Induction of lipids and resultant FAME profiles of microalgae from coastal waters of Pearl River Delta

Bioresource Technology 146 (2013) 192–199

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Induction of lipids and resultant FAME profiles of microalgae from coastal waters of Pearl River Delta Maurycy Daroch a, Congcong Shao a, Ying Liu a, Shu Geng a, Jay J. Cheng 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 Biological and Agricultural Engineering, North Carolina State University, Raleigh, NC 27695, USA

h i g h l i g h t s  Thirty-seven algal strains were isolated from coastal waters of Pearl River Delta.  Hindakia strain native to Pearl River Delta was selected for FAME production.  Two methods of lipid induction were used: salt stress and nitrogen starvation.  Salt stress yields three-fold higher lipid productivity than nitrogen starvation.

a r t i c l e

i n f o

Article history: Received 3 May 2013 Received in revised form 10 July 2013 Accepted 13 July 2013 Available online 19 July 2013 Keywords: Microalgae Diversity Cultivation Oil accumulation Algal biodiesel

a b s t r a c t This article presents a study on identification, cultivation and characterisation of microalgal strains from the coastal waters of the Pearl River Delta in Guangdong, China. Thirty-seven identified strains belong to the families: Chlorellaceae, Scotiellocystoidaceae, Scenedesmaceae,Selenastraceae,Micractiniaceae, Coccomyxaceae, Trebouxiaceae and Chlorococcaceae. Of isolated strains, Hindakia PKUAC 169 was selected for lipid induction using two methods: nitrogen starvation and salt stress. After derivatisation of algal lipids through in situ transesterification, lipid profiles of the alga under the two methods were analysed. The results have shown that both lipid yield and fatty acid profiles vary with the methods. Of the two tested methods of inducing lipid production, salt stress yielded three-fold higher lipid productivity than nitrogen starvation. The lipids are predominantly composed of C14–C18 fatty acids, which are favourable for biodiesel production. Moreover, the content of polyunsaturated fatty acids was below the limit of 12% set by EN14214 biodiesel standard. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Biofuels are a promising solution in addressing global climate change and enhancing energy security. To date, three generations of biodiesel feedstocks have been utilised. First generation biodiesel, i.e., biodiesel produced from refined vegetable oils such as rapeseed, soybean and palm oils, is currently a major contributor to biodiesel production worldwide (OECD-FAO, 2012). However, increased production of biodiesel from these feedstocks is believed to have driven the prices of vegetable oil up and have negative im-

Abbreviations: PKUAC, Peking University Algae Collection; RFLP, restriction fragment length polymorphism; FAME, fatty acid methyl esters. ⇑ Corresponding author at: Shenzhen Engineering Laboratory for Algal Biofuel Technology Development and Application, Peking University-Shenzhen Graduate School, Shenzhen 518055, China. Tel.: +86 0755 2661 1617; fax: +86 0755 2603 5227. E-mail address: [email protected] (J.J. Cheng). 0960-8524/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biortech.2013.07.048

pact on both direct and indirect land use practices as more land is being converted to biofuel production. This problem is often encountered in tropical areas where large hectarage has been converted into oil palm plantations in recent years (Fitzherbert et al., 2008). Second generation biodiesel crops that can be grown on marginal lands such as Jatropha curcas L. have not lived up to initial expectations so far and their oil yields rarely approach claims from early reports (Jongschaap et al., 2007). Biodiesel production from microalgal biomass has long been seen as an alternative to first and second generation of biodiesel crops. Many microalgal strains accumulate their reserves as triglycerides that can be used for biodiesel production. Moreover, algae exhibit a number of advantages when compared with terrestrial crops: they exhibit higher growth rates, do not require soil, and can be grown year-round under certain climate conditions (Borowitzka, 1999). Moreover, many algal strains can be grown in waste or saline water streams, a significant advantage over terrestrial crops that rely on often limited freshwater resource

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(Christenson and Sims, 2011). All these features have made microalgae a very promising biodiesel feedstock of the future (Chisti, 2007; Daroch et al., 2013). Many microalgae contain a small amount of triglycerides during their optimal growth conditions as most of the resources are used for cellular growth. They are capable however, of accumulating considerable amount of triglycerides upon induction of cellular stress (Rodolfi et al., 2009). According to many studies triglycerides accumulated by microalgae are mainly composed of saturated and monounsaturated fatty acids which can be efficiently packed into the cell and provide energy reserves for rebuilding the cell after the stress is withdrawn (Roessler, 1990). These types of fatty acids are, in principle adequate substrate for biodiesel production as they offer higher cetane numbers and better ignition quality. It is known however that different growth conditions and methods of inducing oil accumulation can significantly affect fatty acid composition inside the cell (Knothe, 2011). Algal lipid profiles may vary from those rich in monosaturated fatty acids which are useful for biodiesel production (Knothe, 2008), to long polyunsaturated fatty acids which are unlikely to be a suitable feedstock for biodiesel production due to poor oxidative stability (Knothe, 2011). Induction of triglyceride accumulation usually requires modifying algae growth conditions from nutrient replete growth medium. Nutrient deficiency: nitrogen, silicate and phosphorus in particular, has been regarded as an efficient trigger of oil accumulation in algae since mid twentieth century (Spoehr and Milner, 1949). Despite the higher oil content in nutrient-starved biomass, overall oil productivity is often lower as biomass productivities under starvation conditions drop significantly (Rodolfi et al., 2009). It has been also known that other forms of stress, e.g., salt stress can also affect both growth rate and lipid content in various algae (Ben-Amotz et al., 1985) even between the strains of the same species (Siaut et al., 2011) and these findings are more rare than reports regarding nutrient deficiency studies. There exist two main strategies for cultivating microalgae on a large scale: open systems and photobioreactors (Chisti, 2007). Utilisation of a simple salt trigger to increase lipid production in microalgal strains would benefit both designs since energy intensive centrifugation steps can be replaced by cultivating algae in lipid inducing medium triggered by higher salt concentration. Moreover, utilisation of such a salt trigger in coastal areas can be performed by cultivating appropriately selected microalgal strains in seawater enriched with trace elements, which would significantly lower the freshwater footprint of algae to biofuel process. Pearl River Delta coastal areas have long been known for the diverse of its aquatic life and significant environmental advantages of using local strains for biofuel production (Guo et al., 2013). However, little work has been done to assess the possibility of using the microalgae resources in this local area for biodiesel production. This study explores the utilisation of native biological resource of Pearl River Delta for the production of fatty acid methyl esters under different lipid accumulation inducers: nitrogen starvation and salt trigger.

2. Methods 2.1. Algae sample collection Algae specimens were collected at three locations around the Shenzhen offshore area. At each site samples were collected from surface shallow water and water 5 m deep on 18 July 2011. Site 1 is situated at 22°310 0000 N, 113°590 5900 E, Site 2 at 22°260 0000 N,113°530 1200 E and Site 3 at 22°290 3000 N, 113°570 0100 E. Water samples were collected in sterile glass bottles (1 L) and transported to the laboratory within 1 h. Upon arrival water

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samples were concentrated to a hundred fold, divided into 3 sets, diluted in 2 series (undiluted and ten-fold dilution) and plated onto Kuhl and soil-extract (SE) 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 SE or Kuhl liquid medium and cultivated for additional 14 days with occasional agitation under the same conditions. A total of 285 isolates were obtained using this method. Redundant isolates were excluded by a combination of morphological analysis under bright field microscope at 1000 magnification and restriction fragment length polymorphism (RFLP). The 18S rRNA gene sequence region was amplified with 519 forward/1406 reverse primer pair as described in section 2.4 of Methods and digested using PstI and TaqI enzymes. Unique RFLP patterns yielded 37 isolates. 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 Kuhl and soil extract media prepared as described by Kuhl, 1962 and Starr and Zeikus, 1993, respectively. BG11 medium was prepared as described by Stanier et al., 1971, nitrogen starved BG11 (BG11N) was modified by removing NaNO3 from the recipe and salt induced medium (BG11 + NaCl) contained NaCl at the final concentration of 150 mM. 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 isolated algal strains Isolated 37 algal strains were pre-cultured in BG 11 medium. Cultures were grown for 14 days at 25 °C under constant illumination at 2000 lux with 80 rpm agitation in Moma TS2112B incubator [Chengdu Scientific, China] equipped with fluorescent lamps until they reached the exponential growth phase. The incubator was used throughout the entire experiment, unless specifically stated otherwise. Pre-cultures were used to inoculate in triplicates shake flask cultures to 1.0  106 cells mL1 (calculated by haemocytometer) in 100 mL of BG11 medium and grown in 150 mL conical flasks. Cultures were grown at 25 °C in constant illumination at 2000 lux. 2.4. Genomic DNA extraction and 18S rRNA analysis About 100 mL cultures of each isolate were grown to mid-log phase and pelleted by centrifugation at 3220g. The resultant pellets were flash frozen in liquid nitrogen and ground to a fine powder. Genomic DNA was isolated with a 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 a Ready-to-use PCR Kit [Generay, China]. Amplification was performed in a heated-lid thermocycler with the primers 519 forward, 50 - CAGCAGCCGCGGTAATAC-30 and 1406 reverse, 50 - ACGGGCGGTGTGTRC-30 (Zhu et al., 2008). PCR was performed under the following cycling conditions: initial denaturation 94 °C for 5 min; followed by 30 cycles of 94 °C for 1 min., 56 °C for 1 min., 72 °C for 2.5 min. and final extension 72 °C for 10 min. Resulting PCR products were separated on 1% (w/v) in TAE agarose gel, purified Gel Extraction Kit [Biomiga, China] and sequenced at Beijing Genomics Institute, Shenzhen, China with 519 forward/1406 reverse primer pair. 2.5. Sequence and phylogenetic analysis Resultant DNA sequences were analysed with BLASTN against nucleotide collection for species identification. Maximum

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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. 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 Nanophotometer P300 [Implen, Germany]. For dry biomass weight determination, 100 mL of algal culture was filtered through the pre-weighted glass fibre filter GF/C [Whatman, USA] and washed twice with deionised water. The resultant filter was oven dried at 65 °C for 12 h until a consistent weight was reached. 2.7. Cultivation of Hindakia sp. PKUAC 169, induction and monitoring of oil accumulation Hindakia sp. PKUAC 169 was pre-cultured in BG 11 medium. Cultures were grown in laboratory room temperature at 22 °C, at constant illumination of 2000 lux at 80 rpm until they reached exponential growth phase. Pre-cultures were used to inoculate in triplicates in three cultivation media: unmodified, nitrogen starved and salt induced. For each of these growth media Hindakia sp. PKUAC 169strain was pre-cultured in BG 11 medium, pelleted by centrifugation at 3220g, washed twice with sterile distilled water and resuspended in 900 mL of fresh growth medium (BG11, BG11N and BG11 + NaCl) in 1000 mL Erlenmeyer flask at the final concentration corresponding to 0.035 g (dry weight)/L. Cultures in these three growth media were cultivated under aerated conditions at the approximate air flow-rate of 4.5 L min1 for 12 days at room temperature (approximately 22 °C, maintained by the laboratory air conditioning unit), at constant illumination of 2000 lux provided by fluorescent lamps. Lipid accumulation was monitored by fluorescence microscopy as described by Siaut et al., 2011. 2.8. In situ biodiesel production and FAME determination In situ transesterification of algal lipids was performed with modified protocol of Johnson and Wen (2009). Freeze-dried algal biomass (0.02 g) was placed in a glass test tube and mixed with 1.7 mL of methanol, 0.3 mL of sulphuric acid and 2 mL of hexane. The reaction mixture was heated at 90 °C for 1 h and the samples were well-mixed during heating. After the reaction, the tubes were allowed to cool to room temperature, and 2 mL distilled water was added, vortexed and centrifuged for 30 min at 3220g. The hexane layer that contained FAME was collected and transferred to a preweighed glass vial. The solvent was evaporated using N2, and the mass of FAMEs were determined gravimetrically. FAME profiles were determined using gas chromatography Agilent 7890A [Agilent, US] with auto-sampler, flame ion detector (FID) and DB5-MS column (60 m  250 lm  0.25 lm). The column oven was set to 70 °C equilibration for 2.0 min and then 10 °C min1 increase to 230 °C, 3 °C min1 to 290 °C then hold 5 min; splitless injection; injection temperature 280 °C. Helium was used as the carrier gas. 3. Results and discussion 3.1. Isolation of Pearl River Delta algae and phylogeny Two hundred eighty-five cultivable microalgae were isolated using soil extract and Kuhl growth media supplemented with agar from three sites situated in the Shenzhen Bay area. Redundant isolates 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 a 519 forward/1406 reverse primer pair (Zhu et al., 2008). A total of 37 unique strains were identified and selected for phylogenetic analysis (Figs. 1 and 2). The majority of strains were identified as Chlorella sp. (Fig. 1A) with a percentage similar to the previous study conducted at the Shenzhen Mangrove Nature Reserve (Guo et al., 2013). The main difference between species isolated from the two sites was observed among Nanochloris sp. and Nannochlorum sp., which were major species isolated during the previous study (50% of total isolated strains from Shenzhen Mangrove Nature Reserve). These members were represented by a unique strain in this study. As reported in the previous study, all strains of Nanochloris sp. and Nannochlorum sp. were isolated using F/2 medium which contains a much higher concentration of salt than growth media used in this paper. The difference is most likely due to selectivity of F/2 growth medium towards marine strains and does not represent dominance of these species at the site of study. Analysis of algal families reveals that over 30% of isolated algal strains belong to the family Chlorellaceae, whereas over 20% represent Scotiellocystoidaceae. Other families were present in the minority and include: Scenedesmaceae,Selenastraceae,Micractiniaceae, Coccomyxaceae, and Trebouxiaceae (Fig. 1B). A phylogenetic tree of all 37 unique strains is presented in Fig. 2. Phylogenetic analysis revealed four major groups: Group I: Scotiellocystoidaceae,Chlorococcaceae; Group II: Coccomyxaceae,Selenastraceae; Group III: Scenedesmaceae; and Group IV:Chlorellaceae,Trebouxiaceae,Micractiniaceae. Overall tree topology is similar to the previous study (Guo et al., 2013). Basic morphological features of isolated algae are summarised in Table 1 and the majority of isolated strains are round unicellular organisms ranging from 2 to 11 lm in diameter. 3.2. Growth of selected algal strains in laboratory scale and determination of their biomass productivity Preliminary results suggested that strains isolated with Kuhl and soil extract media performed better when using BG 11 growth medium. Moreover BG 11 medium is better suited to nitrogen starvation modification than either of the growth media used for isolation. All isolated strains have been tested for their growth rates and biomass accumulation in BG 11 medium. The best ten biomass producing strains (Table 1, marked with an asterisk) have been assessed for their lipid accumulation induction using two methods: nitrogen starvation and salt stress. Of isolated microalgal strains Hindakia PKU AC 169, a unicellular round-oval microalga of approximately 3–4 lm in diameter, was the third highest biomass producing strain in BG-11 medium after Chlorella AC 166 and Chlorella AC 138 (Table 1). It was also the only strain capable of producing considerable lipid content in both nitrogen starvation and salt induced modifications of BG-11when analysed with fluorescent microscopy and Nile Red staining (Supplementary Fig. 1). Nile Red is a fluorescent dye that binds to both neutral yielding yellow-gold fluorescence of neutral lipids (Diaz et al., 2008) and red to yellow fluorescence of polar lipids (Diaz et al., 2008). For these reasons Hindakia PKU AC 169 was selected for further analysis. As in the previous study (Guo et al., 2013), biomass cultivation was performed in 1 L flasks aerated at 4.5 L min1 until the early stationary phase was achieved. Hindakia PKU AC 169 was cultivated in three growth media BG-11, BG-11 + NaCl and BG11N at 22 °C (Fig. 3), and was found to produce 0.402, 0.449 and 0.182 g (dry weight) of biomass/litre, respectively, after 12 days of growth, (Table 2). Very little information is available regarding performance of other Hindakia sp. strains in laboratory cultivation. A literature review suggests that a study by Zhou et al. (2011) describing cultivation of Hindakia sp.UM265 in nutrient-rich concentrated municipal wastewater is the unique example of a similar

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Fig. 1. Distribution of algae species (A) and algae families (B) among 37 strains isolated during this study and identified against nucleotide collection with BLASTN on the basis of their 18S rRNA sequences amplified with 519 forward/1406 reverse primer pair.

study. The cell size of Hindakia sp. PKU AC169 varies depending on the growth medium used (2.3–3.8 lm in BG11, 3.7–4.5 lm in BG11 + NaCl, 3.5–4.1 lm in BG11N) but generally is considerably smaller than Hindakia sp. UM265 (6–9 lm). Comparison of growth parameters of two strains of Hindakia sp. is difficult as UM265 was grown in mixotrophic conditions in nutrient rich medium; whereas PKU AC169 under strictly autotrophic conditions. Moreover, exact conditions of cultivation such as like growth temperature and shaker speed are lacking from the former study. Hindakia sp. UM265 grown in concentrated wastewater exhibited growth rate of 0.275 g L1 d1 and lipid productivity of 77.8 mg L1 d1.Hindakia sp. PKU AC169 growth parameters in three growth media are summarised in Table 2; both growth rate and lipid productivity were significantly lower under three growth conditions tested. 3.3. The effect of growth media on lipid production and FAME composition Hindakia PKU AC 169 grown in BG11 and BG11 + NaCl exhibits similar profiles with respect to the OD achieved after 12 days of cultivation, whereas the strain cultivated under nitrogen starvation conditions grows more slowly (Fig. 3). Nitrogen starvation has long been known to increase lipid content inside cells at the expense of biomass productivity (Rodolfi et al., 2009; Sheehan et al., 1998). Very often the decrease of biomass productivity is not compensated by increased oil content of the cells, resulting in lower overall lipid productivity (Rodolfi et al., 2009). This study is consistent with those findings. Cultivation of Hindakia PKU AC 169 in BG11 medium containing 150 mM NaCl results in higher biomass and lipid production and resultant higher lipid productivity than both unmodified and nitrogen starved BG11. In salt induced medium, Hindakia PKU AC 169 produced approximately one-eighth more biomass and almost twice the lipid as FAME content when compared with unmodified BG11. More importantly, the alga produced more lipids and its growth rate was more than three times greater in the salt induced medium than under nitrogen starvation conditions. Performance of Hindakia PKU AC 169 in different growth media is summarised in Table 2. The effect of salinity on algae cultivation performance has been tested in several algal strains to date and effects largely vary depending on the strain and salinity of the growth medium. Dunaliella salina is a well-known halophilic alga that is capable of survival in molar concentrations of salt approaching saturation, i.e., 5.5 M and is

routinely grown in seawater of an approximate NaCl concentration of 0.5 M. A study by Takagi et al., 2006 shows that cultivation of Dunaliella in higher content of NaCl (1 M instead of 0.5 M) resulted in increased accumulation of lipids (including triacylglycerols) from 60.6% (w/w) to 67.8% (w/w) without compromising biomass productivity. Other studies regarding the effect of salt concentration on biomass productivity and lipid productivity have been performed using freshwater algae. A recent study using the freshwater strain Scenedesmus obliquus (Kaewkannetra et al., 2012) confirms that increased lipid accumulation under salt stress that does not compromise biomass yield is achievable. The results have shown that low to medium salinities, up to 50 mM NaCl, were appropriate to promote growth rate and lipid accumulation. Similar results were obtained by Harwati et al. (2012) which found that Chlorococcum sp. grown at 1% NaCl (170 mM) in BG11 medium accumulates 70% more lipids than the same strain grown in BG11 without modification. Growth rates and biomass productivities were similar using both media. A deleterious effect on biomass productivity was observed at 2% NaCl (340 mM) in BG11 (Harwati et al., 2012). An opposite trend has been found for four freshwater strains of Botryococcus isolated in Thailand. Concentration of salt in a range of 88 mM had a deleterious effect on both growth and lipid accumulation of these strains (Yeesang and Cheirsilp, 2011), whereas a study of marine alga Nannochloropsis oculata showed that increasing salinity of the medium from 3.5% NaCl to 4.5– 5.5% NaCl increased lipid accumulation at the expense of growth rate (Gu et al., 2012). Salinity has an important effect on lipid and fatty acid metabolism and previous studies show that increased salinity can induce synthesis of membrane lipids to decrease the membrane permeability and fluidity (Chen et al., 2008; Xu and Beardall, 1997). It has been shown that the diatom Nitzschia laevis cultivated under salt stress conditions responds by increasing the content of total lipids and predominately membrane lipids without any negative effect on growth rate at 2% NaCl (340 mM) (Chen et al., 2008). Increase of lipid content of Hindakia PKU AC 169 can be most likely attributed to the same mechanism and difference in fluorescence between nitrogen starved cultures (bright yellow focused fluorescence, typical for neutral lipids) and salt-induced cultures (dispersed orange fluorescence, typical for polar lipids; Supplementary Fig. 1) are consistent with these findings. Comparison of FAME profiles of Hindakia PKU AC 169 and of Hindakia UM265 (Zhou et al., 2011) shows both similarities and differences. Lipid profiles of both strains are almost exclusively

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Fig. 2. Maximum likelihood phylogenetic tree of 18S rRNA sequences of microalgae isolated from Shenzhen offshore area. Numbers above branches indicate bootstrap values of maximum likelihood analysis from 1000 replicates. Strain Hindakia sp. PKU AC 169 used for detailed study has been marked with asterisk (⁄).

composed from fatty acids C16 and C18.The fraction of palmitic (16:0) and linoleic acids (18:2) contribute to almost 60% of Hindakia UM265 FAME profile and their content is significantly lower in Hindakia PKU AC 169 regardless of growth medium and the overall FAME profile is generally more diverse (Table 3). Analysis of FAME profiles of Hindakia PKU AC 169 grown in three different variants of BG11 shows that the growth medium has a significant effect on resultant FAMEs composition. Lipids produced in unmodified BG11 and BG11 + NaCl are predominantly composed of

unsaturated fatty acids as opposed to nitrogen starved medium that contains more saturated fatty acids. Previous studies have shown that nitrogen starvation results in accumulation of triglycerides rich in saturated fatty acids (Thompson, 1996). Triglycerides are then used as reserves to rebuild the cells once the stress is released (Roessler, 1990). Saturated fatty acids in triglycerides are preferred over unsaturated ones for two reasons. First, saturated fatty acids can be more efficiently packed as storage lipids; second, they provide more energy upon oxidation than unsaturated fatty

Table 1 Summary of morphology and basic growth parameters of thirty-seven algal strains isolated from the Shenzhen offshore area. Ten highest biomass producing strains are marked with an asterisk. Hindakia sp. PKU AC 169 was the highest biomass producing strain showing a positive result in BG-11, BG-11 + NaCl, and BG11N is additionally underlined. Genus

Family

OD

Dry weight (g/L)

Growth rate by OD (day1)

Growth rate by dry weight (g L1 day1)

Shape

Length or diameter (lm)

AC101 ⁄ AC104 AC107 AC110 AC111 ⁄ AC112 ⁄ AC114 AC115 AC116 AC118 ⁄ AC119 AC121 AC122 AC125 AC127 AC128 AC131 AC132 AC135 ⁄ AC137 ⁄ AC138 AC139 AC141 ⁄ AC143 ⁄ AC144 AC150 AC151 ⁄ AC154 ⁄ AC159 AC160 AC163 AC164 ⁄ AC166 ⁄ AC167 AC168

Micractinium Micractinium Parachlorella Chlorella Mychonastes Monoraphidium Mychonastes Chlorella Ourococcus Scenedesmus Mychonastes Scenedesmus Ourococcus Monoraphidium Micractinium Desmodesmus Mychonastes Scenedesmus Chlorella Mychonastes Chlorella Pseudodictyosphaerium Micractinium Mychonastes Chlorella Monoraphidium Mychonastes Chlorella Chlorella Desmodesmus Mychonastes Chlorella Chlorella Monoraphidium Nannochloris Hindakia

Micractiniaceae Micractiniaceae Chlorellaceae Chlorellaceae Scotiellocystoidaceae Selenastraceae Scotiellocystoidaceae Chlorellaceae Coccomyxaceae Scenedesmaceae Scotiellocystoidaceae Scenedesmaceae Coccomyxaceae Selenastraceae Micractiniaceae Scenedesmaceae Scotiellocystoidaceae Scenedesmaceae Chlorellaceae Scotiellocystoidaceae Chlorellaceae Chlorococcaceae Micractiniaceae Scotiellocystoidaceae Chlorellaceae Selenastraceae Scotiellocystoidaceae Chlorellaceae Chlorellaceae Scenedesmaceae Scotiellocystoidaceae Chlorellaceae Chlorellaceae Selenastraceae Chlorellaceae Chlorellaceae

0.25 ± 0.04 2.63 ± 0.82 2.10 ± 0.81 1.8 ± 0.44 1.37 ± 0.17 2.58 ± 0.22 2.81 ± 0.85 2.07 ± 0.61 2.37 ± 0.61 2.42 ± 0.80 2.85 ± 0.31 1.12 ± 0.07 1.90 ± 0.29 2.23 ± 0.25 1.91 ± 0.64 1.75 ± 0.41 2.16 ± 0.26 2.20 ± 0.27 0.18 ± 0.02 3.33 ± 0.15 3.84 ± 0.53 1.06 ± 0.22 0.28 ± 0.03 2.76 ± 0.56 2.53 ± 0.49 2.03 ± 0.18 2.15 ± 0.09 2.98 ± 0.39 3.31 ± 0.77 1.07 ± 0.28 2.12 ± 0.34 2.00 ± 0.36 4.56 ± 1.14 3.50 ± 0.54 2.18 ± 0.14 3.64 ± 0.28

0.034 ± 0.005 0.356 ± 0.111 0.284 ± 0.109 0.244 ± 0.060 0.186 ± 0.023 0.349 ± 0.030 0.381 ± 0.115 0.280 ± 0.082 0.320 ± 0.082 0.327 ± 0.108 0.385 ± 0.042 0.151 ± 0.009 0.257 ± 0.040 0.302 ± 0.033 0.259 ± 0.086 0.237 ± 0.055 0.292 ± 0.035 0.297 ± 0.037 0.025 ± 0.002 0.451 ± 0.020 0.520 ± 0.071 0.143 ± 0.030 0.038 ± 0.00 0.373 ± 0.075 0.343 ± 0.066 0.274 ± 0.024 0.290 ± 0.013 0.404 ± 0.053 0.448 ± 0.104 0.145 ± 0.039 0.288 ± 0.046 0.270 ± 0.049 0.617 ± 0.154 0.474 ± 0.073 0.295 ± 0.020 0.493 ± 0.038

0.015 0.197 0.157 0.128 0.101 0.193 0.210 0.154 0.178 0.180 0.214 0.079 0.141 0.164 0.140 0.129 0.159 0.161 0.009 0.248 0.288 0.077 0.017 0.204 0.188 0.151 0.160 0.222 0.248 0.078 0.159 0.148 0.344 0.262 0.161 0.272

0.002 0.027 0.021 0.017 0.014 0.026 0.028 0.021 0.024 0.024 0.029 0.011 0.019 0.022 0.019 0.018 0.022 0.022 0.001 0.034 0.039 0.010 0.002 0.028 0.025 0.020 0.022 0.030 0.033 0.011 0.022 0.020 0.047 0.035 0.022 0.037

Round Round Round Round Ellipse Ellipse Round Round Selenodont Round Round Spindle Selenodont Selenodont Round Ellipse Round Ellipse Round Round Round Round Round Round Round Spindle Round Round Round Round Round Round Round Ellipse Round Round-oval

6.25 5.54–5.93 4.15–4.54 3.35–3.47 7.57–10.75 3.46–5.1 3.04–3.48 3.38 4.32–5.80 3.55–3.79 2.81–3.36 11.08–11.25 5.37–5.91 5–5.16 2.47–2.49 3.34–4.44 2.47–3.25 6.46–7.93 3.82–4.63 2.61–3.18 2.90–4.90 1.44–1.93 3.02–3.72 2.17–3.28 2.98–3.96 13.57–27.51 2.16–2.17 3.70–3.99 2.83–3.57 3.95–4.71 2.48–2.68 2.81–3.38 2.67–2.99 3.44–3.57 2.76–2.77 2.63–3.78

Parietochloris

Trebouxiaceae

2.51 ± 0.26

0.340 ± 0.035

0.185

0.025

Fan-shaped

5.61–6.39

⁄ ⁄

AC169 AC171

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Fig. 3. Growth curves of Hindakia PKU AC 169 cultivated in three growth media unmodified BG-11 (rhombes), salt induced medium BG-11+ NaCl (squares) and nitrogen depleted medium BG11-N (triangles) at 22 °C for 12 days. Error bars represent standard deviation of three independent biological replicas. Table 2 Summary of basic growth parameters of Hindakia PKU AC 169 cultivated in three growth media BG-11, BG-11 + NaCl and BG11N at 22 °C for 12 days. Hindakia PKU AC 169

Biomass concentration [gDW L1]

Growth rate [gDW L1 day1]

FAME% [gFAME/gDWalgae]

FAME yield [gFAME L1]

FAME productivity [gFAME L1 day1]

BG-11 BG-11 + NaCl BG11N

0.402 ± 0.116 0.449 ± 0.023 0.182 ± 0.024

0.032 ± 0.001 0.034 ± 0.001 0.010 ± 0.001

38.09% ± 1.23% 68.39% ± 6.89% 52.89% ± 4.66%

0.153 ± 0.045 0.307 ± 0.017 0.096 ± 0.013

0.013 ± 0.004 0.026 ± 0.001 0.008 ± 0.001

Table 3 Summary of cellular fatty acid composition as methyl esters (FAMEs) of Hindakia PKU AC 169 cultivated in three growth media BG-11, BG-11 + NaCl and BG11N at 22 °C for 12 days. ND – none detected. Fatty acid composition

C14:0 C16:0 C16:2 C16:3 C17:0 C18:0 C18:1 C18:2 C18:3 R Saturated (Sat) R Unaturated (Un) R Un/Sat

Growth medium BG11

BG11 + NaCl

BG11N

16.29% ± 0.48% 17.17% ± 2.36% 4.57% ± 2.04% 16.78% ± 2.41% ND 10.51% ± 0.28% 3.33% ± 0.24% 11.28% ± 0.77% 20.08% ± 0.90% 43.97% ± 2.20% 56.03% ± 2.20% 1.27

16.78% ± 0.60% 16.91% ± 0.14% 10.06% ± 0.24% 11.35% ± 0.09% ND 10.98% ± 0.40% 5.73% ± 0.12% 17.73% ± 0.34% 10.46% ± 0.14 44.67% ± 0.86% 55.33% ± 0.86% 1.24

17.91% ± 0.58% 21.07% ± 0.03% 1.90% ± 0.09% 10.37% ± 0.29% 3.99% ± 0.54% 12.13% ± 0.06% 7.22% ± 0.15% 11.24% ± 0.09% 14.48% ± 0.29% 55.10% ± 0.02% 44.90% ± 0.02% 0.81

acids (Roessler, 1990). A recent study on Nannochloropsis oceanica IMET1 has shown a positive relation between nitrogen content in the growth medium and the content of polar unsaturated lipids. It has also confirmed that decreasing the nitrogen content in the growth medium results in an accumulation of neutral lipids that are mainly composed of saturated fatty acids (Xiao et al., 2013). They suggest that in a course of nitrogen starvation lipid classes have changed. Polar membrane lipids rich in unsaturated fatty acids were partially consumed, and neutral triglycerides rich in

saturated fatty acids were synthesised (Xiao et al., 2013). A similar mechanism has likely occurred during nitrogen starvation of Hindakia PKU AC 169 and explains the increased content of saturated fatty acids under nitrogen starvation conditions. The degree of fatty acid unsaturation also differs between BG11 and BG11 + NaCl. The former contains a considerable amount of hexadecatrienoic acid (16:3) and a-linolenic acid (18:3), whereas the latter contains more hexadecadienoic acid (16:2) and linoleic acids (18:2). Nitrogen starved cells are rich in saturates: myristic (14:0), palimtic (16:0) and stearic (18:0) acids. These findings are summarised in Table 3. The decrease of fatty acid chain unsaturation in a response to salt stress is also consistent with previous findings (Xu and Beardall, 1997). Decreasing the unsaturation will most likely result in reduced membrane permeability to limit the diffusion of potentially harmful ions into the cell (Xu and Beardall, 1997). Co-existence of high biomass and lipid productivity exhibited by Hindakia PKU AC 169 grown in BG11 medium supplemented with NaCl makes it a promising feedstock for biofuel production, providing that conversion of these lipids to FAMEs is performed with a method that is not sensitive to a high content of membrane lipids. Many methods of biodiesel production such as alkali (Freedman et al., 1984) and lipase (Watanabe et al., 2002) have been found sensitive to high content of polar lipids, mainly phospholipids. Acid catalysed transesterification was used to successfully convert phospholipids to methyl esters without the need to additionally pre-treat the lipids (Lu et al., 2009), and this study is consistent with those findings.

M. Daroch et al. / Bioresource Technology 146 (2013) 192–199

Analysis of the FAME profile of Hindakia PKU AC 169 under reported growth conditions suggests that the strain may be suitable for production of biodiesel by acid catalysed transesterification of algal lipids. Hindakia PKU AC 169 grown in BG11 + NaCl has a favourable chain length of its fatty acids composed exclusively of biodiesel-suitable C14 to C18 fatty acids. Moreover content of highly unsaturated fatty acids such as hexadecatrienoic acid (16:3) and a-linolenic acid (18:3) is limited, compared to the same strain in unmodified BG11 medium and is below the limit of 12% set by EN14214 biodiesel standard. On the other hand, the saturated fatty acids content is lower than the same strain grown in nitrogen starvation conditions which can have a positive effect on utilisation of Hindakia PKU AC 169-derived biodiesel in colder climates. 4. Conclusions The microalgal strain Hindakia PKU AC 169 isolated from coastal waters of the Pearl River Delta region was successfully cultivated in two derivatives of BG11 medium: nitrogen starved and salt induced. Both methods yielded improved lipid accumulation, but only salt induction resulted in increased overall lipid productivity. Derivatisation of algal lipids to FAMEs showed different lipid profiles under selected growth conditions and suggest that salt induced medium is better suited for biodiesel production than nitrogen starved medium due to the higher productivity and more favourable composition of fatty acids. Acknowledgements This project was funded by a Shenzhen Development and Reform Commission Grant [2011] 835. Authors would like to acknowledge Shanfa Zhang for morphological description of algal strains and Zongchao Jia for his lab assistance. References Ben-Amotz, A., Tornabene, T.G., Thomas, W.H., 1985. Chemical profile of selected species of microalgae with emphasis on lipids. J. Phycol. 21 (1), 72–81. Borowitzka, M.A., 1999. Commercial production of microalgae: ponds, tanks, tubes and fermenters. J. Biotechnol. 70 (1–3), 313–321. Chen, G.Q., Jiang, Y., Chen, F., 2008. Salt-induced alterations in lipid composition of diatom Nitzschia laevis (bacillariophyceae) under heterotrophic culture condition. J. Phycol. 44 (5), 1309–1314. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25 (3), 294–306. 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., 2013. Recent advances in liquid biofuel production from algal feedstocks. Appl. Energy 102, 1371–1381. Diaz, G., Melis, M., Batetta, B., Angius, F., Falchi, A.M., 2008. Hydrophobic characterization of intracellular lipids in situ by Nile Red red/yellow emission ratio. Micron 39 (7), 819–824. Fitzherbert, E.B., Struebig, M.J., Morel, A., Danielsen, F., Brühl, C.A., Donald, P.F., Phalan, B., 2008. How will oil palm expansion affect biodiversity? Trends Ecol. Evol. 23 (10), 538–545. Freedman, B., Pryde, E.H., Mounts, T.L., 1984. Variables affecting the yields of fatty esters from transesterified vegetable oils. J. Am. Oil Chem. Soc. 61 (10), 1638– 1643.

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