Bioresource Technology 161 (2014) 297–303
Contents lists available at ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Enhanced lipid production in Chlorella pyrenoidosa by continuous culture Xiaobin Wen a,b, Yahong Geng a, Yeguang Li a,⇑ a b
Key Laboratory of Plant Germplasm Enhancement and Speciality Agriculture, Wuhan Botanical Garden, Chinese Academy of Sciences, Wuhan 430074, China University of Chinese Academy of Sciences, Beijing 100049, China
h i g h l i g h t s Chlorella pyrenoidosa XQ-20044 is able to accumulate lipids in growing cells. One step production of algal lipid was achieved in chemostat culture. Proper SNI was the key for simultaneous algal growth and lipid accumulation. Lipid productivity was significantly enhanced by continuous culture.
a r t i c l e
i n f o
Article history: Received 24 January 2014 Received in revised form 12 March 2014 Accepted 16 March 2014 Available online 25 March 2014 Keywords: Chlorella Growth Lipid productivity Continuous culture Chemostat
a b s t r a c t Usually microalgae growth and lipid accumulation do not run in parallel throughout cultivation, which necessarily lowers overall lipid productivity. However, we show through batch and feed-batch studies of Chlorella pyrenoidosa XQ-20044 that by varying the nitrate concentration, conditions which produce fairly high lipid content could be achieved without sacrificing algal growth. Simultaneous microalgae growth and lipid production was achieved in continuous chemostat culture when the specific nitrate input rate was in the range of 0.78–4.56 mmol g 1 d 1. Moreover, the maximum lipid productivity (144.93 mg L 1 d 1) in the continuous culture was significantly higher than in batch culture (96.28 mg L 1 d 1), thus indicating the feasibility and great advantage of one-step production of microalgal lipids. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Aquatic microalgae have a high capacity for photosynthesis, and many are able to use excess photosynthetically-fixed carbon for synthesis of neutral lipids, which are also known as triacylglycerols (TAGs) (Chisti, 2007). Oleaginous microalgae are thus considered to be ideal raw materials for biodiesel production, especially if their growth is coupled to the direct bio-fixation of waste CO2 (Kwak et al., 2006). Over the past 50 years the concept and feasibility of microalgal biodiesel have been discussed extensively (Wijffels and Barbosa, 2010), but this renewable energy source has yet to be exploited. Limiting factors include the lack of appropriate microalgal strains, less-than optimal lipid productivity and ineffective culturing techniques for lipid accumulation.
⇑ Corresponding author. Address: Wuhan Botanical Garden, Chinese Academy of Sciences, Wuhan 430074, Hubei Province, China. Tel.: +86 27 87510542; fax: +86 27 87510251. E-mail address: [email protected] (Y. Li). http://dx.doi.org/10.1016/j.biortech.2014.03.077 0960-8524/Ó 2014 Elsevier Ltd. All rights reserved.
To date, several hundred oleaginous species have been isolated and characterized (Gouveia et al., 2009). A common thread in these studies is that cell growth and lipid accumulation do not happen at the same time during cultivation (Lourenco et al., 2002; Merzlyak et al., 2007), which results in lower overall lipid productivity. In order to overcome this, researchers have explored two-stage cultivation strategies to enhance microalgal lipid production. In such strategies, which have been used mainly with batch cultures, the microalgal cells first grow rapidly under growth-optimized conditions, and then are transferred to conditions where light irradiance (Zhang et al., 2009), nutrition (Su et al., 2011), culture pH (Han et al., 2013), as well as other factors (Das et al., 2011; Liu et al., 2008) are adjusted to promote lipid accumulation at the expense of cell growth. However, detailed study of the stress response of oleaginous microalgae under nitrogen deficiency may provide a foundation for the production of biomass and lipids in one step. Recent data showed that while Neochloris oleoabundans UTEX #1185 accumulated lipid under relatively high nitrogen stress conditions (i.e., low nitrogen levels), its growth was not severely limited (Adams
298
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
et al., 2013). In contrast, growth of Chlorella vulgaris UTEX #265 was severely limited under high nitrogen stress, but lipid accumulation was triggered ahead of the growth limitation (Adams et al., 2013). These may or may not be species-specific characteristics, but the results have inspired the authors to explore the threshold values of nitrogen concentration and/or supply rate, in order to establish a continuous cultivation system for concurrent production of biomass and lipids. It is hypothesized that by properly regulating the nitrate concentration (e.g., severe stress or moderate stress) in continuous culture, the algal cells could grow at a reasonable rate and accumulate lipid simultaneously, thus leading to higher lipid productivity over batch culture. Along this line, one-step astaxanthin production was achieved with continuous culture of Haematococcus pluvialis, where >0.8% (DW) astaxanthin accumulated in the green vegetative cells (Del Rio et al., 2005). However, only a few studies have been published on the continuous cultivation of microalgae for lipid production. Sobczuk and Chisti (2010) investigated the effects of dilution rate on lipid productivity of the freshwater microalga Choricystis minor in chemostat culture, and concluded that the lipid content did not change significantly with various dilution rates. Very similar results were reported for chemostat cultures of Chlorella minutissima and Dunaliella tertiolecta by Tang et al. (2012). In contrast, Klok et al. (2013) reported that excess light combined with a growth-limiting nitrogen supply resulted in TAG accumulation (up to 12.4%, w/w) and cell replication in turbidostat cultures of Neochloris oleoabundans. Given the quite different results of Klok et al., it is an open question how other microalgal species will respond in continuous culture. In this report, batch and feed-batch cultures of C. pyrenoidosa were used to study its response, in terms of growth and lipid accumulation, to different nitrogen concentrations. The results were used to establish chemostat cultures with different dilution rates, in order to verify the assumption that microalgal cultures can be regulated to grow and accumulate lipids concurrently, thus enhancing lipid productivity. 2. Methods 2.1. Strains and pre-culture conditions C. pyrenoidosa XQ-20044 was used in this study. It was provided by the Algae Culture Collection of Wuhan Botanical Garden, Chinese Academy of Sciences. This fast growing strain was originally isolated from a Spirulina culture pond in Sichuan province, China. Its lipid content in a column photobioreactor was >45% of dry biomass (unpublished results), thus showing its potential as a biodiesel feedstock. For seed cultures, the algal cells were grown photoautotrophically in Erlenmeyer flasks at 25 °C. The flasks contained 600 mL of medium, and were placed on a shaker (100 rpm) for 48 h of cultivation before further experiments. Continuous illumination was provided by fluorescent lamps, and the light intensity on the surface of the flask was adjusted to 50 lmol m 2 s 1. The basal growth medium for seed culture and culture experiments was a modified BG11 medium which had the following composition (per liter): NaNO3, 100 mg; K2HPO43H2O, 40 mg; MgSO47H2O, 75 mg; CaCl22H2O, 36 mg; Citric acid, 6 mg; Fe-Ammonium citrate, 6 mg; EDTANa2, 1 mg; Na2CO3, 20 mg; H3BO3, 2.86 lg; MnCl24H2O 1.8 lg; ZnSO47H2O, 0.22 lg; CuSO45H2O, 0.08 lg; Na2MoO42H2O, 0.391 lg; Co(NO3)26H2O, 0.0494 lg (Grobbelaar, 2007). Sodium nitrate concentration was modified as indicated in the text. 2.2. Batch cultures Batch cultures of C. pyrenoidosa XQ-20044 were grown in aerated-column photobioreactors. The seed cultures were
centrifuged, rinsed, and resuspended in nitrate-free BG-11 medium to an optical density of 0.5 ± 0.05 at 540 nm. Then aliquots of this suspension were transferred to the glass columns (inner diameter 3 cm), which were submerged in a water tank. A thermostatic water circulator was used to provide 30 °C water bath for the culture columns. CO2-enriched (1%, v/v) air was passed through a sterile filter and then bubbled into the bottom of each column at a flow rate of 250 ml min 1. Light (300 lmol m 2 s 1 irradiance impinging on the surface of each column) was continuously provided by ten Phillips 36 W cool white fluorescent lamps. A concentrated sodium nitrate stock solution (autoclaved and stored at 4 °C) was used to adjust the columns to initial nitrate concentrations that ranged from 0.24 mM to 17.65 mM. And for each nitrate concentration, three replicate cultures were conducted in parallel. After 8 d of cultivation, cells were collected by centrifugation (5000 rpm at 15 °C for 5 min) and lyophilized ( 56 °C cryotrapping, 10–14 Pa vacuum) for further analysis. 2.3. Feed-batch cultures Feed-batch culturing of C. pyrenoidosa XQ-20044 was carried out similar to the batch cultures, except nitrate (in concentrations of 0.24, 0.48, 0.72, 0.96, 1.18, 2.35 and 3.53 mM) was added to the photobioreactors at 0, 24, 48, 72, and 96 h of cultivation. In the batch cultures with an initial nitrate concentration >3.53 mM, the residual nitrate level after 24 h was >50% of the initial value. Thus, only nitrate concentrations ranging from 0.24 mM to 3.53 mM were investigated in these feed-batch cultures. 2.4. Chemostat cultures A peristaltic pump with a ten-roller pump head was used for the continuous cultures. It delivered a maximum flow rate of 32 mL min 1 and a minimum flow rate of 0.1 mL min 1. Exactly 2000 mL of algae were grown in a closed column bioreactor with an inner diameter of 10 cm. A silicone tube was connected to the bottom of the column to form a U-shaped overflow tube. Air enriched with CO2 (1%, v/v) was passed through a sterile Millex syringe filter (0.22 lm) and then bubbled into the bottom of the culture at a flow rate of 2 L min 1; this continuously agitated the cells, so they received equivalent light. The culture pH was maintained in the range of 7–8, and continuous illumination (600 lmol m 2 s 1 photon flux) was provided by four fluorescent lamps (Philips MASTER PL-L 36 W) on one side of the bioreactor. The whole cultivation unit was placed in a thermostatic room to maintain the culture temperature at 30 °C. Cultivations (in triplicate) were carried out initially in batch mode by inoculating the seed culture into BG-11 medium to an optical density of 0.1 (540 nm). After 30 h of cultivation, when the optical density of the algal suspension had increased to 1.0 or more, the bioreactor was switched to continuous mode by feeding BG-11 medium (0.71 mM sodium nitrate). Cultivations with different dilution rates (ranged from 0.24 d 1 to 2.4 d 1) were carried out successively. Achievement of steady-state condition at each dilution rate was monitored by daily measuring of the optical density, biomass dry weight, and residual nitrate concentration. The steady state was maintained at least for three days before further determinations. 2.5. Analytical methods Biomass dry weight was measured to evaluate microalgal growth. About 10 mL algal suspension was filtered through a predried GF/C glass microfiber filter paper (0.45 lm), which was dried at 80 °C under vacuum for 4 h (Lee et al., 1996) and re-weighed to
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
calculate biomass dry weight (DW). A spectrophotometric method, described by Collos et al. (1999), was used to monitor residual nitrate concentration. Briefly, the absorbance of culture filtrate (0.22 lm filter) at 220 nm and a pre-constructed standard curve were used to determine residual nitrate. Pigments were extracted from live cells with hot DMSO (70 °C) and quantified spectrophotometrically using coefficients determined by Merzlyak et al. (2007). The photosynthetic status of the cells was evaluated by measuring the net photosynthetic oxygen evolution rate (Zhang et al., 2013) using a liquid phase oxygen measurement system with white light. For biochemical composition analysis, the microalgal cells were harvested and vacuum lyophilized. For lipid quantification, 50 mg of dry algal biomass was fully ground, transferred to a covered centrifuge tube and extracted with a mixture of n-hexane and ethyl acetate (1:1, v:v). The extraction was repeated 3 times and all extracts were combined into a pre-weighed glass tube, and then dried with a stream of nitrogen (Wen et al., 2012). The lipids were determined gravimetrically. Neutral lipid, glycolipids, and phospholipids were fractionated from the lipid extracts (100 mg) by column chromatography using a 2 cm 20 cm column packed with 4 g silica gel 60 (Wang and Wang, 2012). The lipids were differentially eluted, first with 20 mL of chloroform to obtain the neutral lipids, and then with 15 mL of acetone:methanol (9:1, v:v) to obtain the glycolipids; phospholipids were eluted last with 15 mL of methanol. The fractions were confirmed by TLC separation, and then quantified gravimetrically. In order to analyze the fatty acids, about 20 mg of the lipid extract was dissolved in 3 mL of n-hexane, and then transmethylated by adding 3 mL of methanol–KOH (0.5% KOH) and heating at 50 °C for 60 min. After cooling to room temperature, the hexane layer was separated and dried with anhydrous sodium sulfate. Fatty acid methyl esters were analyzed by gas chromatography (Agilent 7890A) using an HP-5 Phenyl Methyl Siloxan column (30 m 0.32 mm 0.25 lm) and a flame ionization detector. 1 lL samples were injected with a splitting ratio of 5:1 at 250 °C. The heating program was 150 °C for 2 min, followed by an increase in the temperature of 10 °C/min to 250 °C, where it was held for 8 min. The detector temperature was set at 240 °C. A Supelco 37Component FAME Mix (47885-U) was used as external standard for fatty acid identification. All of the above analytical experiments were done in triplicate, and the results were analyzed for variance (Zar, 1999) with SAS 8.01 at a significance level a = 0.05. Tukey’s multiple comparison tests were done where applicable.
299
3. Results and discussion 3.1. Non-parallel occurrence of cell growth and lipid accumulation in batch culture Nitrogen starvation has frequently been reported to induce lipid accumulation in microalgal cells (Hu et al., 2008). To clarify the growth and lipid responses of C. pyrenoidosa XQ-20044 to varying nitrate levels, a series of batch cultures were carried out at different initial nitrate concentrations (Fig. 1). The results showed that biomass dry weight (DW) of C. pyrenoidosa XQ-20044 increased almost linearly with increasing initial nitrate from 0.24 mM to 2.35 mM. Further increases in the nitrate concentration did not affect biomass accumulation, possibly due to light limitation (Fig. 1A). The lipid content (% DW), however, exhibited different trends in response to increasing nitrate concentration (Fig. 1A). Specifically, the lipid content first increased from 38.79% to 52.36% (DW), and then declined to 39.59% (DW), as the initial nitrate concentration was increased from 0.24 mM to 5.88 mM. Further increasing the initial nitrate concentration (above 5.88 mM) had no significant influence on lipid content. Changes of the nitrate concentration during culture are also indicated in Fig. 1B. These results showed that cell growth and lipid accumulation did not always change in parallel, and moreover, they suggested relatively high lipid content could be achieved without the expense of algal growth. As shown in Fig. 1A, decreasing the nitrate concentration from 5.88 mM to 2.35 mM significantly increased the lipid content (p < 0.05), while the biomass was not affected (1.8 g L 1, p > 0.05). 3.2. Effects of intermittent nitrate feeding on cell growth and lipid accumulation To confirm that there is a culture status under which C. pyrenoidosa XQ-20044 moderately accumulates lipids in growing cells, feed-batch culture was conducted with initial nitrate concentrations of 0.24, 0.47, 0.71, 0.94, 1.18, 2.35 and 3.53 mM, and the same amount of nitrate (as the initial concentration) was fed respectively into each culture every 24 h. A similar response of biomass dry weight and lipid content to varying nitrate was observed (Fig. 2A). In particular, the cultures with initial nitrate concentration of 0.71, 0.94 and 1.18 mM reached maximal biomass density of about 2.2 g L 1 when nitrate was fed intermittently, and all of them showed fairly high lipid content (35–45% DW). The nitrate concentration of the 3 cultures fluctuated only within a small range during the feed-batch culture
Fig. 1. (A) Effects of initial nitrate concentration on biomass dry weight (j) and lipid content (N) in batch culture of C. pyrenoidosa XQ-20044. (B) Changes of nitrate concentration during batch culture of C. pyrenoidosa XQ-20044. More than 2 mM nitrate was detected at the end of the culture with initial nitrate concentrations higher than 6 mM (data not shown).
300
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Fig. 2. (A) Effects of intermittent nitrate feeding on biomass dry weight (j) and lipid content (N) of C. pyrenoidosa XQ-20044. (B) Changes of nitrate concentration during batch culture of C. pyrenoidosa XQ-20044.
Fig. 3. Effects of dilution rate on steady-state biomass concentration (j), lipid content (N) and nitrate input rate (s) in chemostat culture of C. pyrenoidosa XQ20044. Experiments were carried out with constant feeding medium composition (BG-11 with 0.71 mM nitrate) and illumination (600 lmol m 2 s 1), but with varying dilution rate between 0.24 and 2.4 d 1.
(Fig. 2B). These results suggested that microalgal growth and lipid accumulation could both be optimized by using continual feeding of small amounts of nitrate. The nitrate concentration and biomass density of culture changes over time in both batch and feed-batch mode, thus the optimum status for concurrent microalgae growth and lipid accumulation cannot be sustained using these two modes. However, continuous cultivation of C. pyrenoidosa XQ-20044 in chemostat mode could potentially overcome this problem, and thus was investigated. 3.3. Concurrent microalgae growth and lipid accumulation in continuous culture Steady-state studies of C. pyrenoidosa XQ-20044 were carried out using a chemostat system with constant medium composition (BG-11 with 0.71 mM nitrate), illumination and air flow rate, but with dilution rates that varied between 0.24 and 2.4 d 1. Steadystates were achieved with all the tested dilution rates. The cultures with high dilution rate showed a green color while the cultures with low dilution rate had a yellow-green appearance. 3.3.1. Microalgal growth As shown in Fig. 3, the nitrate input rate, which varied from 0.17 to 1.69 mol L 1 d 1, correlated linearly with dilution rate. Also, since the nitrate level in all steady-state cultures was below detection limit (0.01 mM), the nitrate uptake rate was equal to the
Fig. 4. Effects of dilution rate on pigment content (Chla h, Total Car e) and net photosynthetic oxygen evolution rate (N) in chemostat culture of C. pyrenoidosa XQ20044.
nitrate input rate. Dilution rate also had a remarkable impact on the biomass concentration at each steady-state (Fig. 3). With increasing dilution rates from 0.25 d 1 to 2.4 d 1, the biomass dry weight was decreased 86%, from 0.57 g L 1 to 0.08 g L 1. The pigment content and net photosynthetic oxygen evolution rate of C. pyrenoidosa XQ-20044 as a function of dilution rate at each steady-state are illustrated in Fig. 4. The lowest contents of Chlorophyll a (1.33 mg g 1) and total carotenoids (1.00 mg g 1) in the cells were observed at the lowest dilution rate of 0.25 d 1. However, with increasing dilution rate, both Chlorophyll a and total carotenoids increased, and reached their maximum values (24.16 mg g 1 and 6.24 mg g 1, respectively), at the steady-state with the dilution rate of 2.4 d 1. The net photosynthetic oxygen evolution rate, which is a substantial growth indicator for autotrophic organisms, was calculated from the measured changes in dissolved oxygen per unit time and the Chlorophyll a content of the culture. As shown in Fig. 4, the rapidly growing cells (with high dilution rate) presented high photosynthetic oxygen evolution rate. However, when the dilution rates (growth rates) were decreased from 1.44 d 1 to 0.48 d 1, the net photosynthetic oxygen evolution rate was only decreased by 13%. Statistic analysis showed that the cells maintained a relative stable photosynthetic activity when the dilution rates were 0.48, 0.72, 0.96, and 1.44 d 1. Out of this range, the photosynthetic oxygen evolution changed significantly (p > 0.05) when changing the dilution rates. Photosynthetic activity of nitrate-stressed microalgae is essential for carbon storage metabolism, especially for triglyceride accumulation (Dillschneider et al., 2013; Pan et al., 2011). In the chemostat
301
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Table 3 Comparison of fatty acid profiles of C. pyrenoidosa XQ-20044 in chemostat and batch culture. Fatty acid
Fig. 5. Biomass (j) and lipid (N) productivity as affected by the dilution rate in chemostat culture of C. pyrenoidosa XQ-20044.
C14:0 C15:0 C16:0 C16:1 C17:0 C17:1 C18:0 C18:1n9c C18:2n6t C18:3n3 C20:0 C20:1n9 C22:0 C22:1n9
% of Total fatty acids Batch culture
Chemostat
0.31 ± 0.06 0.21 ± 0.03 25.53 ± 0.12 9.4 ± 0.16 0.67 ± 0.05 0.4 ± 0.03 7.39 ± 0.56 27.45 ± 0.40 12.69 ± 0.34 11.22 ± 0.49 2.05 ± 0.09 2.21 ± 0.42 ND 0.45 ± 0.09
0.38 ± 0.05 0.18 ± 0.02 22.91 ± 0.13 9.72 ± 0.23 0.53 ± 0.05 0.42 ± 0.03 6.1 ± 0.19 29.2 ± 0.54 13.42 ± 0.05 12.57 ± 0.36 2.19 ± 0.09 2.01 ± 0.42 0.33 ± 0.12 ND
ND: not detected. Table 1 Lipid content and composition (means + SD) of C. pyrenoidosa XQ-20044 at steadystate with various dilution rates. Dilution rate
Lipid content (% DW)
Lipid class distribution (% total lipids) Neutral lipids
Glycolipids
Phospholipids
0.24 0.48 0.72 0.96 1.44 1.92 2.40
30.04 ± 1.35 34.69 ± 0.68 25.51 ± 1.06 23.32 ± 1.31 19.97 ± 0.13 14.92 ± 1.18 14.60 ± 1.28
76.19 ± 2.15 81.62 ± 1.47 78.06 ± 1.45 69.65 ± 0.77 48.51 ± 2.31 37.04 ± 0.87 36.96 ± 1.55
15.06 ± 0.97 11.55 ± 0.12 16.43 ± 0.83 22.15 ± 1.01 43.05 ± 1.05 52.76 ± 0.09 52.17 ± 1.02
4.71 ± 0.52 4.06 ± 0.77 4.56 ± 0.04 5.45 ± 0.13 5.42 ± 0.13 8.15 ± 0.43 7.85 ± 0.42
Fig. 6. Nitrate uptake (line) and specific TAG accumulation rate (column) as affected by specific nitrate input rate in chemostat culture of C. pyrenoidosa XQ20044.
culture of this study, the photosynthetic activity sustained, rather than declined, when the dilution rate (growth rate) was decreased from 1.44 d 1 to 0.48 d 1 (Fig. 4). It will be shown below that lipid
productivity at these dilution rates (from 0.48 to 1.44 d 1) was significantly higher (p < 0.05) than at other dilution rates (Fig. 5). The sustained photosynthetic activity of C. pyrenoidosa XQ-20044 might contribute to triglyceride accumulation. 3.3.2. Lipid accumulation The dilution rate had differing impacts on lipid content in different steady-state conditions (Fig. 3). The highest lipid content (34.69% DW) was observed with the dilution rate of 0.48 d 1. Increasing the dilution rate (i.e., from 0.48 to 1.92 d 1) led to a decline in the lipid content. When the dilution rate was further increased from 1.92 d 1 to 2.4 d 1, the lipid content (14.92% DW) did not change due to sufficient nitrate supply. A similar phenomenon was reported before (Sobczuk and Chisti, 2010; Tang et al., 2012); in those studies, the chemostat culture had relatively high nitrate concentration (feeding medium) and no significant changes in lipid content were observed under different dilution rates. The results from this study also suggested that severe nitrogen limitation (the lowest dilution rate) reduced lipid content as well as growth (Fig. 3), which was different from previous studies reported that lipid accumulation varies inversely in response to nitrate concentration (Lourenco et al., 2002; Merzlyak et al., 2007). Fractionation of the lipids by column chromatography (Table 1) revealed that neutral lipids (mainly TAG) were the main component with dilution rate less than or equal to 0.96 d 1. Glycolipids had higher percentage (about 50%) of the lipids when the dilution rate was increased to 1.92 d 1 or more. Apparently cellular TAG accumulation was triggered when steady-state was attained at dilution rate of 0.24, 0.48, 0.72, 0.96, and 1.44 d 1 respectively. 3.3.3. Productivity of biomass and lipid The biomass productivity of chemostat culture increased dramatically in response to increasing dilution rates (from 0.24 d 1 to 1.44 d 1), reaching a peak of 641.52 mg L 1 d 1 at the dilution
Table 2 Comparison of growth and lipid accumulation in chemostat and batch culture of C. pyrenoidosa XQ-20044. Culture mode
Dilution rate (d 1)
Nitrate consumed (mM/ 8 d)
Biomass productivity (mg L 1 d 1)
Lipid content (% DW)
Lipid productivity (mg L 1 d 1)
C16 + C18 (% total FFA)
Chemostat Chemostat Chemostat Chemostat Batch
0.48 0.72 0.96 1.44 /
2.71 4.07 5.42 8.13 2.71
417.81 ± 26.88 482.40 ± 13.15 539.87 ± 30.73 641.52 ± 24.37 215.81 ± 8.08
34.69 ± 0.68 25.51 ± 1.06 23.32 ± 1.31 19.97 ± 063 44.07 ± 6.04
144.93 ± 5.16 124.25 ± 5.65 125.89 ± 8.50 128.12 ± 4.28 96.28 ± 4.55
93.68 ± 1.15 95.22 ± 0.97 93.47 ± 2.04 92.12 ± 1.42 93.92 ± 1.17
302
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Table 4 Biomass and lipid productivity reported in literatures dealing with microalgal lipid production in chemostat culture. Species
Dilution rate (d 1)
Nitrate in feeding medium (mM)
Biomass productivity (mg L 1 d 1)
Lipid productivity (mg L 1 d 1)
References
Chlorella pyrenoidosa XQ-20044 Chlorella minutissima (UTEX 2219) Dunaliella tertiolecta (UTEX LB 999) Choricystis minor B. Fott
0.48 0.328 0.42 0.336
0.71 8.82 2.18 17.6
417 ± 26 137 91 351
144 ± 5 6 10 82
This study Tang et al. (2012) Tang et al. (2012) Sobczuk and Chisti (2010)
rate of 1.44 d 1 (Fig. 5). However, the biomass productivity started to decrease when the dilution rates were further increased from 1.44 to 2.40 d 1. A similar pattern was also observed in lipid productivity in response to dilution rate, except that the maximum lipid productivity (144.93 mg L 1 d 1) was achieved with a dilution rate of 0.48 d 1 (Fig. 5). ANOVA analysis concluded that the lipid productivity at dilution rates of 0.48, 0.72, 0.96 and 1.44 d 1 were significantly higher (p < 0.05) than that achieved with other dilution rates. Few studies has been conducted in C. pyrenoidosa previously but maximal biomass productivity attained in C. minutissima was 137 mg L 1 d 1 with 6 mg L 1 d 1 FAME productivity (Tang et al., 2012). Sobczuk and Chisti (2010) reported the maximal biomass productivity of 351 mg L 1 d 1 with 82 mg L 1 d 1 lipid productivity when Choricystis minor chemostat culture was performed with a dilution rate of 0.336 d 1. The biomass productivity and lipid productivity in this study were both higher than previously reported (Table 4). So, what is the favorable condition for concurrent growth and lipid accumulation for C. pyrenoidosa XQ-20044? When the dilution rate was in the range of 0.48–1.44 d 1, all of the supplied nitrate was assimilated by cells and the chemostat cultures showed moderate cell growth (Fig. 3) and photosynthetic activity (Fig. 4). Importantly, TAG accumulation also took place in the cells under the above-mentioned conditions (Fig. 3 and Table 1). Therefore, the lipid productivity was significantly improved, as indicated in Fig. 5. The specific nitrate input rate, or SNI (mmol g 1 d 1), was one of the key factors proposed by Del Rio et al. (2005) for simultaneous cell growth and astaxanthin accumulation in Haematococcus. As illustrated in Fig. 6, only when the SNI was in the range of 0.78–4.56 mmol g 1 d 1 (with a corresponding nitrate uptake of 0.81–1.59 mmol per gram of biomass) could a high rate of specific TAG accumulation (TAG increase per g biomass and unit time) be attained in the chemostat cultures of C. pyrenoidosa XQ-20044. Meanwhile the net photosynthetic oxygen evolution rate and cell growth were limit to some extent (Figs. 3 and 4) under the same condition. The active accumulation of TAG (specific TAG accumulation rate) in steady-state cells clearly indicated the concurrent microalgae growth and lipid accumulation. It is interesting that similar results were also found by Adams et al. (2013) in batch culture study of six species of oleaginous green algae; they concluded that some species were able to combine their growth and lipid accumulation phases. Why does SNI have the same effects on Chlorella growth and lipid accumulation? The reason may be accompanied accumulation of secondary carotenoids and fatty acids in cells of many Chlorophyta species when they are under stresses (Boussiba, 2000). 3.4. Comparison of chemostat and batch culture Batch culture of C. pyrenoidosa XQ-20044 was carried out in the same photobioreactor used for the continuous cultures, in order to supply basic data for comparing chemostat and batch cultures. Since the maximal lipid productivity in the chemostat was achieved with dilution rate of 0.48 d 1, the same amount of nitrate that was fed into the chemostat culture at dilution rate of 0.48 d 1
in 8 days was used to run an 8-day batch culture. Biomass productivity, lipid content and lipid productivity were determined and compared in Table 2. The chemostat culture with dilution rate of 0.48 d 1 had twice the biomass productivity of the batch culture, while the lipid content was somewhat lower than that in the batch culture. Due to the sustained algal growth and moderate lipid accumulation, the lipid productivity of the chemostat (144.93 mg L 1 d 1) was significantly higher than that of the batch culture (96.28 mg L 1 d 1). However, the lipids from both types of culture had very similar fatty acid profiles (Table 3) and the dominating fatty acid species in raw lipids were C16 and C18 (Table 2). Hence the authors reported here for the first time that, in comparison to batch culture, lipid productivity can be significantly enhanced by continuous cultivation of oleaginous microalgae with proper specific nitrate input rate (SNI). Also, to the best of our knowledge, this is the highest lipid productivity so far recorded in chemostat study for microalgal lipid production (Table 4). The new method proposed in this study provides an alternative way to produce lipids by using simultaneous microalgae growth and lipid accumulation, i.e., a one-step process, which was different from the classical two-step production mode using batch culture. This new strategy has a great advantage over batch culture in both biomass productivity and lipid productivity. When it is applied in mass culture of oleaginous microalgae, the cost of microalgal lipid production could be reduced significantly, thus promoting the commercialization of microalgal biodiesel. 4. Conclusions Batch and feed-batch culture of C. pyrenoidosa XQ-20044 showed the existence of certain conditions under which high lipid content could be achieved without sacrificing cell growth. Simultaneous microalgal growth and lipid production was achieved in continuous culture by properly manipulating the dilution rate. The optimal specific nitrate input rate for the concurrent growth and lipid accumulation was in the range of 0.78–4.56 mmol g 1 d 1 for C. pyrenoidosa XQ-20044 under experimental conditions. Enhanced lipid productivity (144.93 mg L 1 d 1) demonstrated the feasibility of one-step production of microalgal lipids by continuous culture, which provides a new strategy in development of microalgal biodiesel. Acknowledgements This work was supported by Ministry of Science and Technology of China (No. 2013AA065805) and National Natural Science Foundation of China (No. CNSF31272680). The authors thank Dr. David Herrin and Dr. Liming Luo for helpful discussions and careful editing of the manuscript. References Adams, C., Godfrey, V., Wahlen, B., Seefeldt, L., Bugbee, B., 2013. Understanding precision nitrogen stress to optimize the growth and lipid content tradeoff in oleaginous green microalgae. Bioresour. Technol. 131, 188–194.
X. Wen et al. / Bioresource Technology 161 (2014) 297–303 Boussiba, S., 2000. Carotenogenesis in the green alga Haematococcus pluvialis: cellular physiology and stress response. Physiol. Plant. 108, 111–117. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Collos, Y., Mornet, F., Sciandra, A., Waser, N., Larson, A., Harrison, P.J., 1999. An optical method for the rapid measurement of micromolar concentrations of nitrate in marine phytoplankton cultures. J. Appl. Phycol. 11, 179–184. Das, P., Aziz, S.S., Obbard, J.P., 2011. Two phase microalgae growth in the open system for enhanced lipid productivity. Renew. Energy 36, 2524–2528. Del Rio, E., Acien, G., Garcia-Malea, M.C., Rivas, J., Molina-Grima, E., Guerrero, M.G., 2005. Efficient one-step production of astaxanthin by the microalga Haematococcus pluvialis in continuous culture. Biotechnol. Bioeng. 91, 808–815. Dillschneider, R., Steinweg, C., Rosello-Sastre, R., Posten, C., 2013. Biofuels from microalgae: photoconversion efficiency during lipid accumulation. Bioresour. Technol. 142, 647–654. Gouveia, L., Marques, A.E., da Silva, T.L., Reis, A., 2009. Neochloris oleabundans UTEX #1185: a suitable renewable lipid source for biofuel production. J. Ind. Microbiol. Biotechnol. 36, 821–826. Grobbelaar, J.U., 2007. Algal nutrition: mineral nutrition. In: Richmond, A. (Ed.), Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell Publishing Ltd., Oxford, pp. 97–115. Han, F., Huang, J., Li, Y., Wang, W., Wan, M., Shen, G., Wang, J., 2013. Enhanced lipid productivity of Chlorella pyrenoidosa through the culture strategy of semicontinuous cultivation with nitrogen limitation and pH control by CO2. Bioresour. Technol. 136, 418–424. Hu, Q., Sommerfeld, M., Jarvis, E., Ghirardi, M., Posewitz, M., Seibert, M., Darzins, A., 2008. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J. 54, 621–639. Klok, A.J., Martens, D.E., Wijffels, R.H., Lamers, P.P., 2013. Simultaneous growth and neutral lipid accumulation in microalgae. Bioresour. Technol. 134, 233–243. Kwak, K.O., Jung, S., Chung, S.Y., Kang, C.M., Huh, Y.I., Bae, S.O., 2006. Optimization of culture conditions for CO2 fixation by a chemoautotrophic microorganism, strain YN-1 using factorial design. Biochem. Eng. J. 31, 1–7. Lee, Y.K., Ding, S.Y., Hoe, C.H., Low, C.S., 1996. Mixotrophic growth of Chlorella sorokiniana in outdoor enclosed photobioreactor. J. Appl. Phycol. 8, 163–169.
303
Liu, Z.Y., Wang, G.C., Zhou, B.C., 2008. Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresour. Technol. 99, 4717–4722. Lourenco, S.O., Barbarino, E., Mancini-Filho, J., Schinke, K.P., Aidar, E., 2002. Effects of different nitrogen sources on the growth and biochemical profile of 10 marine microalgae in batch culture: an evaluation for aquaculture. Phycologia 41, 158–168. Merzlyak, M.N., Chivkunova, O.B., Gorelova, O.A., Reshetnikova, I.V., Solovchenko, A.E., Khozin-Goldberg, I., Cohen, Z., 2007. Effect of nitrogen starvation on optical properties, pigments, and arachidonic acid content of the unicellular green alga Parietochloris incisa (Trebouxiophyceae, Chlorophyta). J. Phycol. 43, 833–843. Pan, Y.Y., Wang, S.T., Chuang, L.T., Chang, Y.W., Chen, C.N., 2011. Isolation of thermotolerant and high lipid content green microalgae: oil accumulation is predominantly controlled by photosystem efficiency during stress treatments in Desmodesmus. Bioresour. Technol. 102, 10510–10517. Sobczuk, T., Chisti, Y., 2010. Potential fuel oils from the microalga Choricystis minor. J. Chem. Technol. Biotechnol. 85, 100–108. Su, C.H., Chien, L.J., Gomes, J., Lin, Y.S., Yu, Y.K., Liou, J.S., Syu, R.J., 2011. Factors affecting lipid accumulation by Nannochloropsis oculata in a two-stage cultivation process. J. Appl. Phycol. 23, 903–908. Tang, H., Chen, M., Simon Ng, K.Y., Salley, S.O., 2012. Continuous microalgae cultivation in a photobioreactor. Biotechnol. Bioeng. 109, 2468–2474. Wang, G., Wang, T., 2012. Characterization of lipid components in two microalgae for biofuel application. J. Am. Oil Chem. Soc. 89, 135–143. Wen, X.B., Jiang, L.L., Geng, Y.H., Shen, X., Li, Y.G., 2012. Comparative study of methods for microalgal total lipid quantification (in Chinese with English title and abstracts). China Oils Fats 37, 80–85. Wijffels, R.H., Barbosa, M.J., 2010. An outlook on microalgal biofuels. Science 329, 796–799. Zar, J.H., 1999. Biostatistical Analysis, fourth ed. Prentice-Hall, New Jersey. Zhang, B.Y., Geng, Y.H., Li, Z.K., Hu, H.J., Li, Y.G., 2009. Production of astaxanthin from Haematococcus in open pond by two-stage growth one-step process. Aquaculture 295, 275–281. Zhang, Y.M., Chen, H., He, C.L., Wang, Q., 2013. Nitrogen starvation induced oxidative stress in an oil-producing green alga Chlorella sorokiniana C3. PLoS ONE 8, e69225.
Contents lists available at ScienceDirect
Bioresource Technology journal homepage: www.elsevier.com/locate/biortech
Enhanced lipid production in Chlorella pyrenoidosa by continuous culture Xiaobin Wen a,b, Yahong Geng a, Yeguang Li a,⇑ a b
Key Laboratory of Plant Germplasm Enhancement and Speciality Agriculture, Wuhan Botanical Garden, Chinese Academy of Sciences, Wuhan 430074, China University of Chinese Academy of Sciences, Beijing 100049, China
h i g h l i g h t s Chlorella pyrenoidosa XQ-20044 is able to accumulate lipids in growing cells. One step production of algal lipid was achieved in chemostat culture. Proper SNI was the key for simultaneous algal growth and lipid accumulation. Lipid productivity was significantly enhanced by continuous culture.
a r t i c l e
i n f o
Article history: Received 24 January 2014 Received in revised form 12 March 2014 Accepted 16 March 2014 Available online 25 March 2014 Keywords: Chlorella Growth Lipid productivity Continuous culture Chemostat
a b s t r a c t Usually microalgae growth and lipid accumulation do not run in parallel throughout cultivation, which necessarily lowers overall lipid productivity. However, we show through batch and feed-batch studies of Chlorella pyrenoidosa XQ-20044 that by varying the nitrate concentration, conditions which produce fairly high lipid content could be achieved without sacrificing algal growth. Simultaneous microalgae growth and lipid production was achieved in continuous chemostat culture when the specific nitrate input rate was in the range of 0.78–4.56 mmol g 1 d 1. Moreover, the maximum lipid productivity (144.93 mg L 1 d 1) in the continuous culture was significantly higher than in batch culture (96.28 mg L 1 d 1), thus indicating the feasibility and great advantage of one-step production of microalgal lipids. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction Aquatic microalgae have a high capacity for photosynthesis, and many are able to use excess photosynthetically-fixed carbon for synthesis of neutral lipids, which are also known as triacylglycerols (TAGs) (Chisti, 2007). Oleaginous microalgae are thus considered to be ideal raw materials for biodiesel production, especially if their growth is coupled to the direct bio-fixation of waste CO2 (Kwak et al., 2006). Over the past 50 years the concept and feasibility of microalgal biodiesel have been discussed extensively (Wijffels and Barbosa, 2010), but this renewable energy source has yet to be exploited. Limiting factors include the lack of appropriate microalgal strains, less-than optimal lipid productivity and ineffective culturing techniques for lipid accumulation.
⇑ Corresponding author. Address: Wuhan Botanical Garden, Chinese Academy of Sciences, Wuhan 430074, Hubei Province, China. Tel.: +86 27 87510542; fax: +86 27 87510251. E-mail address: [email protected] (Y. Li). http://dx.doi.org/10.1016/j.biortech.2014.03.077 0960-8524/Ó 2014 Elsevier Ltd. All rights reserved.
To date, several hundred oleaginous species have been isolated and characterized (Gouveia et al., 2009). A common thread in these studies is that cell growth and lipid accumulation do not happen at the same time during cultivation (Lourenco et al., 2002; Merzlyak et al., 2007), which results in lower overall lipid productivity. In order to overcome this, researchers have explored two-stage cultivation strategies to enhance microalgal lipid production. In such strategies, which have been used mainly with batch cultures, the microalgal cells first grow rapidly under growth-optimized conditions, and then are transferred to conditions where light irradiance (Zhang et al., 2009), nutrition (Su et al., 2011), culture pH (Han et al., 2013), as well as other factors (Das et al., 2011; Liu et al., 2008) are adjusted to promote lipid accumulation at the expense of cell growth. However, detailed study of the stress response of oleaginous microalgae under nitrogen deficiency may provide a foundation for the production of biomass and lipids in one step. Recent data showed that while Neochloris oleoabundans UTEX #1185 accumulated lipid under relatively high nitrogen stress conditions (i.e., low nitrogen levels), its growth was not severely limited (Adams
298
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
et al., 2013). In contrast, growth of Chlorella vulgaris UTEX #265 was severely limited under high nitrogen stress, but lipid accumulation was triggered ahead of the growth limitation (Adams et al., 2013). These may or may not be species-specific characteristics, but the results have inspired the authors to explore the threshold values of nitrogen concentration and/or supply rate, in order to establish a continuous cultivation system for concurrent production of biomass and lipids. It is hypothesized that by properly regulating the nitrate concentration (e.g., severe stress or moderate stress) in continuous culture, the algal cells could grow at a reasonable rate and accumulate lipid simultaneously, thus leading to higher lipid productivity over batch culture. Along this line, one-step astaxanthin production was achieved with continuous culture of Haematococcus pluvialis, where >0.8% (DW) astaxanthin accumulated in the green vegetative cells (Del Rio et al., 2005). However, only a few studies have been published on the continuous cultivation of microalgae for lipid production. Sobczuk and Chisti (2010) investigated the effects of dilution rate on lipid productivity of the freshwater microalga Choricystis minor in chemostat culture, and concluded that the lipid content did not change significantly with various dilution rates. Very similar results were reported for chemostat cultures of Chlorella minutissima and Dunaliella tertiolecta by Tang et al. (2012). In contrast, Klok et al. (2013) reported that excess light combined with a growth-limiting nitrogen supply resulted in TAG accumulation (up to 12.4%, w/w) and cell replication in turbidostat cultures of Neochloris oleoabundans. Given the quite different results of Klok et al., it is an open question how other microalgal species will respond in continuous culture. In this report, batch and feed-batch cultures of C. pyrenoidosa were used to study its response, in terms of growth and lipid accumulation, to different nitrogen concentrations. The results were used to establish chemostat cultures with different dilution rates, in order to verify the assumption that microalgal cultures can be regulated to grow and accumulate lipids concurrently, thus enhancing lipid productivity. 2. Methods 2.1. Strains and pre-culture conditions C. pyrenoidosa XQ-20044 was used in this study. It was provided by the Algae Culture Collection of Wuhan Botanical Garden, Chinese Academy of Sciences. This fast growing strain was originally isolated from a Spirulina culture pond in Sichuan province, China. Its lipid content in a column photobioreactor was >45% of dry biomass (unpublished results), thus showing its potential as a biodiesel feedstock. For seed cultures, the algal cells were grown photoautotrophically in Erlenmeyer flasks at 25 °C. The flasks contained 600 mL of medium, and were placed on a shaker (100 rpm) for 48 h of cultivation before further experiments. Continuous illumination was provided by fluorescent lamps, and the light intensity on the surface of the flask was adjusted to 50 lmol m 2 s 1. The basal growth medium for seed culture and culture experiments was a modified BG11 medium which had the following composition (per liter): NaNO3, 100 mg; K2HPO43H2O, 40 mg; MgSO47H2O, 75 mg; CaCl22H2O, 36 mg; Citric acid, 6 mg; Fe-Ammonium citrate, 6 mg; EDTANa2, 1 mg; Na2CO3, 20 mg; H3BO3, 2.86 lg; MnCl24H2O 1.8 lg; ZnSO47H2O, 0.22 lg; CuSO45H2O, 0.08 lg; Na2MoO42H2O, 0.391 lg; Co(NO3)26H2O, 0.0494 lg (Grobbelaar, 2007). Sodium nitrate concentration was modified as indicated in the text. 2.2. Batch cultures Batch cultures of C. pyrenoidosa XQ-20044 were grown in aerated-column photobioreactors. The seed cultures were
centrifuged, rinsed, and resuspended in nitrate-free BG-11 medium to an optical density of 0.5 ± 0.05 at 540 nm. Then aliquots of this suspension were transferred to the glass columns (inner diameter 3 cm), which were submerged in a water tank. A thermostatic water circulator was used to provide 30 °C water bath for the culture columns. CO2-enriched (1%, v/v) air was passed through a sterile filter and then bubbled into the bottom of each column at a flow rate of 250 ml min 1. Light (300 lmol m 2 s 1 irradiance impinging on the surface of each column) was continuously provided by ten Phillips 36 W cool white fluorescent lamps. A concentrated sodium nitrate stock solution (autoclaved and stored at 4 °C) was used to adjust the columns to initial nitrate concentrations that ranged from 0.24 mM to 17.65 mM. And for each nitrate concentration, three replicate cultures were conducted in parallel. After 8 d of cultivation, cells were collected by centrifugation (5000 rpm at 15 °C for 5 min) and lyophilized ( 56 °C cryotrapping, 10–14 Pa vacuum) for further analysis. 2.3. Feed-batch cultures Feed-batch culturing of C. pyrenoidosa XQ-20044 was carried out similar to the batch cultures, except nitrate (in concentrations of 0.24, 0.48, 0.72, 0.96, 1.18, 2.35 and 3.53 mM) was added to the photobioreactors at 0, 24, 48, 72, and 96 h of cultivation. In the batch cultures with an initial nitrate concentration >3.53 mM, the residual nitrate level after 24 h was >50% of the initial value. Thus, only nitrate concentrations ranging from 0.24 mM to 3.53 mM were investigated in these feed-batch cultures. 2.4. Chemostat cultures A peristaltic pump with a ten-roller pump head was used for the continuous cultures. It delivered a maximum flow rate of 32 mL min 1 and a minimum flow rate of 0.1 mL min 1. Exactly 2000 mL of algae were grown in a closed column bioreactor with an inner diameter of 10 cm. A silicone tube was connected to the bottom of the column to form a U-shaped overflow tube. Air enriched with CO2 (1%, v/v) was passed through a sterile Millex syringe filter (0.22 lm) and then bubbled into the bottom of the culture at a flow rate of 2 L min 1; this continuously agitated the cells, so they received equivalent light. The culture pH was maintained in the range of 7–8, and continuous illumination (600 lmol m 2 s 1 photon flux) was provided by four fluorescent lamps (Philips MASTER PL-L 36 W) on one side of the bioreactor. The whole cultivation unit was placed in a thermostatic room to maintain the culture temperature at 30 °C. Cultivations (in triplicate) were carried out initially in batch mode by inoculating the seed culture into BG-11 medium to an optical density of 0.1 (540 nm). After 30 h of cultivation, when the optical density of the algal suspension had increased to 1.0 or more, the bioreactor was switched to continuous mode by feeding BG-11 medium (0.71 mM sodium nitrate). Cultivations with different dilution rates (ranged from 0.24 d 1 to 2.4 d 1) were carried out successively. Achievement of steady-state condition at each dilution rate was monitored by daily measuring of the optical density, biomass dry weight, and residual nitrate concentration. The steady state was maintained at least for three days before further determinations. 2.5. Analytical methods Biomass dry weight was measured to evaluate microalgal growth. About 10 mL algal suspension was filtered through a predried GF/C glass microfiber filter paper (0.45 lm), which was dried at 80 °C under vacuum for 4 h (Lee et al., 1996) and re-weighed to
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
calculate biomass dry weight (DW). A spectrophotometric method, described by Collos et al. (1999), was used to monitor residual nitrate concentration. Briefly, the absorbance of culture filtrate (0.22 lm filter) at 220 nm and a pre-constructed standard curve were used to determine residual nitrate. Pigments were extracted from live cells with hot DMSO (70 °C) and quantified spectrophotometrically using coefficients determined by Merzlyak et al. (2007). The photosynthetic status of the cells was evaluated by measuring the net photosynthetic oxygen evolution rate (Zhang et al., 2013) using a liquid phase oxygen measurement system with white light. For biochemical composition analysis, the microalgal cells were harvested and vacuum lyophilized. For lipid quantification, 50 mg of dry algal biomass was fully ground, transferred to a covered centrifuge tube and extracted with a mixture of n-hexane and ethyl acetate (1:1, v:v). The extraction was repeated 3 times and all extracts were combined into a pre-weighed glass tube, and then dried with a stream of nitrogen (Wen et al., 2012). The lipids were determined gravimetrically. Neutral lipid, glycolipids, and phospholipids were fractionated from the lipid extracts (100 mg) by column chromatography using a 2 cm 20 cm column packed with 4 g silica gel 60 (Wang and Wang, 2012). The lipids were differentially eluted, first with 20 mL of chloroform to obtain the neutral lipids, and then with 15 mL of acetone:methanol (9:1, v:v) to obtain the glycolipids; phospholipids were eluted last with 15 mL of methanol. The fractions were confirmed by TLC separation, and then quantified gravimetrically. In order to analyze the fatty acids, about 20 mg of the lipid extract was dissolved in 3 mL of n-hexane, and then transmethylated by adding 3 mL of methanol–KOH (0.5% KOH) and heating at 50 °C for 60 min. After cooling to room temperature, the hexane layer was separated and dried with anhydrous sodium sulfate. Fatty acid methyl esters were analyzed by gas chromatography (Agilent 7890A) using an HP-5 Phenyl Methyl Siloxan column (30 m 0.32 mm 0.25 lm) and a flame ionization detector. 1 lL samples were injected with a splitting ratio of 5:1 at 250 °C. The heating program was 150 °C for 2 min, followed by an increase in the temperature of 10 °C/min to 250 °C, where it was held for 8 min. The detector temperature was set at 240 °C. A Supelco 37Component FAME Mix (47885-U) was used as external standard for fatty acid identification. All of the above analytical experiments were done in triplicate, and the results were analyzed for variance (Zar, 1999) with SAS 8.01 at a significance level a = 0.05. Tukey’s multiple comparison tests were done where applicable.
299
3. Results and discussion 3.1. Non-parallel occurrence of cell growth and lipid accumulation in batch culture Nitrogen starvation has frequently been reported to induce lipid accumulation in microalgal cells (Hu et al., 2008). To clarify the growth and lipid responses of C. pyrenoidosa XQ-20044 to varying nitrate levels, a series of batch cultures were carried out at different initial nitrate concentrations (Fig. 1). The results showed that biomass dry weight (DW) of C. pyrenoidosa XQ-20044 increased almost linearly with increasing initial nitrate from 0.24 mM to 2.35 mM. Further increases in the nitrate concentration did not affect biomass accumulation, possibly due to light limitation (Fig. 1A). The lipid content (% DW), however, exhibited different trends in response to increasing nitrate concentration (Fig. 1A). Specifically, the lipid content first increased from 38.79% to 52.36% (DW), and then declined to 39.59% (DW), as the initial nitrate concentration was increased from 0.24 mM to 5.88 mM. Further increasing the initial nitrate concentration (above 5.88 mM) had no significant influence on lipid content. Changes of the nitrate concentration during culture are also indicated in Fig. 1B. These results showed that cell growth and lipid accumulation did not always change in parallel, and moreover, they suggested relatively high lipid content could be achieved without the expense of algal growth. As shown in Fig. 1A, decreasing the nitrate concentration from 5.88 mM to 2.35 mM significantly increased the lipid content (p < 0.05), while the biomass was not affected (1.8 g L 1, p > 0.05). 3.2. Effects of intermittent nitrate feeding on cell growth and lipid accumulation To confirm that there is a culture status under which C. pyrenoidosa XQ-20044 moderately accumulates lipids in growing cells, feed-batch culture was conducted with initial nitrate concentrations of 0.24, 0.47, 0.71, 0.94, 1.18, 2.35 and 3.53 mM, and the same amount of nitrate (as the initial concentration) was fed respectively into each culture every 24 h. A similar response of biomass dry weight and lipid content to varying nitrate was observed (Fig. 2A). In particular, the cultures with initial nitrate concentration of 0.71, 0.94 and 1.18 mM reached maximal biomass density of about 2.2 g L 1 when nitrate was fed intermittently, and all of them showed fairly high lipid content (35–45% DW). The nitrate concentration of the 3 cultures fluctuated only within a small range during the feed-batch culture
Fig. 1. (A) Effects of initial nitrate concentration on biomass dry weight (j) and lipid content (N) in batch culture of C. pyrenoidosa XQ-20044. (B) Changes of nitrate concentration during batch culture of C. pyrenoidosa XQ-20044. More than 2 mM nitrate was detected at the end of the culture with initial nitrate concentrations higher than 6 mM (data not shown).
300
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Fig. 2. (A) Effects of intermittent nitrate feeding on biomass dry weight (j) and lipid content (N) of C. pyrenoidosa XQ-20044. (B) Changes of nitrate concentration during batch culture of C. pyrenoidosa XQ-20044.
Fig. 3. Effects of dilution rate on steady-state biomass concentration (j), lipid content (N) and nitrate input rate (s) in chemostat culture of C. pyrenoidosa XQ20044. Experiments were carried out with constant feeding medium composition (BG-11 with 0.71 mM nitrate) and illumination (600 lmol m 2 s 1), but with varying dilution rate between 0.24 and 2.4 d 1.
(Fig. 2B). These results suggested that microalgal growth and lipid accumulation could both be optimized by using continual feeding of small amounts of nitrate. The nitrate concentration and biomass density of culture changes over time in both batch and feed-batch mode, thus the optimum status for concurrent microalgae growth and lipid accumulation cannot be sustained using these two modes. However, continuous cultivation of C. pyrenoidosa XQ-20044 in chemostat mode could potentially overcome this problem, and thus was investigated. 3.3. Concurrent microalgae growth and lipid accumulation in continuous culture Steady-state studies of C. pyrenoidosa XQ-20044 were carried out using a chemostat system with constant medium composition (BG-11 with 0.71 mM nitrate), illumination and air flow rate, but with dilution rates that varied between 0.24 and 2.4 d 1. Steadystates were achieved with all the tested dilution rates. The cultures with high dilution rate showed a green color while the cultures with low dilution rate had a yellow-green appearance. 3.3.1. Microalgal growth As shown in Fig. 3, the nitrate input rate, which varied from 0.17 to 1.69 mol L 1 d 1, correlated linearly with dilution rate. Also, since the nitrate level in all steady-state cultures was below detection limit (0.01 mM), the nitrate uptake rate was equal to the
Fig. 4. Effects of dilution rate on pigment content (Chla h, Total Car e) and net photosynthetic oxygen evolution rate (N) in chemostat culture of C. pyrenoidosa XQ20044.
nitrate input rate. Dilution rate also had a remarkable impact on the biomass concentration at each steady-state (Fig. 3). With increasing dilution rates from 0.25 d 1 to 2.4 d 1, the biomass dry weight was decreased 86%, from 0.57 g L 1 to 0.08 g L 1. The pigment content and net photosynthetic oxygen evolution rate of C. pyrenoidosa XQ-20044 as a function of dilution rate at each steady-state are illustrated in Fig. 4. The lowest contents of Chlorophyll a (1.33 mg g 1) and total carotenoids (1.00 mg g 1) in the cells were observed at the lowest dilution rate of 0.25 d 1. However, with increasing dilution rate, both Chlorophyll a and total carotenoids increased, and reached their maximum values (24.16 mg g 1 and 6.24 mg g 1, respectively), at the steady-state with the dilution rate of 2.4 d 1. The net photosynthetic oxygen evolution rate, which is a substantial growth indicator for autotrophic organisms, was calculated from the measured changes in dissolved oxygen per unit time and the Chlorophyll a content of the culture. As shown in Fig. 4, the rapidly growing cells (with high dilution rate) presented high photosynthetic oxygen evolution rate. However, when the dilution rates (growth rates) were decreased from 1.44 d 1 to 0.48 d 1, the net photosynthetic oxygen evolution rate was only decreased by 13%. Statistic analysis showed that the cells maintained a relative stable photosynthetic activity when the dilution rates were 0.48, 0.72, 0.96, and 1.44 d 1. Out of this range, the photosynthetic oxygen evolution changed significantly (p > 0.05) when changing the dilution rates. Photosynthetic activity of nitrate-stressed microalgae is essential for carbon storage metabolism, especially for triglyceride accumulation (Dillschneider et al., 2013; Pan et al., 2011). In the chemostat
301
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Table 3 Comparison of fatty acid profiles of C. pyrenoidosa XQ-20044 in chemostat and batch culture. Fatty acid
Fig. 5. Biomass (j) and lipid (N) productivity as affected by the dilution rate in chemostat culture of C. pyrenoidosa XQ-20044.
C14:0 C15:0 C16:0 C16:1 C17:0 C17:1 C18:0 C18:1n9c C18:2n6t C18:3n3 C20:0 C20:1n9 C22:0 C22:1n9
% of Total fatty acids Batch culture
Chemostat
0.31 ± 0.06 0.21 ± 0.03 25.53 ± 0.12 9.4 ± 0.16 0.67 ± 0.05 0.4 ± 0.03 7.39 ± 0.56 27.45 ± 0.40 12.69 ± 0.34 11.22 ± 0.49 2.05 ± 0.09 2.21 ± 0.42 ND 0.45 ± 0.09
0.38 ± 0.05 0.18 ± 0.02 22.91 ± 0.13 9.72 ± 0.23 0.53 ± 0.05 0.42 ± 0.03 6.1 ± 0.19 29.2 ± 0.54 13.42 ± 0.05 12.57 ± 0.36 2.19 ± 0.09 2.01 ± 0.42 0.33 ± 0.12 ND
ND: not detected. Table 1 Lipid content and composition (means + SD) of C. pyrenoidosa XQ-20044 at steadystate with various dilution rates. Dilution rate
Lipid content (% DW)
Lipid class distribution (% total lipids) Neutral lipids
Glycolipids
Phospholipids
0.24 0.48 0.72 0.96 1.44 1.92 2.40
30.04 ± 1.35 34.69 ± 0.68 25.51 ± 1.06 23.32 ± 1.31 19.97 ± 0.13 14.92 ± 1.18 14.60 ± 1.28
76.19 ± 2.15 81.62 ± 1.47 78.06 ± 1.45 69.65 ± 0.77 48.51 ± 2.31 37.04 ± 0.87 36.96 ± 1.55
15.06 ± 0.97 11.55 ± 0.12 16.43 ± 0.83 22.15 ± 1.01 43.05 ± 1.05 52.76 ± 0.09 52.17 ± 1.02
4.71 ± 0.52 4.06 ± 0.77 4.56 ± 0.04 5.45 ± 0.13 5.42 ± 0.13 8.15 ± 0.43 7.85 ± 0.42
Fig. 6. Nitrate uptake (line) and specific TAG accumulation rate (column) as affected by specific nitrate input rate in chemostat culture of C. pyrenoidosa XQ20044.
culture of this study, the photosynthetic activity sustained, rather than declined, when the dilution rate (growth rate) was decreased from 1.44 d 1 to 0.48 d 1 (Fig. 4). It will be shown below that lipid
productivity at these dilution rates (from 0.48 to 1.44 d 1) was significantly higher (p < 0.05) than at other dilution rates (Fig. 5). The sustained photosynthetic activity of C. pyrenoidosa XQ-20044 might contribute to triglyceride accumulation. 3.3.2. Lipid accumulation The dilution rate had differing impacts on lipid content in different steady-state conditions (Fig. 3). The highest lipid content (34.69% DW) was observed with the dilution rate of 0.48 d 1. Increasing the dilution rate (i.e., from 0.48 to 1.92 d 1) led to a decline in the lipid content. When the dilution rate was further increased from 1.92 d 1 to 2.4 d 1, the lipid content (14.92% DW) did not change due to sufficient nitrate supply. A similar phenomenon was reported before (Sobczuk and Chisti, 2010; Tang et al., 2012); in those studies, the chemostat culture had relatively high nitrate concentration (feeding medium) and no significant changes in lipid content were observed under different dilution rates. The results from this study also suggested that severe nitrogen limitation (the lowest dilution rate) reduced lipid content as well as growth (Fig. 3), which was different from previous studies reported that lipid accumulation varies inversely in response to nitrate concentration (Lourenco et al., 2002; Merzlyak et al., 2007). Fractionation of the lipids by column chromatography (Table 1) revealed that neutral lipids (mainly TAG) were the main component with dilution rate less than or equal to 0.96 d 1. Glycolipids had higher percentage (about 50%) of the lipids when the dilution rate was increased to 1.92 d 1 or more. Apparently cellular TAG accumulation was triggered when steady-state was attained at dilution rate of 0.24, 0.48, 0.72, 0.96, and 1.44 d 1 respectively. 3.3.3. Productivity of biomass and lipid The biomass productivity of chemostat culture increased dramatically in response to increasing dilution rates (from 0.24 d 1 to 1.44 d 1), reaching a peak of 641.52 mg L 1 d 1 at the dilution
Table 2 Comparison of growth and lipid accumulation in chemostat and batch culture of C. pyrenoidosa XQ-20044. Culture mode
Dilution rate (d 1)
Nitrate consumed (mM/ 8 d)
Biomass productivity (mg L 1 d 1)
Lipid content (% DW)
Lipid productivity (mg L 1 d 1)
C16 + C18 (% total FFA)
Chemostat Chemostat Chemostat Chemostat Batch
0.48 0.72 0.96 1.44 /
2.71 4.07 5.42 8.13 2.71
417.81 ± 26.88 482.40 ± 13.15 539.87 ± 30.73 641.52 ± 24.37 215.81 ± 8.08
34.69 ± 0.68 25.51 ± 1.06 23.32 ± 1.31 19.97 ± 063 44.07 ± 6.04
144.93 ± 5.16 124.25 ± 5.65 125.89 ± 8.50 128.12 ± 4.28 96.28 ± 4.55
93.68 ± 1.15 95.22 ± 0.97 93.47 ± 2.04 92.12 ± 1.42 93.92 ± 1.17
302
X. Wen et al. / Bioresource Technology 161 (2014) 297–303
Table 4 Biomass and lipid productivity reported in literatures dealing with microalgal lipid production in chemostat culture. Species
Dilution rate (d 1)
Nitrate in feeding medium (mM)
Biomass productivity (mg L 1 d 1)
Lipid productivity (mg L 1 d 1)
References
Chlorella pyrenoidosa XQ-20044 Chlorella minutissima (UTEX 2219) Dunaliella tertiolecta (UTEX LB 999) Choricystis minor B. Fott
0.48 0.328 0.42 0.336
0.71 8.82 2.18 17.6
417 ± 26 137 91 351
144 ± 5 6 10 82
This study Tang et al. (2012) Tang et al. (2012) Sobczuk and Chisti (2010)
rate of 1.44 d 1 (Fig. 5). However, the biomass productivity started to decrease when the dilution rates were further increased from 1.44 to 2.40 d 1. A similar pattern was also observed in lipid productivity in response to dilution rate, except that the maximum lipid productivity (144.93 mg L 1 d 1) was achieved with a dilution rate of 0.48 d 1 (Fig. 5). ANOVA analysis concluded that the lipid productivity at dilution rates of 0.48, 0.72, 0.96 and 1.44 d 1 were significantly higher (p < 0.05) than that achieved with other dilution rates. Few studies has been conducted in C. pyrenoidosa previously but maximal biomass productivity attained in C. minutissima was 137 mg L 1 d 1 with 6 mg L 1 d 1 FAME productivity (Tang et al., 2012). Sobczuk and Chisti (2010) reported the maximal biomass productivity of 351 mg L 1 d 1 with 82 mg L 1 d 1 lipid productivity when Choricystis minor chemostat culture was performed with a dilution rate of 0.336 d 1. The biomass productivity and lipid productivity in this study were both higher than previously reported (Table 4). So, what is the favorable condition for concurrent growth and lipid accumulation for C. pyrenoidosa XQ-20044? When the dilution rate was in the range of 0.48–1.44 d 1, all of the supplied nitrate was assimilated by cells and the chemostat cultures showed moderate cell growth (Fig. 3) and photosynthetic activity (Fig. 4). Importantly, TAG accumulation also took place in the cells under the above-mentioned conditions (Fig. 3 and Table 1). Therefore, the lipid productivity was significantly improved, as indicated in Fig. 5. The specific nitrate input rate, or SNI (mmol g 1 d 1), was one of the key factors proposed by Del Rio et al. (2005) for simultaneous cell growth and astaxanthin accumulation in Haematococcus. As illustrated in Fig. 6, only when the SNI was in the range of 0.78–4.56 mmol g 1 d 1 (with a corresponding nitrate uptake of 0.81–1.59 mmol per gram of biomass) could a high rate of specific TAG accumulation (TAG increase per g biomass and unit time) be attained in the chemostat cultures of C. pyrenoidosa XQ-20044. Meanwhile the net photosynthetic oxygen evolution rate and cell growth were limit to some extent (Figs. 3 and 4) under the same condition. The active accumulation of TAG (specific TAG accumulation rate) in steady-state cells clearly indicated the concurrent microalgae growth and lipid accumulation. It is interesting that similar results were also found by Adams et al. (2013) in batch culture study of six species of oleaginous green algae; they concluded that some species were able to combine their growth and lipid accumulation phases. Why does SNI have the same effects on Chlorella growth and lipid accumulation? The reason may be accompanied accumulation of secondary carotenoids and fatty acids in cells of many Chlorophyta species when they are under stresses (Boussiba, 2000). 3.4. Comparison of chemostat and batch culture Batch culture of C. pyrenoidosa XQ-20044 was carried out in the same photobioreactor used for the continuous cultures, in order to supply basic data for comparing chemostat and batch cultures. Since the maximal lipid productivity in the chemostat was achieved with dilution rate of 0.48 d 1, the same amount of nitrate that was fed into the chemostat culture at dilution rate of 0.48 d 1
in 8 days was used to run an 8-day batch culture. Biomass productivity, lipid content and lipid productivity were determined and compared in Table 2. The chemostat culture with dilution rate of 0.48 d 1 had twice the biomass productivity of the batch culture, while the lipid content was somewhat lower than that in the batch culture. Due to the sustained algal growth and moderate lipid accumulation, the lipid productivity of the chemostat (144.93 mg L 1 d 1) was significantly higher than that of the batch culture (96.28 mg L 1 d 1). However, the lipids from both types of culture had very similar fatty acid profiles (Table 3) and the dominating fatty acid species in raw lipids were C16 and C18 (Table 2). Hence the authors reported here for the first time that, in comparison to batch culture, lipid productivity can be significantly enhanced by continuous cultivation of oleaginous microalgae with proper specific nitrate input rate (SNI). Also, to the best of our knowledge, this is the highest lipid productivity so far recorded in chemostat study for microalgal lipid production (Table 4). The new method proposed in this study provides an alternative way to produce lipids by using simultaneous microalgae growth and lipid accumulation, i.e., a one-step process, which was different from the classical two-step production mode using batch culture. This new strategy has a great advantage over batch culture in both biomass productivity and lipid productivity. When it is applied in mass culture of oleaginous microalgae, the cost of microalgal lipid production could be reduced significantly, thus promoting the commercialization of microalgal biodiesel. 4. Conclusions Batch and feed-batch culture of C. pyrenoidosa XQ-20044 showed the existence of certain conditions under which high lipid content could be achieved without sacrificing cell growth. Simultaneous microalgal growth and lipid production was achieved in continuous culture by properly manipulating the dilution rate. The optimal specific nitrate input rate for the concurrent growth and lipid accumulation was in the range of 0.78–4.56 mmol g 1 d 1 for C. pyrenoidosa XQ-20044 under experimental conditions. Enhanced lipid productivity (144.93 mg L 1 d 1) demonstrated the feasibility of one-step production of microalgal lipids by continuous culture, which provides a new strategy in development of microalgal biodiesel. Acknowledgements This work was supported by Ministry of Science and Technology of China (No. 2013AA065805) and National Natural Science Foundation of China (No. CNSF31272680). The authors thank Dr. David Herrin and Dr. Liming Luo for helpful discussions and careful editing of the manuscript. References Adams, C., Godfrey, V., Wahlen, B., Seefeldt, L., Bugbee, B., 2013. Understanding precision nitrogen stress to optimize the growth and lipid content tradeoff in oleaginous green microalgae. Bioresour. Technol. 131, 188–194.
X. Wen et al. / Bioresource Technology 161 (2014) 297–303 Boussiba, S., 2000. Carotenogenesis in the green alga Haematococcus pluvialis: cellular physiology and stress response. Physiol. Plant. 108, 111–117. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294–306. Collos, Y., Mornet, F., Sciandra, A., Waser, N., Larson, A., Harrison, P.J., 1999. An optical method for the rapid measurement of micromolar concentrations of nitrate in marine phytoplankton cultures. J. Appl. Phycol. 11, 179–184. Das, P., Aziz, S.S., Obbard, J.P., 2011. Two phase microalgae growth in the open system for enhanced lipid productivity. Renew. Energy 36, 2524–2528. Del Rio, E., Acien, G., Garcia-Malea, M.C., Rivas, J., Molina-Grima, E., Guerrero, M.G., 2005. Efficient one-step production of astaxanthin by the microalga Haematococcus pluvialis in continuous culture. Biotechnol. Bioeng. 91, 808–815. Dillschneider, R., Steinweg, C., Rosello-Sastre, R., Posten, C., 2013. Biofuels from microalgae: photoconversion efficiency during lipid accumulation. Bioresour. Technol. 142, 647–654. Gouveia, L., Marques, A.E., da Silva, T.L., Reis, A., 2009. Neochloris oleabundans UTEX #1185: a suitable renewable lipid source for biofuel production. J. Ind. Microbiol. Biotechnol. 36, 821–826. Grobbelaar, J.U., 2007. Algal nutrition: mineral nutrition. In: Richmond, A. (Ed.), Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell Publishing Ltd., Oxford, pp. 97–115. Han, F., Huang, J., Li, Y., Wang, W., Wan, M., Shen, G., Wang, J., 2013. Enhanced lipid productivity of Chlorella pyrenoidosa through the culture strategy of semicontinuous cultivation with nitrogen limitation and pH control by CO2. Bioresour. Technol. 136, 418–424. Hu, Q., Sommerfeld, M., Jarvis, E., Ghirardi, M., Posewitz, M., Seibert, M., Darzins, A., 2008. Microalgal triacylglycerols as feedstocks for biofuel production: perspectives and advances. Plant J. 54, 621–639. Klok, A.J., Martens, D.E., Wijffels, R.H., Lamers, P.P., 2013. Simultaneous growth and neutral lipid accumulation in microalgae. Bioresour. Technol. 134, 233–243. Kwak, K.O., Jung, S., Chung, S.Y., Kang, C.M., Huh, Y.I., Bae, S.O., 2006. Optimization of culture conditions for CO2 fixation by a chemoautotrophic microorganism, strain YN-1 using factorial design. Biochem. Eng. J. 31, 1–7. Lee, Y.K., Ding, S.Y., Hoe, C.H., Low, C.S., 1996. Mixotrophic growth of Chlorella sorokiniana in outdoor enclosed photobioreactor. J. Appl. Phycol. 8, 163–169.
303
Liu, Z.Y., Wang, G.C., Zhou, B.C., 2008. Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresour. Technol. 99, 4717–4722. Lourenco, S.O., Barbarino, E., Mancini-Filho, J., Schinke, K.P., Aidar, E., 2002. Effects of different nitrogen sources on the growth and biochemical profile of 10 marine microalgae in batch culture: an evaluation for aquaculture. Phycologia 41, 158–168. Merzlyak, M.N., Chivkunova, O.B., Gorelova, O.A., Reshetnikova, I.V., Solovchenko, A.E., Khozin-Goldberg, I., Cohen, Z., 2007. Effect of nitrogen starvation on optical properties, pigments, and arachidonic acid content of the unicellular green alga Parietochloris incisa (Trebouxiophyceae, Chlorophyta). J. Phycol. 43, 833–843. Pan, Y.Y., Wang, S.T., Chuang, L.T., Chang, Y.W., Chen, C.N., 2011. Isolation of thermotolerant and high lipid content green microalgae: oil accumulation is predominantly controlled by photosystem efficiency during stress treatments in Desmodesmus. Bioresour. Technol. 102, 10510–10517. Sobczuk, T., Chisti, Y., 2010. Potential fuel oils from the microalga Choricystis minor. J. Chem. Technol. Biotechnol. 85, 100–108. Su, C.H., Chien, L.J., Gomes, J., Lin, Y.S., Yu, Y.K., Liou, J.S., Syu, R.J., 2011. Factors affecting lipid accumulation by Nannochloropsis oculata in a two-stage cultivation process. J. Appl. Phycol. 23, 903–908. Tang, H., Chen, M., Simon Ng, K.Y., Salley, S.O., 2012. Continuous microalgae cultivation in a photobioreactor. Biotechnol. Bioeng. 109, 2468–2474. Wang, G., Wang, T., 2012. Characterization of lipid components in two microalgae for biofuel application. J. Am. Oil Chem. Soc. 89, 135–143. Wen, X.B., Jiang, L.L., Geng, Y.H., Shen, X., Li, Y.G., 2012. Comparative study of methods for microalgal total lipid quantification (in Chinese with English title and abstracts). China Oils Fats 37, 80–85. Wijffels, R.H., Barbosa, M.J., 2010. An outlook on microalgal biofuels. Science 329, 796–799. Zar, J.H., 1999. Biostatistical Analysis, fourth ed. Prentice-Hall, New Jersey. Zhang, B.Y., Geng, Y.H., Li, Z.K., Hu, H.J., Li, Y.G., 2009. Production of astaxanthin from Haematococcus in open pond by two-stage growth one-step process. Aquaculture 295, 275–281. Zhang, Y.M., Chen, H., He, C.L., Wang, Q., 2013. Nitrogen starvation induced oxidative stress in an oil-producing green alga Chlorella sorokiniana C3. PLoS ONE 8, e69225.
