Bioresource Technology 156 (2014) 117–122
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Scale-up cultivation of Chlorella ellipsoidea from indoor to outdoor in bubble column bioreactors Shi-Kai Wang a,c, Yi-Ru Hu a,c, Feng Wang a, Amanda R. Stiles b, Chun-Zhao Liu a,⇑ a National Key Laboratory of Biochemical Engineering & Key Laboratory of Green Process and Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing 100190, PR China b Department of Plant and Microbial Biology, University of California, Berkeley, CA 94720, USA c University of Chinese Academy of Sciences, Beijing 100049, PR China
h i g h l i g h t s Chlorella ellipsoidea cells were cultivated in bubble column bioreactors. Microalgal cells were able to quickly adapt to the outdoor conditions. Biomass production cost in outdoor culture was lower than that in indoor culture.
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Article history: Received 28 November 2013 Received in revised form 4 January 2014 Accepted 6 January 2014 Available online 17 January 2014 Keywords: Chlorella ellipsoidea Bubble column bioreactor Outdoor cultivation Fatty acid composition Economic evaluation
a b s t r a c t The cultivation of Chlorella ellipsoidea in bubble column bioreactors was investigated at different scales under indoor and outdoor conditions. The algal cells were able to quickly adapt to the outdoor conditions and achieved a growth rate of 31.55 mg L1 day1. Due to differences in light and temperature, the outdoor culture produced a higher percentage of unsaturated fatty acids compared to the indoor cultures, while the amino acid composition was unaffected. The overall cost of the biomass produced by the 200 L outdoor cultivation (58.70 US$/kg-dry weight) was estimated to be more than 7 times lower than that of the 20 L indoor cultivation (431.39 US$/kg-dry weight). Together these results provide a basis for the cultivation of C. ellipsoidea for the large-scale production of biofuels, high-value nutrients and/or recombinant proteins. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Microalgae are photoautotrophic sunlight-driven cell factories that can convert carbon dioxide to various products such as lipids, carbohydrates, proteins, fatty acids, vitamins, antibiotics, and antioxidants (Chisti, 2007). In addition to these naturally-produced compounds, microalgae are also a promising platform for the production of recombinant proteins. Microalgae have unique advantages, including a rapid growth rate, ease of cultivation, and the ability to make the same post-transcriptional and post-translational modifications as plants (Potvin and Zhang, 2010). The feasibility of utilizing microalgae as a production system for therapeutic or industrial proteins has been previously demonstrated (Specht et al., 2010). Chlorella ellipsoidea is a single-celled eukaryotic green algae that has been extensively studied. As with other Chlorella sp,
⇑ Corresponding author. Tel./fax: +86 10 82622280. E-mail address: [email protected] (C.-Z. Liu). 0960-8524/$ - see front matter Ó 2014 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2014.01.023
it can be found in both fresh and marine water systems, and it can be cultured under autotrophic, heterotrophic, mixotrophic, or photoheterotrophic growth conditions (Mata et al., 2010). It is rich in high-quality proteins, vitamins, lipid-soluble compounds, glycolipids, sulfolipids, and compounds used as food additives. It is also beneficial for human health due to its ability to lower blood sugar levels and increase hemoglobin concentrations, and it is used to enhance animal growth when it is applied as an aquaculture or animal feed (Mata et al., 2010; Kay, 1991). Furthermore, C. ellipsoidea is a well-established model organism and a promising bioreactor for the production of complex foreign proteins for pharmaceutical and industrial use (Wang et al., 2003; Walker et al., 2005). The genetic transformation of C. ellipsoidea is feasible, and it has been previously utilized for the recombinant production of flounder growth hormone (FGH) and rabbit neutrophil peptide-1 (NP-1) (Liu et al., 2013; Kim et al., 2002; Chen et al., 2001; Bai et al., 2013). For the industrial production of products from microalgae, it is necessary to evaluate the feasibility of scaled-up outdoor culture systems. The utilization of natural sunlight and uncontrolled
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outdoor culture conditions can reduce the overall costs of the cultivation process (Feng et al., 2011). Raceway ponds and other open-culture systems are commonly used for large-scale outdoor cultivation due to their low costs for construction and operation; however, they often perform poorly due to contamination risk, lack of control of the culture conditions, and problems with mixing and light utilization efficiency (Mata et al., 2010). Closed bioreactors can provide a suitable environment in terms of light, nutrients, CO2, and temperature, and they have the potential to produce higher rates of biomass production and improved culture quality (Brennan and Owende, 2010). The feasibility of Chlorella sp. outdoor cultivation has been confirmed by many studies using different types of bioreactors at scales ranging from 10 to 70 L, including a flat glass plate bioreactor, a cylindrical glass bottle bioreactor, and a nylon membrane column bioreactor. The results have demonstrated that many Chlorella strains can adapt to the fluctuating temperatures and irradiance levels inherent in outdoor cultivation, and that these systems can be used for biodiesel production (Feng et al., 2011, 2012; Zhou et al., 2013). In addition, the outdoor production of Chlorella sp. can also utilize the waste produced by other processes for nutrients, such as organic fertilizer derived from food waste or manure, dairy and piggery wastewater, or flue gas generated by the combustion of natural gas (Chisti, 2013). This can decrease the cost of the cultivation process (Doucha et al., 2005; Lam and Lee, 2012; Huo et al., 2012; Zhu et al., 2013). However, the growth rate is often reduced by outdoor cultivation; for example, the growth rate of Calluna vulgaris was reduced by 27% in a study by Lam and Lee comparing outdoor cultivation with cultivation in a controlled environment (Lam and Lee, 2012). In addition, the lipid content and fatty acid composition also varied in response to climatic variation (Olofsson et al., 2012), which may influence the quality of the final products. Although there are numerous reports describing the outdoor cultivation of microalgae, studies focusing on C. ellipsoidea are rare and the culture scale in published studies is limited. It is important to understand the variations in growth and metabolism between indoor and outdoor cultivation conditions in order to determine the feasibility of outdoor cultivation for this species. In the current study, a 200 L outdoor cultivation system was constructed using a bubble column bioreactor made of a polyethylene membrane. The cultivation of C. ellipsoidea in bubble column bioreactors was scaled-up from the 20 L indoor culture to a 200 L outdoor culture, and the biomass, fatty acid content and composition, protein content, and amino acid composition were investigated and compared with that of 2 L sterilized flanged glass bioreactor. The effect of controlled and uncontrolled light intensity and temperature on C. ellipsoidea’s growth and metabolism was analyzed, and the cost of each cultivation system was calculated.
constructed from transparent polyethylene (PE) bag with a thickness of 0.2 mm (Yangpu Packaging Material Co., Ltd., Hebei, China). The bottom of the plastic culture bags with the desired dimensions was heat-sealed by a SF-B 800 pedal sealing machine (Xingye Machinery Equipment Co., Ltd., Wenzhou, China). The size of all bioreactors was shown in Supplementary material-Table 1. Air was supplied using an air sparger (compressed stainless steel particles) at the bottom of each bioreactor. Each 20 L bioreactor was handed at a shelf, and each 200 L bioreactor was supported by a circular wire mesh. The indoor cultivations were performed at 25 °C under a light/ dark cycle of 16/8 h with 25,000 lux illumination intensity, and the outdoor cultivations were conducted under the natural temperature and light conditions in Haidian district of Beijing, China (latitude 39°590 N, longitude 116°190 E), in autumn (September 29th, 2012 to October 19th, 2012). The initial biomass concentration was 0.075 g/L, and CO2-enriched air (1%) was bubbled into the reactor at a flow rate of 0.1 v/v/min through a 0.22 lm filter (Millipore, MA, USA) for the 2 L indoor cultivation and directly aerated into the reactor for the 20 L indoor and 200 L outdoor cultures. For the 2 L indoor culture, the reactor and the air supply systems were sterilized by autoclaving at 121 °C for 40 min prior to starting the cultivation. 2.2. Microalgae growth measurement To determine the relationship between the optical density at 680 nm (OD680) and the algal biomass (g/L), the algal culture broth was diluted or concentrated to a certain OD680, then harvested by centrifugation for 5 min at 10,000 rpm, 4 °C. The algal pellets were then washed three times with distilled water and dried at 105 °C to a constant weight. The dry weight (DW, g/L) of the algal cells was determined gravimetrically. The relationship between the OD680 and the C. ellipsoidea biomass was as follows:
DWðg=LÞ ¼ 0:38352 OD680 0:02647 ðR2 ¼ 0:9996Þ
The algal growth was measured every two days by measuring the OD680 of the culture broth using a UV-2100 spectrophotometer (Unico, Shanghai, China) and the biomass was calculated using Eq. (1). The maximum specific growth rate (lmax, day1) at the exponential stage was calculated as follows:
lmax ðday1 Þ ¼ ðln DW2 ln DW1 Þ=ðt2 t1 Þ
ð2Þ
where DW1 and DW2 were the dry biomass weight (g/L) at time t1 and t2, respectively. The doubling time (TD, days) was calculated as follows:
T D ðdaysÞ ¼ lnð2Þ=lmax 2. Methods 2.1. Microalgae strain and culture systems
ð3Þ 1
The rate of biomass production (P, mg L according to the following equation: 1
C. ellipsoidea (UTEX 20), kindly provided by Prof. Zan-Min Hu (Institute of Genetics and Developmental Biology, Chinese Academy of Sciences), was grown on BG-11 medium. The cells were maintained in 250 mL Erlenmeyer flasks containing 100 mL liquid medium and incubated at 25 °C in an orbital shaker at 100 rpm. Algal cells were grown photoautotrophically under a 16 h light period per day with 35 lmol m2 s1 for 15 days. The pre-cultured cells were used as the inoculum for all bubble-column cultivation experiments. The 2 L bubble column bioreactor was constructed from a flanged glass column (Beijing Glass Group Company, Beijing, China), and the 20 and 200 L bubble column bioreactors were
ð1Þ
Pðmg L1 day Þ ¼ ðDWx DW0 Þ=t x
1
day
) was calculated
ð4Þ
where DW0 and DWx were the initial dry biomass weight (mg/L) and the dry biomass weight (mg/L) at time tx, respectively. 2.3. Lipid analysis For the extraction and transesterification of the fatty acids, 50 mg lyophilized algal biomass was ground for 10 min and then dispersed in 3 mL of a 7.5% (w/v) KOH/CH3OH solution with 200 mg of heptadecanoic acid added as an internal standard. After the solution was incubated at 70 °C for 4 h, 2 mL HCl/CH3OH (1:1, v/v) and 2 mL 14% BF3/CH3OH (ANPEL, Shanghai, China) were
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2.4. Algal protein and amino acid analysis The total intracellular protein was determined using the Kjeldahl method as described by Lynch and Barbano (Lynch and Barbano, 1999), and the total algal protein content was calculated by multiplying by a factor of 6.38, a value previously determined for the measurement of Chlorella sp. protein content (Li et al., 2013). For the amino acid composition analysis, the protein was hydrolyzed using 4.2 M NaOH for tryptophan, performic acid for cysteine, and 6 M HCl for the other amino acids. The hydrolyzed amino acid samples were separated using ion exchange chromatography and quantified using an automatic amino acid analyzer (Hitachi L-8900, Japan).
2.5. Measurement of climatic conditions The light intensity and culture temperature were recorded at 8:00, 12:00, 16:00, and 20:00 every two days. The light intensity was measured using a JD-3 luxmeter (Jiading Xuelian Instrument Factory, Shanghai, China), and the temperature was measured using a spirit thermometer. The samples were collected every two days at 9:00 am.
2.6. Economic assumptions The total cost of the cultivation process includes both a fixed capital investment and operational costs (Li et al., 2011). In this study, the fixed capital investments are listed in Supplementary material-Table 2. The costs were calculated based on one batch cultivation and the estimated lifetime of the equipment. The bioreactors used for the 20 and 200 L cultivations utilized a single-use PE membrane which cost US$0.33/m (width: 0.75 m, double layer) resulting in a cost of US$0.165 and US$0.654 for the 20 and 200 L cultivation, respectively. The shelf used for the outdoor culture cost US$130 with an estimated lifetime of 10 years. The cost of the air compressor used for the 20 L indoor cultivation was US$32.68 with an estimated lifetime of 5 years of continuous operation. The cost of the air compressor used for the outdoor cultivation was US$620.9 and estimated to last for 7 years. The light used for the indoor cultivation was US$1.63 with an estimated lifetime of 3000 h under continuous use. Four lights were utilized as a unit to illuminate for a 20 L bioreactor. The investment capital and the power consumption of the cooling system for the indoor cultivations were based on the ratio of the bioreactor volume to the total volume of the room. The electricity consumption was calculated based on the power of the instruments and their operation time. The price of the electricity and water was based on the local price (US$0.078/kW h and US$0.65/ton, respectively), and the cost of the medium was estimated based on the market price at Fang-Dou chemical web (http://www.16ds.com/). The local price for CO2 was US$0.245/kg.
3. Results and discussion 3.1. Scale-up cultivation of C. ellipsoidea from indoor to outdoor After a two day lag period, the algal cells were able to survive and adapt to the outdoor conditions in the 200 L bioreactor (Supplementary material-Fig. 1). Following the lag period, the cells entered the exponential phase, and they reached the stationary phase at 14 days (Fig. 1). As shown in Table 1, the maximum specific growth rates (lmax) of the indoor cultures at the exponential stage (0.168 and 0.162 day1 for the 2 and 20 L cultures, respectively) were higher than that of the outdoor culture (0.145 day1), and this was accompanied by a shorter doubling time (4.13 and 4.28 days for the indoor cultures versus 4.78 days for the outdoor culture). This resulted in a rate of biomass production as high as 47.71 and 39.85 mg L1 day1 in the indoor cultures while the biomass production rate of the outdoor culture reached only 31.55 mg L1 day1 (Table 1). Similar results have been reported by Zhu et al. (2013). This different in growth rate is due to the fact that the algal cells cultured indoors are cultured in optimal controlled conditions rather than the uncontrolled outdoor natural environment. As illustrated in Supplementary material-Fig. 2, the varied weather conditions in the uncontrolled outdoor environment resulted in dramatic fluctuations in the temperature and light intensity during cultivation, and the natural day length (approximately 1213 h day1) was shorter than the illumination time applied to the indoor culture (16 h day1). In addition, algae cultivated outdoors are more vulnerable to contamination by foreign microalgae and microorganisms than the indoor cultures, which may also have a negative impact on the algal growth. The fatty acid (FA) compositions of C. ellipsoidea cultures under indoor and outdoor conditions are listed in Table 2. There were eight to nine detectable fatty acids, and the fatty acids with 16 and 18 carbon atoms accounted for more than 99% of the total fatty acids. Fatty acids with 16 and 18 carbon atoms are the ideal components for biodiesel, indicating that C. ellipsoidea could function as a potential species for the production of biodiesel (Xu et al., 2006). In addition, linolenic acid (18:3), which is beneficial for the treatment of cardiovascular diseases (Huo et al., 2012), was high in all culture conditions (more than 15% of the total FA), especially in the outdoor cultivation (as high as 40.26% of total FA).
0.8
0.6
Biomass (g/L)
added and transesterification occurred at 70 °C for 1.5 h. The mixture was centrifuged at 2000 rpm for 3 min at 4 °C following the addition of 1 mL 0.9% NaCl and 4 mL n-hexane. The organic phase was volatilized under a nitrogen atmosphere and re-dissolved in 4 mL chloroform for GC analysis. The fatty acid methyl esters (FAMEs) were analyzed using a gas chromatogarph equipped with an FID detector (Agilent, 7890 A, America) and an Agilent DB-17 column (30 m 0.25 mm). The injector and detector temperatures were 250 °C. The oven temperature was set at 180 °C for 1 min, increased from 180 to 230 °C at a rate of 6 °C/min, and then remained at 230 °C for 25 min.
0.4
2 L Indoor 20 L Indoor 200 L Outdoor
0.2
0.0
0
4
8 Time (day)
12
16
Fig. 1. The growth curve of C. ellipsoidea under different cultivation conditions.
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Table 1 Growth kinetic parameters of the algal cells under different culture conditions. Culture condition
Specific growth rate lmax (day1)
Doubling time (days)
Final biomass concentration (g L1)
Rate of biomass production (mg L1 day1)
2 L Indoor 20 L Indoor 200 L Outdoor
0.168 ± 0.006 0.162 ± 0.011 0.145 ± 0.026
4.13 ± 0.04 4.28 ± 0.15 4.78 ± 0.24
0.77 ± 0.05 0.66 ± 0.04 0.53 ± 0.10
47.71 ± 0.9 39.85 ± 1.6 31.55 ± 7.5
Values in a column are significantly different (P < 0.05) from each other according to t-test.
Table 2 The fatty acid composition of the algal cells cultured in different conditions. FA composition
14:0 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 C16 C18 SFA MUFA PUFA Total (% of DW)
FA content (% of total FA) 2 L indoor
20 L indoor
200 L outdoor
0.97 ± 0.09 19.68 ± 1.23 – 4.95 ± 0.11 4.71 ± 0.88 1.61 ± 0.05 10.60 ± 1.34 38.12 ± 2.04 19.37 ± 0.99 29.34 ± 0.08 69.70 ± 1.22 22.26 ± 0.92 10.60 ± 1.09 67.15 ± 1.78 8.94 ± 0.21
0.92 ± 0.11 18.64 ± 1.09 0.92 ± 0.10 3.46 ± 1.08 4.59 ± 0.15 1.94 ± 0.03 10.59 ± 0.20 41.99 ± 1.75 16.94 ± 0.68 27.61 ± 0.38 71.46 ± 1.15 21.50 ± 0.35 11.51 ± 0.77 66.98 ± 1.11 8.71 ± 0.22
0.99 ± 0.14 14.62 ± 1.50 2.14 ± 1.23 11.12 ± 0.88 14.77 ± 2.27 2.18 ± 0.54 2.77 ± 0.99 10.00 ± 1.74 40.26 ± 1.97 42.64 ± 0.80 55.22 ± 1.63 17.79 ± 1.84 4.90 ± 1.93 76.15 ± 2.83 7.44 ± 0.58
a a a a a a
a a a a a a
b b b b b b
Values in a row with different letters are significantly different (P < 0.05) according to t-test.
There were no obvious differences in the FA composition between the 2 and 20 L indoor cultures. However, the FA composition of the outdoor culture was distinct from the indoor cultures. Similar to a report by Zhu et al. (2013), the proportion of unsaturated fatty acids (including monounsaturated fatty acids (MUFA) and polyunsaturated fatty acids (PUFA)) in the algal cells cultivated outdoors was higher than that from the indoor cultures, especially the PUFA. In addition, fatty acids containing 16 carbon atoms were higher in the outdoor culture (42.64% of the total FA) compared to the indoor cultures (29.34% and 27.61% of the total FA for the 2 and 20 L cultures, respectively). Fatty acids containing 18 carbon atoms were higher in the indoor cultures (69.70% and 71.46% of total FA in 2 and 20 L indoor cultures, respectively) compared to the outdoor culture (55.22% of the total FA). Hexadecadienoic acid (16:2), hiagonic acid (16:3), and linolenic acid (18:3) were higher in the outdoor culture, while oleic acid (18:1) and linoleic acid (18:2) were higher in the indoor cultures. The differences in the fatty acid composition between the indoor and outdoor cultivated cells are likely due to the differences in culture conditions. It has been previously demonstrated that the fatty acid composition varies with temperature and irradiance, and a higher temperature and irradiance level will result in more saturated fatty acids (Olofsson et al., 2012). The highest temperature during the last few days of the outdoor cultivation was less than 20 °C (Supplementary material-Fig. 2), which was cooler than the indoor cultivation (25 °C). The high unsaturated fatty acid content in the outdoor culture was a likely a response to the low temperature; a high unsaturated fatty acid content can enhance membrane fluidity at low temperatures (Thompson et al., 1992). Although the light intensity outdoors at midday was higher than the light intensity for the indoor cultures, the photoperiod was much shorter and the light intensity outdoors had greater fluctuations (Supplementary material-Fig. 2). The increase in the percentage of unsaturated fatty acids in response to the poor irradiance and short photoperiod is consistent with a report by Seyfabadi et al. that unsaturated fatty acids increase as the irradiance and light
duration decrease, and this is accompanied by a decrease in saturated fatty acids (SFA) (Seyfabadi et al., 2011). This is due to a key role of PUFA for acclimation and maintenance of the photosynthetic membrane (Klyachko-Gurvich et al., 1999). The changes in the fatty acid composition of the algal cells may be an adaptive response to the varying environmental conditions (Ogbonda et al., 2007). The protein contents and amino acid compositions of C. ellipsoidea in response to indoor and outdoor cultivation are listed in Table 3. The algal cells cultivated indoors had protein contents as high as 45.43% and 46.65% of the DW, while the protein content in the outdoor culture, 41.75% of DW, was slightly lower. The protein contents in this study were consistent with that reported by Hempel et al. for Chlorella sp. (44.3% of DW) (Hempel et al., 2012). However, the rate of protein production was much lower in the outdoor culture (13.17 mg L1 day1) compared to the indoor cultures (21.67 and 18.59 mg L1 day1 at the 2 and 20 L scale, respectively) due to the low biomass production of the outdoor culture. Several studies have previously demonstrated that a high irradiance level and a long photoperiod help to increase the protein content in Chlorella sp. and other algal species (Ogbonda et al., 2007; Seyfabadi et al., 2011). As previously discussed, it is likely that the short photoperiod and the poor irradiance in the
Table 3 Amino acid and protein of the algal cells cultured in different conditions. Amino acid
Content (% of DW) 2 L indoor
20 L indoor
200 L outdoor
Essential amino acids
Phe Thr Met Val Ile Leu Trp Lys Total
2.04 ± 0.16 1.87 ± 0.11 0.35 ± 0.08 1.88 ± 0.26 1.41 ± 0.09 3.33 ± 0.18 0.30 ± 0.12 2.32 ± 0.18 13.50 ± 0.86
2.11 ± 0.08 1.92 ± 0.28 0.38 ± 0.02 1.93 ± 0.13 1.41 ± 0.09 3.41 ± 0.41 0.54 ± 0.07 2.38 ± 0.25 14.08 ± 1.03
1.91 ± 0.28 1.77 ± 0.13 0.54 ± 0.11 1.98 ± 0.29 1.27 ± 0.15 3.06 ± 0.54 0.43 ± 0.17 1.95 ± 0.22 12.91 ± 1.63
Non-essential amino acids
Ala Gly Tyr Ser His Arg Glu Asp Pro Cys Total
3.12 ± 0.13 2.17 ± 0.12 1.54 ± 0.15 1.76 ± 0.27 0.68 ± 0.09 2.70 ± 0.08 5.94 ± 0.20 3.89 ± 0.16 1.45 ± 0.09 0.60 ± 0.10 23.85 ± 1.82
3.51 ± 0.14 2.18 ± 0.10 1.59 ± 0.28 1.79 ± 0.11 0.67 ± 0.15 2.62 ± 0.17 5.44 ± 0.18 4.02 ± 0.09 1.63 ± 0.05 0.58 ± 0.04 24.03 ± 2.18
3.00 ± 0.49 2.00 ± 0.16 1.34 ± 0.32 1.63 ± 0.06 0.51 ± 0.31 2.34 ± 0.49 3.96 ± 0.28 3.51 ± 0.17 1.40 ± 0.20 0.59 ± 0.13 20.28 ± 2.05
Essential amino acid percentage (% of total fatty acids) Total amino acid content (% of DW) Total protein content (% of DW) Protein productivity (mg L1 day1)
36.14 ± 1.63
36.95 ± 1.83
38.90 ± 3.21
37.35 ± 0.98 a
38.11 ± 1.05 a
33.19 ± 1.53 b
45.43 ± 1.74 a
46.65 ± 1.39 a
41.75 ± 3.25 b
21.67 ± 2.24
18.59 ± 1.98
13.17 ± 2.84
Values in a row with different letters are significantly different (P < 0.05) according to t-test.
S.-K. Wang et al. / Bioresource Technology 156 (2014) 117–122 Table 4 The cost of each cultivation process (US$).
Capital investment Power Medium CO2 Final biomass concentration (g/L) Total volume (L) Total biomass (g) Total cost (US$/batch) Cost (US$/kg DW)
20 L indoor
200 L outdoor
1.478 3.970 0.0264 0.22 0.66 20 13.20 5.694 431.394
1.546 2.212 0.264 2.2 0.53 200 106.00 6.222 58.698
final growth stage resulted in the low protein content of the outdoor culture. However, the amino acid composition showed few differences in response to the different culture conditions, with the exception that the total essential amino acid percentage was slightly higher in the outdoor culture. The abundant amino acid composition included alanine, arginine, aspartic acid, glutamic acid, lysine, glycine, methionine, and leucine (more than 2% of DW), which was in accordance with reports by Li et al. (2013); Hempel et al. (2012). In addition, Hempel et al. found that the relative amounts of single amino acids were very similar when different strains of Chlorella sp. were screened (Hempel et al., 2012). Combined with this study, it can be concluded that the amino acid composition of the algal cells among different Chlorella sp. strains is relatively similar and it is also stable under different culture conditions. There were no differences between the sterilized flanged glass bioreactor (2 L) and the 20 L open polyethylene bag bioreactor on the fatty acid content and composition, protein content, and amino acid composition under the same culture conditions. This indicated that the bioreactor material and the cultivation scale had little influence on the composition of the algal cells. These parameters were primarily affected by the culture conditions. The scale-up of C. ellipsoidea cultivation from the 20 L indoor culture to the 200 L outdoor culture was feasible and helped to decrease the overall cost of the algal biomass. In the large-scale culture, the bioreactor was able to maintain the same level of aeration and mixing efficiency. In addition, compared with a raceway pond, which is most commonly used for the outdoor cultivation of microalgae, the outdoor culture system tested in this study can greatly decrease the evaporation rate and the risk of contamination (Chisti, 2007). This system has the potential to be utilized for the industrial production of other algal species on a large scale. However, the growth and metabolism of the algal cells grown outdoors were greatly affected by the climate. The outdoor cultivation can not operate throughout the year due to the seasonal changed climate. In Beijing, the temperature is too high in summer and too low in winter, and the operation time is only about half year. Therefore, there are geographic restrictions for the site selection. In addition, the fluctuations in climate conditions also affected the continuous quality of the products; this could be avoided by utilizing temperature control techniques and by the use of shading or the addition of artificial light for illumination control.
3.2. Economic analysis of indoor and outdoor for the scale-up cultivation of C. ellipsoidea The total power consumption and costs are listed in Supplementary material-Table 2 and Table 4. The outdoor cultivation required less equipment and consumed less energy because it is unnecessary to control the light and temperature or to ensure sterile conditions. Compared with the indoor cultivation, the cost of producing algal biomass using outdoor cultivation was far less
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expensive, with an estimated cost 7 times lower than the cost of the 20 L indoor cultures. In the 20 L indoor culture, the low-cost bioreactor significantly decreased the capital investment; the capital investment represented only 28% of the total cost, while the cost of power (68% of the total cost) is the main investment. For the outdoor cultivation, the uncontrolled culture conditions and the large culture scale greatly decreased the operational costs. In addition, both the capital investment and power consumption were low, resulting in an overall cost much lower than that of indoor cultivation. However, compared to studies using different algal species (López-Elías et al., 2005; Li et al., 2011; Acién et al., 2012), the cost of biomass production was still high. This was mainly due to the low density of C. ellipsoidea; the final biomass concentration of C. ellipsoidea and C. vulgaris is generally approximately 0.8 g/L (Lam and Lee, 2012; Li et al., 2013), while the final biomass concentration of Chlorella zofingiensis can achieve concentrations of up to 3 g/L (Feng et al., 2012; Zhu et al., 2013). The cost for biomass would decrease to US$10.37/kg DW if a final biomass concentration of 3 g/L can be achieved. In this sense, the outdoor culture system tested in this study has the potential for the industrial production of other algal species on a large scale. This design can also be combined with thermal power stations in order to use flue gas as the CO2 source and the biomass production cost can be decreased by 24.80–26.76% with flue gas (Nagarajan et al., 2013). 4. Conclusion The scale-up cultivation of C. ellipsoidea was studied in bag bioreactors from indoor to outdoor conditions. The microalgae could adapt to the outdoor conditions, although with a lower biomass production. The fatty acid composition was distinct between indoor and outdoor cultures, while the amino acid composition was stable. The cost of outdoor biomass production was far lower compared to the indoor cultures. It demonstrates that the outdoor cultivation system of C. ellipsoidea has the potential for the largescale production of fatty acids or foreign proteins, and will also be used for the cultivation of other algal species for large-scale biomass production. Acknowledgements This work was financially supported by the Major State Basic Research Development Program (973 Project) of China (No. 2011CB200903), the Knowledge Innovation Program of the Chinese Academy of Sciences (No. YZ200947), and the Chinese Academy of Sciences Fellowships for Young International Scientists (No. 2011Y1GA01). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.biortech.2014.01. 023. References Acién, F.G., Fernández, J.M., Magán, J.J., Molina, E., 2012. Production cost of a real microalgae production plant and strategies to reduce it. Biotechnol. Adv. 30, 1344–1353. Bai, L.L., Yin, W.B., Chen, Y.H., Niu, L.L., Sun, Y.R., Zhao, S.M., et al., 2013. A new strategy to produce a defensin: stable production of mutated NP-1 in nitrate reductase deficient Chlorella ellipsoidea. PLoS One 8, e54966. Brennan, L., Owende, P., 2010. Biofuels from microalgae – a review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sustain. Energy Rev. 14, 557–577.
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Chen, Y., Wang, Y.Q., Sun, Y.R., Zhang, L.M., Li, W.B., 2001. Highly efficient expression of rabbit neutrophil peptide-1 gene in Chlorella ellipsoidea cells. Curr. Genet. 39, 365–370. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Chisti, Y., 2013. Constraints to commercialization of algal fuels. J. Biotechnol. 167, 201–214. Doucha, J., Straka, F., Livansky, K., 2005. Utilization of flue gas for cultivation of microalgae (Chlorella sp.) in an outdoor open thin-layer photobioreactor. J. Appl. Phycol. 17, 403–412. Feng, P.Z., Deng, Z.Y., Hu, Z.Y., Fan, L., 2011. Lipid accumulation and growth of Chlorella zofingiensis in flat plate photobioreactors outdoors. Bioresour. Technol. 102, 10577–10584. Feng, P.Z., Deng, Z.Y., Fan, L., Hu, Z.Y., 2012. Lipid accumulation and growth characteristics of Chlorella zofingiensis under different nitrate and phosphate concentrations. J. Biosci. Bioeng. 114, 405–410. Hempel, N., Petrick, I., Behrendt, F., 2012. Biomass productivity and productivity of fatty acids and amino acids of microalgae strains as key characteristics of suitability for biodiesel production. J. Appl. Phycol. 24, 1407–1418. Huo, S.H., Wang, Z.M., Zhu, S.N., Zhou, W.Z., Dong, R.J., Yuan, Z.H., 2012. Cultivation of Chlorella zofingiensis in bench-scale outdoor ponds by regulation of pH using dairy wastewater in winter, South China. Bioresour. Technol. 121, 76–82. Kay, R.A., 1991. Microalgae as food and supplement. Crit. Rev. Food Sci. Nutr. 30, 555–573. Kim, D.H., Kim, Y.T., Cho, J.J., Bae, J.H., Hur, S.B., Hwang, I., et al., 2002. Stable integration and functional expression of flounder growth hormone gene in transformed microalga, Chlorella ellipsoidea. Mar. Biotechnol. 4, 63–73. Klyachko-Gurvich, G., Tsoglin, L.N., Doucha, J., Kopetskii, J., Shebalina, B.I., Semenenko, V.E., 1999. Desaturation of fatty acids as an adaptive response to shifts in light intensity. Physiol. Plant. 107, 240–249. Lam, M.K., Lee, K.T., 2012. Potential of using organic fertilizer to cultivate Chlorella vulgaris for biodiesel production. Appl. Energy 94, 303–308. Li, J., Zhu, D.L., Niu, J.F., Shen, S.D., Wang, G.C., 2011a. An economic assessment of astaxanthin production by large scale cultivation of Haematococcus pluvialis. Biotechnol. Adv. 29, 568–574. Li, C., Yang, H., Li, Y., Cheng, L., Zhang, M., Zhang, L., et al., 2013. Novel bioconversions of municipal effluent and CO2 into protein riched Chlorella vulgaris biomass. Bioresour. Technol. 132, 171–177. Liu, L.L., Wang, Y.Q., Zhang, Y.C., Chen, X.Y., Zhang, P., Ma, S.W., 2013. Development of a new method for genetic transformation of the green alga Chlorella ellipsoidea. Mol. Biotechnol. 54, 211–219.
López-Elías, J.A., Voltolina, D., Enríquez-Ocaña, F., Gallegos-Simental, G., 2005. Indoor and outdoor mass production of the diatom Chaetoceros muelleriin a mexican commercial hatchery. Aquacult. Eng. 33, 181–191. Lynch, J.M., Barbano, D.M., 1999. Kjeldahl nitrogen analysis as a reference method for protein determination in dairy products. J. AOAC Int. 82, 1389–1398. Mata, T.M., Martins, A.A., Caetano, N.S., 2010. Microalgae for biodiesel production and other applications: a review. Renew. Sustain. Energy Rev. 14, 217–232. Nagarajan, S., Chou, S.K., Cao, S., Wu, C., Zhou, Z., 2013. An updated comprehensive techno-economic analysis of algae biodiesel. Bioresour. Technol. 145, 150–156. Ogbonda, K., Aminigo, R.E., Abu, G., 2007. Influence of aeration and lighting on biomass production and protein biosynthesis in a Spirulina sp. isolated from an oil-polluted brackish water marsh in the Niger Delta. Nigeria. Afr. J. Biotechnol. 6, 2596–2600. Olofsson, M., Lamela, T., Nilsson, E., Bergé, J.P., del Pino, V., Uronen, P., et al., 2012. Seasonal variation of lipids and fatty acids of the microalgae Nannochloropsis oculata grown in outdoor large-scale photobioreactors. Energies 5, 1577–1592. Potvin, G., Zhang, Z.S., 2010. Strategies for high-level recombinant protein expression in transgenic microalgae: a review. Biotechnol. Adv. 28, 910–918. Seyfabadi, J., Ramezanpour, Z., Amini Khoeyi, Z., 2011. Protein, fatty acid, and pigment content of Chlorella vulgaris under different light regimes. J. Appl. Phycol. 23, 721–726. Specht, E., Miyake-Stoner, S., Mayfield, S., 2010. Micro-algae come of age as a platform for recombinant protein production. Biotechnol. Lett. 32, 1373–1383. Thompson, P.A., Guo, M.X., Harrison, P.J., Whyte, J.N.C., 1992. Effects of variation in temperature II. On the fatty-acid composition of eight species of marine phytoplankton. J. Phycol. 28, 488–497. Walker, T.L., Purton, S., Becker, D.K., Collet, C., 2005. Microalgae as bioreactors. Plant Cell Rep. 24, 629–641. Wang, P., Wang, Y.Q., Geng, D.G., Li, W.B., Sun, Y.R., 2003. Molecular cloning and characterization of a cDNA encoding nitrate reductase from Chlorella ellipsoidea (Chlorophyta). J. Appl. Phycol. 15, 457–463. Xu, H., Miao, X., Wu, Q., 2006. High quality biodiesel production from microalga Chlorella protothecoides by heterotrophic growth in fermenters. J. Biotechnol. 126, 499–507. Zhou, X.P., Xia, L., Ge, H.M., Zhang, D.L., Hu, C.X., 2013. Feasibility of biodiesel production by microalgae Chlorella sp. (FACHB-1748) under outdoor conditions. Bioresour. Technol. 138, 131–135. Zhu, L.D., Wang, Z.M., Takala, J., Hiltunen, E., Qin, L., Xu, Z.B., et al., 2013. Scale-up potential of cultivating Chlorella zofingiensis in piggery wastewater for biodiesel production. Bioresour. Technol. 137, 318–325.
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Scale-up cultivation of Chlorella ellipsoidea from indoor to outdoor in bubble column bioreactors Shi-Kai Wang a,c, Yi-Ru Hu a,c, Feng Wang a, Amanda R. Stiles b, Chun-Zhao Liu a,⇑ a National Key Laboratory of Biochemical Engineering & Key Laboratory of Green Process and Engineering, Institute of Process Engineering, Chinese Academy of Sciences, Beijing 100190, PR China b Department of Plant and Microbial Biology, University of California, Berkeley, CA 94720, USA c University of Chinese Academy of Sciences, Beijing 100049, PR China
h i g h l i g h t s Chlorella ellipsoidea cells were cultivated in bubble column bioreactors. Microalgal cells were able to quickly adapt to the outdoor conditions. Biomass production cost in outdoor culture was lower than that in indoor culture.
a r t i c l e
i n f o
Article history: Received 28 November 2013 Received in revised form 4 January 2014 Accepted 6 January 2014 Available online 17 January 2014 Keywords: Chlorella ellipsoidea Bubble column bioreactor Outdoor cultivation Fatty acid composition Economic evaluation
a b s t r a c t The cultivation of Chlorella ellipsoidea in bubble column bioreactors was investigated at different scales under indoor and outdoor conditions. The algal cells were able to quickly adapt to the outdoor conditions and achieved a growth rate of 31.55 mg L1 day1. Due to differences in light and temperature, the outdoor culture produced a higher percentage of unsaturated fatty acids compared to the indoor cultures, while the amino acid composition was unaffected. The overall cost of the biomass produced by the 200 L outdoor cultivation (58.70 US$/kg-dry weight) was estimated to be more than 7 times lower than that of the 20 L indoor cultivation (431.39 US$/kg-dry weight). Together these results provide a basis for the cultivation of C. ellipsoidea for the large-scale production of biofuels, high-value nutrients and/or recombinant proteins. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Microalgae are photoautotrophic sunlight-driven cell factories that can convert carbon dioxide to various products such as lipids, carbohydrates, proteins, fatty acids, vitamins, antibiotics, and antioxidants (Chisti, 2007). In addition to these naturally-produced compounds, microalgae are also a promising platform for the production of recombinant proteins. Microalgae have unique advantages, including a rapid growth rate, ease of cultivation, and the ability to make the same post-transcriptional and post-translational modifications as plants (Potvin and Zhang, 2010). The feasibility of utilizing microalgae as a production system for therapeutic or industrial proteins has been previously demonstrated (Specht et al., 2010). Chlorella ellipsoidea is a single-celled eukaryotic green algae that has been extensively studied. As with other Chlorella sp,
⇑ Corresponding author. Tel./fax: +86 10 82622280. E-mail address: [email protected] (C.-Z. Liu). 0960-8524/$ - see front matter Ó 2014 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2014.01.023
it can be found in both fresh and marine water systems, and it can be cultured under autotrophic, heterotrophic, mixotrophic, or photoheterotrophic growth conditions (Mata et al., 2010). It is rich in high-quality proteins, vitamins, lipid-soluble compounds, glycolipids, sulfolipids, and compounds used as food additives. It is also beneficial for human health due to its ability to lower blood sugar levels and increase hemoglobin concentrations, and it is used to enhance animal growth when it is applied as an aquaculture or animal feed (Mata et al., 2010; Kay, 1991). Furthermore, C. ellipsoidea is a well-established model organism and a promising bioreactor for the production of complex foreign proteins for pharmaceutical and industrial use (Wang et al., 2003; Walker et al., 2005). The genetic transformation of C. ellipsoidea is feasible, and it has been previously utilized for the recombinant production of flounder growth hormone (FGH) and rabbit neutrophil peptide-1 (NP-1) (Liu et al., 2013; Kim et al., 2002; Chen et al., 2001; Bai et al., 2013). For the industrial production of products from microalgae, it is necessary to evaluate the feasibility of scaled-up outdoor culture systems. The utilization of natural sunlight and uncontrolled
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outdoor culture conditions can reduce the overall costs of the cultivation process (Feng et al., 2011). Raceway ponds and other open-culture systems are commonly used for large-scale outdoor cultivation due to their low costs for construction and operation; however, they often perform poorly due to contamination risk, lack of control of the culture conditions, and problems with mixing and light utilization efficiency (Mata et al., 2010). Closed bioreactors can provide a suitable environment in terms of light, nutrients, CO2, and temperature, and they have the potential to produce higher rates of biomass production and improved culture quality (Brennan and Owende, 2010). The feasibility of Chlorella sp. outdoor cultivation has been confirmed by many studies using different types of bioreactors at scales ranging from 10 to 70 L, including a flat glass plate bioreactor, a cylindrical glass bottle bioreactor, and a nylon membrane column bioreactor. The results have demonstrated that many Chlorella strains can adapt to the fluctuating temperatures and irradiance levels inherent in outdoor cultivation, and that these systems can be used for biodiesel production (Feng et al., 2011, 2012; Zhou et al., 2013). In addition, the outdoor production of Chlorella sp. can also utilize the waste produced by other processes for nutrients, such as organic fertilizer derived from food waste or manure, dairy and piggery wastewater, or flue gas generated by the combustion of natural gas (Chisti, 2013). This can decrease the cost of the cultivation process (Doucha et al., 2005; Lam and Lee, 2012; Huo et al., 2012; Zhu et al., 2013). However, the growth rate is often reduced by outdoor cultivation; for example, the growth rate of Calluna vulgaris was reduced by 27% in a study by Lam and Lee comparing outdoor cultivation with cultivation in a controlled environment (Lam and Lee, 2012). In addition, the lipid content and fatty acid composition also varied in response to climatic variation (Olofsson et al., 2012), which may influence the quality of the final products. Although there are numerous reports describing the outdoor cultivation of microalgae, studies focusing on C. ellipsoidea are rare and the culture scale in published studies is limited. It is important to understand the variations in growth and metabolism between indoor and outdoor cultivation conditions in order to determine the feasibility of outdoor cultivation for this species. In the current study, a 200 L outdoor cultivation system was constructed using a bubble column bioreactor made of a polyethylene membrane. The cultivation of C. ellipsoidea in bubble column bioreactors was scaled-up from the 20 L indoor culture to a 200 L outdoor culture, and the biomass, fatty acid content and composition, protein content, and amino acid composition were investigated and compared with that of 2 L sterilized flanged glass bioreactor. The effect of controlled and uncontrolled light intensity and temperature on C. ellipsoidea’s growth and metabolism was analyzed, and the cost of each cultivation system was calculated.
constructed from transparent polyethylene (PE) bag with a thickness of 0.2 mm (Yangpu Packaging Material Co., Ltd., Hebei, China). The bottom of the plastic culture bags with the desired dimensions was heat-sealed by a SF-B 800 pedal sealing machine (Xingye Machinery Equipment Co., Ltd., Wenzhou, China). The size of all bioreactors was shown in Supplementary material-Table 1. Air was supplied using an air sparger (compressed stainless steel particles) at the bottom of each bioreactor. Each 20 L bioreactor was handed at a shelf, and each 200 L bioreactor was supported by a circular wire mesh. The indoor cultivations were performed at 25 °C under a light/ dark cycle of 16/8 h with 25,000 lux illumination intensity, and the outdoor cultivations were conducted under the natural temperature and light conditions in Haidian district of Beijing, China (latitude 39°590 N, longitude 116°190 E), in autumn (September 29th, 2012 to October 19th, 2012). The initial biomass concentration was 0.075 g/L, and CO2-enriched air (1%) was bubbled into the reactor at a flow rate of 0.1 v/v/min through a 0.22 lm filter (Millipore, MA, USA) for the 2 L indoor cultivation and directly aerated into the reactor for the 20 L indoor and 200 L outdoor cultures. For the 2 L indoor culture, the reactor and the air supply systems were sterilized by autoclaving at 121 °C for 40 min prior to starting the cultivation. 2.2. Microalgae growth measurement To determine the relationship between the optical density at 680 nm (OD680) and the algal biomass (g/L), the algal culture broth was diluted or concentrated to a certain OD680, then harvested by centrifugation for 5 min at 10,000 rpm, 4 °C. The algal pellets were then washed three times with distilled water and dried at 105 °C to a constant weight. The dry weight (DW, g/L) of the algal cells was determined gravimetrically. The relationship between the OD680 and the C. ellipsoidea biomass was as follows:
DWðg=LÞ ¼ 0:38352 OD680 0:02647 ðR2 ¼ 0:9996Þ
The algal growth was measured every two days by measuring the OD680 of the culture broth using a UV-2100 spectrophotometer (Unico, Shanghai, China) and the biomass was calculated using Eq. (1). The maximum specific growth rate (lmax, day1) at the exponential stage was calculated as follows:
lmax ðday1 Þ ¼ ðln DW2 ln DW1 Þ=ðt2 t1 Þ
ð2Þ
where DW1 and DW2 were the dry biomass weight (g/L) at time t1 and t2, respectively. The doubling time (TD, days) was calculated as follows:
T D ðdaysÞ ¼ lnð2Þ=lmax 2. Methods 2.1. Microalgae strain and culture systems
ð3Þ 1
The rate of biomass production (P, mg L according to the following equation: 1
C. ellipsoidea (UTEX 20), kindly provided by Prof. Zan-Min Hu (Institute of Genetics and Developmental Biology, Chinese Academy of Sciences), was grown on BG-11 medium. The cells were maintained in 250 mL Erlenmeyer flasks containing 100 mL liquid medium and incubated at 25 °C in an orbital shaker at 100 rpm. Algal cells were grown photoautotrophically under a 16 h light period per day with 35 lmol m2 s1 for 15 days. The pre-cultured cells were used as the inoculum for all bubble-column cultivation experiments. The 2 L bubble column bioreactor was constructed from a flanged glass column (Beijing Glass Group Company, Beijing, China), and the 20 and 200 L bubble column bioreactors were
ð1Þ
Pðmg L1 day Þ ¼ ðDWx DW0 Þ=t x
1
day
) was calculated
ð4Þ
where DW0 and DWx were the initial dry biomass weight (mg/L) and the dry biomass weight (mg/L) at time tx, respectively. 2.3. Lipid analysis For the extraction and transesterification of the fatty acids, 50 mg lyophilized algal biomass was ground for 10 min and then dispersed in 3 mL of a 7.5% (w/v) KOH/CH3OH solution with 200 mg of heptadecanoic acid added as an internal standard. After the solution was incubated at 70 °C for 4 h, 2 mL HCl/CH3OH (1:1, v/v) and 2 mL 14% BF3/CH3OH (ANPEL, Shanghai, China) were
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2.4. Algal protein and amino acid analysis The total intracellular protein was determined using the Kjeldahl method as described by Lynch and Barbano (Lynch and Barbano, 1999), and the total algal protein content was calculated by multiplying by a factor of 6.38, a value previously determined for the measurement of Chlorella sp. protein content (Li et al., 2013). For the amino acid composition analysis, the protein was hydrolyzed using 4.2 M NaOH for tryptophan, performic acid for cysteine, and 6 M HCl for the other amino acids. The hydrolyzed amino acid samples were separated using ion exchange chromatography and quantified using an automatic amino acid analyzer (Hitachi L-8900, Japan).
2.5. Measurement of climatic conditions The light intensity and culture temperature were recorded at 8:00, 12:00, 16:00, and 20:00 every two days. The light intensity was measured using a JD-3 luxmeter (Jiading Xuelian Instrument Factory, Shanghai, China), and the temperature was measured using a spirit thermometer. The samples were collected every two days at 9:00 am.
2.6. Economic assumptions The total cost of the cultivation process includes both a fixed capital investment and operational costs (Li et al., 2011). In this study, the fixed capital investments are listed in Supplementary material-Table 2. The costs were calculated based on one batch cultivation and the estimated lifetime of the equipment. The bioreactors used for the 20 and 200 L cultivations utilized a single-use PE membrane which cost US$0.33/m (width: 0.75 m, double layer) resulting in a cost of US$0.165 and US$0.654 for the 20 and 200 L cultivation, respectively. The shelf used for the outdoor culture cost US$130 with an estimated lifetime of 10 years. The cost of the air compressor used for the 20 L indoor cultivation was US$32.68 with an estimated lifetime of 5 years of continuous operation. The cost of the air compressor used for the outdoor cultivation was US$620.9 and estimated to last for 7 years. The light used for the indoor cultivation was US$1.63 with an estimated lifetime of 3000 h under continuous use. Four lights were utilized as a unit to illuminate for a 20 L bioreactor. The investment capital and the power consumption of the cooling system for the indoor cultivations were based on the ratio of the bioreactor volume to the total volume of the room. The electricity consumption was calculated based on the power of the instruments and their operation time. The price of the electricity and water was based on the local price (US$0.078/kW h and US$0.65/ton, respectively), and the cost of the medium was estimated based on the market price at Fang-Dou chemical web (http://www.16ds.com/). The local price for CO2 was US$0.245/kg.
3. Results and discussion 3.1. Scale-up cultivation of C. ellipsoidea from indoor to outdoor After a two day lag period, the algal cells were able to survive and adapt to the outdoor conditions in the 200 L bioreactor (Supplementary material-Fig. 1). Following the lag period, the cells entered the exponential phase, and they reached the stationary phase at 14 days (Fig. 1). As shown in Table 1, the maximum specific growth rates (lmax) of the indoor cultures at the exponential stage (0.168 and 0.162 day1 for the 2 and 20 L cultures, respectively) were higher than that of the outdoor culture (0.145 day1), and this was accompanied by a shorter doubling time (4.13 and 4.28 days for the indoor cultures versus 4.78 days for the outdoor culture). This resulted in a rate of biomass production as high as 47.71 and 39.85 mg L1 day1 in the indoor cultures while the biomass production rate of the outdoor culture reached only 31.55 mg L1 day1 (Table 1). Similar results have been reported by Zhu et al. (2013). This different in growth rate is due to the fact that the algal cells cultured indoors are cultured in optimal controlled conditions rather than the uncontrolled outdoor natural environment. As illustrated in Supplementary material-Fig. 2, the varied weather conditions in the uncontrolled outdoor environment resulted in dramatic fluctuations in the temperature and light intensity during cultivation, and the natural day length (approximately 1213 h day1) was shorter than the illumination time applied to the indoor culture (16 h day1). In addition, algae cultivated outdoors are more vulnerable to contamination by foreign microalgae and microorganisms than the indoor cultures, which may also have a negative impact on the algal growth. The fatty acid (FA) compositions of C. ellipsoidea cultures under indoor and outdoor conditions are listed in Table 2. There were eight to nine detectable fatty acids, and the fatty acids with 16 and 18 carbon atoms accounted for more than 99% of the total fatty acids. Fatty acids with 16 and 18 carbon atoms are the ideal components for biodiesel, indicating that C. ellipsoidea could function as a potential species for the production of biodiesel (Xu et al., 2006). In addition, linolenic acid (18:3), which is beneficial for the treatment of cardiovascular diseases (Huo et al., 2012), was high in all culture conditions (more than 15% of the total FA), especially in the outdoor cultivation (as high as 40.26% of total FA).
0.8
0.6
Biomass (g/L)
added and transesterification occurred at 70 °C for 1.5 h. The mixture was centrifuged at 2000 rpm for 3 min at 4 °C following the addition of 1 mL 0.9% NaCl and 4 mL n-hexane. The organic phase was volatilized under a nitrogen atmosphere and re-dissolved in 4 mL chloroform for GC analysis. The fatty acid methyl esters (FAMEs) were analyzed using a gas chromatogarph equipped with an FID detector (Agilent, 7890 A, America) and an Agilent DB-17 column (30 m 0.25 mm). The injector and detector temperatures were 250 °C. The oven temperature was set at 180 °C for 1 min, increased from 180 to 230 °C at a rate of 6 °C/min, and then remained at 230 °C for 25 min.
0.4
2 L Indoor 20 L Indoor 200 L Outdoor
0.2
0.0
0
4
8 Time (day)
12
16
Fig. 1. The growth curve of C. ellipsoidea under different cultivation conditions.
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Table 1 Growth kinetic parameters of the algal cells under different culture conditions. Culture condition
Specific growth rate lmax (day1)
Doubling time (days)
Final biomass concentration (g L1)
Rate of biomass production (mg L1 day1)
2 L Indoor 20 L Indoor 200 L Outdoor
0.168 ± 0.006 0.162 ± 0.011 0.145 ± 0.026
4.13 ± 0.04 4.28 ± 0.15 4.78 ± 0.24
0.77 ± 0.05 0.66 ± 0.04 0.53 ± 0.10
47.71 ± 0.9 39.85 ± 1.6 31.55 ± 7.5
Values in a column are significantly different (P < 0.05) from each other according to t-test.
Table 2 The fatty acid composition of the algal cells cultured in different conditions. FA composition
14:0 16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3 C16 C18 SFA MUFA PUFA Total (% of DW)
FA content (% of total FA) 2 L indoor
20 L indoor
200 L outdoor
0.97 ± 0.09 19.68 ± 1.23 – 4.95 ± 0.11 4.71 ± 0.88 1.61 ± 0.05 10.60 ± 1.34 38.12 ± 2.04 19.37 ± 0.99 29.34 ± 0.08 69.70 ± 1.22 22.26 ± 0.92 10.60 ± 1.09 67.15 ± 1.78 8.94 ± 0.21
0.92 ± 0.11 18.64 ± 1.09 0.92 ± 0.10 3.46 ± 1.08 4.59 ± 0.15 1.94 ± 0.03 10.59 ± 0.20 41.99 ± 1.75 16.94 ± 0.68 27.61 ± 0.38 71.46 ± 1.15 21.50 ± 0.35 11.51 ± 0.77 66.98 ± 1.11 8.71 ± 0.22
0.99 ± 0.14 14.62 ± 1.50 2.14 ± 1.23 11.12 ± 0.88 14.77 ± 2.27 2.18 ± 0.54 2.77 ± 0.99 10.00 ± 1.74 40.26 ± 1.97 42.64 ± 0.80 55.22 ± 1.63 17.79 ± 1.84 4.90 ± 1.93 76.15 ± 2.83 7.44 ± 0.58
a a a a a a
a a a a a a
b b b b b b
Values in a row with different letters are significantly different (P < 0.05) according to t-test.
There were no obvious differences in the FA composition between the 2 and 20 L indoor cultures. However, the FA composition of the outdoor culture was distinct from the indoor cultures. Similar to a report by Zhu et al. (2013), the proportion of unsaturated fatty acids (including monounsaturated fatty acids (MUFA) and polyunsaturated fatty acids (PUFA)) in the algal cells cultivated outdoors was higher than that from the indoor cultures, especially the PUFA. In addition, fatty acids containing 16 carbon atoms were higher in the outdoor culture (42.64% of the total FA) compared to the indoor cultures (29.34% and 27.61% of the total FA for the 2 and 20 L cultures, respectively). Fatty acids containing 18 carbon atoms were higher in the indoor cultures (69.70% and 71.46% of total FA in 2 and 20 L indoor cultures, respectively) compared to the outdoor culture (55.22% of the total FA). Hexadecadienoic acid (16:2), hiagonic acid (16:3), and linolenic acid (18:3) were higher in the outdoor culture, while oleic acid (18:1) and linoleic acid (18:2) were higher in the indoor cultures. The differences in the fatty acid composition between the indoor and outdoor cultivated cells are likely due to the differences in culture conditions. It has been previously demonstrated that the fatty acid composition varies with temperature and irradiance, and a higher temperature and irradiance level will result in more saturated fatty acids (Olofsson et al., 2012). The highest temperature during the last few days of the outdoor cultivation was less than 20 °C (Supplementary material-Fig. 2), which was cooler than the indoor cultivation (25 °C). The high unsaturated fatty acid content in the outdoor culture was a likely a response to the low temperature; a high unsaturated fatty acid content can enhance membrane fluidity at low temperatures (Thompson et al., 1992). Although the light intensity outdoors at midday was higher than the light intensity for the indoor cultures, the photoperiod was much shorter and the light intensity outdoors had greater fluctuations (Supplementary material-Fig. 2). The increase in the percentage of unsaturated fatty acids in response to the poor irradiance and short photoperiod is consistent with a report by Seyfabadi et al. that unsaturated fatty acids increase as the irradiance and light
duration decrease, and this is accompanied by a decrease in saturated fatty acids (SFA) (Seyfabadi et al., 2011). This is due to a key role of PUFA for acclimation and maintenance of the photosynthetic membrane (Klyachko-Gurvich et al., 1999). The changes in the fatty acid composition of the algal cells may be an adaptive response to the varying environmental conditions (Ogbonda et al., 2007). The protein contents and amino acid compositions of C. ellipsoidea in response to indoor and outdoor cultivation are listed in Table 3. The algal cells cultivated indoors had protein contents as high as 45.43% and 46.65% of the DW, while the protein content in the outdoor culture, 41.75% of DW, was slightly lower. The protein contents in this study were consistent with that reported by Hempel et al. for Chlorella sp. (44.3% of DW) (Hempel et al., 2012). However, the rate of protein production was much lower in the outdoor culture (13.17 mg L1 day1) compared to the indoor cultures (21.67 and 18.59 mg L1 day1 at the 2 and 20 L scale, respectively) due to the low biomass production of the outdoor culture. Several studies have previously demonstrated that a high irradiance level and a long photoperiod help to increase the protein content in Chlorella sp. and other algal species (Ogbonda et al., 2007; Seyfabadi et al., 2011). As previously discussed, it is likely that the short photoperiod and the poor irradiance in the
Table 3 Amino acid and protein of the algal cells cultured in different conditions. Amino acid
Content (% of DW) 2 L indoor
20 L indoor
200 L outdoor
Essential amino acids
Phe Thr Met Val Ile Leu Trp Lys Total
2.04 ± 0.16 1.87 ± 0.11 0.35 ± 0.08 1.88 ± 0.26 1.41 ± 0.09 3.33 ± 0.18 0.30 ± 0.12 2.32 ± 0.18 13.50 ± 0.86
2.11 ± 0.08 1.92 ± 0.28 0.38 ± 0.02 1.93 ± 0.13 1.41 ± 0.09 3.41 ± 0.41 0.54 ± 0.07 2.38 ± 0.25 14.08 ± 1.03
1.91 ± 0.28 1.77 ± 0.13 0.54 ± 0.11 1.98 ± 0.29 1.27 ± 0.15 3.06 ± 0.54 0.43 ± 0.17 1.95 ± 0.22 12.91 ± 1.63
Non-essential amino acids
Ala Gly Tyr Ser His Arg Glu Asp Pro Cys Total
3.12 ± 0.13 2.17 ± 0.12 1.54 ± 0.15 1.76 ± 0.27 0.68 ± 0.09 2.70 ± 0.08 5.94 ± 0.20 3.89 ± 0.16 1.45 ± 0.09 0.60 ± 0.10 23.85 ± 1.82
3.51 ± 0.14 2.18 ± 0.10 1.59 ± 0.28 1.79 ± 0.11 0.67 ± 0.15 2.62 ± 0.17 5.44 ± 0.18 4.02 ± 0.09 1.63 ± 0.05 0.58 ± 0.04 24.03 ± 2.18
3.00 ± 0.49 2.00 ± 0.16 1.34 ± 0.32 1.63 ± 0.06 0.51 ± 0.31 2.34 ± 0.49 3.96 ± 0.28 3.51 ± 0.17 1.40 ± 0.20 0.59 ± 0.13 20.28 ± 2.05
Essential amino acid percentage (% of total fatty acids) Total amino acid content (% of DW) Total protein content (% of DW) Protein productivity (mg L1 day1)
36.14 ± 1.63
36.95 ± 1.83
38.90 ± 3.21
37.35 ± 0.98 a
38.11 ± 1.05 a
33.19 ± 1.53 b
45.43 ± 1.74 a
46.65 ± 1.39 a
41.75 ± 3.25 b
21.67 ± 2.24
18.59 ± 1.98
13.17 ± 2.84
Values in a row with different letters are significantly different (P < 0.05) according to t-test.
S.-K. Wang et al. / Bioresource Technology 156 (2014) 117–122 Table 4 The cost of each cultivation process (US$).
Capital investment Power Medium CO2 Final biomass concentration (g/L) Total volume (L) Total biomass (g) Total cost (US$/batch) Cost (US$/kg DW)
20 L indoor
200 L outdoor
1.478 3.970 0.0264 0.22 0.66 20 13.20 5.694 431.394
1.546 2.212 0.264 2.2 0.53 200 106.00 6.222 58.698
final growth stage resulted in the low protein content of the outdoor culture. However, the amino acid composition showed few differences in response to the different culture conditions, with the exception that the total essential amino acid percentage was slightly higher in the outdoor culture. The abundant amino acid composition included alanine, arginine, aspartic acid, glutamic acid, lysine, glycine, methionine, and leucine (more than 2% of DW), which was in accordance with reports by Li et al. (2013); Hempel et al. (2012). In addition, Hempel et al. found that the relative amounts of single amino acids were very similar when different strains of Chlorella sp. were screened (Hempel et al., 2012). Combined with this study, it can be concluded that the amino acid composition of the algal cells among different Chlorella sp. strains is relatively similar and it is also stable under different culture conditions. There were no differences between the sterilized flanged glass bioreactor (2 L) and the 20 L open polyethylene bag bioreactor on the fatty acid content and composition, protein content, and amino acid composition under the same culture conditions. This indicated that the bioreactor material and the cultivation scale had little influence on the composition of the algal cells. These parameters were primarily affected by the culture conditions. The scale-up of C. ellipsoidea cultivation from the 20 L indoor culture to the 200 L outdoor culture was feasible and helped to decrease the overall cost of the algal biomass. In the large-scale culture, the bioreactor was able to maintain the same level of aeration and mixing efficiency. In addition, compared with a raceway pond, which is most commonly used for the outdoor cultivation of microalgae, the outdoor culture system tested in this study can greatly decrease the evaporation rate and the risk of contamination (Chisti, 2007). This system has the potential to be utilized for the industrial production of other algal species on a large scale. However, the growth and metabolism of the algal cells grown outdoors were greatly affected by the climate. The outdoor cultivation can not operate throughout the year due to the seasonal changed climate. In Beijing, the temperature is too high in summer and too low in winter, and the operation time is only about half year. Therefore, there are geographic restrictions for the site selection. In addition, the fluctuations in climate conditions also affected the continuous quality of the products; this could be avoided by utilizing temperature control techniques and by the use of shading or the addition of artificial light for illumination control.
3.2. Economic analysis of indoor and outdoor for the scale-up cultivation of C. ellipsoidea The total power consumption and costs are listed in Supplementary material-Table 2 and Table 4. The outdoor cultivation required less equipment and consumed less energy because it is unnecessary to control the light and temperature or to ensure sterile conditions. Compared with the indoor cultivation, the cost of producing algal biomass using outdoor cultivation was far less
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expensive, with an estimated cost 7 times lower than the cost of the 20 L indoor cultures. In the 20 L indoor culture, the low-cost bioreactor significantly decreased the capital investment; the capital investment represented only 28% of the total cost, while the cost of power (68% of the total cost) is the main investment. For the outdoor cultivation, the uncontrolled culture conditions and the large culture scale greatly decreased the operational costs. In addition, both the capital investment and power consumption were low, resulting in an overall cost much lower than that of indoor cultivation. However, compared to studies using different algal species (López-Elías et al., 2005; Li et al., 2011; Acién et al., 2012), the cost of biomass production was still high. This was mainly due to the low density of C. ellipsoidea; the final biomass concentration of C. ellipsoidea and C. vulgaris is generally approximately 0.8 g/L (Lam and Lee, 2012; Li et al., 2013), while the final biomass concentration of Chlorella zofingiensis can achieve concentrations of up to 3 g/L (Feng et al., 2012; Zhu et al., 2013). The cost for biomass would decrease to US$10.37/kg DW if a final biomass concentration of 3 g/L can be achieved. In this sense, the outdoor culture system tested in this study has the potential for the industrial production of other algal species on a large scale. This design can also be combined with thermal power stations in order to use flue gas as the CO2 source and the biomass production cost can be decreased by 24.80–26.76% with flue gas (Nagarajan et al., 2013). 4. Conclusion The scale-up cultivation of C. ellipsoidea was studied in bag bioreactors from indoor to outdoor conditions. The microalgae could adapt to the outdoor conditions, although with a lower biomass production. The fatty acid composition was distinct between indoor and outdoor cultures, while the amino acid composition was stable. The cost of outdoor biomass production was far lower compared to the indoor cultures. It demonstrates that the outdoor cultivation system of C. ellipsoidea has the potential for the largescale production of fatty acids or foreign proteins, and will also be used for the cultivation of other algal species for large-scale biomass production. Acknowledgements This work was financially supported by the Major State Basic Research Development Program (973 Project) of China (No. 2011CB200903), the Knowledge Innovation Program of the Chinese Academy of Sciences (No. YZ200947), and the Chinese Academy of Sciences Fellowships for Young International Scientists (No. 2011Y1GA01). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.biortech.2014.01. 023. References Acién, F.G., Fernández, J.M., Magán, J.J., Molina, E., 2012. Production cost of a real microalgae production plant and strategies to reduce it. Biotechnol. Adv. 30, 1344–1353. Bai, L.L., Yin, W.B., Chen, Y.H., Niu, L.L., Sun, Y.R., Zhao, S.M., et al., 2013. A new strategy to produce a defensin: stable production of mutated NP-1 in nitrate reductase deficient Chlorella ellipsoidea. PLoS One 8, e54966. Brennan, L., Owende, P., 2010. Biofuels from microalgae – a review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sustain. Energy Rev. 14, 557–577.
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