Bhaskar Singh

Department of Applied Chemistry Indian Institute of Technology (BHU)

Varanasi, India

Yun Liu

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

Yogesh C. Sharma

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

CONTENTS

8.1 Introduction………………………………………………………………………………………………… 99

8.2 Esterification/Transesterification……………………………………………………………… 101

8.3 Thermochemical………………………………………………………………………………………. 103

8.4 World Scenario on Production and Application of Biodiesel……………………. 108

8.5 Biofuel/Biodiesel from Microalgal Oil as a Potential Alternative to

Other Fuels………………………………………………………………………………………………. 109

8.6 Conclusion………………………………………………………………………………………………… 110

Acknowledgments……………………………………………………………………………………………… 110

References…………………………………………………………………………………………………………. 110

8.1 INTRODUCTION

Microalgae have emerged as a potential feedstock for the production of biodiesel. The steps involved in the production of bio-oil and biodiesel from microalgae include cul­tivation, harvesting, dewatering and concentrating microalgae, extraction of oil/lip — ids from the microalgae, and separation of triglycerides and free fatty acids from the crude lipids (for the synthesis of biodiesel). The final step consists of pyrolysis or ther­mochemical catalytic liquefaction for the production of bio-oil and esterification or transesterification for the synthesis of biodiesel. The species of microalgae employed for the production of biofuel include Chlorella vulgaris, Chlorella sorokiniana, Sargassumpatens C., and Spirulina. The method of biodiesel production from algal biomass can be done either by direct transesterification or in two steps involving the extraction of oil from algae followed by transesterification. Economic in situ transesterification of the microalgae has been adopted that involves combining the two steps of lipid extraction and transesterification into a single step. Direct trans­esterification of the microalgae after cell disruption by sonication resulted in a high conversion of biodiesel (97.25%). The fuel properties of the biodiesel synthesized from the microalgal oil derived from Chlorella protothecoides showed high fuel quality with a cold filter plugging point of -13°C. A high composition of unsaturated fatty acid methyl ester content in the microalgal oil methyl esters (MOME) (i. e., 90.7 wt%) led to a low oxidation stability of the fuel (4.5 h). The chemical treatment, pyrolysis, or thermochemical catalytic liquefaction of microalgal oil for the synthe­sis of bio-oil eliminates the dewatering and drying steps. The major constituents of bio-oil obtained from brown microalgae Sargassum patens C. Agardh by hydrother­mal liquefaction consist of carbon (64.64%), followed by oxygen (22.04%), hydrogen (7.35%), nitrogen (2.45%), and sulfur (0.67%).

The alga belongs to the third-generation feedstock for the synthesis of a renewable fuel, biodiesel, or bio-oil. The first generation of feedstock was the crop species, and second-generation feedstock consisted of grasses and trees, which principally consisted of lignocellulosic biomass. With the limited availability of crop species, the focus of recent research has been on second — and third-generation feedstocks (Stephenson et al., 2011). Second-generation feedstocks have certain constraints that involve breaking the complex structure of lignin and converting the crystalline cellulose to amorphous cellulose. The process involved in second-generation biofuels makes it quite energy intensive. Hence, the focus of the research to a large extent in recent years has been on third-generation feedstocks, that is, microalgae (Lam and Lee, 2012). The conversion of microalgal lipids into biodiesel is a holistic approach that begins with the identifica­tion of an appropriate microalgal species that has a high potential to accumulate oil within the cells. The oil consists of crude lipids and neutral lipids. Neutral lipids con­sist of triglycerides, free fatty acids, hydrocarbons, sterols, wax and sterol esters, and free alcohols. Among these, only triglycerides and free fatty acids are saponifiable, and hence can be converted to biodiesel by esterification or transesterification. Crude lipids consist of neutral lipids along with pigments (Sharma et al., 2011). The triglycerides and free fatty acids are the part of microalgal lipids that can be converted to biodiesel or bio-oil. The microalgal biomass can be used for the production of biofuel, either by pyrolysis or through direct combustion or thermochemical liquefaction in which bio­oil is produced. Alternatively, the lipid can be derived from microalgal biomass and converted to biodiesel via transesterification (Kao et al., 2012).

In general, microorganisms that accumulate more than 20% to 25% of their weight as lipid are called oleaginous species (Kang et al., 2011). As these oleaginous micro­organisms can accumulate a large amount of oil within their cells, they can be very good feedstocks from which to extract oil and lipids that can be converted to biofuel (bio-oil or biodiesel). In some of the algae, the lipid content may be as high as 75% of their dry biomass. Most species of algae produce triglycerides (that can be utilized for biofuel production) and alkanes. A few algal species may also contain long-chain hydrocarbons that are formed via the terpenoid pathway (Sivakumar et al., 2012). Hence, the characterization of the algae is prerequisite to assessing the potential of the microalgae extract to be converted to biofuel. Once the triglyceride content in the microalgal species is determined, depending on their feasibility, the microalgae extract can be utilized for its conversion to either bio-oil or biodiesel. This chapter deals with the conversion of microalgal lipids into biodiesel or bio-oil, taking into account the following aspects: esterification/transesterification, and chemical method.

The consumption of microalgae is restricted to very few species, for example Spirulina, Chlorella, and Dunaliella (Jensen, 1993; Pulz and Gross, 2004). The market value of microalgal products (health foods) is estimated at US$20-25 million and it is by far the largest commercial application of microalgae (Metting, 1996). Spirulina and Chlorella are currently dominating the microalgal market. Spirulina is a source of protein that is comparable to meat and dairy products. Spirulina also contains high amounts of vitamin A and B12 (Metting, 1996).

Microalgal products to be used for human nutrition are usually sold in the form of tablets or powders (Metting, 1996; Radmer, 1996; Pulz and Gross, 2004). The pack­aged food industry, valued at US$2 trillion, is on the hunt for sustainable and natu­ral sources of fiber and healthy fats as ingredients for nutritionally high-value and — quality foods (Singh et al., 1996). Microalgal foods pioneer Solazyme-Roquette has created “high-lipid algal flour” (Daniells, 2011), intended for use as a main ingredi­ent alternative to make healthier processed foods such as chocolate milk (4.5% algal flour), frozen desserts, and even low-calorie salad dressings.

Extracts from microalgae are creating a new sector for microalgal products (Pulz and Gross, 2004). Products made from algal extracts include Chlorella health drinks and Spirulina liquid CO2 extracted antioxidant capsules. The microalgal biomass from Spirulina and Chlorella is not only used in human nutrition, but also in animal feed (Pulz and Gross, 2004), as it has been proven to support the immune system of animals. The market value of Spirulina and Chlorella is estimated at US$80 and $100 million, respectively (Radmer, 1996).

Biodiesel and
Value-Added Products

Microalgae are an invaluable biomass source with potential uses that could lead to environmental and economic benefits for society. Biotechnological Applications of Microalgae: Biodiesel and Value-Added Products

presents the latest developments and recent research trends with a focus on potential biotechnologically related uses of microalgae. It gives an analysis of microalgal biology, ecology, biotechnology, and biofuel production capacity as well as a thorough discussion of the value-added products that can be generated from diverse microalgae.

The book provides a detailed discussion of microalgal strain selection for biodiesel production, a key factor in successful microalgal cultivation and generation of desired biofuel products. It also describes microalagal enumeration methods, harvesting and dewatering techniques, and the design, and the pros and cons, of the two most common methods for cultivation—open raceway ponds and photobioreactors. Chapters cover lipid extraction and identification, chemical and biological methods for transesterification of microalgal lipids, and procedures involved in life cycle analysis of microalgae. They also examine the importance of microalgal cultivation for climate change abatement through CO2 sequestration and microalgae involvement in phycoremediation of domestic and industrial wastewaters.

The book concludes with a general discussion of microalgal biotechnology and its potential as a modern “green gold rush.” The final chapter provides an overview of advanced techniques such as genetic engineering of microalgae to increase lipid yield. This book provides a one-stop benchmark reference on microalgal biotechnology, considering all aspects, from microalgal screening to production of biofuels and other value — added products.

6000 Broken Sound Parkway, NW Suite 300, Boca Raton, FL 33487 711 Third Avenue New York, NY 1001 7 2 Park Square, Milton Park Abingdon, Oxon 0X14 4RN, UK

Although they are more expensive to build and run than open systems, the promise of improved yields, and the possibility of growing a wider range of species, has led to significant interest in closed reactors. It is much easier to control contamination and environmental parameters in closed systems, allowing the cultivation of more sensitive strains and expanding the potential product range. Biomass concentra­tions obtained are higher than in open systems, thus reducing the cost of harvesting. However, the capital and operating costs of closed reactors are higher than those of open ponds (Carvalho et al., 2006).

A large variety of PBR designs have been proposed, only a few of which have been commercialized (Greenwell et al., 2010). Most designs are based on the premise of optimizing light provision by maximizing the area-to-volume ratio, while ensuring a reasonable working volume, cost of reactor material, and mixing pattern (Carvalho et al., 2006). One of the major problems with closed reactors is temperature control, and the larger the area-to-volume ratio, the more susceptible the temperature of the medium is to changes in environmental temperature. The optimum light path length is 2 to 4 cm (Borowitzka, 1999), but most closed reactors have a larger diameter for ease of mixing, cleaning, temperature regulation, and to increase the working volume while reducing the cost of construction materials. Sedimentation is prevented by maintaining turbulent flow through mixing mechanically or by airlift.

An important and often overlooked feature of closed reactors is the ease with which they can be cleaned and sanitized (Chisti, 2007). Closed microalgal reactors are often presented as having the advantage of a decreased risk of contamination. Contamination can be avoided in closed reactors, but only if they are operated under sterile conditions, which adds to the cost (Scott et al., 2010). Due to their large size and surface area, closed reactors cannot be effectively sterilized by heat, and therfore require chemical steriliz­ers. These are not always 100% effective and sometimes require large volumes of sterile water to flush out the chemical agent. Most closed PBRs do not satisfy the good manu­facturing practice requirements for production of pharmaceutical products (Lee, 2001).

The most common designs are tubular (Miyamoto et al., 1988; Richmond et al., 1993; Borowitzka, 1996; Vonshak, 1997) or flat-plate (Hu et al., 1996; Vonshak, 1997) reactors. Both types usually operate with culture circulated between a light-harvesting unit, consisting of narrow tubes or plates, to provide a high surface area, and a reser­voir or gas exchange unit in which CO2 is supplied, O2 removed, and harvesting car­ried out. The circulating pump must be carefully designed so as to avoid shear forces disrupting algal cells. A variety of microalgae, including Chlorella and Spirulina, have been successfully maintained in both tubular and flat-plate PBRs (Molina Grima, 1999; Lee, 2001; Pulz, 2001; Carvalho et al., 2006).

Scale-up of any PBR design is challenging due to the difficulty in maintaining optimum light, temperature, mixing, and mass transfer in large volumes (Ugwu et al., 2008). Large-scale closed systems will likely be based on the integration of multiple units rather than increasing the size of a single reactor (Brennan and Owende, 2010).

Dewatering of the dilute algal suspensions required to minimize light limitation in the bioreactor is recognized as a key challenge in large-scale algal processes. This is aggravated by small algal cell size and a density only slightly greater than that of water. An initial dewatering step is required to achieve a solids concentration of 1% to 2.5% by mass prior to concentration using an energy-intensive operation such as centrifugation to a solids concentration of 5% to 20% (Benemann and Oswald, 1996). Ideally, flocculation and sedimentation are used for the primary dewatering, facilitated by the algal species selected (Lardon 2009; Stephenson et al., 2010). Other options include natural settling (Collet et al., 2010), filtration (Yang et al., 2011), and dissolved air flotation (Campbell et al., 2010). The decanter centrifuge, spiral-plate centrifuge, and rotary press are most typically proposed for the second dewatering step (Lardon et al., 2009; Campbell et al., 2010; Clarens et al., 2010; Collet et al., 2010; Stephenson et al., 2010). Stephenson et al. (2010) report a relative electricity consumption for flocculation of 1 unit versus 4 units for centrifugation and 14.4 units for the cultivation. On using a more dilute algal culture, as typically found with the raceway, both pumping energy and the need to increase flocculant supplied to main­tain its volumetric concentration impact the process.

Wastewater is one of the main sources of increasing water pollutant levels glob­ally (Gomec, 2010). An understanding of wastewater characteristics is essential in the design and operational processes of wastewater treatment and deserves ample research efforts. Wastewater is divided into two types: (1) municipal wastewater and (2) industrial wastewater. Wastewater is generally a combination of household and industrial waste, depending on the treatment collection system. These are generated during different human activities and are mixed together. Figure 12.1 provides a few

2. Agricultural and food processing:

a) Sugar refinery

b) Confectionery

c) Diary

d) Yeast fermentation

a) Shrimp production

b) Fish production

c) Salmon trout production

d) Trout production

FIGURE 12.1 Types of domestic and industrial wastewater.

TABLE 12.1

Chemical Composition and Characteristics of Untreated Wastewater

Contaminants

Total solids

Total dissolved solids

Fixed

Volatile

Suspended solids

Fixed

Volatile

Settleable solids BOD5, 20°C TOC COD

Nitrogen (total as N)

Organic

Free ammonia

Nitrites

Nitrates

Phosphorus (total as P)

Organic

Inorganic

Chlorides

Sulfate

Alkalinity (as CaCO3) Grease

Total coliforms

Volatile organic compounds

Source: From Rawat et al. (2011).

examples of municipal and industrial wastewater types. Municipal and industrial wastewater treatment by socio-economic biological treatment is a highly efficient technology (Metcalf and Eddy, 1991), and reclaimed wastewater can be used for various purposes. Table 12.1 shows the evaluation of weak, medium, and strong typical composition levels of domestic wastewater. A complete assessment of waste­water quality can be broadly classified into three main characteristics by its physical, chemical, and biological constituents and their sources (Figure 12.2).

Flocculation is used to separate microalgal cells from broth by the addition of one or more chemicals. Microalgal cell walls carry a negative charge that prevents self­aggregation within the suspension. This negative charge is countered by the addition of polyvalent ions called flocculants. These can be cationic, anionic, or nonionic, and they flocculate the cells without affecting their composition and/or being toxic. These flocculants have been classified into two groups, namely (1) inorganic agents, including polyvalent metal ions such as Al3+ and Fe+3 that form polyhydroxy com­plexes at suitable pH; and (2) polymeric flocculants, including ionic, nonionic, natural, and synthetic polymers. Among the former group, aluminum sulfate, ferric chloride, and ferrous sulfate are commonly used multivalent flocculants whose efficiency is directly proportional to the ionic charge. Fe3+ has been reported to be 80% effi­cient in harvesting different types of algae (Knuckey et al., 2006). The mechanism of polymer flocculation involves ionic interaction between polyelectrolyte and algal cells, resulting in the bridging of algae and formation of flocs. The extent of aggrega­tion depends on the charge, molecular weight, and concentration of polymers. It has been observed that binding capability increases with an increase in molecular weight and charge on the polymer. Algal properties such as the pH of broth, concentration of biomass, and its charge are equally important to consider when selecting a polymer. Tenney et al. (1969) found effective flocculation in Chlorella when using a cationic polyelectrolyte, whereas an anionic polyelectrolyte failed to do so. Divakaran and Pillai (2002) successfully used chitosan as a bioflocculant for Spirulina, Oscillatoria, Chlorella, and Synechocystis spp. The efficiency of the method is affected by media pH, and best results were recorded at pH 7.0 for freshwater and a lower pH for marine species. Organic flocculants are reported to be beneficial in terms of their lower sensitivity to media pH, low dosage requirements, and wider range of applications. Heasman et al. (2000) also studied chitosan as a flocculant for Tetraselmis chui, Thalassiosira pseudonana, and Isochrysis sp., and they found that only 40 mg L-1 of chemical was needed for complete aggregation, whereas 150 mg L-1 was needed for Chaetoceros muelleri. Microbial flocculants (AM49) were also studied by Oh et al. (2001) for the harvesting of Chlorella vulgaris. This flocculant was found to be bet­ter than other commonly used flocculants. Recovery of more than 83% solids when operating in the pH range 5 to 11 was recorded; this is higher than that when using aluminum sulfate (72%) or the cationic polymer polyacrylamide (78%).

Algae also have the property of auto-flocculation when supplemental CO2 supply is removed. Disruption of the CO2 supply during photosynthesis increases the pH, which results in the precipitation of magnesium, calcium, phosphate, and carbonate salts along with algal cells. The positively charged ions interact with the negatively charged algal surfaces and bind them, resulting in auto-flocculation. Sukenik and Shelef (1984) conducted a study on auto-flocculation in pond and laboratory-scale experiments, and reported some very promising results. The unavailability of condu­cive conditions—especially light and CO2—can, however, limit this process.

There is a demand for natural pigments to be applied in the food, pharmaceuti­cal, and aquaculture industries (Dufosse et al., 2005). The use of synthetic dyes in these industries is slowly declining due to their toxic effects (Dufosse et al., 2005). Compared to synthetic alternatives, microalgal carotenoids have the advantage of supplying natural isomers in their natural ratios (Pulz and Gross, 2004; Milledge, 2011). Microalgal pigments have been used as alternatives to synthetic pigments in various industries (Table 10.2).

Other than for coloring purposes, carotenoids have recently been used as anti­oxidants. Carotenoids have antioxidant effects that can be beneficial in countering diseases such as cancer, obesity, and hypertension (Inbaraj et al., 2006; Murthy et al., 2005). Table 10.3 indicates the applications of commercially exploited carotenoids in various industries.

The worldwide demand for carotenoids has been increasing at an average yearly rate of 2.2% (Guedes et al., 2011). Among the 400 known carotenoids, so far only a few have been exploited (Cosgrove, 2010; Milledge, 2011). The two most com­monly exploited algal carotenoids are P-carotene and astaxanthin, which are mainly produced by Dunaliella salina and Haematococcus pluvialis, respectively (Pulz and Gross, 2004; Spolaore et al., 2006; Del Campo, 2007; Cosgrove 2010;

TABLE 10.3 Commercially Available Carotenoids and Their Applications Food Coloring of Animals and of Supplements Carotenoid Type Coloring Animal Products (anti-oxidants) Annatto X Apocarotenal X Apocarotenal ester X Astaxanthin X X X Beta-carotene X X X

Vitamin A Source Cosmetic Ref. Vital Solutions market report, 2010 Vital Solutions market report, 2010 Vital Solutions market report, 2010 X Pulz and Gross, 2004; Vital Solutions market report, 2010; Milledge, 2011 X X Pulz and Gross, 2004; Spolaore et al., 2006; Del Campo, 2007; Vital

Solutions market report, 2010; Milledge, 2011

Inbaraj et al., 2006; Vital Solutions market report, 2010 Vital Solutions market report, 2010 Del Campo, 2007; Vital Solutions market report 2010

X Vital solutions market report, 2010

Del Campo, 2007; Vital Solutions market report 2010

Milledge, 2011). The market size for P-carotene is estimated at 1,200 tonnes per year and greater than US$280 million in sales volume per year (Pulz and Gross, 2004). The market price of natural P-carotene is much higher than that of synthetic P-carotene ($1,000 to $2,000 kg-1 for natural P-carotene versus $400 to $800 kg-1 for synthetic P-carotene). Although the price of natural P-carotene is higher than that of the synthetic form, preference is still given to the natural form because it has physical properties that make it superior to the synthetic form. The annual worldwide market of astaxanthin is estimated at US$200 million (Spolaore et al., 2006). The market size of astaxanthin is estimated at just below 300 tonnes per year, with a sales volume of less than US$150 million per year (Pulz and Gross, 2004; Spolaore et al., 2006).

Lutein and zeaxanthin are the other two carotenoids with great potential. These carotenoids are commonly derived from petals of Tagetes erecta and Tagetespatula, commonly referred to as marigold flowers (Del Campo et al., 2007). Microalgae also have an ability to accumulate these carotenoids. Lutein and zeaxanthin are known to selectively accumulate in the macula of the human retina. They protect the eyes from light and oxidative stresses (Kotake-Nara and Nagao, 2011).

The lutein extracted from other sources is usually 95% esterified, whereas in microalgae, lutein is found in the free nonesterified form. Muriellopsis sp. recorded lutein yields of 75 mg m-2d-1 in an outdoor open pond system (Del Campo et al., 2007). These values are similar to those obtained in a closed system (Harun et al., 2010; Del Campo et al., 2007). Overall, the free lutein content of Muriellopsis sp. biomass varies between 0.4% and 0.6%; which represents a higher content of esterified lutein than found in crown petals of Tagetes plants. The global market size of lutein is expected to hit $124.5 million by 2013 (Heller, 2008).

Zeaxanthin is mainly produced synthetically due to the fact that its content in natural sources (including microalgae) is considered very low for industrial produc­tion. The major problem underlying the commercial exploitation of zeaxanthin is the development of production processes that will result in the extraction of high amounts of zeaxanthin (Weiss et al., 2008). In 2006, the global market of zeaxanthin was estimated at $2 million (Heller, 2008). The awareness of this carotenoid still remains lower than that of lutein.

Some researchers have suggested that the production of algal biodiesel would produce greenhouse emissions, increase the water footprint, and require more energy than the production of biofuels from corn and canola feedstocks (Park et al., 2011a). These could, however, offset the use of wastewater as a nutrient source. Other researchers concluded that algal biodiesel production would be energetically viable but feedstock inputs would account for almost half of the energy produced, thus making the process not economically viable. The use of wastewater will offset this, giving a net increase in the net energy ratio (NER) (Sturm and Lamer, 2011). Conversion of algal biomass to energy via a multistage biorefinery process, including lipid extraction for biodiesel, utilization of residual biomass for combustion, and anaerobic digestion of biosolids, has the poten­tial to provide a significant amount of energy in the region of 4,610 kW-h d-1 to 48,000 kW-h d-1 (Sheehan et al., 1998; Sturm and Lamer, 2011). The energy requirement of conventional wastewater treatment is significantly higher than that of high-rate algal ponds. The Advanced Integrated Wastewater Ponds System (AIWPS), designed by Oswald and Green, LLC, requires up to 91% less energy (kW-h kg-1 BOD removed) than conventional systems (Olguin, 2003; Rawat et al., 2011).

Microalgal oxygen release provides the oxygen required for the proliferation of heterotrophic bacteria, thus negating the requirement for mechanical aera­tion as in conventional wastewater treatment. Conventional wastewater treatment costs approximately four times more than the use of HRAPs (Rawat et al., 2011). The AIWPS consists of advanced facultative ponds with anaerobic digestion pits, HRAPs, algal settling ponds, and maturation ponds in series (Craggs, 2005). This system requires 50 times more land area than conventional wastewater treatment viz. activated sludge, not taking into account the land area required for waste activated sludge disposal. Capital costs and operational costs of the AIWPS are half and less than one-fifth that of activated sludge, respectively (Park et al., 2011a). The supply of nutrients, water, and CO2 contributes from 10% to 30% of the total cost of commercial algal production (Benemann, 2008). Much of the cost of wastewater HRAPs is covered by the cost of wastewater treatment (Table 12.2). The costs of algal production and harvesting using wastewater treatment HRAPs have less environmental impact in terms of water footprint, energy, and tertilizer use. Recycling of growth media is used as a method of minimizing costs. Recycling can, however, cause a reduction in algal productiv­ity due to the increase in contamination and/or the accumulation of inhibitory metabolites (Park et al., 2011a).

12.2 CONCLUSION

Researchers are in general agreement that the use of wastewater treatment HRAPs is the only economical method currently available for algal production of biofuels. There are significant benefits to the use of wastewater HRAPs for the effective, low — cost treatment of wastewaters and algal biomass production for biofuels generation. There is, however, still a great need to optimize conditions for algal growth and nutrient removal under prevailing climatic conditions. Large-scale lipid optimiza­tion and harvesting of algal biomass still remains a challenge, and improvements in this area will subsequently decrease the overall cost of algae production and reme­diation of wastewater.

ACKNOWLEDGMENTS

The authors hereby acknowledge the National Research Foundation (South Africa) for financial contribution.

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Flow cytometry (FCM) is an automated cell counting technique that captures the fluorescence and scatter properties of the microalgal cells. The major advantage of automated cell counting techniques over optical microscopy is that they mini­mize the errors associated with human counting (Marie et al., 2005). The use of this highly sophisticated technique is hindered by the cost of the equipment as well as the requirement of highly skilled and trained personnel to operate the instrument. In addition, this technique is also plagued by limited sensitivity at lower microalgal cell concentrations. The solid-phase cytometer method for conducting total direct counts of bacteria is less biased and has performed significantly better than any of the microscopic methods (Lisle et al., 2004). Basic image analysis methods do not generally discriminate between phytoplankton and other material such as detritus and sediment in samples, thereby presenting a problem in the application to routine field samples. This technique may be more useful for the analysis of cultures and mono-specific high-density blooms (Karlson et al., 2010).

1.5 CONCLUSION

Microalgal enumeration can be tedious and cumbersome due to the small size of microalgal cells. Furthermore, this is exacerbated by the prohibitive cost of the available sophisticated equipment. To date, however, microalgal enumeration has been accomplished by gravimetric analysis, counting chambers, and flow cytometry.

Sedgewick-Rafter

Cultures and high cell numbers

1000 cells L-1 Limit of Detection (LOD) A rapid estimate of cell concentrations

Accurate results only when sample contains high algal cell densities Method-easy to learn and use; highly trained taxonomist needed for verification of species identification Compound Microscope Cover slips Pipettes Sedgewick-Rafter slides Compound microscope: £2500/US$3250 Sedgewick-Rafter slides: Perspex: £50/ US$65 Glass: £166/US$213 £1/US$1.3 20 min

This depends on the sample density

This depends on the sample density

Only one sample at a time Dependent on preservative used

Palmer-Maloney

Cultures and high cell numbers as in bloom situations 10,000 cells L-1 (LOD)

A rapid estimate of high cell concentrations

Accurate results only when sample contains very high algal cell densities Method-easy to learn and use; highly trained taxonomist needed for verification of species identification Compound Microscope Cover slips Pipettes Palmer-Maloney slides Compound microscope: £2500/US$3250 Palmer-Maloney slides: Ceramic-£60/ US$80 Stainless steel-£170/US$230 £1/US$1.3 5 min

10-30 min/sample depending on cell density

14-30 min/sample depending on the sample density Only one sample at a time Dependent on preservative used

Haemocytometer

Cultures and extremely high cell concentrations of small organisms

10,000,000 cells L-1 (LOD)

A rapid estimate of extremely high cell concentrations

Accurate results only when sample contains extremely high algal cell densities

Method-easy to learn and use; highly trained taxonomist needed for verification of species identification

Compound Microscope Pipettes Haemocytometer slide with cover glass.

Compound microscope: £2500/US$3250 Haemocytometer slide: £200/US$230

£1/US$1.3

5 min

<20 min/sample depending on the sample density

<30 min dependent on target species

Only one sample at a time

Dependent on preservative used

Source: Adapted from LeGresley and McDermott, 2010.

The invention and development of advanced techniques such as fluid imaging

alleviates the drawbacks of flow cytometry.

ACKNOWLEDGMENTS

The authors hereby acknowledge the National Research Foundation (South Africa)

for financial contribution.

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