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When a fluid is subjected to temperatures and pressures above its critical temperature and pressure, it becomes a supercritical fluid. Supercritical fluid extraction (SFE) is the process of extracting oil from oil-containing materials using a supercritical fluid as the extraction solvent. The advantage of supercritical fluids used in oil extraction is their increased solvating power (Mercer and Armenta, 2011). Factors to consider when selecting an SFE solvent include that the solvent is nonflammable, nontoxic, has low critical parameters, good solvating properties, is easily separated from product, and is environmentally friendly and inexpensive. The added advantages of SFE over conventional solvent extraction are that it provides simple and flexible process control of temperature, shorter extraction times, low cost, and solvent-free product. The SFE consists of an SFE solvent tank, solvent and feed pumps, a high-pressure pump, extraction vessels, and restrictor and absorbent vessels. Carbon dioxide (CO2) is widely used as a solvent in SFE due to its moderate critical temperature (31.1°C) and pressure (72.9 atm) (Cooney et al., 2009). In this method, CO2 is used as the extracting solvent when it is in a supercritical state (i. e., it has both gas and liquid properties). The supercritical state of CO2 can be achieved by liquefying CO2 under higher pressure and heating to a particular temperature. The important operating parameters considered for optimizing the extraction efficiency of this method are operating temperature and pressure, quantity of CO2 supplied, feed particle size, and residence time. Dried algae paste must be used for supercritical extraction; this helps in increasing the contact time between the SFE solvent and the algae paste. CO2 acts as a gas in air at ambient temperature, and can be removed after the
Comparison of different oil extraction methods
TABLE 7.2
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extraction and reused again for further extractions. A comparison of three different oil extraction methods used for lipid recovery is provided in Table 7.2.
Microalgae are becoming more attractive feedstocks for biodiesel production as higher oil-yielding algae have the potential to replace conventional biodiesel feedstocks. The viability of microalgae oil-based biodiesel production primarily depends on the identification of appropriate higher lipid producing algal strains. From preliminary studies on lipid analysis for identifying algae, important fuel properties of the algal oil can be predicted and compared with biodiesel standards. The simplest method of assessing the fuel quality of biodiesel is predicting its fuel properties based on the fatty acid composition of algal oil, thereby allowing us to ascertain the suitability of selected algae strains for biodiesel production. The economical/technical viability of microalgal oil-based biodiesel production depends on implementation of the suitable technologies used in the downstream processing of microalgae.
Allard, B., Rager M. N., and Templier, J. (2002). Occurrence of high molecular weight lipids (c80+) in the trilaminar outer cell walls of some freshwater microalgae. A reappraisal of algaenan structure. Organic Geochemistry, 33: 789-801.
Anderson, V. D. (1900). Press, U. S. Patent 647, 354.
ASTM Standard Specification for Biodiesel Fuel (B100) Blend Stock for Distillate Fuels, (2008). In Annual Book of ASTM Standards, ASTM International, West Conshohocken, PA, Method D6751-08.
Belarbi, E. H., Molina, E., and Chisti, Y. (2000). A process for high yield and scalable recovery of high purity eicosapentaenoic acid esters from microalgae and fish oil. Enzyme Microbiology and Technology, 26: 516-529.
Brennan, L., and Owende, P. (2010). Biofuels from microalgae. A review of technologies for production, processing, and extractions of biofuels and co-products. Renewable and Sustainable Energy Review, 14: 557-577.
Chisti, Y. (2007). Biodiesel from microalgae. Biotechnology Advances, 25: 294-306.
Cooney, M., Young, G., and Nagle, N. (2009). Extraction of bio-oils from microalgae. Separation and Purification Reviews, 38: 291-325.
De Boer, K., Moheimani, N. R., Borowitzka, M. A., and Bahri, P. A. (2012) Extraction and conversion pathways for microalgae to biodiesel: A review focused on energy consumption. Journal of Applied Phycology, 24: 1681-1698.
Demirbas, A. (2009). Production of biodiesel from algae oils. Energy Sources, Part A, 31: 163-168.
Demirbas, A., and Demirbas, M. F. (2010). Algae Energy: Algae as a New Source of Biodiesel. Springer London Ltd., United Kingdom, p. 143.
EN Committee for Standardization Automotive Fuels (2003). Fatty Acid Methyl Esters (FAME) for Diesel Engines—Requirements and Test Methods. European Committee for Standardization, Brussels. Method EN 14214.
Erickson, R. D., Pryde, H. E., Brekke, L. O., Mounts, L. T., and Falb, A. R. (1984). Handbook of Soy Oil Processing and Utilization. American Oil Chemists Society, Champaign, IL, p. 598.
Hamm, W., and Hamilton, J. R. (2000). Edible Oil Processing. Boca Raton, FL: CRC Press LLC, p. 281.
Harwood, J. L., and Guschina, I. A. (2009). The versatility of algae and their lipid metabolism. Biochimie, 91: 679-684.
Hu, Q., Sommerfeld, M., Jarvis, E., Ghirardi, M. L., Posewitz, M. C., and Seibert, M. (2008). Microalgal triacylglycerols as feedstocks for biofuel production. The Plant Journal, 54: 621-639.
Johnson, L. A., and Lusas, E. W. (1983). Comparison of alternative solvents for oils extraction. Journal of the American Oil Chemists Society, 60: 229-242.
Kalayasiri, P., Jayashke, N., and Krisnangkura, K. (1996). Survey of seed oils for use as diesel fuels. Journal of the American Oil Chemists Society, 73: 471-474.
Krisnangkura, K. (1986). A simple method for estimation of cetane index of vegetable oil methyl esters. Journal of the American Oil Chemists Society, 63: 552-553.
Lee, J., Yoo, C., Jun, S., Ahn, C., and Hee-Mock, O. (2010). Comparison of several methods for effective lipid extraction from microalgae. Bioresource Technology, 101(Suppl. 1): S75-S77.
Letellier, M., and Budzinski, H. (1999). Microwave assisted extraction of organic compounds. Analusis, 27: 259.
Lewis, T., Nichols, P. D., and McMeekin, T. A. (2000). Evaluation of extraction methods for recovery of fatty acids from lipid producing micro-heterotrophs. Journal of MicrobiologicalMethods, 43: 107-116.
Lin, Y. H., Chang, F. L., Tsao, C. Y., and Leu, J. Y. (2007). Influence of growth phase and nutrient source on fatty acid composition of Isochrysis galbana CCMP 1324 in a batch photoreactor. Biochemical Engineering Journal, 37: 166-176.
Macias-Sanchez, M. D., Mantell, C., Rodriguez, M., Martinez de la Ossa, E., Lubian, L. M., and Montero, O. (2005). Supercritical fluid extraction of carotenoids and chlorophyll a from Nannochloropsis gaditana. Journal of Food Engineering, 66: 245-251.
Mansour, M. P. (2005). Reversed-phase high-performance liquid chromatography purification of methyl esters of C16-C28 polyunsaturated fatty acids in microalgae, including octa- cosaoctaenoic acid [28:8(n-3)]. Journal of Chromatography A, 1097: 54-58.
Mercer, P., and Armenta, R. E. (2011). Developments in oil extraction from microalgae. European, Journal of Lipid Science and Technology, 113: 539-547.
Mittelbach, M. (1996). Diesel fuel derived from vegetable oils. VI. Specifications and quality control of biodiesel. Bioresource Technology, 56: 7-11.
Mutanda, T., Ramesh, D., Karthikeyan, S. Kumari, S., Anandraj A., and Bux, F. (2011). Bioprospecting for hyper-lipid producing microalgal strains for sustainable biofuel production. Bioresource Technology, 102: 57-70.
Paik, M. J., Kim, H., Lee, J., Brand, J., and Kim, K. R. (2009). Separation of triacylglycer — ols and free fatty acids in microalgal lipids by solid phase extraction for separate fatty acid profiling analysis by gas chromatography. Journal of Chromatography A, 1216: 5917-5923.
Pawliszyn, J. (1993). Kinetic model of supercritical fluid extraction. Journal of Chromatography Science, 31: 31-37.
Popoola, T. O.S., and Yangomodou, O. D. (2006). Extraction, properties and utilization potentials of cassava seed oil. Biotechnology, 5: 38-41.
Ramos, M. J., Fernandez, C. M., Casas, A., Rodriguez, L., and Perez, A. (2009). Influence of fatty acid composition of raw materials on biodiesel properties. Bioresource Technology, 100: 261-268.
Richmond A. (2004). Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell Science Ltd., Malden, MA.
Sahena, F., Zaidul, I. S.M., Jinap, S., Karim, A. A., Abbas, K. A., Norulaini, N. A.N., and Omar, A. K.M. (2009). Application of supercritical CO2 in lipid extraction — A review. Journal of Food Engineering, 95: 240-253.
Sander, K., and Murthy G. S. (2009). Enzymatic degradation of microalgal cell walls. ASABE Annual International Meeting, Sponsored by ASABE Grand Sierra Resort and Casino, Reno, Nevada, June 21-June 24, 2009. Paper number 1035636.
Singh, J., and Gu, S. (2010). Commercialization potential of microalgae for biofuels production. Renewable and Sustainable Energy Reviews, 14: 2596-2610.
Versteegh, G. J.M., and Blokker, P. (2004). Resistant macromolecules of extant and fossil microalgae. Phycology Research, 52: 325-339.
Apart from the key algal compound groupings, mentioned above, there are new market sectors and applications emerging in algal biotechnology.
10.2.4.1 Cosmetic Extracts
Marine microalgae contribute to a range of extracts rich in proteins, vitamins, and minerals, which are incorporated as active ingredients into a number of cosmetic products (Kim et al., 2008). In addition to carotenoids, phycobiliproteins, and PUFAs, microalgae produce a number of other compounds (that exhibit a range of benefits) appealing to cosmetic formulators (Table 10.8).
These compounds prevent blemishes, repair damaged skin, aid in the treatment of seborrhoea (greasy skin caused by excess sebum), and inhibit the inflammation process (Kim et al., 2008). They are formulated into face and skin care products,
TABLE 10.8
Microalgal Compounds and Their Cosmeceutical Properties
Cosmeceutical Properties
Skin protection against UV radiation Antioxidant
Protection against UV irradiation or oxidative damage
Prevention of light-induced pathologies of the human skin and eyes
Prevention of degenerative disorders (atherosclerosis, cardiovascular disease, and cancer)
Antioxidative action
Protection against UV irradiation
Antioxidant
Emmolient
Blood stimulant
Diuretic
Moisturizing activities
Source: Adapted from Kim et al. (2008).
such as anti-aging creams and moisturizers, sun protection products, hair care products, refresherant or regenerant care products, emollients, and anti-irritant skin peels (Spolaore et al., 2006; Carlsson et al., 2007). Arthrospira (Spirulina) and Chlorella sp. are the two main genera that have established positions in the skin care market (Table 10.9).
The LVMH Group (Louis Vuitton and Moet Hennessey) (Paris, France) and Danial Jouvance (Carnac, France) have both invested in microalgal production systems (Spolaore et al., 2006; and Kim et al., 2008). It is evident that the largest market for micro — and macroalgal cosmetics is in France, with a demand estimated at 5,000 tonnes (Kim et al., 2008). This demand will continue to escalate, with the cosmetic industry evincing more interest as research and extensive studies progressively highlight the benefits of microalgal extracts on skin health.
TABLE 10.9 Cosmetic Companies Producing Commercial Products Formulated with
Source: Adapted from Spolaore et al. (2006) and Kim et al. (2008). |
10.2.4.2 Stable Isotope Biochemicals
Microalgae are also well suited to produce isotopically labeled compounds due to their ability to incorporate stable isotopes from inexpensive inorganic molecules into high-value isotopic organic chemicals. The ability to cultivate phototropic algae under strictly controlled conditions enables the easy incorporation of stable isotopes from inorganic carbon, hydrogen, and nitrogen sources (Pulz and Gross, 2004; Spolaore et al., 2006; Milledge, 2011). These stable isotopic compounds are used to facilitate the structural determination (at atomic level) of proteins, carbohydrates, and nucleic acids. In addition to metabolic studies (Spolaore et al., 2006), they can also be employed for clinical purposes such as gastrointestinal or breath diagnosis tests (Radmer, 1996; Pulz and Gross, 2004). Table 10.10 indicates some of the isotopically labeled microalgal products.
The market value of these compounds is estimated at US$13 million per year. A major distributor of such isotopic compounds is Spectra Stable Isotopes (Andover, MA; acquired by Cambridge Isotope Laboratories [CIL] in 2008) (Spolaore et al., 2006).
Innovative ways to optimize maximum microalgal biomass production and technological advances for transesterification would be necessary to make microalgae more cost effective for biodiesel production and to sustain an economically viable microalgal biotechnological industry (Figure 13.1). Improvements at various intermediary stages of culturing, selection of strains of algae, harvesting, and extraction of bio-fuel production and co-products could bring down the production costs. Norsker et al. (2011) state that by optimizing irradiation conditions, mixing, photosynthetic
FIGURE 13.1 Schematic model for microalgal biotechnology. |
efficiency, growth media, and CO2 costs, the overall cost of production could be reduced to Euro 0.68 per kilogram, which would be economically acceptable for using algae as feedstock for biodiesel and chemicals. Alternative metabolic pathways such as heterotrophy and mixotrophy should be explored to maximize algal growth without a shift to energetically inefficient metabolism. Service (2008) states that algae grown in dark stainless steel fermenters convert sugars to oils more efficiently. Heterotrophic and mixotrophic cultivation of microalgae in fermentation systems for commercial viability should be explored (Gladue and Maxey, 1994; Xu et al., 2006; Chi et al., 2007).
Microalgae hold great potential as a source of cheap and environmentally friendly biofuel. The total annual production of microalgae in 2004 was 5,000 tons, with global sales worth about US$1.25 billion (Pulz and Gross, 2004). However, we believe that comprehensive evaluation of select species from an integrated perspective would be of greatest benefit to commercial operations. Although Serrano’s quote (Serrano, 2010) that, “We are still like the Wright Brothers, putting pieces of wood and paper together” is in a different context, it is apt here. The rigor of microalgal biofuel research, coupled with its interdisciplinary nature, suggests that a comprehensive modeling strategy, one that accounts for numerous culture and harvest parameters and optimizes industrial processes from a perspective of cost, would be of great value. Simulation models that incorporate elements of nutrient systems, ideal culture conditions, and harvest of multiple products such as fuels and high-value nutraceu — ticals and/or recombinant proteins would be instrumental in the development of a viable bio-economy. Brown (2009) pointed out that as mass cultivation of algae for biofuels per se may not sustain microalgal technology, attention should be paid to non-fuel products and co-products as well. These co-products include carotenoids, phycolbiliproteins, astaxanthin, and eicosapentaenoic acid; additionally, algal biomass waste could be used as fertilizer (Donovan and Stowe, 2009).
Various processes are involved in this modeling activity. As Malcata (2010) observed, modeling exercises, instead of empirical approaches, should have biological meaning for which specific experimental data should be obtained on the optimum versus enhanced growth, metabolic cycles, assimilation efficiencies, that is, conversion of substrate into reserves, accumulation, and product sysnthesis/excretion. Scott et al. (2010) commented that there is an inadequacy of established background knowledge in this area, and there is a need to integrate biology and engineering.
The central theme rests on the predictive aspects of modeling that enable one to determine the exact quantities of the envisaged end product together with coproducts. To estimate the actual quantities, we require appropriate input data regarding culture conditions, harvest efficiencies, and yield of co-products, as outlined above. The effective price for the microalgae-derived biofuels can be calculated by optimizing the cost functional involving several variables under appropriately formulated constraints. Results obtained from all stages of the process constitute the vital parameters in the mathematical model. As the process is dynamic in character, time delays do occur in a natural way, and these delays account for the process lead-time. We need to estimate these time delays, maintaining the stability of the corresponding delay-free systems. Division rates of the reaction mechanisms play a vital role in the process of restoring and/or maintaining the stability of the processes. Simulations based on realistic data will grossly help in the validation of these models. Thoroughly validated models are utilized for predicting the optimal cost of biofuel under conditions where lipid yields are maximum.
Williams and Laurens (2010) argued that a fundamental change in the approaches to production is needed, and that “biofuel-only” options may not be economically viable. They showed that 30% to 50% of primary production is lost in the production of protein and lipid, and that if lipid production is increased, then production of other valuable co-products is reduced. These authors argue that the availability of nutrients such as phosphorus and nitrogen, delivery of CO2, and the energy costs associated with sterilization and recycling of spent culture water and removal of biological contaminants, pathogens, and predators would escalate production of microalgal biomass and could be “show-stoppers.”
In conclusion, microalgal biotechnology has made rapid advances in the mass cultivation of algae and their application toward biofeed, biopharmacy, biofuel, bioremediation, bioactive compounds, and space research. However, fewer than fifty species are utilized, while thousands remain unexplored. The potential roles of microalgae in genetic engineering and nanotechnology have increased the prospects for the next generation of “designer microalgae.” To establish algal biotechnology as an economically viable enterprise, concerted research is needed to (1) develop inexpensive media through enrichment of wastewater; (2) isolate and culture new strains of high-yielding microalgae, preferably a consortium of extremophiles; (3) improve production systems; (4) enhance biochemical and metabolic pathways through genetic engineering; and (5) improve harvesting techniques.
Additionally, attention should be given to high-value natural and recombinant products that can be extracted from algae to enhance the profitability of biofuel operations. Simulation models will serve as the foundation for industrial processes that optimize wastewater treatment systems, nutrient levels, and strategies for harvest and extraction of bioactive compounds. A robust bio-economy built on a platform of innovative microalgal technologies requires a cross-disciplinary approach among biologists, biotechnologists, molecular biologists, biochemists, engineers, chemists, bioreactor manufacturers, aquaculturists, and modelers.
We are grateful to Professor Faizal Bux and Dr. Taurai Mutanda, Institute for Water and Wastewater Technology, Durban University of Technology, South Africa, for inviting us to contribute this chapter. We are most grateful to Professor John Beardall, Monash University, Clayton, Victoria, Australia, for constructive review of the manuscript. We thank Bala T. Durvasula and Dr. Ivy Hurwitz for their help with formatting the manuscript.
The research of V. Sree Hari Rao is supported by the Foundation for Scientific Research and Technological Innovation (FSRTI), a Constituent Division of the Sri Vadrevu Seshagiri Rao Memorial Charitable Trust, Hyderabad, India.
Alabi, A. O. (2009). Microalgal Technologies and Processes for Biofuels/Bioenergy Production in British Columbia. The British Columbia Innovation Council, Vancouver, pp. 1-74.
Allen, E. J., and Nelson, E. W. (1910). On the artificial culture of marine plankton organisms. Journal of the Marine Biological Association of the United Kingdom, 8: 421-474.
Andersen, R. A. (2005). Algal Culturing Techniques. Academic Press, Amsterdam.
AquaticBiofuel. com (2008). 2008 the Year of Algae Investments (December 5, 2008), accessed June 30, 2009, http://aquaticbiofuel. com/2008/12/05/2008-the-year-of-algae-investments/.
Banerjee, A., Sharma, R., Chisti, Y., and Banerjee, U. C. (2002). Botryococcus braunii: A renewable source of hydrocarbons and other chemicals. Critical Reviews in Biotechnology, 22: 245-279.
Basova, M. K. (2005). Fatty acid composition of lipids in microalgae. International Journal on Algae, 7: 33-57.
Benemann, J. R. (2008). NREL-AFOSR Workshop, Algal Oil for Jet Fuel Production; Arlington, VA, 19 February 2008.
Benemann, J. R. (1993). Utilization of carbon dioxide from fossil fuel-burning plants with biological systems. Energy Conversion and Management, 34: 999-1004.
Ben-Amoz, A. (2009). Bioactive compounds: Glycerol production, carotenoid production, fatty acids production. In The Alga Dunaliella Biodiversity, Physiology, Genomics and Biotechnology (Eds. Ben-Amotz, A., Polle, J. E.W., and Subba Rao, D. V.). Science Publishers, Enfield, NH, pp. 189-207.
Brown, M. R. (1991). The amino acid and sugar composition of 16 species of microalgae used in mariculture. Journal of Experimental Marine Biology and Ecology, 145: 79-99.
Brown, P. (2009). Algal Biofuels Research, Development, and Commercialization Priorities: A Commercial Economics Perspective. Energy Overviews. [Online] ep Overviews Publishing, Inc, 22. 06. 2009. http://www. epoverviews. com/oca/Algae%20Biofuel%20 Development%20Prioritie.
Cardon, Z. G., Gray, D. W., and Lewis, L. A. (2008). The green algal underground: Evolutionary secrets of desert cells. Bioscience, 58: 114-122.
Casal, C. C., Cauresma, M. M., Vega, M. J.M., and Vilches, C. C. (2011). Enhanced productivity of a lutein-enriched novel acidophile microalga grown on urea. Marine Drugs, 9: 29-42.
Cauresma, M., Casal, C., Forjanb, E., and Vilches, C. C. (2011). Productivity and selective accumulation of carotenoids of the novel extremophile microalga Chlamydomonas acidophila grown with different carbon sources in batch systems. Journal of Industrial Microbiology and Biotechnology, 38: 167-177.
Chang, E. H., and Yang, S. S. (2003). Some characteristics of microalgae isolated in Taiwan for biofixation of carbon dioxide. Botanical Bulletin of Academia Sinica, 44: 43-52.
Chen, C. Y., Yeh, K. L., Aisyah, R., Lee, D. J., and Chang, J. S. (2011). Cultivation, photobioreactor design and harvesting of microalgae for biodiesel production: A critical review. Bioresource Technology, 10: 71-81.
Chi, Z. Y., Pyle, D., Wen, Z. Y., Frear, C., and Chen, S. L. (2007). A laboratory study of producing docosahexaenoic acid from biodiesel-waste glycerol by microalgal fermentation. Process Biochemistry, 42: 1537-1545.
Chisti, Y. (2007). Biodiesel from microalgae. Biotechnology Advances, 25: 294-306.
Chiu, S., Kao, C. C., Tsai, M., Ong, S., Chen, C., and Lin, C. (2009). Lipid accumulation and CO2 utilization of Nannochloropsis oculata in response to CO2 aeration. Bioresource Technology, 100: 833-838.
Coesel, S. N., Baumgartner, A. C., Teles, U. M., Ramo, A. A., and Henriques, N. M. (2008). Nutrient limitation is the main regulatory factor for carotenoid accumulation and for Psy and Pds steady state transcript levels in Dunaliella salina (Chlorophyta) exposed to high light and salt stress. Marine Biotechnology, 10: 602-611.
Converti, A., Casazza, A., Ortiz, E., Perego, P., and Borghi, M. (2009). Effect of temperature and nitrogen concentration on the growth and lipid content of Nannochloropsis oculata and Chlorella vulgaris for biodiesel production. Chemical Engineering and Processing, 48: 1146-1151.
Coronet, J. F. (2010). Calculation of optimal design and ideal productivities of volumetrically lightened photo bioreactors using the constructal approach. Chemical Engineering Science, 65: 985-998.
Courchesne, N. M.D., Parisien, A., Wang, B., and Lan, C. Q. (2009). Enhancement of lipid production using biochemical, genetic and transcription factor engineering approaches. Journal of Biotechnology, 141: 31-41.
Craggs, R. J., Sukias, J. P., Tanner, C. T., and Davies-Colley, R. J. (2004). Advanced pond system for diary-farm effluent treatment. New Zealand Journal of Agricultural Research, 47: 449-460.
Craggs, R. J., Heubeck, S., Lundquist, T. J., and Benemann, J. R. (2011). Algal biofuels from wastewater treatment high rate algal ponds. Water Science and Technology, 63: 660-665.
Cysewski, G. R., and Lorenz, R. T. (2004). Industrial production of microalgal cell-mass and secondary products—Species of high potential: Haematococcus. In Handbook of Microalgal Culture. Wiley-Blackwell, United Kingdom, pp. 281-288.
Day, J. G., and Harding, K. K. (2008). Cryopreservation of algae. In Plant Cryopreservation: A Practical Guide Biomedical and Life Sciences. Plant Cryopreservation Section II (Ed. B. M. Reed), Springer, New York, pp. 95-116. doi: 10.1007/978-0-387-72276-4_6.
De Pauw, N., Morales, J., and Persoone, G. (1984). Mass culture of microalgae in aquaculture systems: Progress and constraint. Hydrobiologia, 116/117: 121-134.
Duarte, P., and Subba Rao, D. V. (2009). Photosynthesis — Energy relationships in Dunaliella. In The Alga Dunaliella Biodiversity, Physiology, Genomics and Biotechnology (Eds. Ben-Amotz A., Polle, J. E.W., and Subba Rao, D. V.), Science Publishers, Enfield, NH, pp. 209-229.
Dunahay, T. G., Jarvis, E. E., and Roessler, P. G. (1995). Genetic transformation of the diatoms Cyclotella cryptica and Navicula saprophila. Journal of Phycology, 31: 1004-1012.
Dunahay, T. G., Jarvis, E. E., Dai, S. S., and Roessler, P. G. (1996). Manipulation of microalgal lipid production using genetic engineering. Applied Biochemistry and Biotechnology, 57-58(1): 223-231.
Danquah, M. K., Ang, L., Uduman, N., Moheimani, N., and Forde, G. M. (2009). Dewatering of microalgal culture for biodiesel production: Exploring polymer flocculation and tangential flow filtration. Journal of Chemical Technology and Biotechnology, 84: 1078-1083.
Del Campo, J. A., Garcia-Gonzalez, M., and Guerrero, M. G. (2007). Outdoor cultivation of microalgae for carotenoid production: Current state and perspectives. Applied Microbiology and Biotechnology, 74: 1163-1174.
Donovan, J., and Stowe, N. (2009). Is the Future of Biofuels in Algae? [Online] Renewable Energy World, 12 06 2009. http://www. renewableenergyworld. com/rea/news/article/2009/06/ is-the-futu.
Eroglu, E., and Melis, A. (2010). Extracellular terpenoid hydrocarbon extraction and quantitation from the green microalgae Botryococcus braunii var. Showa. Bioresource Technology, 101: 2359-2366.
Esperanza, D. R.F., Gabriel, A. M., Carmen, G. M., Joaquin, R., Emilio, M. G., and Miguel, G. G. (2005). Efficient one-step production of astaxanthin by the microalga Haematococcus pluvialis in continuous culture. Biotechnology and Bioengineering, 91: 808-815.
Fairley, P. (2011). Introduction: Next generation biofuels. Nature,474: S2-S5. doi:10.1038/474S02a
Fernandez-Reiriz, M. J., Perez-Camacho, A., Ferreiro, M. J., Blanco, J., Planas, M., Campos, J., and Labarta, U. (1989). Biomass production and variation in the biochemical profile (total protein, carbohydrates, RNA, lipids and fatty acids) of seven species of marine microalgae. Aquaculture, 83: 17-37.
Flynn, K. J., Greenwell, H. C., Lovitt, R. W., and Shields, R. J. (2010). Selection for fitness at the individual or population levels: Modelling effects of genetic modifications in microalgae on productivity and environmental safety. Journal of Theoretical Biology, 263: 269-280.
Garcia-Malea, M. C., Brindley, C., Del Rio, E., Acien, F. G., Fernandez, J. M., and Molina, E. (2005). Modeling of growth and accumulation of carotenoids in Haematococcus pluvialis as a function of irradiance and nutrients supply. Biochemical Engineering Journal, 26: 107-114.
Garcia, M. L., Del Rio Sanchez, E., Casas Lopez, J. L., Acien, F. G., Fernandez, J. M., Fernandez Sevilla, J. M., Rivas, J., Guerrero, M. G., and Molina Grima, E. (2006). Comparative analysis of the outdoor culture of Haematococcus pluvialis in tubular and bubble column photobioreactors. Journal of Biotechnology, 29: 329-342.
Gladue, R. M., and Maxey, J. E. (1994). Microalgal feeds for aquaculture. Journal of Applied Phycology, 6: 131-141.
Gomez, P. I., Barriga, A., Silvia Cifuentes, A., and Gonzalez, M. A. (2003). Effect of salinity on the quantity and quality of carotenoids accumulated by Dunaliella salina (strain CONC-007) and Dunaliella bardawil (strain ATCC 30861) Chlorophyta. Biological Sciences, 36: 185-192.
Gouveia, L., and Olieveira, A. C. (2009). Microalgae as a raw material for biofuel production. Journal of Industrial Microbiology and Biotechnology, 36: 269-274.
Gouveia, L., Marques, A. E., Da Silva, T. L., and Reis, A. (2009). Neochloris oleoabundans UTEX#1185: A suitable renewable lipid source for biofuel production. Journal of Industrial Microbiology and Biotechnology, 36: 821-826.
Gressel, J. (2008). Transgenics are imperative for biofuel crops. Plant Science, 174: 246-263.
Grima, E. M., Belarbia, E. H., Acieen Fernandeza, F. G., Robles Medina, A., and Chistib, Y. (2003). Recovery of microalgal biomass and metabolites: Process options and economics. Biotechnology Advances, 20: 491-515.
Grobbelaar, J. U. (2009). Factors governing algal growth in photobioreactors: The “open” versus “closed” debate. Journal of Applied Phycology, 21: 489-492.
Grobbelaar, J. U. (2010). Microalgal biomass production: challenges and realities. Photosynthesis Research, 106: 135-144.
Haag, A. L. (2007). Algae bloom again. Nature, 447: 520-521.
Harel, Y., Ohad, I., and Kaplan, A. (2004). Activation of photosynthesis and resistance to photoinhibition in cyanobacteria with biological desert crust. Plant Physiology, 136: 3070-3079.
Harun, R., Singh, M., Forde, G. M., and Danquah, M. K. (2010). Bioprocess engineering of microalgae to produce a variety of consumer products. Renewable and Sustained Energy Reviews, 14: 1037-1047.
Hejazi, M. A., Holwerda, E., and Wijffels, R. H. (2004). Milking microalga Dunaliella salina for P-carotene production in two-phase bioreactors. Biotechnology and Bioengineering, 85: 475-481.
Heredia-Arroyo, T., Wei, W., and Hu, B. (2010). Oil accumulation via heterotrophic/mixo — trophic Chlorella protothecoides. Applied Biochemistry and Biotechnology, 162: 1978-1995. http://techtransfer. universityofcalifornia. edu/NCD/21141.html. A Hybrid Pond-Bioreactor for Mass Algal Culture. Tech ID: 21141/UC Case 2010-280-0.
Huerlimann, R., Nys, R., and Heimann, K. (2010). Growth, lipid content, productivity, and fatty acid composition of tropical microalgae for scale-up production. Biotechnology and Bioengineering, 107: 245-257.
Huesmann, M. H. (2000). Can advances in science and technology prevent global warming? A critical review of limitations and challenges. Mitigation and Adaptation Strategies for Global Change, 11: 539-577.
Hunt, R. W., Chinnaswamy, S., Bhatnager, A., and Das, K. C. (2010). Effect of biochemical stimulants on biomass productivity and metabolite content of microalgae, Chlorella sorokiniana. Applied Biochemistry and Biotechnology, doi: 10.1007/s2010-010-9012-2.
Huntley, M. E., and Redalje, D. G. (2007). CO2 mitigation and renewable oil from photosynthetic microbes: A new appraisal. Mitigation and Adaptation Strategies for Global Climate Change, 12: 573-608.
Janssen, M., Tramper, J., Mur, L. R., and Wijffels, R. H. (2003). Enclosed outdoor photobioreactors: Light regime, photosynthetic efficiency, scale-up, and future prospects. Biotechnology and Bioengineering, 81: 193-210.
Jin, E., Feth, B., and Melis, A. (2003). A mutant of the green alga Dunaliella salina constitutively accumulates zeaxanthin under all growth conditions. Biotechnology Bioengineering, 81: 116-124.
Johnson, M. B. (2009). Microalgal Biodiesel Production through a Novel Attached Culture System and Conversion Parameters. M. Sc. thesis in Biological Systems Engineering, Blacksburg, VA, p. 83.
Kaeriyama, H., Katsuki, E., Otsubo, M., Yamada, M., Ichimi, K., Tada, K., and Harrison, P. J. (2011). Effects of temperature and irradiance on growth of strains belonging to seven Skeletonema species isolated from Dokai Bay, southern Japan. European Journal of Phycology, 46: 113-124.
Kanellos, M. (2009). Algae Biodiesel: It’s $33 a Gallon. Greentech Media. [Online] Greentech Media, 03 02 2009. [Cited: 10 12 2009.] http://www. greentechmedia. com/articles/ algae-biodiesel-its-33-a-gallon-5.
Kebede-Westhead, E., Pizarro, C., and Mulbry, W. W. (2006). Treatment of swine manure effluent using freshwater algae; Production, nutrient recovery, and elemental composition of algal biomass at four effluent loading rates. Journal of Applied Phycology, 18: 41-46.
Keerthi, S., Uma Devi, K., Subba Rao, D. V., Sarma, N. S. (in press). Exogenous vitamin dependency in two carotenogenic Dunaliella strains isolated from coastal Bay of Bengal.
Khan, S. A., Hussain, M. Z., Prasad, S., and Banerjee, U. C. (2009). Prospects of biodiesel production from microalgae in India. New Delhi. India 2009. Renewable and Sustainable Energy Reviews, 13: 2361-2372.
Khozin-Goldberg, I., and Cohen, Z. (2006). The effect of phosphate starvation on the lipid and fatty acid composition of the freshwater eustigmatophyte Monodus subterraneus. Phytochemistry, 67: 696-701.
Kleinegris, D. M.M., Janssen, M., Brandenburg, W. A., and Wiiffels, R. H. (2011). Two-phase systems: Potential for in situ extraction of microalgal products. Biotechnology Advances, 29: 502-507.
Kong, Q. X., Li, L., Martinez, B., Chen, P., and Ruan, R. (2010). Culture of microalgae Chlamydomonas reinhardtii in wastewater for biomass feedstock production. Applied Biochemistry and Biotechnology, 160: 9-18.
Lebeau, T., and Robert, R. M. (2006). Biotechnology of immobilized microalgae. A culture technique for future? In Algal Cultures, Analogues of Blooms and Applications (Ed. Subba Rao, D. V.), Science Publishers, Enfield, NH, pp. 801-839.
Lee, Y. K., Ding, S. Y., Hoe, C. H., and Low, C. S. (1996). Mixotrophic growth of Chlorella soro — kiniana in outdoor enclosed photobioreactor. Journal of Applied Phycology, 8: 163-169.
Leon, R., and Fernandez, E. (2007). Nuclear transformation of eukaryotic microalgae. In Transgenic Microalgae as Green Cell Factories (Eds. Leon, R., Galvan, A., and Fernandez, E.). Landes Bioscience and Springer Science+ Business Media, New York,
pp. 1-11.
Li, S., and Tsai, H. (2008). Transgenic microalgae as a non-antibiotic bactericide producer to defend against bacterial pathogen infection in the fish digestive tract. Fish & Shellfish Immunology, 26: 316-325.
Li, Y., Horsman, M., Wang, B., Wu, N., and Lan, C. Q. (2008). Effects of nitrogen sources on cell growth and lipid accumulation of green alga Neochloris oleoabundans. Applied Microbiology and Biotechnology, 81: 629-636.
Li, H., Xu, H., and Wu, Q. (2007). Large-scale biodiesel production from microalga Chlorella protothecoides through heterotrophic cultivation in bioreactors. Biotechnology and Bioengineering, 98: 764-771.
Liu, Z. Y., Wang, G. C., and Zhou, B. C. (2008). Effect of iron on growth and lipid accumulation in Chlorella vulgaris. Bioresource Technology, 99: 4717-4722.
Major, K. M., and Henley, W. (2008). Influence of salinity and temperature on growth and photosynthesis in the extremophilic chlorophyte, Nannochloris sp. Journal of Phycology, 37: 32.
Malcata, F. X. (2010). Microalgae and biofuels: A promising partnership? Trends in Biotechnology, 29: 542-559.
Mandal, S., and Mallick, N. (2009). Microalga Scenedesmus obliquus as a potential source for biodiesel production. Applied Microbiology and Biotechnology, 84: 281-291.
Mata, T. M., Martins, A. A., and Caetanao, N. S. (2010). Microalgae for biodiesel production and other applications: A review. Renewable and Sustainable Energy Reviews, 14: 217-232.
Matsumoto, M., Sugiyama, H., Maeda, Y., Sato, R., Tanaka, T., and Matsunaga, T. (2009). Marine diatom, Navicula sp. strain JPCC DA0580 and marine green alga, Chlorella sp. strain NKG400014 as potential sources for biodiesel production. Applied Biochemistry and Biotechnology, 16: 483-490.
McGinnis, K. M., Dempster, T. A., and Sommerfeld, M. R. (1997). Characterization of the growth and lipid content of the diatom Chaetoceros muelleri. Journal of Applied Phycology, 9: 19-24.
Miquel, P. (1893). De la culture artificielle des diatoms. Introduction. Le Diatomiste, 1: 73-75.
Misumi, O., Yoshida, Y., Nishida, K., Fujiwara, T., and Sekajin, T. (2008). Genome analysis and its significance in four unicellular algae, Cyanidioshyzon merolae, Ostreococcus tauri, Chlamydomonas reinhardtii, and Thalassiosira pseudonana. Journal of Plant Research, 121: 3-17.
Mitra, M., and Melis, A. (2008). Optical properties of microalgae for enhanced biofuels production. Optics Express, 16: 21807-21820.
Morgan-Kiss, R. M., Ivanov, A. G., Modia, S., Czymmek, K., and Huner, N. P. (2008). Identity and physiology of a new psychrophilic eukaryotic green alga, Chlorella sp. strain BI, isolated from a transitory pond near Bratina Island, Antarctica. Extremophiles, 12: 701-711.
Muradyan, E. A., Klyachko-Gurvich, G. L., Tsoglin, L. N., Sergeyenko, T. V., and Pronina, N. A. (2004). Changes in lipid metabolism during adaptation of the Dunaliella salina photosynthetic apparatus to high CO2 concentration. Russian Journal of Plant Physiology, 51: 53-62.
Natural Resources Defense Council (2009). The Promise of Algae Biofuels. Catie Ryan, Terrapin Bright Green, LLC, New York, p. 81.
Norsker, N., Barbosa, M. J., Vermue, M. H., and Wijffels, R. H. (2011). Microalgal production — A close look at the economics. Biotechnology Advances, 29: 24-27.
Olaizola, M. (2000). Commercial production of astaxanthin from Haematococcus pluvialis using 25,000-liter outdoor photobioreactors. Journal of Applied Phycology, 12: 499-506.
Orpez, R., Martinez, M. E., Hodaifa, G., Yousfi, F. El., Sanchez, S., and Jbari, N. (2009). Growth of the microalga Botryococcus braunii in secondarily treated sewage. Desalination, 246: 625-630.
Park, J. B.K., Craggs, R. J., and Shilton, A. N. (2011). Wastewater treatment high rate algal ponds for biofuel production. Bioresource Technology, 102: 71-81.
Perez-Garcia, O., De-Basham, L., Hernandez, J., and Bashan, Y. (2010). Efficiency of growth and nutrient uptake from wastewater by heterotrophic, autotrophic, and mixo- trophic cultivation of Chlorella vulgaris with Azospirillum brasiliense. Phycology, 46: 800-812.
Pienkos, P. T., and Darzins, A. (2009). The promise and challenges of microalgal-derived biofuels. Biofuels, Bioproducts and Biorefining, 3: 431-440.
Pienkos, P. T., Laurens, L., and Aden, A. (2011). Making biofuel from microalgae. American Scientist, 99: 474-481.
Pitman, J. K., Dean, A. P., and Osundeko, O. (2011). The potential of sustainable algal biofuel production using wastewater resources. Bioresource Technology, 102: 17-25.
Pocock, T., Vetteril, A., and Falk, S. (2011). Evidence of phenotypic plasticity in the Antarctic extremophile Chlamydomonas raudenis Ettl. UWO 241. Journal of Experimental Botany, 62: 1169-1177.
Pruvost, J., Van Vooren, G., and Legrand, C. (2009). Investigation of biomass and lipids production with Neochloris oleoabundans in photobioreactor. Bioresource Technology, 100: 5988-5995.
Pulz, O. (2007). Performance Summary Report: Evaluation of Green Fuel’s 3D Matrix Algae Growth Engineering Scale Unit, APS Red Hawk Power Plant, AZ. IGV Instut fur Getreideverabeitung GmbH, pp. 1-14.
Pulz, O., and Gross, W. (2004). Valuable products from biotechnology of microalgae. Applied Microbiology and Biotechnology, 65: 635-648.
Radakovits, R., Jinkerrson, R. E., Darzins, A., and Posewitz, M. C. (2010). Genetic engineering of algae for enhanced biofuel production. Eukaryotic Cell, 9: 486-501.
Raven, J. A. (2009). Carbon dioxide fixation by Dunaliella spp. and the possible use of this genus in carbon dioxide mitigation and wastewater reduction. In The Alga Dunaliella Biodiversity, Physiology, Genomics and Biotechnology (Eds. Ben-Amotz, A., Polle, J. E.W., and Subba Rao, D. V.), Science Publishers, Enfield, NH, pp. 359-384.
Riegman, R., Boer, M., and Senerpont Domis, L. (1996). Growth of harmful marine algae in multispecies cultures. Journal of Plankton Research, 18: 1851-1866.
Rivasseau, C., Farhi, E., Gromova, M., Oliver, J., and Bligny, R. (2010). Resistance to irradiation of micro-algae growing in the storage pools of a nuclear reactor investigated by NMR and neutron spectroscopies. Spectroscopy: An International Journal, 24: 381-385.
Rodolfi, L., Zitteli, G. C., Bassi, N., Padovani, G., Biondi, N., Bonini, G., and Tredici, M. R. (2008). Microalgae for oil: Strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnology and Bioengineering, 102: 100-112.
Ronquillo, J. D., Matias, J. R., Saisho, T., and Yamasaki, S. (1997). Culture of Tetraselmis tetrathele and its utilization in the hatchery production of different penaeid shrimps in Asia. Hydrobiologia, 358: 237-244. doi: 10.1023/A:1003128701968.
Rosenberg, J. N., Oyler, G. A., Wilkinson, L., and Betenbaugh, M. J. (2008). A green light for engineered algae: redirecting metabolism to fuel a biotechnology revolution. Biotechnology, 19: 430-436.
Russell, G., and Fielding, A. H. (1974). The competition properties of marine algae. Journal of Ecology, 62: 689-698.
Scott, S. A., Davey, M. P., Dennis, J. S., Horst, I., Howe, C. J., and Smith, A. G. (2010). Biodiesel from algae: challenges and prospects. Current Opinion Biotechnology, 21: 277-286.
Serrano, L. (2010). Five hard truths for synthetic biology. Nature, 463: 288-290.
Service, R. F. (2008). Eyeing oil, synthetic biologists mine microbes for black gold. Science, 322: 522-523.
Spijkerman, E. (2010). High photo synthetic rates under a co-limitation for inorganic phosphorus and carbon dioxide. Journal Phycology, 46: 658-664.
Spijkeman, E., and Wacker, A. (2011). Interactions between P-limitation and different C conditions on the fatty acid composition of an extremophile microalga. Extremophiles, 15: 597-609.
St. John, J. (2009). Algae Company Number 56: Plankton Power. Greentech Media. [Online] Greentech Media Inc, 04 08 2009. [Cited: October 12 2009.] http://www. greentechmedia. com/articles/read/plankton-power-another-algae.
Stephens, E., Ross, I. L., Mussgnug, J. H., Wagner, L. D., Borowitzka, M. A., Posten, C., Kruser, O., and Hankamer, B. (2010). Future prospects of microalgal biofuel production systems. Trends in Plant Science, 15: 554-564.
Sturm, B. S.M., and Lamer, S. L. (2011). An energy evaluation of coupling nutrient removal from wastewater with algal biomass production. Applied Energy, 88: 3499-3506.
Subba Rao, D. V. (2009). Cultivation, growth media, division rates and applications of Dunaliella species. In The Alga Dunaliella Biodiversity, Physiology, Genomics and Biotechnology (Eds. Ben-Amotz, A., Polle, J. E.W., and Subba Rao, D. V.). Science Publishers, Enfield, NH, pp. 45-90.
Subba Rao, D. V., Pan Y., and Al-Yamani, F. (2005). Growth and photosynthetic rates of Chalmydomonas plethora and Nitzschia frustula cultures isolated from Kuwait Bay, Arabian Gulf and their potential as live algal food for tropical mariculture. Marine Ecology, 26: 63-71.
Takagi, M., Karseno, S., and Yoshida, Y. (2006). Effect of salt concentration on intracellular accumulation of lipids and triglyceride in marine microalgae Dunaliella cells. Journal of Bioscience and Bioengineering, 101: 223-226.
Theegala, C. S., Robertson, C., and Suleiman, A. A. (2001). Phytoremediation potential and toxicity of barium to three freshwater microalgae: Scenedesmus subspicatus, Selenastrum capricornutum, and Nannochloropsis sp. Practice Periodical of Hazardous, Toxic, and Radioactive Waste Management, 5: 194-202.
Tsoglin, L. N., and Gabel, B. V. (2000). Potential productivity of microalgae in industrial photobioreactor. Russian Journal of Plant Physiology, 47: 668-673.
Ugwu, C. U., Aoyagi, H., and Uchiyama, H. (2008). Photobioreactors for mass cultivation of algae. Bioresource Technology, 99: 4021-4028.
Wahal, S., and Viamajala, S. (2010). Maximizing algal growth in batch reactors using sequential change in light intensity. Applied Biochemistry and Biotechnology, 161: 511-522.
Weber, A. P.M., Oesterhelt, C., Gross, W., Brautigam, A., Imboden, L. A., Krassovskayal, I., Linka, N., Truchina, J., Zimmermann, M., Jamai, A., Riekhof, W. R., Yu, B., Garavito, R. M., and Benning, C. (2004). EST—Analysis of the thermo-acidophilic red microalga Galderia sulphuraria reveals potential for lipid A biosynthesis and unveils the pathway of carbon export from rhodoplasts. Plant molecular Biology, 55: 17-32.
Wang, L., Li, Y., Chen, P., Min, M., Chen, Y., Zhu, J., and Ruan, R. R. (2010). Anaerobic digested dairy manure as a nutrient supplement for cultivation of oil-rich green microalgae Chlorella sp. Bioresource Technology, 101: 2623-2688.
Weyer, K. M., Bush, D. R., Darzins, A., and Willson, D. B. (2010). Theoretical maximum algal oil production. Bioenergy Research, 3: 204-213.
Wijffels, R. H. (2007). Potential of sponges and microalgae for marine biotechnology. Trends in Biotechnology, 26(1): 26-31.
Williams, PJ. le B., and Laurens L. M.L. (2010). Microalgae as biodiesel and biomass feedstocks. Energy and Environmental Science, 3: 554-590.
Woertz, I., Feffer, A., Lundquist, T., and Nelson, Y. (2009). Algae grown on dairy and municipal wastewater for simultaneous nutrient removal and lipid production for biofuel feedstock, Journal of Environmental Engineering, 135: 1115-1122.
Wu, X., and Merchuk, J. C. (2004). Simulation of algae growth in a bench scale internal loop airlift reactor. Chemical Engineering Science, 59: 2899-2912.
Xu, H., Miao, X. L., and Wu, Q. Y. (2006). High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters. Journal of Biotechnology, 126: 499-507.
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FIGURE 2.4 Morphological diversity of microalgae: (a) Ceratium hirundinella (Muller) Dujardin, (b) Ceratium longipeps (Bailey) Grun., (c) Ceratium trichoceros (Ehrenberg) Kofoid, (d) Gymnodinium sanguineum Hirasaka.
■ P fertilizer
■ Electrical power
■ Perspex tubing
■ N fertilizer
■ P fertilizer
■ Electrical power
■ Concrete walls
■ Steel paddle wheel
■ PVC lining
■ Water supply & treatment
FIGURE 9.2 The relative contribution of fossil energy (left) and GWP (right) to the total requirements for microalgal biodiesel production using a tubular airlift reactor (upper) and a raceway (lower). From the LCA of C. vulgaris conducted by Stephenson et al. (2010) under standard conditions. The total fossil energy requirements of 230 and 29 GJ and GWP of 13,550 and 1,900 kg CO2 per tonne biodiesel formed were estimated for the tubular reactor and raceway, respectively.
Applications and Market Segments
• Human Nutrition
Functional Foods Nutraceuticals
• Cosmeceuticals
• Animal Feed
• Pigments
о Food and Beverage Industry
о Pharmaceuticals Aquaculture Cosmetics
• Clinical and diagnostic research reagents
• Bioremediation
FIGURE 10.1 Applications of algal biomass.
CLEAN TECHNOLOGY
The most commonly used design for commercial microalgal production is the raceway pond. A raceway is an oval-shaped, single — or multiple-loop recirculation channel (Figure 5.1), usually 15 to 20 cm deep, with mixing provided through circulation by a rotating paddlewheel (Pulz, 2001; Brennan and Owende, 2010). Baffles are often placed in the bends of the flow channel to guide the water and facilitate mixing (Chisti, 2007). They are commonly built from concrete or packed earth,
and covered with a plastic lining. The largest raceway pond in operation is 5,000 m2, located at Earthrise Nutritionals, a commercial Spirulina producer in Southern California (Spolaore et al., 2006).
By convention, in continuous production, nutrients are introduced in front of the paddlewheel and harvesting takes place behind the wheel (Brennan and Owende, 2010). CO2 is provided by gas exchange via natural contact with the surrounding air (Singh et al., 2011). Occasionally, submerged aerators are installed to enhance CO2 absorption. Light provision is by natural sunlight. Ponds can be placed inside covered tunnels to aid in temperature regulation.
Raceway ponds incur relatively low capital investment as well as operational costs. Weekly monitoring is usually sufficient, and the main costs are in the media components and the energy consumed for mixing (Singh et al., 2011). Biomass concentrations are normally in the region of 0.5 g L-1, with a biomass productivity of 10 to 25 g m-2d-1 (Sheehan et al., 1998; Lee, 2001). Raceway ponds have been in use since the first commercial microalgal ventures were established, and extensive experience in their design and operation exists. Examples of the productivities obtained with various microalgal species in raceway ponds are given in Table 5.2.
The only open system to achieve very high cell densities sustainably is the cascade system developed in the Czech Republic and used for cultivation of Chlorella (Setlik et al., 1970). With a culture depth of less than 1 cm, cell densities of up to
TABLE 5.2
Examples of Productivities Achieved in Open Ponds with Various Microalgal Species
Highest Productivity
10 g L-1 Chlorella were achieved. However, the biomass productivity was comparable to that of raceways (25 g m-2d-1) (Lee, 2001). The system had a sloping base made of glass, which rendered it very expensive, but the use of cheaper materials could make it price competitive with raceway ponds. A similar system has been used in Western Australia, consisting of a 0.5-ha sloping, plastic-lined pond for the production of Chlorella, achieving similar biomass productivity (Borowitzka, 1999).
CO2 provision to raceway ponds through additional sparging of compressed CO2 contributed some 40% of the energy consumption and 30% GHG emissions (Clarens et al., 2010). Alternatively, co-location with industries or facilities with high CO2 emissions (such as power stations, fermentation plants, or anaerobic digester systems) may facilitate reduced contributions for effective CO2 provision. This has been demonstrated; however, it is noted that the CO2 source may influence the productivity achieved and interacts with the supply of other nutrients. Stephenson et al. (2010) modeled the impact of CO2 concentration in the gas to be compressed for sparging into either the raceway or tubular reactor system. Owing to the influence of CO2 concentration on both concentration driving force (and hence transfer rate) and on the volume of gas to be compressed, its impact is significant, with the fossil energy requirement nearly doubling on decreasing the CO2 concentration from 12.5% (typical of flue gases) to 9% and increasing fourfold on decrease to 5% by volume in the raceway system. The design of a low-depth carbonation sump also favors reduced energy consumption.
It is increasingly recognized that the provision of nutrients, especially combined nitrogen, to bioprocesses affects their life-cycle impact (Harding 2009; Harding et al., 2012). Lardon et al. (2009) illustrated that the provision of fertilizer accounted for 15% to 25% of the energy requirements per unit biodiesel. This was substantially reduced under nitrogen-limited conditions (6% to 9%). Clarens et al. (2010) illustrated that 50% of the energy requirement and GWP for biomass production is attributable to nutrient provision in their raceway system. The potential exists to replace these with wastewaters such as effluent from the conventional activated sludge process or source-separate urine. The former has the potential to provide a water supply simultaneously. Similar benefits can be achieved by maximizing the nutrient recycle (Stephenson et al., 2010; Richardson, 2011) and minimizing the nitrogen input required, either by optimizing the nitrogen limitation or selecting an algal species of low nitrogen content, for example, Phaeodactylum tricornutm at 0.8% N over algal species with a typical nitrogen content of 6% by mass (Richardson et al., 2012b).
Ismail Rawat, Ramanathan Ranjith Kumar, and Faizal Bux
Institute for Water and Wastewater Technology Durban University of Technology Durban, South Africa
12.1 Introduction…………………………………………………………………………………………….. 179
12.2 Wastewater Characteristics…………………………………………………………………….. 181
12.2.1 Physical Characteristics………………………………………………………………. 182
12.2.2 Chemical Characteristics……………………………………………………………. 184
12.2.3 Biological Characteristics…………………………………………………………… 184
12.3 Phycoremediation…………………………………………………………………………………… 185
12.4 Algae Species Used for Phycoremediation……………………………………………… 186
12.5 High-Rate Algal Ponds (HRAPs)…………………………………………………………….. 187
12.5.1 Nutrient Removal……………………………………………………………………….. 188
12.5.2 Factors Affecting High-Rate Algae Ponds………………………………….. 190
12.5.3 Efficiency of Wastewater Treatment and Algal Growth……………. 191
12.6 Wastewater as Feedstock for Biomass Production…………………………………. 192
12.7 Economics and Energy Balance of Phycoremediation Using HRAPs……. 194
12.8 Conclusion………………………………………………………………………………………………. 195
Acknowledgments……………………………………………………………………………………………… 195
References………………………………………………………………………………………………………… 196
The disposal of liquid and solid waste in rivers, streams, lakes, and oceans has been occurring for extended periods of time. Increasing industrialization to serve rapidly expanding urban population needs generates large amounts of wastewater that require treatment before release in order to prevent further environmental deterioration. Point source wastewater contamination has the capacity to “overload” receiving water bodies and is the most widespread threat to environmental water quality. Wastewater generally contains high concentrations of organic and inorganic nutrients, which are among the main causes of irreversible ecological degradation. This disrupts the bio-system and natural recycling processes such as photosynthesis, respiration, nitrogen fixation, evaporation, and precipitation. Effective wastewater treatment and the use of reclaimed wastewater have great potential to help meet fresh water requirements for various domestic and industrial uses, thus somewhat alleviating the need for water in growing urban centers. In industrial and municipal wastewater, reduction of various chemical stacks at sources is not an easy process and is very expensive to treat by conventional treatment methods due to the demand for skilled operators, high capital investment, high operational costs, reliability etc. Complex operation of conventional treatment methods for removing chemicals does not guarantee sludge reduction. Sludge removal is one of the main challenges in sustainable wastewater treatment, but can be accomplished by the Best Available Technique (BAT) to treat the socio-economic aspect of efficient wastewater treatment. This, coupled with potential energy resource recovery, is mandatory and necessary in exploring the feasibility of biological treatment. There has been growing worldwide interest due to decreasing water resources and increasing demand for preservation and the sustainable management of water resources (Garca et al., 2000).
Microalgal cultivation is an attractive biotechnological wastewater treatment method that has potential as an alternative method to conventional treatment. Microalgae are popular bio-resources, as appropriate microalgal technology can add a number of benefits to the treatment process because they have a greater capacity for the treatment of a number of wastewater contaminants. Chinnasamy et al. (2010) observed that a consortium of fifteen native microalgae efficiently reduced more than 96% of carpet mill treated wastewater nutrients within 72 h. Wang et al. (2010) reported rapid decreases in nitrate, phosphate, and metal levels in wastewater treatment over a short period of microalgal cultivation. Microalgal wastewater treatment is an economically viable method of wastewater treatment that has an extensive research history spanning more than 50 years (Oswald et al., 1953; Oswald, 1991; Ruiz et al., 2011). Microalgae-based wastewater removal of nutrients and/or chemicals is achieved by accumulation in, or conversion to, biomass, making it a better biotechnological method for the preservation of freshwater ecosystems (Hoffmann, 1998; Ruiz et al., 2011). Considering that inexpensive effluent can be used as feed for desired microalgal species to produce algae-derived products, while simultaneously removing nutrients, makes it an attractive biological system. Thus, phycoremediation technology is a promising field for applied studies such as in wastewater treatment, and biomass and biofuels production for sustainable energy.
The cultivation of microalgae for wastewater treatment is a high-quality, ecofriendly process with no secondary pollution. Reclaimed effluent produces high-value microalgal metabolites such as lipids, carbohydrates, and proteins. Microalgae are often applied in the tertiary treatment of domestic wastewater in maturation ponds, or in small — to medium-scale municipal wastewater treatment systems (Hanumantha Rao et al., 2011; Rawat et al., 2011). Technologies such as the advanced integrated wastewater pond systems (AIWPS) are commercially available (Oswald, 1991). The most common designs include facultative ponds, which are relatively deep and support surface growth of microalgae. High-rate algal ponds (HRAPs) are a hallmark technology to treat a number of wastewater streams, especially under tropical and subtropical conditions due to the availability of sunlight utilized by microalgae for photosynthesis (Phang et al., 2000; Mustafa et al., 2011). Shallow ponds depend on mechanical mixing for maximum algae production and removal of biological oxygen demand. HRAPs are the most
cost-effective reactors for liquid waste management and capture of solar energy, and the capture of atmospheric carbon dioxide, and are used in the treatment of animal wastes (Narkthon, 1996). HRAP wastewater treatment can be highly efficient in reducing bacteria, biological oxygen demand (BOD), and nutrient levels by integrated approaches to recycling wastes. Phycoremediation can be used in the process as a second step after initial anaerobic treatment of high organic wastewater to yield a significant reduction in influent organic matter, such as nitrogen and phosphorus. Harvested microalgae are rich in nutrients such as nitrogen, potassium, and phosphorus, which can be used for animal feed, etc. (Ogbonna et al., 2000; Olguin, 2003; Rawat et al., 2011). Therefore, HRAPs are very appropriate for sanitation in small rural communities because of their simplicity of operation in comparison to conventional technologies such as the activated sludge process. This chapter critically evaluates phycoremediation HRAPs for removal of high organic strength nutrients by means of enriched microalgae.
Floatation is a separation process based on the attachment of air bubbles to solid particles. The resulting flocs float to the liquid surface and are harvested by skimming and filtration. The success of flotation depends on the nature of suspended particles (microalgal cells in harvesting process). Air bubbles drift up the smaller particles (<500 pm) more easily (Matis et al., 1993). Also, the lower instability of suspended particles results in relatively higher air-particle contact. The attachment of air bubbles also depends on the air, solid, and aqueous phase contact angle. The larger the contact angle, the greater the tendency of air to adhere (Shelef et al., 1984). Dissolved air flotation (DAF), electrolytic flotation, and dispersed air flotation are the commonly used flotation techniques according to the method of bubble production. Dissolved air flotation is the most widely used method for the treatment of industrial effluent. Van Vuuren et al. (1965) performed a study on flotation and reported that flocculation requires several hours of sedimentation, while flotation shortens the duration to only a few minutes. The DAF procedure by chemical flocculation is reported to recover up to 6% (w/v) algal biomass slurries from algae culture (Bare et al., 1975). Although flotation has been used by several researchers as a potential harvesting method, there is only limited evidence of its technical and economic viability.
Carotenoids are colored, lipid-soluble compounds that occur in higher plants, microalgae, as well as in nonphotosynthetic organisms (Del Campo, 2007; Takaichi, 2011). Carotenoids contribute to light harvesting, maintenance of structure, and functioning
Applications and Market Segments
• Human Nutrition Functional Foods Nutraceuticals
• Cosmeceuticals
• Animal Feed
• Pigments
о Food and Beverage Industry
о Pharmaceuticals Aquaculture Cosmetics
• Clinical and diagnostic research reagents
• Bioremediation FIGURE 10.1 (See color insert.) Applications of algal biomass.
TABLE 10.1 Global Production of Algal Biomass for Commercially Relevant Algal Genera
Source: Adapted from Pulz and Gross (2004), Spolaore et al. (2006); Milledge (2011). |
of photosynthetic complexes in plants and microalgae (Pulz and Gross, 2004; Del Campo, 2007). They occur widely in nature and are responsible for many of the brilliant red, orange, and yellow colors of edible vegetables and fruits and some aquaculture animals.
Microalgae combine properties of higher plants with some properties of prokaryotes. This combination represents the rationale for using microalgae for the production of carotenoids and other products (Del Campo, 2007; Guedes et al., 2011) instead of using plants or prokaryotes. Furthermore, the production of carotenoids by microalgae can be easily maximized by manipulating growth conditions. Under unfavorable growth conditions, microalgae produce high amounts of carotenoids, such as P-carotene, astaxanthin, and canthaxanthin (Orosa et al., 2000).
Microalgal wastewater treatment using microalgae with the production of biomass as a by-product is not a new concept. However, it occurs only on a minor scale in waste stabilization ponds and HRAPs. Wastewater treatment using HRAPs has the potential to produce large amounts of biomass that can be used for a variety of applications, including the production of renewable fuels, fertilizer, animal feed, etc. (Rawat et al., 2011). Recent studies have suggested that the use of wastewater as a substrate for biofuel production may make the process economically viable (Brennan and Owende, 2010; Boelee et al., 2011; Cho et al., 2011). Focusing the growth of microalgae on biomass productivity rather than lipid productivity may be beneficial as larger amounts of biomass improve the viability of conversion to alternate fuels (Pittman et al., 2011). Microalgal biomass to biofuels conversion may be carried out by several methods depending on the biomass characteristics (e. g., lipid or carbohydrate content) (Garcia et al., 2006; Rawat et al., 2011). The yields of biomass from HRAPs depend on the type of effluent being treated with specific regard to nutrient content. Table 12.2 summarizes growth and lipid productivity of microalgal species on a variety of wastewater types. Piggery waste effluent treatment by HRAPs has potential productivities of up to 50 t ha-1yr-1 (Rawat et al., 2011).
Maximum algal productivities in HRAPs can be achieved by countering rate — limiting and inhibitory conditions. Carbon is often a rate-limiting substrate and may be alleviated by the addition of CO2. This addition serves a dual role in the provision of carbon and a method of pH control. The addition of CO2 has been shown to double algal productivity at the laboratory scale and increase productivity by 30% in a pilot-scale HRAP (Park et al., 2011a). Biomass grown at the Lawrence wastewater treatment plant showed algal productivities ranging from 5 to 16 g m-2d-1 and average lipid contents of 10% without the addition of CO2. With the addition of CO2, productivities were expected to be 25 g m-2d-1 (Sturm and Lamer, 2011). However, it must be considered that addition of excess CO2 leads to a decrease in pH. A pH maintained at a maximum of 8 inhibits physico-chemical processes of nutrient removal such as volatilization of ammonia and phosphate precipitation (Craggs, 2005). But this is not necessarily a negative point, as the increase in assimilation by biomass production offsets the losses on physico-chemical removal. Furthermore, it enables the recycling of nutrients that would have been otherwise lost. Ammonia volatilization accounts for approximately 24% nitrogen loss in HRAPs without pH control (Park et al., 2011a).
TABLE 12.2
Biomass and Lipid Productivities of Microalgae Grown on Various Wastewater Streams
Biomass (DW) Productivity |
Lipid Content |
Lipid Productivity |
||
Wastewater Type |
Microalgal Species |
(mg L-1d-1) |
(%DW) |
(mg L-1d-1) |
Municipal (primary treated) |
nd |
25a |
nd |
nd |
Municipal (centrate) |
Chlamydomonas reinhardtii (biocoil grown) |
2000 |
25.25 |
505 |
Municipal (secondary treated) |
Scenedesmus obliquus |
26b |
31.4i |
8і |
Municipal (secondary treated) |
Botryococcus braunii |
345.6c |
17.85 |
62 |
Municipal (primary treated + CO2) |
Mix of Chlorella sp., Micractinium sp., Actinastrum sp. |
270.7d |
9 |
24.4 |
Agricultural (piggery manure with high NO3-N) |
Botryococcus braunii |
700e |
nd |
69 |
Agricultural (dairy manure with polystyrene foam support) |
Chlorella sp. |
2.6 g m-2d-1 |
9і |
230і mg m-2d-1 |
Agricultural (fermented swine urine) |
Scenedesmus sp. |
6f |
0.9і |
0.54i |
Agricultural (anaerobically digested diary manure) |
Mix of Microspora willeana, Ulothrix zonata, Ulothrix aequalis, Rhizoclonium hieroglyphicum, Oedogonium sp. |
5.5 g m-2d-1 |
nd |
nd |
Agricultural (swine effluent, maximum manure loading rate) |
R. hieroglyphicum |
10.7 g m-2d-1 |
0.7і |
72і mg m-2d-1 |
Agricultural (swine effluent, +CO2, maximum manure loading rate) |
R. hieroglyphicum |
17.9 |
1.2і |
210 mg m-2d-1 |
Agricultural (digested dairy manure, 20x dilution |
Chlorella sp. |
81.4® |
13.6i |
11і |
TABLE 12.2 (Continued) Biomass and Lipid Productivities of Microalgae Grown on Various Wastewater Streams
Source: From Rawat et al. (2011). Note: nd — Not determined. a Estimated from biomass value of -1000 mg L-1 after 40 days. b Estimated from biomass value of 1.1 mg L-1 h-1. c Estimated from biomass value of 14.4 mg L-1 h-1. d Estimated from biomass value of 812 mg L-1 after 3 days. e Estimated from biomass value of 7 g L-1 after 10 days. f Estimated from biomass value of 197 mg L-1 after 31 days. g Estimated from biomass value of 1.71 g L-1 after 21 days. h Estimated from lipid productivity and lipid content value. i Fatty acid content and productivity determined rather than total lipid. |
The counting chamber methods are well established and frequently applied for microalgal enumeration due to their low cost and easy application. The three common types of counting chamber methods for microalgae enumeration are the (1) Sedgewick — Rafter counting slide, (2) Palmer-Maloney counting slide, and (3) haemocytometer counting slide (LeGresley and McDermott, 2010). The three methods require samples with high cell densities. The presence of contaminating particles in the same size range as the algae and failure of cells to separate after cell division may be possible sources of erroneous counts (Coutteau, 1996). Table 4.1 compares the merits and drawbacks as well as fundamentals of the three counting chamber methods.