Effect of Nitrogen Source in Low-Cost Media on

Biomass and Lipid Productivity of Nannochloropsis

salina for Large-Scale Biodiesel ProductionJunying Liua and Krys BangertbaDepartment of Chemical and Biological Engineering, University of Sheffield, Sheffield, United Kingdom; liuhailinshef@hotmail.com(for correspondence)bDepartment of Molecular Biology and Biotechnology, University of Sheffield, Firth Court, Western Bank, Sheffield, UnitedKingdom

Published online 00 Month 2014 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/ep.11967

A major cost component for large-scale algal cultivation isthe medium. In this study, a significantly higher growth ratewas found for Nannochloropsis salina in an inexpensivemedium of seawater and agricultural fertilizer than wasreached in the f/2 medium. A key difference between thisnew low-cost medium and the f/2 medium is the nitrogensource. In the conditions tested herein, with aeration usingair, biomass productivity reached 0.5496 g L21 day21 after10 days in the low-cost medium compared with 0.1215 gL21 day21 in the f/2 medium. The lipid productivity of algaegrown in the low-cost medium was also higher than that inthe f/2 medium (0.1281 g L21 day21 versus 0.0432 g L21

day21). The strong linear correlation (R2 5 0.9721) betweenchlorophyll content and biomass concentration demonstratedthat this new medium can support healthy and consistentgrowth of algal cells. The results suggest that it would be pref-erable to feed the algae for large-scale biodiesel productionwith agricultural waste stream or untreated swine waste-water containing high levels of ammonia rather than treatedeffluent in which the ammonia/urea has been oxidized tonitrate. VC 2014 American Institute of Chemical Engineers Environ

Prog, 00: 000–000, 2014

Keywords: Nannochloropsis salina, agricultural fertilizer,seawater, lipid productivity, algal biodiesel

INTRODUCTION

Algae are an attractive option for a biodiesel feedstockdue to high biomass productivity and lipid yield comparedwith the first generation of oil crops [1]. Therefore, it is beingseriously considered for the next generation of biofuels [2].Most studies focus on one of two aspects: attempts to isolateand screen algae with high lipid content and attempts toenhance the lipid content, either by genetic modification ofthe rate-limiting enzyme or by environmental stress factors,such as nitrogen starvation and light inhibition [3]. Microal-gae with high oil content are desired for producing biodie-sel. Moreover, the selection of the most adequateindustrialized species needs to take into account other fac-tors, such as the growth rate [4]. Many studies have been

conducted using a genetic engineering approach to enhancethe lipid accumulation in different species [5].

Although little progress appears to have been made onthe mechanism of reactions and the structures of some keyenzymes, scientists have uncovered a trade-off betweengrowth rate and lipid content [6]. Williams and Laurens [2]estimated that an increase in lipid content from 15% to 30%would reduce the growth rate by 50% or more. Therefore,overall lipid production is more complicated than a simpleincrease in lipid content, and enhancing lipid content shouldnot be considered the primary method to increase lipidyields. A key for algal biodiesel production and industrializa-tion is to increase the biomass production [7]. However, themajor obstacle to achieving higher biomass production andmore economical biodiesel production is the high cost ofalgal production [8]. The cost includes the substantial con-sumption of freshwater sources, inorganic nutrients (primar-ily nitrogen and phosphorus), and CO2. Wastewater andseawater (SW) might provide favorable alternatives to fresh-water for growing algae [8]. Wastewater is a relatively goodchoice because it is an economic source of nutrients for algalculture, though it has limitations [9]. Wastewater used in themass cultivation of microalgae also presents biological prob-lems, including contaminating organisms such as bacteria,protozoa, fungi, and other algae [10], and nonbiologicalproblems, including adjustment of element concentrations byadding chemicals [11] and the removal of relatively highheavy-metal concentrations [12]. It is impractical to sterilizethe very large volumes of water needed in mass cultivationof microalgae.

Therefore, the costs associated with the medium are oneof the major considerations in large-scale algal production.The common nitrogen of the medium is nitrate becausenitrate is much more stable than other nitrogen sources (e.g.,urea and ammonia) and is easy to measure and analyze(Table 1). Moreover, the standard f/2 culture medium forNannochloropsis species is extremely expensive. One solu-tion for overcoming the high cost of algal cultivation is toreplace the f/2 medium with a mixture of di-ammoniumphosphate and ammonia or agricultural fertilizer containingthe optimal nitrogen and phosphorus content for cellgrowth. The Aquatic Species Program has attempted outdoorVC 2014 American Institute of Chemical Engineers

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cultivation projects using fertilizer that was not designed forNannochloropsis, but the green algae could not grow consis-tently [25].

Carbon is one of the three most important nutrients influ-encing algal growth; therefore, the source and concentrationof carbon greatly affect the biomass productivity and lipidcontent of algae [26]. The necessary CO2 aeration dependson the algal growth system and the specific conditions of thewastewater from the power plant. Whether the carbonsource is flue gas or liquid, the cost of adding carbon mustbe included within the cost of algae production. Therefore,the wastewater utilization and CO2 supply is circumstance-specific [11].

Cultivating algae by using SW and agricultural fertilizershas the potential to reduce the cost and make algal biodieselcommercially viable. The present study examined the possi-bility of reducing the cost of cultivating algae by the follow-ing methods: using a fertilizer to replace nutrients in the f/2medium, using SW to replace freshwater, and aerating withair but not CO2. In addition, urea was added to the batchcultures in given intervals because the big advantage of ureais the low cost in comparison with sodium nitrate, and moreimportantly the combination with nitrate is more optimal foralgal growth [27]. Nannochloropsis salina is a picoplanktongenus of the marine environment with a small cell (2–4 lmin diameter), a spherical to slightly ovoid shape, and is non-flagellate [26]. The genus N. salina was used in this studybecause it has the best combination of biomass productivityand lipid content (about 30%) in comparison with other algalbiodiesel producers, attaining 60% lipid content after nitro-gen starvation [6]. The experiments were designed to investi-gate the effects of different nitrogen sources (mixed andisolated) and feeding rates of urea.

MATERIALS AND METHODS

Algal Strain CultureN. salina was grown in the f/2 medium for 2 weeks

before transferring to the new low-cost medium as an inocu-lum. The inoculum was grown in different media after reach-ing the late exponential phase. A total of 250 mL cultureswere grown in 340 mL cylindrical glass bottles (12 cm length,5 cm diameter, three replicates) under 24 h of light providedby cool white fluorescent tubes with a photon flux of 70mmol photons m22 s21 at the culture surface at 25�C. Therewere film-lids on the flasks with ventilation holes to allowthe aeration of the cultures while to prevent cross-contamination and to minimize evaporation. Aeration with a

flow rate of 0.3 mL min21 was provided after filtrationthrough a 0.22 mm gas filter. Cultures were aerated by twopumps with 24-way proportioning valve and 3-mm PVCpipes at the bottom of the bottle for 12 h per day.

Experimental Design and Culture MediaThe f/2 medium was prepared with distilled water using

previously described methods [28]. Natural SW from Liver-pool Crosby Beach (Liverpool, UK) was used in this study.The characteristics of the SW (a salinity of 31 g L21) andagricultural fertilizer (Feed&Gro) from local stores are shownin Table 2. The fertilizer (NP) was dissolved in the distilledwater. The SW became very clear after 14 days’ standing.Based on the results of a pre-experiment which was per-formed immediately before this study, the effect of urea oncell growth was investigated here. A stock solution of urea(8.3 mM) was added to cultures according to experimentaldesign (Table 3). The concentration of HCO3

2 solution frombottle sparkling water was 3.5 mM, and its feeding mode isshown in Table 3. The employment of the fed-batch processcan be the best option for this system, which can eliminatethe toxicity of the whole nutrient requirement for a batchprocess from the start. The total amount of NP additiondecreased while the amount of urea addition increased fromS-1 to S-6. N. salina was grown in the lab defined mediumof f/2 as a control. Although an exhaustive investigation ofthe many possible combinations and concentration is beyondthe scope of this study, a manageable set of alternative nitro-gen supply scenarios is chosen above.

Relationship Between Dry Weight and OD680

Cell concentrations were measured every day as opticaldensities (OD) by a Thermo UV–vis spectrophotometer atthe absorbance of 680 nm (OD680) and were plotted versusdry weigh (DW g L21) on a standard curve. Each samplewas diluted to give an absorbance in the range 0.1–1.0 (ifOD was greater than 1.0). The standard curves were pre-pared as follows: 3 mL, 6 mL, 9 mL, 12 mL, 15 mL, 18 mL, 21mL, and 24 mL of microalgal culture with OD of 1.0 werediluted into 24 mL (three replicates), respectively. Microalgalcells were harvested by centrifugation and washed twicewith distilled water at 3000g for 10 min. The sample pelletswere freeze-dried for 2 days, after which the total dry weightwas determined. A linear regression relationship wasobtained between OD and DW. The linear regression wasanalyzed by SPSS 19. The cell density (Y mg L21) was calcu-lated using Eq. (1) for the new low-cost medium and Eq. (2)for the f/2 medium,

Table 1. Nitrogen sources and concentrations for different species.

Medium species Nitrogen source Nitrogen (g L21) References

f/2 medium Nannochloropsis sp. NaNO3 0.012 [4]Dunaliella Medium Dunaliella salina CCAP 19/18 NaNO3 0.070 [13]TAP medium Chlamydomonas reinhardtii NH4Cl 0.098 [14]Zarrouk medium Spirulina platensis NaNO3 0.412 [15]Chu medium — NaNO3 0.014 [16]Walne medium Cyclotella sp. NaNO3 0.016 [4]Walne medium Chlorella sp. Urea 0.100 [17]Fitzgerald medium Chlorella vulgaris NaNO3 0.070 [18]AF6 medium Chlorella vulgaris NaNO3 0.023 [19]Bristol medium Neochloris oleabundans NaNO3 0.041 [20]Erdschreiber Dunaliella tertiolecta NaNO3 0.198 [21]Basal medium Chlorella vulgaris NaNO3 0.041 [22]WC medium — NaNO3 0.014 [23]Wastwater Botryococcus braunii NH4

1, NO32 0.01 [24]

Agricultural fertilizer Nannochloropsis salina NH41, NO3

2 0.549 Current research

Environmental Progress & Sustainable Energy (Vol.00, No.00) DOI 10.1002/ep2 Month 2014

Y 50:3156OD 68020:0396; R250:9750 ðP < 0:001Þ (1)

Y 50:3194OD 68020:0314; R250:9966 P < 0:001ð Þ (2)

Determination of Chlorophyll Content and LipidContent

The chlorophyll content was determined using previouslydescribed methods [29]. The final chlorophyll content YC ðlgchlorophyll ml21Þ was calculated as the weighted average oftwo absorbances (645 nm and 663 nm) using the followingequation,

YC5ð202OD645180:2OD663Þ=4 (3)

Analysis of Specific Growth Rate, Biomass, and LipidProductivity

Growth rate of N. salina was calculated as:

l5ln ðXt=X0Þ=ðt2t0Þ (4)

where Xt and X0 represent cell densities (i.e., dry weight) at theend and beginning of the exponential growth phase, respec-tively. Biomass and lipid productivity during 10 days were calcu-lated in this study. The specific growth rate (m day21) and DWof the 10th day were used to calculate biomass productivity (Pb,g L21 day21) with the following equation [30]:

Pb5DW l (5)

Lipid content (L %) and biomass productivity (Pb g L21

day21) were used to calculate the lipid productivity (Pl , gL21 day21) based on the following equation:

Pl5PbL (6)

The lipid content was analyzed using gravimetric methodsas described by Bligh and Dyer [31]. First, lyophilized algalcells were added to 500 mL of chloroform/methanol (2:1, v/v)and sonicated for 1 min on ice. The supernatant was collectedafter centrifugation at 3000g for 10 min, and the volume wasthen estimated to adjust the ratio of chloroform, methanol,and NaCl water. The mixture of the three solvents was centri-fuged to the separated organic phase. The chloroform layerwas collected and dried in a fume hood to constant weight.The total lipid content was calculated gravimetrically.

Analysis of Seawater and Nutrient Concentrationsin Fertilizer

Salinity was measured using a refractometer according tothe manufacturer’s instructions. The concentrations of

Table 2. Characteristics of seawater, urea, sparkling water, and fertilizer (NP) ingredients.

Nutrients Seawater Fertilizer Sparkling water Urea solution

pH 8.0 — 7.0 —Salinity (g L21) 31 — — —HCO3

2 (mM) — — 3.5 —Ammonia nitrogen (mM) 0.078 12 — —Nitrate-nitrogen (mM) 0.018 — <0.016 —Ureic nitrogen (mM) 5.1 — 8.3Total phosphate (mM) 0.090 12 — —Potassium oxide (mM) 9 — —Trace element (mM) — — — —

Table 3. Feeding modes of fertilizer, urea, and HCO32 for Nannochloropsis salina.

Treatments Basal medium (NP/mL) NP addition* Urea addition** HCO32 addition

S-1 2 1 mL/2 days 0 0S-2 1 1 mL/2 days 0.5 mL/day 5 0S-3 1 0.5 mL/day 0.5 mL/day 0S-4 1 1 mL/day 5 1 mL/2 days 0S-5 1 0 0.5 mL/day 0S-6 1 0 1 mL/2 days 1 mL/dayf/2 0 0 0 0

*Characteristics of the fertilizer, urea, and HCO32 were shown in Table 2.

**The detailed ammonia content in the culture over time seen Figure 1.

Figure 1. Biomass growth of Nannochloropsis salina culti-vated in seawater under various nutrient regimes. The f/2medium served as the control. Nutrient regimes are pre-sented in Table 3. [Color figure can be viewed in the onlineissue, which is available at wileyonlinelibrary.com.]

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ammonium and phosphate in the new low-cost mediumwere analyzed using an API test kit in a DR2800 spectropho-tometer (Hach Lange Company). The standard curves ofammonia nitrogen concentration were established by Ness-ler’s reagent spectrophotometry [32] and the phosphate con-tent by the bismuth phosphomolybdate blue photometricmethod. Nitrate and nitrite were analyzed using a HachLange test kit (LCK340).

FTIR AnalysisTo estimate how major organic compounds functional

groups (lipids, protein, and carbohydrate) in algal cells shiftedunder different nutrient conditions, Fourier transforms infraredspectroscopy (FTIR) was conducted using a ShimadzuIRPrestige-21 Fourier Transformation Infrared Spectrophotome-ter (Shimadzu, UK). The sample and related spectra were ana-lyzed [33]. Principal component analysis (PCA) was performedto compare an entire spectral region of 1800–900 cm21 usingXLSTAT software [34]. Statistical data analysis was carried outusing PASW 19.0 software to run ANOVA (Duncan-tests).

RESULTS

Effects of Ammonium Content on Algal Growth andLipid Content

The initial cell density of N. salina in the seven treatmentswas �0.1 at OD680. The biomass concentrations (g L21) ofalgae in the f/2 medium and other media were significantlydifferent (P< 0.001). The media with 0.5 mL addition of urea(S-3) reached the maximum growth rate (1.1091 day21) dur-ing the short exponential phase. The ammonium concentra-tions in the S-3 and S-4 treatments were highest (189.24 mg

L21 and 226.56 mg L21), whereas the ammonium content inthe S-2 and S-1 groups were the lowest (60.22 mg L21 and18.24 mg L21; Figure 1). The cell growth rates changed onday 7. The S-3 and S-4 group reached the stationary phase.The ammonium content of the S-4 group rose to 625.00 mgL21 after 9 days. The treatments with lower ammonium con-tent had higher growth rates during this phase. Biomass con-centrations of all groups were significantly different from thecontrol (df 5 6, F 5 70.968, P< 0.001). The S-2 and S-1groups reached biomass concentrations of 0.754, and 0.608 gL21, respectively. The highest biomass concentration reachedwas 3.5 times higher than that of the f/2 medium. The dryweight increased with the increasing total NP generally whiledecreased with the increasing urea (Figure 2).

Effect of Ammonium Content and Liquid CarbonSource on Algal Productivity

The specific growth rate, biomass productivity, lipid con-tent, and lipid productivity at the end of the experiment areshown in Table 4. A significant inverse relationship betweenthe growth rate and lipid content was not observed. The cul-ture with the highest ammonium content (S-4) had the low-est specific growth rate but also a relatively low lipid content(22.5%) and then the lowest biomass productivity (0.0671 gL21 day21). The highest lipid content (35.6%) was obtainedin the f/2 medium, which was 56% higher than the averageof the new low-cost medium (22.8%). However, the f/2

Figure 2. Effects of total NP, urea, and HCO3 on dry weight.

Table 4. Specific growth rate (m), biomass productivity (Pb), lipid content (L), and lipid productivity (PL) in fed-batch cultures.

Treatments m (day21) Pb (g L21 day21) L (g g21) PL (g L21 day21)

S-1 0.8007bc 0.4022** 0.215b 0.0850S-2 0.8894b 0.5496** 0.233b 0.1281*S-3 1.1091a 0.1981* 0.235b 0.0471S-4 0.7096cd 0.0671 0.225b 0.0152S-5 0.9038b 0.2768* 0.187b 0.0529S-6 0.9409b 0.3881** 0.272ab 0.1066*f/2 0.6234d 0.1215 0.356a 0.0432

Significant difference at 0.05 level for Duncan test.*The mean difference is significant at the 0.05 level.**The mean difference is significant at the 0.001 level.

Figure 3. Relationship between cell density and chlorophyllcontent of Nannochloropsis salina under different feedingregimes. [Color figure can be viewed in the online issue,which is available at wileyonlinelibrary.com.]

Environmental Progress & Sustainable Energy (Vol.00, No.00) DOI 10.1002/ep4 Month 2014

medium had the second-lowest biomass productivity (0.1215g L21 day21). The treatment (S-6) with a liquid carbonsource (HCO3

2) had the second highest lipid content andshowed a significant difference from S-4. The difference inlipid content of these treatments was confirmed by the PCAof FTIR spectra below.

The treatments (S-2 and S-1) with relatively low ammo-nium content had high biomass productivity (0.5496 g L21

day21 and 0.4022 g L21 day21, respectively), irrespective ofthe type of growth media. Statistically significant differencesin biomass productivity were observed between the fertilizertreatments and the f/2 medium (df 5 6, F 5 51.448, P< 0.001).Furthermore, the S-2 group demonstrated the highest lipidproduction, even though it had the median lipid content. TheS-6 treatment had the second highest lipid productivity. Whencompared with cells in the f/2 medium, both the S-2 and S-6groups had significantly higher lipid productivity (P< 0.05).

Chlorophyll AnalysisThe chlorophyll content was also measured to demon-

strate the effect of fertilizer on cell growth (Figure 3). Thetreatments with low ammonium content had a very high cor-relation with cell density (dry weight) (R 5 94–97%). Signifi-cant differences in the chlorophyll content of groups wereobserved at the end of the experiment (t 5 5.534, df 5 6,P 5 0.001).

Chlorophyll concentration in S-2 treatment increasedalmost fivefold and reached its highest value (22.5 mg L21).The chlorophyll content was strongly correlated with biomassconcentration (dry weight) (R2 5 0.9721), providing directexperimental evidence that the doubling of cell density causesa doubling of chlorophyll content in the new low-costmedium. This suggests that the combination of SW and inex-pensive agricultural fertilizer can support the rapid growth ofN. salina. However, the treatments with relatively high ammo-nium content and the f/2 medium presented different results;in these cases, the chlorophyll content was only weakly corre-lated with biomass concentration (R2 5 0.634–0.794). The highammonium concentration is speculated to be toxic to algalcells. Moreover, lower correlation for S3, S4, and S5 couldreflect increased presence of heterotrophic microorganisms,including remineralization because the slight white stuff wasobserved at the bottom of the bottle in the later period.

Evaluation of the Change in Cellular Composition byPCA

The major changes in cellular composition for a givennutrient condition were observed using FTIR at the end of

the experiment. The bands were assigned to specific molecu-lar groups upon the basis of biochemical standards in previ-ous studies [33]. They are attributed to a range of vibrationalmodes in lipid CH2 (2926 cm21) and carbonyl (1740 cm21),amide I and II (1649 cm21 and 1545 cm21, respectively), car-bohydrate (1014 cm21), and nucleic acid (1245 cm21) [35].Following the normalization to amide II, PCA was carriedout over the 1800–900 cm21 wavenumber region on the finalsampling day for all 21 samples. The PCA clearly resolvedthe data into two principal components (Figure 4). Loadingplots showed the magnitude of each wavenumber in theseparation of cell composition change at the duration of cellgrowth along the PC and the zero loading on the wavenum-ber is of little amount. The PC loading indicated that alongF1 the separation of treatments of algae can be occurred inthe region of carbohydrate, protein, and lipid. The positiveloadings in these regions represent that that spectra situatedpositively along F1 display relatively higher carbohydrateand lipid level than that of other phases, while along F2, thevariation also occurred within the carbohydrate and proteinregions with more protein (Data not shown). F1 accountedfor 64.52% of the variation, and F2 accounted for 23.49% ofthe variation. Therefore, the plot of the PC scores (88.01%)of F1 versus F2 derived from FTIR spectra is greater than85%, indicating that FTIR is an effective means of determin-ing the major changes in cells.

The f/2 and S-6 treatments both had high lipid contentand were on the positive regions along F1. This result is con-sistent with the gravimetric lipid measurement. Moreover, thetreatments with urea clustered in the high-protein region,indicating that N. salina can utilize urea effectively. A corre-lation analysis was performed between lipid contentextracted by the gravimetric method and the FTIR lipid signal(1740 cm21). A good correlation (R2 5 0.918) was observedbetween the gravimetric lipid content and the FTIR lipid sig-nal: amide II at 1740 cm21 (Figure 5).

DISCUSSION

Major Nutrients for Algal GrowthMany studies have shown that nitrogen is one of the limit-

ing factors for algal growth and nitrogen deprivation dam-ages phycobilisome and slows photosynthesis [36].Therefore, many studies have examined the effectiveness ofnitrogen sources for use in algal cultivation, such as nitrate,ammonium, urea, and wastewater [11,37]. Nitrate is muchmore stable than other nitrogen sources and is then easy toanalyze. There has been relatively little research on fertilizersfor algae as biodiesel feedstock [38]. A fertilizer (primarilyNH4NO3) was used to grow blue-green algae and found thatthe mass culture of Tolypothrix tenuis could reach a densityof 1.63 g L21 [39].

Figure 4. Principal component analysis (PCA) of FTIR spec-tra derived from Nannochloropsis salina cells at the end ofthe experiment in different treatments with two componentsF1 against F2. [Color figure can be viewed in the onlineissue, which is available at wileyonlinelibrary.com.]

Figure 5. Relationship of lipid content and FTIR signal (1740cm21). [Color figure can be viewed in the online issue,which is available at wileyonlinelibrary.com.]

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In this study, N. salina grown in the new low-costmedium containing SW and agricultural fertilizer (nitrogenand phosphorus) grew faster and had higher biomass pro-ductivity than in the f/2 medium. The pre-experiment dem-onstrated that it was the urea nitrogen in the fertilizer thatlimited the growth of N. salina (data not shown). Intensiveurea application can serve as the sole nitrogen source forplant growth [40]. Urea has gained general application inlarge-scale algal cultivation because it is less expensive thanother sources of nitrogen, but its instability at high tempera-tures limits its application [41]. Pustizzi et al. [42] suggestedthat there is a slight advantage to growing algae in a mediumcontaining both inorganic and organic nitrogen comparedwith either source alone because Aureococcus physiologyand photosynthesis are different during growth on a mixtureof urea-N and nitrate. There is little difference in the growthrate for most species grown under comparable environmen-tal conditions using NO3

2, NH41, and urea as nitrogen sour-

ces [43]. However, the high urea concentrations in this studycaused the culture to reach the stationary phase rapidly and,subsequently, the death phase. The accumulation of urea inthe medium was speculated to be a stress to the cells andthus caused the algae to enter the death phase [27]. Mecha-nisms behind toxic effects could be urea are stored in thecell, and intracellular urea transporters are essential for mov-ing urea from the cytosol to storage vacuoles, in order toprevent possible toxicity to the cell [44]. Toxicity happenswhen the urea concentration is beyond the storage ability ofvacuoles. Moreover, high urea addition might bring downthe pH. By contrast, the treatments with small quantities ofurea could reach high cell densities. Therefore, urea plays animportant role of a promoter of cell growth, rather thanrestrictor, in conditions of such low concentrations [17].

Lipid Productivity and Biomass ProductivityThe highest lipid content was obtained in the f/2 medium

with nitrate, whereas the lowest lipid content was observed forthe S-4 group with the highest ammonium content. Other lipidcontents fell into the normal range of lipid content for Nanno-chloropsis species (0.19–0.36 g g21). This pattern possiblyresulted from the different biochemical pathways induced bydifferent growth media. An increase of the urea concentrationin the nutrient medium led to a decrease of lipid content [17].

The second highest lipid content was achieved by thetreatment providing HCO3

2. The algal growth affects the pH(e.g., becoming more alkaline due to the assimilation of car-bon sources), and the addition of bicarbonate salts can beused to maintain the pH of medium in an optimum range.The lipid productivity under favorable conditions in thisstudy mostly fell within the range of published lipid produc-tivities [45]. Although the lipid content for the S-2 groupreached the median value, the group yielded the higher totallipid productivity because it had the highest growth ratecompared with other treatments in this study as well as pub-lished result [45]. In comparison with other researches, thehigh lipid yield came from the highest biomass but not fromhigh lipid content. Much research has shown that strategiesto increase lipid content can be highly deleterious to cell via-bility and consequently result in significant losses of primaryproduct mass. Therefore, to optimize lipid productivity forbiodiesel, the trade-off between lipid content, growth rate,and cell density needs to be considered [2]. Lipid contenttaken alone is not a suitable criterion for choosing algal spe-cies for biodiesel production because it fails to account forother factors such as growth rate and biomass concentration.

CONCLUSION

Using SW and agricultural fertilizer may provide a power-ful and cost-effective way to produce biodiesel. Different lev-

els of urea feeding frequency were carried out to investigatethe growth rate and lipid content. Media composed of theoptimal combination of natural SW and agricultural fertilizerwith low ammonium concentration significantly increasedthe biomass productivity, and consequently lipid productiv-ity. Ammonia has been demonstrated to be toxic to Nanno-chloropsis species at high concentration. Therefore, a furtherstudy will be carried out to determine the actual nitrogenrequirement of Nannochloropsis species as well as to confirmthe inhibition threshold of urea/ammonia. Ultimately, anoptimization will be required of agricultural fertilizer, thefeeding mode, and the quantity of urea by applying aresponse surface design. Thus, the microalgae lipid produc-tivity can be maximized while lowered the production cost.

ACKNOWLEDGMENTS

The authors thank the China Scholarship Council (CSC)and Carbon Trust (UK) for supporting this research finan-cially. They also thank Dr. Jim Gilmour, Professor Steve Wil-kinson, and Professor Catherine Biggs for FTIR laboratoryassistance.

LITERATURE CITED

1. Chisti, Y. (2007). Biodiesel from microalgae, Biotechnol-ogy Advances, 25, 294–306.

2. Williams, P.J.B. & Laurens, L.M.L. (2010). Microalgae asbiodiesel and biomass feedstocks: Review and analysis ofthe biochemistry, energetics and economics, Energy andEnvironmental Science, 3, 554–590.

3. Pal, D., Khozin-Goldberg, I., Cohen, Z., & Boussiba, S.(2011). The effect of light, salinity, and nitrogen availabil-ity on lipid production by Nannochloropsis sp., AppliedMicrobiology and Biotechnology, 90, 1429–1441.

4. Doan, T.T.Y., Sivaloganathan, B., & Obbard, J.P. (2011).Screening of marine microalgae for biodiesel feedstock,Biomass and Bioenergy, 35, 2534–2544.

5. Schenk, P., Thomas-Hall, S., Stephens, E., Marx, U.,Mussgnug, J., Posten, C., Kruse, O., & Hankamer, B.(2008). Second generation biofuels: High-efficiencymicroalgae for biodiesel production, BioEnergy Research,1, 20–43.

6. Griffiths, M. & Harrison, S. (2009). Lipid productivity as akey characteristic for choosing algal species for biodieselproduction, Journal of Applied Phycology, 21, 493–507.

7. Delrue, F., Setier, P.A., Sahut, C., Cournac, L., Roubaud,A., Peltier, G., & Froment, A.K. (2012). An economic, sus-tainability, and energetic model of biodiesel productionfrom microalgae, Bioresource Technology, 111, 191–200.

8. Savage, N. (2011). Algae: The scum solution, Nature, 474,15–16.

9. Pittman, J.K., Dean, A.P., & Osundeko, O. (2011). Thepotential of sustainable algal biofuel production usingwastewater resources, Bioresource Technology, 102, 17–25.

10. Shimamatsu, H. (2004). Mass production of Spirulina, anedible alga, Hydrobiologia, 512, 39–44.

11. Jiang, L., Luo, S., Fan, X., Yang, Z., & Guo, R. (2011).Biomass and lipid production of marine microalgae usingmunicipal wastewater and high concentration of CO2,Applied Energy, 88, 3336–3341.

12. Shehata, S.A., Lasheen, M.R., Ali, G.H., & Kobbia, I.A.(1999). Toxic effect of certain metals mixture on somephysiological and morphological characteristics of fresh-water algae, Water, Air, and Soil Pollution, 110, 119–135.

13. Kleinegris, D., Es, M.A., Janssen, M., Brandenburg, W.A.,& Wijffels, R.H. (2010). Carotenoid fluorescence in Duna-liella salina, Journal of Applied Phycology, 22, 645–649.

14. Li, Y., Han, D., Hu, G., Dauvillee, D., Sommerfeld, M.,Ball, S., & Hu, Q. (2010). Chlamydomonas starchlessmutant defective in ADP-glucose pyrophosphorylase

Environmental Progress & Sustainable Energy (Vol.00, No.00) DOI 10.1002/ep6 Month 2014

hyper-accumulates triacylglycerol, Metabolic Engineering,12, 387–391.

15. Sydney, E.B., Sturm, W., de Carvalho, J.C., Thomaz-Soccol,V., Larroche, C., Pandey, A., & Soccol, C.R. (2010). Poten-tial carbon dioxide fixation by industrially importantmicroalgae, Bioresource Technology, 101, 5892–5896.

16. Weis, J.J., Madrigal, D.S., & Cardinale, B.J. (2008). Effectsof algal diversity on the production of biomass in homo-geneous and heterogeneous nutrient environments: Amicrocosm experiment, PLoS One, 3, e2825. doi:10.1371/journal.pone.0002825.

17. Hsieh, C.H. & Wu, W.T. (2009). Cultivation of microalgaefor oil production with a cultivation strategy of urea limi-tation, Bioresource Technology, 100, 3921–3926.

18. Widjaja, A., Chien, C.-C., & Ju, Y.-H. (2009). Study ofincreasing lipid production from fresh water microalgaeChlorella vulgaris, Journal of the Taiwan Institute ofChemical Engineers, 40, 13–20.

19. Illman, A.M., Scragg, A.H., & Shales, S.W. (2000). Increasein Chlorella strains calorific values when grown in lownitrogen medium, Enzyme and Microbial Technology, 27,631–635.

20. Gouveia, L., Marques, A., da Silva, T., & Reis, A. (2009).Neochloris oleabundans UTEX #1185: A suitable renew-able lipid source for biofuel production, Journal of Indus-trial Microbiology and Biotechnology, 36, 821–826.

21. Tang, H., Abunasser, N., Garcia, M.E.D., Chen, M., SimonNg, K.Y., & Salley, S.O. (2011). Potential of microalgaeoil from Dunaliella tertiolecta as a feedstock for biodie-sel, Applied Energy, 88, 3324–3330.

22. Converti, A., Casazza, A.A., Ortiz, E.Y., Perego, P., & DelBorghi, M. (2009). Effect of temperature and nitrogenconcentration on the growth and lipid content of Nanno-chloropsis oculata and Chlorella vulgaris for biodieselproduction, Chemical Engineering and Processing: Pro-cess Intensification, 48, 1146–1151.

23. Gamfeldt, L. & Hillebrand, H. (2011). Effects of totalresources, resource ratios, and species richness on algalproductivity and evenness at both metacommunity andlocal scales, PLoS One, 6, e21972. doi: 10.1371/journal.pone.0021972.

24. Sydney, E.B., da Silva, T.E., Tokarski, A., Novak, A.C., deCarvalho, J.C., Woiciecohwski, A.L., Larroche, C., &Soccol, C.R. (2011). Screening of microalgae with poten-tial for biodiesel production and nutrient removal fromtreated domestic sewage, Applied Energy, 88, 3291–3294.

25. Sheehan, J., Dunahay, T., Benemann, J., & Roessler, P.(1998). A look back at the US Department of Energy’saquatic species program-biodiesel from algae, NationalRenewable Energy Laboratory. NREL/TP-580-24190.

26. Richmond, A. (2004). Handbook of microalgal culture,Oxford: Blackwell Publishing.

27. Ren, M. & Ogden, K.L. Cultivation of Nannochloropsisgaditana on mixtures of nitrogen sources, EnvironmentalProgress and Sustainable Energy, doi: 10.1002/ep.11818.

28. Guillard, R.R.L. (1975). Culture of phytoplankton forfeeding marine invertebrates. In W.L. Smith & M.H. Chan-ley (Eds.), Culture of marine invertebrate animals (pp.29–60), New York: Plenum Book Publishing Corporation.

29. Wellbum, A.R. (1994). The spectral determination of chlo-rophyll a and b, as well as total carotenoids, using vari-ous solvents with spectrophotometers of differentresolution, Journal of Plant Physiology, 144, 307–313.

30. Wood, E.R. & Wingard, L.M. (2005). Measuring growthrates in microalgal cultures. In R.A. Andersen (Ed.), Algalculturing techniques (270p), Burlington, MA: ElsevierAcademic Press.

31. Bligh, E.G. & Dyer, W.J. (1959). A rapid method for totallipid extraction and purification, Canadian Journal of Bio-chemistry and Physiology, 37, 911–917.

32. Liu, H.B., Chen, T.H., Chang, D.Y., Chen, D., Liu, Y., He,H. P., Yuan, P., & Frost, R. (2012). Nitrate reduction overnanoscale zero-valent iron prepared by hydrogen reductionof goethite, Materials Chemistry and Physics, 133, 205–211.

33. Liu, J., Mukherjee, J., Hawkes, J.J., & Wilkinson, S.J.(2013). Optimization of lipid production for algal biodie-sel in nitrogen stressed cells of Dunaliella salina usingFTIR analysis, Journal of Chemical Technology and Bio-technology, 88, 1807–1814.

34. Dean, A.P., Sigee, D.C., Estrada, B., & Pittman, K.J.(2010). Using FTIR spectroscopy for rapid determinationof lipid accumulation in response to nitrogen limitationin freshwater microalgae, Bioresource Technology, 101,4499–4507.

35. Murdock, J. & Wetzel, D.L. (2009). FT-IR microspectro-scopy enhances biological and ecological analysis ofalgae, Applied Spectroscopy Reviews, 44, 335–361.

36. Collier, J.L. & Grossman, A.R. (1992). Chlorosis inducedby nutrient deprivation in Synechococcus sp. strain PCC7942: Not all bleaching is the same, Journal of Bacteriol-ogy, 174, 4718–4726.

37. Sinclair, G., Kamykowski, D., & Glibert, P.M. (2009).Growth, uptake, and assimilation of ammonium, nitrate,and urea, by three strains of Karenia brevis grown underlow light, Harmful Algae, 8, 770–780.

38. Ashraf, M., Javaid, M., Rashid, T., Ayub, M., Zafar, A., &Ali, S. (2011). Replacement of expensive pure nutritivemedia with low cost commercial fertilizers for mass cul-ture of freshwater algae, Chlorella vulgaris, InternationalJournal of Agriculture and Biology, 13, 484–490.

39. Anderson, D.B., Molten, P.M., & Metting, B. (1981).Assessment of blue-green algae in substantially reducingnitrogen fertilizer requirements for biomass fuel crops(5p), Richland: Pacific Northwest Laboratory.

40. Witte, C.P. (2011). Urea metabolism in plants, Plant Sci-ence, 180, 431–438.

41. Danesi, E.D.G., de , O., Rangel-Yagui, C., de Carvalho,J.C.M., & Sato, S. (2002). An investigation of effect ofreplacing nitrate by urea in the growth and production ofchlorophyll by Spirulina platensis, Biomass and Bioen-ergy, 23, 261–269.

42. Pustizzi, F., MacIntyre, H., Warner, M.E., & Hutchins, D.A.(2004). Interaction of nitrogen source and light intensityon the growth and photosynthesis of the brown tide algaAureococcus anophagefferens, Harmful Algae, 3, 343–360.

43. Solomon, C.M., Collier, J.L., Berg, G.M., & Glibert, P.M.(2010). Role of urea in microbial metabolism in aquaticsystems: A biochemical and molecular review, AquaticMicrobial Ecology, 59, 67–88.

44. Wang, W.H., K€ohler, B., Cao, F.Q., & Liu, L.H. (2008).Molecular and physiological aspects of urea transport inhigher plants, Plant Science, 175, 467–477.

45. Huerlimann, R., de Nys, R., & Heimann, K. (2010).Growth, lipid content, productivity, and fatty acid com-position of tropical microalgae for scale-up production,Biotechnology and Bioengineering, 107, 245–257.

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