Bioresource Technology xxx (2014) xxx–xxx
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Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition Tse-Shih Lin, Jane-Yii Wu ⇑ Department of Bioindustry Technology, Da-Yeh University, No. 168, University Rd., Dacun, Changhua 51591, Taiwan
h i g h l i g h t s Reports a novel microalgal isolate Chlorella sp. Y8-1. The high lipid content was obtained under mixotrophic condition. Fatty acid compositions of microalgae Y8-1 are appropriate for biodiesel production.
a r t i c l e
i n f o
Article history: Received 2 September 2014 Received in revised form 2 November 2014 Accepted 3 November 2014 Available online xxxx Keywords: Chlorella sp. Mixotrophic Lipid Fatty acid Biodiesel
a b s t r a c t In order to produce microalgal lipids that can be transformed to biodiesel fuel, one isolate with high lipid content was identified as Chlorella sp. Y8-1. The growth and lipid productivity of an isolated microalga Chlorella sp. Y8-1 were investigated under different cultivation conditions, including autotrophic growth (CO2, with light), heterotrophic growth (sucrose, without light) and mixotrophic growth (organic carbon sources and CO2, with light). Mixotrophic Chlorella sp. Y8-1 showed higher lipid content (35.5 ± 4.2%) and higher lipid productivity (0.01 g/L/d) than Chlorella sp. Y8-1 cultivated under autotrophic and heterotrophic conditions on modified Walne medium. Fatty acid analysis of Chlorella sp. Y8-1 showed the major presence of palmitic acid (C16:0), oleic acid (C18:1), linoleic acid (C18:2) and linolenic acids (C18:3). The main fatty acid compositions of the Chlorella sp. Y8-1 are appropriate for biodiesel production. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction The rapid development of human activity and the precipitous consumption of fossil fuels have caused an energy crisis, constituting a major problem in the twenty-first century. Vasudevan and Briggs (2008) predicted that the crude oil and natural gas reserves on earth will be depleted in 40 and 64 years, respectively (Vasudevan and Briggs, 2008). Moreover, the over-consumption of fossil fuels releases substantial amounts of greenhouse gases, inducing climate change and global warming. Therefore, sustainable, renewable, and carbon–neutral energy sources must be exploited to replace fossil fuels. Biodiesel is a nontoxic, renewable, and environmentally friendly energy source. Compared with conventional oil crops, microalgae are promoted as an ideal bioenergy feedstock because they exhibit a more rapid growth rate, demonstrate higher photosynthetic efficiency, can potentially produce considerably higher areal oil yields, ⇑ Corresponding author. Fax: +886 4 8511323. E-mail address: [email protected] (J.-Y. Wu).
do not compete with food or feed crops, and can be grown on barren land. Therefore, microalgae-based biodiesel has attracted increasing attention worldwide. However, the relatively high cost of microalgae-based biodiesel production is a major obstacle to the commercial application of this biotechnology. Increasing biomass concentration, lipid content per microalgal biomass, and lipid productivity are necessary for the economic feasibility of microalgae-based biodiesel production (Ho et al., 2012). Today, photoautotrophic culture is the most common strategy for cultivating microalgae. But the photoautotrophic culture exists severely limiting biomass production because of cellular self-shading that hinders light availability towards the end of growth. The low cell concentration gained in the photoautotrophic culture increases the biomass harvesting cost. A feasible alternative strategy to improve the efficient use of light or eliminate its requirement by cells and so reduce the costs of microalgal biomass production would be a mixotrophic culture of microalgae. Mixotrophic growth requires relatively low light intensities and, consequently, can reduce energy costs (Fernández Sevilla et al., 2004). The mixo-
http://dx.doi.org/10.1016/j.biortech.2014.11.005 0960-8524/Ó 2014 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx
trophic culture regime is a variant of the heterotrophic culture, where CO2 and organic carbons are simultaneously assimilated and both respiratory and photosynthetic metabolism operates concurrently (Lee, 2004). Many studies have been published in this research field. Previous studies have confirmed that the lipid content in some microalgae can be enhanced by applying various cultivation conditions such as nitrogen or phosphate starvation (Khozin-Goldberg and Cohen, 2006), and high light intensity (Cheirsilp and Torpee, 2012). Among these methods, using carbon and nitrogen sources are known to affect the metabolism of lipids and fatty acids in various microalgae. In addition, applying carbon and nitrogen conditions is simple and more cost-effective than using other methods. Therefore, the types and concentrations of carbon and nitrogen sources are critical in enhancing the lipid productivity of biodiesel production (Jeon et al., 2006). Some microalgal species, including the Chlorella (Zheng et al., 2012), Chlamydomonas (Chen and Johns, 1994), and Nannochloropsis species (Hu and Gao, 2003), have been reported to accumulate amounts of lipid in cells under various culture conditions. However, there are few studies on the effects of carbon resources, especially, the effects of carbon sources on the biomass production and lipid component of algae under mixotrophic cultivation (Andrade and Costa, 2007; Bouarab et al., 2004). Therefore, marine microalgae in seawater around Taiwan that exhibit high lipid productivity were isolated in this study to observe the effects of carbon and nitrogen sources on the growth and lipid productivity of the isolated microalgae under mixotrophic conditions.
2. Methods 2.1. Collection of samples, establishment and identification of algal strains The microalgae samples were collected from seawater around Taiwan, stored in sterile centrifugal tubes, and sent to the laboratory within 3 d for algal cell isolation. Walne medium plates were prepared using full-strength seawater, containing 18 g/L of agar and 1 g/L of glucose. After inoculation, the plates were cultured at 30 °C for 2–7 d. Single colonies composed of spherical cells atypical of either yeast, fungi, or bacteria were extracted and carefully transferred to a new plate. After becoming established, these algal strains were identified according to their 18S rRNA gene sequences, as well as some morphological characteristics. For morphological observation, cells from each strain were observed using a light microscope (ESPA, Taiwan). For obtaining the DNA sequences of the 18S rRNA gene from one strain, a single colony of the strain grown on an agar plate was carefully transferred to a 50-mL tube containing 1 g/L of glucose and 10 mL of a Walne liquid medium prepared using seawater. The culture was then cultivated at 30 °C for 1 wk with continuous aeration (10% CO2, 0.5 vvm). The algal cells were collected using centrifugation (5000 rpm 5 min), rinsed with 5 mL of deionized water, and lyophilized prior to performing DNA sequencing. The amplified 18S rRNA gene in the genomic DNA of algal cells was obtained and sent to Mission Biotech (Taipei, Taiwan) for DNA sequencing. The resulting 18S rRNA gene sequences were aligned and compared to the nucleotide sequences of known microorganisms in the GenBank database of the National Center for Biotechnology Information by using a Basic Local Alignment Search Tool (BLAST). The samples were also analyzed using MEGA 4.1 software (Tamura et al., 2007) and by employing the multiple alignment program CLUSTAL W to construct a neighbor-jointing (NJ) tree. The bootstrap values were obtained from 1000 replications of NJ analyses (Burja et al., 2006).
2.2. Serum bottle cultivation of isolated microalgae Y8-1 The microalga Y8-1 was cultivated in a 1 L serum bottle with a working volume of 0.8 L at 30 °C by using an exponentially growing seed culture, and 4300 lux of light intensity was adopted. Aeration was achieved by sparging air enriched with 10% CO2 at 2 vvm. During microalgal growth, the liquid sample was collected from the serum bottle with respect to time to determine microalgal biomass concentration, pH, residual sugar concentration and lipid content of the microalgal biomass. The microalga Y8-1 was grown in the serum bottle on modified Walne media. The modified Walne media was composed (per liter) of 30 g of malt salt, 2 mg of NaH2PO42H2O, 4.5 mg of Na2EDTA, 3.36 mg of H3BO3, 0.036 mg of MnCl24H2O, 0.13 mg of FeCl36H2O, and nitrogen sources (urea). At the same time, different carbon sources were also used for the growth of microalga Y8-1. Inorganic carbon (CO2) was used as the carbon source in photoautotrophic cultivation, while organic carbon (sucrose) was used in heterotrophic cultivation. Mixotrophic cultivation, which means the microalgae could undergo photosynthesis and simultaneously use both organic (fructose, glucose, glycerol, sucrose, and xylose) and inorganic carbon (CO2) as carbon sources, was also investigated in this study. The effects of these cultivation conditions on microalgae growth and lipid production were investigated. 2.3. Analytical method Biomass was determined by measuring the OD of each sample at 680 nm (OD680). The dry cell weights of the diluted samples were then detected and measured for plotting the standard curve. The amounts of total sugar were estimated by the phenol–sulfuric acid assay method of Dubois et al. (1956) using fructose, glucose, sucrose, and xylose standard calibration curves, respectively (Dubois et al., 1956). The glycerol concentration was determined using high-performance liquid chromatography (HPLC; Young Lin Acme 9000 HPLC). 2.4. Total lipid extraction The total lipid content (dry weight) was measured by employing a modified version of the method used by Bligh and Dyer (1959) (Bligh and Dyer, 1959). After the cultivation was complete, the culture medium was centrifuged at 9000 rpm and 4 °C for 2 min; the cell pellets were then collected for freeze drying. The samples were pulverized after drying by using a homogenizer, and were extracted using a chloroform–methanol mixture (1:2 v/ v). Approximately 15 mL of solvents was used for 50 mg of dried samples in each extraction step. After the samples were mixed using a vortex mixer for 1 min, they were ultrasonicated for 3 h and centrifuged at 3000 rpm for 10 min. The solid phases were separated carefully using Whatman No. 1 filter paper, and the solids were washed using 5 mL of chloroform. After this process, 9 mL of sterilized water was added to a solvent phase, and the solvent was mixed using a vortex mixer. The solvent phase was centrifuged at 3000 rpm for 10 min, and the chloroform layer was collected. The weight of the lipids was measured after removing the solvent by using a nitrogen blowing concentrator; the lipid content was then calculated. 2.5. Fatty methyl esters and fatty acid analysis To observe the saponification/esterification reactions, each of the samples were mixed with 2 mL of NaOH–methanol solution and disrupted using a sonicator, heated in a 100 °C water bath for 10 min, and cooled to room temperature. The samples were
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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Dictyosphaerium ehrenbergianum strain CCAP 222/10 (GQ176857.1) 92 55
Dictyosphaerium ehrenbergianum strain CCAP 222/57 (GQ176855.1) Dictyosphaerium ehrenbergianum strain CB 2008/107 (GQ487213.1)
Parachlorella hussii strain ACOI 939 (HM126551.1) 42 74 Parachlorella hussii strain ACOI 1508 (HM126548.1) 44 Dictyosphaerium ehrenbergianum strain CCAP 222/46 (GQ487202.1)
Dictyosphaerium libertatis strain CB 2008/76 (GQ487211.1) 7328
Dictyosphaerium ehrenbergianum strain CCAP 222/20 (GQ487192.1)
60 Dictyosphaerium ehrenbergianum strain CCAP 222/27 (GQ477062.1)
Chlorella sp. MDL5-18 (AY197632.1) 97
59 80 48
Dictyosphaerium sp. CCAP 222/25 (GQ176862.1) Parachlorella kessleri (FM205885.1) Dictyosphaerium sp. SAG 70.80 (GQ176860.1) Chlorellaceae sp. TP-2008 (FM205842.1) Chlorella sorokiniana (FM205860.1) 64 97
54
Micractinium sp. TP-2008b (FM205852.1) Chlorella sp. EN 2003/25 (HQ111430.1)
97
Coronastrum ellipsoideum strain UTEX LB1382 (GQ507370.1 Chlorella sp. CCAP 211/90 (FM205861.1)
63 Chlorella sp. IFRPD 1018 (AB260898.1)
46
Chlorella sp. IFRPD 1014 (AB260897.1) Chlorella sp. KMMCC C-185 (GQ122357.1) Chlorella vulgaris culture-collection KMMCC:C-204 (GQ122359.1) Chlorella vulgaris culture-collection KMMCC:C-27 (GQ122334.1) 27 55 Chlorella sp. KMMCC C-181 (GQ122354.1)
Chlorella vulgaris culture-collection KMMCC:C-111 (GQ122346.1) 46
Chlorella sp. KMMCC C-235 (GQ122360.1) Chlorella vulgaris culture-collection KMMCC:C-77 (GQ122338.1) Chlorella sp. KAS012 (AB176666.1) Chlorella sp. KMMCC C-239 (GQ122361.1) Chlorella sp. Y 8-1 (KJ855064)
0.01
Fig. 1. The phylogenetic tree of isolated microalgae.
then mixed with 2 mL of HCl in methanol and 1 mL of boron trifluoride solution (Sigma–Aldrich, St. Louis, MO, USA), heated again in a 90 °C water bath for 15 min, and then cooled again. Next, 3 mL of a saturated NaCl aqueous solution and 4 mL of n-hexane were added, and the samples were mixed thoroughly. The upper liquid layer was transferred to a 4-mL amber glass vial, sealed using ParafilmÒ, and stored at 20 °C until analysis was performed. The fatty methyl ester (FAME) samples were analyzed using a gas chromatography machine (YL6100 GC, Young Lin, Korea) equipped with a Carbowax column (fused silica, 30 m 0.25 mm, Quadrex, U.S.A.) and a flame ionization detector according to appropriate reference standards (Sigma, Louisiana, USA). The injector temperature was set at 220 °C, the column temperature was raised from 50 °C to 150 °C in increments of 15 °C/min, and then from 150 °C to 250 °C in increments of 5 °C/min. Methyl esters prepared from fatty acids, including C16:0, C16:1, C18:0, C18:1, C18:2, C18:3, and C20:0, as well as a FAME standard mixture, were all purchased from Sigma– Aldrich and used as the standards for identifying fatty acids in the samples. These fatty acids were quantified according to their peak area relative to the C17:0 fatty acid internal standard, and expressed as a percentage of total fatty acid content.
3. Results and discussion 3.1. Collection of samples, establishment and identification of algal strains One strain of algae-like microorganisms capable of mixotrophic growth was isolated from Tainan (located in Southern Taiwan). Using BLAST software to align and compare the 18S rRNA gene sequences with other known microorganisms in the GenBank database revealed that this strain exhibited a close phylogenic relationship with the Chlorella family (NCBI number KJ855064), which are microalgae well-known for their high lipid content (Fig. 1). According to the phylogenetic analysis, it is reasonable to identify Y8-1 as Chlorella sp. Y8-1. The intracellular lipid droplets of Chlorella sp. Y8-1 were observed using Sudan Black B and Nile Red staining under fluorescence microscopy; the lipids were stained in black and yellow, respectively (data not shown). 3.2. Effects of the carbon sources on the growth and lipid production of Chlorella sp. Y8-1 in mixotrophic culture In mixotrophic cultivation, Chlorella sp. Y8-1 exhibited rapid growth in response to the addition of carbon, except xylose
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx
pH
12.0 (a) Blank (without carbon & nitrogen source) 10.5
(b) Blank 1 (without carbon source)
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(f) sucrose: 1 g/L
(g) xylose: 1 g/L
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) Dried cell weight (g/L)
4.5 0.75
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Fig. 2. The time courses of dried cell weight and sugar consumption of isolated Chlorella sp. Y8-1 microalgae at various carbon sources. Culture condition: light/dark cycle: 24/ 0; light source: fluorescent; light intensity: 4300 lux; nitrogen sources: urea 0.5 g/L; various carbon sources; aeration: 10% CO2; n = 3. Note: (a) and (b): autotrophic cultivation; (c)–(g): mixotrophic cultivation.
0.6
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Bl a B l nk an fru k 1 ct o gl s e uc gl ose yc e su r o l cr o x y se lo se
(
) Dried cell weight (g/L)
(
) Lipid content (%)
(Fig. 2). After a short delay phase of 1 d, the cells underwent exponential growth for 4 d and then entered the stationary phase after 7 d. The cells that contained fructose, glucose, glycerol, and sucrose as the carbon sources grew much more rapidly and higher than did those that contained xylose. The dried biomass concentration increased as high as 0.45 g/L during the late-exponential phase. In addition, the mixotrophic cells cultured in the presence of 10% CO2 (2 vvm), 1 g/L of sucrose and 0.5 g/L of urea reached the stationary phase more rapidly than those cultured in the presence of other carbons did. However, approximately equal dried biomass concentrations were obtained from the samples grown on glucose and glycerol under mixotrophic cultivation (Fig. 2). In mixotrophic cultivation, four carbon sources induced cell growth and lipid accumulation during the period of culture. As shown in Fig. 3, the specific growth rates (l) were significantly higher in the presence of organic carbon sources (fructose, glucose,
Carbon sources (1 g/L) Fig. 3. Lipid content and growth of Chlorella sp. Y8-1 after 14 day culture at various carbon sources (1 g/L). Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; nitrogen sources: urea 0.5 g/L; aeration: 10% CO2; n = 3. Note: Blank: without organic carbon and nitrogen sources (autotrophic cultivation). Blank 1: without organic carbon sources (autotrophic cultivation). Others: mixotrophic cultivation.
glycerol and sucrose) during 7 days. Additionally, the maximal specific growth rate (lm) (0.39 ± 0.04 d 1) and lipid content (24.0 ± 0.61% of dry cell weight) were obtained when sucrose was used as the organic carbon source. However, fructose presumably cannot be used to enhance the lipid content of Chlorella sp. Y8-1 under mixotrophic cultivation (Fig. 3). Adding a carbon source to the medium has been widely reported to increase the growth and lipid content of microalgae (Qlao and Wang, 2009). Chlorella species can grow on various carbon substrates such as sodium acetate (Qlao and Wang, 2009), fructose (Gao et al., 2009), glucose (Yeh and Chang, 2012), glycerol (Heredia-Arroyo et al., 2010), sucrose (Gao et al., 2009), and acetate (Heredia-Arroyo et al., 2010). Fructose, glucose, glycerol, and sucrose were transformed into glyceraldehydes-3-phosphate, which is a crucial intermediate product involved in both glycolysis and the pentose phosphate pathway (Lewin, 1974). However, xylose apparently cannot be used to grow microalgae effectively. Therefore, the growth was considerably slower when xylose was used as the organic carbon source, compared with the growth when the other four organic carbon sources were used. Sucrose can be decomposed to yield a glucose molecule and a fructose molecule, that is why microalgae can utilize sucrose to growth. Therefore, it is possible that Chlorella sp. possesses an inducible transport system for hexoses (Haass and Tanner, 1974), but not for pentose. Fig. 3 indicates that lipid accumulation occurred in mixotrophic cells (simultaneously using both 1 g/L glucose and 10% CO2, glycerol and 10% CO2, sucrose and 10% CO2 as carbon sources, respectively), and the lipid content of the autotrophic cells (Blank 1, using 10% CO2 as carbon sources) was only 6.0 ± 2.5%. It would be reasonable to suggest that TCA cycle is the starting point for many anabolisms, such as lipid, protein and so on. Therefore, this result explains the metabolic basis that lipid content was significantly lower in autotrophy culture than in the heterotrophy and mixtrophy culture. The lipid content of the mixotrophic cells observed in the current study was 9.4 ± 2.6–24.0 ± 3.2% (Fig. 3). It is possible that some carbon sources can lead to enhancement of acetyl CoA/malonyl CoA pool – which represents the central carbon donor for fatty acid synthesis, thereby increasing lipid pool (Ngangkham et al., 2012). Miao and Wu (2006) cultivated Chlorella protothecoides by using glucose and demonstrated that the lipid content was fourfold in response to heterotrophic growth when compared with autotrophic growth (Miao and Wu, 2006). In addition, the present study also found that the cell concentration (>0.40 g/L) of the Chlorella sp. Y8-1 from mixotrophic (using 1 g/L sucrose and 10% CO2 as carbon source) growth was higher than the sum of those from autotrophic (0.15 g/L) (using 10% CO2 as carbon source) and heterotrophic (using 1 g/L sucrose as carbon source) growth (0.17 g/L, data not shown). This indicated that the mixotrophic growth is not a simple combination of photoautotrophic and heterotrophic growth. Although it has been reported that
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx Table 1 Lipid and biomass production of different microalgae under photoautotrophic, heterotrophic and mixotrophic culture conditions, respectively. Microalgae
Autotrophic
Heterotrophic
Mixotrophic
Refs.
Lipid content (%)
Biomass (g/L)
Lipid content (%)
Biomass (g/L)
Lipid content (%)
Biomass (g/L)
Marine Chlorella sp. Y8-1 Chlorella sp. Chlorella vulgaris Chlorella sorokiniana Nannochloropsis sp.
16.5 30.0 – 19.0 28.0
0.22 0.38 – – 0.37
5.9 21.0 2.0 – 20.0
0.17 0.48 3.00 – 0.38
35.5 25.5 0.8 33.0 27.5
0.45 1.45 4.00 – 1.20
This study Cheirsilp and Torpee (2012) EL-Sheekh et al. (2012) Ngangkham et al. (2012) Cheirsilp and Torpee (2012)
Freshwater Chlorella sp. Chlorella vulgaris ESP-31
13.5 20.0
0.60 0.8
13.0 16.0
0.75 0.2
15.0 53.0
1.40 3.0
Cheirsilp and Torpee (2012) Yeh and Chang (2012)
Note: Values were not showed from the results of studies and labeled as ‘‘–‘‘ in the table.
0.8
20
0.6
15
0.4
10
0.2
5 (b) After 15 day
30
0.0 (b) After 15 day 1.0
25
0.8
20
0.6
15
0.4
10
0.2
5 0
0
0.1 0.25 0.5 0.75 1 1.5
2
Urea concentration (g/L)
0 0.1 0.25 0.5 0.75 1 1.5 2
0.0
0.5
0.4 0.3 0.2 0.1 0.0
0.4
-1
0.5
) µ during 7 days (day )
25
0
Lipid content (%)
1.0
(
(a) After 7 day
-1
(a) After 7 day
) µm (day )
Lipid content (%)
30
0.3 0.2 0.1
(
Our earlier studies revealed that urea was a suitable nitrogen source for cultivation of Chlorella sp. Y8-1. Urea was adopted into the medium at 0–2 g/L. The biomass, specific growth rates, and lipid content of Chlorella sp. Y8-1 in the stationary phase are shown in Fig. 4. The cultures had a low biomass concentration, specific growth rate, and lipid content in the N-free medium. The Chlorella sp. Y8-1 grew the most robustly in the medium containing 0.1– 0.25 g/L of urea, reaching a biomass concentration of
( ) Dried cell weight (g/L)
3.3. Effects of urea concentration on the growth and lipid production of Chlorella sp. Y8-1
0.68 ± 0.06 g/L; however, the growth was limited when the urea concentration was higher than 0.5 g/L (data not shown). Within the urea range from 0.5 to 2.0 g/L, the biomass concentration in the urea experiment decreased from 0.6 ± 0.05 to 0.5 ± 0.04 g/L. In the medium with 0.1–2.0 g/L of urea, the specific growth rate during 7 days and maximal specific growth rate slightly decreased as the urea concentration increased (Fig. 4). In the experimental range, the lipid content of Chlorella sp. Y8-1 increased as the culture period increased. In addition, the lipid content increased from 6.7 ± 3.9% to 31.5 ± 3.18% of dry cell weight as the urea was increased from 0 to 0.25 g/L; however, the lipid production was limited when the urea concentration was higher than 0.5 g/L. The maximal lipid content (31.5 ± 3.18% of dry cell weight) was obtained in the medium containing 0.25 g/L of urea (Fig. 4). In the current experiments, the higher the urea concentration was, the higher the lipid content of the Chlorella sp. Y8-1 when the urea ranged from 0 to 0.25 g/L. The results corresponded with those of several reports; for example, the lipid content of Ellipsoidion sp. increased as the nitrate concentration increased, and the maximal dry weight of 32.8% was obtained when the nitrate-N used was 1.92 mmol/L (Xu et al., 2001). Furthermore, Isochrysis galbana decreased markedly from 30% (NaNO3 > 2 mmol/L) to 19% of dry weight at 0.5 mmol/L NaNO3 (Molina and Sánchez,
( ) Dried cell weight (g/L)
photosynthesis and heterotrophic growth occur simultaneously and independently in mixotrophic cultures (Marquez et al., 1993), the presence of organic carbon can alter both the photosynthetic and heterotrophic metabolism of Chlorella (Villarejo et al., 1995). The lipid and biomass production of mixotrophic cultivation are much higher than under autotrophic and heterotrophic cultivation indicated in previous work (Table 1). Thus, these findings indicate that adding organic carbon sources leads to a marked stimulation of lipid accumulation in Chlorella sp. Y8-1, and that the ideal organic carbon source for the production of lipids is sucrose.
0.0
Urea concentration (g/L)
Fig. 4. Lipid content and growth of Chlorella sp. Y8-1 at various urea concentrations under mixotrophic cultivation. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; organic carbon sources: sucrose 1 g/L; aeration: 10% CO2; n = 3.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx
1992). However, for most microalgae, the lipid content increased under nitrogen deficient conditions. In other studies, Nannochloropsis oculata had maximal lipid content per cell under low nitrogen conditions (Converti et al., 2009), and the lipid content of Chlamydomonas reinhardtii CC124 and Chlorella vulgaris increased under nitrogen-deficient conditions (Deng et al., 2011). In general, this behavior is most probably a survival response until restoration of less nutritionally stressing conditions.
and Johns, 1995). Chen (1996) and Chen and Johns (1994) have both indicated that, compared with bacteria and yeasts, microalgae can tolerate only relatively low substrate concentrations (Chen, 1996; Chen and Johns, 1994). In addition, at the initial sucrose concentration of 0.5–1.0 g/L, the sucrose level gradually decreased and was nearly completely depleted at the end of cultivation. When a higher initial sucrose concentration of 5–10 g/L was used, approximately 3.0 g/L of sucrose remained at the end of cultivation. In our experiments, high concentrations of sucrose have been shown to inhibit microalgal growth, at least for a considerable period of time. It was likely that algal cells grown on high concentration of substrates require a lag period to adopt the bioreactor operation (Perez-Garcia et al., 2011). The effects of sucrose concentration on specific growth rate and lipid production are summarized in Fig. 6. As shown in Fig. 6, the specific growth rate during 7 days and maximal specific growth rate were significantly higher in the presence of sucrose. However, the specific growth rate during 7 days and maximal specific growth rate were decreased when the sucrose concentration was 10 g/L. It was also likely that algal cells grown on high concentration of substrates
3.4. Effects of sucrose concentration on the growth and lipid production of Chlorella sp. Y8-1 The effects of carbon source concentrations on the growth and lipid production of isolated Chlorella sp. Y8-1 are shown in Figs. 5 and 6. The growth rate of isolated Chlorella sp. Y8-1 increased when the initial sucrose concentration was increased from 0 to 0.75 g/L (Fig. 5). Increasing the initial sucrose concentration further did not enhance the growth of this strain. The growth rate slightly decreased when the initial sucrose concentration was increased to 1 g/L. This could have been an effect of substrate inhibition (Chen
pH
12.0 (a) Sucrose conc.: 0 g/L 10.5
(b) Sucrose conc.: 0.5 g/L
(c) Sucrose conc.: 0.75 g/L
(e) Sucrose conc.: 2.5 g/L
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Fig. 5. The time courses of dried cell weight and sugar consumption of isolated Chlorella sp. Y8-1 microalgae at various sucrose concentrations. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; urea concentration: 0.25 g/L; various sucrose concentration; aeration: 10% CO2; n = 3. Note: (a): autotrophic cultivation. (b)–(h): mixotrophic cultivation.
0.3 0.2 0.1
0 0.5 0.75 1 2.5 5 7.5 10 Sucrose concentration (g/L)
Fig. 6. Lipid content and growth of Chlorella sp. Y8-1 at various sucrose concentrations. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; urea concentration: 0.25 g/L; various sucrose concentration; aeration: 10% CO2; n = 3. Note: 0 g/L sucrose: autotrophic cultivation. 0.5–10 g/L sucrose: mixotrophic cultivation.
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require a lag period to adopt (Perez-Garcia et al., 2011). The lipid production of Chlorella sp. Y8-1 increased from 16.5 ± 3.6% to 35.5 ± 4.2% when the initial sucrose concentration increased from 0 to 0.5 g/L (Fig. 6). It was likely that in limited N condition, protein, nucleic acid synthesis slow down so cell division and growth are restricted, most assimilated carbon are led to produce storage and stress-resistant materials such as lipid, starch. When the initial sucrose concentration was higher than 0.75 g/L, the lipid production of Chlorella sp. Y8-1 gradually decreased. Cheirsilp and Torpee (2012) also discovered that the lipid content of Nannochloropsis sp. increased from 109.8 to 798.1 mg/L when the initial glucose concentration was increased from 0 to 15 g/L, but the lipid content slightly decreased when the initial glucose concentration was 20 g/L (Cheirsilp and Torpee, 2012). Consequently, these findings indicate that the ideal sucrose concentration for lipid production of Chlorella sp. Y8-1 is 0.5 g/L. 3.5. Fatty acid composition of Chlorella sp. Y8-1 in mixotrophic culture The primary fatty acids in Chlorella sp. Y8-1, as percentages of total fatty acids, were 16:0 (19.2 ± 0.6%), 18:1 (11.2 ± 0.4%), 18:2 (18.0 ± 0.6%), and 18:3 (10.5 ± 0.3%) (data not shown), indicating good agreement with the results observed in the lipid production of C. vulgaris (Yusof et al., 2011). The major saturated fatty acids (SFA) in Chlorella sp. Y8-1 were palmitic acid (16:0) and stearic acid (C18:0). The major monounsaturated fatty acid (MUFA) was oleic acid (18:1). It indicated that Chlorella sp. Y8-1 accumulated a high proportion of SFA and MUFA to 50% of the total lipid content. Compared to the commonly used soybean oil as feedstocks for biodiesel production in the US, the biodiesel derived from microalgae lipid in this study would be more saturated and provide a higher cetane number (CN), lower NOx emissions, shorter ignition delay time, and higher oxidative stability. Numerous fatty acids could not be identified in this study, because unsaturated fatty acids largely prevail in lipids. 4. Conclusions Compared to photoautotrophic and heterotrophic cultures, an abundance of lipids were accumulated when Chlorella sp. Y8-1 was cultivated under mixotrophic conditions, suggesting that it has great potential for renewable biodiesel feedstock applications. Mixotrophic Chlorella sp. Y8-1 showed higher lipid content (35.5 ± 4.2%) and higher lipid productivity (0.01 g/L/d) than Chlorella sp. Y8-1 cultivated under autotrophic and heterotrophic conditions. Additionally, under mixotrophic conditions, the microalgae accumulates lipids up to 35.5 ± 4.2% of dry cell weight, with palmitic acid (16:0) and stearic acid (C18:0) representing the most abundant fatty acid components. Mixotrophic cultivation is much easier to alter conditions to improve the yield of biomass and lipid of microalgal production. Therefore, further study on the influence of environmental factors would facilitate the improvement of algal lipid production. Acknowledgement This work was financially supported in part by a grant from the National Science Council of Republic of China under a contract number of NSC 100-2632-B-212-001-MY3. References Andrade, M.R., Costa, J.A.V., 2007. Mixotrophic cultivation of microalga Spirulina platensis using molasses as organic substrate. Aquaculture 264, 130–134. Bligh, E.G., Dyer, W.J., 1959. A rapid method for total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917.
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Bouarab, L., Dauta, A., Loudiki, M., 2004. Heterotrophic and mixotrophic growth of Micractinium pusillum Fresenius in the presence of acetate and glucose: effect of light and acetate gradient concentration. Water Res. 38, 2706–2712. Burja, A.M., Radianingtyas, H., Windust, A., Barrow, C.J., 2006. Isolation and characterization of polyunsaturated fatty acid producing Thraustochytrium species: screening of strains and optimization of omega-3 production. Appl. Microbiol. Biotechnol. 72, 1161–1169. Cheirsilp, B., Torpee, S., 2012. Enhanced growth and lipid production of microalgae under mixotrophic culture condition: effect of light intensity, glucose concentration and fed-batch cultivation. Bioresour. Technol. 110, 510–516. Chen, F., 1996. High cell density culture of microalgae in heterotrophic growth. Trends Biotechnol. 14, 421–426. Chen, F., Johns, M.R., 1994. Substrate inhibition of Chlamydomonas reinhardtii by acetate in heterotrophic culture. Proc. Biochem. 29, 245–252. Chen, F., Johns, M.R., 1995. A strategy for high cell density culture of heterotrophic microalgae with inhibitory substrates. J. Appl. Phycol. 7, 43–46. Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., Borghi, M.D., 2009. Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem. Eng. Process. 48, 1146–1151. Deng, X., Fei, X., Li, Y., 2011. The effects of nutritional restriction on neutral lipid accumulation in Chlamydomonas and Chlorella. Afr. J. Microbiol. Res. 5, 260–270. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. EL-Sheekh, M.M., Bedaiwy, M.Y., Osman, M.E., Ismail, M.M., 2012. Mixotrophic and heterotrophic growth of some microalgae using extract of fungal-treated wheat bran. Int. J. Recycling Org. Waste Agric. 1, 12–20. Fernández Sevilla, J.M., Cerón García, M.C., Sánchez Mirón, A., Belarbi, E.H., García Camacho, F., Molina Grima, E., 2004. Pilot plant-scale outdoor mixotrophic cultures of Phaeodactylum tricornutum using glycerol in vertical bubble column and airlift photobioreactors: studies in fed-batch mode. Biotechnol. Prog. 20, 728–736. Gao, C., Zhai, Y., Ding, Y., Wu, Q., 2009. Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides. Appl. Energy 87, 756–761. Haass, D., Tanner, W., 1974. Regulation of hexose transport in Chlorella vulgaris. Plant Physiol. 53, 14–20. Heredia-Arroyo, T., Wei, W., Hu, B., 2010. Oil accumulation via heterotrophic/ mixotrophic Chlorella protothecoides. Appl. Biochem. Biotechnol. 162, 1978– 1995. Ho, S.H., Chen, C.Y., Chang, J.S., 2012. Effect of light intensity and nitrogen starvation on CO2 fixation and lipid/carbohydrate production of an indigenous microalga Scenedesmus obliquus CNW-N. Bioresour. Technol. 113, 244–252. Hu, H., Gao, K., 2003. Optimization of growth and fatty acid composition of a unicellular marine picoplankton, Nannochloropsis sp., with enriched carbon sources. Biotechnol. Lett. 25, 421–425. Jeon, Y.C., Cho, C.W., Yun, Y.S., 2006. Combined effects of light intensity and acetate concentration on the growth of unicellular microalga Haematococcus pluvialis. Enzyme Microb. Technol. 39, 490–495. Khozin-Goldberg, I., Cohen, Z., 2006. Effect of phosphate starvation on lipid and fatty acid composition of the Eustigmatophyte Monodus subterraneus. Phytochemistry 67, 696–701. Lee, Y.K., 2004. Algal nutrition: heterotrophic carbon nutrition. In: Richmond, A. (Ed.), Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell, Oxford, pp. 116–124. Lewin, R.A., 1974. Biochemical taxonomy. In: Stewart, W.D.P. (Ed.), Algal Physiology and Biochemistry. Blackwell Scientific Publications, Oxford, UK, pp. 1–25. Marquez, F.J., Sasaki, K., Kakizono, T., Nishio, N., Nagai, S., 1993. Growth characteristics of Spirulina platensis in mixotrophic and heterotrophic conditions. J. Ferment. Bioeng. 76, 408–410. Miao, X., Wu, Q., 2006. Biodiesel production from heterotrophic microalgal oil. Bioresour. Technol. 97, 841–846. Molina, G.E., Sánchez, P.J.A., 1992. EPA from Isochrysis galbana growth conditions and productivity. Process Biochem. 27, 299–305. Ngangkham, M., Ratha, S.K., Prasanna, R., Saxena, A.K., Dhar, D.W., Sarika, C., Prasad, R.B.N., 2012. Biochemical modulation of growth, lipid quality and productivity in mixotrophic cultures of Chlorella sorokiniana. Springerplus 1, 1–13. Perez-Garcia, O., Escalante, F.M.E., de-Bashan, L.E., Bashan, Y., 2011. Heterotrophic cultures of microalgae: metabolism and potential products. Water Res. 45, 11– 36. Qlao, H., Wang, G., 2009. Effect of carbon source on growth and lipid accumulation in Chlorella sorokiniana GXNN01. Chin. J. Oceanol. 27, 762–768. Tamura, K., Dudley, J., Nei, M., Kumar, S., 2007. MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596–1599. Vasudevan, P.T., Briggs, M., 2008. Biodiesel production current state of the art and challenges. J. Ind. Microbiol. Biotechnol. 35, 421–430. Villarejo, A., Orús, M.I., Martínez, F., 1995. Coordination of photosynthetic and respiratory metabolism in Chlorella vulgaris UAM 101 in the light. Physiol. Plant. 94, 680–686. Xu, N., Zhang, X., Fan, X., Han, L., Zeng, C., Tseng, C.K., 2001. Effects of nitrogen source and concentration on growth rate and fatty acid composition of Ellipsoidion sp. (Eustigmatophyta). J. Appl. Phys. 13, 463–469. Yeh, K.L., Chang, J.S., 2012. Effects of cultivation conditions and media composition on cell growth and lipid productivity of indigenous microalga Chlorella vulgaris ESP-31. Bioresour. Technol. 105, 120–127.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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Yusof, Y.A.M., Basari, J.H., Mukti, N.A., Sabuddin, R., Muda, A.R., Sulaiman, S., Makpol, S., Ngah, W.Z.W., 2011. Fatty acids composition of microalgae Chlorella vulgaris can be modulated by varying carbon dioxide concentration in outdoor culture. Afr. J. Biotechnol. 10, 13536–13542.
Zheng, L.S.T., Oh, J.Y., Jeon, B.H., Moon, H.S., Kwon, S.U., Lim, B.K., Kang, C.W., 2012. The dietary effects of fermented Chlorella vulgaris (CBT) on production performance, liver lipids and intestinal microflora in laying hens. AsianAustralas. J. Anim. Sci. 25, 261–266.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition Tse-Shih Lin, Jane-Yii Wu ⇑ Department of Bioindustry Technology, Da-Yeh University, No. 168, University Rd., Dacun, Changhua 51591, Taiwan
h i g h l i g h t s Reports a novel microalgal isolate Chlorella sp. Y8-1. The high lipid content was obtained under mixotrophic condition. Fatty acid compositions of microalgae Y8-1 are appropriate for biodiesel production.
a r t i c l e
i n f o
Article history: Received 2 September 2014 Received in revised form 2 November 2014 Accepted 3 November 2014 Available online xxxx Keywords: Chlorella sp. Mixotrophic Lipid Fatty acid Biodiesel
a b s t r a c t In order to produce microalgal lipids that can be transformed to biodiesel fuel, one isolate with high lipid content was identified as Chlorella sp. Y8-1. The growth and lipid productivity of an isolated microalga Chlorella sp. Y8-1 were investigated under different cultivation conditions, including autotrophic growth (CO2, with light), heterotrophic growth (sucrose, without light) and mixotrophic growth (organic carbon sources and CO2, with light). Mixotrophic Chlorella sp. Y8-1 showed higher lipid content (35.5 ± 4.2%) and higher lipid productivity (0.01 g/L/d) than Chlorella sp. Y8-1 cultivated under autotrophic and heterotrophic conditions on modified Walne medium. Fatty acid analysis of Chlorella sp. Y8-1 showed the major presence of palmitic acid (C16:0), oleic acid (C18:1), linoleic acid (C18:2) and linolenic acids (C18:3). The main fatty acid compositions of the Chlorella sp. Y8-1 are appropriate for biodiesel production. Ó 2014 Elsevier Ltd. All rights reserved.
1. Introduction The rapid development of human activity and the precipitous consumption of fossil fuels have caused an energy crisis, constituting a major problem in the twenty-first century. Vasudevan and Briggs (2008) predicted that the crude oil and natural gas reserves on earth will be depleted in 40 and 64 years, respectively (Vasudevan and Briggs, 2008). Moreover, the over-consumption of fossil fuels releases substantial amounts of greenhouse gases, inducing climate change and global warming. Therefore, sustainable, renewable, and carbon–neutral energy sources must be exploited to replace fossil fuels. Biodiesel is a nontoxic, renewable, and environmentally friendly energy source. Compared with conventional oil crops, microalgae are promoted as an ideal bioenergy feedstock because they exhibit a more rapid growth rate, demonstrate higher photosynthetic efficiency, can potentially produce considerably higher areal oil yields, ⇑ Corresponding author. Fax: +886 4 8511323. E-mail address: [email protected] (J.-Y. Wu).
do not compete with food or feed crops, and can be grown on barren land. Therefore, microalgae-based biodiesel has attracted increasing attention worldwide. However, the relatively high cost of microalgae-based biodiesel production is a major obstacle to the commercial application of this biotechnology. Increasing biomass concentration, lipid content per microalgal biomass, and lipid productivity are necessary for the economic feasibility of microalgae-based biodiesel production (Ho et al., 2012). Today, photoautotrophic culture is the most common strategy for cultivating microalgae. But the photoautotrophic culture exists severely limiting biomass production because of cellular self-shading that hinders light availability towards the end of growth. The low cell concentration gained in the photoautotrophic culture increases the biomass harvesting cost. A feasible alternative strategy to improve the efficient use of light or eliminate its requirement by cells and so reduce the costs of microalgal biomass production would be a mixotrophic culture of microalgae. Mixotrophic growth requires relatively low light intensities and, consequently, can reduce energy costs (Fernández Sevilla et al., 2004). The mixo-
http://dx.doi.org/10.1016/j.biortech.2014.11.005 0960-8524/Ó 2014 Elsevier Ltd. All rights reserved.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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trophic culture regime is a variant of the heterotrophic culture, where CO2 and organic carbons are simultaneously assimilated and both respiratory and photosynthetic metabolism operates concurrently (Lee, 2004). Many studies have been published in this research field. Previous studies have confirmed that the lipid content in some microalgae can be enhanced by applying various cultivation conditions such as nitrogen or phosphate starvation (Khozin-Goldberg and Cohen, 2006), and high light intensity (Cheirsilp and Torpee, 2012). Among these methods, using carbon and nitrogen sources are known to affect the metabolism of lipids and fatty acids in various microalgae. In addition, applying carbon and nitrogen conditions is simple and more cost-effective than using other methods. Therefore, the types and concentrations of carbon and nitrogen sources are critical in enhancing the lipid productivity of biodiesel production (Jeon et al., 2006). Some microalgal species, including the Chlorella (Zheng et al., 2012), Chlamydomonas (Chen and Johns, 1994), and Nannochloropsis species (Hu and Gao, 2003), have been reported to accumulate amounts of lipid in cells under various culture conditions. However, there are few studies on the effects of carbon resources, especially, the effects of carbon sources on the biomass production and lipid component of algae under mixotrophic cultivation (Andrade and Costa, 2007; Bouarab et al., 2004). Therefore, marine microalgae in seawater around Taiwan that exhibit high lipid productivity were isolated in this study to observe the effects of carbon and nitrogen sources on the growth and lipid productivity of the isolated microalgae under mixotrophic conditions.
2. Methods 2.1. Collection of samples, establishment and identification of algal strains The microalgae samples were collected from seawater around Taiwan, stored in sterile centrifugal tubes, and sent to the laboratory within 3 d for algal cell isolation. Walne medium plates were prepared using full-strength seawater, containing 18 g/L of agar and 1 g/L of glucose. After inoculation, the plates were cultured at 30 °C for 2–7 d. Single colonies composed of spherical cells atypical of either yeast, fungi, or bacteria were extracted and carefully transferred to a new plate. After becoming established, these algal strains were identified according to their 18S rRNA gene sequences, as well as some morphological characteristics. For morphological observation, cells from each strain were observed using a light microscope (ESPA, Taiwan). For obtaining the DNA sequences of the 18S rRNA gene from one strain, a single colony of the strain grown on an agar plate was carefully transferred to a 50-mL tube containing 1 g/L of glucose and 10 mL of a Walne liquid medium prepared using seawater. The culture was then cultivated at 30 °C for 1 wk with continuous aeration (10% CO2, 0.5 vvm). The algal cells were collected using centrifugation (5000 rpm 5 min), rinsed with 5 mL of deionized water, and lyophilized prior to performing DNA sequencing. The amplified 18S rRNA gene in the genomic DNA of algal cells was obtained and sent to Mission Biotech (Taipei, Taiwan) for DNA sequencing. The resulting 18S rRNA gene sequences were aligned and compared to the nucleotide sequences of known microorganisms in the GenBank database of the National Center for Biotechnology Information by using a Basic Local Alignment Search Tool (BLAST). The samples were also analyzed using MEGA 4.1 software (Tamura et al., 2007) and by employing the multiple alignment program CLUSTAL W to construct a neighbor-jointing (NJ) tree. The bootstrap values were obtained from 1000 replications of NJ analyses (Burja et al., 2006).
2.2. Serum bottle cultivation of isolated microalgae Y8-1 The microalga Y8-1 was cultivated in a 1 L serum bottle with a working volume of 0.8 L at 30 °C by using an exponentially growing seed culture, and 4300 lux of light intensity was adopted. Aeration was achieved by sparging air enriched with 10% CO2 at 2 vvm. During microalgal growth, the liquid sample was collected from the serum bottle with respect to time to determine microalgal biomass concentration, pH, residual sugar concentration and lipid content of the microalgal biomass. The microalga Y8-1 was grown in the serum bottle on modified Walne media. The modified Walne media was composed (per liter) of 30 g of malt salt, 2 mg of NaH2PO42H2O, 4.5 mg of Na2EDTA, 3.36 mg of H3BO3, 0.036 mg of MnCl24H2O, 0.13 mg of FeCl36H2O, and nitrogen sources (urea). At the same time, different carbon sources were also used for the growth of microalga Y8-1. Inorganic carbon (CO2) was used as the carbon source in photoautotrophic cultivation, while organic carbon (sucrose) was used in heterotrophic cultivation. Mixotrophic cultivation, which means the microalgae could undergo photosynthesis and simultaneously use both organic (fructose, glucose, glycerol, sucrose, and xylose) and inorganic carbon (CO2) as carbon sources, was also investigated in this study. The effects of these cultivation conditions on microalgae growth and lipid production were investigated. 2.3. Analytical method Biomass was determined by measuring the OD of each sample at 680 nm (OD680). The dry cell weights of the diluted samples were then detected and measured for plotting the standard curve. The amounts of total sugar were estimated by the phenol–sulfuric acid assay method of Dubois et al. (1956) using fructose, glucose, sucrose, and xylose standard calibration curves, respectively (Dubois et al., 1956). The glycerol concentration was determined using high-performance liquid chromatography (HPLC; Young Lin Acme 9000 HPLC). 2.4. Total lipid extraction The total lipid content (dry weight) was measured by employing a modified version of the method used by Bligh and Dyer (1959) (Bligh and Dyer, 1959). After the cultivation was complete, the culture medium was centrifuged at 9000 rpm and 4 °C for 2 min; the cell pellets were then collected for freeze drying. The samples were pulverized after drying by using a homogenizer, and were extracted using a chloroform–methanol mixture (1:2 v/ v). Approximately 15 mL of solvents was used for 50 mg of dried samples in each extraction step. After the samples were mixed using a vortex mixer for 1 min, they were ultrasonicated for 3 h and centrifuged at 3000 rpm for 10 min. The solid phases were separated carefully using Whatman No. 1 filter paper, and the solids were washed using 5 mL of chloroform. After this process, 9 mL of sterilized water was added to a solvent phase, and the solvent was mixed using a vortex mixer. The solvent phase was centrifuged at 3000 rpm for 10 min, and the chloroform layer was collected. The weight of the lipids was measured after removing the solvent by using a nitrogen blowing concentrator; the lipid content was then calculated. 2.5. Fatty methyl esters and fatty acid analysis To observe the saponification/esterification reactions, each of the samples were mixed with 2 mL of NaOH–methanol solution and disrupted using a sonicator, heated in a 100 °C water bath for 10 min, and cooled to room temperature. The samples were
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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Dictyosphaerium ehrenbergianum strain CCAP 222/10 (GQ176857.1) 92 55
Dictyosphaerium ehrenbergianum strain CCAP 222/57 (GQ176855.1) Dictyosphaerium ehrenbergianum strain CB 2008/107 (GQ487213.1)
Parachlorella hussii strain ACOI 939 (HM126551.1) 42 74 Parachlorella hussii strain ACOI 1508 (HM126548.1) 44 Dictyosphaerium ehrenbergianum strain CCAP 222/46 (GQ487202.1)
Dictyosphaerium libertatis strain CB 2008/76 (GQ487211.1) 7328
Dictyosphaerium ehrenbergianum strain CCAP 222/20 (GQ487192.1)
60 Dictyosphaerium ehrenbergianum strain CCAP 222/27 (GQ477062.1)
Chlorella sp. MDL5-18 (AY197632.1) 97
59 80 48
Dictyosphaerium sp. CCAP 222/25 (GQ176862.1) Parachlorella kessleri (FM205885.1) Dictyosphaerium sp. SAG 70.80 (GQ176860.1) Chlorellaceae sp. TP-2008 (FM205842.1) Chlorella sorokiniana (FM205860.1) 64 97
54
Micractinium sp. TP-2008b (FM205852.1) Chlorella sp. EN 2003/25 (HQ111430.1)
97
Coronastrum ellipsoideum strain UTEX LB1382 (GQ507370.1 Chlorella sp. CCAP 211/90 (FM205861.1)
63 Chlorella sp. IFRPD 1018 (AB260898.1)
46
Chlorella sp. IFRPD 1014 (AB260897.1) Chlorella sp. KMMCC C-185 (GQ122357.1) Chlorella vulgaris culture-collection KMMCC:C-204 (GQ122359.1) Chlorella vulgaris culture-collection KMMCC:C-27 (GQ122334.1) 27 55 Chlorella sp. KMMCC C-181 (GQ122354.1)
Chlorella vulgaris culture-collection KMMCC:C-111 (GQ122346.1) 46
Chlorella sp. KMMCC C-235 (GQ122360.1) Chlorella vulgaris culture-collection KMMCC:C-77 (GQ122338.1) Chlorella sp. KAS012 (AB176666.1) Chlorella sp. KMMCC C-239 (GQ122361.1) Chlorella sp. Y 8-1 (KJ855064)
0.01
Fig. 1. The phylogenetic tree of isolated microalgae.
then mixed with 2 mL of HCl in methanol and 1 mL of boron trifluoride solution (Sigma–Aldrich, St. Louis, MO, USA), heated again in a 90 °C water bath for 15 min, and then cooled again. Next, 3 mL of a saturated NaCl aqueous solution and 4 mL of n-hexane were added, and the samples were mixed thoroughly. The upper liquid layer was transferred to a 4-mL amber glass vial, sealed using ParafilmÒ, and stored at 20 °C until analysis was performed. The fatty methyl ester (FAME) samples were analyzed using a gas chromatography machine (YL6100 GC, Young Lin, Korea) equipped with a Carbowax column (fused silica, 30 m 0.25 mm, Quadrex, U.S.A.) and a flame ionization detector according to appropriate reference standards (Sigma, Louisiana, USA). The injector temperature was set at 220 °C, the column temperature was raised from 50 °C to 150 °C in increments of 15 °C/min, and then from 150 °C to 250 °C in increments of 5 °C/min. Methyl esters prepared from fatty acids, including C16:0, C16:1, C18:0, C18:1, C18:2, C18:3, and C20:0, as well as a FAME standard mixture, were all purchased from Sigma– Aldrich and used as the standards for identifying fatty acids in the samples. These fatty acids were quantified according to their peak area relative to the C17:0 fatty acid internal standard, and expressed as a percentage of total fatty acid content.
3. Results and discussion 3.1. Collection of samples, establishment and identification of algal strains One strain of algae-like microorganisms capable of mixotrophic growth was isolated from Tainan (located in Southern Taiwan). Using BLAST software to align and compare the 18S rRNA gene sequences with other known microorganisms in the GenBank database revealed that this strain exhibited a close phylogenic relationship with the Chlorella family (NCBI number KJ855064), which are microalgae well-known for their high lipid content (Fig. 1). According to the phylogenetic analysis, it is reasonable to identify Y8-1 as Chlorella sp. Y8-1. The intracellular lipid droplets of Chlorella sp. Y8-1 were observed using Sudan Black B and Nile Red staining under fluorescence microscopy; the lipids were stained in black and yellow, respectively (data not shown). 3.2. Effects of the carbon sources on the growth and lipid production of Chlorella sp. Y8-1 in mixotrophic culture In mixotrophic cultivation, Chlorella sp. Y8-1 exhibited rapid growth in response to the addition of carbon, except xylose
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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pH
12.0 (a) Blank (without carbon & nitrogen source) 10.5
(b) Blank 1 (without carbon source)
(c) fructose: 1 g/L
(d) glucose: 1 g/L
(e) glycerol: 1 g/L
(f) sucrose: 1 g/L
(g) xylose: 1 g/L
9.0 7.5 6.0
(
0.8
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0.4 0.2
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) Carbon source conc. (g/L)
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4 6 8 10 12 0 Time (Day)
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Time (Day)
0
2
4
6
8 10 12
0.0
(
) Dried cell weight (g/L)
4.5 0.75
Time (Day)
Fig. 2. The time courses of dried cell weight and sugar consumption of isolated Chlorella sp. Y8-1 microalgae at various carbon sources. Culture condition: light/dark cycle: 24/ 0; light source: fluorescent; light intensity: 4300 lux; nitrogen sources: urea 0.5 g/L; various carbon sources; aeration: 10% CO2; n = 3. Note: (a) and (b): autotrophic cultivation; (c)–(g): mixotrophic cultivation.
0.6
20 0.4
15 10
0.2
0.4 0.2
0.0
-1 -1
(
)
0.2
(day )
0.6 0.4
µm
0.0 0.8
0 0.6
(
)
5
during 7 days (day )
25
µ
0.8
30
0.0
Bl a B l nk an fru k 1 ct o gl s e uc gl ose yc e su r o l cr o x y se lo se
(
) Dried cell weight (g/L)
(
) Lipid content (%)
(Fig. 2). After a short delay phase of 1 d, the cells underwent exponential growth for 4 d and then entered the stationary phase after 7 d. The cells that contained fructose, glucose, glycerol, and sucrose as the carbon sources grew much more rapidly and higher than did those that contained xylose. The dried biomass concentration increased as high as 0.45 g/L during the late-exponential phase. In addition, the mixotrophic cells cultured in the presence of 10% CO2 (2 vvm), 1 g/L of sucrose and 0.5 g/L of urea reached the stationary phase more rapidly than those cultured in the presence of other carbons did. However, approximately equal dried biomass concentrations were obtained from the samples grown on glucose and glycerol under mixotrophic cultivation (Fig. 2). In mixotrophic cultivation, four carbon sources induced cell growth and lipid accumulation during the period of culture. As shown in Fig. 3, the specific growth rates (l) were significantly higher in the presence of organic carbon sources (fructose, glucose,
Carbon sources (1 g/L) Fig. 3. Lipid content and growth of Chlorella sp. Y8-1 after 14 day culture at various carbon sources (1 g/L). Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; nitrogen sources: urea 0.5 g/L; aeration: 10% CO2; n = 3. Note: Blank: without organic carbon and nitrogen sources (autotrophic cultivation). Blank 1: without organic carbon sources (autotrophic cultivation). Others: mixotrophic cultivation.
glycerol and sucrose) during 7 days. Additionally, the maximal specific growth rate (lm) (0.39 ± 0.04 d 1) and lipid content (24.0 ± 0.61% of dry cell weight) were obtained when sucrose was used as the organic carbon source. However, fructose presumably cannot be used to enhance the lipid content of Chlorella sp. Y8-1 under mixotrophic cultivation (Fig. 3). Adding a carbon source to the medium has been widely reported to increase the growth and lipid content of microalgae (Qlao and Wang, 2009). Chlorella species can grow on various carbon substrates such as sodium acetate (Qlao and Wang, 2009), fructose (Gao et al., 2009), glucose (Yeh and Chang, 2012), glycerol (Heredia-Arroyo et al., 2010), sucrose (Gao et al., 2009), and acetate (Heredia-Arroyo et al., 2010). Fructose, glucose, glycerol, and sucrose were transformed into glyceraldehydes-3-phosphate, which is a crucial intermediate product involved in both glycolysis and the pentose phosphate pathway (Lewin, 1974). However, xylose apparently cannot be used to grow microalgae effectively. Therefore, the growth was considerably slower when xylose was used as the organic carbon source, compared with the growth when the other four organic carbon sources were used. Sucrose can be decomposed to yield a glucose molecule and a fructose molecule, that is why microalgae can utilize sucrose to growth. Therefore, it is possible that Chlorella sp. possesses an inducible transport system for hexoses (Haass and Tanner, 1974), but not for pentose. Fig. 3 indicates that lipid accumulation occurred in mixotrophic cells (simultaneously using both 1 g/L glucose and 10% CO2, glycerol and 10% CO2, sucrose and 10% CO2 as carbon sources, respectively), and the lipid content of the autotrophic cells (Blank 1, using 10% CO2 as carbon sources) was only 6.0 ± 2.5%. It would be reasonable to suggest that TCA cycle is the starting point for many anabolisms, such as lipid, protein and so on. Therefore, this result explains the metabolic basis that lipid content was significantly lower in autotrophy culture than in the heterotrophy and mixtrophy culture. The lipid content of the mixotrophic cells observed in the current study was 9.4 ± 2.6–24.0 ± 3.2% (Fig. 3). It is possible that some carbon sources can lead to enhancement of acetyl CoA/malonyl CoA pool – which represents the central carbon donor for fatty acid synthesis, thereby increasing lipid pool (Ngangkham et al., 2012). Miao and Wu (2006) cultivated Chlorella protothecoides by using glucose and demonstrated that the lipid content was fourfold in response to heterotrophic growth when compared with autotrophic growth (Miao and Wu, 2006). In addition, the present study also found that the cell concentration (>0.40 g/L) of the Chlorella sp. Y8-1 from mixotrophic (using 1 g/L sucrose and 10% CO2 as carbon source) growth was higher than the sum of those from autotrophic (0.15 g/L) (using 10% CO2 as carbon source) and heterotrophic (using 1 g/L sucrose as carbon source) growth (0.17 g/L, data not shown). This indicated that the mixotrophic growth is not a simple combination of photoautotrophic and heterotrophic growth. Although it has been reported that
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx Table 1 Lipid and biomass production of different microalgae under photoautotrophic, heterotrophic and mixotrophic culture conditions, respectively. Microalgae
Autotrophic
Heterotrophic
Mixotrophic
Refs.
Lipid content (%)
Biomass (g/L)
Lipid content (%)
Biomass (g/L)
Lipid content (%)
Biomass (g/L)
Marine Chlorella sp. Y8-1 Chlorella sp. Chlorella vulgaris Chlorella sorokiniana Nannochloropsis sp.
16.5 30.0 – 19.0 28.0
0.22 0.38 – – 0.37
5.9 21.0 2.0 – 20.0
0.17 0.48 3.00 – 0.38
35.5 25.5 0.8 33.0 27.5
0.45 1.45 4.00 – 1.20
This study Cheirsilp and Torpee (2012) EL-Sheekh et al. (2012) Ngangkham et al. (2012) Cheirsilp and Torpee (2012)
Freshwater Chlorella sp. Chlorella vulgaris ESP-31
13.5 20.0
0.60 0.8
13.0 16.0
0.75 0.2
15.0 53.0
1.40 3.0
Cheirsilp and Torpee (2012) Yeh and Chang (2012)
Note: Values were not showed from the results of studies and labeled as ‘‘–‘‘ in the table.
0.8
20
0.6
15
0.4
10
0.2
5 (b) After 15 day
30
0.0 (b) After 15 day 1.0
25
0.8
20
0.6
15
0.4
10
0.2
5 0
0
0.1 0.25 0.5 0.75 1 1.5
2
Urea concentration (g/L)
0 0.1 0.25 0.5 0.75 1 1.5 2
0.0
0.5
0.4 0.3 0.2 0.1 0.0
0.4
-1
0.5
) µ during 7 days (day )
25
0
Lipid content (%)
1.0
(
(a) After 7 day
-1
(a) After 7 day
) µm (day )
Lipid content (%)
30
0.3 0.2 0.1
(
Our earlier studies revealed that urea was a suitable nitrogen source for cultivation of Chlorella sp. Y8-1. Urea was adopted into the medium at 0–2 g/L. The biomass, specific growth rates, and lipid content of Chlorella sp. Y8-1 in the stationary phase are shown in Fig. 4. The cultures had a low biomass concentration, specific growth rate, and lipid content in the N-free medium. The Chlorella sp. Y8-1 grew the most robustly in the medium containing 0.1– 0.25 g/L of urea, reaching a biomass concentration of
( ) Dried cell weight (g/L)
3.3. Effects of urea concentration on the growth and lipid production of Chlorella sp. Y8-1
0.68 ± 0.06 g/L; however, the growth was limited when the urea concentration was higher than 0.5 g/L (data not shown). Within the urea range from 0.5 to 2.0 g/L, the biomass concentration in the urea experiment decreased from 0.6 ± 0.05 to 0.5 ± 0.04 g/L. In the medium with 0.1–2.0 g/L of urea, the specific growth rate during 7 days and maximal specific growth rate slightly decreased as the urea concentration increased (Fig. 4). In the experimental range, the lipid content of Chlorella sp. Y8-1 increased as the culture period increased. In addition, the lipid content increased from 6.7 ± 3.9% to 31.5 ± 3.18% of dry cell weight as the urea was increased from 0 to 0.25 g/L; however, the lipid production was limited when the urea concentration was higher than 0.5 g/L. The maximal lipid content (31.5 ± 3.18% of dry cell weight) was obtained in the medium containing 0.25 g/L of urea (Fig. 4). In the current experiments, the higher the urea concentration was, the higher the lipid content of the Chlorella sp. Y8-1 when the urea ranged from 0 to 0.25 g/L. The results corresponded with those of several reports; for example, the lipid content of Ellipsoidion sp. increased as the nitrate concentration increased, and the maximal dry weight of 32.8% was obtained when the nitrate-N used was 1.92 mmol/L (Xu et al., 2001). Furthermore, Isochrysis galbana decreased markedly from 30% (NaNO3 > 2 mmol/L) to 19% of dry weight at 0.5 mmol/L NaNO3 (Molina and Sánchez,
( ) Dried cell weight (g/L)
photosynthesis and heterotrophic growth occur simultaneously and independently in mixotrophic cultures (Marquez et al., 1993), the presence of organic carbon can alter both the photosynthetic and heterotrophic metabolism of Chlorella (Villarejo et al., 1995). The lipid and biomass production of mixotrophic cultivation are much higher than under autotrophic and heterotrophic cultivation indicated in previous work (Table 1). Thus, these findings indicate that adding organic carbon sources leads to a marked stimulation of lipid accumulation in Chlorella sp. Y8-1, and that the ideal organic carbon source for the production of lipids is sucrose.
0.0
Urea concentration (g/L)
Fig. 4. Lipid content and growth of Chlorella sp. Y8-1 at various urea concentrations under mixotrophic cultivation. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; organic carbon sources: sucrose 1 g/L; aeration: 10% CO2; n = 3.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx
1992). However, for most microalgae, the lipid content increased under nitrogen deficient conditions. In other studies, Nannochloropsis oculata had maximal lipid content per cell under low nitrogen conditions (Converti et al., 2009), and the lipid content of Chlamydomonas reinhardtii CC124 and Chlorella vulgaris increased under nitrogen-deficient conditions (Deng et al., 2011). In general, this behavior is most probably a survival response until restoration of less nutritionally stressing conditions.
and Johns, 1995). Chen (1996) and Chen and Johns (1994) have both indicated that, compared with bacteria and yeasts, microalgae can tolerate only relatively low substrate concentrations (Chen, 1996; Chen and Johns, 1994). In addition, at the initial sucrose concentration of 0.5–1.0 g/L, the sucrose level gradually decreased and was nearly completely depleted at the end of cultivation. When a higher initial sucrose concentration of 5–10 g/L was used, approximately 3.0 g/L of sucrose remained at the end of cultivation. In our experiments, high concentrations of sucrose have been shown to inhibit microalgal growth, at least for a considerable period of time. It was likely that algal cells grown on high concentration of substrates require a lag period to adopt the bioreactor operation (Perez-Garcia et al., 2011). The effects of sucrose concentration on specific growth rate and lipid production are summarized in Fig. 6. As shown in Fig. 6, the specific growth rate during 7 days and maximal specific growth rate were significantly higher in the presence of sucrose. However, the specific growth rate during 7 days and maximal specific growth rate were decreased when the sucrose concentration was 10 g/L. It was also likely that algal cells grown on high concentration of substrates
3.4. Effects of sucrose concentration on the growth and lipid production of Chlorella sp. Y8-1 The effects of carbon source concentrations on the growth and lipid production of isolated Chlorella sp. Y8-1 are shown in Figs. 5 and 6. The growth rate of isolated Chlorella sp. Y8-1 increased when the initial sucrose concentration was increased from 0 to 0.75 g/L (Fig. 5). Increasing the initial sucrose concentration further did not enhance the growth of this strain. The growth rate slightly decreased when the initial sucrose concentration was increased to 1 g/L. This could have been an effect of substrate inhibition (Chen
pH
12.0 (a) Sucrose conc.: 0 g/L 10.5
(b) Sucrose conc.: 0.5 g/L
(c) Sucrose conc.: 0.75 g/L
(e) Sucrose conc.: 2.5 g/L
(d) Sucrose conc.: 1 g/L
(g) Sucrose conc.: 7.5 g/L
(f) Sucrose conc.: 5 g/L
(h) Sucrose conc.: 10 g/L
9.0 7.5 6.0 4.5
2
) Sucrose conc. (g/L)
8
0.8
0
(
(
) Dried cell weight (g/L)
1.0
6
0.6 4
0.4 0.2 0.0 0
3
6 9 12 15 0 Time (Day)
3
6 9 12 15 0 Time (Day)
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6 9 12 15 0 Time (Day)
3
6
9
12 15 0
Time (Day)
3
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3
6 9 12 15 0 Time (Day)
3
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3
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12 15
Time (Day)
0.6
0.4
0.2
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-1
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during 7 days (day )
0.0
0.3
0.1
0.4
( )
0.0 (b) After 15 day
0.4
-1
0.2
0.5
( ) µm (day )
0.4
) Dried cell weight (g/L)
0.6
(
(a) After 7 day
) Dried cell weight (g/L)
40 (a) After 7 day 35 30 25 20 15 10 5 0 40 (b) After 15 day 35 30 25 20 15 10 5 0 0 0.5 0.75 1 2.5 5 7.5 10 Sucrose concentration (g/L)
(
Lipid content (%)
Lipid content (%)
Fig. 5. The time courses of dried cell weight and sugar consumption of isolated Chlorella sp. Y8-1 microalgae at various sucrose concentrations. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; urea concentration: 0.25 g/L; various sucrose concentration; aeration: 10% CO2; n = 3. Note: (a): autotrophic cultivation. (b)–(h): mixotrophic cultivation.
0.3 0.2 0.1
0 0.5 0.75 1 2.5 5 7.5 10 Sucrose concentration (g/L)
Fig. 6. Lipid content and growth of Chlorella sp. Y8-1 at various sucrose concentrations. Culture conditions: light/dark cycle: 24/0; light source: fluorescent; light intensity: 4300 lux; urea concentration: 0.25 g/L; various sucrose concentration; aeration: 10% CO2; n = 3. Note: 0 g/L sucrose: autotrophic cultivation. 0.5–10 g/L sucrose: mixotrophic cultivation.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
T.-S. Lin, J.-Y. Wu / Bioresource Technology xxx (2014) xxx–xxx
require a lag period to adopt (Perez-Garcia et al., 2011). The lipid production of Chlorella sp. Y8-1 increased from 16.5 ± 3.6% to 35.5 ± 4.2% when the initial sucrose concentration increased from 0 to 0.5 g/L (Fig. 6). It was likely that in limited N condition, protein, nucleic acid synthesis slow down so cell division and growth are restricted, most assimilated carbon are led to produce storage and stress-resistant materials such as lipid, starch. When the initial sucrose concentration was higher than 0.75 g/L, the lipid production of Chlorella sp. Y8-1 gradually decreased. Cheirsilp and Torpee (2012) also discovered that the lipid content of Nannochloropsis sp. increased from 109.8 to 798.1 mg/L when the initial glucose concentration was increased from 0 to 15 g/L, but the lipid content slightly decreased when the initial glucose concentration was 20 g/L (Cheirsilp and Torpee, 2012). Consequently, these findings indicate that the ideal sucrose concentration for lipid production of Chlorella sp. Y8-1 is 0.5 g/L. 3.5. Fatty acid composition of Chlorella sp. Y8-1 in mixotrophic culture The primary fatty acids in Chlorella sp. Y8-1, as percentages of total fatty acids, were 16:0 (19.2 ± 0.6%), 18:1 (11.2 ± 0.4%), 18:2 (18.0 ± 0.6%), and 18:3 (10.5 ± 0.3%) (data not shown), indicating good agreement with the results observed in the lipid production of C. vulgaris (Yusof et al., 2011). The major saturated fatty acids (SFA) in Chlorella sp. Y8-1 were palmitic acid (16:0) and stearic acid (C18:0). The major monounsaturated fatty acid (MUFA) was oleic acid (18:1). It indicated that Chlorella sp. Y8-1 accumulated a high proportion of SFA and MUFA to 50% of the total lipid content. Compared to the commonly used soybean oil as feedstocks for biodiesel production in the US, the biodiesel derived from microalgae lipid in this study would be more saturated and provide a higher cetane number (CN), lower NOx emissions, shorter ignition delay time, and higher oxidative stability. Numerous fatty acids could not be identified in this study, because unsaturated fatty acids largely prevail in lipids. 4. Conclusions Compared to photoautotrophic and heterotrophic cultures, an abundance of lipids were accumulated when Chlorella sp. Y8-1 was cultivated under mixotrophic conditions, suggesting that it has great potential for renewable biodiesel feedstock applications. Mixotrophic Chlorella sp. Y8-1 showed higher lipid content (35.5 ± 4.2%) and higher lipid productivity (0.01 g/L/d) than Chlorella sp. Y8-1 cultivated under autotrophic and heterotrophic conditions. Additionally, under mixotrophic conditions, the microalgae accumulates lipids up to 35.5 ± 4.2% of dry cell weight, with palmitic acid (16:0) and stearic acid (C18:0) representing the most abundant fatty acid components. Mixotrophic cultivation is much easier to alter conditions to improve the yield of biomass and lipid of microalgal production. Therefore, further study on the influence of environmental factors would facilitate the improvement of algal lipid production. Acknowledgement This work was financially supported in part by a grant from the National Science Council of Republic of China under a contract number of NSC 100-2632-B-212-001-MY3. References Andrade, M.R., Costa, J.A.V., 2007. Mixotrophic cultivation of microalga Spirulina platensis using molasses as organic substrate. Aquaculture 264, 130–134. Bligh, E.G., Dyer, W.J., 1959. A rapid method for total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917.
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Bouarab, L., Dauta, A., Loudiki, M., 2004. Heterotrophic and mixotrophic growth of Micractinium pusillum Fresenius in the presence of acetate and glucose: effect of light and acetate gradient concentration. Water Res. 38, 2706–2712. Burja, A.M., Radianingtyas, H., Windust, A., Barrow, C.J., 2006. Isolation and characterization of polyunsaturated fatty acid producing Thraustochytrium species: screening of strains and optimization of omega-3 production. Appl. Microbiol. Biotechnol. 72, 1161–1169. Cheirsilp, B., Torpee, S., 2012. Enhanced growth and lipid production of microalgae under mixotrophic culture condition: effect of light intensity, glucose concentration and fed-batch cultivation. Bioresour. Technol. 110, 510–516. Chen, F., 1996. High cell density culture of microalgae in heterotrophic growth. Trends Biotechnol. 14, 421–426. Chen, F., Johns, M.R., 1994. Substrate inhibition of Chlamydomonas reinhardtii by acetate in heterotrophic culture. Proc. Biochem. 29, 245–252. Chen, F., Johns, M.R., 1995. A strategy for high cell density culture of heterotrophic microalgae with inhibitory substrates. J. Appl. Phycol. 7, 43–46. Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., Borghi, M.D., 2009. Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chem. Eng. Process. 48, 1146–1151. Deng, X., Fei, X., Li, Y., 2011. The effects of nutritional restriction on neutral lipid accumulation in Chlamydomonas and Chlorella. Afr. J. Microbiol. Res. 5, 260–270. Dubois, M., Gilles, K.A., Hamilton, J.K., Rebers, P.A., Smith, F., 1956. Colorimetric method for determination of sugars and related substances. Anal. Chem. 28, 350–356. EL-Sheekh, M.M., Bedaiwy, M.Y., Osman, M.E., Ismail, M.M., 2012. Mixotrophic and heterotrophic growth of some microalgae using extract of fungal-treated wheat bran. Int. J. Recycling Org. Waste Agric. 1, 12–20. Fernández Sevilla, J.M., Cerón García, M.C., Sánchez Mirón, A., Belarbi, E.H., García Camacho, F., Molina Grima, E., 2004. Pilot plant-scale outdoor mixotrophic cultures of Phaeodactylum tricornutum using glycerol in vertical bubble column and airlift photobioreactors: studies in fed-batch mode. Biotechnol. Prog. 20, 728–736. Gao, C., Zhai, Y., Ding, Y., Wu, Q., 2009. Application of sweet sorghum for biodiesel production by heterotrophic microalga Chlorella protothecoides. Appl. Energy 87, 756–761. Haass, D., Tanner, W., 1974. Regulation of hexose transport in Chlorella vulgaris. Plant Physiol. 53, 14–20. Heredia-Arroyo, T., Wei, W., Hu, B., 2010. Oil accumulation via heterotrophic/ mixotrophic Chlorella protothecoides. Appl. Biochem. Biotechnol. 162, 1978– 1995. Ho, S.H., Chen, C.Y., Chang, J.S., 2012. Effect of light intensity and nitrogen starvation on CO2 fixation and lipid/carbohydrate production of an indigenous microalga Scenedesmus obliquus CNW-N. Bioresour. Technol. 113, 244–252. Hu, H., Gao, K., 2003. Optimization of growth and fatty acid composition of a unicellular marine picoplankton, Nannochloropsis sp., with enriched carbon sources. Biotechnol. Lett. 25, 421–425. Jeon, Y.C., Cho, C.W., Yun, Y.S., 2006. Combined effects of light intensity and acetate concentration on the growth of unicellular microalga Haematococcus pluvialis. Enzyme Microb. Technol. 39, 490–495. Khozin-Goldberg, I., Cohen, Z., 2006. Effect of phosphate starvation on lipid and fatty acid composition of the Eustigmatophyte Monodus subterraneus. Phytochemistry 67, 696–701. Lee, Y.K., 2004. Algal nutrition: heterotrophic carbon nutrition. In: Richmond, A. (Ed.), Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell, Oxford, pp. 116–124. Lewin, R.A., 1974. Biochemical taxonomy. In: Stewart, W.D.P. (Ed.), Algal Physiology and Biochemistry. Blackwell Scientific Publications, Oxford, UK, pp. 1–25. Marquez, F.J., Sasaki, K., Kakizono, T., Nishio, N., Nagai, S., 1993. Growth characteristics of Spirulina platensis in mixotrophic and heterotrophic conditions. J. Ferment. Bioeng. 76, 408–410. Miao, X., Wu, Q., 2006. Biodiesel production from heterotrophic microalgal oil. Bioresour. Technol. 97, 841–846. Molina, G.E., Sánchez, P.J.A., 1992. EPA from Isochrysis galbana growth conditions and productivity. Process Biochem. 27, 299–305. Ngangkham, M., Ratha, S.K., Prasanna, R., Saxena, A.K., Dhar, D.W., Sarika, C., Prasad, R.B.N., 2012. Biochemical modulation of growth, lipid quality and productivity in mixotrophic cultures of Chlorella sorokiniana. Springerplus 1, 1–13. Perez-Garcia, O., Escalante, F.M.E., de-Bashan, L.E., Bashan, Y., 2011. Heterotrophic cultures of microalgae: metabolism and potential products. Water Res. 45, 11– 36. Qlao, H., Wang, G., 2009. Effect of carbon source on growth and lipid accumulation in Chlorella sorokiniana GXNN01. Chin. J. Oceanol. 27, 762–768. Tamura, K., Dudley, J., Nei, M., Kumar, S., 2007. MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol. 24, 1596–1599. Vasudevan, P.T., Briggs, M., 2008. Biodiesel production current state of the art and challenges. J. Ind. Microbiol. Biotechnol. 35, 421–430. Villarejo, A., Orús, M.I., Martínez, F., 1995. Coordination of photosynthetic and respiratory metabolism in Chlorella vulgaris UAM 101 in the light. Physiol. Plant. 94, 680–686. Xu, N., Zhang, X., Fan, X., Han, L., Zeng, C., Tseng, C.K., 2001. Effects of nitrogen source and concentration on growth rate and fatty acid composition of Ellipsoidion sp. (Eustigmatophyta). J. Appl. Phys. 13, 463–469. Yeh, K.L., Chang, J.S., 2012. Effects of cultivation conditions and media composition on cell growth and lipid productivity of indigenous microalga Chlorella vulgaris ESP-31. Bioresour. Technol. 105, 120–127.
Please cite this article in press as: Lin, T.-S., Wu, J.-Y. Effect of carbon sources on growth and lipid accumulation of newly isolated microalgae cultured under mixotrophic condition. Bioresour. Technol. (2014), http://dx.doi.org/10.1016/j.biortech.2014.11.005
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Yusof, Y.A.M., Basari, J.H., Mukti, N.A., Sabuddin, R., Muda, A.R., Sulaiman, S., Makpol, S., Ngah, W.Z.W., 2011. Fatty acids composition of microalgae Chlorella vulgaris can be modulated by varying carbon dioxide concentration in outdoor culture. Afr. J. Biotechnol. 10, 13536–13542.
Zheng, L.S.T., Oh, J.Y., Jeon, B.H., Moon, H.S., Kwon, S.U., Lim, B.K., Kang, C.W., 2012. The dietary effects of fermented Chlorella vulgaris (CBT) on production performance, liver lipids and intestinal microflora in laying hens. AsianAustralas. J. Anim. Sci. 25, 261–266.
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