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Chemo-enzymatic saccharification and bioethanol fermentation of lipid-extracted residual biomass of the microalga, Dunaliella tertiolecta
Bioresource Technology 132 (2013) 197–201
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Chemo-enzymatic saccharification and bioethanol fermentation of lipid-extracted residual biomass of the microalga, Dunaliella tertiolecta Ok Kyung Lee a, A Leum Kim a, Dong Ho Seong b, Choul Gyun Lee b, Yeon Tae Jung c, Jin Won Lee c, Eun Yeol Lee a,⇑ a b c
Department of Chemical Engineering, Kyung Hee University, Gyeonggi-do 446-701, Republic of Korea Marine Bioenergy Research Center, Department of Biological Engineering, Inha University, Incheon 402-751, Republic of Korea Department of Chemical and Biomolecular Engineering, Sogang University, Seoul 121-742, Republic of Korea
h i g h l i g h t s " The residual biomass of Dunaliella tertiolecta after lipid extraction was saccharified and used for bioethanol fermentation. " A pretreatment procedure was not required for enzymatic saccharification of the residual biomass. " Saccharification yield based on the total amount of carbohydrates was 80.9% (w/w). " Bioethanol was directly produced with 82% yield from the saccharification solution without additional pretreatment. " The waste residual biomass generated during microalgal biodiesel production could be used for the production of bioethanol.
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Article history: Received 14 August 2012 Received in revised form 29 December 2012 Accepted 4 January 2013 Available online 19 January 2013 Keywords: Bioethanol fermentation Dunaliella tertiolecta Reducing sugar Residual biomass Saccharification
a b s t r a c t Chemo-enzymatic saccharification and bioethanol fermentation of the residual biomass of Dunaliella tertiolecta after lipid extraction for biodiesel production were investigated. HCl-catalyzed saccharification of the residual biomass at 121 °C for 15 min produced reducing sugars with a yield of 29.5% (w/w) based on the residual biomass dry weight. Various enzymes were evaluated for their ability to saccharify the residual biomass. Enzymatic saccharification using AMG 300L produced 21.0 mg/mL of reducing sugar with a yield of 42.0% (w/w) based on the residual biomass at pH 5.5 and 55 °C. Bioethanol was produced from the enzymatic saccharification products without additional pretreatment by Saccharomyces cerevisiae with yields of 0.14 g ethanol/g residual biomass and 0.44 g ethanol/g glucose produced from the residual biomass. The waste residual biomass generated during microalgal biodiesel production could be used for the production of bioethanol to improve the economic feasibility of microalgal biorefinery. Ó 2013 Elsevier Ltd. All rights reserved.
1. Introduction The depletion of fossil fuels has triggered the development of alternative renewable biofuels, such as bioethanol and biodiesel (Lin and Tanaka, 2006; Meher et al., 2006). Various types of biomass have been considered as feedstock for biofuel production (Demirbas, 2006). Starch and sugarcane have been used for commercial-scale production of bioethanol, i.e., first generation biomass. However, crop-based first generation bioethanol causes moral issues of using food for fuel production. Second generation bioethanol uses various cellulosic materials as feedstock. Despite intensive research on cellulosic bioethanol, commercial plants do not exist due to the difficulty and complexity of lignin pretreat-
⇑ Corresponding author. Tel.: +82 31 201 3839; fax: +82 31 204 8114. E-mail address: [email protected] (E.Y. Lee). 0960-8524/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2013.01.007
ment. One of the critical success factors of microbial bioethanol production from biomass is cost-competitiveness of the saccharification process. To address these technical challenges, much effort has been given to the development of cost-effective pretreatment and saccharification processes for cellulosic biomass (Nigam and Singh, 2011). As the third generation biomass, microalgae and macroalgae have been considered as feedstock for biofuel production based on the expectation that large amounts of biomass will soon be available at an acceptable cost (John et al., 2011; Waltz, 2009). Biofuel production using algae has attracted much attention because it can be cultured using CO2 and sunlight (Singh et al., 2011). Another advantage of microalgae and macroalgae is that they do not have lignin, allowing development of a cost-effective pretreatment process (Libessart et al., 1995; Miranda et al., 2012). Microalgae is good feedstock for biodiesel production because it accumulates a high percentage of lipids on the basis of dry cell
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mass (Nigam and Singh, 2011; Wahlen et al., 2011; Zou et al., 2009). Recently, we developed a large-scale cultivation method for the marine microalgae, Dunaliella tertiolecta LB999, with a photobioreactor using a semi-permeable membrane for the production of biodiesel (Kim and Lee, 2010). During the production of biodiesel from D. tertiolecta LB999 biomass, a significant amount of residual biomass is generated as waste after lipid extraction. Biodiesel production from microalgae still suffers from high production cost. Most of the valuable cell components of microalgae need to be utilized to enhance the economic feasibility of microalgal biorefinery. The residual biomass can be used for the production of bioethanol because it has a large amount of polysaccharide. Many investigations on saccharification and bioethanol fermentation from microalgae and macroalgae biomass have been performed (Choi et al., 2010; Harun et al., 2010, 2011; Harun and Danquah, 2011; Kim et al., 2011; Lee et al., 2011; Wargacki et al., 2012). However, investigations on saccharification and bioethanol fermentation of the microalgal residual biomass after lipid extraction have not been reported. The main objective of this study is to produce bioethanol from the residual biomass of D. tertiolecta LP999 resulted from lipid extraction.
units (AGU)/mL and 100 fungal beta-glucanase units (FBG)/g, respectively. All solvents used in this study were of analytical or reagent grade, and purchased from Sigma Co. (USA). All chemicals for making DNS reagent were purchased from Daejung Chemicals Co. (Korea). Glucose, galactose, and xylose were purchased from Sigma Co. (USA). Yeast extract, malt extract, peptone and dextrose were purchased from Merck (Germany). 2.4. Determination of the cellular composition of D. tertiolecta LB999 The lipid content of the dried biomass of D. tertiolecta LB999 at cell harvesting was determined using the Soxhlet method (AOAC, 2000; method 920.39). The carbohydrate content was determined using analytical methods of the National Renewable Energy Laboratory (NREL) (Sluiter, 2006). The protein content was analyzed using a micro-Kjedahl method (AOAC 2000, method 976.05). The ash content was determined by comparing the sample weight before and after heating in a furnace at 550 °C for 12 h (Kim et al. 2011). 2.5. Saccharification of residual microalgae biomass
2. Methods 2.1. Cultivation of microalgae The microalgae strain used in this work was D. tertiolecta LB999. The components of the culture medium were as follow: NaNO3, 1.5 g/L; K2HPO4, 40 mg/L; MgSO4, 75 mg/L; CaCl2, 36 mg/L; HOOCCH2C(OH)COOHCH2COOHH2O, 6 mg/L; C6H5O7Fe3H2O, 6 mg/L; EDTA, 1 mg/L; Na2CO3, 20 mg/L; MnCl2, 1.8 mg/L; H3BO3, 2.8 mg/L; ZnSO4, 2.2 mg/L; Co(NO3)2, 0.5 mg/L; CuSO4, 0.8 mg/L; Na2MoO42H2O, 0.3 mg/L in artificial seawater (NaCl, 24.7 g/L; KCl, 0.66 g/L; MgCl26H2O, 8.48 g/L; CaCl22H2O, 1.9 g/L; MgSO47H2O, 3.07 g/L; NaHCO3, 0.18 g/L). D. tertiolecta LB999 was cultured in a 70 L plate type photobioreactor with fluorescent lighting (60 lE/m2 s) at 20–25 °C for seven days. The culture was bubbled with air containing 2% (v/v) CO2 (2 vvm; volume of air added to the liquid volume per minute). The initial biomass concentration was 0.3 g/L. The cells were harvested after seven-day culture. After centrifugation, we obtained the biomass concentration of 9.1 g fresh cell/L. The harvested cells were freeze-dried and used for lipid extraction. 2.2. Extraction of total lipids from microalgae biomass The total lipids were extracted twice from the freeze-dried cells. Fifteen volumes of chloroform and methanol (1:2 (v/v)) were added to the freeze-dried biomass, and the lipid was extracted with magnetic stirring and reflux at 65 °C for 2 h or at room temperature for overnight. After lipid extraction, the residual biomass was dried at 42 °C for 2 h or at room temperature for overnight. The dried residual biomass was ground into powder using pestle and mortar. Approximately 80% of the residual biomass powder had diameter from 75 to 300 lm. 2.3. Chemicals and enzymes Commercial cellulase (Celluclast 1.5L, Novoprime B957), amyloglucosidase (AMG 300L) and Viscozyme L were purchased from Novozymes (Denmark). Viscozyme L is a multienzyme complex containing arabanase, cellulase, b-glucanase, hemicellulase and xylanase. The enzymes activities of the Celluclast 1.5L, Novoprime B957, AMG 300L and Viscozyme L are 700 endoglucanase units (EGU)/g, 8000 high cellulase units (HCU)/g, 300 amyloglucosidase
For chemical saccharification, 5% (w/v) of the residual biomass was autoclaved at 121 °C for 15 min in the presence of HCl (0.1, 0.3, 0.5, 0.7, 1 M) or H2SO4 (0.05, 0.15, 0.25, 0.35, 0.5 M). Enzymatic saccharification was performed with 5% (w/v) of the residual biomass at various temperatures (35–55 °C) and pH (3.5–6.5). The reaction mixture was incubated with Celluclast 1.5L, Novoprime B957, AMG 300L, and/or Viscozyme L (Lee et al., 2011; Kim et al., 2011). Enzyme in the range of 0.1–1.0 mL/g was used based on the dry mass of the residual biomass. For the chemo-enzymatic saccharification, acid-generated hydrolysates were adjusted to a pH of 5.5 with 0.1 M sodium acetate buffer. Celluclast 1.5L, Viscozyme L, Novoprime B957, or AMG 300L was added for enzymatic saccharification. The samples were centrifuged at 7000 rpm and 4 °C for 10 min before analysis of the reducing sugars. Batch saccharification reaction was performed in 1 L Erlenmeyer flask (at 55 °C, pH 5.5 and 0.4 mL enzyme/g residual biomass) on a rotary shaker (230 rpm). The reaction mixture of 1 mL was taken periodically and then the amount of reducing sugars was determined using DNS method for the analysis of saccharification reaction progress. 2.6. Ethanol production using enzymatic hydrolysates of the residual biomass Saccharomyces cerevisiae YPH500 (ATCC 76626) was used for ethanol fermentation of the enzymatic saccharification products of the residual biomass (Boubekeur et al. 2001). For seed culture, S. cerevisiae YPH500 was cultured in 10 mL culture tubes containing 3 mL of culture medium at 30 °C and 200 rpm for 12 h. The composition of the culture medium was as follows: yeast extract, 3 g/L; malt extract, 3 g/L; peptone 5 g/L; dextrose 10 g/L (Lee et al., 2011). For a flask fermentation, 1 mL of the seed culture was inoculated to 200 mL of the enzymatic saccharification solution containing 17.5 g/L reducing sugars and 12 g/L yeast extract in a 500 mL culture flask at 30 °C and 200 rpm for 12 h. For a fermenter experiment, 10 mL of the seed culture was inoculated to the enzymatic saccharification solution (working volume: 1 L) in a 5 L fermenter with pH (6.5) and temperature (30 °C) control (Biotron, Korea). The glucose consumption and ethanol production were analyzed periodically. As a control, we conducted two independent fermentation experiments. Culture medium containing only 12 g/L yeast extract was used as the negative control experi-
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ment. Culture medium containing 17.5 g/L glucose and 12 g/L yeast extract was used as the positive control experiment. 2.7. Analyses The total reducing sugar concentration in the saccharification products of the microalgal residual biomass was determined using dinitrosalicyclic acid (DNS) reagent (Miller, 1959). The absorbance was measured at 550 nm, and D-glucose was used as a standard. The reducing sugar concentration and composition were also determined by HPLC with a refractive index (RI) detector and Biorad HPX-87-H organic acid column (Lee et al., 2011; Kim et al., 2011). The column temperature was 60 °C, and 5 mM H2SO4 was used as the mobile phase at a flow rate of 0.6 mL/min. All reaction mixtures were filtered with a 0.2 lm membrane before analysis. Saccharification efficiency is calculated as follows: saccharification efficiency (%) = (reducing sugars determined by DNS or HPLC/residual biomass (mg)) 100. During the ethanol fermentation, the concentrations of reducing sugars and bioethanol were determined by HPLC under the same conditions described above. 3. Results and discussion 3.1. Chemical saccharification
Saccharification yields (% w/w residual biomass)
After photoautotrophic culture of D. tertiolecta LB999, the cells were harvested and concentrated. The compositions of carbohydrate, lipid, protein, ash, and moisture in D. tertiolecta biomass were 37.8, 20.6, 25.5, 9.6 and 6.5% (w/w), respectively. We obtained 48 g lipid and 172 g residual biomass, which indicated that the lipids were nearly completely extracted by methanol and chloroform. The residual biomass was analyzed to determine the composition of carbohydrate, protein, moisture and ash using the corresponding methods described in the Section 2. The percentage of total carbohydrates of the residual biomass after lipid extraction was approximately 51.9% (w/w). We conducted a diluted acid-mediated saccharification of the residual biomass. Effects of sulfuric acid and hydrochloric acid concentration on the saccharification yield were evaluated to determine the optimal concentration of diluted acid (Fig. 1). The amount of reducing sugars during saccharification was determined by the DNS method. The maximum amount of reducing sugars was approximately 14.8 mg/mL, represented as the D-glucose equiva-
HCl H2SO4
30 25 20 15 10 5 0
0.1
0.3
0.5
0.7
1
Acid concentration (N) Fig. 1. Effect of diluted acid concentrations on chemical saccharification of the residual biomass after lipid extraction. Acidic saccharification condition: 121 °C, 15 min.
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lent, when 5% (w/v) of the residual biomass was hydrolyzed by 0.5 M HCl with autoclaving at 121 °C for 15 min. A saccharification yield of 29.5% (w/w, based on the residual biomass) was obtained. The yield based on the total carbohydrate amount of the residual biomass after lipid extraction was 56.7% (w/w) (Table 1). 3.2. Enzymatic saccharification Enzymatic saccharification of the residual biomass was investigated. As the first step, various saccharification enzymes were screened for their ability to hydrolyze the residual biomass. AMG 300L (amyloglucosidase), Celluclast 1.5L (endocellulase), Novoprime B957 (endocellulase) and Viscozyme L were tested as the biocatalyst for saccharification. Amyloglucosidase hydrolyzes the terminal 1,4-linked a-D-glucose residue with the release of b-Dglucose. Endocellulase randomly hydrolyzes the internal bonds of 1,4-b-D-glycosidic linkages in cellulose. Viscozyme L contains arabanase, cellulase, beta-glucanase, hemicellulase and xylanase. Arabanase hydrolyzes araban, and hemicellulase degrades hemicellulose. b-Glucanase is an endo-glycosidase. Xylanase degrades linear polysaccharide b-1,4-xylan into xylose. As shown in Table 2, AMG 300L and Viscozyme L exhibited relatively better hydrolysis activity than the other enzymes. At a pH of 5.5 and temperature of 55 °C, enzymatic saccharification yields were 21.9%, 19.2%, 0.3% and 0.2% (w/w on the basis of the residual biomass) for AMG 300L, Viscozyme L, Novoprime B957 and Celluclast 1.5L, respectively. The yields based on the amount of total carbohydrates of the residual biomass after lipid extraction were 42.2%, 37.0%, 0.6% and 0.4% (w/w) for AMG 300L, Viscozyme L, Novoprime B957 and Celluclast 1.5L, respectively. As the major component of total carbohydrates of the residual biomass after lipid extraction was starch, AMG 300L and Viscozyme L containing amyloglucosidase and b-glucanase exhibited better hydrolysis activities than Celluclast 1.5L and Novoprime B957 containing endocellulases. The effects of pH and temperature on the saccharification efficiency were investigated. The pH range was from 3.5 to 6.5, and the reaction temperature ranged from 35 to 55 °C. As shown in Table 2, the pH of 5.5 and temperature of 55 °C were optimal for the AMG 300L and Viscozyme L. The combined use of AMG 300L and Viscozyme L was also evaluated (Supplementary Fig. 1) as some enzymes can work synergistically to degrade polysaccharides. The use of the enzyme mixture of AMG 300L and Viscozyme L (1:1) showed good results but did not substantially enhance the saccharification efficiency. As the major component of total carbohydrates in the residual biomass was starch, the saccharification efficiency of AMG 300L was better than that of other enzymes. Although low-cost saccharification enzymes are available, the saccharification enzyme cost is a burden in the production of bioethanol from biomass. Therefore, the enzyme dosage needs to be optimized. When AMG 300L was used, the saccharification yield increased gradually as the enzyme concentration increased to 0.4 mL/ g of residual biomass for 1 h saccharification reaction (Supplementary Fig. 2). Above concentration of 0.4 mL/g of residual biomass, the saccharification yield was not greatly affected by the enzyme concentration. When 0.4 mL AMG 300L per 1 g of the residual biomass was used, a saccharification yield of 17.5% (w/w) was obtained. The yield based on the amount of total carbohydrates of the residual biomass after lipid extraction was 33.7% (w/w). 3.3. Chemical and enzymatic saccharification Generally, hydrolysis of carbohydrates involves liquefaction and saccharification steps. In order to saccharify the intracellular carbohydrates, such as starch granules (Libessart et al., 1995), the microalgae cell wall needs to be broken down by a pretreatment.
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Table 1 Comparison of the saccharification yield of enzymatic and chemo-enzymatic saccharification based on the residual biomass and the total carbohydrates of the residual biomass.
a b
Methods
Sugar concentration (mg/mL)
Saccharification yielda
Saccharification yieldb
0.5 M HCl AMG Acid pretreatment and enzymatic saccharification
14.8 21.0 21.2
29.5 42.0 42.4
57.1 80.9 81.7
Saccharification yield: w/w, based on the residual biomass. Saccharification yield: w/w, based on the total amount of carbohydrates of the residual biomass.
Table 2 Effects of enzyme, reaction temperature and pH on the enzymatic saccharification yield of the residual biomass after lipid extraction. Enzymatic saccharification condition: 0.3 mL enzyme/g residual biomass and 2 h. 35 °C
AMG Celluclast 1.5 L Novoprime B 957 Viscozyme L
45 °C
55 °C
pH 3.5
pH 4.5
pH 5.5
pH 6.5
pH 3.5
pH 4.5
pH 5.5
pH 6.5
pH 3.5
pH 4.5
pH 5.5
pH 6.5
0.0 0.8 0.0 0.0
15.7 0.1 0.0 10.3
12.0 0.1 0.1 10.7
8.0 0.8 0.2 7.9
0.0 0.0 0.6 0.7
17.7 0.1 0.4 12.5
14.2 0.2 0.9 12.4
12.1 0.1 0.8 12.4
0.0 0.7 0.0 0.1
18.9 0.4 0.3 16.7
21.9 0.2 0.3 19.2
17.2 0.3 0.4 14.3
However, in our case, the autoclaving pretreatment step did not affect the saccharification efficiency (data not shown) because the harvested cells were already disrupted during lipid extraction by methanol and chloroform. As a result, saccharification experiments were performed without pretreatment. In order to enhance the saccharification efficiency, a dilute acid pretreatment step was applied before enzymatic hydrolysis of the residual biomass (Supplementary Fig. 3). When the residual biomass was treated under the optimum conditions of diluted acidmediated treatment (0.5 M HCl, 121 °C, 15 min) and AMG 300Lcatalyzed saccharification (0.4 mL enzyme per 1 g of the residual biomass, 55 °C, pH 5.5, 12 h), the amount of reducing sugar reached 21.2 mg/mL, corresponding to a yield of 42.4% (w/w, based on the residual biomass), after 12 h reaction (Table 1). However, the overall efficiency was nearly the same as that of the enzymatic treatment only, indicating that the microalgal residual biomass after oil extraction could be saccharified without chemical pretreatment. A simple enzymatic saccharification process without chemical treatment would be helpful for improving the cost effectiveness of bioethanol production from the residual biomass. 3.4. Batch saccharification of the microalgal residual biomass and analysis of saccharification products Batch saccharification of the microalgal residual biomass was conducted at the optimal condition (Supplementary Fig. 4). As the batch saccharification by AMG 300L progressed, the amount of reducing sugars increased. The amount of the released reducing sugars reached a maximum at 12 h under the optimal condition. The final amount of reducing sugars represented as the D-glucose equivalent was 21.0 mg/mL. A saccharification yield of 42.0% (w/ w, based on the residual biomass) was obtained (Table 1). The yield based on the total amount of carbohydrates of the residual biomass after lipid extraction was 80.9% (w/w). The degradation products were analyzed by HPLC. The main component in the degradation mixture was glucose. D-glucose was released as the main product as the AMG 300L, an amyloglucosidase, hydrolyzes the terminal 1,4-linked a-D-glucose residues. From the above results, the residual biomass of D. tertiolecta LB999 after lipid extraction was successfully saccharified into fermentable sugars. 3.5. Bioethanol fermentation using enzymatic hydrolysates of the microalgal residual biomass Bioethanol was produced from the enzymatic saccharification products of the residual biomass using S. cerevisiae YPH500. The
Fig. 2. Bioethanol fermentation of the enzymatic hydrolysates of the residual biomass using S. cerevisiae YPH500. (d; Glucose decline profile, s; Bioethanol production profile, j; Cell growth profile).
cell growth, sugar consumption and bioethanol generation profiles are shown in Fig. 2. After 12 h of bioethanol fermentation, the concentration of the reducing sugar decreased with time, and the concentration of bioethanol increase gradually, clearly indicating that the saccharification products of the residual biomass of D. tertiolecta LB999 were used as fermentation sugars. In the negative control experiment, negligible amount of ethanol was produced. On the contrary, when ethanol fermentation was conducted with the culture medium containing 17.5 g/L glucose and 12 g/L yeast extract as the positive control experiment, similar amount of ethanol was produced, supporting that the reducing sugars, mainly consisting of glucose, of the enzymatic saccharification products were successfully used for ethanol fermentation. The fermentation yields of bioethanol based on the residual biomass weight and the saccharified glucose were 0.14 g ethanol/g residual biomass and 0.44 g ethanol/g glucose, respectively. Without additional pretreatment steps after enzymatic saccharification, a high fermentation yield of bioethanol, 82% of the theoretical fermentation yield, was obtained. When taking it consideration that an unidentified monosaccharide existed at approximately 4.3% of total reducing sugars based on the ratio of HPLC peaks, the bioethanol fermentation yield from glucose was almost up to 85.7% of the theoretical fermentation yield. The unidentified monosaccharide was not used for ethanol fermentation by S. cerevisiae YPH500. Direct bioethanol fermentation without additional pretreatment
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steps would be helpful for decreasing the process cost by using the simplified bioethanol production process. 4. Conclusion The saccharification of microalgal residual biomass was successfully performed. AMG 300L-catalyzed saccharification produced 21.0 mg/mL of reducing sugar with 42.0% yield based on the residual biomass mass. The saccharification yield based on the total amount of carbohydrates of the residual biomass was 80.9% (w/w). Bioethanol was produced from the enzymatic saccharification products by S. cerevisiae YPH500, and the fermentation yield was 0.14 g ethanol/g residual biomass and 82.0% of the theoretical fermentation yield. This study clearly showed that the residual biomass of microalgae after lipid extraction for biodiesel production could be used for saccharification and subsequent bioethanol fermentation. Acknowledgements This work was supported by a grant from the Development of Marine-Bioenergy program funded by the Ministry of Land, Transport and Maritime Affairs of the Korean government. 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.2013.01. 007. References AOAC (Association of Analytical Chemists), 2000. Official Methods of Analysis of Association of Analytical Chemist. Horwitz, W., Gaithersburg, Maryland, USA. Boubekeur, S., Camougrand, N., Bunoust, O., Rigoulet, M., Guerin, B., 2001. Participation of acetaldehyde dehydrogenases in ethanol and pyruvate metabolism of the yeast Saccharomyces cerevisiae. Eur. J. Biochem. 268, 5057– 5065. Choi, S.P., Nguyen, M.T., Sim, S.J., 2010. Enzymatic pretreatment of Chlamydomonas reinhardtii biomass for ethanol production. Bioresour. Technol. 101, 5330–5336.
201
Demirbas, A., 2006. Global biofuel strategies. Energy Educ. Sci. Technol. 17, 27–63. Harun, R., Danquah, M.K., 2011. Influence of acid pre-treatment on microalgal biomass for bioethanol production. Process Biochem. 46, 304–309. Harun, R., Danquah, M.K., Forde, G.M., 2010. Microalgal biomass as a fermentation feedstock for bioethanol production. J. Chem. Technol. Biotechnol. 85, 199–203. Harun, R., Jason, W.S.Y., Cherrington, T., Danquah, M.K., 2011. Exploring alkaline pre-treatment of microalgal biomass for bioethanol production. Appl. Energy 88, 3464–3467. John, R.P., Anisha, G.S., Nampoothiri, K.M., Pandey, A., 2011. Micro and macroalgal biomass: a renewable source for bioethanol. Bioresour. Technol. 102, 186–193. Kim, Z., Lee, C.G., 2010. Photobioreactor for a large-scale marine microalgal culture using semi-permeable membrane. Korean Pat. 10-0991373. Kim, N.J., Li, H., Jung, K.S., Chang, H.N., Lee, P.C., 2011. Ethanol production from marine algal hydrolysates using Escherichia coli KO11. Bioresour. Technol. 102, 7466–7469. Lee, S.J., Oh, Y.H., Kim, D.H., Kwon, D.Y., Lee, C.G., Lee, J.W., 2011. Converting carbohydrates extracted from marine algae into ethanol using various ethanolic Escherichia coli strains. Appl. Biochem. Biotechnol. 164, 878–888. Libessart, N., Maddelein, M.L., Koornhuyse, N., Decq, A., Delrue, B., Mouille, G., D’Hulst, C., Ball, S., 1995. Storage, photosynthesis and growth: the conditional nature of mutations affecting starch synthesis and structure in Chlamydomonas. Plant Cell 7, 1117–1127. Lin, Y., Tanaka, S., 2006. Ethanol fermentation from biomass resources: current state and prospects. Appl. Microbiol. Biotechnol. 69, 627–642. Meher, L.C., Sagar, D.V., Naik, S.N., 2006. Technical aspects of biodiesel production by transesterification – a review. Renew. Sustain. Energy Rev. 10, 248–268. Miller, G.L., 1959. Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal. Chem. 31, 426–428. Miranda, J.R., Passarinho, P.C., Gouveia, L., 2012. Pre-treatment optimization of Scenedesmus obliquus microalga for bioethanol production. Bioresour. Technol. 104, 342–348. Nigam, P.S., Singh, A., 2011. Production of liquid biofuels from renewable resources. Prog. Energy Combust. 37, 52–68. Singh, A., Nigam, P.S., Murphy, J.D., 2011. Mechanism and challenges in commercialisation of algal biofuels. Bioresour. Technol. 102, 26–34. Sluiter, A., 2006. Determination of Structural Carbohydrates and Lignin in Biomass, Version 2006. National Renewable Energy Laboratory, USA.. Wahlen, B.D., Willis, R.M., Seefeldt, L.C., 2011. Biodiesel production by simultaneous extraction and conversion of total lipids from microalgae, cyanobacteria, and wild mixed-cultures. Bioresour. Technol. 102, 2724–2730. Waltz, E., 2009. Biotech’s green gold? Nat. Biotechnol. 27, 15–18. Wargacki, A.J., Leonard, E., Win, M.N., Regitsky, D.D., Santos, C.N.S., Kim, P.B., Cooper, S.R., Raisner, R.M., Herman, A., Sivitz, A.B., Lakshmanaswamy, A., Kashiyama, Y., Baker, D., Yoshikuni, Y., 2012. An engineered microbial platform for direct biofuel production from brown macroalgae. Science 335, 308–313. Zou, S., Wu, Y., Yang, M., Li, C., Tong, J., 2009. Thermochemical catalytic liquefaction of the marine microalgae Dunaliella tertiolecta and characterization of bio-oils. Energy Fuels 23, 3753–3758.
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Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Chemo-enzymatic saccharification and bioethanol fermentation of lipid-extracted residual biomass of the microalga, Dunaliella tertiolecta Ok Kyung Lee a, A Leum Kim a, Dong Ho Seong b, Choul Gyun Lee b, Yeon Tae Jung c, Jin Won Lee c, Eun Yeol Lee a,⇑ a b c
Department of Chemical Engineering, Kyung Hee University, Gyeonggi-do 446-701, Republic of Korea Marine Bioenergy Research Center, Department of Biological Engineering, Inha University, Incheon 402-751, Republic of Korea Department of Chemical and Biomolecular Engineering, Sogang University, Seoul 121-742, Republic of Korea
h i g h l i g h t s " The residual biomass of Dunaliella tertiolecta after lipid extraction was saccharified and used for bioethanol fermentation. " A pretreatment procedure was not required for enzymatic saccharification of the residual biomass. " Saccharification yield based on the total amount of carbohydrates was 80.9% (w/w). " Bioethanol was directly produced with 82% yield from the saccharification solution without additional pretreatment. " The waste residual biomass generated during microalgal biodiesel production could be used for the production of bioethanol.
a r t i c l e
i n f o
Article history: Received 14 August 2012 Received in revised form 29 December 2012 Accepted 4 January 2013 Available online 19 January 2013 Keywords: Bioethanol fermentation Dunaliella tertiolecta Reducing sugar Residual biomass Saccharification
a b s t r a c t Chemo-enzymatic saccharification and bioethanol fermentation of the residual biomass of Dunaliella tertiolecta after lipid extraction for biodiesel production were investigated. HCl-catalyzed saccharification of the residual biomass at 121 °C for 15 min produced reducing sugars with a yield of 29.5% (w/w) based on the residual biomass dry weight. Various enzymes were evaluated for their ability to saccharify the residual biomass. Enzymatic saccharification using AMG 300L produced 21.0 mg/mL of reducing sugar with a yield of 42.0% (w/w) based on the residual biomass at pH 5.5 and 55 °C. Bioethanol was produced from the enzymatic saccharification products without additional pretreatment by Saccharomyces cerevisiae with yields of 0.14 g ethanol/g residual biomass and 0.44 g ethanol/g glucose produced from the residual biomass. The waste residual biomass generated during microalgal biodiesel production could be used for the production of bioethanol to improve the economic feasibility of microalgal biorefinery. Ó 2013 Elsevier Ltd. All rights reserved.
1. Introduction The depletion of fossil fuels has triggered the development of alternative renewable biofuels, such as bioethanol and biodiesel (Lin and Tanaka, 2006; Meher et al., 2006). Various types of biomass have been considered as feedstock for biofuel production (Demirbas, 2006). Starch and sugarcane have been used for commercial-scale production of bioethanol, i.e., first generation biomass. However, crop-based first generation bioethanol causes moral issues of using food for fuel production. Second generation bioethanol uses various cellulosic materials as feedstock. Despite intensive research on cellulosic bioethanol, commercial plants do not exist due to the difficulty and complexity of lignin pretreat-
⇑ Corresponding author. Tel.: +82 31 201 3839; fax: +82 31 204 8114. E-mail address: [email protected] (E.Y. Lee). 0960-8524/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2013.01.007
ment. One of the critical success factors of microbial bioethanol production from biomass is cost-competitiveness of the saccharification process. To address these technical challenges, much effort has been given to the development of cost-effective pretreatment and saccharification processes for cellulosic biomass (Nigam and Singh, 2011). As the third generation biomass, microalgae and macroalgae have been considered as feedstock for biofuel production based on the expectation that large amounts of biomass will soon be available at an acceptable cost (John et al., 2011; Waltz, 2009). Biofuel production using algae has attracted much attention because it can be cultured using CO2 and sunlight (Singh et al., 2011). Another advantage of microalgae and macroalgae is that they do not have lignin, allowing development of a cost-effective pretreatment process (Libessart et al., 1995; Miranda et al., 2012). Microalgae is good feedstock for biodiesel production because it accumulates a high percentage of lipids on the basis of dry cell
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mass (Nigam and Singh, 2011; Wahlen et al., 2011; Zou et al., 2009). Recently, we developed a large-scale cultivation method for the marine microalgae, Dunaliella tertiolecta LB999, with a photobioreactor using a semi-permeable membrane for the production of biodiesel (Kim and Lee, 2010). During the production of biodiesel from D. tertiolecta LB999 biomass, a significant amount of residual biomass is generated as waste after lipid extraction. Biodiesel production from microalgae still suffers from high production cost. Most of the valuable cell components of microalgae need to be utilized to enhance the economic feasibility of microalgal biorefinery. The residual biomass can be used for the production of bioethanol because it has a large amount of polysaccharide. Many investigations on saccharification and bioethanol fermentation from microalgae and macroalgae biomass have been performed (Choi et al., 2010; Harun et al., 2010, 2011; Harun and Danquah, 2011; Kim et al., 2011; Lee et al., 2011; Wargacki et al., 2012). However, investigations on saccharification and bioethanol fermentation of the microalgal residual biomass after lipid extraction have not been reported. The main objective of this study is to produce bioethanol from the residual biomass of D. tertiolecta LP999 resulted from lipid extraction.
units (AGU)/mL and 100 fungal beta-glucanase units (FBG)/g, respectively. All solvents used in this study were of analytical or reagent grade, and purchased from Sigma Co. (USA). All chemicals for making DNS reagent were purchased from Daejung Chemicals Co. (Korea). Glucose, galactose, and xylose were purchased from Sigma Co. (USA). Yeast extract, malt extract, peptone and dextrose were purchased from Merck (Germany). 2.4. Determination of the cellular composition of D. tertiolecta LB999 The lipid content of the dried biomass of D. tertiolecta LB999 at cell harvesting was determined using the Soxhlet method (AOAC, 2000; method 920.39). The carbohydrate content was determined using analytical methods of the National Renewable Energy Laboratory (NREL) (Sluiter, 2006). The protein content was analyzed using a micro-Kjedahl method (AOAC 2000, method 976.05). The ash content was determined by comparing the sample weight before and after heating in a furnace at 550 °C for 12 h (Kim et al. 2011). 2.5. Saccharification of residual microalgae biomass
2. Methods 2.1. Cultivation of microalgae The microalgae strain used in this work was D. tertiolecta LB999. The components of the culture medium were as follow: NaNO3, 1.5 g/L; K2HPO4, 40 mg/L; MgSO4, 75 mg/L; CaCl2, 36 mg/L; HOOCCH2C(OH)COOHCH2COOHH2O, 6 mg/L; C6H5O7Fe3H2O, 6 mg/L; EDTA, 1 mg/L; Na2CO3, 20 mg/L; MnCl2, 1.8 mg/L; H3BO3, 2.8 mg/L; ZnSO4, 2.2 mg/L; Co(NO3)2, 0.5 mg/L; CuSO4, 0.8 mg/L; Na2MoO42H2O, 0.3 mg/L in artificial seawater (NaCl, 24.7 g/L; KCl, 0.66 g/L; MgCl26H2O, 8.48 g/L; CaCl22H2O, 1.9 g/L; MgSO47H2O, 3.07 g/L; NaHCO3, 0.18 g/L). D. tertiolecta LB999 was cultured in a 70 L plate type photobioreactor with fluorescent lighting (60 lE/m2 s) at 20–25 °C for seven days. The culture was bubbled with air containing 2% (v/v) CO2 (2 vvm; volume of air added to the liquid volume per minute). The initial biomass concentration was 0.3 g/L. The cells were harvested after seven-day culture. After centrifugation, we obtained the biomass concentration of 9.1 g fresh cell/L. The harvested cells were freeze-dried and used for lipid extraction. 2.2. Extraction of total lipids from microalgae biomass The total lipids were extracted twice from the freeze-dried cells. Fifteen volumes of chloroform and methanol (1:2 (v/v)) were added to the freeze-dried biomass, and the lipid was extracted with magnetic stirring and reflux at 65 °C for 2 h or at room temperature for overnight. After lipid extraction, the residual biomass was dried at 42 °C for 2 h or at room temperature for overnight. The dried residual biomass was ground into powder using pestle and mortar. Approximately 80% of the residual biomass powder had diameter from 75 to 300 lm. 2.3. Chemicals and enzymes Commercial cellulase (Celluclast 1.5L, Novoprime B957), amyloglucosidase (AMG 300L) and Viscozyme L were purchased from Novozymes (Denmark). Viscozyme L is a multienzyme complex containing arabanase, cellulase, b-glucanase, hemicellulase and xylanase. The enzymes activities of the Celluclast 1.5L, Novoprime B957, AMG 300L and Viscozyme L are 700 endoglucanase units (EGU)/g, 8000 high cellulase units (HCU)/g, 300 amyloglucosidase
For chemical saccharification, 5% (w/v) of the residual biomass was autoclaved at 121 °C for 15 min in the presence of HCl (0.1, 0.3, 0.5, 0.7, 1 M) or H2SO4 (0.05, 0.15, 0.25, 0.35, 0.5 M). Enzymatic saccharification was performed with 5% (w/v) of the residual biomass at various temperatures (35–55 °C) and pH (3.5–6.5). The reaction mixture was incubated with Celluclast 1.5L, Novoprime B957, AMG 300L, and/or Viscozyme L (Lee et al., 2011; Kim et al., 2011). Enzyme in the range of 0.1–1.0 mL/g was used based on the dry mass of the residual biomass. For the chemo-enzymatic saccharification, acid-generated hydrolysates were adjusted to a pH of 5.5 with 0.1 M sodium acetate buffer. Celluclast 1.5L, Viscozyme L, Novoprime B957, or AMG 300L was added for enzymatic saccharification. The samples were centrifuged at 7000 rpm and 4 °C for 10 min before analysis of the reducing sugars. Batch saccharification reaction was performed in 1 L Erlenmeyer flask (at 55 °C, pH 5.5 and 0.4 mL enzyme/g residual biomass) on a rotary shaker (230 rpm). The reaction mixture of 1 mL was taken periodically and then the amount of reducing sugars was determined using DNS method for the analysis of saccharification reaction progress. 2.6. Ethanol production using enzymatic hydrolysates of the residual biomass Saccharomyces cerevisiae YPH500 (ATCC 76626) was used for ethanol fermentation of the enzymatic saccharification products of the residual biomass (Boubekeur et al. 2001). For seed culture, S. cerevisiae YPH500 was cultured in 10 mL culture tubes containing 3 mL of culture medium at 30 °C and 200 rpm for 12 h. The composition of the culture medium was as follows: yeast extract, 3 g/L; malt extract, 3 g/L; peptone 5 g/L; dextrose 10 g/L (Lee et al., 2011). For a flask fermentation, 1 mL of the seed culture was inoculated to 200 mL of the enzymatic saccharification solution containing 17.5 g/L reducing sugars and 12 g/L yeast extract in a 500 mL culture flask at 30 °C and 200 rpm for 12 h. For a fermenter experiment, 10 mL of the seed culture was inoculated to the enzymatic saccharification solution (working volume: 1 L) in a 5 L fermenter with pH (6.5) and temperature (30 °C) control (Biotron, Korea). The glucose consumption and ethanol production were analyzed periodically. As a control, we conducted two independent fermentation experiments. Culture medium containing only 12 g/L yeast extract was used as the negative control experi-
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ment. Culture medium containing 17.5 g/L glucose and 12 g/L yeast extract was used as the positive control experiment. 2.7. Analyses The total reducing sugar concentration in the saccharification products of the microalgal residual biomass was determined using dinitrosalicyclic acid (DNS) reagent (Miller, 1959). The absorbance was measured at 550 nm, and D-glucose was used as a standard. The reducing sugar concentration and composition were also determined by HPLC with a refractive index (RI) detector and Biorad HPX-87-H organic acid column (Lee et al., 2011; Kim et al., 2011). The column temperature was 60 °C, and 5 mM H2SO4 was used as the mobile phase at a flow rate of 0.6 mL/min. All reaction mixtures were filtered with a 0.2 lm membrane before analysis. Saccharification efficiency is calculated as follows: saccharification efficiency (%) = (reducing sugars determined by DNS or HPLC/residual biomass (mg)) 100. During the ethanol fermentation, the concentrations of reducing sugars and bioethanol were determined by HPLC under the same conditions described above. 3. Results and discussion 3.1. Chemical saccharification
Saccharification yields (% w/w residual biomass)
After photoautotrophic culture of D. tertiolecta LB999, the cells were harvested and concentrated. The compositions of carbohydrate, lipid, protein, ash, and moisture in D. tertiolecta biomass were 37.8, 20.6, 25.5, 9.6 and 6.5% (w/w), respectively. We obtained 48 g lipid and 172 g residual biomass, which indicated that the lipids were nearly completely extracted by methanol and chloroform. The residual biomass was analyzed to determine the composition of carbohydrate, protein, moisture and ash using the corresponding methods described in the Section 2. The percentage of total carbohydrates of the residual biomass after lipid extraction was approximately 51.9% (w/w). We conducted a diluted acid-mediated saccharification of the residual biomass. Effects of sulfuric acid and hydrochloric acid concentration on the saccharification yield were evaluated to determine the optimal concentration of diluted acid (Fig. 1). The amount of reducing sugars during saccharification was determined by the DNS method. The maximum amount of reducing sugars was approximately 14.8 mg/mL, represented as the D-glucose equiva-
HCl H2SO4
30 25 20 15 10 5 0
0.1
0.3
0.5
0.7
1
Acid concentration (N) Fig. 1. Effect of diluted acid concentrations on chemical saccharification of the residual biomass after lipid extraction. Acidic saccharification condition: 121 °C, 15 min.
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lent, when 5% (w/v) of the residual biomass was hydrolyzed by 0.5 M HCl with autoclaving at 121 °C for 15 min. A saccharification yield of 29.5% (w/w, based on the residual biomass) was obtained. The yield based on the total carbohydrate amount of the residual biomass after lipid extraction was 56.7% (w/w) (Table 1). 3.2. Enzymatic saccharification Enzymatic saccharification of the residual biomass was investigated. As the first step, various saccharification enzymes were screened for their ability to hydrolyze the residual biomass. AMG 300L (amyloglucosidase), Celluclast 1.5L (endocellulase), Novoprime B957 (endocellulase) and Viscozyme L were tested as the biocatalyst for saccharification. Amyloglucosidase hydrolyzes the terminal 1,4-linked a-D-glucose residue with the release of b-Dglucose. Endocellulase randomly hydrolyzes the internal bonds of 1,4-b-D-glycosidic linkages in cellulose. Viscozyme L contains arabanase, cellulase, beta-glucanase, hemicellulase and xylanase. Arabanase hydrolyzes araban, and hemicellulase degrades hemicellulose. b-Glucanase is an endo-glycosidase. Xylanase degrades linear polysaccharide b-1,4-xylan into xylose. As shown in Table 2, AMG 300L and Viscozyme L exhibited relatively better hydrolysis activity than the other enzymes. At a pH of 5.5 and temperature of 55 °C, enzymatic saccharification yields were 21.9%, 19.2%, 0.3% and 0.2% (w/w on the basis of the residual biomass) for AMG 300L, Viscozyme L, Novoprime B957 and Celluclast 1.5L, respectively. The yields based on the amount of total carbohydrates of the residual biomass after lipid extraction were 42.2%, 37.0%, 0.6% and 0.4% (w/w) for AMG 300L, Viscozyme L, Novoprime B957 and Celluclast 1.5L, respectively. As the major component of total carbohydrates of the residual biomass after lipid extraction was starch, AMG 300L and Viscozyme L containing amyloglucosidase and b-glucanase exhibited better hydrolysis activities than Celluclast 1.5L and Novoprime B957 containing endocellulases. The effects of pH and temperature on the saccharification efficiency were investigated. The pH range was from 3.5 to 6.5, and the reaction temperature ranged from 35 to 55 °C. As shown in Table 2, the pH of 5.5 and temperature of 55 °C were optimal for the AMG 300L and Viscozyme L. The combined use of AMG 300L and Viscozyme L was also evaluated (Supplementary Fig. 1) as some enzymes can work synergistically to degrade polysaccharides. The use of the enzyme mixture of AMG 300L and Viscozyme L (1:1) showed good results but did not substantially enhance the saccharification efficiency. As the major component of total carbohydrates in the residual biomass was starch, the saccharification efficiency of AMG 300L was better than that of other enzymes. Although low-cost saccharification enzymes are available, the saccharification enzyme cost is a burden in the production of bioethanol from biomass. Therefore, the enzyme dosage needs to be optimized. When AMG 300L was used, the saccharification yield increased gradually as the enzyme concentration increased to 0.4 mL/ g of residual biomass for 1 h saccharification reaction (Supplementary Fig. 2). Above concentration of 0.4 mL/g of residual biomass, the saccharification yield was not greatly affected by the enzyme concentration. When 0.4 mL AMG 300L per 1 g of the residual biomass was used, a saccharification yield of 17.5% (w/w) was obtained. The yield based on the amount of total carbohydrates of the residual biomass after lipid extraction was 33.7% (w/w). 3.3. Chemical and enzymatic saccharification Generally, hydrolysis of carbohydrates involves liquefaction and saccharification steps. In order to saccharify the intracellular carbohydrates, such as starch granules (Libessart et al., 1995), the microalgae cell wall needs to be broken down by a pretreatment.
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Table 1 Comparison of the saccharification yield of enzymatic and chemo-enzymatic saccharification based on the residual biomass and the total carbohydrates of the residual biomass.
a b
Methods
Sugar concentration (mg/mL)
Saccharification yielda
Saccharification yieldb
0.5 M HCl AMG Acid pretreatment and enzymatic saccharification
14.8 21.0 21.2
29.5 42.0 42.4
57.1 80.9 81.7
Saccharification yield: w/w, based on the residual biomass. Saccharification yield: w/w, based on the total amount of carbohydrates of the residual biomass.
Table 2 Effects of enzyme, reaction temperature and pH on the enzymatic saccharification yield of the residual biomass after lipid extraction. Enzymatic saccharification condition: 0.3 mL enzyme/g residual biomass and 2 h. 35 °C
AMG Celluclast 1.5 L Novoprime B 957 Viscozyme L
45 °C
55 °C
pH 3.5
pH 4.5
pH 5.5
pH 6.5
pH 3.5
pH 4.5
pH 5.5
pH 6.5
pH 3.5
pH 4.5
pH 5.5
pH 6.5
0.0 0.8 0.0 0.0
15.7 0.1 0.0 10.3
12.0 0.1 0.1 10.7
8.0 0.8 0.2 7.9
0.0 0.0 0.6 0.7
17.7 0.1 0.4 12.5
14.2 0.2 0.9 12.4
12.1 0.1 0.8 12.4
0.0 0.7 0.0 0.1
18.9 0.4 0.3 16.7
21.9 0.2 0.3 19.2
17.2 0.3 0.4 14.3
However, in our case, the autoclaving pretreatment step did not affect the saccharification efficiency (data not shown) because the harvested cells were already disrupted during lipid extraction by methanol and chloroform. As a result, saccharification experiments were performed without pretreatment. In order to enhance the saccharification efficiency, a dilute acid pretreatment step was applied before enzymatic hydrolysis of the residual biomass (Supplementary Fig. 3). When the residual biomass was treated under the optimum conditions of diluted acidmediated treatment (0.5 M HCl, 121 °C, 15 min) and AMG 300Lcatalyzed saccharification (0.4 mL enzyme per 1 g of the residual biomass, 55 °C, pH 5.5, 12 h), the amount of reducing sugar reached 21.2 mg/mL, corresponding to a yield of 42.4% (w/w, based on the residual biomass), after 12 h reaction (Table 1). However, the overall efficiency was nearly the same as that of the enzymatic treatment only, indicating that the microalgal residual biomass after oil extraction could be saccharified without chemical pretreatment. A simple enzymatic saccharification process without chemical treatment would be helpful for improving the cost effectiveness of bioethanol production from the residual biomass. 3.4. Batch saccharification of the microalgal residual biomass and analysis of saccharification products Batch saccharification of the microalgal residual biomass was conducted at the optimal condition (Supplementary Fig. 4). As the batch saccharification by AMG 300L progressed, the amount of reducing sugars increased. The amount of the released reducing sugars reached a maximum at 12 h under the optimal condition. The final amount of reducing sugars represented as the D-glucose equivalent was 21.0 mg/mL. A saccharification yield of 42.0% (w/ w, based on the residual biomass) was obtained (Table 1). The yield based on the total amount of carbohydrates of the residual biomass after lipid extraction was 80.9% (w/w). The degradation products were analyzed by HPLC. The main component in the degradation mixture was glucose. D-glucose was released as the main product as the AMG 300L, an amyloglucosidase, hydrolyzes the terminal 1,4-linked a-D-glucose residues. From the above results, the residual biomass of D. tertiolecta LB999 after lipid extraction was successfully saccharified into fermentable sugars. 3.5. Bioethanol fermentation using enzymatic hydrolysates of the microalgal residual biomass Bioethanol was produced from the enzymatic saccharification products of the residual biomass using S. cerevisiae YPH500. The
Fig. 2. Bioethanol fermentation of the enzymatic hydrolysates of the residual biomass using S. cerevisiae YPH500. (d; Glucose decline profile, s; Bioethanol production profile, j; Cell growth profile).
cell growth, sugar consumption and bioethanol generation profiles are shown in Fig. 2. After 12 h of bioethanol fermentation, the concentration of the reducing sugar decreased with time, and the concentration of bioethanol increase gradually, clearly indicating that the saccharification products of the residual biomass of D. tertiolecta LB999 were used as fermentation sugars. In the negative control experiment, negligible amount of ethanol was produced. On the contrary, when ethanol fermentation was conducted with the culture medium containing 17.5 g/L glucose and 12 g/L yeast extract as the positive control experiment, similar amount of ethanol was produced, supporting that the reducing sugars, mainly consisting of glucose, of the enzymatic saccharification products were successfully used for ethanol fermentation. The fermentation yields of bioethanol based on the residual biomass weight and the saccharified glucose were 0.14 g ethanol/g residual biomass and 0.44 g ethanol/g glucose, respectively. Without additional pretreatment steps after enzymatic saccharification, a high fermentation yield of bioethanol, 82% of the theoretical fermentation yield, was obtained. When taking it consideration that an unidentified monosaccharide existed at approximately 4.3% of total reducing sugars based on the ratio of HPLC peaks, the bioethanol fermentation yield from glucose was almost up to 85.7% of the theoretical fermentation yield. The unidentified monosaccharide was not used for ethanol fermentation by S. cerevisiae YPH500. Direct bioethanol fermentation without additional pretreatment
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steps would be helpful for decreasing the process cost by using the simplified bioethanol production process. 4. Conclusion The saccharification of microalgal residual biomass was successfully performed. AMG 300L-catalyzed saccharification produced 21.0 mg/mL of reducing sugar with 42.0% yield based on the residual biomass mass. The saccharification yield based on the total amount of carbohydrates of the residual biomass was 80.9% (w/w). Bioethanol was produced from the enzymatic saccharification products by S. cerevisiae YPH500, and the fermentation yield was 0.14 g ethanol/g residual biomass and 82.0% of the theoretical fermentation yield. This study clearly showed that the residual biomass of microalgae after lipid extraction for biodiesel production could be used for saccharification and subsequent bioethanol fermentation. Acknowledgements This work was supported by a grant from the Development of Marine-Bioenergy program funded by the Ministry of Land, Transport and Maritime Affairs of the Korean government. 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.2013.01. 007. References AOAC (Association of Analytical Chemists), 2000. Official Methods of Analysis of Association of Analytical Chemist. Horwitz, W., Gaithersburg, Maryland, USA. Boubekeur, S., Camougrand, N., Bunoust, O., Rigoulet, M., Guerin, B., 2001. Participation of acetaldehyde dehydrogenases in ethanol and pyruvate metabolism of the yeast Saccharomyces cerevisiae. Eur. J. Biochem. 268, 5057– 5065. Choi, S.P., Nguyen, M.T., Sim, S.J., 2010. Enzymatic pretreatment of Chlamydomonas reinhardtii biomass for ethanol production. Bioresour. Technol. 101, 5330–5336.
201
Demirbas, A., 2006. Global biofuel strategies. Energy Educ. Sci. Technol. 17, 27–63. Harun, R., Danquah, M.K., 2011. Influence of acid pre-treatment on microalgal biomass for bioethanol production. Process Biochem. 46, 304–309. Harun, R., Danquah, M.K., Forde, G.M., 2010. Microalgal biomass as a fermentation feedstock for bioethanol production. J. Chem. Technol. Biotechnol. 85, 199–203. Harun, R., Jason, W.S.Y., Cherrington, T., Danquah, M.K., 2011. Exploring alkaline pre-treatment of microalgal biomass for bioethanol production. Appl. Energy 88, 3464–3467. John, R.P., Anisha, G.S., Nampoothiri, K.M., Pandey, A., 2011. Micro and macroalgal biomass: a renewable source for bioethanol. Bioresour. Technol. 102, 186–193. Kim, Z., Lee, C.G., 2010. Photobioreactor for a large-scale marine microalgal culture using semi-permeable membrane. Korean Pat. 10-0991373. Kim, N.J., Li, H., Jung, K.S., Chang, H.N., Lee, P.C., 2011. Ethanol production from marine algal hydrolysates using Escherichia coli KO11. Bioresour. Technol. 102, 7466–7469. Lee, S.J., Oh, Y.H., Kim, D.H., Kwon, D.Y., Lee, C.G., Lee, J.W., 2011. Converting carbohydrates extracted from marine algae into ethanol using various ethanolic Escherichia coli strains. Appl. Biochem. Biotechnol. 164, 878–888. Libessart, N., Maddelein, M.L., Koornhuyse, N., Decq, A., Delrue, B., Mouille, G., D’Hulst, C., Ball, S., 1995. Storage, photosynthesis and growth: the conditional nature of mutations affecting starch synthesis and structure in Chlamydomonas. Plant Cell 7, 1117–1127. Lin, Y., Tanaka, S., 2006. Ethanol fermentation from biomass resources: current state and prospects. Appl. Microbiol. Biotechnol. 69, 627–642. Meher, L.C., Sagar, D.V., Naik, S.N., 2006. Technical aspects of biodiesel production by transesterification – a review. Renew. Sustain. Energy Rev. 10, 248–268. Miller, G.L., 1959. Use of dinitrosalicylic acid reagent for determination of reducing sugar. Anal. Chem. 31, 426–428. Miranda, J.R., Passarinho, P.C., Gouveia, L., 2012. Pre-treatment optimization of Scenedesmus obliquus microalga for bioethanol production. Bioresour. Technol. 104, 342–348. Nigam, P.S., Singh, A., 2011. Production of liquid biofuels from renewable resources. Prog. Energy Combust. 37, 52–68. Singh, A., Nigam, P.S., Murphy, J.D., 2011. Mechanism and challenges in commercialisation of algal biofuels. Bioresour. Technol. 102, 26–34. Sluiter, A., 2006. Determination of Structural Carbohydrates and Lignin in Biomass, Version 2006. National Renewable Energy Laboratory, USA.