-batch procedure contained two stages of feeding: three times of feeding with glucose-containing medium and four times of feeding with glucose. The feeding time, time course of cell
Fig. 3. Transcript levels biotin carboxylase (BC) and stearoyl ACP desaturae (SAD) in heterotrophic C. zofingiensis in batch culture with 50 g L1 of various sugars. The expression of actin gene was used as control. CTR, control cultured without sugar.
increase the biomass concentration by fed with key medium components or fresh medium at time intervals. It was indicated that C. zofingiensis could reach high specific growth rate (ca. 0.03 h1) and a high cell-density (ca. 53 g L1) in a fed-batch culture system (Sun et al., 2008). To further evaluate the lipid production of C. zofingiensis, a 3.7-L fermenter was used to investigate the effect of glucose
Fig. 4. Cell biomass, glucose and nitrate consumption and lipid production in a two-stage fed-batch fermentation of C. zofingiensis in a 3.7-L fermenter. (j) glucose concentration; (s) cell biomass; (column) lipid content; (h) lipid yield; (}) NO 3 concentration; (;) glucose-containing medium feeding; (;;) glucose feeding.
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growth and lipid production are shown in Fig. 4. In the fed-batch fermentation a high lipid yield of 20.7 g L1 was obtained, which was about 3.9-fold of that (e.g., 5.27 g L1) obtained by classical batch culture (Fig. 1, Fig. 4). At the stage-fed with glucose-containing medium, the algal cells grew fast but the cellular lipid content was low, possibly due to the relatively low C/N ratio as indicated by the remaining concentrations of glucose and nitrate in the medium (Fig. 4). When stage-fed with glucose, due to the consumption of nitrogen and the new addition of glucose, a high C/N ratio formed in the culture medium, resulting in a favorable condition for lipid accumulation within cells (Fig. 4). Under high-light stress condition, photoautotrophic C. zofingiensis produced lipids up to 2.21 g L1; while heterotrophic C. zofingiensis could produce lipids up to 20.7 g L1 with high lipid productivity of 1.38 g d1 L1 in a well-developed fermentation system. Chlorella protothecoides was previously reported to accumulate high levels of oil under heterotrophic growth condition (Xiong et al., 2008). Thus heterotrophic Chlorella species may have high potential for biodiesel production. The major drawback of using heterotrophic cultures for oils is the need of glucose or other sugars to feed the cells. Glucose, which takes up about 60% of total cost, depends on food-based agriculture. The price of industrial glucose can be as low as 100 USD per ton (www.alibada.com). According to Fig. 4, the conversion ratio of sugar to lipid is about 21.0%. Thus the roughly estimated cost for one liter oil produced by C. zofingiensis is around $0.9. Our study showed that C. zofingiensis could also grow well with other sugars as sole carbon sources (Fig. 1). Thus, by feeding C. zofingiensis with cheaper sugar sources from industrial or agricultural waste (Xu et al., 2006; Jiang et al., 2009; Cheng et al., 2009), the cost of algal oil could be further reduced. For example, cane molasses is a by-product of sugar industry, which consists of approximate 40–50% (w/w) of total sugars (Najafpour and Poi Shan, 2003) and costs around one fifth of glucose. Theoretically the cost of oil by C. zofingiensis can be greatly reduced by feeding the algal cells with cane molasses. Moreover, C. zofingiensis could yield high amount of the high-value astaxanthin under heterotrophic conditions (Ip and Chen, 2005). Thus, by feeding algae with ‘‘waste” sugars and tying algal oil production to other products (e.g. astaxanthin), profitable biodiesel from microalgae would not remain a research project for long. Other heterotrophic organisms, such as bacteria and yeasts, may be potential sources for biodiesel. Bacteria are less used for biodiesel production since they have low contents of oils and produce mostly complex lipid (Kalscheuer et al., 2006; Gouda et al., 2008). Yeasts have the ability of accumulating high levels of cellular lipids, ranging from 5.32 to 37.1 g L1 (Li et al., 2008b). Generally speaking, fermentation-based biodiesel production contributes the emission of CO2, which might limit the application of the microorganisms for biodiesel production to some extent. Unlike yeasts, C. zofingiensis can grow well photoautotrophically, mixotrophically, and heterotrophically. Thus, by switching its growth modes, C. zofingiensis may release much less CO2 during the accumulation of lipids, as has been demonstrated by Xiong et al. (2010).
4. Conclusions The effects of various sugars on fatty acid production by C. zofingiensis were investigated in heterotrophic culture systems. Glucose was found to be the best carbon source for cell growth and lipid production. Compared with batch culture of C. zofingiensis, fed-batch fermentation of C. zofingiensis allowed much higher yield (20.7 g L1) and productivity (1.38 g d1 L1) of lipids. C. zofingiensis has high potential for biodiesel production.
Acknowledgements This work was supported by a Seed Funding Program from the University of Hong Kong and the GRF Grant of the Research Grants council of Hong Kong.
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Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Production potential of Chlorella zofingienesis as a feedstock for biodiesel Jin Liu a, Junchao Huang a,*, King Wai Fan a, Yue Jiang b,**, Yujuan Zhong a, Zheng Sun a, Feng Chen a a b
School of Biological Science, The University of Hong Kong, Hong Kong, China Department of Biology, Hong Kong Baptist University, Kowloon Tong, Hong Kong, China
a r t i c l e
i n f o
Article history: Received 12 January 2010 Received in revised form 20 May 2010 Accepted 25 May 2010 Available online 7 July 2010 Keywords: Biodiesel Chlorella zofingiensis Fed-batch fermentation Lipid Microalgae
a b s t r a c t The lipid contents and fatty acid profiles of Chlorella zofingiensis cultured in the dark with various carbon sources were investigated. Glucose was found to be the best carbon source for the growth and lipid production. When cultivated with 50 g L1 glucose, C. zofingiensis accumulated lipids up to 52% of the dry biomass, with triacylglycerols (TAGs) accounting for 72.1% of the total lipids. Fatty acid profiles revealed that glucose contributed to the highest yield of total fatty acids (TFAs) and proportion of oleic acid (35.7% of TFAs), which corresponded to the strongest up-regulation of biotin carboxylase (BC) and stearoyl ACP desaturase (SAD) genes. In fed-batch cultivation based on glucose, the lipid yield and productivity of C. zofingiensis were further increased to 20.7 g L1 and 1.38 g d1 L1 respectively, representing 3.9-fold of those achieved in batch culture. We conclude that C. zofingiensis has great potential for biodiesel production. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction Petroleum-based fuels are recognized as unsustainable energy source due to their depleting supplies and contribution to global warming (Chisti, 2008). Renewable biofuels are promising alternatives to petroleum-based fuels, among which biodiesel has attracted the most attention in recent years (Knothe et al., 1997; Hu et al., 2008; Huang et al., 2010). Biodiesel is a diesel-equivalent fuel derived from biological feedstocks and is chemically referred to as a fatty acid methyl ester (FAME). Compared with traditional fuels, biodiesel is carbon neutral, contributes less emission of gaseous pollutants and hence is environmentally beneficial (Ma and Hanna, 1999). Currently biodiesel is mainly derived from vegetable oils, animal fats and waste cooking oils (Knothe et al., 1997; Lang et al., 2001; Zhang et al., 2003). However, those sources cannot even meet the growing need for transportation energy as large land areas have to be used for oil crops cultivation (Chisti, 2008). In addition, the relatively low productivity by oil plants also restricts the commercialization of biodiesel from plants (Lang et al., 2001). There is the need therefore, to search for a cost-effective source of biodiesel to fulfil industrial sustainability. Microalgae are currently being considered as feedstocks for biodiesel production because of their rapid growth rates and high yield of lipids (Chisti, 2008; Hu et al., 2008; Huang et al., 2009). Furthermore, unlike oil crops, microalgae can be easily cultured in outdoor ponds or indoor well-established bioreactors or fermentors (Chen,
* Corresponding author. Tel.: +852 22990309; fax: +852 22990311. ** Corresponding author. Tel.: +852 34117062; fax: +852 34115995. E-mail addresses: [email protected] (J. Huang), [email protected] (Y. Jiang). 0960-8524/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biortech.2010.05.082
1996; Del Campo et al., 2007; Pruvost et al., 2009). Thus microalgae are superior to oil crops for the sustainable production of biomass in an environmental-friendly operation mode. When a microalga is considered for mass biodiesel production, two key factors, namely cell biomass and lipid content are essential for the initial assessment. The green microalgae in the genus Chlorella consist of about 10 species that can grow photoautotrophically, mixotrophically and heterotrophically with high biomass concentration (Chen, 1996; Shi and Chen, 2002; Ip et al., 2004; Miao and Wu, 2004). Considering their lipid contents, C. vulgaris and C. protothecoides have been reported to be the candidates for biodiesel production under photoautotrophic or heterotrophic culture conditions (Miao and Wu, 2004; Liu et al., 2008; Hsieh and Wu, 2009). Certain Chlorella species was reported to attain very high biomass concentration in heterotrophic culture (Sun et al., 2008). In addition, lipids extracted from heterotrophically grown cells had similar properties to fuel oil in terms of oxygen content, heating value, density and viscosity (Miao and Wu, 2004). Moreover, cultivating Chlorella heterotrophically could eliminate the light requirement that could definitely reduce the cost of the final product (Chen, 1996). We reported previously that C. zofingiensis could grow rapidly in the dark with glucose as the sole carbon and energy source (Ip and Chen, 2005). Furthermore, by using fed-batch culture strategy, up to 53 g L1 biomass could be reached for this alga (Sun et al., 2008). In this study, we investigated the production potential of C. zofingiensis as a biodiesel feedstock, focusing on its growth and lipid contents under heterotrophic culture conditions with various organic carbon sources. The regulation of two key genes involved in fatty acid biosynthesis was also surveyed in order to deepen our fundamental understanding of biodiesel formation.
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2. Methods 2.1. Microalga strain C. zofingiensis (ATCC 30412) was obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). This alga was maintained at 4 °C on an agar slant of Kuhl medium (Kuhl and Lorenzen, 1964) consisting of (per liter) 1.01 g KNO3; 0.62 g NaH2PO4H2O; 0.089 g Na2HPO42H2O; 0.247 g MgSO47H2O; 14.7 mg CaCl22H2O; 6.95 mg FeSO47H2O; 0.061 mg H3BO3; 0.169 mg MnSO4H2O; 0.287 mg ZnSO47H2O; 0.0025 mg CuSO45H2O; and 0.01235 mg (NH4)6MO7O244H2O. The pH of the medium was adjusted to pH 6.5 prior to autoclaving. 2.2. Batch and fed-batch culture Inocula were prepared by growing the microalga in 250 mL Erlenmeyer flask containing 50 ml of the Kuhl medium at 25°C for 4 days with orbital shaking at 150 rpm and illuminated with continuous light of 30 lmol photon m2 s1. For batch culture, Erlenmeyer flasks, each containing 90 mL medium supplemented with various carbon sources (lactose, galactose, sucrose, fructose, mannose and glucose), were inoculated with 10% (v/v) of exponentially growing inoculum and then incubated at 25 °C in an orbital shaker at 150 rpm in the dark. The two-stage fed-batch strategy previously developed by us for high-biomass of C. zofingiensis (Sun et al., 2008) was used to investigate if the well-controlled fermentation system is superior to the classical batch culture system. The working volume of the fermentor (3.7-L, Bioengineering Ag, Wald, Switzerland) was 3.0 L. The cultivation conditions in the fermenter were controlled as follows: pH 6.5; temperature 25 °C; agitation 450 rpm; and dissolved oxygen concentration at 50% saturation. The pH value of medium was regulated by phosphate buffer which maintained the value near 6.5. During fed-batch cultivation, the sterilized stock nutrient solution was fed into the fermenter to maintain the glucose concentration at 5–20 g L1. 2.3. Determination of glucose concentration, nitrate concentration, dry cell weight and specific growth rate The cells were centrifuged at 3800g for 5 min. Glucose and nitrate concentrations in the supernatant were determined according to Miller (1959) and Elton-Bott (1979), respectively. The pellet was re-suspended in distilled water and filtered through a pre-dried Whatman GF/C filter paper (1.2 lm pore size). The algal cells on the filter paper discs were dried at 70 °C in a vacuum oven until constant weight and were cooled down to room temperature in a desiccator before weighting. The specific growth rate (l) at the exponential phase was calculated according to the equation, l = (ln X2 ln X1)/(t2 t1), where X2 and X1 are the dry cell weight concentration (g L1) at time t2 and t1, respectively.
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(GLs), and phospholipids (PLs) on silica cartridges (Waters, Milford, MA, USA) by sequential elution with chloroform, acetone, and methanol, as previously described (Christie, 2003). Neutral lipids were further resolved to subclasses by TLC (silica gel 60, 20 20 cm plates, 0.25 mm thickness; Merck, Whitehouse Station, NJ, USA) using a solvent system of petroleum ether:diethyl ether:acetic acid (70:30:1, v/v/v). Lipids were visualized by brief exposure to 2,7-dichlorofluorescein (Sigma, St. Louis, MO, USA) vapors and were identified by comparison with the standards (Sigma). Fatty acid methyl esters (FAMEs) were prepared by direct transmethylation of freeze-dried cells, lipid extracts, or individual lipids with sulphuric acid in methanol (Christie, 2003). The FAMEs were analyzed by using a HP 6890 capillary gas chromatograph (Hewlett–Packard, Palo Alto, CA) equipped with a flame ionization detector (FID) and a HP-INNOwax capillary column (30 m 0.32 mm) (Agilent Technologies, Inc., Wilmington, DE). Nitrogen was used as carrier gas. Initial column temperature was set at 170 °C, which was subsequently raised to 230 °C at 1 °C min1. The injector was kept as 250 °C with an injection volume of 2 lL under splitless mode. The FID temperature was set at 270 °C. FAMEs were identified by chromatographic comparison with authentic standards (Sigma). The quantities of individual FAMEs were estimated from the peak areas on the chromatogram using heptadecanoic acid as the internal standard. 2.5. RNA isolation and RT-PCR assay RNA was isolated from aliquots of about 108 cells using the TRI Reagent (Molecular Research Center, Cincinnati, OH, USA) according to the manufacturer’s instructions. The concentration of total RNA was determined spectrophotometrically at 260 nm. Total RNA (1 lg) extracted from different samples was reverse transcribed to cDNA by using a SuperScript III First-Strand Synthesis System (Invitrogen, Carlsbad, CA, USA) for reverse transcriptionPCR (RT-PCR) primed with oligo(dT) according to the manufacturer’s instructions. PCR amplification was carried out according to Li et al. (2008a) using specific primers of BC (forward, 50 GTGCGATTGGGTATGTGGGGGTG-30 , and reverse, 50 -CGACCAGGAC CAGGGCGGAAAT-30 ) and SAD (forward, 50 -TCCAGGAACGTGCCAC CAAG-30 , and reverse, 50 -GCGCCCTGTCTTGCCCTCATG-30 ). C. zofingiensis actin (ACT) primers (forward, 50 -TGCCGAGCGTGAAATTGT GAG-30 , and reverse, 50 -CGTGAATGCCAGCAGCCTCCA-30 ) were used to demonstrate equal amounts of templates and loading. The GenBank accession numbers for BC and SAD were GQ996717 and GQ996719, respectively. Amplification of the cDNA was done by conventional PCR [94 °C for 2 min followed by 24 cycles (for ACT gene) or 26 cycles (for BC and SAD genes) of 94 °C for 15 s, 58 °C for 20 s, 72 °C for 30 s]. PCR products were separated on a 2% agarose gel and stained with ethidium bromide for photography (Biorad, Hercules, CA, USA). 3. Results and discussion
2.4. Lipid extraction and fractionation and fatty acid analysis Cells were harvested and lyophilized for lipid extraction and fatty acids analysis. Lipids were extracted with chloroform–methanol (2:1, v/v), and then separated into chloroform and aqueous methanol layers by addition of methanol and water to give a final solvent ratio of chloroform:methanol:water of 1:1:0.9. The chloroform layer was washed with 20 mL of a 5% NaCl solution, and evaporated to dryness. Thereafter, the total lipids were measured gravimetrically and stored at 20 °C under nitrogen gas to prevent lipid oxidation or used directly for subsequent analysis. Total lipid extracts were fractionated into neutral lipids (NLs), glycolipids
3.1. Heterotrophic growth and lipid production of C. zofingiensis with various carbon sources We have previously reported that C. zofingiensis could grow well in the dark with various sugars as carbon sources (Sun et al. 2008). To investigate the influences of carbon sources on lipid production in C. zofingiensis cells, we cultured the algal cells in the dark with lactose, galactose, sucrose, fructose, mannose or glucose as a carbon source, respectively. Among the tested sugars, glucose gave the highest growth rate (0.03 h1), cell biomass (10.1 g L1), lipid content (0.52 g g1), and lipid yield (5.27 g L1). Algal cells cultured
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with mannose, fructose, or sucrose produced slightly lower amounts of cell biomass and lipids. In contrast, lactose and galactose were observed to be poor carbon sources for lipid production since low biomass and lipid contents were obtained (Fig. 1). These findings demonstrated a close relationship between lipid biosynthesis and cell growth for the heterotrophic alga. As shown in Fig. 2, based on lipid class analysis of C. zofingiensis cultured with glucose, neutral lipids (NLs) were found to be the major constituent that accounted for 80.9% of the total lipids, while phospholipids (PLs) and glycolipids (GLs) together accounted for 19.1%. In NLs, TAGs were the predominant component, accounting for 72.1% of the total lipids. In the batch culture, up to 0.375 g g1 TAGs were accumulated in heterotrophic C. zofingiensis cells. Unlike PLs and GLs that are membrane-bounded lipids, TAGs serve primarily as a storage form of carbon and energy (Hu et al., 2008). In addition, TAGs are superior to phospholipids (PLs) or glycolipids (GLs) for biodiesel due to their higher content of fatty acids (Pruvost et al., 2009). Hence C. zofingiensis may act as a promising host for biodiesel production due to its fast cell growth and high content of TAGs.
3.2. Fatty acid profiles of dark-grown C. zofingiensis cultures Fatty acid composition considerably influences the properties of biodiesel such as cetane number, heat of combustion, oxidative stability, cloud point, lubricity, which will finally influence the quality of biodiesel (Knothe, 2009). In this study, the fatty acid compositions of the heterotrophically cultured C. zofingiensis with various sugars were investigated (Table 1). It was found that C16:0, C16:2, C18:1, C18:2 and C18:3 (n-3) were the major fatty acids, which accounted for more than 84% of the total fatty acids (TFAs). The highest amounts of TFAs (45.4% of cell biomass) and C18:1 (35.7% of TFAs) were achieved in C. zofingiensis cultured with glucose as the carbon source, which were about five and three times, respectively of those given by lactose. Furthermore, the fatty acid profiles of NLs, GLs and PLs from algal cells cultivated with glucose were also investigated. Compared with GLs and PLs, NLs had a lower proportion of C16:0 but a much higher percentage of oleic acid (Table 2). As ideal biodiesel, the fatty acids should be oxidative and low-temperature stable. Generally, saturated fatty acids are oxidative stable, while unsaturated fatty acids give low-temperature stability (Knothe, 2008). The enhanced proportion of oleic acid (C18:1) in the total fatty acids has been considered as a feasible ap-
Fig. 2. Distribution of NLs, GLs and PLs in total lipids extracted from heterotrophic C. zofingiensis cells in batch culture with 50 g L1 of glucose for 14 days. The horizontal line inside the NLs bar marks the portion of TAGs in this fraction.
proach to balance the oxidative and low-temperature stability with retaining the cetane number at an acceptable level (Knothe, 2009). The high content of oleic acid in the heterotrophic cells further supports that C. zofingiensis is a favorable host for producing high quality biodiesel. 3.3. Sugars up-regulate the transcription of BC and SAD genes of C. zofingiensis The de novo biosynthesis of fatty acid initially catalyzed in chloroplast by the key enzyme acetyl-CoA carboxylase (ACCase); while the introduction of the first double bond to acyl chain is carried out by stearoyl ACP desaturae (SAD), an enzyme playing an important role in determining the ratio of unsaturated and saturated fatty acids (Hu et al., 2008). To reveal the relationship between the fatty acid profiles and the regulation of ACCase and SAD, we investigated the transcript levels of SAD and biotin carboxylase (BC, a subunit of ACCase) in dark-grown C. zofingiensis cells cultured with various sugars using the RT-PCR approach. The transcription of the two genes was shown to be differentially regulated by various carbon sources (Fig. 3). In coincidence with the higher contents of TFAs in cells cultured with sucrose, fructose, mannose or glucose, the transcript levels of BC in the cells were strongly up-regulated. In contrast, galactose moderately enhanced the transcription of BC whereas lactose, the poor carbon source for the cell growth and fatty acid accumulation, caused a slight up-regulation of BC. These results are consistent with the findings of Ohlrogge and Jaworski (1997) that the first reaction step of fatty acid biosynthesis catalyzed by ACCase is a major point of flux control for this pathway. A similar pattern induced by sugars was also found for SAD at mRNA levels, although much stronger up-regulation was observed when compared with the expression of BC (Fig. 3). Similarly, lactose induced low-level expression of SAD while glucose triggered high amounts of SAD transcripts, which is consistent with the unsaturation values of fatty acids in the algal cells cultured with the tested sugars (Table 1). 3.4. Fed-batch fermentation enhances lipid production by C. zofingiensis
Fig. 1. Cell biomass (black column), lipid content (gray column) and yield (white column) of C. zofingiensis in batch culture with 50 g L1 of various sugars. Lac, lactose; Gal, galactose; Suc, sucrose; Fru, fructose; Man, mannose; Glu, glucose.
As shown from the above results, the productivity of lipid is closely related to the growth rate, cell density and cellular lipid content. Compared with batch culture, fed-batch cultivation mode can extend the exponential growth phase of the cell and further
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J. Liu et al. / Bioresource Technology 101 (2010) 8658–8663 Table 1 Fatty acid profiles of dark-grown C. zofingiensis cultured with 50 g L1 of various sugars for 14 days. Fatty acids
Sugars
C16:0 C16:1 C16:2 C16:3 C16:4 C18:0 C18:1 C18:2 C18:3 (n-6) C18:3 (n-3) C18:4 Others Mounsatda Pounsatdb Unsatdc DUS (r/mol)d TFAe a b c d e
Lactose
Galactose
Sucrose
Fructose
Mannose
Glucose
28.73 ± 1.22 3.42 ± 0.12 9.02 ± 0.35 5.28 ± 0.27 0.92 ± 0.05 0.95 ± 0.06 12.59 ± 0.45 20.87 ± 0.87 1.39 ± 0.03 12.86 ± 0.39 1.24 ± 0.04 2.72 ± 0.09 16.01 ± 0.55 51.59 ± 2.11 67.59 ± 2.95 1.43 ± 0.05 8.48 ± 0.33
27.81 ± 0.99 2.52 ± 0.13 11.08 ± 0.61 2.52 ± 0.11 0.65 ± 0.03 1.61 ± 0.09 25.15 ± 0.89 16.48 ± 0.83 1.17 ± 0.05 7.74 ± 0.31 1.10 ± 0.06 2.17 ± 0.11 27.67 ± 0.82 40.74 ± 2.05 68.42 ± 2.31 1.24 ± 0.06 20.30 ± 0.71
23.16 ± 0.83 1.49 ± 0.06 8.48 ± 0.39 1.96 ± 0.07 0.23 ± 0.02 2.80 ± 0.09 31.99 ± 1.21 19.51 ± 0.75 0.55 ± 0.01 7.36 ± 0.35 0.46 ± 0.01 2.00 ± 0.07 33.48 ± 1.12 38.57 ± 1.28 72.05 ± 3.72 1.22 ± 0.06 40.48 ± 1.29
23.62 ± 1.12 1.71 ± 0.08 8.31 ± 0.44 2.12 ± 0.09 0.17 ± 0.01 2.34 ± 0.12 32.06 ± 0.93 19.27 ± 0.49 0.52 ± 0.03 7.48 ± 0.18 0.45 ± 0.01 1.94 ± 0.04 33.78 ± 1.03 38.33 ± 1.49 72.10 ± 3.11 1.22 ± 0.03 43.01 ± 1.11
23.38 ± 1.03 1.49 ± 0.05 7.94 ± 0.29 1.83 ± 0.06 0.19 ± 0.01 2.44 ± 0.10 33.30 ± 1.45 19.36 ± 0.98 0.52 ± 0.03 7.31 ± 0.23 0.45 ± 0.02 1.78 ± 0.08 34.79 ± 1.37 37.61 ± 1.44 72.40 ± 3.13 1.21 ± 0.03 43.10 ± 1.79
22.62 ± 0.77 1.97 ± 0.08 7.38 ± 0.33 1.94 ± 0.04 0.22 ± 0.02 2.09 ± 0.08 35.68 ± 1.23 18.46 ± 0.66 0.51 ± 0.02 7.24 ± 0.30 0.49 ± 0.02 1.40 ± 0.08 37.64 ± 1.26 36.24 ± 1.63 73.89 ± 2.99 1.21 ± 0.05 45.38 ± 1.83
Mounsatd: percentage of monounsaturated fatty acids (% of total fatty acids). Pounsatd: percentage of polyunsaturated fatty acids (% of total fatty acids). Unsatd: percentage of unsaturated fatty acids (% of total fatty acids). DUS (r/mol): degree of fatty acid unsaturation = [1.0 (% monoenes) + 2.0 (% dienes) + 3.0 (% trienes) + 4.0 (% tetraenes)]/100. TFA: total fatty acids (g)/cell dry weight (g) 100%.
Table 2 Fatty acid profiles of neutral lipids, glycolipids and phospholipids from heterotrophic C. zofingiensis cells cultivated with 50 g L1 glucose for 14 days. Fatty acids
NLs
GLs
PLs
C16:0 C16:1 C16:2 C16:3 C16:4 C18:0 C18:1 C18:2 C18:3 (n-6) C18:3 (n-3) C18:4 Others Mounsatd Pounsatd Unsatd DUS (r/mol)
20.10 ± 0.83 1.65 ± 0.06 6.38 ± 0.35 1.80 ± 0.07 0.25 ± 0.02 2.76 ± 0.11 38.41 ± 2.15 19.20 ± 1.1 0.45 ± 0.02 6.86 ± 0.28 0.42 ± 0.02 1.71 ± 0.05 40.06 ± 2.13 35.37 ± 1.57 75.42 ± 4.19 1.21 ± 0.05
26.45 ± 1.28 2.18 ± 0.12 18.42 ± 1.06 3.23 ± 0.19 0.76 ± 0.04 8.03 ± 0.51 14.29 ± 0.63 5.07 ± 0.19 1.26 ± 0.01 16.42 ± 0.91 0.36 ± 0.01 3.53 ± 0.12 16.47 ± 0.66 45.52 ± 2.45 61.99 ± 2.88 1.31 ± 0.04
42.40 ± 1.87 1.53 ± 0.05 6.93 ± 0.26 0.99 ± 0.03 0.35 ± 0.01 1.82 ± 0.06 16.51 ± 0.71 17.50 ± 0.96 0.96 ± 0.04 8.97 ± 0.64 0.49 ± 0.02 1.55 ± 0.04 18.04 ± 1.12 36.19 ± 2.24 54.23 ± 2.07 1.03 ± 0.06
on cell growth and lipid accumulation in a fed-batch culture system. The fed-batch procedure contained two stages of feeding: three times of feeding with glucose-containing medium and four times of feeding with glucose. The feeding time, time course of cell
Fig. 3. Transcript levels biotin carboxylase (BC) and stearoyl ACP desaturae (SAD) in heterotrophic C. zofingiensis in batch culture with 50 g L1 of various sugars. The expression of actin gene was used as control. CTR, control cultured without sugar.
increase the biomass concentration by fed with key medium components or fresh medium at time intervals. It was indicated that C. zofingiensis could reach high specific growth rate (ca. 0.03 h1) and a high cell-density (ca. 53 g L1) in a fed-batch culture system (Sun et al., 2008). To further evaluate the lipid production of C. zofingiensis, a 3.7-L fermenter was used to investigate the effect of glucose
Fig. 4. Cell biomass, glucose and nitrate consumption and lipid production in a two-stage fed-batch fermentation of C. zofingiensis in a 3.7-L fermenter. (j) glucose concentration; (s) cell biomass; (column) lipid content; (h) lipid yield; (}) NO 3 concentration; (;) glucose-containing medium feeding; (;;) glucose feeding.
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growth and lipid production are shown in Fig. 4. In the fed-batch fermentation a high lipid yield of 20.7 g L1 was obtained, which was about 3.9-fold of that (e.g., 5.27 g L1) obtained by classical batch culture (Fig. 1, Fig. 4). At the stage-fed with glucose-containing medium, the algal cells grew fast but the cellular lipid content was low, possibly due to the relatively low C/N ratio as indicated by the remaining concentrations of glucose and nitrate in the medium (Fig. 4). When stage-fed with glucose, due to the consumption of nitrogen and the new addition of glucose, a high C/N ratio formed in the culture medium, resulting in a favorable condition for lipid accumulation within cells (Fig. 4). Under high-light stress condition, photoautotrophic C. zofingiensis produced lipids up to 2.21 g L1; while heterotrophic C. zofingiensis could produce lipids up to 20.7 g L1 with high lipid productivity of 1.38 g d1 L1 in a well-developed fermentation system. Chlorella protothecoides was previously reported to accumulate high levels of oil under heterotrophic growth condition (Xiong et al., 2008). Thus heterotrophic Chlorella species may have high potential for biodiesel production. The major drawback of using heterotrophic cultures for oils is the need of glucose or other sugars to feed the cells. Glucose, which takes up about 60% of total cost, depends on food-based agriculture. The price of industrial glucose can be as low as 100 USD per ton (www.alibada.com). According to Fig. 4, the conversion ratio of sugar to lipid is about 21.0%. Thus the roughly estimated cost for one liter oil produced by C. zofingiensis is around $0.9. Our study showed that C. zofingiensis could also grow well with other sugars as sole carbon sources (Fig. 1). Thus, by feeding C. zofingiensis with cheaper sugar sources from industrial or agricultural waste (Xu et al., 2006; Jiang et al., 2009; Cheng et al., 2009), the cost of algal oil could be further reduced. For example, cane molasses is a by-product of sugar industry, which consists of approximate 40–50% (w/w) of total sugars (Najafpour and Poi Shan, 2003) and costs around one fifth of glucose. Theoretically the cost of oil by C. zofingiensis can be greatly reduced by feeding the algal cells with cane molasses. Moreover, C. zofingiensis could yield high amount of the high-value astaxanthin under heterotrophic conditions (Ip and Chen, 2005). Thus, by feeding algae with ‘‘waste” sugars and tying algal oil production to other products (e.g. astaxanthin), profitable biodiesel from microalgae would not remain a research project for long. Other heterotrophic organisms, such as bacteria and yeasts, may be potential sources for biodiesel. Bacteria are less used for biodiesel production since they have low contents of oils and produce mostly complex lipid (Kalscheuer et al., 2006; Gouda et al., 2008). Yeasts have the ability of accumulating high levels of cellular lipids, ranging from 5.32 to 37.1 g L1 (Li et al., 2008b). Generally speaking, fermentation-based biodiesel production contributes the emission of CO2, which might limit the application of the microorganisms for biodiesel production to some extent. Unlike yeasts, C. zofingiensis can grow well photoautotrophically, mixotrophically, and heterotrophically. Thus, by switching its growth modes, C. zofingiensis may release much less CO2 during the accumulation of lipids, as has been demonstrated by Xiong et al. (2010).
4. Conclusions The effects of various sugars on fatty acid production by C. zofingiensis were investigated in heterotrophic culture systems. Glucose was found to be the best carbon source for cell growth and lipid production. Compared with batch culture of C. zofingiensis, fed-batch fermentation of C. zofingiensis allowed much higher yield (20.7 g L1) and productivity (1.38 g d1 L1) of lipids. C. zofingiensis has high potential for biodiesel production.
Acknowledgements This work was supported by a Seed Funding Program from the University of Hong Kong and the GRF Grant of the Research Grants council of Hong Kong.
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