Bioresource Technology 127 (2013) 422–428
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Biochemical features and bioethanol production of microalgae from coastal waters of Pearl River Delta Hui Guo a,1, Maurycy Daroch a,1, Lei Liu a, Guoyu Qiu a, Shu Geng a,c, Guangyi Wang a,b,⇑ a Shenzhen Engineering Laboratory for Algal Biofuel Technology Development and Application, School of Environment and Energy, Peking University Shenzhen Graduate School, Shenzhen 518055, China b Department of Microbiology, University of Hawaii at Manoa, Honolulu, HI 96822, USA c Department of Plant Sciences, University of California at Davis, CA 95616, USA
h i g h l i g h t s " Two native to Pearl River Delta algae were used for bioethanol production. " Both strains exhibited highest growth rates in aerated cultures. " Accumulation of cellular carbohydrates was highest in early stationary phase. " Highest bioethanol yield from S. abundans was obtained after two step hydrolysis.
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Article history: Received 29 June 2012 Received in revised form 1 October 2012 Accepted 4 October 2012 Available online 16 October 2012 Keywords: Microalgae diversity Microalgae cultivation Carbohydrate accumulation Algal bioethanol
a b s t r a c t This study describes identification, cultivation, monitoring of carbohydrate accumulation and bioethanol production from microalgal strains from the coastal waters of Pearl River Delta. Eighteen identified strains belong to the families Chlorellaceae, Scotiellocystoidaceae, Neochloridaceae, Selenastraceae and Scenedesmaceae. Of isolated strains Mychonastes afer PKUAC 9 and Scenedesmus abundans PKUAC 12 were selected for further biomass and ethanol production analysis. Comparison of three cultivation modes (stationary, shaken and aerated) resulted in the highest biomass productivity obtained for aerated cultures that yielded 0.09 g and 0.11 g dry weight per day per litre of medium for M. afer PKUAC 9 and S. abundans PKUAC 12, respectively. Carbohydrate accumulation monitored by FTIR showed that early stationary phase is optimal for biomass harvest. Microalgal biomass was successfully used as a carbohydrate feedstock for fermentative bioethanol production. S. abundans PKUAC 12 was superior feedstock for bioethanol production when pre-treated with the combination of dilute acid treatment and cellulase. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Biofuels hold much promise in mitigating climate change and resource depletion as well as enhancing energy security. First generation bioethanol seemed to be the most feasible short-term alternative to fossil fuels due to well established production technology. However, increased production of ethanol from edible crops (mainly sugarcane and corn) raised concerns about the Abbreviations: FTIR, Fourier transformed infrared spectroscopy; PKUAC, Peking University algae collection; GOD-POD, glucose oxidase-peroxidise; RFLP, restriction fragment length polymorphism. ⇑ Corresponding author. Address: Department of Microbiology, University of Hawaii at Manoa, Oceanography, 1000 Pope Road, MSB 205, Honolulu, HI 96822, USA. Tel.: +86 0755 26611617/+1 808 956 8021; fax: +86 0755 26035227/+1 808 956 5339. E-mail addresses: gywang@pkusz.edu.cn, guangyi@hawaii.edu (G. Wang). 1 These authors contributed equally to the publication. 0960-8524/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2012.10.006
impact of first generation bioethanol on food prices and increased deforestation (Cassman and Liska, 2007; Fargione et al., 2008). As a result attention turned into second generation bioethanol produced from lignocellulose feedstocks (e.g., waste biomass) and dedicated lignocellulose crops like Miscanthus that can be grown on marginal lands (Daroch and Mos, 2011; Hattori and Morita, 2010). So far biochemical conversion of these feedstocks to ethanol did not live up to the expectations. The main issue that hinders lignocellulose conversion to biofuels is its resistance to saccharification caused by high content of lignin. Despite years of research it still remains an area for optimisation and development (Agbor et al., 2011). Bioethanol produced from microalgal feedstock can be an alternative, as algal biomass is less resistant to conversion into simple sugars than plant biomass. Moreover, several microalgal species e.g. Chlorella vulgaris (Branyikova et al., 2011), Chlamydomonas reinhardtii UTEX 90 (Choi et al., 2010) accumulate their energy reserves in starch, which is an efficient carbohydrate
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feedstock and can be easily hydrolysed into glucose with both chemical and/or enzymatic methods. Algae also exhibit a number of other advantages when compared with terrestrial plants: they exhibit higher growth rates, do not require soil and can be grown all year around in some climates (Borowitzka, 1999). Moreover certain algal species can be grown in the absence of fresh water and perform very well in waste or saline water streams (Christenson and Sims, 2011). All those features enable algae to become a promising bioethanol feedstock of the future (Daroch et al., 2012). Autotrophic growth of algal biomass for fermentative ethanol production is most likely to succeed when native strains, already selected by evolution to dominate in certain site, are utilised. Two potential problems arise when considering utilisation of non-native strains in large-scale outdoor cultivation. Firstly, nonnative, especially genetically improved strains could potentially seriously harm ecosystems when released to the environment as contamination of neighbouring sites is unavoidable in open systems. Secondly, utilisation of non-native algal strains for large scale-cultivation, often results in instantaneous culture contamination with native strains that are already evolutionary better adapted for cultivation sites. Inability of maintaining a stable algae population and protecting it from predators and local strains are some of the biggest challenges for algae producers (Christenson and Sims, 2011; Singh and Dhar, 2011). To address these problems two methods are currently in use. Firstly, large volume inoculums are grown in photobioreactors to minimise risk of contamination before releasing them to open system for short-term outdoor cultivation (Christenson and Sims, 2011; Singh and Dhar, 2011). Major problem associated with this method is the cost of photobioreactors and water sterilisation system. Secondly, utilisation of extremophillic algae like Dundaliella that tolerate higher salinities (or other combination of extemophile-environmental pressure) can result in specie dominance (Borowitzka, 1999; Singh and Dhar, 2011). The problem associated with this approach is that although many algal strains can survive in such conditions their growth rates are usually lower compared with algae grown in milder conditions (Yeesang and Cheirsilp, 2011). Nevertheless, considering the diversity of microalgae in natural environment, microalgal collections from local habitats will greatly benefit algal biofuel industry development and can also alleviate environmental concerns related to large-scale production of algal biofuels. Although the coastal areas of Pearl River Delta have long been known for huge diversity of aquatic life surprisingly only little work has been done to assess the possibility of using local microalgae resources for biofuel production. Our study aimed to fill this niche and test the feasibility of producing renewable fuel ethanol from local algal strains. Here we describe isolation, identification, and utilisation of marine microalgal resource of Pearl River Delta as a carbohydrate feedstock for bioethanol production.
2. Methods 2.1. Algae sample collection Algae specimens were collected from surface shallow water at two sites in Shenzhen Mangrove Nature Reserve on 21.01.2011. Site 1 is situated at 22° 310 22.799400 N, 113° 590 35.700 E and Site 2 at 22° 310 23.62800 N, 113° 590 40.487400 E. Water samples were collected to sterile glass bottles (1 L) and transported to the laboratory within 1 h. Upon arrival water samples were concentrated to a hundred fold; divided into 3 sets, diluted in 3 series and plated onto BG11 and F/2 agar plates. Plates were incubated at 25 °C in 12 L:12 D photoperiod at 2000 lux for 14 days. Single colonies were picked by sterile toothpick, transferred to 100 mL of either BG11 or F/2 liquid medium and cultivated for additional 12 days with occasional agita-
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tion in the same conditions. A total of 214 isolates were obtained using this method. Redundant strains were excluded by a combination of morphological analysis under bright field microscope at 1000 magnification and restriction fragment length polymorphism (RFLP) of 18S rRNA gene as described in Section 2.3 to yield 18 unique isolates. Three strains (PKUAC 3, PKUAC 4, and PKUAC 9) were isolated from Shenzhen Mangrove Nature Reserve Site 1 using BG 11 medium and one strain, PKUAC 20 was isolated using F/2 medium from the same location. Six strains (PKUAC 1, PKUAC 2, PKUAC 10, PKUAC 12, PKUAC 13, and PKUAC 14) were isolated from Shenzhen Mangrove Nature Reserve Site 2 using BG 11 medium, whereas strains: PKUAC 5, PKUAC 6, PKUAC 7, PKUAC 15, PKUAC 17, PKUAC 21, PKUAC22 were isolated using F/2 medium from the same location. Unique strains were stored in Peking University Algae Collection (PKUAC) at 4 °C with limited lighting conditions. 2.2. Growth media Algae were grown in BG11 and F/2 media prepared essentially as described by Stanier et al. (1971) and Guillard and Ryther (1962), respectively. Agar plates were additionally supplemented with 1% (w/v) agar. All chemicals used for growth media preparation were of analytical grade. 2.3. Cultivation of selected algal strains Two algal strains Mychonastes afer PKUAC 9 and Scenedesmus abundans PKUAC 12 selected on the basis of preliminary screening for growth rate and cell size were pre-cultured in BG 11 medium. Cultures were grown for 7 days in 28 °C, 12 h of light and 12 h of dark (12 L:12 D) photoperiod at 2000 lux with occasional agitation until they reached exponential growth phase. Pre-cultures were used to inoculate in triplicates under three cultivation conditions: stationary, shaken (150 rpm) and aerated at approximate air flowrate of 4.5 L min 1. Seed cultures were diluted to 1.0 106 cell mL 1 (calculated by haemocytometer) in 450 mL of BG11 medium and grown in 500 mL conical flasks. Cultures were grown in 28 °C in 14 L: 10 D photoperiod at 2000 lux during the first 48 h and then at 4000 lux until stationary phase were achieved. 2.4. Genomic DNA extraction and 18S rRNA analysis About 100 mL of each alga was grown to mid-log phase and pelleted by centrifugation at 4000g. The resulting pellet was flash frozen in liquid nitrogen and ground to a fine powder. Genomic DNA was isolated with Plant Genomic DNA Isolation Kit [Generay, China] according to manufacturer’s guidelines. DNA (15 ng) was used as a template for 18S rRNA gene PCR amplification performed with Ready-to-use PCR Kit [Generay, China]. Amplification was performed in a heated-lid thermocycler with the primers 16S1N forward, 50 -TCCTGCCAGTAGTCATATGC-30 and 16S2N reverse, 50 -TGATCCTTCTCGCAGGTTCAC-30 (Grzebyk and Sako, 1998). PCR was performed under following cycling conditions: initial denaturation 94 °C for 5 min; followed by 40 cycles of 94 °C for 30 s, 50 °C for 30 s, 72 °C for 2 min; final extension 72 °C for 7 min. Resulting PCR products were separated on 1% (w/v) in TAE agarose gel, purified with TIANgel Midi DNA Purification Kit [TIANGEN, China] and TA cloned to pMD 18-T Vector [Takara, China]. Resultant plasmids were transformed into E. coli DH5a and plated on LB agar medium supplemented with ampicillin, IPTG and X-gal. White colonies were picked and grown in LB medium supplemented with 100 lg mL 1 ampicillin for plasmid isolation. Plasmids were isolated with AxyPrep™ Plasmid Miniprep Kit [Axygen, China] and sequenced at Beijing Genomics Institute, Shenzhen, China with following set of primers BcaBEST Primer RV-M and BcaBEST Primer M13–47 [Takara, China].
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2.5. Sequence and phylogenetic analysis Resultant sequences were analysed with BLASTN against nucleotide collection for species identification. Maximum likelihood phylogenetic tree of microalgae 18S rRNA gene was constructed with Mega 5 software. Numbers above branches indicate bootstrap values of maximum likelihood analysis from 1000 replicates. 2.6. Carbohydrate profiles of algal biomass and hydrolysates Carbohydrate profiles of lyophilised algal biomass were determined using following methods; starch was determined according to Branyikova et al. (2011); cellulose was estimated according to procedures of Association of Official Analytical Chemists (Horwitz, 1980); hemicellulose was determined as the difference between neutral detergent fibre (NDF) and acid detergent fibre (ADF) as described by Rezaeian et al. (2005). Soluble sugars were determined according to method described by Li (2000). Total carbohydrate content of algal hydrolysates was determined with anthrone method (McCready et al., 1950). Glucose content was analysed with enzymatic GOD-POD test [Biochem, China] against the standard curve prepared with glucose. 2.7. Determination of algal growth and dry weight Cell growth was monitored by measuring the optical density at 680 nm (OD680) using a UV–Vis spectrophotometer UV1800 [Shimadzu, Japan]. For dry biomass weight determination, 5 mL of algal culture was filtered through the pre-weighted glass fibre filter GF/C [Whatman, USA] and washed twice with deionised water. Resultant filter was oven dried at 80 °C for 24 h until the consistent weight is reached. 2.8. FTIR spectroscopy and determination of total protein and carbohydrates For FTIR spectroscopy analysis 2 mL of algal culture was collected, pelleted by centrifugation at 12,000g and washed twice with deionised water. Resultant pellets were oven dried at 40 °C for 24 h. Dry algal biomass was ground to a fine powder with KBr and pressed into tablets. FTIR spectroscopy was performed in quintuplicate in the range 4000–400 cm 1 using IR Prestige-21 [Shimadzu, Japan] spectrometer. Peak areas corresponding to carbohydrate (1190–950 cm 1) and protein absorption bands (1710–1620 cm 1) and were calculated with Shimadzu IRsolution 1.10 software. Carbohydrate and protein data obtained with FTIR spectroscopy were correlated with experimental data obtained with anthrone method (McCready et al., 1950) and bicinchoninic acid method (Smith et al., 1985) respectively. Protein content was determined with BCA Protein Assay Reagent [Pierce, USA] according to manufacturer’s instructions using a standard curve prepared with BSA [Pierce, USA]. 2.9. Algae saccharification and ethanol fermentation 2.9.1. Algae pre-treatment with acid Pre-treatment of algae was carried out by mixing 2.5 g of lyophilised algae with 50 mL of 3% sulphuric acid at 170 rpm for 5 min at room temperature. Mixture was subsequently incubated for 30 min at 110 °C in the autoclave. Prior to fermentation cellular debris was removed by centrifugation (1800g, 5 min) and pH of the supernatant was adjusted to 5.0 with solid CaCO3. 2.9.2. Amylase catalysed hydrolysis of acid pre-treated algal biomass After acid pre-treatment, pH of algal biomass was adjusted to pH 5.5 with solid NaOH and 10 mg of a-amylase [Solaribio, China]
(approx. 15 U/g algae) and 10 mg glucoamylase [NewTopBio, China] (approx. 0.4 U/g algae) were added to acid pre-treated samples. Enzymatic hydrolysis was performed for another 30 min at 55 °C with mixing. Prior to fermentation cellular debris was removed by centrifugation (1800g, 5 min) and pH of the supernatant was adjusted to 5.0 with solid CaCO3. 2.9.3. Cellulase catalysed hydrolysis of acid pre-treated algal biomass After acid pre-treatment, pH of algal biomass was adjusted to pH 5.0 with solid NaOH and 250 mg of cellulase from Trichoderma reesei ATCC 26921 [Sigma, China] (approx. 0.100 U per gram algae were added to acid pre-treated samples. Enzymatic hydrolysis was performed for another 30 min at 37 °C with mixing. Prior to fermentation cellular debris was removed by centrifugation (1800g, 5 min) and pH of the supernatant was adjusted to 5.0 with solid CaCO3. 2.9.4. Ethanol fermentation with S. cerevisiae Yeast Saccharomyces cerevisiae Hansen pre-culture was grown to mid-log phase in YPD (1% yeast extract, 2% peptone, and 2% glucose) medium at 30 °C, 200 rpm for 48 h. Algal hydrolysate was inoculated with 3% (v/v) pre-cultured yeast and cultured anaerobically at 30 °C, 200 rpm for 48 h. Fermentation was performed in sealed fermentation flasks sparged with nitrogen for 20 min. before autoclaving to remove oxygen. For the determination of ethanol, total sugars and glucose content, fermentation broths were sampled at 8, 16, 24 and 48 h. 2.9.5. Ethanol determination Ethanol concentration in the fermentation broth was determined using gas chromatography GC 2010 [Shimadzu, Japan] with autosampler and flame ion detector (FID) and HP-INNOWAX column (15 m 0.530 mm 1.00 m). The column oven was set to 70 °C equilibration for 2.0 min and then 20 °C min 1 increase to 120 °C, and 40 °C min 1 to 200 °C; split ratio 10:1; injection temperature 250 °C, FID temperature 250 °C. Nitrogen was used as the carrier gas. Ethanol retention time was determined as 0.784 min. 3. Results and discussion 3.1. Isolation of Pearl River Delta algae A total of 214 cultivable microalgal strains were isolated using BG-11 and F/2 growth media supplemented with agar from two sites situated in Shenzhen Mangrove Nature Reserve. Redundant strains were excluded by a combination of morphological analysis with bright field microscopy and restriction fragment length polymorphism (RFLP) of 18S rRNA gene sequence amplified with 16S1N and 16S2N primers (Grzebyk and Sako, 1998). A total of 18 unique strains were identified and selected for phylogenetic analysis (Fig. 1). Two thirds of isolated algal strains belong to family Chlorellaceae, two representatives of Scotiellocystoidaceae were also identified, the remaining individual representatives of other taxons belonged to Neochloridaceae, Selenastraceae, Scenedesmaceae. Single strain (Chlorophyceae sp. PKUAC 10) could not be classified at family level on the basis of its 18S rRNA gene sequence analysis. Phylogenetic tree of all 18 unique strains is presented in Fig. 1. 3.2. Growth of selected algal species in laboratory scale and determination of their carbohydrate accumulation Initial screening of all isolated algae indicated two strains M. afer PKUAC 9 and S. abundans PKUAC 12 as promising feedstocks for fermentative production of bioethanol due to high growth
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Fig. 1. Maximum likelihood phylogenetic tree of 18S rRNA sequences of microalgae isolated from Shenzhen Mangrove Natural Reserve. Numbers above branches indicate bootstrap values of maximum likelihood analysis from 1000 replicates. Underlined strains were selected for bioethanol production study.
rates; cell size and sugar content (data not shown). To test different methods of strain cultivation algae growth was monitored for
24 days in three cultivation modes: stationary, shaken at 150 rpm and aerated (Fig. 2). Aerated cultures yield more biomass
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Fig. 2. Growth curves and determination of carbohydrate accumulation by selected strains of Pearl River Delta algae. Scenedesmus abundans PKUAC 12 (A) and Mychonastes afer PKUAC 9 (B) cultivated using 3 different modes (aerated (rhombes), shaken (squares) and stationary (triangles)) for 24 days at 28 °C. Determination of carbohydrate accumulation was performed with FTIR. S. abundans PKUAC 12 (C) and M. afer PKUAC 9 (D). Growth curve of aerated cultures (rhombes), ratio of FTIR peak areas corresponding to sugar and protein (S/P) as described in Section 2.8.
than other cultivation modes with the average yields of 0.09 g and 0.11 g dry weight per day per litre of medium for M. afer PKUAC 9 and S. abundans PKUAC 12, respectively. Their respective biomass concentrations achieved 1.95 g L 1 and 1.7 g L 1. Growth rates for S. abundans PKUAC 12 strains are generally within the range of other Scenedesmaceae reported to be 0.10 to 0.26 g L 1 d 1 (de Morais and Costa, 2007; Griffiths and Harrison, 2009; Rodolfi et al., 2009). Within this range higher productivities were obtained using CO2 enriched air during cultivation. Biomass concentration for the strain reported in this study S. abundans PKUAC 12 cultivated in atmospheric concentration of CO2 was reported to achieve 1.7 g L 1 after 24 days of cultivation a value very similar to S. obliquus (1.8 g L 1) which was achieved in photobioreactor fed with 12% CO2 after 21 days of cultivation. Little is known about productivities of Mychonastes strains. Similar biomass concentrations, approximately 1.2 g L 1 were obtained in unmodified BG 11 medium for HSO-3–1 strain (Yuan et al., 2011) and 1.95 g L 1 for PKUAC 9 strain. Higher biomass yield of M. afer HSO-3–1 strain was achieved (3.29 g L 1) when cultivated in optimised BG11 medium containing 3.15 g L 1 NaNO3. M. afer PKUAC 9 strain has not been grown in these conditions. Cellular protein and carbohydrates of these two strains were monitored by FTIR spectroscopy at: 72, 96, 120, 144, 240, 360, 480 h. Dried biomass mixed with KBr was analysed by FTIR spectroscopy at 4000–400 cm 1. Resultant data were correlated with those obtained for carbohydrate content and protein content determined with anthrone reagent and BCA method, respectively. The correlation between FTIR and chemical methods was 0.999 and 0.911 for M. afer and S. abundans, respectively. Correlation coefficient between FTIR and chemical methods computed as a ratio of total cellular sugar to total cellular protein was found optimal for both strains when carbohydrate peak area was integrated in the range of 1190–950 cm 1 for carbohydrate and 1710–1620 cm 1 for protein. These ranges were previously ascribed to carbohydrates and amide I absorption regions, respectively (Dean et al., 2010; Stehfest et al., 2005). Changes of sugar/
protein ratio during algae cultivation are summarised in Fig. 2. Monitoring of sugar accumulation indicated that carbohydrate content decreased in newly-started cultures of both strains. The decrease corresponds to lag phase in algal growth and is likely ascribed to adaptation to new growth conditions. Sugar accumulation starts rising after algal culture reaching mid exponential phase. Both strains start accumulating sugars again until they reach early stationary phase when they reach stable values and are harvested. Early stationary phase was previously found optimal for starch productivity using C. reinharditi (Nguyen et al., 2009; Thyssen et al., 2001) and our results are consistent with these findings. Results of this study also support previous report of FTIR spectroscopy as a tool to track changes of cellular composition of algal feedstock for biofuel production (Dean et al., 2010). Carbohydrate profiles of both algae were determined to identify optimal methods of biomass saccharification (Table 2). S. abundans PKUAC 12 biomass is mostly composed of hemicellulose at 32.07% (w/w). Each of other carbohydrates i.e. cellulose, starch and soluble sugars are below 5% of dry cellular weight. M. afer PKUAC 9 biomass is composed of hemicellulose 17.10% (w/w) and starch 7.54% (w/w). It was expected that S. abundans PKUAC 12 biomass will be efficiently hydrolysed with a two stage treatment of sulphuric acid and cellulase mixture from T. reesei ATCC 26921. Cellulases and hemicellulases in nature act in synergy to degrade lignocellulose and both activities were found to be exhibited simultaneously in T. reesei (HerpoelGimbert et al., 2008). Higher content of starch in M. afer PKUAC 9 suggested efficient hydrolysis with a combination of sulphuric acid and amylases. 3.3. Biomass hydrolysis and ethanol fermentation This work is one of few that focus in exploration of native biological resource for bioethanol production (Hon-Nami, 2006; Ueno et al., 1998) rather than utilising laboratory strains (Choi et al.,
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H. Guo et al. / Bioresource Technology 127 (2013) 422–428 Table 1 Concentration of carbohydrates in hydrolysates of S. abundans PKUAC 12 and M. afer PKUAC 9. Strain
Pretreatment method H2SO4
H2SO4 + amylases
H2SO4 + cellulase
S. abundans PKUAC 12
7.409 4.422
7.734 4.016
10.752 5.730
Total sugars [gL 1] Glucose [gL 1]
M. afer PKUAC 9
5.133 1.206
6.057 2.766
6.223 2.047
Total sugars [gL 1] Glucose [gL 1]
Table 2 Summary of basic parameters of S. abundans PKUAC 12 and M. afer PKUAC 9 biomass and their carbohydrate profiles upon harvest. Strain
Cell size [lm]
Biomass concentration [gL 1]
Growth rate [g/day]
Soluble sugars [% (w/w)]
Starch [% (w/w)]
Cellulose [% (w/w)]
Hemicellulose [% (w/w)]
Protein content [% (w/w)]
S. abundans PKUAC 12 M. afer PKUAC 9
4.5
1.70
0.11
3.04
1.51
4.35
32.07
35.98
2.4
1.95
0.09
3.10
7.54
0.64
17.10
20.93
Fig. 3. Fermentative production of ethanol using Scenedesmus abundans PKUAC 12 and Mychonastes afer PKUAC 9 biomass. Diluted H2SO4 treatment of S. abundans PKUAC 12 (A) and M. afer PKUAC 9 (D); amylase treatment of acid hydrolysed fraction of S. abundans PKUAC 12 (B) and M. afer PKUAC 9 (E); cellulase treatment of acid hydrolysed fraction of S. abundans PKUAC 12 (C) and M. afer PKUAC 9 (F). Concentration of: total sugars (rhombes), glucose (squares) and ethanol (triangles) from fermentation broth are presented in g L 1.
2010; Nguyen et al., 2009) or biomass obtained from algae-producing companies (Kim et al., 2012). Biomass cultivation for fermentative ethanol production was performed in 10 L laboratory photobioreactor until early stationary phase was achieved for both strains. Algal biomass was harvested by centrifugation, lyophilised and then pre-treated with dilute sulphuric acid. Hydrolysis was enhanced by two different enzymatic treatments: with amylolytic or cellulolytic enzymes. Results of algal biomass saccharification are presented in Table 1. Release of sugars was the most effective when S. abundans was sequentially hydrolysed with dilute sulphuric acid and cellulase mixture to yield 10.752 g L 1 of total sugars and 5.730 g L 1 of glucose in hydrolysate. The most efficient method of saccharifying M. afer was sequential treatment with dilute sulphuric acid and a combination of a- and glucoamylases. This hydrolysate contained 6.057 g L 1 of total sugars and 2.766 g L 1
of glucose. Efficient biomass hydrolysis by amylolytic enzymes indicates higher starch accumulation by that strain than that of S. abundans what is also confirmed by analysing carbohydrate profiles of both algae (Table 2). Both enzymatic treatments increase the concentration of total sugars and glucose; however the overall impact of two-step treatment is rather low at these enzyme loadings and reaction times. Previous studies also indicated that two-step hydrolysis enhanced algal biomass saccharification (Kim et al., 2011; Wang et al., 2011). Different methods of biomass hydrolysis had an effect on overall efficiency of bioethanol production (Table 1 and Figs. 3 and 4). The highest yield of bioethanol production (0.103 g of ethanol per gram of dry weight algae) was obtained after treating biomass of S. abundans with dilute sulphuric acid and cellulase. The reported yield is average with respect to experiments elsewhere. Similar studies report between 0.05 g (Kim
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Fig. 4. Yield of bioethanol production (gram of ethanol per gram of dry algae) after 24 h of fermentation with Saccharomyces cerevisiae Hansen from Scenedesmus abundans PKUAC 12 and Mychonastes afer PKUAC 9 biomass using different biomass pre-treatments.
et al., 2012) and 0.4 g (Lee et al., 2011) of ethanol per gram of dry weight algae, it should be noted, however that latter result was obtained using E. coli SJL2526 strain genetically engineered with additional sugar uptake proteins. Typical ethanol fermentation yields from microalgal feedstock using S. cerevisiae do not exceed 0.3 g ethanol per gram of dry weight algae (Choi et al., 2010; Nguyen et al., 2009). Both of these studies utilised laboratory strain C. reinhardtii UTEX 90 as carbohydrate feedstock which may not perform equally well in open systems in sites remote to its natural environment. The course of fermentation is presented in Fig. 3. Glucose as a preferred source of energy was utilised relatively fast, usually within 6–9 h, the subsequent raise in ethanol production must have resulted from utilisation of other carbohydrates from hemicellulose fraction of biomass and resulted in slower production of ethanol. 4. Conclusions Microalgal strains M. afer PKUAC 9 and S. abundans PKUAC 12 isolated from coastal waters of Pearl River Delta region were successfully used for saccharification and subsequent fermentative bioethanol production. Saccharification of algal biomass resulted in the highest release of total sugars of 10.752 g L 1 hydrolysate and glucose 5.730 g L 1 hydrolysate from S. abundans PKUAC 12 biomass harvested in early stationary phase. Dilute acid and cellulase treated S. abundans PKUAC 12 biomass was the best feedstock for fermentative bioethanol production yielding 0.103 g of ethanol per gram of dry weight algae. Acknowledgements This project was partially funded by Shenzhen Development and Reform Commission grant # 835 (2011)(GYW) and NSFC grant # 31170109 (GYW). References Agbor, V.B., Cicek, N., Sparling, R., Berlin, A., Levin, D.B., 2011. Biomass pretreatment: fundamentals toward application. Biotechnol. Adv. 29, 675–685. Borowitzka, M.A., 1999. Commercial production of microalgae: ponds, tanks, tubes and fermenters. J. Biotechnol. 70, 313–321. Branyikova, I., Marsalkova, B., Doucha, J., Branyik, T., Bisova, K., Zachleder, V., Vitova, M., 2011. Microalgae-novel highly efficient starch producers. Biotechnol. Bioeng. 108, 766–776. Cassman, K.G., Liska, A.J., 2007. Food and fuel for all: realistic or foolish? Biofuel Bioprod. Biorefin. 1, 18–23. Choi, S.P., Nguyen, M.T., Sim, S.J., 2010. Enzymatic pretreatment of Chlamydomonas reinhardtii biomass for ethanol production. Bioresour. Technol. 101, 5330–5336.
Christenson, L., Sims, R., 2011. Production and harvesting of microalgae for wastewater treatment, biofuels, and bioproducts. Biotechnol. Adv. 29 (6), 686–702. Daroch, M., Geng, S., Wang, G., 2012. Recent advances in liquid biofuel production from algal feedstocks. Appl. Energy, <http://dx.doi.org/10.1016/j.apenergy. 2012.07.031>. Daroch, M., Mos, M., 2011. Role of biotechnology in developing the bioenergy sector in Poland. BioTechnologia 92, 23–32. de Morais, M.G., Costa, J.A.V., 2007. Biofixation of carbon dioxide by Spirulina sp. and Scenedesmus obliquus cultivated in a three-stage serial tubular photobioreactor. J. Biotechnol. 129 (3), 439–445. Dean, A.P., Sigee, D.C., Estrada, B., Pittman, J.K., 2010. Using FTIR spectroscopy for rapid determination of lipid accumulation in response to nitrogen limitation in freshwater microalgae. Bioresour. Technol. 101, 4499–4507. Fargione, J., Hill, J., Tilman, D., Polasky, S., Hawthorne, P., 2008. Land clearing and the biofuel carbon debt. Science 319, 1235–1238. Griffiths, M.J., Harrison, S.T.L., 2009. Lipid productivity as a key characteristic for choosing algal species for biodiesel production. J. Appl. Phycol. 21 (5), 493–507. Grzebyk, D., Sako, Y., 1998. Phylogenetic analysis of nine species of Prorocentrum (Dinophyceae) inferred from 18S ribosomal DNA sequences, morphological comparisons, and description of Prorocentrum panamensis, sp. nov. J. Phycol. 34, 1055–1068. Guillard, R.R., Ryther, J.H., 1962. Studies of marine planktonic diatoms. I. Cyclotella nanaHustedt, and Detonula confervacea (cleve). Gran. Can. J. Microbiol. 8, 229– 239. Hattori, T., Morita, S., 2010. Energy crops for sustainable bioethanol production; which, where and how? Plant Prod. Sci. 13, 221–234. Herpoel-Gimbert, I., Margeot, A., Dolla, A., Jan, G., Molle, D., Lignon, S., Mathis, H., Sigoillot, J.-C., Monot, F., Asther, M., 2008. Comparative secretome analyses of two Trichoderma reesei RUT-C30 and CL847 hypersecretory strains. Biotechnol. Biofuels 1 (1), 18. Hon-Nami, K., 2006. A unique feature of hydrogen recovery in endogenous starchto-alcohol fermentation of the marine microalga, Chlamydomonas perigranulata. Appl. Biochem. Biotechnol. 131 (1–3), 808–828. Horwitz, W., 1980. Official methods of analysis of the association of official analytical chemists, thirteenth ed. Association of Official Analytical Chemists, Washington. Kim, J., Um, B.-H., Kim, T., 2012. Bioethanol production from micro-algae, Schizocytrium sp., using hydrothermal treatment and biological conversion. Korean J. Chem. Eng. 29, 209–214. Kim, N.J., Li, H., Jung, K., Chang, H.N., Lee, P.C., 2011. Ethanol production from marine algal hydrolysates using Escherichia coli KO11. Bioresour. Technol. 102, 7466– 7469. Lee, S., Oh, Y., Kim, D., Kwon, D., Lee, C., Lee, J., 2011. Converting carbohydrates extracted from marine algae into ethanol using various ethanolic Escherichia coli Strains. Appl. Biochem. Biotechnol. 164, 878–888. Li, H., 2000. Principles and techniques of plant physiological biochemical experiment, first ed. Higher Education Press, Beijing. McCready, R.M., Guggolz, J., Silviera, V., Owens, H.S., 1950. Determination of starch and amylose in vegetables. Anal. Chem. 22, 1156–1158. Nguyen, M.T., Choi, S.P., Lee, J., Lee, J.H., Sim, S.J., 2009. Hydrothermal acid pretreatment of Chlamydomonas reinhardtii biomass for ethanol production. J. Microbiol. Biotechnol. 19, 161–166. Rezaeian, M., Beakes, G.W., Chaudhry, A.S., 2005. Relative fibrolytic activities of anaerobic rumen fungi on untreated and sodium hydroxide treated barley straw in in vitro culture. Anaerobe 11 (3), 163–175. Rodolfi, L., Zittelli, G.C., Bassi, N., Padovani, G., Biondi, N., Bonini, G., Tredici, M.R., 2009. Microalgae for oil: strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol. Bioeng. 102 (1), 100–112. Singh, N.K., Dhar, D.W., 2011. Microalgae as second generation biofuel. A review. Agron. Sustain. Dev. 31 (4), 605–629. Smith, P.K., Krohn, R.I., Hermanson, G.T., Mallia, A.K., Gartner, F.H., Provenzano, M.D., Fujimoto, E.K., Goeke, N.M., Olson, B.J., Klenk, D.C., 1985. Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76–85. Stanier, R.Y., Kunisawa, R., Mandel, M., Cohen-Bazire, G., 1971. Purification and properties of unicellular blue–green algae (order Chroococcales). Bacteriol. Rev. 35, 171–205. Stehfest, K., Toepel, J., Wilhelm, C., 2005. The application of micro-FTIR spectroscopy to analyze nutrient stress-related changes in biomass composition of phytoplankton algae. Plant Physiol. Biochem. 43, 717–726. Thyssen, C., Schlichting, R., Giersch, C., 2001. The CO2-concentrating mechanism in the physiological context: lowering the CO2 supply diminishes culture growth and economises starch utilisation in Chlamydomonas reinhardtii. Planta 213, 629–639. Ueno, Y., Kurano, N., Miyachi, S., 1998. Ethanol production by dark fermentation in the marine green alga, Chlorococcum littorale. J. Ferment. Bioeng. 86 (1), 38–43. Wang, X., Liu, X.H., Wang, G.Y., 2011. Two-stage hydrolysis of invasive algal feedstock for ethanol fermentation. J. Integr. Plant Biol. 53, 246–252. Yeesang, C., Cheirsilp, B., 2011. Effect of nitrogen, salt, and iron content in the growth medium and light intensity on lipid production by microalgae isolated from freshwater sources in Thailand. Bioresour. Technol. 102 (3), 3034–3040. Yuan, C., Liu, J.H., Fan, Y., Ren, X.H., Hu, G.R., Li, F.L., 2011. Mychonastes afer HSO-3-1 as a potential new source of biodiesel. Biotechnol. Biofuels 4.