Aquaculture 392–395 (2013) 69–76

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Modulating lipid accumulation and composition in microalgae bybiphasic nitrogen supplementation

Sachitra K. Ratha a, Radha Prasanna a,⁎, Rachapudi B.N. Prasad b, Chandragiri Sarika b,Dolly W. Dhar a, Anil K. Saxena a

a Division of Microbiology, Indian Agricultural Research Institute (IARI), New Delhi 110012, Indiab Centre for Lipid Research, CSIR-Indian Institute of Chemical Technology, Hyderabad 500 007, India

⁎ Corresponding author. Tel.: +91 11 25847649; fax:E-mail address: radhapr@gmail.com (R. Prasanna).

0044-8486/$ – see front matter © 2013 Elsevier B.V. Allhttp://dx.doi.org/10.1016/j.aquaculture.2013.02.004

a b s t r a c t

a r t i c l e i n f o

Article history:Received 19 July 2012Received in revised form 3 February 2013Accepted 4 February 2013Available online 12 February 2013

Keywords:BiomassMicroalgaeNitrogenTotal lipidsα-Linolenic acid

A biphasic nutritional strategy was employed for enhancing lipid accumulation in the three selectedmicroalgae Chlorella sp. MIC-G4, Chlorella sp. MIC-G6 and Scenedesmus sp. MIC-G8, involving supplementationwith different nitrogen sources (NaNO3, KNO3, urea and (NH4)2CO3), followed by their limitation. NaNO3 wasfound to be the best for enhancing lipid accumulation. Palmitic, linoleic and alpha linolenic acids were themost abundant fatty acids. MIC-G6 recorded the highest content of α- linolenic acid (43.7% of total fattyacid) after 32 d growth in nitrogen rich medium and 36.0%, after 24 d growth in nitrogen rich mediumfollowed by 8 d in nitrogen limited medium. An enhancement of 40–50% in lipids on dry weight basis wasrecorded in Chlorella sp. MIC-G4 and Scenedesmus sp. MIC-G8, in the treatment involving 24 d growth innitrogen rich medium, and 8 d in nitrogen limited medium. An abundance of 92–97% (% of total fatty acids)of C16–C18 fatty acids was recorded. Our study illustrates the promise of MIC-G4 and MIC-G8 as a source oflipids, triggered by the biphasic mode of cultivation, and the need to undertake in-depth gene expressionprofiling to enhance further the lipid accumulation.

© 2013 Elsevier B.V. All rights reserved.

1. Introduction

Microalgae have the ability to grow very fast, synthesize and accu-mulate high amounts of lipids which can be modulated by severalbiotic and abiotic factors; hence, attracted a lot of attention for biodieselproduction and as a potential renewable resource for essential fattyacids in the last decade. The lipid content of some microalgal species,which is the most important aspect for biofuel production, is muchhigher (50–60% of dry weight) than that of terrestrial plants, such asrice and soybean (2–20% of dry weight). Oil productivity of microalgalcultures exceed the yield of the best oilseed crops, e.g. 12,000 l ha−1

of biodiesel for microalgae cultivated in open pond production systemas compared with 1190 l ha−1 for rapeseed (Schenk et al., 2009).Most biological systems, including microorganisms, animals, higherplants and insects are capable of de novo synthesis of fatty acids fromacetate to short chain fatty acids, with oleic acid (18:1 ω-9) as themajor product. Oleic acid is further desaturated to form linoleic acid(18:2 ω-6) and further to α-linolenic acid (18:3 ω-3, ALA). These threeparent fatty acids compete with each other for the 6 desaturase to formthe essential fatty acids (EFA) such as EPA (eicosa pentaenoic acid) andDHA (docosahexaenoic acid) which play a significant therapeutic role inthe regulation of physiological functions, especially being essential forthe proper development of brain and nervous system. Recent work has

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also shown their protective role in the prevention of cardiovascular dis-eases (Davis and Kris-Etherton, 2003). Althoughmarine fish oil is consid-ered as the richest source of EPA, it is envisaged that this is obtained viabioaccumulation in the food chain, through consumption of microalgae,which are known to be the primary producers of ω-3 PUFAs (Wen andChen, 2003). Most fungi, algae, bacteria, mosses and some invertebratespossess the desaturase and elongase for the synthesis of long chainPUFAs, but several higher plants and animals lack the requisite enzymes,and thereby rarely contain PUFAs above C18 (Gill and Valivety, 1997).

The accumulation of lipids, especially TAGs (triacylglycerols) inmicroalgae, especially under stress conditions is considered an adap-tive mechanism (Hu et al., 2008; Liu et al., 2012), as these compoundscan serve as energy/carbon compounds, besides being an importantelectron sink. Many microalgae can accumulate lipids due to excessphotosynthesis and some species can accumulate high amount oflipids under heterotrophic or environmental stress, such as nutrientdeficiency or salt stress (Takagi et al., 2000). Most reports on enhanc-ing lipid accumulation have demonstrated that nitrogen limitationconditions lead to enhanced lipid content, but this is generally con-comitant with lower biomass productivity, i.e. significantly lowerlipid productivity. In order to avoid this problem, a two stage “fatten-ing cultivation strategy” has been proposed (Li et al., 2008; Rodolfi etal., 2009). This involves growth under nutrient rich conditions leadingto high levels of biomass concentration, followed by triggering lipidaccumulation by shifting to a “fattening” medium (specific nutrientN/P depleted). Breur et al. (2012) were able to demonstrate the

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significance of selecting suitable microalgal strains for TAGs accumula-tion through screening of available germplasm and using nitrogen star-vation as a trigger.

The present study focused on the lipid accumulating potential ofindigenous microalgae from selected extreme habitats of India, ana-lyzing the effect of different sources of nitrogen and their limitation atdifferent stages of growth, as a strategy to enhance lipid productivityof three selected microalgae, Chlorella sp. MIC-G4, Scenedesmus sp.MIC-G8, and Chlorella sp. MIC-G6, and also focus towards identifyingpromising α-linolenic producers.

2. Materials and methods

2.1. Microalgae and their growth conditions

A set of ten microalgal isolates were observed under microscope(Zeiss Model Axio Scope. A1) and identified with the help of standardtaxonomic keys (Komárek and Fott, 1983; Prescott, 1962). The detailsof the taxonomic enumeration of isolated microalgae with theirhabitats are presented in Table 1. These isolates were obtained fromdifferent ecological regions of India (details of locations sampledgiven as Supplementary Table 1; Ratha et al. (2012)). Prior to use, thestock cultures were grown in Bold's Basal medium (Starr and Zeikus,1993) at pH 6.6 and incubated at 25±2 °C under a photoperiod of16:8 h light dark cycle at light intensity of 33 μmol photon m−2 s−1

PAR and hand shaken two to three times daily (without sparging withair or CO2), to avoid adherence to the sides of the flasks.

2.2. Screening of microalgal strains

A time course study was carried out for the ten microalgal species,listed in Table 1, along with their place of collection, which werescreened in terms of their growth rate, chlorophyll, carbohydrates andtotal lipid content, as well as biomass productivity. The experimentwas carried out in Erlenmeyer flasks of 100 ml capacity, containing40 ml modified Bold's Basal medium, under the optimal growth condi-tions, as described above. For all experiments, 10% inoculumwas addedfrom two weeks old cultures having Abs750=0.2. Growth rate wasmonitored bymeasuring the changes of turbidity in the culturemedium

Table 1Taxonomic enumeration of isolated microalgae and details of habitat.

Name of organism Place of isolation Habitat D

Chlorococcum sp. MIC-G1 Leh, Jammu and Kashmir Epilithic in pool CChlorococcum sp. MIC-G2 Guwahati, Assam Epipelic in lake C

toch

Chlorococcum sp. MIC-G3 Rohtang Pass,Himachal Pradesh

Planktic in lake Cchzo3

Chlorella sp. MIC-G4 Rohtang Pass, Himachal Pradesh Planktic and epipelicin stream

Cce

Chlorella sp. MIC-G5 Dal Lake, Jammuand Kashmir

Planktic in lake CS

Chlorella sp. MIC-G6 Chilka Lake, Odisha Planktic in lake Cp

Kirchneria sp. MIC-G7 Chilka Lake, Odisha Planktic in lake Coch

Scenedesmus sp. MIC-G8 Dal Lake, Jammu and Kashmir Planktic in lake Cap

Chlamydomonas sp. MIC-G9 Umian Lake, Barapani, Meghalaya Planktic in lake Cce

Ulothrix sp. MIC-G10 Bahuda river, Odisha Free floating Fv1sp

(Absorbance at 750 nm: Abs750) by using a UV–vis spectrophotometer(Perkin Elmer model Lambda) and dry cell weight (DCW) at threedays interval up to 32 days. All the experiments and analyses werecarried out using triplicate samples.

2.3. Effect of nitrogen sources

Out of the ten isolates, two — Chlorella sp. MIC-G4, Chlorellasp. MIC-G6 and one Scenedesmus sp. MIC-G8 were selected on thebasis of their high growth rate and total lipid content for further exper-imentation. Four treatments, in Bold's Basal medium, were evaluated,which included T1 — NaNO3 (2.94 mM), equimolar concentration ofKNO3 (T2), urea (T3) and ammonium bicarbonate (T4) respectively.

2.4. Nutrient limitation experiment

The cells from 8th, 16th and 24th day old cultures grown in thedifferent nitrogen sources were centrifuged at 3000 g for 10 min. Cellpellets were re-suspended in modified Bold's basal medium, withoutnitrogen and analyzed on 8th day. The treatments are denoted as NL1T1/T2/T3/T4–8 d with nitrogen+8 d without nitrogen; NL2 T1/T2/T3/T4–16 d with nitrogen+8 d without nitrogen; NL3 T1/T2/T3/T4–24 dwith nitrogen+8 d without nitrogen. The dry biomass, chlorophyll,carbohydrates and lipid content were measured.

2.5. Estimation of dry weight

Growth rate was determined by measuring the absorbance ofsamples at 750 nm (Abs750). Biomass concentration was calculatedby multiplying Abs750 values with predetermined conversion factorconverting the Abs750 value to dry cell weight (DCW). The conversionfactorwas established by plotting Abs750 versus DCWof a series of sam-ples, from a broad absorbance range of 0.2 to 2.8. Dry cell weight (DCW)was determined gravimetrically using a known amount of algal cultureafter centrifugation at 5000 g for 10 min. The algal pellet was washedtwice with distilled water, and then the harvested biomass was driedat 75 °C in an oven until it reached a constant weight. The specificgrowth rate (μ) was calculated by the following equation:

escription

ells spherical; solitary, colonial, chloroplast hollow sphere; cells 8–11 μm in diameter.ells spherical, ensheathed, solitary or number of cells crowedgether to form stratum; cells 22.3–68.5 μm diameter; sheath 6–9 μm;loroplast hollow sphere, pyrenoids single, prominent.ells spherical, solitary or number of cells crowded together to form a stratum;loroplast parietal with central prominent pyrenoids; cells 14.4–22.5 μm in diameter;ospore mother cell thick walled, 21–29 μm in diameter; spores elliptical,.2–4.7 μm broad, 2.2–4.4 μm long.ells spherical, solitary with thin cell wall; chloroplast parietal with distinct,ntral pyrenoids; cells 2.3–5.2 μm in diameter.ells spherical, solitary with thin cell wall; chloroplast parietal; cells 2.6–7.6 μm broad;pore mother cells 11.4–14.5 μm diameter.ells spherical, solitary with thin cell wall; chloroplast parietal with distinct,yrenoids not cleared, cells 4.3–7.2 μm in diameter.ells solitary, slightly sigmoid or semicircular, broadly lunate, ends bluntly pointed ends,uter side convex, cells 2.4–4.3 μm broad, 6.8–14.6 μm long; cell wall smooth, hyline,loroplast parietal, without pyrenoid; Reproduction by autospores.oenobia 2–4 celled; arranged in liner to sub alternating series, outer cells lunate,pices attenuated; cells 4.2–6.2 μm broad, 14.2–37.5 μm long; chloroplast single,arietal with central prominent pyrenoids.ells elliptical, motile, biflagellated; chloroplast basal cup shaped with single pyrenoids;lls 4.8–7.2 μm broad, 7.8–12.6 μm long.ilaments uniseriate; light green, cells quadrate or cylindrical or ellipsoidal,arying in dimension, terminal cell rounded; septa stratified, cells 9–12 μm broad,1–28 μm long; chloroplast parietal, plate like, with conspicuous pyrenoids; spores,herical or elliptical, 8–12 μm in diameter.

Table 2Evaluation of chlorophylls, carbohydrates and lipids (%, on dry cell weight basis) of tenmicroalgal isolates (n=3)a.

Name of isolate Chlorophyll (%) Carbohydrates (%) Lipids (%)

Chlorococcum sp. MIC-G1 2.56cd±0.10 17.86c±0.61 11.83c±0.65Chlorococcum sp. MIC-G2 3.06ab±0.20 29.03c±0.64 10.46d±0.86Chlorococcum sp. MIC-G3 2.76abcd±0.08 39.23b±0.13 8.43e±0.65Chlorella sp. MIC-G4 2.53cd±0.27 28.66c±0.24 14.16b±0.21Chlorella sp. MIC-G5 3.10a±0.07 25.16d±0.27 13.20b±0.27Chlorella sp. MIC-G6 2.60cd±0.16 37.60b±0.40 18.13a±0.37Kirchneria sp. MIC-G7 2.56cd±0.02 66.4a±0.35 10.96cd±0.50Scenedesmus sp. MIC-G8 2.46d±0.34 30.85c±0.46 13.50b±0.44Chlamydomonas sp. MIC-G9 2.66abcd±0.23 24.40d±2.95 11.83c±0.97Ulothrix sp. MIC-G10 2.95abc±0.16 19.46e±1.50 10.43d±0.76SEM 0.08 0.44 0.22C.D. (0.05) 0.22 1.21 0.61

a Mean n=3 replicates of algal cultures; SEM: standard error of mean; differentletters in each column indicate significant differences at 5% probability level.

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Ln C/C0=μδt; where C0 is the initial biomass concentration (g l−1)and C is the biomass concentration (g l−1) at any time t (Yeesang andCheirsilp, 2011).

Lipids (%) were expressed in terms of dry cell weight (DCW). Toestimate chlorophyll, 10 ml of algal culture was centrifuged at3000 g for 10 min and the pellet was treated with known volume ofmethanol and kept in water bath for 30 min at 60 °C, absorbance ofthe pooled extracts was measured at 652 and 665 nm and chlorophyll(a+b) was estimated using standard equations (Lichtenthaler, 1987).The total carbohydrate content was measured spectrophotometrically,according to the method of Spiro (1966), using D-glucose as a standard.Chlorophyll and carbohydrates are expressed (%), in terms of dry cellweight (DCW).

2.6. Extraction of lipid from algal biomass

The total lipids were extracted from microalgal biomass using amodified method of Dittmer and Wells (1969), using a mixture ofchloroform and methanol (2:1, v/v). The chloroform and aqueousmethanol layers were separated by the addition of methanol andwater to give a final solvent ratio of chloroform: methanol: water(2:1:0.8). The chloroform layer containing the lipids was washed with1% NaCl solution, collected and evaporated to dryness under vacuum.The lipid content was determined gravimetrically. All the experimentswere carried in triplicate.

2.7. FAME analysis

The dried biomass (0.25 g) was taken in a 50 ml flask and 15 ml of2% sulfuric acid in methanol solution was added and the contentsrefluxed for 4 h. The contents were diluted with water and the organicphase was extracted with ethyl acetate (3×25 ml). The ethyl acetatephase was thoroughly washed with water and dried over anhydroussodium sulfate. Ethyl acetate was evaporated on a rotary evaporatorto recover fatty acid methyl esters (0.12 g).

Gas Chromatographic (GC) analysis was performed using the fattyacid methyl esters on an Agilent 6890 series gas chromatograph using1 μl injections, flame ionization detector (FID), equippedwith a DB-225capillary column (30 m, 0.25 mm i.d. 0.5 mm). The column tempera-ture was initially maintained at 160 °C for 2 min, increased to 230 °Cat 5 °C min−1 and held for 20 min. The injector and detector tempera-tures were maintained at 230 and 270 °C, respectively with a split ratioof 50:1. Nitrogen was used as carrier gas with flow rate of 1.0 ml/min.This GC method allows for quantification of hydrocarbons, free fattyacids, fatty acid methyl esters in a single analysis. To identify individualfree fatty acids and hydrocarbons of the cultures, the GC–MS analysiswas performed with Agilent 6890 gas chromatograph connected toAgilent 5973 Nmass selective detector at 70 ev (m/z 50–600; sourceat 230 °C and quadruple at 150 °C) in the EI mode with a HP-1 mscapillary column (30 m×0.25 mm×0.5 μm). The oven temperaturewas initially maintained at 160 °C for 2 min, increased to 230 °C at5 °C min−1 and held for 20 min. Helium was used as carrier gaswith flow rate of 1.0 ml/min. the inlet pressure was maintained at230 °C and split ratio was 50:1. The composition and relative abun-dance of the different fatty acids were determined using standards.GC with mass spectrometry as described above was used for freefatty acids, fatty acid methyl esters identification.

2.8. Statistical analyses

Experiments for the determination of dry weight, growth rates, aswell as the biochemical assays were all repeated three times to ensurethe clarity of the data. The statistical analyses were performed usingthe software SPSS 10. One-way analysis of variance (ANOVA) wasused to evaluate the differences among the treatments. If ANOVAeffects were significant, comparisons between the different means

were made using post hoc least significant differences (LSD). Standarddeviation of values and error bar shown in the figures were determinedby number of replicates (3) and the probability values were less than0.05.

3. Results

3.1. Screening of microalgae on the basis of growth attributes andlipid accumulation

A set of ten microalgal isolates, belonging to five genera (three spe-cies of each of Chlorella and Chlorococcum, one each of Scenedesmus,Chlamydomonas, Kirchneria and Ulothrix) were screened under standardlaboratory conditions for their growth rate, and biochemical composi-tion. The summary of chlorophyll, carbohydrate and lipid percentage(on dry weight basis) is given in Table 2. All the species were initiallygrown under NaNO3 as nitrogen source in Bold's Basal medium, to iden-tify the promising strains in relation to biomass and lipid production. Theresults show that the total chlorophyll content was highest in Chlorellasp. MIC-G5 (3.10%) and lowest in Scenedesmus sp. MIC-G8 (2.46%),whereas the highest carbohydrate content was recorded in Kirchneriasp. MIC-G7 (66.4%) and lowest in Chlamydomonas sp. MIC-G9 (24.4%),after 32 days of cultivation. The highest accumulation of lipids of 18%was recorded in Chlorella sp. MIC-G6, followed by Chlorella sp. MIC-G4(14%) and Scenedesmus sp. MIC-G8 (13.48%), which were selected forfurther analyses.

3.2. Growth kinetics of microalgae

Fig. 1 show the growth kinetics of all the ten isolates, Chlorococcumsp. MIC-G1, Chlorococcum sp. MIC-G2, Chlorococcum sp. MIC-G3,Chlorella sp.MIC-G4, Chlorella sp.MIC-G5, Chlorella sp.MIC-G6,Kirchneriasp. MIC-G7, Scenedesmus sp. MIC-G8, Chlamydomonas sp. MIC-G9, andUlothrix sp. MIC-G10 grown in normal strength of nitrogen in Bold'sBasal medium. The curves representing the variation in the biomassconcentration over time indicates a very short lag or adaptation phase,followed by exponential growth (24 d) and stationary phase.

3.3. Effect of different sources of nitrogen

The growth kinetics of all the three microalgae under four nitrogensources, are shown in Supplementary Table 2 and Fig. 2. As shown inFig. 2(A), the cell growth of Chlorella sp. MIC-G4 showed a significantenhancement up to 20th day, on all the sources of nitrogen, but onlyNaNO3, followed by KNO3 supported growth in the post exponentialphase. The highest dry biomass (0.53 g l−1) was obtained in culturesfed with NaNO3 on 32nd day. NaNO3 was the only N source supportiveof growth up to 32 d of incubation. Fig. 2(B) depicts the growth rate of

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Fig. 1. The growth kinetics of microalgae in Bold's basal medium. Chlorococcum sp.MIC-G1, Chlorococcum sp. MIC-G2, Chlorococcum sp. MIC-G3, Chlorella sp. MIC-G4,Chlorella sp. MIC-G5, Chlorella sp. MIC-G6, Kirchneria sp. MIC-G7, Scenedesmus sp.MIC-G8, Chlamydomonas sp. MIC-G9, Ulothrix sp. MIC-G10.

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Chlorella sp. MIC-G6, in which ammonium bicarbonate supported thebest growth followed by urea, NaNO3 and KNO3 up to 16 days ofincubation. The highest value of 0.43 g l−1 was achieved in the cellsfed with NaNO3 on 32nd day. In Scenedesmus sp. MIC-G8, all the four

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Fig. 2. Effect of different nitrogen sources, NaNO3 (T1), KNO3 (T2), Urea (T3) and(NH4)2CO3 (T4) on growth kinetics of three microalgae, (A) Chlorella sp. MIC-G4,(B) Chlorella sp. MIC-G6 and (C) Scenedesmus sp. MIC-G8 during 32 days of study. Meann=3 replicates.

nitrogen sources supported growth up to 32 days, but cells fed withsodium nitrate (Fig. 2(C)) recorded the highest biomass of 0.62 g l−1

dry biomass. Therefore, on an overall basis, NaNO3, followed by KNO3

were more proficient in enhancing growth.Chlorella sp. MIC-G4 showed a specific growth rate of 0.211 d−1

which corresponds to a doubling time of 4.7 d; this isolate reachedthe stationary phase with a DCW of 0.53 g l−1. Lipid content slightlyincreased in stationary phase and values of 14% lipids was recorded(Fig. 3). Chlorella sp. MIC-G6 showed a growth rate of 0.234 d, corre-sponding to doubling time of 4.3 d and the maximum DCW obtainedwas 0.433 g l−1 with 18% lipid. Scenedesmus sp. MIC-G8 showed agrowth rate of 0.461 d and doubling time 2.2 d; the maximum con-centration of lipid with respect to dry biomass was 13%. The brackishwater species, Chlorella sp. MIC-G6, was found to be the highest lipidproducer (18% DCW) under normal laboratory conditions.

The time course profiles in terms of chlorophyll content of cells incultures with different nitrogen sources also showed a progressiveincrease with period of incubation. A doubling in the level of carbohy-drates was recorded in Chlorella sp. MIC-G4 on 32d of incubation. Apositive correlation was recorded between the accumulation ofcarbohydrates and chlorophyll content (Supplementary Table 3), inall the treatments (T1, T2, T3 and T4).

3.4. Effect of nitrogen limitation

The impact of biphasic strategy, involving nitrogen limitation onlipid production is shown in Fig. 4. In terms of chlorophyll, NL3recorded significantly lower values in Chlorella sp. MIC-G4, while inChlorella sp. MIC-G6, no significant differences were recorded in theNL1, NL2 or NL3 treatments. However, lipid % (based on DCW)showed a gradual enhancement with period of incubation in nitrogensupplemented medium. Chlorella sp. MIC-G4 (Fig. 4(A)) exhibited thehighest lipid content in the NL3 treatment involving growth for 24 dayswith nitrogen supplementation, followed by 8 d in nitrogendeficientme-dium. The lipid content in Chlorella sp. MIC-G4 in the various N sources,after NL3 treatment were — T1 (28%), T2 (18%), T3 (21%) and T4 (16%)respectively (Table 3; control values given in parentheses correspond tothe same culture periods, but without nitrogen limitation). Fig 4(B) and(C) illustrate the effect of nitrogen limitation on % lipid of Chlorella sp.MIC-G6 and Scenedesmus sp. MIC-G8, in which a similar trend wasrecorded. The values were much lower in MIC-G8. In terms of chloro-phyll, NL3 recorded significantly lower values in Chlorella sp. MIC-G4(Table 3), while Chlorella sp. MIC-G6 was not significantly affected bynitrogen sources, both in terms of chlorophyll or % lipids.

FAME profiles generated revealed a high percentage of C16-C18(92.6–96.8%) fatty acids (Table 4). Palmitic acid (C16:0) ranged from19.5 to 27.3%, while linoleic acid (18:2) ranged from 31.1 and 32.1% inMIC-G4, 5.2 and 6.5 in MIC-G6 and 26.7 and 29.7% in MIC-G8.α-Linolenic acid (ALA) ranged from14.5 to 43.7%. In general, saturatedfatty acids ranged from30.2 to 34.9% vis a vis 69.8 to 65.1% of unsaturatedfatty acids. The ratio between unsaturated/saturated ranged from 1.9 to2.3%.

4. Discussion

Microalgae, a group of fast-growing unicellular or simple multi-cellular microorganisms are found in very wide range of environmentsandoffer several advantages, includinghigher photosynthetic efficiency,enhanced growth rates and higher biomass production compared toother energy crops (Li et al., 2008; Rodolfi et al., 2009). Oleaginousalgae produce only small quantities of Triacylglycerols (TAGs) under op-timal growth or favorable environmental conditions (Hu et al., 2008)and their metabolism can bemanipulated through simplemodificationsof the chemical composition of the culture medium (Behrens and Kyle,1996). Under stress conditions (chemical stimuli, physical stimuli),microalgae alter their lipid production and accumulate more amount

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Fig. 3. Effect of different sources of nitrogen, NaNO3 (T1), KNO3 (T2), Urea (T3), Ammonium bicarbonate (T4) on lipid accumulation during 32 days of culture. (A) Chlorella sp.MIC-G4, (B) Chlorella sp. MIC-G6, and (C) Scenedesmus sp. MIC-G8. Data expressed as percent dry cell weight (DCW). Mean n=3 replicates.

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of TAGs. Themajor chemical stimuli are nutrient starvation, salinity andgrowth-medium pH, while the major physical stimuli are temperatureand light intensity (Breur et al., 2012; Hu et al., 2008). The lipid contentincreased from 18% to 40% (in terms of dry cell weight) in Chlorellavulgaris, when grown in nitrogen deprivation medium (Illman et al.,2000). The possible reason for this enhancement is the diversion ofnutrients for cell growth in the first stage, followed by biomass enrich-ment with lipids, in the absence of nutrient(s) in the second stage (Linand Lin, 2011). Recent reports (Breur et al., 2012; Li et al., 2012; Liuet al., 2012; Msanne et al., 2012) have analyzed the metabolic changesoccurring during such stress regimes, especially for nitrogen starvation,and illustrated the interdependence of catabolic and anabolic processes,besides the more significant role of carbohydrate metabolism overphotosynthesis in TAG accumulation in microalgae.

Microalgal oil content can seldom exceed 80% by weight of dry bio-mass (Chisti, 2007).Most of the commercially employed algae (Chlorella,Crypthecodinium, Cylindrotheca, Dunaliella, Isochrysis, Nannochloris,Nannochloropsis, Neochloris, Nitzschia, Phaeodactylum, Porphyridium,Schizochytrium, Tetraselmis) have oil levels between 20 and 50%, buthigher productivities can be obtained under certain stress conditions—nutrients, temperature and light (Mata et al., 2010; Schenk et al.,2009). Another alternative is to tap the innate capacity of microalgae,by screening a wide range of habitats and improving the potential ofpromising isolates by different methods, including genetic engineering.Although the lipid content of algal species will remain consistent pro-vided it is grown under the same set of conditions, every algal specieshas its own lipid profile which generally remains constant whengrown under a specified set of conditions. Therefore, strain selection is

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NL1 NL2 NL3

Fig. 4. Influence of different sources of nitrogen and their limitation on lipid accumulation.(A) Chlorella sp. MIC-G4, (B) Chlorella sp. MIC-G6, (C) Scenedesmus sp. MIC-G8. NL1— 8 dnitrogen source+8 d without nitrogen; NL2 — 16 d nitrogen source+8 d withoutnitrogen;NL3— 24 d nitrogen source+8 dwithout nitrogen. C1— 16 dN supplemented;C2— 24 dN supplemented and C3— 32 dN supplemented. Data expressed as percent drycell weight (DCW). Mean n=3 replicates.

Table 3Influence of different sources of nitrogen and their deficiency on lipid accumulation(% on dry cell weight basis).

Treatments T1(NaNO3) T2 (KNO3) T3(Urea) T4 (NH4CO3)

MIC-G4 Chlorella sp.NL1(C1) 14 (11) 11 (11) 13 (9) 10 (10)NL2 (C2) 15 (12) 13 (9) 15 (8) 14 (8)NL3 (C3) 28 (14) 18 (11) 21 (9) 16 (9)SEM 0.192 0.176 0.351 0.401C.D. (0.05) 0.532 0.487 0.972 1.111

MIC-G6 Chlorella sp.NL1 (C1) 14 (11) 12 (8) 10 (9) 14 (8)NL2 (C2) 12 (14) 12 (11) 11 (10) 12 (10)NL3 (C3) 25 (18) 12 (14) 18 (12) 17 (12)SEM 0.778 0.451 0.703 0.588C.D. (0.05) 2.156 1.249 1.948 1.629

MIC-G8 Scenedesmus sp.NL1 (C1) 12 (11) 12 (9) 10 (8) 14 (8)NL2 (C2) 16 (12) 14 (10) 15 (8) 16 (7)NL3 (C3) 27 (13) 20 (10) 15 (10) 17 (8)SEM 1.179 0.801 0.754 0.940C.D. (0.05) 3.267 2.219 2.089 2.605

NL1 — 8 d nitrogen source+8 d without nitrogen; NL2 — 16 d nitrogen source+8 dwithout nitrogen; NL3 — 24 d nitrogen source+8 d without nitrogen. C1 — 16 d Nsupplemented; C2 — 24 d N supplemented and C3 — 32 d N supplemented. Dataexpressed as percentage of dry cell weight. Mean n=3 replicates.

Table 4Qualitative analysis of FAME profiles, in terms of fatty acids (% of total fatty acids).

Fattyacid

MIC-G4(+N)

MIC-G4(−N)

MIC-G6(+N)

MIC-G6(−N)

MIC-G8(+N)

MIC-G8(−N)

12:0 0.5 0.5 1.3 1.6 1.3 1.114:0 1.2 1.9 0.9 0.9 0.6 0.515:0 0.1 0.1 0.4 0.3 0.2 0.316:0 20.3 19.5 25.2 27.3 23.2 22.316:1 1.7 1.6 3.3 2.5 1.7 3.316:2 2.8 2.5 4.1 3.1 2.3 2.316:3 11.5 11.4 0.5 0.4 12.6 9.517:0 6.4 6.7 ND ND 6.7 6.218:0 1.4 1.1 1.1 1.8 1.4 3.318:1 3.6 3.1 11.2 15.0 3.8 5.318:2 31.1 32.1 5.2 6.5 29.7 26.718:3 18.0 18.0 43.7 36.0 14.5 15.520:0 0.2 0.1 0.3 0.5 0.2 0.520:2 0.8 0.8 1.0 2.0 1.4 1.622:0 0.1 0.1 1.0 1.1 0.1 0.722:1 0.2 0.3 0.3 0.3 0.3 0.924:0 0.1 0.2 0.5 0.7 ND NDC16–C18 96.8 96.0 94.3 92.6 95.9 94.4SFAa 30.3 30.2 30.7 34.2 33.7 34.9MUFAb 5.5 5.0 14.8 17.8 5.8 9.5PUFAc 64.2 64.8 54.5 48.0 60.5 55.6TLd 14 28 18 25 13 27USF:SFAe 2.3 2.3 2.3 1.9 2.0 1.9

−N denotes (+N) 24 d+(−N) 8 d; +N, 32 d (+N).a SFA, saturated fatty acids.b MUFA, monounsaturated fatty acids.c PUFA—polyunsaturated fatty acids.d TL—total lipids.e USF:SFA—ratio of unsaturated and saturated fatty acids; ND not detected.

74 S.K. Ratha et al. / Aquaculture 392–395 (2013) 69–76

critical for biodiesel production. Under favorable growth conditions,algae synthesize fatty acids principally for esterification into glycerol-based polar lipids (e.g., glycolipids and phospholipids), the major con-stituents of intracellularmembranes. Under unfavorable environmentalor stress conditions, however, many algae alter their lipid biosyntheticpathways for the formation and accumulation of neutral lipids, about20–50% (of cell dry weight), mainly in the form of TAGs, along withsmall amounts of monoacylglycerols (MAGs), diacylglycerols (DAGs)and sterol esters, typically stored in cytosolic lipid bodies (Hu et al.,

2008). Neutral lipids can be readily converted to biodiesel or othertypes of fuels through the existing and emerging oil refining processes(Hu et al., 2008).

A set of ten microalgal isolates were evaluated in terms of theirgrowth kinetics, which is known to influence the lipid accumulationin microalgal cells. Observations revealed that three isolates Chlorellasp. MIC-G4, Chlorella sp. MIC-G6 and Scenedesmus sp. MIC-G8 werethe top three ranking strains, with 13.48–18% lipids (in terms ofDCW). These three strains were compared for their performance inmedia supplemented with different nitrogen sources (T1 — NaNO3,

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T2 — KNO3, T3 — urea and T4 — (NH4)2CO3). Among the nitrogensources, NaNO3 was found to be the best for enhancing lipid accumu-lation and sustained growth up to 32d. Earlier studies with Neochlorisoleoabundans by Li et al. (2008) showed that the highest lipid cellcontent of 40% was obtained with NaNO3 (3 mM), while in ourstudy, NaNO3 supplementation at the rate of 2.94 mM concentrationled to highest lipid percentage in all the three microalgae evaluated.Interestingly, chlorophyll values also showed a steady increase upto16–20d in all the nitrogen sources, and in NaNO3 supplementedcultures, growth was sustained upto the 32nd day. Nitrate and ureaare known to be better than ammoniacal sources of nitrogen forgrowth and lipid accumulation in selected microalgae, includingNeochloris sp. and Chlorella sp. (Li et al., 2008).

Lourenco et al. (2002) observed that the role of nitrogen starvationin enhancing lipid accumulation needs to be based on selection of suit-able nitrogen source. Navarro-Pérez et al. (2001) observed that lipidcontent tends to increase when cultures reach stationary phase, espe-cially when silicate and nitrogen become limiting, mainly in diatoms.Rebolloso-Fuentes et al. (2001) found that long lasting growth stagesled to increased PUFA levels. Amongmicroalgae, the Eustigmatophyceaemembers were found to be the best producers of ALA. In our study,Chlorella sp. (MIC-G6) proved to be a promising source of ALA, althoughunder nitrogen deprivation condition, there was a reduction in ALA by9%. However, this can be compensated by the 7% enhancement in totallipids under nitrogen deprivation conditions.

In order to fulfil the economical viability of biodiesel projections,there is a need to build up biomass, which can provide high rates oflipid productivity. It was first reported by Rodolfi et al. (2009) that inorder to increase lipid content and lipid productivity, a two phase culti-vation process (a nutrient sufficient phase to produce high algae biomassfollowed by a nitrogen deprived phase to boost lipid synthesis) was asuitable strategy. It has been postulated that the deficiency of nutrientsleads to cessation of growth and the organism channels the metabolicflux generated in photosynthesis to lipid / fatty acid biosynthesis.Microalgae also known to accumulate lipids as a storage product(Behrens and Kyle, 1996; Chisti, 2007).

Nitrogen is the most commonly reported nutritional limiting fac-tor triggering lipid accumulation (Breur et al., 2012; Courchesne etal., 2009; Li et al., 2012; Lin and Lin, 2011; Liu et al., 2012; Msanneet al., 2012). Certain researchers suggest that cells accumulate largequantities of chlorophyll molecules when nitrogen source is abun-dantly available, and under conditions of exhaustion of nutrients,the cells utilize chlorophylls as a nitrogen source (Li et al., 2008;Rodolfi et al., 2009). In Botryococcus braunii, oil content can reach75% by weight of dry biomass, but it is associated with low productiv-ities due to slow growth rates (Dayananda et al., 2007). Nitrogen lim-itation is known to bring about three changes: decrease of the cellularcontent of thylakoid membrane, activation of acyl hydrolase andstimulation of the phospholipids hydrolysis (Mandal and Mallick,2009) or activation of diacylglycerol acyltransferase, which convertsacyl-CoA to triglyceride (Takagi et al., 2000). Recent reports showedthat most of the ESTs in global expression profiles of N starvedgreen alga Micractinium pusillum were involved in carbohydratemetabolism, Pyruvate and acetyl CoA synthesis etc. (Li et al., 2012).

In our studies, the nitrogen deprived culture, after a nitrogensufficient phase, exhibited a relatively high biomass of 0.5 g l−1 andduring the N deprivation, an increase in lipid content from 14% to28% in Chlorella sp. MIC-G4, 18% to 25% in Chlorella sp. MIC-G6 and13 to 27% in Scenedesmus sp. MIC-G8 respectively was recorded. Anenhancement in lipid percentage (16–28% higher than their respec-tive controls) on dry weight basis was recorded in NL3 condition,involving 24 d growth in nitrogen rich medium, followed by 8 d innitrogen limited medium. In general, C16 and C18 fatty acid residueswere dominant accompanied by a high degree of unsaturation. It hasbeen reported that each microalga gears its biosynthetic pathwaytowards a preferential fatty acid (Guedes et al., 2011) and similar

observations were recorded in our study. Guedes et al. (2011) foundthat Nannochloropsis sp., belonging to Eumastigmatophyceae was theprominent ALA producer in a set of 27 microalgal cultures screenedand deprivation of nitrogen strongly influenced the FAME profiles.

Lipid accumulation in Scenedesmus obliquus was studied undervarious culture conditions, and a significant increase in lipid up to43% and 30% DCW under N-deficiency and P-deficiency respectivelywas recorded, as against 12.7% under control condition (Mandal andMallick, 2009). In our study, N deficiency led to an enhancement of40–50% over control in treatment involving 24 d growth in NaNO3

followed by 8 d under N-limitation. The quantitative analysis of thelipids through generation of FAME profiles revealed that 92–96% ofthe fatty acids belonged to C16–C18 type with USFA/SFA ratio beingin the range of 1.9–2.3% in the strains. N limitation exhibited a differ-ential effect, with MIC-4 recording an enhancement, while otherstrains showed reduced values. The abundance of saturated fatty acidsis of extreme significance for consideration as a source of biodiesel,because such oils have higher cetane number (CN), decreased NOxemissions, a shorter ignition delay time and oxidative stability(Antolin et al., 2002). Highest quantities of saturated fatty acids wererecorded in MIC-6, which showed as 3.5% enhancement underN-limitation. Interestingly, lignoceric acid (24:0) was recorded inMIC-G4 and MIC G6, which was enhanced from 0.1 to 0.2% and from0.5 to 0.7% in the two strains respectively, under N limitation. Thisfatty acid is very common in fish and peanut oil (1.1–1.7%) and itspresence is a positive quality trait for the algal oil in our cultures. Nosignificant changes in C16–C18, saturated or PUFA was recordedunder N limitation.

The predominance of palmitic acid (16:1) ranging from 20 to 27%is a common feature in several algae, which can be as high as 49–50%(Yeesang and Cheirsilp, 2011). Other researchers have pointed outthe predominance of palmitic acid, oleic acids and linoleic acid inBotryococcus sp. This is similar to those found in plant oils (Fang et al.,2004). Another aspect of significance is that in MIC-G6 and MIC-G8,N-limitation led to a significant reduction in PUFA, a concurrentincrease in SFA which makes them more promising candidates for indepth research as biofuel sources.

Microalgae are known to show a decline in photosynthetic perfor-mance and nitrogen deprived cells undergo physiological changesthat reduce the efficiency of light harvesting energy transductionand CO2 fixation (Berges et al., 1996). But in our study, no significantdecrease in the growth attributes such as chlorophyll was recorded inNL1 and NL2 treatments, and Scenedesmus sp. MIC-G8 was not affectedeven in the NL3 treatments with all different N sources. This illustratesthe potential of the “fattening strategy” in enhancing lipid productivityin the strains being investigated, which has now been further strength-ened by molecular analyses involving global gene expression profiles(Li et al., 2012). A combination of nitrogen deficiency, high levels ofiron and light intensity was shown to enhance lipid accumulation by15–35% in four microalgae (Yeesang and Cheirsilp, 2011).

The limited success of algal lipid production stems primarily fromthe lack of understanding of metabolic pathways and the regulationof algal lipid metabolism in general and neutral lipid synthesis andaccumulation in particular, and failure to develop cost-affordableand energy-efficient algal feedstock production systems and processes(Davis and Kris-Etherton, 2003; Hsieh and Wu, 2009; Li et al., 2008;Xu et al., 2001). Lipid productivity is generally reduced in comparisonwith nitrogen sufficient growth; however, in our study, a significantenhancement in lipid productivity vis a vis nutrient limitation wasclearly evident. Our earlier study using carbon precursors/aminoacids etc. have shown that enhancement of lipids, based on ourhypothesis that in the presence of sodium thiosulfate, diversion of Cskeletons to fatty acid synthesis occurs (Ngangkham et al., 2012).This is strengthened by the observations of Li et al. (2012) who dem-onstrated by differential gene expression analyses that the transcriptlevels of genes related to pyruvate and acetyl-CoA are dramatically

76 S.K. Ratha et al. / Aquaculture 392–395 (2013) 69–76

increased after N starvation and therefore may play a key role in driv-ing carbon flow into TAG biosynthesis. Hence, metabolic flux analysescan be a useful tool to understand flux distribution and manipulateAcetyl CoA/Malonyl CoA reaction for enhanced lipid synthesis.

The major obstacle for microalgal oil production is their relativelyhigh costs, which may be overcome by the technology developments.Optimization of process is a very important aspect for such technologydevelopment (Schenk et al., 2009). Our study clearly highlighted adifferential response of the algal species to sources of nitrogen,which was variable both in terms of growth and lipid accumulation.A novel aspect of our study was that the microalga Chlorella sp.MIC-G6 exhibited very high content of ALA, which is comparable tothat in flax seeds, in which 57% ALA has been recorded (Davis andKris-Etherton, 2003). Further, metabolic engineering and a deeperunderstanding of the interactions of anabolic and catabolic pathwaysin this organism can further enhance the lipid content in thisorganism.

5. Conclusions

A biphasic nutritional strategy was employed for enhancing lipidaccumulation in three selected microalgae — Chlorella sp. MIC-G4,Chlorella sp. MIC-G6 and Scenedesmus sp. MIC-G8. The lipid content ofChlorella sp. MIC-G4 and Scenedesmus sp. MIC-G8 were significantlyenhanced from ~40 to 50% over their controls by this strategy, involvingN limitation. Enhanced lipid production was recorded in NaNO3

followed by ammoniumbicarbonate. In depth analyses of FAMEprofilesrevealed the predominance of palmitic, linoleic and linolenic acids and92.6%–96.8% C16–C18 fatty acids. MIC-G6 was identified as a promisingALA producer, which warrants in depth studies and upscaling for itspromise as a nutraceutical.

Acknowledgments

The authors are grateful to the Division of Microbiology, IndianAgricultural Research Institute (IARI), New Delhi and the IndianCouncil of Agricultural Research for providing the facilities and financialsupport, to undertake the investigations.

Appendix A. Supplementary data

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.aquaculture.2013.02.004.

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