Research Article
Effect of Carbon Dioxide Concentration on Cell Growthand Oil Yield of Freshwater and Marine Microalgae
Mariam Al Hattab and Abdel Ghaly*
Corresponding Author: Professor A. E. Ghaly, Department of Process Engineering and Applied Science, Faculty of Engineering, Dalhousie University, Halifax, Nova Scotia, Canada; Tel::902-494-6014; Email: abdel.ghaly@dal.ca
Received: March 26, 2015; Accepted: April 6, 2015; Published: April 15, 2015;
Citation: Al Hattab M& A Ghaly(2015) Effect of carbon dioxide concentration on cell growth and oil yield of freshwater and marine microalgae. International Journal of Bioprocess and Biotechnological Advancements, 1(1):31-44
Copyrights: ©2015 Al HattabM, Ghaly A. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
 

The Chlorella saccharophila (freshwater) and Tetraselmissuecica (marine) microalgae were investigated for their potential as feedstock for the production of biodiesel.  Assessment was based on their ability to produce biomass and lipid accumulation.  Varying concentrations of CO2 (3, 6 and 9%) were used at 24 h light exposure.  The results indicated that T. suecica produced higher cell yields compared to the C. saccharophila under all parameters tested.  Statistical analysis indicated that the biomass yields achieved using CO2 at varying concentrations were significantly different from one another.  However, varying CO2 concentrations over the range of 3 to 9% did not significantly affect the oil yields for both species over the elapsed time.  Thus, from an economic stand point it is much more suitable to use CO2 at a concentration of 3% as opposed to higher concentrations.  The use of NaHCO3 as a carbon source on the biomass and oil yields were also evaluated using the same experimental parameters.  Results indicated that the CO2 carbon source resulted in higher biomass and oil yields for the marine microalgae species when compared to NaHCO3 carbon source.  However, the Chlorella saccharophila species resulted in higher biomass and lipid yields using the sodium bicarbonate carbon source.  This suggests that different species have a preferred carbon source.  Some are better at the uptake of one source over the other.  Lower oil yields were achieved using the 3% CO2 as the carbon source compared to NaHCO3 for the Chlorella saccharophilaspecies.  The Tetraselmissuecicaspecies resulted in a slight increase in oil yield using 3% CO2 as opposed to NaHCO3.  The optimal growth conditions for Chlorella saccharophilaare the combination of nutrients, with 24 h light exposure and NaHCO3 as a carbon source and those for Tetraselmissuecicaare the ammonium nitrate, the 24 h light exposure and 3% CO2 as a carbon source.

Keywords
Freshwater, Marine Water, Microalgae, Cell yield, Oil Yield, NaHCO3, CO2, Carbon Source.

INTRODUCTION

The demonising fossil fuel reserves and the environmental concerns associated with burning fossil fuels have accelerated the need for a renewable energy source that is environmentally friendly.  Increased carbon dioxide emissions have been correlated with the amount of fossil fuel being burnt [1].  Biofuels such as biodiesel and bioethanol are promising substitution for petroleum fuel source [2].  Numerous feedstocks can be used as biomass for biofuel generation which include food waste, agricultural waste, municipal waste and both edible and nonedible oilseeds [3].  Currently, the best crops for biofuel production are oilseeds, but they are considered a food source for many people around the world [2,3].  However, microalgae have been noted to store oil that is 10 folds higher than the leading plant crop [4]. 

Microalgae are abundant microorganisms in nature, able to convert carbon dioxide into biomass which can then be used for biodiesel production via a transesterification reaction process [5].  Microalgae as an alternatefuel source is ideal because of their high growth rates and their ability to store higher lipids than the leading crop plants [6].  In addition, the waste that is generated after oil extraction can be used for other value added products, such as animal feed, organic fertilizers, and other biofuel products such as methane and ethanol via fermentation [7].  The amount of lipids stored in the microalgal cells can be manipulated by changing the environmental parameters such as temperature, pH, nutrient source, carbon source and light duration [8-10]. 

Biodiesel is a liquid fuel that is biodegradable and nontoxic [11].  It generates the same amount of energy (calorific value) as that produced using petroleum diesel without the resale of harsh compounds such as NOx, SOx and hydrocarbons into the atmosphere [3,12].  Biodiesel can be used in existing diesel engines without the need for much modification [2].  It is for these reasons that biodiesel is regarded as the best renewable energy source that is environmentally friendly and a viable source for replacement of the currently used petroleum source. 

OBJECTIVES

The aim of this study was to investigate the possibility of increasing the microalgae cell growth rate and oil yield by exposing the microalgae to various carbon sources (NaHCO3 and CO2) in a specially designed pilot scale open pond system.  The specific objectives were: (a) to study the effect of CO2 concentration in the air on biomass yield and oil content at three levels (3:97, 6:94 and 9:91 v/v CO2 to air) and (b) to evaluate the effect of carbon source (NaHCO3 and CO2) on the microalga biomass yield and oil content.

MATERIALS AND METHODS

Experimental Apparatus

A fully automated multiple open pond system (Figure 1) consisted of a frame, 18 open pond units, a cooling unit, a lighting unit, a supernatant collection unit and control unit was used in this study. 

The frame (244 cm in width x 41 cm in depth x 283 cm in height) consisted of three shelves (76 cm apart) and housed the open pond, light, cooling, water collection and control units.  Each shelf was divided vertically into two sides by a 1.2 cm thick plywood sheet to provide a better control of light and feed.  The open pond unit consisted of six ponds, each was made of galvanized steel and was divided into three compartments (each was 38 cm in length x 38 cm in width x 12.5 cm in height and can hold up to 18 L).  The lighting unit provided 430 hectolux of illumination per shelf (480 µmol m-2 s-1) using a mixture of fluorescent and incandescent lamps (six 40 W cool white fluorescent lamps 122 cm in length and four 100 W incandescent bulbs) mounted on each shelf, that sit 100 cm away from the ponds.  A cooling unit was designed to continuously remove the heat produced by the lamps to avoid heating of the algae on the upper and middle shelves.  A 5 cm diameter PVC pipe (having 6 mm diameter holes spaced 6 cm apart and facing out) was placed under the backside of the ponds.
Two metal blocks placed under each pond provided a 5 cm space between the pond and the lighting system of the shelf below it.  A 5 cm diameter PVC pipe was attached vertically to the left side of the frame and acted as a manifold through which air was blown by means of a motor driven fan (Model AK4L143A Type 821, Franklin Electric, Bluffton, Indiana, United States of America). The supernatant from each tray was collected in a separate container (2.7 L each) located at the bottom of the system.  The outlets were connected to plastic tubes of 1 cm outside diameter, which were passed through a solenoid valve. 

A computer was used to operate and control the various components of the open pond system and record the various measurements.  The light intensity was measured using a Quantum Sensor, SQ-316 Series (Apogee, Logan, Utah, United States of America). The pH was measured using pH electrodes (EW-59001-65, Cole Parmer, Montreal, Quebec, Canada). The temperature was measured using thermocouples (WD-08541-12, Nova-Tech International, Houston, Texas, United States of America). A basic computer program (BASIC Stamp Editor v2.5) allowed the configuration of the operating frequency and duration of the light, aeration unit and collection system. The computer was connected to a data coordinator (cDAQ-9178, National Instruments) which had 24 digital output ports and 24 digital input ports. The digital output ports were connected to electronic circuits which were responsible for the lighting, cooling and collection systems.

Microalgae

One freshwater microalgae (Chlorella saccharophila) and one marine (Tetraselmissuecica)were selected based on their ability to yield high biomass and store lipids [13]. 

The freshwater strain Chlorella saccharophilawas selected for study because of its high lipid content (45%).  This strain is capable of achieving a biomass yield of 3.88 g/L, which is not the highest among the freshwater species, but can however be offset by the fact that it achieves the highest lipid content.  This results in a lipid yield of 1.75 g/L.  The highest biomass yielding algae Scenedesmusobliqus of 4.34 g/L only achieves a lipid content of 38%, which intern results in a lipid yield of 1.69 g/L. Chlorella saccharophilais a green unicellular microalga belonging to the Chlorella genus [14].  The cells have an average size of 7.3 μm[15]. The cells contain a single chloroplast enclosed in a spherical or subspherical form.  These cells reproduce asexually through production of non-motile autospores[16].  This species is able to use glucose [17],bicarbonateandcarbondioxideasthecarbon source for growth [18]. The optimal temperature and pH for growth are 20-24°C and 7.5-8, respectively.

The marine microalgae strain Tetraselmissuecicawas selected for this study because of its high biomass yield of 4.48 g/L and comparatively high lipid content.  This species achieves a lipid content of 23% which is not the highest among the other species but can, however, be offset by the fact that it achieves the highest biomass yield.  This results in a lipid yield of 1.03 g/L, while the Chaetocerosmuelleri, with the highest lipid content of 34%, only achieves a biomass yield of 0.98 g/L, which results in a lipid yield of 0.33 g/L.  Tetraselmissuecicagrows as single cells.  They are motile and can be compressed or curved, but they are never twisted [19].  The cells are spherical or elliptic with a length of 35 μm and a width of 14 μm.  This species is able to use both sodium bicarbonate [20] and carbon dioxide [21] as the carbon source for growth. The optimal temperature and pH for growth are 18-24°C and 7-9, respectively [22]. 

Experimental Design

Two set of experiments were carried out. In the first set of experiments, the selected freshwater (Chlorella saccharophila) and marine (Tetraselmissuecica) microalgae species were grown in an open pond system using NaHCO3 as a carbon source.  The sodium bicarbonate (NaHCO3) was administered at a concentration of 1300 mg/L.  Ammonium nitrate was used for the marine microalgae and a combination of nutrients (ammonium nitrate, ammonium sulfate and ammonium phosphate) was used for the freshwater algae as sources of nitrogen at the optimum light exposure (24 h) as recommended by [13].  The light intensity was kept at 480 μmol/m2 s1 and the nitrogen content, pH and temperature were kept constant at 70 mg/L, 8.3-8.9 and 22°C, respectively.  In the second set of experiments the effects of carbon dioxide concentration on the algae biomass and oil content were evaluated. Carbon dioxide was administered at concentrations of 3, 6 and 9% (v/v in air).  The algae wereexposed to full light exposure (24 hand the light intensity was kept at 480 μmol/m2 s1).  A combination of nutrients (ammonium nitrate, ammonium phosphate and ammonium sulfate) was used as nutrient for the freshwater microalgae and ammonium nitrate was used for the marine microalgae.  The nitrogen content, pH and temperature were the same as in the first set of experiment.  The best results obtained with CO2 were compared with those obtained with NaHCO3

Preparation of Liquid Medium for Inoculum Growth

The freshwater microalgae medium was prepared on algal proteose medium (ATCC Catalog Medium No. 847, American Type Culture Collection, Manassas, Virginia, United States of America) and was made up by adding 1 g of proteose peptone (Difco 0120) to 1 L of Bristols solution (Table 1). Bristols solution was prepared by adding the following amounts from the prepared stock solutions: 10mL of NaNO3, 10 mL of CaCl2, 10 mL of MgSO4 7H2O, 10 mL of K2HPO4, 10 mL of KH2PO4, 10 mL ofNaCl,0.05 mLofFeCl3and940 mL of distilled water.  The stock solutions were prepared as follows: 10 g of NaNO3 in 400 mL of distilled water, 1g of CaCl2 in 400 mL of distilled water, 3 g of MgSO4 7H2O in 400 mL of distilled water, 3 g of K2HPO4 in 400 mL of distilled water, 7 g of KH2PO4 in 400 mL of distilled water and 1 g of NaCl in 400 mL of distilled water. 

The marine microalgae medium was prepared in F/2 medium [23]. The trace element liquid medium stock solution (Table 2) was prepared by the addition of 4.16 g of Na2 EDTA, 3.15 g of FeCl3•6H2O, 0.01 g of CuSO4•5H2O, 0.022 g of ZnSO4•7H2O, 0.01 g of CoCl2•6H2O, 0.18 g of MnCl2•4H2O and 0.006 g of Na2MoO4•2H2O into 1 L of autoclaved seawater (HalifaxWaterfront, Halifax, Nova Scotia, Canada). The vitamin mix stock solution was prepared by theaddition of 0.1 g of Thiamine HCl and 0.0005 g of biotin into 1 L of autoclaved seawater. The liquid medium was prepared by the addition of 0.075 g of NaNO3, 0.00565 g of NaH2PO4•2H2O, 1.0 ml of trace element stock solution and 1 ml of vitamin mix stock solution.

Preparation of Solid Medium for Inoculum Growth

The marine micro algae grow only in a marine liquid medium.  The fresh water micro algae solid medium was prepared on algal proteose agar medium (ATCC Catalog Medium No. 847, American Type Culture Collection, Manassas, Virginia, United States of America). The solid medium (Table 1) was made up by the addition of 1 g of proteose peptone (Difco 0120) and 15 g of agar to 1 L of Bristols solution.

Preparation of Inoculum

Sufficient amounts of inoculum were prepared for all the experimental runs for both freshwater and marine microalgae in order to maintain consistency. The procedures for preparing the inoculaare depicted in Figure 2.

The freeze dried Chlorella saccharophila sample (ATCC® 30408TM, Catalog Medium No. 847, AmericanType Culture Collection, Manassas, Virginia, United States of America) was revived in 5 mL of Bristols liquid media.  Using an inoculating loop, cells were transferred from the liquid media onto 3 petri dishes containing proteose agar medium. The plates were incubated for 3 days at room temperature and a photocycle of 14 h light and 10 h dark. The cells were then transferred, by scraping them off the solid media using an inoculating loop and submerging them into a 125 mL Erlenmeyer flask containing 25 mL of Bristols liquid medium. These cells were then left togrow for 2 weeks at a photocycle of 14 h light and 10 h dark. The sample was then transferred to a 500 mL Erlenmeyer flask containing 250 mL of Bristols liquid media which was left to grow for 2 weeks at a photocycle of 14 h light and 10 h dark. Finally, the media was transferred from the 500 mL flask into a 30 L bioreactor containing 25 L of Bristols liquid media and left to grow for 2 more weeks at a photocycle of 14 h light and 10 hour dark.  

The inoculum for Tetraselmissuecica microalga was prepared by taking 5 mL of the liquid sample (UTEX LB 2286, Cedarlane, Burlington, Ontario, Canada)and adding it to 125 mL Erlenmeyer flask containing 25 mL of F/2 liquid media and then left to grow at room temperature for 2 weeks at a photocycle of 14 h light and 10 h dark.  The mixture was then transferred to a 500 mL Erlenmeyer flask containing 250 mL of F/2 liquid media and was left to grow for 2 weeks at a photocycle of 14 h light and 10 h dark.  Finally, the media was transferred from the 500 mL flask into a 30 L bioreactor containing 25 L of F/2 liquid media and left to grow for 2 additional weeks at a cycle of 14 h light and 10 hour dark.

Preparation of Algae Production Media

The freshwater production medium is a modification of the Fitzgerlad medium [24].  The preparation of the stock solutions for this media is shown in Table 3.  The medium was made up by the addition of 1 mL of each of the stock solutions A, B, C and D to 1 L distilled water (Table 4).  

A modification of the F/2 media [23] was used as the production medium for the marine microalga.  The medium was modified by eliminating the addition of sodium nitrate.  The medium consists primarily of autoclaved ocean water (Halifax Waterfront, Halifax, Nova Scotia, Canada). Table 5 shows the elemental analysis of the components present in the marine water which was performed at the Mineral Engineering Center of Dalhousie University.

Experimental Protocol

To each compartment in the open pond system a total of 4.75 L of freshwater production media was added.  The amount of nutrient was added to the production medium.  This solution was enriched with the desired carbon source (1.3 g/L of sodium bicarbonate, 3% CO2, 6% CO2 or 9% CO2).  To this, 250 mL of Chlorella saccharophila inoculum was added to each compartment.  The cells were exposed to 24 h light and left to grow for 10 days.  Every otherday, 100 mL sample was taken for experimental analyses.  The samples were analyzed for pH and biomass yield.  At the end of the run the biomass was harvested from the liquid media using a Sorvall T1 Centrifuge (Thermo Scientific, Marietta, Ohio, United States of America). The supernatant from the centrifuge tubes was decanted and the cells were collected for biomass yield and oil content analyses.  The marine medium was used with marine algae and the same procedure was followed using the ammonium nitrate nutrient system.

Microalgae Biomass Determination

The freshwater mciroagla yield was determined by measuring the optical density at 484 nm from a standard curve between the cell count and optical density. The number of Colony Forming Units (CFU) for Chlorella saccharophila was determined using a series of dilutions. A test tube containing 9 mL of autoclaved distilled water and a 1 mL aliquot sample was added to the tube. The contents of the tube were vortexed (Thermolyne Maxi Mix, Thermolyne Corporation, Hampton, New Hampshire, United States of America) to distribute the cells. A 1 mL aliquot of this solution was added to another tube that had been autoclaved with 9 mL of distilled water.  This tube was again vortexed to distribute the cells.  This wasrepeated7 times to obtain dilutions of 1:1, 1:10, 1:100, 1:1000, 1:10 000, 1:100 000, 1: 1 000 000.  For each of the dilutions made, 0.1 mL of the solution was added to a petri dish containing solid freshwater medium.  The plates were sealed withparafilm, invertedand incubated atroomtemperature (~24°C) at a photocycle of 14 hours light and 10 hours dark for 3 days.  The plates were then removed and the colonies were counted using a colony counter (Model No. 7-910, Fisher Scientific, Ottawa, Ontario, Canada).  The plates consisting of 30-300 CFU were used for calculating the CFU of the sample and the standard curve was prepared by plotting the optical density against the plate count (Figure 3a). The following equation was used to calculate Chlorella saccharophila cell yield:

Cell Yield=((Optical Density)/(5x〖10〗^(-7) ))  x〖 10〗^3                     (1)


The marine microalgae yield was also determined byoptical density measurements. The cell counts were determined using a hemocytometer under a light microscope.  In a 250 mL Erlenmeyer flask, 175 ml of F/2 media was prepared and inoculated with Tetraselmissuecica.The flask was then incubated at room temperature (~24°C) for 2 weeks with 14 h light and 10 h dark periods.  At 2, 5, 8, 11 and 14 days 0.01 mL of sample was taken to determine cell counts using Hemocytometer slide. A standard curve between the cell count and optical density (measured at 750 nm) was developed (Figure 3b) and the followingequation was used to calculate Tetraselmissuecica cell yield: 

     Cell Yield=((Optical Density)/(23x〖10〗^(-4) ))  x〖 10〗^4           (2)

Oil Content Determination

The oil content in the biomass was determined according to Bligh and Dyer method described by Araujo et al. [25] using ultrasound assisted solvent extraction. Firstly, the algae biomass were homogenized and mixed with 25 mL of methanol, 12.5 mL of chloroform and 5 mL of distilled deionized water. This mixture was exposed to ultrasonic energy (Branson 2510R-DTH, Branson Ultrasonics Corporation, Danbury, United States of America) for 40 min. Then, an additional 12.5 mL of chloroform and 12.5 mL of 1.5% (w/v) sodium sulfate solution were added and sonicated for another 20 min.  The solid biomass particles were filtered out of the solution and the liquid fraction was transferred to a separatory funnel with the addition of 75 mL of 0.88% (w/v) KCl.  The mixture was vigorously shaken and left to separate for 24 h.  The solubility of oils in the chloroform solvent and the insolubility of solvents in water allowed for separation to occur into two phases (organic and aqueous). The oil containing phase (on the bottom) was drained out of the separatory funnel and collected into a pre-weighed distill flask.  The flask was distilled using rotary evaporator (HiTEC RE-51, Yamato Scientific America, Santa Clara, California, United States of America) set at 45°C.  The oil left behind was weighed in the flask and the yield was determined as follows:
Oil Yield (%) = (weight of Oil (g))/(weight of Algae Biomass (g)) x 100     (3)

RESULTS AND DISCUSSIONS


Microalgae biomass and oil content were determined for the freshwater (Chlorella saccharophila)and marine (Tetraselmissuecica) microalgae species suing NaHCO3 and 
CO2 as carbon sources with the recommended optimum light exposure and nitrogen source by Al hattab and Ghaly [13].  The results are shown in Table 6. 

Microalgae Biomass Using CO2 as a Carbon Source

Analysis of variance (ANOVA) was performed on the cell yield data as shown in Table 7 using Minitab statistics software (Minitab® 16.2.2., Minitab Inc., Canada).  The effect of microalgae type and the carbon dioxide concentration on the cell yield and the interactions between them are significant at the 0.01 level.  Tukey’s grouping was used to test the differences among the levels of each parameter as shown in Table 8.  The two microalgae were significantly different from one another at the 0.05 level.  The highest mean cell yield (1.76x106 cells/mL)was obtained from the marine microalgae Tetraselmissuecica.  The CO2 concentrations of 6 and 9% were not significantly different form one another but both were, however, significantly different from the 3% concentration at the 0.05 level.  The highest mean cell yield (1.47x106 cells/mL) was obtained with 9% CO2 concentration.  The CO2 concentrations 6% and 9% produced 108% and 122% more cells than that produced with the 3% CO2 concentration, respectively. 

Effect of Microalgae Type

The effects of microalgae type on the cell yield are shown in Figure 4.  Tetraselmissuecica achieved higher cell yields than the Chlorella saccharophila at all carbon dioxide concentrations.  Tetraselmissuecica resulted in cell numbers of 1.148x106, 2.213x106 and 1.930x106 cells/mL while Chlorella saccharophila resulted in cell numbers of 0.181x106, 0.553x106 and 1.019x106 cells/mL at the CO2 concentrations of3%, 6% and 9%, respectively. 

Reports in the literature indicate that the Tetraselmissuecica species produces much higher cell yields (1290-4900 mg/L) than the Chlorella saccharophila species (138-1543 mg/L). Singh et al. [17] reported that Chlorella saccharophila produced a dry cell yield of 378 mg/L.  Isleten-Hosoglu et al. [26] reported a dry cell weight of 138 mg/L for Chlorella saccharophila.  Herrera-Valencia et al. [27] reported a biomass yield of 1.54 g/L for the freshwater species Chlorella saccharophila.  Bondioli et al. [28] noted a biomass yield of 1.66 g/L for theTetraselmissuecica species.  Danquah et al.[29] investigated the Tetraselmissuecica species and noted a biomass  yield  of  1.29 g/L.Michles et al. [30] reported a biomass yield of 4.9 g/L for the marine species Tetraselmissuecica. 

Chinnasamy et al. [31] reported a biomass productivity of 23 mg/L/d for Chlorella saccharophila.  Herrera-Valencia et al. [27] achieved biomass productivity of 154.3 mg/L/dfor Chlorella saccharophila.  Hempel et al. [32] noted a biomass productivity of 310 mg/L/d for the freshwater species Chlorella saccharophila.  Bondioli et al. [28] noted a biomass productivity of 237 mg/L/d for Tetraselmissuecica.  Moheimani [33] achieved a biomass productivity of 320 mg/L/d for Tetraselmissuecica. 

These findings agree with the findings of this study in which case the biomass yield and productivity obtained for the Tetraselmissuecica are higher than those obtained from the Chlorella saccharophila.  In this study the dry cell yield and biomass productivity for the Chlorella saccharophilaranged from 280.5 mg/L to 1579.45 mg/L and 28 mg/L/d to 154 mg/L/d, while those for Tetraselmissuecica species 1779.4-2991.5 mg/L and 177.9-299.2 mg/L/d, respectively.  These values are within the reported range in the literature. 

Effect of Carbon Dioxide Concentration

The effect of carbon dioxide concentration on the biomass yield of the marine and freshwater microalgae is illustrated in Figure 5.  As the carbon dioxide concentration increased from 3% to 9%, a gradual increase in cell yield for Chlorella saccharophila was observed.However, an increase in CO2 concentration from 3 to 6% resulted in increased yield for the Tetraselmissuecica species, but a further increase to 9% resulted in decreased cell yields. 

CO2 provides the cells with the necessary carbon source for cell growth and maintenance.  Low levels of COretard the growth of the cells due to the low carbon availability [34,35]. However, excess CO2 present in the media would alter the pH (Figure 6) which would in turn affect the growth of the cells [36,37].  The algae tolerance to H2CO3 is specific to the species, thus the varying trends of cell yield seen in Figure 5 are the result of species tolerance to the change in pH of the mediumSalih et al. [4]stated that the microalgae growth efficiency and productivity on affected by the CO2 concentration and noted that higher COconcentration resulted in better growth and productivity. Schippers et al. [38] noted that doubling atmospheric carbon dioxide in nutrient rich media resulted in increased microalgae biomass of up to 40% and 50% for saltwater and freshwater species, respectively.  Widjaja[36] stated that an optimal CO2 concentration exists for each species since excess CO2 presents a toxic environment for the cells as a result of changing pH.

Yue and Chen [39] reported that the freshwater microalgae Chlorella species resulted in an increase of 200% in algal growth rate when 1% CO2 was used in the medium as opposed to the ambient air.  They also noted that higher concentrations of CO2 resulted in a decline in algal growth as a result of increased acidity.  Nakano et al. [40]reported that the microalgae species Euglena gracilis exposed to varying concentrations of CO2 (5-45%) grew best at 5% CO2 while concentrations greater than 45% inhibited growth. Maeda et al. [41] noted that Chlorella sp. microalgae was capable of growing at 100% CO2 concentrations, but the highest growth rate was achieved at 10% COconcentration.  Hanagata et al. [42] reported that the algae Scenedesmus sp. can grow at 80% CO2 concentration, but the maximum growth was achieved at 10-20% CO2 concentration.  Studies performed by Ho et al. [43] and Jacob-Lopes et al. [44] found that excess CO2 absorbed in the media has a negative effect on the microalgae growth. 

Kaewkannetra et al. [34] noted that an increase in CO2 concentration from 5% to 15% resulted in an increase in biomass yield from 1.7 g/L to 2.3 g/L.  However, further increase in CO2 concentration resulted in a decreased yield.  Yue and Chen [39] noted that the microalgae species Chlorella grown in CO2 concentration over the range of 0.035-70% and noted the highest cell concentration of 5.9 g/L at 10%CO2 concentration.  Ho et al. [43] and de Morris and Costa [45] noted that an optimum CO2 concentration for the species Scenedesmus exists between 10% and 15%. 

Goswami et al. [35] noted that the microalgae Selenastrum sp. resulted in biomass productivity of 0.667, 0.889, 0.797 and 0.778 mg/L/d at carbon dioxide concentrations of 4,400, 5,200, 7,500 and 8,200 ppm, respectively.  Devgoswami et al. [46] noted that upon an increase in CO2 concentration from 4,400 ppm to 4,758 ppm, the biomass productivity increased from 161 to 188%(g/L/d) while further increase in CO2 concentration (to 7,929 ppm) showed a decrease of 163% for the green microalgae Chlorella.  Similarly, the Haematococcus and Scenedesmus species showed increased growth rates as the concentration of COwas increased from 4,400 ppm to 4,758 ppm,but a further increase to 7,929 ppm resulted in lower growth rates. These findings as well as the findings of this study show that an optimal carbon dioxide concentration exists.

Microalgae Oil Content Using CO2as a Carbon Source

The oil yield results are depicted in Table 6.  Analysis of the variance (ANOVA) was performed on the oil yield data as shown in Table 9.  The effects of microalgae type on oil yield were significant at the 0.003 level.  However, the effect of CO2 concentration and the interactions between microalgae type and CO2 concentration were not significant.  Tukey’s grouping was used to test the differences among the levels of each parameter as shown in Table 10.  The two microalgae Chlorella saccharophilaand Tetraselmissuecica were significantly different from one another at the 0.05 level.  The highest mean oil yield (4.18%)was obtained from the freshwater microalgae species.  The CO2 concentrations were not significantly different from one another at the 0.05 level.  The highest mean oil yield (3.16%) was obtained with the 3% CO2 concentration.

Effect of Microalgae Type

The effect of the microalgae type on the oil content is illustrated in Figure 7.  Chlorella saccharophilaachieved the highest oil yields at all CO2 concentration.  It produced average oil yields of 4.71, 3.90 and 3.59% while Tetraselmissuecica produced average oil yields of 2.09, 1.01 and 2.43% at the 3%, 6% and 9% COconcentrations, respectively.  These results are similar to those of Pittman et al. [47] which indicated that the marine microalgae produce much lower oil yields compered to freshwater microalgae.  The oil yields obtained from Chlorella saccharophilawere 4 times higher than those obtained from Tetraselmissuecica, despite the higher biomass yields obtained from the Tetraselmissuecicaspecies

Sharma et al. [48] stated that the occurrence and extent to which lipids are produced by microalgae is species/strain specific.  Pittman et al. [47] stated that different species use their energy for different metabolic pathways.  In this study a trade-off between cell generation and lipid accumulation was seen among the species.  The marine species used most of its energy for cell generation as opposed to lipid while the freshwater spices used most of its energy for oil accumulation as opposed to cell generation.

Demirbas [7], Moheimani, [33], Sobczuk et al., [49], Sukenik et al., [50], Wagenan et al., [51] and Pagnanelli et al., [52] reported a lipid content in the range of 36-47% and 15-23% for Chlorella saccharophilaand Tetraselmissuecicaspecies, respectivelyThe differences in the lipid content are attributed to the different nutrient systems used and the culture age before harvest. 

Effect of Carbon Dioxide Concentration

The effect of carbon dioxide concentration on the oil yield is illustrated by Figure 8.  As the carbon dioxide concentration was increased from 3% to 9%, the oil yield decreased from 4.7% to 3.6%for the freshwater microalgae (Chlorella saccharophila).  On the other hand as the carbon dioxide concentration was increased from 3% to 6% the oil yield decreased from 2.1% to 1.0%for the marine microalgae (Tetraselmissuecica).  However, a further increase in CO2 concentration to 9% increased the oil yield to 2.43%. 

It should be noted that the trends in Figure 9 are the opposite of the trends of cell yield shown in Figure 5.  The higher oil yield obtained for the freshwater microalgae at the 3% CO2 concentration can be attributed to the trade-off of lower cell generation, and the lower oil yields obtained when the CO2 concentration was increased to 9% is a result of increased cell division.  Similarly, the variation in lipid content for the marine (Tetraselmissuecica) microalgae species can also be attributed to the variation in biomass yield caused by variation in CO2 concentration and the tolerance of the species to the acidity caused by higher CO2 concentrations.  The results showed that from an economic stand point, the 3% CO2 concentration is the most optimal condition for lipid accumulation in both species.  

Similar results were reported in the literature.  Widjaja et al. [36] noted that the microalgae species Chlorella vulgaris in lipid yields of 20%, 28% and 25% at the CO2 concentration of 0, 0.33 and 0.83%, respectively.  Huang and Su [53] noted that for the microalgae species Chlorella vulgaris grown using 0%, 15% and 50% CO2 concentrations resulted in lipid yields of 34%, 35% and 36%, respectively.  The findings are similar to those obtained in this study since they indicate that varying the CO2 concentration does not significantly influence the lipid content.  The variation in oil yield can be attributed to the varying cultivation periods, variation in nutrient systems and the effectiveness of the oil extraction methods used.

Effect of Carbon Source

Biomass

The cell yields shown in Table 6 for Chlorella saccharophila species which was obtained using NaHCO3 as a carbon source (0.689x106 cells/mL) were 74% higher than those achieved using 3% CO2 (0.181x106 cells/mL).  The cell yield for Tetraselmissuecica species that resulted while using NaHCO3 (0.750x106 cells/mL) were 53% lower than those achieved using 3% CO2 (1.148x106 cells/mL).  These results can be attributed to the cells ability to convert the carbon source into the preferred form for uptake and the abundance of the carbon source.  The direct uptake of bicarbonate through an active transport system has only been noted in certain species [54].  In addition, some species have better extracellular carboanhydrase activities which allow them to convert the carbon source into different forms [54-56]. Devgoswami et al. [46] studied the Chlorella microalgae species and noted biomass productivity of 82 and 189 mg/L/d using sodium bicarbonate and CO2, as the carbon source,respectively.  Moheimani[33] noted that the Chlorella sp. and Tetraselmissuecica grown using CO2as a carbon source resulted in biomass yields that were 6 and 23% higher than those obtained using NaHCO3 as the carbon source.  Goswami et al. [35] reported that the Selenastrum sp. grown using NaHCO3 (20-100 mg/L) and CO2 (4.4-8.2 g/L) resulted in biomass productivity in the range of 689-1102 mg/L/d and 667-889 mg/L/d, respectively.  In this study, the biomass productivity achieved using NaHCO3 and 3% CO2 for Chlorella saccharophilawas 106.8 mg/L/d and 28.1 mg/L/d, respectively.  The biomass productivity achieved using NaHCO3 and 3% CO2 for Tetraselmissuecicawas 116.3 mg/L/d and 177.9 mg/L/d, respectively.  Variation in the values achieved in this study and those of the literature are attributed to the different species and different cultivation methods used.

Oil yield

The oil yields obtained with NaHCO3 and 3% CO2 as carbon sources are shown in Figure 9.  The oil yield achieved by Chlorella saccharophila 3% CO2 as a carbon source (4.71%) concentration was significantly lower than that obtained using NaHCO3 (12.91%) as the carbon source.  This suggests that sodium bicarbonate is the preferred carbon source for Chlorella saccharophila that prompts lipid accumulation in the cells.  Whereas, the marine Tetraselmissuecica species is better at utilizing the CO2 as a carbon source for both the generation of cells and the accumulation of lipids.  Tetraselmissuecica resulted in higher cell and oil yields using the CO2 carbon source as opposed to NaHCO3.  However, the values are not significantly different from one another.  The media continuing CO2 can result in pH fluctuations and cause conditions that are not suitable for proper cell function (lipid accumulation).  The presence of NaHCO3 in the media helps regulate the pH so that the environment is not too acidic.   

Mukund et al. [57] tested Chlorella, Chlorococcum sp. and Desmococcus sp. and noted that the photosynthetic activity is strongly influenced by bicarbonate and that it results in increased lipid accumulation.  Nayak et al. [58] reported a lipid yield of 16% in Scenedesmussp IMMTCC-6 when it was supplied with CO2 only, while addition of NaHCO3 to the medium resulted in an increase in lipid yield to 22%.  Dhakal et al. [59] noted that the freshwater microalgae species Chlorella vulgaris produces a lipid yield of 22% using NaHCO3 as the carbon source.  The oil yields obtained in this study were lower than those reported in the literature.  This may be attributed to the different cultivation parameters, cultivation growth period and the fact that different species respond differently to the nutrients supplied. 

CONCLUSION

The cell growth and oil yields of Chlorella saccharophila (freshwater) and Tetraselmissuecica (marine) microalga were investigated using various carbon dioxide concentrations (3, 6, and 9%).  Statistical analysis indicated that the biomass yields achieved using CO2 at varying concentrations were significantly different from one another.  However, varying the CO2 concentration over the range of 3 to 9% did not significantly affect the oil yields for both species over the elapsed time.  Thus, from an economic stand point it is much more suitable to use CO2 at a concentration of 3% as opposed to higher concentrations.  The use of NaHCO3 as a carbon source on the biomass and oil yields were also evaluated using the same experimental parameters.  Results indicated that the CO2 carbon source resulted in higher biomass and oil yields for the Tetraselmissuecica species when compared to NaHCO3 carbon source.  However, the Chlorella saccharophila species resulted in higher biomass and lipid yields using the sodium bicarbonate carbon source.  This suggests that different species have a preferred carbon source.  Some are better at the uptake of one source over the other.  Lower oil yields were achieved using the 3% CO2 as the carbon source compared to NaHCO3 for the Chlorella saccharophila species. The Tetraselmissuecica species resulted in a slight increase in oil yield using 3% CO2 as opposed to NaHCO3.  These findings indicate that the optimal growth conditions for Chlorella saccharophilais the combination of nutrients, with 24 h light exposure and NaHCO3as a carbon source and those for Tetraselmissuecicaare the ammonium nitrate, the 24 h light exposure and 3% CO2 as a carbon source.

1.   Stepan, D.J., R.E. Shockey, T.A. Moe and R. Dorn, 2001.  SUBTASK, 2.3-Carbon dioxide dequestering using microalgal systems. U.S. Department of Energy, National Energy Technology Laboratory, 2001. Accessed on August 14, 2014.  http://www.osti.gov/scitech/servlets/purl/882000

2.   Singh, J. and S. Gu, 2010.  Commercialization potential of microalgae for biofuels production.  Renewable and Sustainable Energy Review, 14: 2596-2610. 

3.   Demirbas, A., 2005.  Biodiesel production form vegetable oils via catalytic and non-catalytic supercritical methanol transesterification methods.  Progress in Energy and Combustion Science, 31(5-6): 466-487. 

4.   Salih, F.M., 2011. Microalgae tolerance to high concentrations of carbon dioxide: A review. Journal of Environmental Protection, 2: 648-654. DOI:10.4236/jep.2011.25074

5.   Jeong, M.L., J.M. Gillis and J.Y. Hwang, 2003. Carbon dioxide mitigation by microalgal photosynthesis. Bulletin of Korean Chemical Society 24: 1763–1766.  ISSN:0253-2964

6.   Song, D., J. Fu and D. Shi, 2008. Exploitation of oil-bearing microalgae for biodiesel. Chinese Journal of Biotechnology, 24(3): 341-348. ISSN 1872-2075

7.   Demirbas, M.F., 2011. Biofuels from algae for sustainable development. Applied Energy, 88: 3473-3480.  DOI: 10.1016/j.apenergy.2011.01.059

8.   Guschina, I.A. and J.L. Harwood, 2006. Lipids and lipid metabolism in eukaryotic algae. Progress in Lipids Research, 45(2): 160-186.  DOI: 10.1016/j.plipres.2006.01.001

9.   Hu, Q., 2004.  Environmental effects on cell composition. In: Richmond A, editor. Handbook of Microalgal Culture: Biotechnology and Applied Phycology.  Oxford: Blackwell Science Ltd: 83–93.

10.          Hu, Q., M. Sommerfeld, E. Jarvis, M. Ghirardi, M. Posewitz, M. Seibert, A. Darzins, 2008. Microalgaltriacylglycerols as feedstocks for biofuel production: perspectives and advances. The Plant Journal, 54: 621-639. ISSN:0960-7412

11.          Ulusoy, Y., Y. Tekin, M. Cetinkaya and F. Karaosmanoglu, 2004.  The engine tests of biodiesel from used frying oil.  Energy Sources, 26(10): 927-932. 

12.          Kalam, M.A. and H.H. Masjuki, 2005. Recent developments on biodiesel in Malaysia.  Journal of Scientific and Industrial Research, 64(11): 920-927. 

13.          Alhattab, M. and A. Ghaly, 2014.  Effects of light exposure and nitrogen source on the production of oil from freshwater and marine water microalgae.  American Journal of Biochemistry and Biotechnology, In Press. 

14.          Lewis, L. A. 1997. Diversity and phylogenetic placement of BracteacoccusTereg (Chlorophyceae, Chlorophyta) based on 18S ribosomal RNA gene sequence data. J. Phycol. 33:279–85.

15.          Bock, C., L. Krienitz and T. Proschold, 2011.  Taxonomic reassessment of the genus Chlorella (Trebouxiophyceae) using molecular signatures (barcodes), including description of seven new species.  Fottea: Journal of the Czech Psychological Society, 11(2): 293-312.  ISSN: 1802-5439. 

16.          John, D.M., 2002.  The freshwater algal flora of the British Isles: An identification Guide to freshwater and terrestrial algae.  Cambridge University Press, 1:1890. 

17.          Singh, D., M. Puri, S. Wilkens, A.S. Mathur, D.K. Tuli and C.J. Barrow, 2013.  Characterization of a new zeaxanthin producing strain of Chlorella saccharophila isolated from New Zealand marine waters. Bioresource Technology, 143:308-314.  http://dx.doi.org/10.1016/j.biortech.2013.06.006

18.          Matsuda, Y. and B. Colman, 1996.  Active uptake of inorganic carbon by Chlorella saccharophila is not repressed by growth in high CO2.  Journal of Experimental Botany, 47(305): 1951-1956. 

19.          Acuna, J.L. and M. Kiefer, 2000. Functional response of the appendicularianOikopleuradioica. Limnology and Oceanography, 45, 608–618.  DOI:10.4319/lo.2000.45.3.0608

20.          White, D.A., A. Pagarette, P. Rooks, S.T. Ali, 2012.  The effect of sodium bicarbonate supplementation on growth and biochemical composition of marine microalgae cultures.  Journal of applied Phycology, 25(1): 153-165.  ISSN: 0921-8971

21.          De Castro Araujo, S. and V.M.T. Garcia, 2005.  Growth and biochemical composition of the diatom Chaetoceros cf. wighmiibrightwell under different temperature, salinity and carbon dioxide levels. I. Protein, carbohydrates and lipids.  Aquaculture, 246(1-4): 405-412.  ISSN 0044-8486

22.          Lavens, P. and P. Sorgeloos, 1996.  Manual on the production and use of live food for aquaculture.  FAO Fisheries Technical Paper. No. 361. Rome, FAO, pp. 295. ISBN: 92-5-103934-8

23.          Guillard, R.R.L. and J.H. Ryther, 1962. Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt and Detonulaconfervaceae (Cleve) Gran. Canadian Journal of Microbiology, 8, 229-239.  DOI:10.1139/m62-029

24.          Hughes, E. O., P. R. Gorham, and A. Zehnder, 1959. Toxicity of a unialgal culture of microcystis aeruginosa. Canadian Journal of Microbiology, 4, 225. DOI:10.1139/m58-024

25.          Araujo, G.S., L.J.B.L. Matos, J.O. Fernandes, S.J.M. Cartaxo, L.R.B. Goncalves, F.A.N. Fernandes and W.R.L. Farias, 2013.  Extraction of lipids from microalgae by ultrasound application: Prospection of the optimal extraction method.  UltrasonicsSonochemistry, 20: 95-98.  http://dx.doi.org/10.1016/j.ultsonch.2012.07.027

26.          Isleten-Hosoglu, M., I. Gultepe and M. Elibol, 2012. Optimization of carbon and nitrogen sources for biomass and lipid production by Chlorella saccharophila under heterotrophic conditions and development of Nile red fluorescene based method for quantification of its neutral lipid content. Biochemical Engineering Journal, 61:11-19.DOI:10.1016/j.bej.2011.12.001

27.          Herrera-Valencia, V., P.Y. Contreras-Pool, S.J. Lopez-Adrain, S. Peraza-Echeverria and L.F. Barahona-Perez, 2011.  The green microalga Chlorella saccharophila as a suitable source of oil for biodiesel production.  Current Microbiology, 63:151-157.  DOI: 10.1007/s0028-011-9956-7

28.          Bondioli, P., L. Della Bella, G. Rivolta, G. ChiniZittelli, N. Bassi, L. Rodolfi , D. Casini, M. Prussi, D. Chiaramonti, and M.R. Tredici, 2012. Oil production by the marine microalgae Nannochloropsis sp. F&M-M24 and Tetraselmissuecica F&M-M33. Bioresourse Technology, 114: 567–672. DOI: 10.1016/j.biortech.2012.02.123

29.          Danquah, M.K., R. Harun, R., R. Halim and G.M. Forde, 2010.  Cultivation medium design via elemental balancing for Tetraselmissuecica.  Chemical and Biochemical Engineering Quaterly, 24(3):361-369. ISSN:0352-9568

30.          Michels, M.H.A., P.M. Slegers, M.H. Vermue and R.H. Wijffels, 2013.  Effect of biomass concentration on the productivity of Tetraselmissuecica in a pilot-scale tubular photobioreactor using natural sunlight.  Algal Research.  http://dx.doi.org/10.1016/j.algal.2013.11.011

31.          Chinnasamy, S., A. Bhatnagar, R.W. Hunt, K.C. Das, 2010. Microalgae cultivation in a wastewater dominated by carpet mill effluents for biofuel applications. Bioresourse Technology, 101:3097–3105. DOI: 10.1016/j.biortech.2009.12.026

32.          Hempel, N., I. Petrick and F. Behrendt, 2012.  Biomass productivity and productivity of fatty acids and amino acids of microalgae strains as key characteristics of suitability for biodiesel production.  Journal of Applied Phycology, 24:1407-1418.  DOI: 10.1007/s10811-012-9795-3.

33.          Moheimani, N.R., 2013.  Inorganic carbon and pH effect on growth and lipid productivity of Tetraselmissuecica and Chlorella sp. (Chlorophyta) grown outdoors in bag photobioreactors.  Journal of Applied Phycology, 25:387-398.  ISSN: 1573-5176

34.          Kaewkannetra, P., P. Enmak and T. Chiu, 2012.  The effect of CO2 and salinity on the cultivation of Scenedesmusobliquus for biodiesel production.  Biotechnology and Bioprocess Engineering, 17:591-597.  DOI 10.1007/s12257-011-0533-5

35.          Goswami, R.C.D, N. Kalita and M.C. Kalita, 2012.  A study on growth and carbon dioxide mitigation by microalgae Selenastrum sp. its growth behavior under different nutrient environments and lipid production.  Annals of Biological Research, 3(1):499-510.  http://scholarsresearchlibrary.com/archive.html

36.          Widjaja, A., 2009. Lipid production from microalgage as a promising candidate for biodiesel production. Makara Journal of Technology Series, 13(1):47-51. DOI: 10.7454/mst.v13i1.496

37.          Sorensen, B. H., N. Nyholm, and A. Baun, 1996. Algal toxicity tests with volatile and hazardous compounds in air-tight test flasks with CO2 enriched headspace. Chemosphere, 32 (8): 1513.  ISSN:0045-6535

38.          Schippers, P., M. Lürling and M. Scheffer, 2004.  Increase of atmospheric CO2 Promotes Phytoplankton Productivity. Ecology Letters, 7(6): 446-451. DOI:10.1111/j.1461-0248.2004.00597.x

39.          Yue, L. and W. Chen, “Isolation and Determination of Cultural Characteristics of a New Highly CO2 Tolerant Fresh Water Microalgae,” Energy Conversion and Man-agement, Vol. 46, No. 11-12, 2005, pp. 1868-1876. DOI:10.1016/j.enconman.2004.10.010

40.          Nakano, Y., K. Miyatake, H. Okuno, K. Hamazaki, S. Takenaka, N. Honami, M. Kiyota, I. Aiga and J. Kondo, 1996. Growth of photosynthetic algae euglena in high CO2 conditions and its photosynthetic characteristics. ActaHorticulturae, 440(9): 49-54.

41.          Maeda, K., M. Owada, N. Kimura, L. Omata, and I. Ka-rube, 1995.  CO2 fixation from the flue gas on coalfired thermal power plant by microalgae. Energy conversion Management, 36(6-9): 717-720. DOI:10.1016/0196-8904(95)00105-M

42.          Hanagata, N., T. Takeuchi and Y. Fukuju, “Tolerance of Microalgae to High CO2 and High Temperature,” Phyto-chemistry, Vol. 31, No. 10, 1992, pp. 3345-3348. doi:10.1016/0031-9422(92)83682-O

43.          Ho, S., Chen W, Chang J. 2010. Scenedesmusobliquus CNW-N as a potential candidate for CO2 mitigation and biodiesel production. Bioresource Technology, 101: 8725-8730.  doi: 10.1016/j.biortech.2010.06.112.

44.          Jacob-Lopes E, Lacerda LMCF, Franco TT. 2008. Biomass production and carbon dioxide fixation by AphanothecemicroscopicaNageli in a bubble column photobioreactor. Biochemical Engineering Journal, 40(1): 27-34.  http://dx.doi.org/10.1016/j.bej.2007.11.013

45.          De Morris, M.G. and J.A. Costa, 2007. Carbon dioxide fixation by Chlorella kessleri, C. vulgaris, Scenedesmusobliquus and Spirulina sp. cultivated in flasks and vertical tubular photobioreactor. Biotechnology Letters, 29:1349–1352. ISSN:0141-5492

46.          Devgoswami, Ch.R., M.C. Kalita, J. Talukdar, R. Bora and P. Sharma, 2011.  Studies on the growth bechavior of Chlorella, Haematococcus and Scenedesmus sp. in culture media with different concentrations of sodium bicarbonate and carbon dioxide gas.  African Journal of Biotechnology, 10(61):13128-13138.  ISSN 1684–5315

47.          Pittman, J.K., A.P. Dean and O. Osundeko, 2011. The potential of sustainable algal biofuel production using wastewater resources. Bioresource Technology, 102:17–25. ISSN 0960-8524

48.          Sharma, K.K., H. Schuhmann and P.M. Schenk, 2012.  High lipid induction in microalgae for biodiesel production.  Energies, 5:1532-1553.  ISSN: 1996-1073. 

49.          Sobczuk, T.M., F.G. Camacho, F.C. Rubio, F.G.A. Fernandez and E.M. Grima, 2000.  Carbon dioxide uptake efficiency by outdoor microalgae cultures in tubular airlift photobioreactors.  ISSN:0006-3592

50.          Sukenik, A., J. Beardall, J.C. Kromkamp, J. Kopecky, J. Masojidek, S. van Bergeijk, S. Gabai, E. Shaham and A. Yamshon, 2009. Photosynthetis performance of outdoor Nannochloropsis mass cultures under a wide range of environmental conditions. Aquatic Microbial Ecology: Preprint. DOI: 10.3354/ame01309

51.          Wagenen, J.V., T.W. Miller, S. Hobbs, P. Hook, B. Crowe and M. Huesemann, 2012. Effects of light and temperature on fatty acid production in Nannochloropsissalina. Energies, 5: 731-740. DOI:10.3390/en5030731

52.          Pagnanelli, F., P. Altimari, F. Trabucco and L. Toro, 2013. Mixotrophic growth of Chlorella vulgaris and Nannochloropsisoculata: interaction between glucose and nitrate. Journal of Chemical Technology and Biotechnology. DOI 10.1002/jctb.4179

53.          Huang, Y.T. and C.P Su, 2013.  High lipid content and productivity of microalgae cultivating under elevated carbon dioxide.  International Journal of Environmental Science and Technology.  DOI: 10.1007/s13762-013-0251-y

54.          Merrett, M.J., N.A. Nimer and L.F. Dong, 1996. The utilization of bicarbonate ions by the marine microalga Nannochloropsisoculata (Droop) Hibberd. Plant, Cell and Environment, 19: 478-484.

55.          Huertas, I.E., G.S. Espie, B. Colman and L.M. Lubian, 2000. Light-dependent bicarbonate uptake and CO2 efflux in the marine microalga Nannochloropsisgaditana. Planta, 211(1): 43-49. ISSN:0032-0935

56.          Ginzberg, A., M. Cohen, U.A. Sod-Moriah, S. Shany, A. Rosenshtrauch and S. Arad, 2000. Chickens fed with biomass of the red microalga Porphyridium sp. have reduced blood cholesterol level and modified fatty acid composition in egg yolk. Journal of Applied Phycology, 12: 325-330.  ISSN: 0921-8971

57.          Mukund, J.G., N.S. Senthilkumar and R.R. Vallinayagam, 2013.  Influence of different concentrations of sodium bicarbonate on growth rate and biochemical composition of microalgae.  Journal of Algal Biomass Utilization, 4(4): 81-87.  ISSN: 2229-6905. 

58.          Nayak, M., S.S. Rath, M. Thirunavoukkarasu, P.K. Panda, B.K. Mishra and R.C. Mohanty, 2013.  Maximizing biomass productivity and CO2biofixation of microalga, Scenedesmus sp. by using sodium hydroxide.  Journal of Microbiology and Biotechnology, 23(9): 1260-1268.  http://dx.doi.org/10.4014/jmb.1302.02044

59.          Dhakal, N. S. Lama, A. Shrestha, T.B. Karki and P.M. Timilsina, 2013.  Study of growth rate and lipid content of various microalgae species from Nepal.  Research Symposium Compendium, 3: 43-47.  http://www.ku.edu.np/renewablenepal/images/rentech3/10-nirpeshdhakal.pdf

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