In the case of mechanical extraction, the feedstock (oil seed or algae biomass) is subjected to high pressure for rupture and release of the oil. The added advantages of mechanical extractions are that (1) no chemicals are used for extraction, (2) the process is free of chemicals in the products, and (3) the product is safe for storage. The major drawbacks of mechanical extraction are an inadequacy in complete lipid recovery from the feedstock, and the high energy inputs.

7.3.1.1 Oil Expeller

The first successful, continuously operating mechanical expeller was invented and patented by Anderson in the year 1900. An oil expeller is also called a screw press. The screw press is well suited for feedstocks having more than 30% oil content. The working principle of this machine is introducing pressure for crushing and break­ing the cells, followed by squeezing out the algal oil. An oil press is the simplest method used for oil extraction from algae biomass (Popoola and Yangomodou, 2006; Demirbas, 2009). The oil cake obtained from a screw press may contain some amount of residual oil (4% to 5% by weight). The oil extraction efficiency improves with applied pressure in a particular range, but too much pressure leads to less lipid recovery, more heat generation, and choking problems. Increased heat produced due to excessive pressure applied inside the screw press leads to darkening of the oil and a low-quality oil cake. The major drawbacks of this method are the high energy consumption, high maintenance cost with low capacities, labor intensiveness, long extraction time and less efficiency than other methods.

Despite the strength of algal oils as products, they comprise a low share in the omega-3 ingredients market, purely because of the lack of competitors in the mar­ketplace. Martek’s microalgal cultivation occurs in fermenters, ranging from 80 to 260 m3 in size (Ratledge, 2004; Spolaore et al., 2006; life’s DHA website, 2012). The microalgae are grown heterotrophically utilizing glucose and yeast extracts as carbon sources (Lee, 1997). Advancing microalgal technology has been the thrust
that keeps Martek highly competitive as an algal omega-3 ingredient manufacturer. Crypthecodinium cohnii was identified and commercially exploited by Martek as “a rich source of docosahexaenoic acid (DHA)” (Martek Corporation website, 2012), producing algal oil containing 40% to 50% DHA but no EPA (Ratledge, 2004; Ward and Singh, 2005; Spoloare et al., 2006).

Many argue that it is not imperative to consume both EPA and DHA as the human body efficiently converts EPA to DHA (Halliday, 2006). As a result, the potential new players on the algal market are focusing their efforts on increased production of high-purity EPA. The annual global demand for EPA is around 300 tonnes (Molina-Grima, 2003; Milledge, 2011). The current market value of fish oil EPA ethyl ester (95% pure) in bulk quantities is about $650 kg-1 (Belarbi et al., 2000); thus, a new source such as microalgal EPA is expected to be market competitive.

Because of their geometry, secretion of mucilage and variations in cell weight, micro­algae must be harvested in a species-specific, nongeneric manner (Benemann, 2008). Considerable process-oriented research is needed to optimize methods of algal har­vest, lipid extraction, and purification of by-products. Flocculation is widely used for algal harvest using various salts (Grima et al., 2003) such as aluminum, iron, potassium, zinc, chitosan, extracellular polymeric substances, bioflocculants such as Paenibacillus + aluminum sulfate, and organic cationic polymers. Co-bioflocculation (Nannochloris + diatoms) is also used, but necessitates extra effort to grow another alga. Pressure filtration (10 PSIG [pounds per square inch gage]) through four coni­cal felt media bags (1 pm), ultrasonication and grinding, cross-flow microfiltration/ ultrafiltration, and continuous foam separation are some of the other methods used for algal harvest. These techniques are labor intensive, expensive, and inefficient, with yields in the range of 30% biomass. Coagulation in the presence of chemicals is an alternative method for harvesting algae. Scenedesmus subspicatus, Selenastrum capricornutum, and Nannochloropsis sp. exposed to 5 mg L-1 barium concentra­tions bio-accumulate up to 88% to 99% of barium within 10 days (Theegala et al., 2001). Further treatment with 200 mg L-1 ferric chloride facilitated harvest of the metal-laden microalgae with an efficiency of nearly 99%.

Tetraselmis suecica can be concentrated up to 148 times using tangential flow filtration (TFF) and up to 357 times with polymer flocculation (PF); TFF requires a high initial capital investment and consumes 2.06 kWh m-3 while PF requires low initial investment with energy consumption in the range of 14.81 kWh m-3 (Danquah et al., 2009). The payback period, an important criterion for the investor, is 1.5 years for TFF and 3 years for PF. Passive and active immobilization techniques hold promise for harvesting high-value molecules such as storage products, antibiotics, hydrocarbons, hydrogen, and glycerol, and should be explored (Lebeau and Robert, 2006). Algaeventure Systems (AVS) has been developing an AVS Harvester that uses conveyor belts of capillaries to concentrate and dry Chlorella cultures. The esti­mated processing cost is $1.92 per ton compared to $875 per ton by centrifugation. One of the drawbacks is the required algal concentration of 3 g L-1; improvements are being pursued to increase harvesting efficiency.

An aqueous and a biocompatible organic phase (dodecane) bioreactor exist to extract P-carotene from Dunaliella salina cells. The organic phase continuously removes P-carotene (“milked”) from the cells with greater than 55% efficiency, and productivity is 2.45 mg m-2d-1, which is much higher than that of commercial plants (Hejazi et al., 2004). Several other methods of harvesting microalgae from liquid cultures are being developed, looking for a breakthrough to drastically reduce har­vesting and dewatering costs. Although details have not been published, mention should be made of the following:

1. Pretreatment of algae that involves application of 10 to 30 kV cm-1 electri­cal pulses for 2 to 20 ps to an algal slurry to rupture the cell walls and to release biodiesel compounds such as methyl hexadecanoate (Diversified Technologies at the University of Galway, Ireland).

2. Usage of amphiphilic solvents, such as acetone, methanol, ethanol, isopro­panol, butanone, dimethyl ether, or propionaldehyde, to separate out the proteins and carbohydrates from the lipids (Aurora Algae).

3. Harvesting, dewatering, and drying system utilizing surface physics and low-energy capillary action (Algaeventure Systems).

4. Single-step and live extraction of lipids (Origin Oil, James Cook University).

5. Hydrothermal liquefaction or thermal depolymerization (New Oil Resources).

6. A two-step catalyst-free algal biodiesel production process, using wet algal biomass and bypassing the drying and solvent extraction steps (University of Michigan).

7. Acoustic-focusing technology that generates ultrasonic fields that concen­trate algal cells into a dense sludge and extract oil (Solix).

Additionally, Kleinegris et al. (2011) have discussed product excretion, cell permea — bilization, and cell death as mechanisms to extract microalgal products. They pro­pose using two-phase systems that could circumvent the step of harvesting algal cells while the product is extracted in situ and prepared for downstream processing.

Historically, the vast majority of commercial production has been carried out in open ponds, and they are still the most widely applied reactor system in industrial microalgal processes (Carvalho et al., 2006). Open systems include natural water bodies, circular ponds, raceway ponds, and cascade systems. The main constraints on growth in open ponds are that it is impossible to control contamination, difficult to keep the culture environment constant, and expensive to harvest the dilute biomass (Carvalho et al., 2006). To maintain a monoculture in open ponds, highly selective culture conditions are necessary in order to guarantee dominance by the desired strain. For this reason, a limited number of species able to tolerate extreme condi­tions (e. g., Spirulina, which grows at high pH, and Dunaliella, which requires high salt concentrations) have been successfully grown. Open systems are susceptible to changes in temperature and irradiance due to local weather and climatic conditions,

thus making it difficult to maintain optimal growth conditions. Low cell densities are usually obtained, requiring large volumes to be processed during harvesting, and thus increasing the cost of product recovery (Carvalho et al., 2006).

9.3.1 Overview of Environmental Assessment of Algal Biodiesel

While prior studies on biofuels were largely limited to feedstocks of terrestrial plant origin, over the past 4 years, a number of studies have been published on the envi­ronmental analysis of algal energy processes (Lardon et al., 2009; Batan et al., 2010; Clarens et al., 2010; Jorquera et al., 2010; Sander and Murthy, 2010; Stephenson et al., 2010; Razon and Tan, 2011; Richardson et al., 2012a). Prior to this, environ­mental analyses of algal energy processes were limited to the work of Kadam (2002) and Sazdanoff (2006). The former considered the benefit of co-combustion of coal and algae in electricity generation, while the latter presented a model for the algal-to — biodiesel fuel cycle, including climatic data to simulate production at four locations in the United States. In most studies, analyses of the energy and global warming potential (GWP) have formed the key assessment criteria, with net energy recovery (NER) and LCA being the most frequently used approaches. In all cases, the absence of commercial-scale inventory data implies that scale-up estimates from laboratory — and pilot-scale data inform these analyses, requiring that a range of assumptions must be made on large-scale performance within these systems. In Table 9.3, the systems analyzed in each of the studies reported are summarized. These can be positioned within the context of Figure 9.1, which provides a block flowsheet of the integrated process for the production of biodiesel from microalgae and demonstrates the manner in which different studies focus on different components of the process. The findings in these studies are discussed in the following sections.

It must be emphasized that the immature nature of the microalgal biofuels process implies that the data have mostly been sourced from laboratory studies, modeling, and some pilot-scale research. Variability in data is a source of variation in results presented in the literature. As an example, Collet et al. (2010) estimated the energy for paddlewheels and pump of water at 0.2 and 0.153 kW-h per kilogram algae, respectively, whereas Clarens et al. (2010) suggested values of 0.035 and 0.029 kW-h per kilogram algae, respectively. Substitution with the lower values assumed by Clarens et al. (2010) leads to a 44% reduction in total energy demand. Further, lipid productivity has been recognized as a key factor in selecting condi­tions for biofuel production (Griffiths and Harrison, 2009; Rodolfi et al., 2009). Lipid productivity is the product of lipid content and specific growth rate. It is recognized that nutrient limitation results in compromised growth rates and high lipid con­tent; hence, care must be exercised to utilize compatible data when estimating lipid productivity. Examples are found in the literature where the high specific growth

TABLE 9.3 Studies of the Environmental Benefit or Burden of Processes for Algal Biofuels Algal Species; System Components Environmental Investigators Primary Product Objective and Process and Boundaries Reactor System Analysis Source of Data Lardon et at.. Chlorella To assess the environmental Infrastructure, Open raceways LCA of virtual Laboratory data 2009 vulgaris; impact of biodiesel from cultivation, facility, using biodiesel microalgae as dewatering, extract CML. Allocation technologically immature & convert, transport based on energetics process and to identify & distribute, barriers to large-scale environmental sustainability. Nutrient supply was compared with nutrient starvation; wet and dry extraction was tested; AD and nutrient recycle were used combustion. Batan et at.. Nannochlo ropsis To evaluate an industrial-scale Cultivation, Polyethylene PBR GREET model, Colorado State 2010 salina; model for microalgal dewatering, AD, bags NER, LCA based University pilot Microalgal biodiesel production in extract & convert, on GHG, allocation plant biodiesel photobioreactors and transport & by substitution compare microalgal biodiesel with soybean-based diesel and petroleum diesel distribute.

Open raceway ponds; flocculation and centrifugation	LCA, no allocation	Laboratory data; meteorological data
LCA — GHG, no allocation
Raceway pond
Raceway ponds; tubular reactors; flat-plate reactors Inoculum in photobioreactors or indoor ponds, open ponds for process	NER; energy LCA, no allocation	Laboratory-scale data from literature USLCI database
(Continued)

KJ о

TABLE 9.3 (Continued) Studies of the Environmental Benefit or Burden of Processes for Algal Biofuels Algal Species; System Components Environmental Investigators Primary Product Objective and Process and Boundaries Reactor System Analysis Source of Data Stephenson Chlorella To compare open raceways Infrastructure, Raceways; airlift LCA using EDIP, Laboratory — (own) et al., 2010 vulgaris; and tubular photobioreactors cultivation, tubular reactors market value and pilot-scale Biodiesel for algal biomass production dewatering, AD, allocation and (lit.) data used in a two-stage process with extract & convert, substitution (Rodolfi et ah, nitrogen limitation (phase 2) transport & 2009) for lipid accumulation; distribute, considered a range of DSP options, including AD of residual biomass combustion Collet et al., Chlorella vulgaris, To assess the environmental Infrastructure, Raceway pond LCA using CML, 2010 biogas, biodiesel impact of biogas generation cultivation, allocation by from microalgae, with nutrient recycle, and compare this to microalgal biodiesel dewatering, AD substitution Yang et al., Chlorella vulgaris To determine the water Cultivation, Raceway pond Water, no allocation 2011 footprint and nutrient usage dewatering, AD, of microalgal biodiesel extract & convert, production, considering use transport & of wastewater, nutrient recycle distribute

Photobioreactor NER Literature data

inoculum; raceway pond; gravitational settling

Tubular airlift reactor, NER and LCA horizontal tubular reactor, raceway

Tubular airlift reactor NER and LCA Own data from

lab

KJ KJ

Stephenson et al.

FIGU RE 9.1 Block flow diagram of the integrated microalgal biodiesel production process, indicating system boundaries of a selection of LCA studies.

rates of nutrient-replete growth are combined with high lipid content of nitrogen limitation to calculate productivity—such productivities are typically unobtainable. These examples highlight the importance of data quality and the challenges found owing to the absence of reliable production data. Clearly, sensitivity analysis forms a key component of these studies. Such sensitivity analyses are presented by, among others, Stephenson et al. (2010), Richardson et al. (2012b), Williams and Laurens (2010), and Norsker et al. (2011).

Microalgae have also been recognized as potential sources of various other VAPs for food applications. Green microalgae and cyanobacteria could be used as a rich source of the photosynthetic pigment chlorophyll. Chlorophyll acts as a che­lating agent and can be used in ointments, and in the treatment of liver recovery and ulcers (Puotinen, 1999). Chlorophyll can also be used as a natural colorant in foods (Humphrey, 2004). Microalgae that have already been grown in large volumes (e. g., Chlorella and Spirulina) could be explored for the commercial production of chlorophylls for food application.

Microalgae might also be a potential source for the commercial production of lutein. Lutein usually occurs in microalgae in its free nonesterified form. Microalga Muriellopsis sp. has been shown to accumulate higher contents of lutein with high productivities of biomass under photo-autotrophic conditions (Del Campo et al., 2000). Studies on the cultivation of Muriellopsis in closed photobioreactors as well as open-pond systems have been carried out (Del Campo et al., 2001, 2007; Blanco et al., 2007). The free lutein content of Muriellopsis biomass was found to be in the range of 0.4% to 0.6% on a dry weight basis (Del Campo et al., 2007). Scenedesmus sp. and Chlorella sp. have also been reported to accumulate lutein (Del Campo et al., 2000, 2004; 2007; Shi et al., 2006).

The red microalga Porphyridium has been shown to be a potential source of sulfated polysaccharides that form thermally reversible gels similar to macroalgae — derived polysaccharides, agar, and carrageenan. These gels have various commercial applications, including in foods, as gelling agents, thickeners, stabilizers, and emul­sifiers (Raja et al., 2008). As a microalga, Porphyridium may offer an advantage over macroalgae due to its relatively faster growth rate. Small-scale outdoor cultivation studies with Porphyridium have been carried out (Arad et al., 1985).

The chlorophyll, lutein, and polysaccharides from microalgae could be commer­cially important VAPs for food applications. However, processes for their commer­cial production from microalgae have not yet been developed and require further R&D studies as well as the development of markets for these products. Table 11.4 provides a list of some of the microalgal species with relevance for biotechnologi­cal applications in food. From Table 11.4, it is very clear that some progress has been made, and there are commercial microalgae applications, including pigments, fatty acids, and health foods. However, studies on the characterization of indigenous microalgae from natural habitats as potential sources of food, feed, and VAPs have remained relatively limited, and there is the need to tap into the vast biodiversity of microalgae growing in natural habitats under diverse climatic conditions for finding suitable candidate microalgae for various applications.

Centrifugation is similar to sedimentation, wherein gravitational force is replaced by centrifugal acceleration to enhance the concentration of solids. Particle size and den­sity difference are the key factors in centrifugal separation. Once separated, the algae concentrate can be obtained by simply draining the supernatant. Many research­ers have advocated this method for reliable recovery of microalgae (Mohn, 1980; Benemann and Oswald, 1996; Girma et al., 2003). Different types of centrifuges have been used, and their respective reliability and efficiency have been documented by several researchers. For example, Heasman et al. (2000) reported that 90% to 100% harvesting efficiency can be achieved via centrifugation. Sim et al. (1988) compared centrifugation, chemical flocculation followed by dissolved air flotation (DAF), and membrane filtration processes for harvesting algae from pilot-scale ponds treating piggery wastewater, and they found that none of these processes were completely satisfactory. Centrifugation was reported to be very effective but too cost and energy intensive to be applied on a commercial scale. This kind of harvesting is usually recommended in the production of high-value metabolites or as a second — stage dewatering technique for concentrating algal slurries from 1% to 5% solids to >15% solids, as it does have some limitations. Undoubtedly, it is an efficient and reliable technique for microalgal recovery but one should also keep in mind its high operational cost.

Terisha Naidoo, Nodumo Zulu,

Dheepak Maharajh, and Rajesh Lalloo CSIR Biosciences Pretoria, South Africa

CONTENTS

10.1 Introduction…………………………………………………………………………………………….. 137

10.2 Commercially Exploited Microalgae, Products, and Applications………….. 138

10.2.1 Carotenoids…………………………………………………………………………………. 138

10.2.1.1 Commercial Applications…………………………………………….. 140

10.2.1.2 Production Processes……………………………………………………. 142

10.2.1.3 Foresight………………………………………………………………………. 143

10.2.2 Phycobiliproteins………………………………………………………………………… 143

10.2.2.1 Commercial Applications of Phycobiliproteins…………….. 144

10.2.2.2 Production Process……………………………………………………….. 145

10.2.2.3 Future Potential……………………………………………………………. 146

10.2.3 Lipids………………………………………………………………………………………….. 146

10.2.3.1 Eicosapentaenoic Acid (EPA) and Docosahexaenoic

Acid (DHA)……………………………………………………………………. 147

10.2.3.2 DHA Production Process……………………………………………… 148

10.2.3.3 EPA Production Process……………………………………………….. 149

10.2.4 Other Potential Applications of Algal Biomass…………………………… 150

10.2.4.1 Cosmetic Extracts………………………………………………………… 150

10.2.4.2 Stable Isotope Biochemicals……………………………………….. 152

10.2.4.3 Human Nutrition………………………………………………………….. 152

10.2.4.4 Biofertilizers…………………………………………………………………. 153

10.2.4.5 Bioremediation/Phycoremediation……………………………… 153

10.3 Conclusion………………………………………………………………………………………………. 154

References…………………………………………………………………………………………………………. 156

10.1 INTRODUCTION

Microalgae represent a biodiverse resource (Metting, 1996; Pulz and Gross, 2004). The complexity of their chemical composition and range of biochemical products make these organisms exploitable resources for valuable and novel products in the food, feed, pharmaceutical, and research industries (Pulz and Gross, 2004). The market for these
applications is still emerging, but there have already been new areas of research in microalgal biotechnology to satisfy the new product demands of industry and consumers.

The use of microalgae as food for human consumption is an age-old tradition. Over 2,000 years ago, the Chinese used Nostoc to survive famine (Milledge, 2011). Species such as Arthrospira (Spirulina) and Aphanizomenon have also been utilized for decades as a source of food (Spolaore et al., 2006). Despite the plethora of historical usage by humans, microalgal culture is a fairly new area of biotechnology research, and its commercial application is virtually untapped (Spolaore et al., 2006; Milledge, 2011).

The interest in algal biomass came about in the 1950s as a result of an increase in the world’s population, and a forecast of insufficient protein supply triggered the search for alternative novel protein sources (Spolaore et al., 2006). Today, the use of algae for food still continues in many parts of the world; however, the large-scale production of algae to eradicate the food calorie and protein shortage has not fully materialized (Milledge, 2011).

There are a limited number of revolutionary companies that have persevered to large-scale production of algal biomass and products. The algal products are nor­mally marketed by the dominant players in the food and pharmaceutical industries. However, there is a significant gap in the microalgal market for expansion of existing products and the introduction of new products (Luiten et al., 2003; Pulz and Gross, 2004; Becker, 2007).

The efficiency of HRAPs depends on a variety of factors. Microalgal growth in HRAPs is similar to the production of biomass on artificial media. CO2, mixing, good light availability and penetration, and essential nutrient content, pH, and tem­perature are among the most important factors in achieving high biomass production and effective nutrient removal (Garcia et al., 2006; Pittman et al., 2011). Biotic factors such as synergistic bacteria, predatory zooplankton, and pathogenic bacteria may also affect the growth of microalgae. The variables will differ, depending on the type of wastewater and from one wastewater treatment site to another (Pittman et al., 2011). Nutrient content (nitrogen and phosphorus) in wastewater can be significantly higher than in conventional media. Nitrogen in wastewater is generally in the form of ammo­nia, which can inhibit algal growth at high concentrations (Pittman et al., 2011).

Carbon is assimilated from the atmosphere and CO2 produced by the oxidation of organic matter. The photosynthetic growth of algae utilizes CO2 as a carbon source for growth while producing oxygen as a by-product, which is utilized by bacteria to mineralize organic matter and produce CO2, which is consumed by algal photosyn­thesis. This aids in the reduction of greenhouse gas emissions (Munoz and Guieysse, 2006; Ansa et al., 2011; Park et al., 2011a). HRAPs are generally carbon limited and must be supplemented, potentially by utilization of flue gas for the improvement of nutrient removal efficiency (De Godos et al., 2010). The diurnal cycle affects pho­tosynthetic activity and thereby pH and nutrient removal efficiency (Garcia et al., 2006). The dissolved CO2 concentration has a direct effect on the pH of the sys­tem, as it is acidic in nature when dissolved in water. The cultivation pH directly affects the bioavailability of nutrients such as ammonia and phosphate. It may also aid in the proliferation of nitrifying bacteria (Craggs, 2005; De Godos et al., 2010). Both the pH and dissolved oxygen (DO) peak at midday due to the maximization of photosynthetic efficiency and thereby the removal of CO2 and an increase in DO of >200% saturation (Garcia et al., 2006; Park et al., 2011a). The consumption of CO2 and carbonic acid by photosynthesis increases the pH to basic levels (>11), thereby enhancing nutrient removal via the volatilization of ammonia and phosphorus pre­cipitation (Craggs, 2005; Su et al., 2012). At night, the removal efficiency decreases and may cease due to inadequate oxygen for aerobic respiration. Furthermore, the lower pH at night decreases nitrogen and phosphorus removal due to pH-dependant processes (Garcia et al., 2006; De Godos et al., 2010). High pH may also reduce nutrient utilization via significant inhibition of algal growth due to ammonia toxicity. Furthermore, a pH above 8.3 increasingly inhibits the bacterial activity and thereby the oxidation of organic matter by heterotrophic bacteria (Craggs, 2005; Ansa et al., 2011). The optimal pH for many freshwater algal species is 8, above or below which productivity decreases (Kong et al., 2010). Some algae are, however, capable of growth at pH > 10, such as Amphora sp. and Ankistrodesmus sp. (Park et al., 2011a). The pH stability in HRAPs is brought about by the balance of CO2 capture from the air, bacterial respiration, and algal CO2 uptake (Su et al., 2012).

The productivity of algal cultures—and thus nutrient removal—is light and tem­perature dependent. Photosynthesis increases with an increase in light intensity until the maximum rate is achieved at light saturation in the absence of nutrient limitation (Park et al., 2011a). Damage to the light receptors (photo-inhibition) occurs beyond the point of light saturation, thereby reducing productivity (Richmond, 2004). The potential for photo-inhibition to occur is more prevalent during the summer months, resulting in photosynthesis ceasing at midday (Olguin, 2003). With an increase in culture density, there is an increase in the shading effect. An algal concentration of 300 g TSS m-3 will absorb all the available light in the top 15 cm of the pond. Mixing is thus essential in reducing this effect (Ansa et al., 2011; Park et al., 2011a). Algal productivity increases with an increase in temperature. For most species of microal­gae under optimal culture conditions, optimal temperatures vary between 28°C and 35°C. The optimal temperature varies with nutrient and light limitation. An increase in temperature above the optimal level results in photorespiration, which reduces the overall productivity (Sheehan et al., 1998). Sudden changes in temperature can result in a substantial decline in algal growth. Temperature also affects the pH, oxygen, and CO2 solubility, as well as the ionic equilibrium (Park et al., 2011a).

HRAPs are susceptible to contamination by native algae and grazing by zooplank­ton and other algal pathogens. Attempts to grow algae as monocultures in HRAPs have failed due to said contamination (Sheehan et al., 1998; Park et al., 2011a). Protozoa and rotifers have the ability to reduce algal concentrations to very low levels in a period of just a few days (Benemann, 2008). Daphnia has the ability to reduce chlorophyll by 99% within a few days. Fungal parasites and viral infections have the ability to induce algal cell structure changes, and changes in diversity and succession, thereby reducing algal populations significantly (Park et al., 2011a; Rawat et al., 2011). Control of grazers and parasites may be achieved by physical methods such as filtration, low DO concentration and high organic loading rates, and chemical treatments such as the application of chemicals that mimic invertebrate hormones, increase the pH, and increase the free ammonia concentration. The most practical method of zooplankton control is the adjustment of the pH to 11, as many zooplanktons have the ability to tolerate low DO levels for extended periods of time. The toxic effects of high pH are augmented by the increase in free ammonia brought about by volatilization of ammo­nia at high pH. The effects of inhibitory substances on parasitic fungi require elucida­tion, and no general treatments for fungal control currently exist (Park et al., 2011a).

The use of a spectrophotometer for indirectly measuring microalgal cell density is done in conjunction with other methods, for example, gravimetric and counting chambers for the calibration curve (see below). The wavelength for the determina­tion of microalgal biomass is in the visible range of the electromagnetic spectrum (400 to 700 nm). It is recommended to do a calibration curve with samples of known microalgal cell numbers so as to extrapolate cell numbers from the standard curve. Spectrophotometric analysis is an easy quantification method although not very accu­rate because some other non-microalgal particles such as artifacts and dissolved and suspended solids may contribute to light absorption. However, it is recommended to ensure that the samples to be analyzed are free of other contaminants prior to analy­sis because this method does not discriminate noncellular materials from microalgal cells. One major drawback of spectrophotometric analysis is that culture conditions greatly affect chlorophyll content, which in turn determines absorbance. Microalgal cells grown in dark conditions have higher chlorophyll contents as compared to cells grown under high light intensity.

The use of light absorption is suitable for the estimation of microalgal population size rather than the determination of the actual number of individual cells.