Influence of nutrient status on the accumulation of biomass and lipid in Nannochloropsis salina and Dunaliella salina

Energy Conversion and Management 106 (2015) 61–72

Contents lists available at ScienceDirect

Energy Conversion and Management journal homepage: www.elsevier.com/locate/enconman

Influence of nutrient status on the accumulation of biomass and lipid in Nannochloropsis salina and Dunaliella salina Yimin Chen a,b,⇑,1, Xu Tang a,1, Rahul Vijay Kapoore b,1, Changan Xu a,⇑, Seetharaman Vaidyanathan b,⇑ a b

Third Institute of Oceanography, State Oceanic Administration, Xiamen 361005, People’s Republic of China ChELSI Institute, Advanced Biomanufacturing Centre, Department of Chemical and Biological Engineering, The University of Sheffield, Sheffield S1 3JD, UK

a r t i c l e

i n f o

Article history: Received 28 July 2015 Accepted 9 September 2015

Keywords: Nutrient stress Microalgae Physiology Biofuel Lipid profile

a b s t r a c t Microalgae have been proposed as carbon-neutral and sustainable fuel feedstock producers due to their fast growth and high lipid content. In this investigation, we examined the effect of different nutrient stresses on overall lipid productivities and physiology, in two microalgae species, namely Nannochloropsis salina and Dunaliella salina. For both the species in semi-continuous cultures, the dilution ratio was found to affect only the harvest period and not the biomass or lipid productivity. Harvest at different growth phases was influenced by the temporal nutrient status of the medium leading to different productivity of biomass and lipids. These two species also showed different cellular biochemistry and lipid metabolism in response to the different nutrient stresses. More efficient stimulators of lipid production than the commonly employed nitrogen limitation were identified, for e.g. deficiency of trace element. Significant influence of nutrient deficiency on lipid profiles was observed in N. salina, such as chain length and degree of unsaturation, with desirable characteristics for biodiesel production. The comparison of lipid profiles between these two species revealed phenomenal differences in some fatty acids, e.g. a higher level of c-linolenic acid (C18:3n6) in D. salina and the exclusive presence of palmitoleic acid in N. salina. The biochemical composition with respect to the cellular function of the fatty acids is discussed with an attempt to better understand the differences in the lipid profiles. Ó 2015 Published by Elsevier Ltd.

1. Introduction Global warming and energy crisis due to the combustion and depletion of conventional fossil fuel have called for substantial interests and efforts in exploring and developing economically and environmentally sustainable renewable fuels [1]. Lipidderived biodiesel from microalgae, the third generation biofuel, provides a promising alternative to petroleum fuels that is potentially sustainable, carbon neutral, and can have a significantly smaller footprint in land area demand, compared to plant-based first and second generation biofuels [2–4]. Several critical culture conditions, including light intensity, temperature, and nutrient concentration, have been found to significantly influence both growth of microalgae and lipid accumulation [5–7]. Amongst these conditions, the nutrient status, in particular nutrient limitation, is

⇑ Corresponding authors at: Third Institute of Oceanography, State Oceanic Administration, Xiamen 361005, People’s Republic of China (Y. Chen). E-mail addresses: [email protected] (Y. Chen), [email protected] (C. Xu), [email protected] (S. Vaidyanathan). 1 The authors contribute equally to the article. http://dx.doi.org/10.1016/j.enconman.2015.09.025 0196-8904/Ó 2015 Published by Elsevier Ltd.

known to have a considerable influence on physiology, and both the quantity and quality of lipids accumulated [8,9]. Microalgae can be cultured in either batch [10,11], continuous [12] or fed-batch/semi-continuous [13] modes. Each cultivation mode has its own advantages and disadvantages [2]. In continuous systems, cellular metabolism can be maintained in a pseudosteady state with the continuous feeding of nutrients, leading to steady biomass productivity. In addition, in chemostatic continuous cultures with successive reductions in influent nitrogen, Chlorella sorokiniana and Oocystis polymorpha has been shown to exhibit drastic reduction in photosynthesis with decreased cellular nitrogen content from about 10–4%, but with little change in lipid content [14]. In contrast, an increase in total fatty acids and remarkable variations in the composition of the fatty acid fraction were induced in batch culture where cellular nitrogen could be decreased to
3% of dry weight. Therefore, these results suggest that nitrogen limitation in batch-cultured cells (or others) is more useful than in continuous culture systems for lipid accumulation [10,11,14]. In spite of this, a recent research proposed an integrated process scheme of continuous and batch wastewater treatment at industrial scale due to the good performance of Chlorella protothecoides in both systems [12]. On the other hand, algal cells in

62

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

semi-continuous cultures are also able to produce high lipid content with reported lipid productivities of 0.139 g L1 d1 [15]. Regardless of the culturing mode, however, there is one common issue that needs to be addressed; that of when to harvest the cells and at which phase of growth to maximise product yields since the nutrient status is different. The answer may vary for different microalgal strains (depending on their physiological characteristics) and for different purposes (depending on the target products like biomass, starch or lipids) [6,16]. Nutrient control like nitrogen starvation has been widely reported to increase lipid concentration in algae on the basis of dry cell weight, but this is at the cost of growth [17]. Because lipid productivity takes into account both the lipid concentration per unit cell and the cell concentration (biomass), it may have only a slight increase or may even show a decrease in comparison to the normal growth condition [18]. Taking a freshwater microalga, Scenedesmus sp., as an example, limited supply of either nitrogen or phosphorus has been shown to improve lipid content up to 30% and 53%, respectively, without enhancement of the total lipid productivity [19]. This raises the question as to whether other nutrient conditions can increase not just the lipid content (%) but more importantly the total lipid productivity. Nannochloropsis is renowned for high lipid content especially the ability to stimulate lipid accumulation under stress conditions (such as nitrogen starvation) [20–22]. This universally distributed microalga plays a significant role in the global carbon and mineral cycles, especially in oligotrophic seawater [23]. It has been used as a feed source in aquaculture and is proposed as an alternative source of fish oil due to its high content of PUFAs (polyunsaturated fatty acids), especially EPA (Eicosapentaenoic acid) [24]. Dunaliella is a green marine microalga belonging to the family Chlorophyceae [25]. It is extremely salt-tolerant with a capability to grow in salinities from 0.05 to 5.0 M NaCl, whilst maintaining a relatively low intracellular sodium concentration [26]. For this reason, D. salina can be grown under high salinity to limit contamination ensuring the dominance of this species in the culture. The algal cells do not contain rigid cell walls but surrounded by a thin elastic membrane and are known to grow fast and accumulate carotenoids and proteins under various stress conditions [27]. It is the first microalgal species used for commercial production of b-carotene [28]. D. salina can also be used for bioethanol fermentation or biogas production due to the high biomass productivities [29,30]. We examined the dilution ratio (i.e. volume ratio of new medium to total culture) and the harvest stage to work out the optimal operation conditions for Nannochloropsis salina and Dunaliella salina with the requirement for maximising biomass or lipid yield. In addition, N. salina and D. salina were grown under four different nutrient-deficient conditions, namely conditions deficient in nitrogen (N), phosphorous (P), vitamins (V) or trace elements (T), to examine the effect of the nutrient status on the corresponding growth rate, lipid productivity as well as the physiology. We demonstrate that more efficient nutrient conditions can be found for enhanced lipid production than the commonly employed nitrogen deficiency, under carbon limited growth conditions. In addition, we analyse and discuss the connection between fatty acid profiles and their metabolic roles, in the two species investigated.

2. Materials and methods

two species in the two experiments (as detailed below) were all cultivated in the same system (Fig. 1). Cells were grown in Erlenmeyer flasks, aerated and incubated at 24 ± 2 °C. Cultures were exposed to light intensity of
80 lE m2 s1, using fluorescent lamps. The microalgal cells were harvested by centrifugation (3000g for 3 min coupled with 8500g for another 5 min) and the cell pellets were frozen at 20 °C until analysis. The f/2 medium used in the cultivations throughout is composed of (per litre) 33.6 g artificial seawater salts (Ultra Marine Synthetica Sea Salt, Waterlife), 75 mg NaNO3, 5.65 mg NaH2PO4 2H2O, 1 ml trace elements stock and 1 ml vitamin mix stock. The trace elemental solution (per litre) includes 4.16 g Na2EDTA, 3.15 g FeCl36H2O, 0.18 g MnCl24H2O, 10 mg CoCl26H2O, 10 mg CuSO45H2O, 22 mg ZnSO47H2O, 6 mg Na2MoO42H2O. The vitamin mix solution (per litre) includes 100 mg vitamin B1, 0.5 mg vitamin B12 and 0.5 mg biotin. Growth curves of both species were monitored by measuring the optical density at 680 nm (OD680) with a UV/Visible spectrophotometer (Ultrospec 2100 pro). The OD680 was translated to dry cell weight (DCW) using pre-calibrated equations. For N. salina,

y ¼ 0:193x þ 0:0078 R2 ¼ 0:9914

ð1Þ

and for D. salina,

y ¼ 0:2895x þ 0:0011 R2 ¼ 0:9968

ð2Þ

where y and x are biomass concentration (g L1) and OD680, respectively. 2.2. Influence of harvest stage and dilution ratio in semi-continuous cultures Harvest stage of algae and medium dilution ratio (DR, the volume proportion of freshly added medium to the original total culture) can affect the nutrient status of the algae remaining in the medium for ongoing cultivation after partial harvest, and in turn the biomass and lipid productivities. To investigate the effect of these two parameters on biomass and lipid productivity, an aliquot of culture (corresponding to the DR) was harvested in three growth phases: middle active growth phase, late active growth phase and stationary phase. An equivalent amount of fresh medium was added to the old culture to maintain a constant culture volume. Two dilution ratios were studied: 0.25 and 0.5. 2.3. Influence of nutrient deficiency The two experimental species of N. salina and D. salina were grown at four nutrient-deficient conditions, including deficiency in nitrogen, phosphorous, vitamins, and trace elements, to investigate the effect of nutrient deficiency. To do this, some of the stock cultures in the middle active growth phase without any operation were directly used as reference (control 1). Cells from other stock cultures were harvested, centrifuged (3000g for 3 min coupled with 8500g for another 5 min), and the cell pellets were resuspended in complete fresh medium (i.e. f/2 medium, control 2), nitrate-free medium, phosphate-free medium, vitamin-free medium, and trace elements-free medium, separately, with the same volume as original. Each of the nutrient-deficient medium was prepared by completely removing the corresponding nutrient from the f/2 medium.

2.1. Microalgae cultivation, harvest and growth measurement 2.4. Biochemical compositional analysis Two marine microalgae, N. salina CCAP 849/2 and D. salina CCAP 19/30 were investigated in this study. Both were grown in the f/2 medium with three biological replicates in each experiment. The

Since the nutrient status can have influence on the fixed carbon distribution, cells were harvested for the assay of major

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

63

Fig. 1. Experimental set-up used in the study. It consists of (1) an air pump; (2) moisturizers containing sterilized water; (3) enclosed incubator; (4) microalgal cultures; and (5) a fluorescence lamp.

biochemical composition as cellular indicators to the changes in nutrients and assayed as detailed elsewhere [31,32]. These indicators include pigments, carbohydrates, proteins and lipids. Briefly, harvested cell pellets (from 2 ml of culture) were lysed in an alkaline solution by glass bead-beating using a cell disruptor (DISRUPTOR GENIEÒ, USA). An aliquot of this lysate was used for carbohydrate assay. The remaining sample was saponified by heating at 100 °C for 30 min and an aliquot of the saponified sample was used for protein assay, after it was brought to room temperature. Another aliquot of the sample was mixed with an organic solvent (chloroform:methanol, 2:1). Followed by vortex and centrifugation, the supernatant aqueous phase was used for chlorophyll assay; whist the bottom organic phase was used for total carotenoids and lipid assay. 2.5. Lipid profiles Both the lipid concentration and the lipid composition can be modified as a result of the changes in nutrients [6,33]. To find out the influence, algal samples (5 ml each) from the second experiment (Section 2.3) were harvested for lipid profiling by using GC–FID (gas chromatography–flame ionised detector). The lipid profile was carried out by three major steps: lipid extraction, lipid derivatisation, and spectral acquisition by GC–FID. The lipid extraction and derivatisation forming fatty acid methyl esters (FAMEs) followed Vandenbrouck’s protocol [34]. The FAMEs were then analysed by GC–FID using a capillary TR-FAME column (25 m  0.32 mm ID  0.25 lm film, Thermo Scientific, UK). Helium was used as the carrier gas at a flow rate of 1 ml/min. The injector temperature was maintained at 250 °C. The derivatised sample (1 lL) was injected in split mode at an oven temperature of 150 °C and the split flow was 120 ml/min at a ratio of 100:1. The temperature gradient was 150 °C for 3 min, 150–240 °C at 8 °C/min and 240 °C for 6 min. The FAMEs were identified according to the retention time of the corresponding peaks in the standard ‘‘F.A.M.E. Mix C8-C24” (Supelco Analytical, USA), which was injected before and after the real samples.

3. Results and discussions 3.1. Influence of dilution ratio and harvest stage in the semicontinuous system In a semi-continuous system, a proportion of the culture is harvested for desired products and supplemented by the same amount of new medium. There are two key questions that need consideration: the amount of algae that can be harvested and the time point for harvest. In other words, the dilution ratio and harvest stage are two critical parameters in that they can affect the nutrient status and thus, affect biomass and lipid production. Both of these are in fact dependent on the purpose of the microalgal cultivation, i.e., whether for accumulation of biomass, for lipids or for other products. The dilution ratio and the harvest stage in microalgae semi-continuous systems have been optimised to produce different target products, such as hydrogen [35] and lipids [36]. N. salina and D. salina are two promising candidates in industry and have been cultivated for producing lipids [37] and b-carotene or lutein [27], respectively. In order to optimise the operational parameters, N. salina and D. salina cultures were harvested at three different growth stages with two dilution ratios (Table 1). The active growth phase of each species that indicates a high rate of biomass production was separated to two sub-phases: middle active growth phase and late active growth phase. Besides, the stationary phase was also studied since lipid accumulation usually occurs in this phase. Regarding the dilution ratio, too low a value (e.g. 0.1) would imply frequent harvest and low efficiency, whilst too high a value (e.g. 0.9) would elongate the course of harvest and lead to low biomass productivity (as the remaining cell concentration is very low). Therefore, a low or high value of dilution ratio should be avoided. The two dilution ratios tested were 0.25 and 0.5 in this study. The time profiles for biomass under the different test conditions are plotted in Fig. 2, and include a control culture where cells kept growing without any disturbance. As shown, N. salina had three clear physiological growth phases: a lag phase (
0–50 h), an active growth phase (
50–110 h), and a stationary phase (
110–150 h).

64

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

Table 1 Effects of dilution ratio and harvest stages on the harvest period, biomass productivity, medium replacement rate and lipid productivity. The values shown are average of 3–5 determinations. Harvest period indicates the time interval required for twice successive harvests (also indicates harvest frequency). Dilution ratio N. salina 0.25

0.5

D. salina 0.25

0.5

Biomass harvest stage (g L1)

Growth phase

Lipid content (%)

Harvest period (h)

Biomass harvest rate (mg L1 h1)

Fresh medium replacment rate (ml h1)

Lipid harvest rate (mg L1 h1)

0.12–0.14 0.16–0.18 0.20–0.22 0.12–0.14 0.16–0.18 0.20–0.22

Middle active growth Late active growth Stationary Middle active growth Late active growth Stationary

16.92 ± 0.31 24.10 ± 0.82 40.48 ± 1.78 16.42 ± 0.15 24.43 ± 0.55 41.13 ± 1.57

12.00 ± 1.47 12.25 ± 1.33 23.83 ± 0.29 24.00 ± 0.5 30.50 ± 0.77 45.75 ± 2.47

2.83 ± 0.15 2.74 ± 0.09 2.15 ± 0.12 2.72 ± 0.04 2.94 ± 0.18 2.41 ± 0.33

3.13 ± 0.39 2.46 ± 0.35 1.57 ± 0.02 3.13 ± 0.07 2.45 ± 0.78 1.64 ± 0.09

0.48 ± 0.03 0.66 ± 0.02 0.87 ± 0.05 0.45 ± 0.01 0.72 ± 0.04 0.99 ± 0.13

0.17–0.22 0.23–0.26 0.29–0.32 0.17–0.22 0.23–0.26 0.29–0.32

Middle active growth Late active growth Stationary Middle active growth Late active growth Stationary

20.83 ± 0.67 24.44 ± 0.93 26.50 ± 1.03 20.92 ± 0.79 24.35 ± 0.84 26.21 ± 0.98

11.00 ± 0.51 23.67 ± 1.61 40.83 ± 2.03 31.50 ± 1.23 48.00 ± 2.50 70.25 ± 3.35

3.40 ± 0.20 2.51 ± 0.01 1.90 ± 0.12 3.12 ± 0.14 2.52 ± 0.02 2.10 ± 0.06

2.70 ± 0.26 1.58 ± 0.10 0.92 ± 0.08 2.38 ± 0.26 1.56 ± 0.05 1.07 ± 0.01

0.71 ± 0.05 0.61 ± 0.01 0.50 ± 0.03 0.65 ± 0.03 0.61 ± 0.01 0.55 ± 0.02

Fig. 2. Effect of dilution ratio and harvest stage on biomass productivity: (a) and (b) are N. salina at dilution ratio (DR) of 0.25 and 0.5, respectively; (c) and (d) are D. salina at dilution ratio of 0.25 and 0.5, respectively. HB: harvested biomass; three stages with different HB were investigated corresponding to the middle growth phase, late growth phase and early stationary phase. The biomass concentration at each harvest point was derived from the measured OD680 (refer Table 1).

Although these two species started from a similar biomass concentration, only two apparent phases were observed in D. salina: an active growth phase (
0–90 h) and a stationary phase (
90–165 h). This is because these two species have different growth cycles. D. salina started to grow at a relatively higher cell concentration relative to its own growth cycle and thus, the lag phase was not present. For the same reason, it can be seen that N. salina reached stationary phase (at
110 h) later than D. salina (at
90 h). In spite of this, the third harvest stage point of N. salina was a little early (at
90 h) before reaching stationary phase as it was found to accumulate lipids from the late active growth phase. The detailed effects of dilution ratio and harvest stage on the harvest period, biomass productivity, rate of medium replacement,

and lipid productivity are elaborated in Table 1. For N. salina, the harvest periods (or harvest frequencies) were similar between middle and late active growth phases due to their similar growth rates. However, the harvest period in stationary phase was nearly twofold compared to that in active growth phase. Generally, the harvest period at the dilution ratio of 0.5 was nearly twice longer than that at the dilution ratio of 0.25 indicating the approximately linear relationship between the harvest period and the dilution ratio. As shown in Table 1, as the harvest stage extended, the fresh medium replacement rate decreased, suggesting decreased requirement for the amount of fresh medium. Nevertheless, the medium replacement rate was nearly unchanged for the same growth phase at different dilution ratios. Hence, it implies that

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

the requirement of fresh medium does not relate to the dilution ratio but only relates to the growth phase when cells were harvested (i.e. harvest stage). It should be pointed out that a low medium replacement rate can conserve consumption of medium and the cost of nutrient inputs. For N. salina, the rate of lipid harvest accelerated with extension in biomass harvest stage due to lipid accumulation especially in stationary phase. As such, although the biomass harvest rate was the slowest in stationary phase, the lipid harvest rate (or lipid productivity) was the fastest in this phase indicating the status of nutrient deficiency. For D. salina, the changing trends of the harvest period, biomass productivity and medium replacement rate were similar to those in N. salina. However, the lipid harvest rate in D. salina decreased with extension in biomass harvest stage, i.e. contrary to N. salina. This is because no lipid accumulation was observed in D. salina and the lipid content (%) was nearly unchanged in different growth phases. Therefore, if microalgae are cultured for collecting biomass, cells should generally be harvested in active growth phase when the growth rate is the highest. This requires frequent harvest due to the fast growth rate, compared to harvest at other growth phases. In turn, the frequent harvest reduces contamination risk. However, the requirement for frequent replacement of medium would increase the cost. Regarding the lipid production for biodiesel, it is better to harvest cells in the stationary phase but only for those oleaginous species with the ability to accumulate significant lipids like N. salina. It should be pointed out that there has to be a trade-off between the biomass/biofuel productivity and the cost. It was estimated that the minimum production cost of USD 73.5/L for indoor microalgae biodiesel was extremely higher than the petrol diesel price (USD 1.1/L in Malaysia); whilst cultivation under outdoor environment could reduce the production cost of dried microalgae biomass from USD 34.4/kg by using chemical fertilisers to USD 14.3/kg [38]. As such, to make it economically feasible, the nutrients should derive from a free or waste source like wastewater containing required nutrients or CO2 from exhaust gases. For instance, Gonçalves et al. [7] cultivated several microalgal species for the integration of biofuel production with CO2 sequestration and nutrient removal from wastewater. The results for N. salina are in agreement with an earlier report [36], in which three harvest stages (1, 2 and 3 d intervals) and two harvest ratios (25% and 50%, v/v) were examined in semicontinuous bench-scale reactors for growing N. salina. They found that the 2-d harvesting interval and 50% harvest ratio resulted in the highest lipid productivity of 38.7 mg L1 d1. In this study, we obtained around 24 mg L1 d1 of lipid productivity at also 50% harvest ratio and
2-d harvesting interval. The difference in lipid productivity may be owing to the different strains and biomass productivity. The species of D. salina is often used for pigment production. Prieto et al. [39] cultivated D. salina in three different culture systems for carotenoid production, including batch, semicontinuous, and a two-stage approach systems. They observed that the highest carotenoid content (10% of dry weight) was achieved in the closed and two-stage system with 90% b-carotene abundance of total carotenoids. In addition, the high carbohydrate content in D. salina makes it a good candidate for bioethanol or biogas production via anaerobic fermentation [40]. 3.2. Influence of nutrient deficiency on growth and lipid production 3.2.1. Biochemical composition Many microalgal species have been observed to accumulate lipids under nutrient-deficient conditions like nitrogen starvation. Nevertheless, the lipid accumulation appears to be in conflict with the growth rate. As a result, the total lipids might not be much higher than that under normal condition under which the cells

65

grow very fast and the lipids content is relatively low. Herein, four nutrient-deficient conditions were examined to look for conditions that induce lipid accumulation with minimal inhibition of growth (i.e., those conditions that would result in higher lipid productivities), under conditions where CO2 is supplied by passing air (0.04% CO2). The conditions tested were deficiency in nitrogen (N), phosphorous (P), vitamins (V) and trace elements (T). The response to these conditions were also compared with two control conditions, one replenished with fresh medium (New) and the other where the original medium was retained (Old). Interestingly, the growth curves for N. salina under the six conditions (Fig. 3a) fall into three groups in the decreasing order of growth: New, T (group A) > P, V (group B) > Old, N (group C), with average active growth rate of 0.08, 0.054 and 0.02 g L1 d1, respectively. The two-tail t-test showed significant difference between group A and C (p = 0.0001), and group B and C (p = 0.02). The fastest growth rate was found in the new medium and the trace elements-deficient medium. This demonstrates that sufficient nutrients ensure the healthy growth of cells except the deficiency of trace elements that does not have as much of an effect on the growth. Lack of phosphorous or vitamins led to a decrease in the growth rate to a similar degree. The growth rate was significantly lagged by the limited nutrient status in the old medium and in the nitrogen-deficient medium. Similar to the growth curves, time profiles of the pigment content based on DCW, including chlorophyll a (Ca%) and total carotenoids (Ct%), were also separated into three groups (Fig. 3b and c). The production of chlorophyll a and total carotenoids (no chlorophyll b in N. salina [31]) decreased with time under the old and the nitrogen-deficient media although the biomass still grew slowly. The decline in pigment production under these two conditions illustrates that the synthesis of pigments is closely related to nitrogen availability. The production of chlorophyll a under other conditions (excluding Old and N conditions) all increased with time after 20 h of cultivation. In contrast, the production of total carotenoids increased with time but dropped after
45 h. This might also be a result of the gradual decrease in available nitrogen towards the end of cultivation. Variations in the biochemical composition of N. salina are shown in Fig. 3d–f. In general, carbohydrates and proteins as a percentage of DCW declined with time under almost all conditions. Nonetheless, a clear enhancement in lipid content was observed under almost all the conditions. The reverse correlation between lipids and carbohydrates (or proteins) indicates that the cells under most nutrient-deficient conditions tend to store the sequestrated carbon and energy in the form of lipids, with increase in the time of exposure to the condition. It might also indicate that carbohydrates or proteins are partially converted to lipids. Carbohydrates as a percentage of DCW increased slightly towards the end of the cultivation showing that the carbohydrates were still slightly accumulated by the cells under all these conditions. Significant drop in Pro% can be seen under most of the tested conditions except for the T condition under which the protein content had an increase towards the end. Regarding the lipid productivity, the highest value on the third day was observed in the T medium (
0.08 g L1 d1) and the P medium (
0.064 g L1 d1), higher than that in the N medium (
0.032 g L1 d1). The difference of the lipid productivity was significant between the T medium and the N medium (p = 0.0004), and the P medium and the N medium (p = 0.0008), as seen by the two-tail t-test. It indicates that the T and P conditions were more efficient than N condition for lipid production as inhibition of growth was minimal. As displayed in Fig. 4, correlation of the growth curves for D. salina amongst these conditions was similar to that for N. salina with the same three groups observed. The average active growth rates of these three groups were 0.096, 0.042 and 0.012 g L1 d1,

66

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

Fig. 3. Effect of different nutrient limitations on growth, pigment and biochemical composition of N. salina with respect to time (three biological replicates were employed). Ca: Chlorophyll a; Ct: total carotenoids; Car: carbohydrates; Pro: proteins; Lipid: lipids – all expressed as a percentage of DCW.

respectively. The two-tail t-test showed significant difference between group A and B (p = 0.01), and group A and C (p = 0.013). There is no significant difference between the new media and the trace element-deficient medium in which the growth rate of cells was the fastest. Compared to the new medium, the P medium exhibited a larger drop in the growth rate than the V medium. Cells hardly grew under both the old and N media. Like with N. salina, the pigment content of D. salina including chlorophylls (a and b) and total carotenoids generally increased with time in almost all media, except the old and N media, which were in accordance with the growth curves. It confirms that nitrogen is a requirement for pigment synthesis. Nevertheless, the time point of
45 h seems to be a turning point after which pigment content under most conditions nearly ceased to increase and even declined. One particular exception was the cells growing in the V medium where the pigment content kept increasing all the time even when its biomass hardly grew after
45 h. Content of total carotenoids under most conditions slightly increased on the first day and hardly changed on the second day until 45 h when their difference started to be clearly distinguishable with a decreasing order: V, T > New, P > N, Old. The highest content of pigments was present in the V medium instead of the new medium indicating that the vitamin deficiency can be used to stimulate the production of pigments. As shown in Fig. 4e–g, the primary biochemical indicators of D. salina responded very differently to those nutrient conditions. The carbohydrate content under most conditions only increased on the first day and started to decline or remained nearly constant (Fig. 4e). It is worth noting that the carbohydrate content was

relatively stable, since the second day, in those media having higher biomass like the new, T and V media but exhibited a dramatic drop in the old, N and P media. The protein productivity in the old and N media was also found to be the lowest and even decreased on the third day (not shown). The highest protein content (%) was present in the P medium and remained at around 55% since the second day (Fig. 4f). Pro% in the other media decreased slightly. Lipid content (%) in the old and N media reached up to around 22 and 27%, respectively (Fig. 4g). Lipid% was also very high in the P medium (up to 28% on the third day) but slightly decreased (25%) towards the end of cultivation. However, the highest lipid productivity was observed in the New medium (
0.039 g L1 d1) and the T medium (
0.033 g L1 d1), higher than that in the N medium (
0.019 g L1 d1). The difference of the lipid productivity was significant between the New medium and the N medium (p = 0.007), the T medium and the N medium (p = 0.002), as seen by the two-tail t-test. It indicates that for the strain of D. salina employed in the study, nutrient deficient conditions do not seem to induce lipid accumulation. Many species have exhibited positive growth only under nitrogen sufficient conditions and nitrogen deficiency has been shown to significantly inhibit growth [41]. Regarding lipid content, lipid accumulation by nitrogen starvation has been shown in some species (e.g. Nannochloropsis [21], Pavlova viridis [41]) but this is not true for all microalgal species (e.g. Tetraselmis subcordiformis [41], D. salina [42] do not appear to show this trend). Deficient conditions of other nutrients have also been studied. For instance, C. reinhardtii can produce more hydrogen in sulphur deprived condition [43]. Low nitrogen and phosphate concentrations are able to

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

67

Fig. 4. Effect of different nutrient limitations on growth, pigment and biochemical composition of D. salina with respect to time (three biological replicates were employed). Ca: Chlorophyll a; Cb: Chlorophyll b; Ct: total carotenoids; Car: carbohydrates; Pro: proteins; Lipid: lipids – all expressed as a percentage of DCW.

stimulate more production of fatty acid methyl esters in Scenedesmus rubescens-like microalga [44]. Vanucci et al. [45] found that in Prorocentrum lima, protein concentration significantly decreased under N condition but nearly no changes were noted under P condition, which is similar to what is observed for D. salina in the present study. Carbohydrate content was significantly increased under both N and P conditions, whilst in the present study, the increase in carbohydrate content was only observed in the last day of cultivation for N. salina under almost all tested conditions. Therefore, it can be seen that the biochemical composition of cells can vary between species and strains. The influence of trace elements on the growth and lipid production has also been reported. However, most of those studies were on the basis of regulating the concentration of a single trace element and the co-effects of different trace elements were rarely reported. Amongst these trace elements, iron is one of the most often studied. It was found that a mild addition of iron could

increase both final cell densities and lipid content but further addition could be toxic [46,47]. On the contrary, iron limitation decreased chlorophyll content and esterase activity but maintained (severe limitation) or even increased (mild limitation) lipid content [48]. In contrast, copper starvation increased chlorophyll content, esterase activity and lipid content [48]. Therefore, the effects of iron and copper on chlorophyll content and esterase activity may be counteracted to some extent when these two ions are both deficient but leading to lipid accumulation like N. salina in the present study. This might be the reason that, in this study, the deficiency of trace elements did not cause large decreases in the growth and chlorophyll content. There is a study on the proteome of Cu, Fe, Zn, and Mn micronutrient deficiency in Chlamydomonas reinhardtii that can enhance our understanding on the influence of the trace element deficiency on the microalgal physiology [49]. Besides, Singh et al. [50] found that a certain combination of nitrogen, phosphorus and iron stresses could result in the

68

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

Table 2 Fatty acids detected in N. salina and D. salina. S and U stand for saturated and unsaturated. No.

FAs

S/U

No.

FAs

S/U

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

Butyric acid (C4:0) Capric acid (C10:0) Undecanoic acid (C11:0) Lauric acid (C12:0) Tridecanoic acid (C13:0) Myristic acid (C14:0) Myristoleic acid (C14:0) Pentadecanoic acid (C15:0) cis-10-Pentadecenoic acid (C15:1) Palmitic acid (C16:0) Palmitoleic acid (C16:1) Heptadecanoic acid (C17:0) cis-10-Heptadecanoic acid (C17:1) Stearic acid (C18:0) Elaidic acid (C18:1n9t) Oleic acid (C18:1n9c) Linolelaidic acid (C18:2n6t)

S S S S S S S S U S U S U S U U U

18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34

Linoleic acid (C18:2n6c) Arachidic acid (C20:0) c-Linolenic acid (C18:3n6) cis-11-Eicosenoic acid (C20:1) Linolenic acid (C18:3n3) Heneicosanoic acid (C21:0) cis-11, 14-Eicosadienoic acid (C20:2) Behenic acid (C22:0) cis-8, 11, 14-Eicosatrienoic acid (C20:3n6) Erucic acid (C22:1n9) cis-11, 14, 17-Eicosatrienoic acid (C20:3n3) Arachidonic acid (C20:4n6) Tricosanoic acid (C23:0) cis-13, 16-Docosadienoic acid (C22:2) Lignoceric acid (C24:0) cis-5, 8, 11, 14, 17-Eicosapentaenoic acid (C20:5n3) Nervonic acid (C24:1)

U S U U U S U S U U U U S U S U U

highest lipid productivity. Therefore, simultaneous management on various nutrients or elements can provide an efficient strategy for lipid production. 3.2.2. Lipid profiles The composition of the fatty acids in the biodiesel predominantly determines the quality of biodiesel, including properties such as cetane number, cold flow characteristics, heat of combustion, oxidative stability, viscosity, and lubricity [51]. Therefore, the lipids were extracted from the cells of N. salina and D. salina and profiled by using GC–FID and FAME analysis. Thirty-four fatty acids were identified in each species as listed in Table 2, consisting of sixteen saturated and eighteen unsaturated fatty acids. Most of the fatty acids identified were long carbon chain with only seven relatively short chains (shorter than 15 carbons), which accounted for around 15–25% of the total fatty acids under the various nutrient conditions. From Table 3, it can be seen that N. salina under all of the nutrient conditions exhibited a decrease in short chain fatty acids (%SCFA) (less than 20% of total FAs), compared to the initial nutrient state (condition I, i.e.
26%), with significant (two tailed t test) decrease relative to I observed for Old (p = 0.02), P (p = 0.03), V (p = 0.01) and T (p = 0.03) conditions. The %SCFA content under nitrogen deficient condition was significantly higher compared to Old (p = 0.03) and V (p = 0.03) conditions. This suggests that vitamins, which are absent in V condition and depleted in the Old medium, possibly contribute to maintaining SCFA content in N. salina. In contrast, for D. salina, a statistically significant increase in %SCFA was noted for the Old (p = 0.02) compared to I, and significant increases noted for N (p = 0.03) and P (p = 0.01) conditions compared to W (replete condition) and for P (p = 0.01) compared to T. This suggests that in D. salina, under the depletion of the key nutrients (N and P) the organism increases lipids with a relatively higher SCFA content. The dominance of long chain fatty acids (LCFA) in these two species was similar to Chlorella sp. that could reach more than 90% of LCFA under particular conditions [52]. Contents of unsaturated fatty acids (UFA) under the varied nutrient-deficient conditions are displayed in Table 4. An average composition of around 45% UFA suggesting that both algal species contained more saturated fatty acids (around 10% higher) than unsaturated fatty acids. The degree of unsaturation in biodiesel has a strong influence on the combustion characteristics and emissions [51]. A higher degree of unsaturation of biodiesel fuels extends the ignition delay as well as the start of combustion. Although a higher degree of unsaturation of biodiesel can decrease

viscosity, it also strongly decreases oxidative stability. For all the above reasons, the relatively less unsaturated fatty acids present in these two algal species appear to be a good characteristic for generating biodiesel. No significant difference of %UFA was observed between the conditions, in N. salina. However, it can be seen that cells of D. salina showed a significantly lower content of %UFA under N condition (p = 0.01), compared to I. The %UFA was also slightly (but significant statistically) lower under, P condition compared to the replete condition (W) (p = 0.002) or T conditions (p = 0.01). This suggests that phosphate depletion induces a decrease in %UFA in D. salina. Exploratory analysis on replicate measurements (three biological replicates) was carried out to obtain a better understanding of the influence of nutrient stress on the lipid production. The cultures in the end of cultivation (68 h) were chosen for the investigation as physiological difference had been accumulated with time and become clearly discernible. Principal component analysis (PCA) was carried out on the compositional information of the lipids. The scores and loadings biplot from the first two principal components (PCs), which explain 96% of the variance in the data, is given in Fig. 5. The nutrient-deficient conditions derived from N. salina (prefixed N) are grouped to the left in the scores plot, whereas those from D. salina (prefixed D) are grouped to the right. It indicates the significantly different responses of these two species to the changes in the nutrient content. Moreover, three major groups can also be seen in each species. For N. salina, the three groups are initial cultures (NI), nitrogen deficient cultures (NN) and other cultures, respectively. It indicates that the culturing time and the nitrogen deficiency are the major reasons that cause the difference amongst these conditions. For D. salina, in contrast, the three groups are relatively scattered. These are (a) initial (DI)/vitamin deficient (DV) cultures, (b) trace elements deficient (DT)/new (DW) cultures, and (c) nitrogen deficient (DN)/phosphate deficient (DP)/old (DO) cultures, respectively. This indicates that D. salina might be more sensitive in the lipid composition than N. salina to the varied nutrient conditions. The loadings demonstrate that changes in butyric acid (C4:0, No. 1), palmitic acid (C16:0, No. 10), palmitoleic acid (C16:1, No. 11), stearic acid (C18:0, No. 14) and c-linolenic acid (C18:3n6, No. 20) predominantly contribute to the variance in the data, between the two species. Butyric acid (C4:0), palmitic acid (C16:0), palmitoleic acid (C16:1) and oleic acid (C18:1n9c) represent the four major fatty acids in N. salina, taking up around 7.8%, 30.5%, 29.4% and 9.9%, respectively, of total FA under N condition (Fig. 6). In D. salina,

69

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

Table 3 Percentage of relatively short chain fatty acids (containing less than 15 carbons) based on the total fatty acids. I – initial medium, W – complete new medium, O – old medium, N – nitrogen deficient medium, P – phosphor deficient medium, V – vitamin deficient medium and T – trace elements deficient medium. Two-tailed Student t test was carried out for paired samples between any two nutrient-deficient conditions. The values in bold indicate significant difference (p < 0.05) between the two compared conditions. %

I

O

W

N

P

V

T

26.37 ± 2.39

14.60 ± 0.48 0.02

15.96 ± 3.93 0.10 0.57

19.78 ± 2.17 0.12 0.03 0.11

13.97 ± 2.63 0.03 0.74 0.52 0.13

13.09 ± 0.87 0.01 0.09 0.35 0.03 0.71

15.07 ± 1.4 0.03 0.59 0.65 0.06 0.44 0.24

20.09 ± 3.22

24.51 ± 2.04 0.04

14.47 ± 3.94 0.30 0.09

24.77 ± 0.83 0.18 0.89 0.03

25.40 ± 4.27 0.34 0.83 0.01 0.79

23.64 ± 0.10 0.20 0.55 0.05 0.12 0.54

15.99 ± 5.42 0.50 0.18 0.25 0.09 0.01 0.13

N. salina p value of t-test

I O W N P V

D. salina p value of t-test

I O W N P V

Table 4 Percentage of unsaturated fatty acids based on the total fatty acids. I – initial medium, W – complete new medium, O – old medium, N – nitrogen deficient medium, P – phosphor deficient medium, V – vitamin deficient medium and T – trace elements deficient medium. Two-tailed Student t test was carried out for paired samples between any two nutrient-deficient conditions. Values in bold indicate the significant difference (p < 0.05). %

I

O

W

N

P

V

T

42.52 ± 2.32

47.76 ± 1.76 0.06

42.19 ± 3.62 0.88 0.19

43.11 ± 2.00 0.72 0.15 0.43

44.91 ± 1.96 0.43 0.22 0.45 0.48

45.44 ± 1.02 0.07 0.17 0.21 0.13 0.79

43.95 ± 0.57 0.40 0.10 0.42 0.42 0.54 0.12

42.50 ± 0.66

42.90 ± 1.3 0.53

46.12 ± 4.94 0.38 0.46

40.41 ± 0.23 0.04 0.10 0.18

38.69 ± 4.96 0.36 0.36 0.00 0.60

39.32 ± 0.93 0.07 0.10 0.10 0.15 0.81

45.50 ± 3.66 0.35 0.45 0.50 0.13 0.01 0.06

N. salina p value of t-test

I O W N P V

D. salina p value of t-test

I O W N P V

butyric acid (C4:0), palmitic acid (C16:0), stearic acid (C18:0), oleic acid (C18:1n9c) and c-linolenic acid (C18:3n6) are the five major fatty acids, taking up around 11.2%, 18.5%, 10.4%, 11.1% and 18.6%, respectively, of total FA under N condition. In N. salina, compared to the initial culture (I) cells under all the conditions at later stages tended to produce longer fatty acids, as butyric acid (C4:0) proportion decreased. Besides, saturated FAs (C4:0 and C16:0) did not change much over time but unsaturated FAs (C16:1 and C18:1n9c) increased slightly. Compared to the new medium (W), cells in the depleted media (except T) presented a slight increase in the two unsaturated FAs. In D. salina, the changes weren’t large, but an increase in C18:0 was noted for the deplete conditions (except T) compared to the replete medium (W), and a slight decrease in C4:0 and an increase in C18:1n9c was noted for T compared to the replete medium (W). The profile of fatty acids in the two tested species was in agreement with other studies. In N. salina, palmitic acid (C16:0), palmitoleic acid (C16:1), and eicosapentaenoic acid (C20:5) were observed to be the three major fatty acids in the lipids, accounting for 32.1%, 26%, and 15.7% of the total fatty acids, respectively [36]. In Dunaliella species, those C16 and C18 fatty acids were also the major fatty acids, as studied by Zhukova and Aizdaicher [53]. For other species like Scenedesmus obliquus CNW-N, under nitrogen starvation, the microalgal lipids were also primarily composed of

C16 and C18 fatty acids (around 90%) [33]. This has been pointed out to be suitable for biodiesel synthesis [33]. Moreover, it was found that long chain mono-unsaturated fatty acids (C16:1, C18:1) have positive impact on cetane number. This indicates that N. salina is more suitable for biodiesel production than D. salina [51]. To a certain extent, the influence of environmental conditions on the distribution of FAs in an algal species can be explained by the roles of FAs in the cellular structure and metabolism. A better understanding of the roles of FAs may enable our prediction on the potential influence caused by a change in the environment. Locations of various fatty acids can be found in chloroplast, membrane and thylakoids, etc. [54,55]. In eukaryotic algae and vascular plants, most of the chloroplast fatty acids are polyenoic acids [54]. Amongst these polyenoic acids, linolenic acid (C18:3), almost exclusively present in chloroplasts, is the major constituent in the leaf tissues of higher plant [54]. High content of pigments in D. salina indicates the richness in chloroplasts and might be the reason that it expresses a higher level of c-linolenic acid (C18:3n6) than N. salina. Palmitic acid (C16:0) has been found to be predominant in both plasma membranes and microsomes [55]. Therefore, it is a primary fatty acid in both species. In contrast, palmitoleic acid (C16:1) was only detected in N. salina. A several-fold increase in palmitoleic acid was observed in tobacco

70

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

Fig. 5. Biplot (scores and loadings) of PCA analysis on the GC–FID spectra of fatty acids, showing clustering of the lipids profile information associated with different nutrientdeficient conditions. The first letter indicates the species (N for N. salina and D for D. salina); the second letter indicates the nutrient conditions: I – initial medium, W – complete new medium, O – old medium, N – nitrogen deficient medium, P – phosphor deficient medium, V – vitamin deficient medium and T – trace elements deficient medium; the suffix numbers indicate the biological replicate. Numbers in the centre loadings plot indicate the fatty acid type that contributes to the variation in the groups (refer Table 2).

Fig. 6. Principal fatty acids detected in N. salina (a) and D. salina (b) under different nutrient deficient conditions. I – initial medium, W – complete new medium, O – old medium, N – nitrogen deficient medium, P – phosphor deficient medium, V – vitamin deficient medium and T – trace elements deficient medium.

when a rat or a yeast gene for the enzyme stearoyl-CoA desaturase was introduced [56,57]. It was also found that a high-palmitoleic acid phenotype of sunflower mutant was associated with a concerted reduction in the fatty acid synthase II activity and an increase of stearoyl-ACP desaturase activity with respect to a high-palmitate mutant [58]. Therefore, it can be deduced that the enzyme stearoyl-CoA desaturase is only present or more active at least in N. salina. Fatty acid elongation appears to be taking place in N. salina under all the conditions compared to the initial cultures (decrease in %SCFA seen in Fig. 5a) but this is not the case in D. salina. Synthesis of fatty acid initiates with the condensation of acetyl CoA and malonyl CoA catalysed by the microsomal enzyme b-ketoacylCoA synthase (Kcs) to yield b-ketoacyl-CoA and CO2. This is the first and also the major rate-limiting reaction triggering the elongation of fatty acid by sequential addition of C2 units to acyls of at least C12 [59]. The Kcs enzymes might have been additionally activated under all the nutrient-stressed conditions in N. salina. However, the Kcs enzymes in D. salina appear to be irrelevant in the

formation of very long chain fatty acids (scarcely present in the algal membranes) but significantly contribute to those comparatively shorter chain acyl-CoA substrates [60]. From this experiment, we can see that the extreme deficiency of nutrients had remarkable but distinct influence on physiology of the two studied species, including biomass production, biochemical composition as well as lipid profile. The influence on biomass production was significant for both species. However, significant influence, on the basis of statistical analysis, on both lipid productivity and fatty acid composition was only observed in N. salina. In a sense, the role and function of different fatty acids in cellular structure and metabolism explain the difference in lipid classes of the two species. 4. Conclusions In this investigation, we show that for microalgal lipid production cell harvest at the stationary phase is desirable, and an active growth phase harvest is desirable for biomass or growth related

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

products. Dilution ratio hardly affected the biomass and lipid productivities but affected the harvest period. Amongst nutrient deficient conditions tested, nitrogen deficiency resulted in the strongest inhibition of cell growth such that the total lipid productivity was not the highest for both N. salina and D. salina, grown photoautotrophically at atmospheric CO2 levels. For N. salina, a higher lipid productivity was reached in T or P condition, whilst for D. salina, the nutrient replete (New) and T conditions produced higher lipid productivities. Lipid profiles showed remarkable difference of fatty acids between the two species. D. salina showed a higher level of c-linolenic acid (C18:3n6), almost exclusively present in chloroplasts. Fatty acid elongation was observed in N. salina under all the conditions but this was not significant in D. salina. The presence of palmitoleic acid in only N. salina implies that the enzyme stearoyl-CoA desaturase is present or is more active in N. salina compared to D. salina. The high lipid content and high level of long chain mono-unsaturated fatty acids (C16:1, C18:1) in N. salina suggests it be a more promising biodiesel producer than D. salina. We believe these observations to be novel and will help us in enhancing our knowledge of nutrient status towards the accumulation of lipids or biomass-dependent products in the two microalgal species, N. salina and D. salina. Acknowledgements We gratefully acknowledge Chinese Scholarship Council (CSC) scheme, Xiamen Blue Environment Tech. Co., Ltd., Special programs of Marine Public Welfare Profession (201405038) and ChELSI (EPSRC EP/E036252/1) for the funding support that made this work possible. References [1] Borowitzka MA, Moheimani NR. Sustainable biofuels from algae. Mitig Adapt Strat Glob Change 2013;18:13–25. [2] Mata TM, Martins AA, Caetano NS. Microalgae for biodiesel production and other applications: a review. Renew Sustain Energy Rev 2010;14:217–32. [3] Demirbas A, Demirbas MF. Importance of algae oil as a source of biodiesel. Energy Convers Manage 2011;52:163–70. [4] Kiran B, Kumar R, Deshmukh D. Perspectives of microalgal biofuels as a renewable source of energy. Energy Convers Manage 2014;88:1228–44. [5] Cade-Menun BJ, Paytan A. Nutrient temperature and light stress alter phosphorus and carbon forms in culture-grown algae. Mar Chem 2010;121:27–36. [6] Chen M, Tang H, Ma H, Holland TC, Ng K, Salley SO. Effect of nutrients on growth and lipid accumulation in the green algae Dunaliella tertiolecta. Bioresour Technol 2011;102:1649–55. [7] Gonçalves AL, Simões M, Pires JCM. The effect of light supply on microalgal growth, CO2 uptake and nutrient removal from wastewater. Energy Convers Manage 2014;85:530–6. [8] Griffiths MJ, van Hille RP, Harrison ST. Lipid productivity, settling potential and fatty acid profile of 11 microalgal species grown under nitrogen replete and limited conditions. J Appl Phycol 2012;24:989–1001. [9] Davis RW, Volponi JV, Jones HD, Carvalho BJ, Wu H, Singh S. Multiplex fluorometric assessment of nutrient limitation as a strategy for enhanced lipid enrichment and harvesting of Neochloris oleoabundans. Biotechnol Bioeng 2012;109:2503–12. [10] Roleda MY, Slocombe SP, Leakey RJG, Day JG, Bell EM, Stanley MS. Effects of temperature and nutrient regimes on biomass and lipid production by six oleaginous microalgae in batch culture employing a two-phase cultivation strategy. Bioresour Technol 2012;129:439–49. [11] Samorì G, Samorì C, Guerrini F, Pistocchi R. Growth and nitrogen removal capacity of Desmodesmus communis and of a natural microalgae consortium in a batch culture system in view of urban wastewater treatment (Part I). Water Res 2012;47:791–801. [12] Tercero EAR, Sforza E, Morandini M, Bertucco A. Cultivation of Chlorella protothecoides with urban wastewater in continuous photobioreactor: biomass productivity and nutrient removal. Appl Biochem Biotechnol 2014;172: 1470–85. [13] Ashokkumar V, Agila E, Sivakumar P, Salam Z, Rengasamy R, Ani FN. Optimization and characterization of biodiesel production from microalgae Botryococcus grown at semi-continuous system. Energy Convers Manage 2014;88:936–46. [14] Richardson B, Orcutt D, Schwertner H, Martinez CL, Wickline HE. Effects of nitrogen limitation on the growth and composition of unicellular algae in continuous culture. Appl Microbiol 1969;18:245–50.

71

[15] Hsieh CH, Wu WT. Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresour Technol 2009;100:3921–6. [16] Dragone G, Fernandes BD, Abreu AP, Vicente AA, Teixeira JA. Nutrient limitation as a strategy for increasing starch accumulation in microalgae. Appl Energy 2011;88:3331–5. [17] Xia L, Yang H, He Q, Hu C. Physiological responses of freshwater oleaginous microalgae Desmodesmus sp. NMX451 under nitrogen deficiency and alkaline pH-induced lipid accumulation. J Appl Phycol 2014;27:649–59. [18] Brennan L, Owende P. Biofuels from microalgae—a review of technologies for production, processing, and extractions of biofuels and co-products. Renew Sustain Energy Rev 2010;14:557–77. [19] Xin L, Hong-ying H, Ke G, Ying-xue S. Effects of different nitrogen and phosphorus concentrations on the growth, nutrient uptake, and lipid accumulation of a freshwater microalga Scenedesmus sp. Bioresour Technol 2010;101:5494–500. [20] Simionato D, Sforza E, Carpinelli E Corteggiani, Bertucco A, Giacometti GM, Morosinotto T. Acclimation of Nannochloropsis gaditana to different illumination regimes: effects on lipids accumulation. Bioresour Technol 2011;102:6026–32. [21] Pal D, Khozin-Goldberg I, Cohen Z, Boussiba S. The effect of light, salinity, and nitrogen availability on lipid production by Nannochloropsis sp.. Appl Microbiol Biotechnol 2011;90:1429–41. [22] Teo SH, Taufiq-Yap YH, Ng FL. Alumina supported/unsupported mixed oxides of Ca and Mg as heterogeneous catalysts for transesterification of Nannochloropsis sp. microalga’s oil. Energy Convers Manage 2014;88:1193–9. [23] Hamidi N, Yanuhar U, Wardana ING. Potential and properties of marine microalgae Nannochloropsis oculata as biomass fuel feedstock. Int J Energy Environ Eng 2014;5:279–90. [24] Mitra M, Patidar SK, George B, Shah F, Mishra S. A euryhaline Nannochloropsis gaditana with potential for nutraceutical (EPA) and biodiesel production. Algal Res 2015;8:161–7. [25] Zhang X, Tang X, Zhou B, Hu S, Wang Y. Effect of enhanced UV-B radiation on photosynthetic characteristics of marine microalgae Dunaliella salina (Chlorophyta, Chlorophyceae). J Exp Mar Biol Ecol 2015;469:27–35. [26] Chen X-J, Wu M-J, Jiang Y, Yang Y, Yan Y-B. Dunaliella salina Hsp90 is halotolerant. Int J Biol Macromol 2015;75:418–25. [27] Tavallaie S, Emtyazjoo M, Rostami K, Kosari H, Assadi MM. Comparative studies of b-carotene and protein production from Dunaliella salina isolated from Lake Hoze-soltan, Iran. J Aquatic Food Product Technol 2015;24:79–90. [28] Borowitzka LJ, Borowitzka MA. Commercial production of-carotene by Dunaliella salina in open ponds. Bull Mar Sci 1990;47:244–52. [29] Doan QC, Moheimani NR, Mastrangelo AJ, Lewis DM. Microalgal biomass for bioethanol fermentation: Implications for hypersaline systems with an industrial focus. Biomass Bioenergy 2012;46:79–88. [30] Deniz I, Vardar-Sukan F, Yüksel M, Saglam M, Ballice L, Yesil-Celiktas O. Hydrogen production from marine biomass by hydrothermal gasification. Energy Convers Manage 2015;96:124–30. [31] Chen Y, Vaidynanthan S. Simultaneous assay of pigments, carbohydrates, proteins and lipids in microalgae. Anal Chim Acta 2013;776:31–40. [32] Chen Y, Vaidyanathan S. A simple, reproducible and sensitive spectrophotometric method to estimate microalgal lipids. Anal Chim Acta 2012;724:67–72. [33] Ho S-H, Chen C-Y, Chang J-S. Effect of light intensity and nitrogen starvation on CO2 fixation and lipid/carbohydrate production of an indigenous microalga Scenedesmus obliquus CNW-N. Bioresour Technol 2012;113:244–52. [34] Vandenbrouck T, Jones OAH, Dom N, Griffin JL, De Coen W. Mixtures of similarly acting compounds in Daphnia magna: from gene to metabolite and beyond. Environ Int 2010;36:254–68. [35] Oncel S, Vardar-Sukan F. Photo-bioproduction of hydrogen by Chlamydomonas reinhardtii using a semi-continuous process regime. Int J Hydrogen Energy 2009;34:7592–602. [36] Cai T, Park SY, Racharaks R, Li Y. Cultivation of Nannochloropsis salina using anaerobic digestion effluent as a nutrient source for biofuel production. Appl Energy 2013;108:486–92. [37] Van Wagenen J, Miller TW, Hobbs S, Hook P, Crowe B, Huesemann M. Effects of light and temperature on fatty acid production in Nannochloropsis salina. Energies 2012;5:731–40. [38] Lam MK, Lee KT. Cultivation of Chlorella vulgaris in a pilot-scale sequentialbaffled column photobioreactor for biomass and biodiesel production. Energy Convers Manage 2014;88:399–410. [39] Prieto A, Pedro Cañavate J, García-González M. Assessment of carotenoid production by Dunaliella salina in different culture systems and operation regimes. J Biotechnol 2011;151:180–5. [40] Mussgnug JH, Klassen V, Schlüter A, Kruse O. Microalgae as substrates for fermentative biogas production in a combined biorefinery concept. J Biotechnol 2010;150:51–6. [41] Huang X, Huang Z, Wen W, Yan J. Effects of nitrogen supplementation of the culture medium on the growth, total lipid content and fatty acid profiles of three microalgae (Tetraselmis subcordiformis, Nannochloropsis oculata and Pavlova viridis). J Appl Phycol 2013;25:129–37. [42] Lamers PP, Janssen M, De Vos RC, Bino RJ, Wijffels RH. Carotenoid and fatty acid metabolism in nitrogen-starved Dunaliella salina, a unicellular green microalga. J Biotechnol 2012. [43] Antal TK, Krendeleva TE, Rubin AB. Acclimation of green algae to sulfur deficiency: underlying mechanisms and application for hydrogen production. Appl Microbiol Biotechnol 2011;89:3–15.

72

Y. Chen et al. / Energy Conversion and Management 106 (2015) 61–72

[44] Tan Y, Lin J. Biomass production and fatty acid profile of a Scenedesmus rubescens-like microalga. Bioresour Technol 2011;102:10131–5. [45] Vanucci S, Guerrini F, Milandri A, Pistocchi R. Effects of different levels of Nand P-deficiency on cell yield, okadaic acid, DTX-1, protein and carbohydrate dynamics in the benthic dinoflagellate Prorocentrum lima. Harmful Algae 2010;9:590–9. [46] Wan M, Jin X, Xia J, Rosenberg JN, Yu G, Nie Z, et al. The effect of iron on growth, lipid accumulation, and gene expression profile of the freshwater microalga Chlorella sorokiniana. Appl Microbiol Biotechnol 2014;98:9473–81. [47] Che R, Huang L, Yu X. Enhanced biomass production, lipid yield and sedimentation efficiency by iron ion. Bioresour Technol 2015. [48] Lelong A, Bucciarelli E, Hégaret H, Soudant P. Iron and copper limitations differently affect growth rates and photosynthetic and physiological parameters of the marine diatom Pseudo-nitzschia delicatissima. Limnol Oceanogr 2013;58:613–23. [49] Hsieh SI, Castruita M, Malasarn D, Urzica E, Erde J, Page MD, et al. The proteome of copper, iron, zinc, and manganese micronutrient deficiency in Chlamydomonas reinhardtii. Mol Cell Proteomics 2013;12:65–86. [50] Singh P, Guldhe A, Kumari S, Rawat I, Bux F. Investigation of combined effect of nitrogen, phosphorus and iron on lipid productivity of microalgae Ankistrodesmus falcatus KJ671624 using response surface methodology. Biochem Eng J 2015;94:22–9. [51] Islam MA, Brown RJ, Brooks PR, Jahirul MI, Bockhorn H, Heimann K. Investigation of the effects of the fatty acid profile on fuel properties using a multi-criteria decision analysis. Energy Convers Manage 2015;98:340–7.

[52] Han F, Pei H, Hu W, Song M, Ma G, Pei R. Optimization and lipid production enhancement of microalgae culture by efficiently changing the conditions along with the growth-state. Energy Convers Manage 2015;90: 315–22. [53] Zhukova NV, Aizdaicher NA. Fatty acid composition of 15 species of marine microalgae. Phytochemistry 1995;39:351–6. [54] Kenyon C. Fatty acid composition of unicellular strains of blue-green algae. J Bacteriol 1972;109:827–34. [55] Azachi M, Sadka A, Fisher M, Goldshlag P, Gokhman I, Zamir A. Salt induction of fatty acid elongase and membrane lipid modifications in the extreme halotolerant alga Dunaliella salina. Plant Physiol 2002;129:1320–9. [56] Grayburn WS, Collins GB, Hildebrand DF. Fatty acid alteration by a D9 desaturase in transgenic tobacco tissue. Nat Biotechnol 1992;10:675–8. [57] Polashock JJ, Chin C-K, Martin CE. Expression of the yeast D-9 fatty acid desaturase in Nicotiana tabacum. Plant Physiol 1992;100:894–901. [58] Salas JJ, Martínez-Force E, Garcés R. Biochemical characterization of a highpalmitoleic acid Helianthus annuus mutant. Plant Physiol Biochem 2004;42: 373–81. [59] Millar AA, Kunst L. Very long chain fatty acid biosynthesis is controlled through the expression and specificity of the condensing enzyme. Plant J 2003;12:121–31. [60] Lassner MW, Lardizabal K, Metz JG. A jojoba beta-Ketoacyl-CoA synthase cDNA complements the canola fatty acid elongation mutation in transgenic plants. The Plant Cell Online 1996;8:281–92.