Oscillatory flow bioreactors contain equally spaced orifice plate baffles in a tubular style reactor. This reactor has improved heat and mass transfer due to the oscillatory motion that is imposed on the net flow of the fluid, which means that the degree of mixing is independent of the net flow. This results in long residence times in rela­tively low length-to-diameter ratios. This reactor design, therefore, has the potential to decrease the energy required for mixing algae cultures, due to decreased pumping requirements, and also leads to decreased capital costs (Harvey et al., 2003).

5.3.3.2 Hybrid Production Systems

As open and closed systems offer different advantages and disadvantages, it seems practical that a combination of the two could provide the best of both worlds. This idea has been investigated in various configurations, either by circulating culture between open and closed reactors through a single growth stage, or by having a two-stage culture regime where cells are moved from one to the other at a certain point. A simple single-stage intermediate design is produced by enclosing or semi­enclosing open ponds in tunnels or greenhouses to improve temperature control and reduce evaporation and contamination. This is very effective in improving produc­tivity, particularly in colder seasons, but comes at increased capital cost (Vonshak, 1997). Pushparaj et al. (1997) described a system where an alveolar panel oriented toward the sun was coupled with an open raceway for gas transfer. Adding the panels improved the productivity of the pond from 0.18 to 0.31 g L-1d-1.

In two-stage configurations, culture is usually grown initially in closed PBRs to optimize the growth rate and minimize contamination of the inoculum, which is then moved to an open pond for the second growth stage. Integration of open and closed PBRs in this way could provide sufficiently large, clean inoculants to limited — duration culture in outdoor raceways in order to significantly limit adverse events (Greenwell et al., 2010).

The second cultivation stage often involves nutrient stress for accumulation of a metabolite such as lipids or pigments. The nutrient stress stage is suited to open ponds because the growth rate is naturally low and therefore not affected by the low light availability (Brennan and Owende, 2010). Initial culture in closed reactors also implies that the culture entering the pond is relatively dense and therefore less likely to be contaminated, particularly in a nutrient-deprived environment (Singh et al., 2011). This sort of system has been used for the production of astaxanthin from Haematococcus (Huntley and Redalje, 2006) and described for the production of biodiesel from Nannochloropsis (Rodolfi et al., 2009).

During the production of algal biodiesel, an algal cake remains and can form a feedstock for further product formation. Potential commodity by-products include an algal biomass suitable for animal feed, fermentation of the carbohydrate portion of the algal biomass to ethanol (Sandler and Murthy, 2010), or anaerobic digestion of the algal cake to biogas. Here we consider the co-production of algal biogas to demonstrate the multi-product approach (Campbell et al., 2010; Stephenson et al., 2010; Richardson et al., 2012). Stephenson et al. (2010) investigated two modes of cell disruption and assumed that only disrupted cells were digested. While the anaero­bic digestion of Spirulina platensis was similar in the presence and absence of cell disruption, disruption was essential for Scenedesmus, which also exhibited signifi­cant resistance to disruption. Collet et al. (2010) demonstrated anaerobic digestion of the algae to biogas without prior recovery of the lipid fraction for biodiesel. This approach allowed avoidance of the concentration and cell disruption steps, thereby reducing the energy and economic costs. The digestate following methane produc­tion has potential to provide a source of N and P for further algal growth. Partial recycle has been demonstrated in the Spirulina system.

Generally in wastewater, millions of microscopic and macroscopic organisms are widely distributed, originating from discharged domestic wastewater. These include bacteria, protozoa, viruses, and limited algal species. Many of these micro — and macroorganisms are considered harmless, and the large diversity of organisms is highly adapted to their conditions and effective in wastewater treatment and acti­vated sludge treatment within the treatment facility. Several recent publications have reported that wastewater provides an ideal medium for potential microbial growth (Kong et al., 2010; Cho et al., 2011; Christenson and Sims, 2011; Park et al., 2011a; Pittman et al., 2011; Rawat et al., 2011), irrespective of anaerobic or aerobic waste­water treatment (Abeliovich, 1986).

The detection of new and rare species is made easier due to the accessibility of classifications based on genotypic and phenotypic data. This will be valuable in the challenges facing systematic classification and the need for establishing well — defined taxa, a stable nomenclature, and enhanced identification procedures. Large — scale screening for bioactive compounds of industrial application necessitates rapid and unequivocal characterization of enormous numbers of algal isolates. Because these biocatalytic compounds hold persistent value as an input for the biotechnology industry, the conservation of microbial gene pools is critical. Ex-situ collections are and will continue to be an essential cradle for warranting that a source of living cells is available for research and manufacturing purposes. It is well documented that exploring the same or similar environments fails to reveal the same organisms again or even, if found, they would not exhibit the desired characteristics exhibited by the earlier strains. Nevertheless, maintenance of representatives of all identified species of algae and cell lines in ex-situ collections is unrealistic. Hence, it is suggested that future researchers and repositories should ensure the provision of the DNA rather than the organisms themselves. We are still largely in the hunter-and-gatherer stage of exploiting algae for food, bioactive compounds, and energy. Hence, further challenges in bioprospecting may includee the protection of intellectual property rights of original owners, a policy for strain distribution, and sharing and material transfer agreements.

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For the isolation of new strains from natural habitats, traditional cultivation tech­niques may be used, such as enrichment cultures (Andersen and Kawachi, 2005). A preferred preliminary step toward single-cell isolations would be enrichment cultures with growth media, soil and/or water extract, or supplementing with nutri­ents such as nitrate, ammonium, and phosphate or a trace metal. Alternatively, the proximate nutrient composition of the source samples may be analyzed and supplemented to the growth media. Algae survive under natural environments despite the fact that natural samples are often deficient in one or more nutrients. This may be due to the fact that bacterial action, grazing, and death of organ­isms recycle those nutrients. Sampling reduces the population of specific spe­cies that help recycle nutrients, and this nutrient stress leads to a decline of the target species. Enrichment can also be disadvantageous if the target species is unable to compete with the other autochthonous flora. Hence, selective culturing is a unique tool suited for enrichment culturing of lipid-producing microalgae. Once enriched, suspensions containing algae from samples may be centrifuged to increase the biomass concentration of desired cell density; then diluted in sterile water and passed through a 60-pm plankton net to remove zooplankton, and col­lected again using a 0.45-pm glass filter. The cells on the filter should be rinsed several times with sterile saline to remove bacteria and then inoculate to BG-11 medium (Rippka et al., 1979) for enrichment. However, some algal strains take weeks to months to be isolated by traditional methods (Anderson, 2005). For large-scale sampling and isolation efforts, high-throughput automated isolation techniques involving fluorescence-activated cell sorting (FACS) have proven extremely useful (Sieracki et al., 2005). Because of morphological similarities when comparing many algal species, actual strain identification should be based on molecular methods such as rRNA sequence comparison or, in the case of closely related strains, other gene markers.

Professor Faizal Bux serves as director at the Institute for Water and Wastewater Technology at the Durban University of Technology. He has 17 years of experience in the fields of water and wastewater treatment, bioremediation and biotransformation of industrial effluents for the production of valuable by-products, and biotechnological applications of microalgae including biodiesel. Professor Bux has a BSc (Honors) in microbiology, an MS in technology (biotechnology), and a doctorate in environmental biotechnology. He has published more than sixty scientific papers in leading Science Citation Index (ISI) journals and contributed to six book chapters, nine technical reports, and more than seventy conference presentations, both nationally and internationally. He also serves as an editor for ISI-listed journals and books. Professor Bux serves on the Management Committee of the International Water Association (IWA) specialist group, Microbial Ecology and Water Engineering. He is a member of many professional bodies and is a Fellow of the Water Institute of Southern Africa. He also serves as a scientific advisor for various NGOs, both in South Africa and internationally, especially with regard to water quality issues.

Xi

Contributors

Faizal Bux

Institute for Water and Wastewater Technology

Durban University of Technology Durban, South Africa

Tapan Chakrabarti

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Vikas S. Chauhan

CSIR-Central Food Technological Research Institute (CFTRI)

Mysore, India

Keshav Das

Biorefining and Carbon Cycling Program

College of Engineering The University of Georgia Athens, Georgia

Sivanesan S. Devi

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Ravi V. Durvasula

Department of Internal Medicine Center for Global Health University of New Mexico School of Medicine and

The Raymond G. Murphy VA Medical Center

Albuquerque, New Mexico

Sanniyasi Elumalai

Presidency College Chennai, India

Melinda J. Griffiths

Centre for Bioprocess Engineering Research

University of Cape Town, South Africa

Susan T. L. Harrison

Department of Chemical Engineering Centre for Bioprocess Engineering Research

University of Cape Town, South Africa

Subburamu Karthikeyan

Department of Agricultural Microbiology

Tamil Nadu Agricultural College Directorate of Natural Resource Management Coimbatore, India

Kannan Krishnamurthi

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Ramanathan Ranjith Kumar

Institute for Water and Wastewater Technology

Durban University of Technology Durban, South Africa

Rajesh Lalloo

CSIR Biosciences Pretoria, South Africa

Yun Liu

College of Life Science and Technology Beijing University of Chemical Technology Beijing, China

Dheepak Maharajh

CSIR Biosciences Pretoria, South Africa

Sandeep N. Mudliar

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Taurai Mutanda

Institute for Water and Wastewater Technology

Durban University of Technology Durban, South Africa

Terisha Naidoo

CSIR Biosciences Pretoria, South Africa

Renganathan Rajkumar

Department of Chemical and Process Engineering

Faculty of Engineering and Built Environment

University Kebangsaan Malaysia Bangi, Malaysia

Desikan Ramesh

Deparment of Farm Machinery Agricultural Engineering College and Research Institute Tamil Nadu Agricultural University Coimbatore, India

Durvasula V. Subba Rao

Center for Global Health, Department of Internal Medicine University of New Mexico School of Medicine and

The Raymond G. Murphy VA Medical Center

Albuquerque, New Mexico

Vadrevu S. Rao

Department of Mathematics,

Jawaharlal Nehru Technological University Hyderabad Hyderabad, India

Ismail Rawat

Institute for Water and Wastewater Technology

Durban University of Technology Durban, South Africa

Christine Richardson

Department of Chemical Engineering Centre for Bioprocess Engineering Research

University of Cape Town, South Africa Ravi Sarada

CSIR-Central Food Technological Research Institute (CFTRI)

Mysore, India

Yogesh C. Sharma

Department of Applied Chemistry Indian Institute of Technology (BHU) Varanasi, India

Ajam Y. Shekh

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Rheka Shukla

Biorefining and Carbon Cycling Program

College of Engineering The University of Georgia Athens, Georgia

Bhaskar Singh

Department of Applied Chemistry Indian Institute of Technology (BHU) Varanasi, India

Manjinder Singh

Biorefining and Carbon Cycling

Program

College of Engineering

The University of Georgia Athens, Georgia

Zahira Yaakob

Department of Chemical and Process Engineering

Faculty of Engineering and Built Environment

University of Kebangsaan Malaysia Bangi, Malaysia

Raju R. Yadav

CSIR-National Environmental Engineering Research Institute (NEERI)

Nagpur, India

Nodumo Zulu

CSIR Biosciences Pretoria, South Africa

Introduction

Taurai Mutanda

Institute for Water and Wastewater Technology Durban University of Technology Durban, South Africa

CONTENTS

1.1 General Overview…………………………………………………………………………………………….. 1

Acknowledgments…………………………………………………………………………………………………. 4

References……………………………………………………………………………………………………………… 4

The distribution of algal species is facilitated by both the selective action of the chemo-physical environment and by the organism’s ability to colonize a particular habitat. Numerous culture media have been developed for the isolation and cultiva­tion of microalgae. Some of them are modifications formulated based on the nutri­ent requirements of the organism. For instance, better growth of marine algae can be achieved by adding small quantities of natural seawater (less than 1% to 4%) rather than supplementing with artificial seawater. Likewise, Schreiber solution, a mixture of nitrate and phosphate, was based on the minimum requirement of the two elements by a diatom culture. Soil extract is amended to Schreiber’s medium for cultivating green dasycladalean Acetabularia and some unicellular benthic marine algae. Earlier algal media were formulated, to include antibiotics, vitamins, trace metals, and organic chelators such as citrate, which was later replaced with EDTA. Likewise, Chu’s medium No. 10 was composed based on proximate analyses of natu­ral samples such as eutrophic lakes. Antibiotics are generally added to the growth medium to inhibit contaminating protists. Germanium dioxide is suggested to inhibit the growth of diatoms. Antibiotics are also helpful as extensive cleansing agents. McDaniel et al. (1962) were able to purify algal cultures free of bacterial contami­nation using a procedure involving treatment with a detergent and carbolic acid. Various media configured for isolation and cultivation of algae are given in Table 3.4.

Microalgae are single-celled, ubiquitous, prokaryotic and eukaryotic primary photosynthetic microorganisms that are taxonomically and phylogenetically diverse. The advanced plant life of today is thought to have evolved from these simple microscopic plant-like entities. In general, the algae are a heterogeneous group of polyphyletic photosynthetic organisms with an estimated 350,000 known species (Brodie and Zuccarella, 2007). There are predominantly two prokaryotic divisions (Cyanophyta and Prochlorophyta) and nine eukaryotic divisions (Glaucophyta, Rhodophyta, Heterokontophyta, Haptophyta, Cryptophyta, Dinophyta, Euglenophyta, Chlorarachniophyta, and Chlorophyta). The biology of microalgae is interesting, and their enigma is due to their wide diversity as well as their plethora of habitats. The biology of microalgae is discussed extensively in Chapter 2 of this book.

Interest in microalgal cultivation is currently blossoming globally for a number of reasons. Microalgae are not extremely fastidious microorganisms but are found in diverse aquatic habitats. Microalgae can be found almost anywhere on Earth, in freshwater, marine, and hyper-saline environments (Williams and Laurens, 2010). The nutritional requirements of a wide array of microalgal strains are known, and the technology for microalgal cultivation is developing at a fast pace. The advent of genetic engineering protocols has brought new vistas to algal molecular systemat — ics. Recently, the general study of microalgae using genomics and molecular biol­ogy tools has attained phenomenal dimensions. The sheer number of microalgal strains from extreme environments that are yet to be discovered and identified is enormous (Brodie and Lewis, 2007). However, microalgal culture collection banks have been established as repository centers for these microorganisms (e. g., UTEX at The University of Texas at Austin).

The importance of microalgae in day-to-day life cannot be overemphasized. As the main primary producers, microalgal biomass is used for food and feed supplements (Lewis et al., 2000). Microalgae are important sources of commercial products such as polyunsaturated fatty acid (PUFA) oils (e. g., y-linolenic acid (GLA), arachidonic acid (AA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA)) (Spolaore et al., 2006). In addition, microalgae such as Dunaliella and Haematococcus are important sources of carotenoids such as P-carotene and astaxanthin, respectively (Spolaore et al., 2006). Furthermore, the cyanobacterium Anthrospira and the rhodophyte Porphyridium are the main commercial producers of phycobiliproteins (i. e., phycoerythrin and phycocyanin), which are used as natu­ral dyes and for pharmaceutical applications (Spolaore et al., 2006). Potential bio­technological applications and value-added products generated from microalgae are discussed in Chapter 10 of this volume.

Chlorella, Arthrospira, and Nostoc are cultivated worldwide for human and animal nutrition, owing to their chemical composition (Spolaore et al., 2006). Microalgae have been hailed as the panacea for the dwindling petroleum-based fuels, and the preponderance of shorter-chain fatty acids has significance for their potential as diesel fuels (Chisti, 2007; Williams and Laurens, 2010). The efficacy of using microalgal biomass and lipids as alternative biofuels is currently a topical issue. Biofuels such as biodiesel, biomethane, biohydrogen, biobutanol, etc., can be generated from microalgae (Chisti, 2007). Current research is targeting other novel potential biotechnological applications in aquaculture, cosmetics, pharmaceuticals, and animal and human nutrition. It is envisaged that future research should focus on microalgal strain improvement through genetic engineering, in order to diversify and economically improve product competitiveness (Spolaore et al., 2006). Microalgal genetic manipulation is still in its infancy and is a pertinent area of investigation in order to improve the quality and quantity of products generated from microalgae. However, the development of nondestructive product recovery techniques from con­tinuous cultivation systems will greatly improve product yield.

Successful microalgal cultivation and generation of these products calls for metic­ulous and rigorous microalgal strain selection. Two important steps in obtaining a robust and suitable microalgal candidate are (1) bioprospecting of target microalgal strain samples from diverse habitats, and (2) strain selection, isolation, and purifica­tion using conventional and advanced microbiological methods (Grobbelaar, 2009; Mutanda et al., 2011). Suitable microalgal strains can be obtained commercially from registered authentic culture collection centers. The microalgal strain of choice is maintained under laboratory conditions, either as a freeze-dried sample or as a slant on solid media at 4°C with routine subculturing. The ever-growing field of phy — cology has introduced new, exciting, and efficient techniques for maintaining micro­algal cultures at ultra-low temperatures (i. e., cryopreservation). Microalgal strain selection for biodiesel production is discussed in detail in Chapter 3 of this volume.

The enumeration of microalgae poses a real challenge due to the requirement of sophisticated equipment such as flow cytometers. The use of optical microscopes for cell counting is relatively cheaper, although not very accurate as compared to faster automated cell counting techniques (Guillard and Sieracki, 2005; Marie et al., 2005). Microalgal cells are counted in order to estimate the size of the cultured pop­ulation and to estimate the rate of culture growth (i. e., determination of the rate of population increase) (Guillard and Sieracki, 2005). Microalgal enumeration methods are described in detail in Chapter 4 of this volume.

The important factors affecting microalgal growth are light intensity, tempera­ture, nutrients, CO2 availability, pH, and salinity (Bhola et al., 2011; Rosenberg et al., 2011). Other factors such as conductivity, oxidation/reduction potential (ORP), total dissolved solids (TDS), and biological factors such as protozoa are also important. These factors must be closely monitored to prevent failure of the cultivation system, especially when growing microalgae on a large commercial scale.

There are essentially two commonly used methods for microalgal cultivation, namely open raceway ponds and photobioreactors. The design, and the pros and cons, of these cultivation systems are discussed in detail in Chapter 5. The open raceway system is amenable to large-scale microalgal cultivation because it is simple and cost effective to operate. Despite these attractive features, microalgal biomass harvesting still remains a huge challenge. Harvesting microalgal biomass is technically difficult because the biomass exists as a dilute aqueous suspension. Furthermore, microalgal cells are very difficult to remove due to their miniscule size (<20 pm), similar in density to water (Lavoie and De la Noue, 1986), and strong negative surface charge, particularly during exponential growth (Moraine et al., 1979; Park et al., 2011). It is a relatively daunting task to surmount these drawbacks.

Several methods are available for dewatering and recovering microalgal biomass, such as centrifugation, flocculation, gravity settling, microfiltration, and dissolved air floatation (DAF) inter alia (Lavoie and de la Noue, 1986; Molina Grima et al., 2003). The technology for microalgal biomass harvesting is still in its infancy, and trials on suitable combinations of these methods are currently underway (Williams and Laurens, 2010). The use of the centrifugation technique on a large scale is not cost effective due the colossal amounts of power consumption (Mutanda et al., 2011). The techniques available for microalgal harvesting and dewatering are discussed at length in Chapter 6.

There are several techniques that are used for extracting lipids from microal­gal biomass (Lewis et al., 2000). Most of these methods are destructive; however, it is desirable to develop nondestructive methods for continuous extraction of l ipids from live microalgal cells. The solvent extraction system using a mixture of solvents such as hexane and methanol are commonly used. Other methods are sonication and microwave-assisted extraction. The Bligh and Dyer method (1959) has been commonly used in many applications, whereby lipids are extracted from biological material using a combination of chloroform and methanol (Lewis et al., 2000). Extracting lipids from microalgal biomass is a real challenge because it is intracellular and therefore requires a cell disruption step. Currently, research is ongoing to develop cost-effective and efficient lipid extraction strategies (Molina — Grima et al., 2003; Williams and Laurens, 2010). Subsequent to lipid extraction, it is desirable to accurately identify the lipid and characterize the lipids using highly analytical techniques. This is done to establish whether the lipids extracted are suit­able for application to biodiesel production. Techniques that are widely used for the analysis of lipids are gas chromatography with mass spectrometry (GC-MS), liquid chromatography (LC), matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF), thin-layer chromatography (TLC), etc. Chapter 7 explores in detail the lipid extraction and identification techniques that are commonly used.

Microalgal lipids are converted into biodiesel through transesterification steps. Transesterification of microalgal lipids into biodiesel is accomplished either chemically or biologically using lipolytic enzymes. These methods are outlined in Chapter 8. To establish the feasibility of biodiesel production from microalgae, it is prudent to perform a life cycle analysis (LCA). The procedures involved in LCA are discussed in Chapter 9. Apart from generating biofuels and other value-added prod­ucts, microalgae cultivation is also profoundly involved in climate change abatement through CO2 sequestration. This important application of microalgae is discussed in Chapter 11. Microalgae can use wastewater rich in nitrates and phosphates as sub­strates for growth while simultaneously removing these macronutrients and thereby arresting eutrophication. Therefore, microalgae are involved in the phycoremedia — tion of domestic and industrial wastewaters, and this is achieved in high-rate algal ponds (Chapter 12). Finally, Chapter 13 discusses general microalgal biotechnology in terms of its potential as today’s “green gold rush.” The chapter gives an overview of advanced techniques such as genetic engineering of microalgae so as to increase lipid yield.

ACKNOWLEDGMENTS

The author hereby acknowledges the National Research Foundation (South Africa) for financial assistance.

REFERENCES

Bhola, V., Ramesh, D., Kumari-Santosh, S., Karthikeyan, S., Elumalai, S., and Bux, F. (2011). Effects of parameters affecting biomass yield and thermal behavior of Chlorella vulgaris. Journal of Bioscience and Bioengineering, 111: 377-382.

Bligh, E. G., and Dyer, W. J. (1959). A rapid method for total lipid extraction and purification.

Canadian Journal of Biochemistry and Physiology, 37: 911-917.

Brodie, J., and Lewis, J. (2007). Unravelling the Algae, the Past, Present, and Future of Algal Systematics. CRC Press, London.

Brodie, J., and Zuccarello, G. C. (2007). Systematics of the species-rich algae: Red algal classification, phylogeny and speciation. In The Taxonomy and Systematics of Large and Species-Rich Taxa: Building and Using the Tree of Life (Eds. T. R. Hodkinson and J. Parnell), Systematics Association Series, CRC Press, London, pp. 317-330.

Chisti, Y. (2007). Biodiesel from microalgae. Biotechnology Advances, 25: 294-306. Grobbelaar, J. U. (2009). From laboratory to commercial production: A case study of a Spirulina (Arthrospira) facility in Musina, South Africa. Journal of Applied Phycology, 21: 523-527.

Guillard, R. R.L., and Sieracki, M. S. (2005). Counting cells in cultures with the light micro­scope. In R. A. Andersen (Ed.). Algal Culturing Techniques (pp. 239-252). Elsevier Academic, Burlington, MA.

Lavoie, A., and De la Noue, J., (1987). Harvesting of Scenedesmus obliquus in wastewaters: Auto — or bioflocculation. Biotechnology and Bioengineering, 30: 852-859.

Lewis, A., Nichols, P. D., and McMeekin, T. A. (2000). Evaluation of extraction methods for recovery of fatty acids from lipid-producing microheterotrophs. Journal of Microbiological Methods, 43: 107-116.

Marie, D, Simon, N., and Vaulot, D. (2005). Phytoplankton cell counting by flow cytometry. In R. A. Andersen (Ed.). Algal Culturing Techniques (pp. 253-268). Elsevier Academic, Burlington, MA.

Molina Grima, E., Belarbi, E. H., Acien Fernandez, F. G., Robles Medina, A., and Chisti, Y. (2003). Recovery of microalgal biomass and metabolites: Process options and econom­ics. Biotechnology Advances, 20: 491-515.

Moraine, R., Shelef, G., Meydan, A., and Levi, A., (1979). Algal single cell protein from wastewater treatment and renovation process. Biotechnology and Bioengineering, 21: 1191-1207.

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.

Park, J. B.K., Craggs, R. J., and Shilton, A. N. (2011). Wastewater treatment high rate algal ponds for biofuel production. Bioresource Technology, 102: 35-42.

Rosenberg, J. N., Mathias, A., Korth, K., Betenbaugh, M. J., and Oyler, G. A. (2011). Microalgal biomass production and carbon dioxide sequestration from an integrated ethanol biore­finery in Iowa: A technical appraisal and economic feasibility evaluation. Biomass and Bioenergy, 35: 3865-3876.

Spolaore, P., Joannis-Cassan, C., Duran, E., and Isambert, A. (2006). Commercial applications of microalgae. Journal of Bioscience and Bioengineering, 101, 87-96.

Williams, P. J.B., and Laurens, L. M.L. (2010). Microalgae as biodiesel and biomass feed­stocks: Review and analysis of the biochemistry, energetics and economics. Energy and Environmental Science, 3: 554-590.

Using a micropipette or Pasteur pipette, or a glass capillary having a straight, bent, or curved tip is handy for single-cell isolation. Micropipettes enable fishing out a single cell from the sample after a series of transfers into sterile rinsing droplets, without the cell being damaged in the process. Finally, the single cell can be pipetted and transferred to the culture medium after microscopic examination. Lewin (1959) recommended placing the droplets on agar to reduce evaporation, but this depends on the size of the cells. Technical skill and expertise are important in order not to shear or damage the cell. The damage may be apparent as cessation of swimming in flagellates or a difference in light refraction due to broken frustules as in diatoms, and severe damage is evident by leakage of protoplasm. The traditional method of micropipette isolation can be successfully attempted with the use of ultra-pure sterile droplets for rinsing, as marine samples hold suspended particles.

TABLE 3.4

Common Media Used for Microalgal Strains from Diverse Aquatic Environments

TABLE 3.4 (Continued)

Common Media Used for Microalgal Strains from Diverse Aquatic Environments

Alternatively, many coccoid algae and most soil algae can be isolated on agar plates. It is the preferred isolation method because it is simple and requires no further processing. Streak or pour plating on suitable agar growth medium enables suc­cessful isolation, although few algae grow embedded in agar (Brahamsha, 1996). An improvised procedure is to make a fine or atomized spray of cells, usually a liquid cell suspension atomized with sterile air under pressure, which can then be used to inoculate or spread on agar plates. Similarly, a dilution method can be used, wherein a single cell is deposited in a test tube, flask, or well of a multiwell plate (Throndsen, 1978). Selection of the appropriate maximum dilution for plating depends on the probable cell density in natural samples. The dilutions can be effected in several ways, such as dilution with sterile culture medium, distilled water, seawater, and filtered water from the sample site, or some combination of these. Also, where neces­sary, salts of ammonium, selenium, or another element can be added as supplements to specifically isolate selected species.

When samples contain a wide variety of cells, centrifugation or settling can be foreseen. The target of concentrating the cells instead of obtaining an axenic culture can easily be achieved by gravity. Also, gravity comes in handy when the goal is to separate the larger and heavier cells from smaller algae and bacteria. Specifically for large dinoflagellates and diatoms, moderate centrifugation for a short duration is enough to pelletize them, and smaller cells can be decanted. Density gradient cen­trifugation with silica sol, Percoll™, etc. has been successfully employed to separate mixed laboratory cultures so that individual species can be separated into a sharp band (Reardon et al., 1979). Large, nonmotile algal cells can be effectively separated by settling. Hence, gravimetric settling is the choice if one aims for concentrating larger cells; however, it is not effective to obtain unialgal culture and hence suggests some combination with other procedures.