Table of Contents Author Guidelines Submit a Manuscript
Journal of Marine Biology
Volume 2016, Article ID 6832847, 8 pages
http://dx.doi.org/10.1155/2016/6832847
Research Article

Growth, Fatty Acid, and Lipid Composition of Marine Microalgae Skeletonema costatum Available in Bangladesh Coast: Consideration as Biodiesel Feedstock

1Department of Biochemistry & Molecular Biology, University of Chittagong, Chittagong 4331, Bangladesh
2Institute of Marine Sciences and Fisheries, University of Chittagong, Chittagong 4331, Bangladesh
3Bangladesh Council of Scientific and Industrial Research (BCSIR), Dr. Qudrat-i-Khuda Road Dhanmondi, Dhaka 1205, Bangladesh

Received 20 October 2015; Revised 30 January 2016; Accepted 2 February 2016

Academic Editor: Garth L. Fletcher

Copyright © 2016 Tania Sharmin et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

Among the various potential sources of renewable energy, biofuels are of most interest. Marine microalgae are the most promising oil sources for making biofuels, which can grow very rapidly and convert solar energy to chemical energy via CO2 fixation. The fatty acid profile of almost all the microalgal oil is suitable for the synthesis of biofuel. In this research, fatty acid and lipid contents of Bangladeshi strains of marine microalgae Skeletonema costatum were performed. For this, the crude oil was extracted by Soxhlet extraction method, using three most common solvent systems, pure hexane and mixture of CHCl3 : MeOH (2 : 1) and hexane : EtOH (3 : 1) one by one. Highest oil recovery (15.37%) came from CHCl3 : MeOH (2 : 1) solvent system from dry biomass whereas the lowest (2.49%) came from n-hexane from wet biomass. The qualitative analysis of the extracted oil by GC/MS analysis revealed that it contained significant amount of myristic acid (C14:0), palmitic acid (C16:0), stearic acid (C18:0), and palmitoleic acid (C16:1). It also indicated presence of hexadecatrienoic acid, benzenedicarboxylic acid, oleic acid, arachidonic acid, eicosapentaenoic acid (EPA), 9-Octadecenoic acid methyl ester (C19H36O2), and so forth. The obtained fatty acid profile indicates high potentiality of S. costatum species to be used as promising biofuel feedstock a little improvisation and substantially it can replace diesel in near future.

1. Introduction

Energy demand worldwide is increasing continuously at a rapid rate with the increase of urbanization in developing countries. Majority of the world’s energy needs are supplied through petrochemical sources, coal, and natural gases, with the exception of hydroelectricity and nuclear energy, and these sources are finite and at current usage rates will be consumed shortly [1]. Heavy dependence on petroleum-based fuels is not sustainable due to increasing fuel costs, diminishing crude oil reserves, and the environmental impact of fossil fuel usage [24]. Crude oil reserves are being depleted at a rate of approximately 85–90 million barrels of oil per day and it is possible that the reserve will be completely depleted within the next 50 years [5]. Therefore, alternative sources of energy will need to be researched and developed within the next 50 years. Continued research will facilitate the development and implementations of renewable fuels to help lessen the world’s dependence on fossil fuels.

Biodiesel is a biofuel consisting of monoalkyl esters that are derived from organic oils, plant, or animal, through the process of trans-esterification [6]. It is also biodegradable and nontoxic and has low emission profile as compared to petroleum diesel [7]. Biodiesel can be burned in existing diesel engines with no modifications and can be blended in any proportion with petroleum diesel. This allows for its use in the existing fuel distribution infrastructure and giving it value as a fuel extender. Shay [8] reported that algae are one of the best sources of biodiesel. In fact, algae are the highest yielding feedstock for biodiesel. It can produce 250 times more than the amount of oil per acre as soybeans and 7 to 31 time greater oil than palm oil [9]. In fact, biodiesel from algae may be the only way to produce enough automobile fuels to replace current gasoline usage. The best algae for biodiesel would be microalgae [10]. Microalgae have much more oil than macroalgae and it is much faster and easier to grow and harvest [8].

Microalgae can also provide several different types of renewable biofuels. These include methane produced by anaerobic digestion of the algal biomass [11]; biodiesel derived from microalgal oil [1217]; and photobiologically produced biohydrogen [1822]. The idea of using microalgae as a source of fuel is not new [2, 16, 23], but it is now being taken seriously because of the escalating price of petroleum and, more significantly, the emerging concern about global warming that is associated with burning fossil fuels [14].

Depending on species, microalgae produce many different kinds of lipids, hydrocarbons, and other complex oils [12, 24]. Not all algal oils are satisfactory for making biodiesel, but suitable oils occur commonly. Another important fact is that using microalgae to produce biodiesel will not compromise with the production of food, fodder, and other products derived from crops. The objective of this investigation was to identify locally available microalgal strains with sustainable lipid content for biofuel production and evaluate the biofuel productivity of that strain. For this purpose, the present work was carried out with marine microalgae, S. costatum, that are dominant in the coastal area, especially in Cox’s Bazar coast line of Bangladesh shown in Figure 1.

Figure 1: Location of the S. costatum sample collection area in the southeastern region, Bangladesh.

2. Materials and Methods

The present investigation was carried out in the Centre for Research Excellence (CRE), Department of Biochemistry and Molecular Biology, University of Chittagong, Chittagong, Bangladesh.

2.1. Sample Collection

S. costatum sample water was collected from Bakkhali River of Cox’s Bazar (Figure 1), Bangladesh, in January 2014. Microscopic isolation and identification were done at Institute of Marine Sciences and Fisheries, University of Chittagong.

2.2. Culture of Skeletonema costatum

For isolation, standard f/2 medium [25] nutrients were added with the sample water and left under light until diatom bloom. Bloomed S. costatum was then isolated and purified by serial dilution and micropipette method [26, 27]. In micropipette method, single cells or filaments were picked up under a dissecting microscope from the enriched sample water, by using 5 mL syringe. The individual cell was transferred to fresh sterile f/2 medium containing 100 mL flasks and kept under light at 25°C for 1 week. For serial dilution, 5 sterilized test tubes, filled with 9 mL sterilized seawater enriched with f/2 medium, were marked as 10−1 up to 10−5 serially and 1 mL of sample containing bloomed diatom was transferred to 10−1 test tube, then 1 mL from 10−1 to 10−2, and so on. The clones were kept in 24 hours photoperiod to develop for 10 to 15 days as stock culture. For large scale S. costatum production 200 liters FRP tanks were used according to the procedure of Aftab Uddin and Zafar [28]. Cell counts were observed by a compound microscope and concurrently salinity (25–28 ppt), temperature (25–27°C), and pH (7.6–8.2) were maintained during the entire culture period and sufficient aeration was provided.

2.3. Biomass Processing

At first, the biomass was dewatered by filtration with a Whatman filter paper. Then the wet algal biomass was collected from the filter paper with a spatula and taken in Petri dishes. The biomass was then dried at 80°C for 2 hours and kept in the desiccator for whole night so that there is no water left in the sample. The dry mass was then grounded properly and weighed immediately. The drying step was skipped in case of wet extraction process.

2.4. Soxhlet Extraction of Fatty Acid and Lipids

Extensive study and research were conducted earlier to determine the best extraction method over wet and dry algae biomass. In this experiment, the effect of three different solvents, that is, n-hexane [29], the mixture of chloroform and methanol at 2 : 1 v/v ratio [30], and n-hexane and ethanol at 3 : 1 v/v ratio [31], had been studied over wet and dry biomass of S. costatum by using Soxhlet apparatus [32, 33]. For this, 5 g of dry weight and 10 g of wet fresh algae paste were taken into the extraction chamber separately. Both ends of the chamber were enclosed with cotton balls to withstand any solid algae discharge. The bulb was filled with 180 mL of one of the three organic solvents and heated at 80°C. The solvent gradually started to vaporize and after condensation, it fell into the extraction chamber containing the algae and release lipids into the chamber. The solvent-lipid mixture was then reached to a critical height within the chamber and the siphoning process was initialized; that is, the mixture was drawn back to the bulb. The system recirculates the solvent by constantly boiling and condensing it. At the end of this extraction process, a mixture of solvent and oil was left in the bulb and the algae biomass was left in the chamber. The processing before evaporation was performed as described by Kumar et al. [34]. The crude extract was taken into a separating funnel and washed with 1% aqueous sodium chloride solution (50 mL) twice. The aqueous layer was removed and the solvent layer was passed through a layer of anhydrous sodium sulfate by taking it in a glass funnel, blocked with cotton plug.

2.5. Evaporation

The solvent-oil mixture was placed into a preweighed flask for evaporation. In the laboratory setup, the solvent and oil mixture was exposed to a vacuum and then heated at 60°C by using a vacuum pump Rotary Evaporator (RE 200, Bibby Sterling, UK) in order to remove all the solvent. After evaporation, the flask was weighed again and compared with the first weight. In this way, the weight of the oil content was calculated. The oil was recovered by using dichloromethane and was collected into a small vial. The percentage of oil extraction was calculated by the following formula:

2.6. Quantitative Analysis of Algal Oil
2.6.1. TLC Analysis

The components of extracted lipids were separated by modified thin-layer chromatography (TLC) method [35]. This was done by TLC silica gel plates (50 mm × 100 mm in size) and was diffused by two mixed solvents [36]. The first eluent was the mixture of ethyl acetate, isopropanol, chloroform, methanol, and KCl (0.25% solution) in a ratio of 25 : 25 : 25 : 10 : 9 (v/v/v/v/v) running to a height of 10 cm from the origin and the second eluent was the mixture of hexane, diethyl ether, and acetic acid in a ratio of 80 : 20 : 2 (v/v/v) to a height of 18 cm from the origin. After dried, the plate was kept in a TLC jar with iodine powder. The individual lipid bands became detectable as yellow bands within 5 min. In iodine vapor exposure, four individual bands were found for CHCl3 : MeOH (2 : 1) extract, six bands for hexane : EtOH (3 : 1) extract, and a single band for pure hexane were found. For the recovery of individual lipid component, the corresponding silica gel was removed from the plates and washed it with chloroform/methanol (2 : 1 v/v) [36]. The lipid component, separated by TLC as described above, was then kept for the fatty acid composition analysis by GC/MS.

2.6.2. Sample Preparation for GC/MS

The fatty oils were esterified with methanol prior to Gas Chromatography-Mass Spectroscopy (GC/MS) analysis to make the fatty oils more volatile and to avoid the acidic attack to the stationary phase/column. For this, 10 mg (2-3 drops) of fatty oil was taken in a screw capped glass tube and 1 mL of Boron trifluoride methanol complex was added. The mixture was then heated at 100°C for 1 hr using water bath and cooled at 25°C. Then 1 mL of deionized water and 2 mL hexane were added and vortexed for 1 min. Then the mixture was centrifuged at low rpm for 2 min, and upper layer was collected, diluted with hexane, and filtered for GC/MS analysis.

2.6.3. GC/MS Analysis

Thin-layer chromatographic (TLC) separation was analyzed by GC-MS with electron impact ionization (EI) method on a gas chromatograph (GC-17A, Shimadzu Corporation, Kyoto, Japan) coupled to a mass spectrometer (GC-MS QP 5050A, Shimadzu Corporation, Kyoto, Japan). A fused silica capillary column (30 m × 2.5 mm; 0.25 m film thickness) is coated with DB-1 (J&W). The inlet temperature was set at 260°C and the oven temperature was programmed as 70°C (0 min); 10°C, 150°C (5 min); 12°C, 200°C (15 min); 12°C, 220°C (5 min). The column flow rate was 0.6 mL/min He gas at a constant pressure of 90 KPa. The aux (GC to MS interface) temperature was set to 280°C. The MS was set in scan mode with a scanning range of 40–350 amu; the ionization mode was EI (electron ionization) type. The mass range was set in the range of 50–550 m/z. The prepared sample was then run for GC/MS analysis. One μL sample was injected in spilt fewer modes. Vial containing lipid was used for GC/MS analysis to make lipid profile. This lipid profile indicates the fatty acids, which are important to evaluate the biomass for biofuel production.

2.7. Statistical Analysis

All the data were expressed as mean ± SD and they were analyzed by using Microsoft Excel 2007. The significance of difference was calculated by Student’s test and the values were considered to be significant.

3. Results

The result of total oil recovery by three different solvents is shown in Table 1. It showed that every solvent system gives maximum oil recovery for dry sample and relatively low recovery for wet sample. The total oil recoveries obtained from dry and wet biomass by using pure hexane were 11.24% and 2.49%, by CHCl3 : MeOH were 15.37% and 6.70%, and by hexane : EtOH were 11.67% and 5.32%, respectively. The result showed that, in both wet and dry conditions, CHCl3 : MeOH (2 : 1) solvent system gives the highest yield.

Table 1: Percentage of total oil recovered by the three different solvents.

The comparison of oil content and % oil contents from dry and wet sample of different solvent extracts is shown in Figure 2. It is clearly evident that both oil content and % oil content of dry wet from CHCl3 : MeOH extract are significantly () different from those of two other solvent systems. Similarly, their contents in wet sample are also higher in CHCl3 : MeOH although the values were not statistically significant (Figure 2).

Figure 2: Comparative oil contents in dry and wet sample of S. costatum due to solvent variation. Data are shown as mean ± SD (a, b, c, and d) with different superscript letters over the bars for both dry and wet samples are significantly different from each other (paired -test, ).

The fatty acid methyl ester (FAME) composition analysis of the extracted oil was very important for the evaluation of a biofuel feedstock. For this purpose, the extracted algae oil was analyzed by GC/MS to obtain the fatty acid profile of S. costatum. The result showed the fatty acid and lipid profile from pure n-hexane extract presented in Table 2, CHCl3 : MeOH extract in Table 3, and hexane : ethanol extract in Table 4. Most importantly, the fatty acids suitable for biodiesel production such as myristic acid (C14:0), palmitic acid (C16:0), palmitoleic acid (C16:1), and stearic acid (C18:0) were the most frequent entities encountered in S. costatum profile.

Table 2: Fatty acid profile of pure n-hexane extract.
Table 3: Fatty acid profile of lipid from CHCl3 : MeOH extract.
Table 4: Fatty acid profile of lipid band no. 1 of Hexane : EtOH extract.

Table 2 showed that myristic acid (C14:0) and palmitic acid (C16:0) were the dominants in pure hexane extract about 35.93% and 27.21%, respectively. The fatty acid profile of CHCl3 : MeOH extract (presented in Tables 3 and 4) showed that the main constituents of this extract are myristic acid (C14:0), palmitic acid (C16:0), and stearic acid (C18:0) up to 36.39%, 25%, and 27.72%, respectively. Palmitoleic acid, oleic acid, arachidonic acid, pentadecanoic acid, eicosapentaenoic acid, and other fatty acids were also present in low content.

4. Discussion

Bioenergy is one of the most important sources that are concerning the scientists and industrial sector. In view of ever increasing global demand for energy, there has been substantial interest in developing renewable biologically produced fuel. Marine algae are one such emerging resource considered as an alternative for biofuel production. Oil extracted from algal biomass can be turned into biodegradable and carbon neutral biodiesel through transesterification reaction [2, 37].

To be an ideal source of sustainable biodiesel, selected microalgal species should contain sufficient lipid with suitable fatty acids for good biodiesel properties. Several works were carried out to extract algal oil for biofuel production, commonly using different organic solvents, and the lipid contents were often reported as the weight ratio of the crude extract and the dry biomass. In this study, the algal strains were cultured and their oil extraction was done by using three different solvent systems, pure hexane [29], CHCl3 : MeOH [35], and hexane : EtOH [31] to compare their oil recovery. The result showed that every solvent system recovered their highest yield from dry biomass and in both dry and wet conditions, CHCl3 : MeOH (2 : 1) and n-hexane : EtOH (3 : 1) were comparatively more effective than pure n-hexane recovering about 15.367% and 11.673% oil, respectively. However, the oil content in the dry sample through CHCl3 : CH3OH is significantly () different from that of two other solvent systems. This may be because CHCl3 : MeOH (2 : 1) and n-hexane : EtOH (3 : 1) both are the mixture of polar (MeOH, EtOH) and nonpolar (CHCl3, n-hexane) solvents; thus both neutral and polar lipids could be extracted by these two solvent systems. On the other hand, the nonpolar solvent n-hexane could preferably dissolve only nonpolar lipids in the microalgae. However, it could not dissolve all the oil content from the cells alone.

A systematic analysis of the fatty acid methyl ester (FAME) composition and comparative fuel properties is very important for species selection for biodiesel production. It has been suggested that the higher the degree of unsaturation of the FAMEs of a biodiesel, the higher the tendency of the biodiesel to oxidize. There are, however, other parameters which also define the oxidation stability of the fuel, for example, natural antioxidant and free fatty acid content [3840]. The most common fatty acids of microalgae are palmitic-(hexadecanoic-C16:0), stearic (octadecanoic-C18:0), oleic (octadecenoic-C18:1), linoleic-(octadecadienoic-C18:2), and linolenic-(octadecatrienoic-C18:3) acids [41]. Most algae have only small amounts of eicosapentaenoic acid (EPA) (C20:5) and docosahexaenoic acid (DHA) (C22:6); however, in some species of particular genera, these PUFAs can accumulate in appreciable quantities depending on cultivation conditions [42]. Consistently the oil extract of S. costatum showed the prevalence of methyl palmitate, methyl stearate, and methyl myristate when methyl myristate, methyl palmitate, and methyl 6,9,12-hexadecatrienoate were 35.93%, 27.21%, and 12.36% in the n-hexane fraction respectively. While CHCl3 : MeOH fraction showed methyl palmitate (22.41), methyl N-[(1-naphthyl)methyl]carbamate (14.99), and 1,2-Benzenedicarboxylic acid as prevalent fatty acids. However, in hexane : EtOH fraction, methyl stearate (20.64%) is only the fatty acid present in considerable quantity.

The abovementioned FAs of marine microalgae S. costatum found from the lab-scale examination are dominant FAs which are known to be most common fatty acids with good biofuel properties [39]. Other FAs detected were hexadecatrienoic acid, benzenedicarboxylic acid, oleic acid, arachidonic acid, and eicosapentaenoic acid (EPA). Indeed, most of the microalgae investigated have similar fatty acid profile, but the percentage content of fatty acid for each microalga is very different. This mainly depends on the strain used and culture conditions [43, 44]. All of the microalgae have the same fatty acid profile in chain C16 and C18. Palmitic fatty acid (C16:0) is a predominant fatty acid in most microalgae research. Some critical fuel parameters like oxidation stability, cetane number, iodine value, and viscosity were correlated with the methyl ester composition and structural configuration. From the current literature, it was found that the FAs indicated by the fatty acid profile of S. costatum are of better fuel properties and thus it is likelihood to be a viable fuel source for internal combustion engines.

The present study offers a scientific basis of the use of this microalga as a feedstock for biofuel. Still it is in a preliminary stage that requires further study on other parameters. It also indicates that it may be possible in the future to improve the properties of biodiesel by means of genetic engineering of the parent oils, which could eventually lead to a fuel enriched with a combination of improved fuel properties.

Conflict of Interests

The authors declare that there is no conflict of interests.

Authors’ Contribution

Chowdhury Md. Monirul Hasan and Tania Sharmin have designed the study and performed the experimental works in laboratory. Sheikh Aftabuddin has assisted in the collection and identification of microalgae strains and prepared their growth environment. Mala Khan has provided her laboratory facilities in analyzing the compounds. Chowdhury Md. Monirul Hasan, Sheikh Aftabuddin, Md. Atiar Rahman, and Tania Sharmin drafted the paper. Md. Atiar Rahman has endeavored in data analysis and interpretation of the results. All authors read and approved the final version of the paper.

Acknowledgments

The authors thank the authority of BCSIR, Dhaka, Chairman of the Department of Biochemistry and Molecular Biology, and Director of the Institute of Marine Sciences and Fisheries, University of Chittagong, for providing laboratory facilities and their kind support in augmentation of the research.

References

  1. A. Srivastava and R. Prasad, “Triglycerides-based diesel fuels,” Renewable and Sustainable Energy Reviews, vol. 4, no. 2, pp. 111–133, 2000. View at Publisher · View at Google Scholar · View at Scopus
  2. M. Y. Chisti, “An unusual hydrocarbon,” Journal of Ramsay Society, vol. 27-28, pp. 24–26, 1981. View at Google Scholar
  3. A. Demirbas and M. F. Demirbas, “Importance of algae oil as a source of biodiesel,” Energy Conversion and Management, vol. 52, no. 1, pp. 163–170, 2011. View at Publisher · View at Google Scholar · View at Scopus
  4. P. T. Pienkos and A. Darzins, “The promise and challenges of microalgal-derived biofuels,” Biofuels, Bioproducts and Biorefining, vol. 3, no. 4, pp. 431–440, 2009. View at Publisher · View at Google Scholar · View at Scopus
  5. A. Z. Abdullah, N. Razali, H. Mootabadi, and B. Salamatinia, “Critical technical areas for future improvement in biodiesel technologies,” Environmental Research Letters, vol. 2, no. 3, Article ID 034001, 6 pages, 2007. View at Publisher · View at Google Scholar · View at Scopus
  6. A. Demirbas, “Importance of biodiesel as transportation fuel,” Energy Policy, vol. 35, no. 9, pp. 4661–4670, 2007. View at Publisher · View at Google Scholar · View at Scopus
  7. M. J. Ramos, C. M. Fernández, A. Casas, L. Rodríguez, and Á. Pérez, “Influence of fatty acid composition of raw materials on biodiesel properties,” Bioresource Technology, vol. 100, no. 1, pp. 261–268, 2009. View at Publisher · View at Google Scholar · View at Scopus
  8. E. G. Shay, “Diesel fuel from vegetable oils: status and opportunities,” Biomass and Bioenergy, vol. 4, no. 4, pp. 227–242, 1993. View at Publisher · View at Google Scholar · View at Scopus
  9. A. B. M. S. Hossain, A. Salleh, A. N. Boyce, P. Chowdhury, and M. Naqiuddin, “Biodiesel fuel production from algae as renewable energy,” American Journal of Biochemistry and Biotechnology, vol. 4, no. 3, pp. 250–254, 2008. View at Publisher · View at Google Scholar · View at Scopus
  10. A. K. Bajhaiya, S. K. Mandotra, M. R. Suseela, K. Toppo, and S. Ranade, “Algal biodiesel: the next generation biofuel for India,” Asian Journal of Experimental Biology, vol. 1, no. 4, pp. 728–739, 2010. View at Google Scholar
  11. P. Spolaore, C. Joannis-Cassan, E. Duran, and A. Isambert, “Commercial applications of microalgae,” Journal of Bioscience and Bioengineering, vol. 101, no. 2, pp. 87–96, 2006. View at Publisher · View at Google Scholar · View at Scopus
  12. A. Banerjee, R. Sharma, Y. Chisti, and U. C. Banerjee, “Botryococcus braunii: a renewable source of hydrocarbons and other chemicals,” Critical Reviews in Biotechnology, vol. 22, no. 3, pp. 245–279, 2002. View at Publisher · View at Google Scholar · View at Scopus
  13. T. G. Dunahay, E. E. Jarvis, S. S. Dais, and P. G. Roessler, “Manipulation of microalgal lipid production using genetic engineering,” Applied Biochemistry and Biotechnology, vol. 57-58, pp. 223–231, 1996. View at Publisher · View at Google Scholar · View at Scopus
  14. M. Gavrilescu and Y. Chisti, “Biotechnology—a sustainable alternative for chemical industry,” Biotechnology Advances, vol. 23, no. 7-8, pp. 471–499, 2005. View at Publisher · View at Google Scholar · View at Scopus
  15. P. G. Roessler, L. M. Brown, T. G. Dunahay et al., “Genetic engineering approaches for enhanced production of biodiesel fuel from microalgae,” in Enzymatic Conversion of Biomass for Fuels Production, vol. 566 of ACS Symposium Series, pp. 255–270, American Chemical Society, 1994. View at Publisher · View at Google Scholar
  16. S. Sawayama, S. Inoue, Y. Dote, and S.-Y. Yokoyama, “CO2 fixation and oil production through microalga,” Energy Conversion and Management, vol. 36, no. 6-9, pp. 729–731, 1995. View at Publisher · View at Google Scholar · View at Scopus
  17. J. Sheehan, T. Dunahay, J. Benemann, and P. Roessler, “A look back at the U.S. Department of Energy's Aquatic Species Program-biodiesel from algae,” Report NREL/TP-580-24190, National Renewable Energy Laboratory, Golden, Colo, USA, 1998. View at Google Scholar
  18. I. Akkerman, M. Janssen, J. Rocha, and R. H. Wijffels, “Photobiological hydrogen production: photochemical efficiency and bioreactor design,” International Journal of Hydrogen Energy, vol. 27, no. 11-12, pp. 1195–1208, 2002. View at Publisher · View at Google Scholar · View at Scopus
  19. A. S. Fedorov, S. Kosourov, M. L. Ghirardi, and M. Seibert, “Continuous hydrogen photoproduction by Chlamydomonas reinhardtii: using a novel two-stage, sulfate-limited chemostat system,” Applied Biochemistry and Biotechnology, vol. 121, no. 1–3, pp. 403–412, 2005. View at Publisher · View at Google Scholar · View at Scopus
  20. M. L. Ghirardi, L. Zhang, J. W. Lee et al., “Microalgae: a green source of renewable H2,” Trends in Biotechnology, vol. 18, no. 12, pp. 506–511, 2000. View at Publisher · View at Google Scholar · View at Scopus
  21. I. K. Kapdan and F. Kargi, “Bio-hydrogen production from waste materials,” Enzyme and Microbial Technology, vol. 38, no. 5, pp. 569–582, 2006. View at Publisher · View at Google Scholar · View at Scopus
  22. A. Melis, “Green alga hydrogen production: progress, challenges and prospects,” International Journal of Hydrogen Energy, vol. 27, no. 11-12, pp. 1217–1228, 2002. View at Publisher · View at Google Scholar · View at Scopus
  23. N. Nagle and P. Lemke, “Production of methyl-ester fuel from microalgae,” Applied Biochemistry and Biotechnology, vol. 24-25, pp. 355–361, 1990. View at Publisher · View at Google Scholar · View at Scopus
  24. P. Metzger and C. Largeau, “Botryococcus braunii: a rich source for hydrocarbons and related ether lipids,” Applied Microbiology and Biotechnology, vol. 66, no. 5, pp. 486–496, 2005. View at Publisher · View at Google Scholar · View at Scopus
  25. R. R. L. Guillard, “Culture of phytoplankton for feeding marine invertebrates,” in Culture of Marine Invertebrate Animals, pp. 29–60, Plenum Press, New York, NY, USA, 1975. View at Google Scholar
  26. E. J. Allen and E. W. Nelson, “The artificial culture of marine plankton organisms,” Journal of the Marine Biological Association of the UK, vol. 8, no. 5, pp. 421–474, 1910. View at Google Scholar
  27. M. R. Droop, “A note on the isolation of small marine algae and flagellates for pure cultures,” Journal of the Marine Biological Association of the United Kingdom, vol. 33, no. 2, pp. 511–514, 1954. View at Publisher · View at Google Scholar
  28. S. Aftab Uddin and M. Zafar, “Live feed production in different nutritive condition as diet for Penaeus monodon in Shrimp Hatchery, Bangladesh,” International Journal of Agriculture and Biology, vol. 8, no. 4, pp. 493–495, 2006. View at Google Scholar
  29. X. Miao and Q. Wu, “Biodiesel production from heterotrophic microalgal oil,” Bioresource Technology, vol. 97, no. 6, pp. 841–846, 2006. View at Publisher · View at Google Scholar · View at Scopus
  30. J. Folch, M. Lees, and G. H. Sloane Stanley, “A simple method for the isolation and purification of total lipides from animal tissues,” Journal of Biology and Chemistry, vol. 226, no. 1, pp. 497–509, 1956. View at Google Scholar · View at Scopus
  31. M. Chen, T. Liu, X. Chen et al., “Subcritical co-solvents extraction of lipid from wet microalgae pastes of Nannochloropsis sp.,” European Journal of Lipid Science and Technology, vol. 114, no. 2, pp. 205–212, 2012. View at Publisher · View at Google Scholar
  32. R. G. Ackman, “Extraction and analysis of omega-3 fatty acids: procedures and pitfalls,” in Omega-3 Fatty Acids: Metabolism and Biological Effects, C. A. Drevon, I. Baksaas, and H. E. Krokan, Eds., pp. 11–20, Birkhäuser, Basel, Switzerland, 1993. View at Google Scholar
  33. H. Gunnlaugsdottir and R. G. Ackmanj, “Three extraction methods for determination of lipids in fish meal: evaluation of a hexane/isopropanol method as an alternative to chloroform-based methods,” Journal of the Science of Food and Agriculture, vol. 61, no. 2, pp. 235–240, 1993. View at Publisher · View at Google Scholar · View at Scopus
  34. P. Kumar, M. R. Suseela, and K. Toppo, “Physico-chemical characterization of algal oil: a potential biofuel,” Asian Journal of Experimental Biological Sciences, vol. 2, no. 3, pp. 493–497, 2011. View at Google Scholar
  35. A. Vieler, C. Wilhelm, R. Goss, R. Süß, and J. Schiller, “The lipid composition of the unicellular green alga Chlamydomonas reinhardtii and the diatom Cyclotella meneghiniana investigated by MALDI-TOF MS and TLC,” Chemistry and Physics of Lipids, vol. 150, no. 2, pp. 143–155, 2007. View at Publisher · View at Google Scholar · View at Scopus
  36. E. Ryckebosch, K. Muylaert, and I. Foubert, “Optimization of an analytical procedure for extraction of lipids from microalgae,” Journal of the American Oil Chemists' Society, vol. 89, no. 2, pp. 189–198, 2012. View at Publisher · View at Google Scholar · View at Scopus
  37. J. Jones, S. Manning, M. Montoya, K. Keller, and M. Poenie, “Extraction of algal lipids and their analysis by HPLC and mass spectrometry,” Journal of the American Oil Chemists' Society, vol. 89, no. 8, pp. 1371–1381, 2012. View at Publisher · View at Google Scholar · View at Scopus
  38. L. C. Meher, D. V. Sagar, and S. N. Naik, “Technical aspects of biodiesel production by transesterification—a review,” Renewable and Sustainable Energy Reviews, vol. 10, no. 3, pp. 248–268, 2006. View at Publisher · View at Google Scholar · View at Scopus
  39. S. K. Hoekman, A. Broch, C. Robbins, E. Ceniceros, and M. Natarajan, “Review of biodiesel composition, properties, and specifications,” Renewable and Sustainable Energy Reviews, vol. 16, no. 1, pp. 143–169, 2012. View at Publisher · View at Google Scholar · View at Scopus
  40. G. Knothe, “Dependence of biodiesel fuel properties on the structure of fatty acid alkyl esters,” Fuel Processing Technology, vol. 86, no. 10, pp. 1059–1070, 2005. View at Publisher · View at Google Scholar · View at Scopus
  41. M. Lapuerta, J. Rodríguez-Fernández, and E. Font de Mora, “Correlation for the estimation of the cetane number of biodiesel fuels and implications on the iodine number,” Energy Policy, vol. 37, no. 11, pp. 4337–4344, 2009. View at Publisher · View at Google Scholar · View at Scopus
  42. R. Huerlimann, R. de Nys, and K. Heimann, “Growth, lipid content, productivity, and fatty acid composition of tropical microalgae for scale-up production,” Biotechnology and Bioengineering, vol. 107, no. 2, pp. 245–257, 2010. View at Publisher · View at Google Scholar · View at Scopus
  43. I. Tzovenis, N. De Pauw, and P. Sorgeloss, “Optimation of T-ISO biomass production rich in essential fatty acids, II. Effect of different light regimes on growth and biomass production,” Aquaculture, vol. 216, no. 1–4, pp. 223–242, 2003. View at Publisher · View at Google Scholar
  44. S. M. Renaud, L.-V. Thinh, G. Lambrinidis, and D. L. Parry, “Effect of temperature on growth, chemical composition and fatty acid composition of tropical Australian microalgae grown in batch cultures,” Aquaculture, vol. 211, no. 1–4, pp. 195–214, 2002. View at Publisher · View at Google Scholar · View at Scopus