Ranganathan Rajkumar and Zahira Yaakob

Department of Chemical and Process Engineering University of Kebangsaan, Bangi, Malaysia

CONTENTS

2.1 Taxonomy……………………………………………………………………………………………………. 7

2.1.1 General Characteristics…………………………………………………………………….. 7

2.2 Morphological Identification………………………………………………………………………… 8

2.3 Molecular Identification……………………………………………………………………………….. 9

2.4 Habitats………………………………………………………………………………………………………. 12

2.4.1 Freshwater………………………………………………………………………………………. 12

2.4.2 Marine…………………………………………………………………………………………… 13

References…………………………………………………………………………………………………………… 14

2.1 TAXONOMY

Algae are a diverse group of organisms that can perform photosynthesis efficiently. On the basis of morphology and size, algae can be subdivided and are classified into two main categories: macroalgae and microalgae. Macroalgae consist of mul­tiple cells that organize into structures resembling the roots, stems, and leaves of higher plants (e. g., kelp). Microalgae are an extremely diverse group of primary producers present in almost all ecosystems on Earth, ranging from marine, fresh­water, desert sands, and hot springs, to snow and ice (Guschina and Harwood, 2006). They are colonial or single-celled organisms that have garnered increasing amounts of attention and interest for industrial purposes. They are categorized into divisions based on various characteristics such as morphological features, pig­mentation, the chemical nature of photosynthetic storage products, and the orga­nization of photosynthetic membranes. The four most important algal groups in terms of abundance are green algae (Chlorophyceae), diatoms (Bacillariophyceae), blue-green algae (Cyanophyceae), and golden algae (Chrysophyceae) (Khan et al., 2009). According to estimations reported by Cardozo et al. (2007), they include between 200,000 and 800,000 species, of which about only 35,000 species have been described.

Although single-celled, colonial, or filamentous algae growing on the agar surface can be isolated by streak plate or spraying, any flagellates as well as other types of algae require the use of advanced techniques. A unialgal culture would contain only one kind of alga, usually a clonal population, but may contain other life forms such as bacteria, fungi, or protozoa. Alternatively, cultures may be axenic, in that they contain only one species of alga. Unialgal cultures are best isolated by targeting the isolation of the zoospores immediately after release from parent cell walls as those cells that begin attaching to surfaces are likely to add contaminants. The algal isola­tion techniques involving cell separation pose limitations with highly heterogeneous samples or when the cells are suspended in a solution of different chemicals, biomol­ecules, and cells. This can be overcome by employing a micromanipulator, which successfully permits the separation of a single cell from a liquid culture. The single cell can be easily separated from an enriched environmental sample and grown in liquid medium as monoculture or in agar plates, thus facilitating a significant time saving over the conventional plating technique. The micromanipulator is the ideal tool for algal screening and isolation, provided the person handling it has acquired skill in handling the equipment (Kacka and Donmez, 2008; Moreno-Garrido, 2008). Using micromanipulation techniques requires expertise and skill. It requires the handling of an inverted microscope or stereo zoom microscope with a magnification up to 200x. Phase contrast or dark-field microscopy offers advantages. Capillary tubes or hematocrit tubes of approximately 1 mm diameter x 100 mm long are used for picking individual cells (Godhe et al., 2002; Knuckey et al., 2002).

High-throughput cell sorting is possible when coupled with flow cytometry, which facilitates the rapid and efficient screening of microalgal strains. Microalgae possess different photosynthetic pigments, emitting various auto-fluorescence, which can be applied in flow cytometry to identify algae (Davey and Kell, 1996). Literature on the isolation of microalgae from natural waters employing flow cytometric cell sort­ing is available (Reckermann, 2000; Crosbie et al., 2003). Chlorophyll is used as a fluorescent probe to distinguish different strains of microalgae. Reckermann (2000) and Sensen et al. (1993) used the chlorophyll auto-fluorescence (CAF) properties of eukaryotic phytoplankton, diatoms, and pico-autotrophic cells for isolation of axenic cultures, whereas Crosbie et al. (2003) used both red and orange auto-fluorescence to differentiate species of algae. Similarly, green auto-fluorescence (GAF), which is common in both autotrophic and heterotrophic dinoflagellates, is also a valuable taxonomic consideration (Tang and Dobbs, 2007). Hence, flow cytometry, coupled with cell sorting, can signify a vital tool for screening and exploiting microalgal strains for specific drives, including biodiesel feedstock development. As compared to fluorescence microscopy, flow cytometry helps the investigator perform rapid and quantitative experimentation. Fluorescence-activated cell sorting (FACS) permits cells with a specific characteristic—or indeed a combination of characteristics—to be separated from the sample. Sinigalliano et al. (2009) compared electronic cell sorting and conventional methods of micropipette cell isolation with dinoflagellates and other marine eukaryotic phytoplankton. Fragile dinoflagellates such as Karenia brevis (Dinophyceae) were distressed upon conventional micropipette procedures while cells were viable on electronic sorting. However, electronic single-cell sorting combined with automated techniques for growth screening has the possibility of screening novel algal strains (Sinigalliano et al., 2009). The benefits and shortcom­ings of the microalgal isolation and purification protocols described in this section are summarized in Table 3.5.

In addition, several immunological and nonimmunological methods to isolate desired unicellular algal cells exist. The immunologic reaction of a specific inte­grated protein on the membrane decides the protocol for cell separation. Large-scale commercialized cell separation involves techniques such as FACS (Takahashi et al., 2004), magnetic-activated cell sorting (Han and Frazier, 2005), and affinity-based cell sorting (Chang et al., 2005), all of which are highly specific and selective. But the limitation is that the immunologically isolated cells may undergo trauma and the inclusive separation system involves high cost. Further, immunoreactions and follow-up elution with capturing antibodies are quite complicated processes. Alternatively, nonimmunological techniques such as dielectrophoresis (Doh and Cho, 2005), hydrodynamic separation (Shevkoplyas et al., 2005), aqueous two-phase system (Yamada et al., 2002), and ultrasound separation (Petersson et al., 2004) have also been employed. These methods work based on the interactive physico-chemical property of a cell with that of the surrounding media, and lack specificity.

The main characteristics of microalgae are primarily simple morphological features that can easily be observed under a light microscope. Cyanophyceae consist of prokary­otic cells commonly called the blue-green algae. They are similar to Gram-negative bacteria, based on the nature of the cell wall, cell structure, and capacity to fix atmo­spheric nitrogen; hence they are called cyanobacteria. However, they possess the pho­tosynthetic system chlorophyll-а, accessory pigments, and thallus organization that resembles other algae. Cyanophyceae members can be broadly divided into coccoid forms and filamentous forms. The coccoid has various forms, from single cell to aggre­gates of unicellular cells; regular or irregular colonies; and pseudofilamentous and pseudoparenchymatous conditions. The filamentous forms exist as simple uniseriate filaments to heterotrichous filaments, which may be differentiated into heterocysts and akinetes. These are ubiquitous in nature, occurring in several habitats with extreme conditions (i. e., temperature, light, pH, and nutritional resources). They are found abun­dantly in a variety of natural and artificial aquatic ecosystems. Cyanophyceae members can be easily identified within a mixture of other algae by their distinct blue-green color.

The Chlorophyceae constitute a major group of algae occurring in various habi­tats. The cells are usually green in color due to the presence of pigments such as chlorophyll-а and — b. The cells contain chloroplasts of various shapes that are located differently in each group of organisms. In addition, the chloroplasts also contain pyrenoids. The nucleus of this group may present either singly or in multiples but in major organisms occurs singly although some genera are multinucleate. Flagellated cells are common either in the vegetative or reproductive phase. There is at least one group without any flagellated cells.

Euglenophyceae members are unicellular, motile, and usually contain one promi­nent flagellum and in some cases two flagella. The anterior position of a cell has a visible gullet, and many dissimilar chloroplasts are found in autotrophic forms and are absent in other forms. Euglenoid cells are enclosed by a proteinacious pellicle and help the organisms achieve pleomorphism. These are widely distributed in all types of water bodies, particularly in organic-rich aquatic ecosystems.

Bacillariophyceae members are popularly known as diatoms. They are basically unicellular, and also occur as pseudofilaments or aggregated in colonies. The cell wall of a diatom is impregnated with silica and they have been well preserved as microfossils. The diatom cell is also called a frustule, and the classification of dia­toms is based on the pattern of ornamentation on their wall. The cells have either radial or bilateral symmetry. The frustules consist of two halves (epitheca and hypotheca) and connecting girdle bands. The surface of the valve has typical mark­ings. Punctae are regularly or irregularly arranged to form striae. Areolae are pores or chambers within the valve wall. Costae are elongated thickenings of the valve wall due to heavy deposition of silica. The valves of some diatoms have an open­ing or fissure along the apical axis called the raphe. The presence of the raphe or its absence on the walls of the diatoms has been one of the features in the identification of diatoms and distinguishing different genera. The radially symmetric forms are grouped as Centracles and the bilaterally symmetric ones are Pennales.

An ideal screen would consider growth physiology, including cell size and numbers, and metabolite production of algal strains. The algal growth physiology for biofuel encompasses a number of parameters, such as maximum cell density, maximum spe­cific growth rate, and tolerance to environmental variables such as temperature, pH, salinity, oxygen levels, CO2 levels, and nutrient requirements (Chisti, 2007; Brennan and Owende, 2010). Because all these parameters require significant experimental effort, the development of automated systems that provide information regarding all parameters simultaneously would be helpful. Screening for metabolite production may involve determining the cellular composition of proteins, lipids, and carbohy­drates, and measuring the productivity of the organism regarding metabolites useful for biofuel generation. The exact screenings employed would depend on the cultiva­tion approaches and fuel precursor desired. For example, a helpful screening for oil production would allow for distinguishing between neutral and polar lipids, and would provide fatty acid profiles. Furthermore, many strains also secrete metabolites

Advantages and Disadvantages of Microalgal Purification Techniques

into the growth medium. Some of these could prove valuable as co-products, and new approaches are needed to develop screening methods for extracellular mate­rials. For mass culture of a given algal strain, it is also important to consider the strain’s robustness, which includes parameters such as culture consistency, resil­ience, community stability, and susceptibility to predators present in a given envi­ronment. Previous studies revealed that algal strains tested in the laboratory do not always perform similarly in outdoor mass cultures (Sheehan et al., 1998). Therefore, to determine a strain’s robustness, small-scale simulations of mass culture conditions must be performed.

At this time, the bottleneck in screening large numbers of algae stems from a lack of high-throughput methodologies that would allow simultaneous screening of multiple phenotypes, such as growth rate and metabolite productivity. Solvent extraction, for example, is the most common method for the determination of lipid content in algae, but it requires a significant quantity of biomass (Bligh and Dyer, 1959; Ahlgren and Merino, 1991). Fluorescent methods using lipid-soluble dyes have also been described, and although these methods require much less biomass (as little as a single cell), it has not yet been established if these methods are valid across a wide range of algal strains (De la Jara et al., 2003; Elsey et al., 2007). Further improvements in analytical methodology could be made through the development of solid-state screening methods. Not only are rapid screening procedures necessary for the biofuels field, but they also could prove extremely useful for the identifica­tion of species, particularly in mixed field samples necessary for the future of algal ecology. They could also reduce the number of redundant screens of algal species.

Understanding biodiversity is critical in ecological research because it unravels the role of each single species in the ecosystem in mediating the environment for the entire biological community. The microalgal biodiversity of a region has economic value and, hence, any loss in biodiversity is of serious economic concern. This emphasizes the need and importance of biodiversity conservation. A survey of the literature on microalgae diversity during the past four decades has revealed that ecosystems harbor a large number of algae belonging to various groups. Despite the availability of elaborate monographs on specific groups of algae, any significant amounts of literature on the taxonomy of several algal species and genera remain scarce and scattered.

Proper identification of the algal taxa has always been considered “not an easy task.” Biochemical investigations of, and research into, environmentally important organisms have diversified phycological research and allowed it to enter a whole new phase altogether. Discovery of the potential of microalgae for industrial production of certain chemicals of pharmaceutical value led to the involvement of multifari­ous types of scientists in understanding the algae. Researchers on the threshold of algal taxonomic study and nonbotanists who are intrigued to know about the algae for pursuing research of their own interest are overwhelmed by the huge literature available. The present work therefore is an attempt to bring together a basic way of identifying all microalgae belonging to many of the genera of various groups of algae that occur abundantly and commonly in all ecosystems. Photographs were prepared for various groups of algae as described here. These photographs con­tain taxonomic information on the individual groups of organisms, especially the identification of genera and species. Different species of microalgae were recorded from water samples. All species of microalgae belonged to two major groups of algae, namely Bacillariophyceae and Dinophyceae. The Bacillariophyceae mem­bers were represented by Mastogloia paradoxa Grun., Rhabdonema adriatium Ktz., Synedra gruvei Grunow, Chaetoceros orientalis Schiller, Nitzschia draveillensis Coste & Ricard, Pleurosigma formosum Wm. Smith, Coscinodiscus janischii var. arafurensis Grun., Cocconeis scutellum Ehrenb., Podocystis spathulata (Shadbolt) Van Heurck, Actinocylus octonarius Ehrenb., Biddulphia biddulphiana (Smith) Boyer, Thalassionema nitzshioides Grun., Rhizosolenia setigera Brightwell, and Thalassiothrix longissima Cleve & Grun. The Dinoflagellates are represented by Ceratium hirundinella (Muller) Dujardin, Ceratium longipeps (Bailey) Grun., Ceratium trichoceros (Ehrenberg) Kofoid, and Gymnodinium sanguineum Hirasaka (Figures 2.1, 2.2, 2.3, and 2.4; see color insert) (Rajkumar, 2010).

Conventional methods of solvent extraction and gravimetric determination for lipid quantification (Bligh and Dyer, 1959) are laborious and time consuming. Moreover, approximately 10 to 15 mg wet weight of cells (Akoto et al., 2005) must be cultured for any appreciable extraction and derivatization. However, in-situ lipid content mea­surements would significantly reduce the quantity of sample as well the preparation time required. Accordingly, there is greater interest in a rapid in-situ measurement of the lipid content of algal cells (Cooksey et al., 1987). Nile Red (9-diethylamino — 5H-benzo[a]phenoxazine-5-one), a lipid-soluble fluorescent dye, has been com­monly used to evaluate the lipid content of animal cells and microorganisms such as yeasts and fungi (Genicot et al., 2005) and specifically extended to microalgae (Cooksey et al., 1987; Elsey et al., 2007). Nile Red is relatively photostable and pro­duces intense fluorescence in organic solvents and hydrophobic environments, which makes them a better candidate for in-situ screening for lipids. Furthermore, neutral and polar lipids can be clearly differentiated due to polarity changes in the medium as evinced by a blue shift in the emission maximum of Nile Red (Greenspan and Fowler, 1985; Laughton, 1986; Cooksey et al., 1987; Lee et al., 1998). The solvent system used for Nile Red would determine the emission spectra of the dye (Elsey et al., 2007). However, the thick cell walls of microalgae inhibit the permeation of Nile Red, and this is variable among algal species, requiring the use of high levels of solvents such as DMSO (20% to 30% v/v) and elevated temperatures (40°C) (Chen et al., 2009). Then again, Chen et al. (2011) developed a two-step microwave-assisted staining method for in vivo quantification of neutral lipids in green algae with thick, rigid cell walls that prevents penetration of the Nile Red dye into the cell. This may also be appropriate for other classes of algae that do not stain properly with Nile Red. Hence, a Nile Red assay can be used as a tool for screening oleaginous algal strains as well as quantitatively determining the neutral lipids in algal cells (Figure 3.2; see color insert).

Recently, another class of lipophilic fluorescent dye BODIPY® 505/515 (4,4-difluoro-1,3,5,7-tetramethyl-4-bora-3a,4a-diaza-s-indacene) has been used to potentially stain microalgal lipids. BODIPY staining lets the lipid droplets stain green and the chloroplasts stain red in live algal cells (Cooper et al., 2010). BODIPY 505/515 is advantageous over Nile Red in emitting a narrower spectrum (Cooper et al., 2010;

Govender et al., 2012). This facilitates the fluorescence distinction of lipid bodies, resulting in better resolution and thus is important for seamless confocal imaging (Cooper et al., 1999). Furthermore, unlike Nile Red, BODIPY 505/515 does not fix to cytoplasmic constituents other than lipid bodies and chloroplasts. This discerning property of BODIPY 505/515 to bind to lipid bodies alone offers rapid screening and isolation of hyper-lipid producing algal strains. Bigelow et al. (2011) developed a rapid, single-step, laboratory-scale in-situ protocol for GC-MS (gas chromatogra­phy with mass spectroscopy) lipid analysis that requires only 250 qg dry mass per sample. When coupled with fluorescent techniques using Nile Red or BODIPY dyes and flow cytometry for cell sorting, the aforesaid GC-MS analysis allows throughput screening of lipid-producing algal strains from varied environments. Upon isolation, purification, and identification of a hyper-lipid producing algal strain, the researcher would be interested in the physiological traits such as the photosynthetic efficiency, carbon fixation rate, growth rate, etc. Alternatively, infrared analysis, which does not depend on stain application but rather detects specific molecular absorption bands to give approximate concentrations, can be used for the detection of many metabolites, including lipids. This method has recently been applied to detecting changes in algal cell composition during nitrogen starvation (Dean et al., 2010).

Spectroscopic methods such as near-infrared (NIR) and Fourier transform infra­red (FTIR) spectroscopies have been established to predict the levels of spiked polar and neutral lipids in algal cells based on multivariate calibration models (Laurens and Wolfrum, 2011). The above infrared spectroscopic techniques are rapid, high — throughput, and non-destructive means of algal screening for lipids. Hence, this cal­ibration model serves as a short-time, high-throughput method of quantifying cell lipids compared to time-consuming traditional wet chemical methods. The NIR and FTIR spectra of biomass of various species accurately predicted the levels of lipids. This fast, high-throughput spectroscopic lipid fingerprinting method is pragmatic in real-time monitoring of lipid accumulation or a multitude of screening efforts that are ongoing in the microalgal research community. Coherent anti-Stokes Raman scatter­ing microscopy is also an associated technique that creates an image of whole cells based on the vibrational spectra of a specific cellular constituent. Huang et al. (2010) demonstrated that Stokes Raman spectroscopy could accomplish detection and identi­fication of cellular storage lipids, specifically triglycerides. Further, similar to infrared spectroscopic techniques, Raman scattering microscopy is also prospective as a rapid, noninvasive compositional analysis method that enables imminent in-line or at-line lipid monitoring. Recently, a single-cell, laser-trapping Raman spectroscopic method that is direct and in vivo has been described as an efficient tool for profiling microbial cellular lipids (Wu et al., 2011). This method is proven in the quantitative estimation of the degree of unsaturation and transition temperatures of algal cellular lipids.

Recent research in microalgal ecology, physiology, systematics, and genomics has revealed a vast, unexpected diversity. The estimation of microalgal biodiversity has been hindered by cultivating microalgae for commercial products. Molecular iden­tification serves as a prominent tool to distinguish inter — and intra-specific morpho­logically similar species (Olmos et al., 2000) and mixed populations (Olsen et al., 1986). The developments of modern biotechnological tools, such as polymerase chain reaction (PCR)- and rDNA-based technologies facilitate in detecting small numbers of microalgae in complex natural populations and are widely applied to ascertain the systematic position of species. Sequence analysis has been used to clarify the taxonomic affinities of a wide range of taxa (McInnery et al., 1995;

FIGURE 2.1 (See color insert.) Morphological diversity of microalgae: (a) Mastogloia paradoxa Grun., (b) Rhabdonema adriatium Ktz., (c) Synedra gruvei Grun.

Baker et al., 1999) and as a powerful tool for assessing the genetic diversity of environmental samples (Van Waasberngen et al., 2000; Baker et al., 2001). Apart from detecting genetic diversity, molecular tools may also help in detecting the spatial repartition of an organism, both in marine and freshwater microalgae. PCR — based methods are more commonly utilized due to their rapid results. They have been successfully used to detect the genetic make-up of various natural samples.

FIGURE 2.4 (See color insert.) Morphological diversity of microalgae: (a) Ceratium hirun — dinella (Muller) Dujardin, (b) Ceratium longipeps (Bailey) Grun., (c) Ceratium trichoceros (Ehrenberg) Kofoid, (d) Gymnodinium sanguineum Hirasaka.

However, a problem with several commonly used primers is that they are con­structed theoretically and from an incomplete database of 18S rRNA sequences from cultured organisms. Therefore, experiential testing is pivotal to confirm PCR primer specificity prior to their use in environmental samples. Thus far, a large number of primer sequences have been produced for amplification and sequencing of ssu-rRNA genes. Some of them have been designed as taxa specific, whereas others, designed to amplify all prokaryotic ssu-rRNA genes, are referred to as uni­versal primers. Because the database of 18S rRNA gene sequences has grown, new
taxonomic groups have been revealed. Formal analyses of species borders have been made possible due to modern advances in techniques for sequence-based spe­cies delimitation (Wiens, 2007; Zhang et al., 2008). A variety of methods for detect­ing species are based on analytical character variation limits from DNA sequence data. Hence, these methods are rooted in phylogenetic species insight, aggregating a population that lacks separate variations into a single species, and distinguish­ing other species by distinct nucleotide differences (Wiens and Penkrot, 2002; Monaghan et al., 2005). Among these methods, statistical parsimony (Templeton et al., 1992) segregates a group of sequences if genotypes are connected by long branches that are affected by homoplasy. In recent times, the maximum likeli­hood approaches aim to connect between statistics and sequence data by analyz­ing the dynamics of lineage branching in phylogenetic trees for determining the species boundaries. This t echnique attempts to determine the point of transition from species-level (speciation) to population-level (coalescent) evolutionary pro­cesses (Pons et al., 2006; Fontaneto et al., 2007). The similar morphological or polymorphic traits of many algal groups that can be achieved by sequence-based identification are, in fact, valuable discoveries in this field (Harvey and Goff, 2006; Lilly et al., 2007; Vanormelingen et al., 2007). The taxonomy of the most common microalgal species is sorted using available distributional information that is reli­able. The distribution records, along with physiological information, will allow for designing ecological models incorporating the special effects of climatic param­eters, which in turn would be very useful to predict transfer in distribution due to climatic changes. At present, such models are nonexistent for microalgae. As a general conclusion, the biogeography of algae is a poorly explored area, but holds great potential for exciting research and is definitely worthy of much greater inter­est than received thus far.

3.6.1 Transfer Techniques

The accomplishment of bioprospecting rests with the successful long-term mainte­nance of the algal strains. The most common method used to preserve microalgal cultures is perpetual maintenance under a controlled environment. Periodic serial sub-culturing of the mother culture onto agar slants is done to maintain the strains (Day, 1999; Warren et al., 2002; Richmond, 2004). This provides metabolically active cultures that retain a vigorous, morphologically, physiologically, and geneti­cally representative population. A crucial factor to consider is that different dura­tions of sub-cultures may provide different stages of the life cycle. Proper labeling and careful checking are required before starting a serial transfer. Manipulations and transfer should be carried out under aseptic conditions. Rigorous microbiologi­cal methods, following standard guidelines for aseptic techniques and maintenance procedures, are crucial (Isaac and Jennings, 1995). The revival of preserved cultures can be successfully accomplished with 1% to 10% (v/v) of the original culture, but some dinoflagellates, Synechococcus and Prochlorococcus, may require a higher inocula level of up to 25% (v/v). Another issue with agar cultures is that some benthic diatom colonies may stick rather firmly to the agar surface or that some filamentous cyanobacteria may even grow into the agar. These can be transferred by removal of agar along with algal material. If older agar cultures are to be revived, the slant may be over-layered with fresh liquid medium for several hours prior to transfer. Another issue that needs bearing in mind is that not all species form appreciable colonies on agar, specifically many flagellates and other planktonic species; likewise, few edaphic and aquatic benthic microalgae grow well in liquid medium. Regardless of that, slant cultures are preferred because of easy and minimal handling during transfer and hence a lower risk of contamination. Mixing the algal liquid culture is customary during transfer to fresh medium or plates. However, uncontrolled mix­ing bears the risk of damage to delicate coccoid green algae, cyanobacteria, and some fragile diatoms such as Thalassiosira and Rhizosolenia. But mixing may be mandatory in some instances or, as in case of Polytoma, where resting cells settle to the bottom, the cell transfer should effect from the bottom of the culture vessel (Anderson, 2005). In contrast, certain colonial flagellate and coccoid green algae (Eudorina, Pediastrum) need agitation and aeration to obtain typical morphology.

Microalgal biodiversity or the variation in life forms within a given ecological region is often considered the measure of a specified ecosystem. Biodiversity indicates not only the species abundance of the area, but also the sum of the genera, species, and ecosystems of a region. The water bodies in these diverse habitats harbor a wide vari­ety of microalgae that appear, disappear, and reappear during the changing seasons.

2.4.1 Freshwater

Algae present in various freshwater habitats such as ponds, puddles, lakes, agricultural lands, oxidation ponds, streams, canals, springs, water storage tanks, reservoirs, and rivers are described and enumerated. Sub-aerial algae that transpire on moist tree barks, the walls of buildings, and dripping rocks are also considered freshwa­ter algae by various algologists. The information on freshwater algae is vast, yet remains scattered. Blue-green algae, green algae, diatoms, and euglenoid flagellates are the main components of freshwater habitats. There are also other types, such as planktons (free floating), benthons (attached to sediments), epiphytic algae (attached to larger algae and hydrophytes), epilithic algae (on stones and rocks of reservoirs and lakes), epipelic algae (attached to sand and mud), endophytes (living inside the tissue of other plants), epizoic algae (on shell of snails), and endozoic forms (inside sponges). Considerable research has been carried out on various water bodies with reference to hydrological conditions and the periodicity, abundance, and seasonal variation of algae. Members of the green algae dominate in summer and become less dominant in winter. High temperatures and organic matter support the occurrence of euglenoids, while high phosphates and low organic matter facilitate the abundance of diatoms. An increase in pH, organic matter, and nutrients sometimes leads to the for­mation of blooms. Microcystis aeruginosa forms blooms in several highly alkaline and nutrient-rich ponds. Diurnal variations of microalgae have also been noticed. Lakes and reservoirs have been found to show highest algae occurrence during sum­mer and winter. The shallow waters near the shores contain epiphytic filamentous green algae, and the deeper waters support only deep euplanktonic organisms such as desmids and diatoms. Blue-green algae occur in low dissolved oxygen, abundant organic matter, and high temperature, and become possible indicators of pollution, while green algae occur with exactly the opposite conditions. Diatoms show seasonal periodicity, always reaching a maximum during summer, and correlate with the sili­cates. Desmids are very sensitive to pollution. Hydrobiological studies on polluted waters have led to spotting microalgal indicators by calculating the species diversity index. Studies on the relationship of microalgae and their nutrient requirements in some lakes have in certain cases led to developing approaches for conservation and prevention from pollution.

The maintenance of metabolically active algae is essential because of the conserva­tion of stock cultures, attainment of explicit morphological and physiological sta­tus, or mass production. As described earlier, conservation of stock cultures is by routine, serial sub-culture and storage, preferably under suboptimal temperature and light regimes that may be similar for most algae. In addition, the nature of the media also plays a role in the frequency of transfer interval. But, for the achievement of desirable physiological cultures or for mass cultures, optimal growth conditions are vital, and this varies greatly with strains. In fact, algae poorly adapted to a specific medium may alter morphological features, as in the case of Chlamydomonas, where loss of functional flagella and some cyanobacteria may lose cell surface features. Another concern needing emphasis, specifically in continuous culture systems, is that culture conditions such as pH, nutrient content, oxygen level, etc. tend to change over time, despite the fact that the external environment remains unchanged and the limiting substrate concentration is at the required concentration. Some microalgae having an absolute requirement for vitamin B12 at very low concentrations can be grown without supplementing vitamin B12 in the culture medium for a number of generations. Complementing medium with vitamins B1 or B12 helps in stimulating the growth of certain algae. Another intrinsic phenomenon of some diatoms is that the cell size eventually becomes too small during continuous vegetative propagation to remain viable. A better alternative is to allow sexual reproduction of the culture to regenerate large, new vigorous cells. To propagate indefinitely, some Dasycladales are subjected to undergo periodic sexual reproduction.

One should appraise whether a particular alga strain would be best maintained for long periods in liquid medium or in agar slants. This is influenced by many envi­ronmental factors, including the habitat of the strain. A soil-water biphasic medium favors the growth of filamentous green algae and euglenoids. In fact, the addition of a soil phase directs the coccoid green algae to retain morphology, and a medium without soil extract promotes the accumulation of starch granules or lipid droplets. Hence, a choice of suitable culture medium specific to the strain is crucial. Second, light intensity must also be considered. For long-term culturing and maintenance of most microalgae, coupling subdued temperature with light intensities between 10 and 30 mmol photons m-2s-1 is vital. Excessive light can cause photo-oxidative stress in some algae. That is one of the reasons that some marine algae of tropical open-ocean are killed by continuous light (Graham and Wilcox, 2000). Furthermore, low light intensities are usually preferred by algae with phycobilisomes, while most dinoflagellates often need higher light intensities (60 to 100 mmol photons m-2s-1). This directs most culture collections to vary the light:dark regimens between 12:12 and 16:8 hours light:dark. However, preserving algae from extreme environments needs specific insight, as suggested by Elster et al. (2001). Third, the temperature of storage is vital. Variations in temperature can more easily influence marine strains than freshwater strains. In general, microalgal cultures are successfully conserved at temperatures between 15°C and 20°C. Indeed, some larger service repositories such as the Culture Collection of Algae at the University of Texas (UTEX) preserve algal strains at 20°C. Prolonged maintenance at 20°C leads to cellular damage resulting from photo-inhibition. And, alternatively, increased light intensities coupled with incubation temperatures higher than 20°C can be employed. However, temperatures above 20°C are mostly incorrect for conserving stocks at comprehensive transfer cycles. One should note that the evaporation of the medium effectively regulates the interval of their transfer cycles. Fourth, the frequency of transfer cycles is considered a key factor. For routine maintenance, sub-culturing is done toward the end of the exponential growth phase of the culture. The shortest transfer cycle is about 1 to 2 weeks for sensitive strains, while some green algae and cyanobacteria, on agar slants at low light and 10°C, is sub-cultured only once every 6 months. However, a safe transfer interval for a specific strain can be predictable to one quarter the time a strain can survive maximally. Usually, a post-transfer period at higher light and temperature regime is valuable in regular quality control assessment. Moreover, perpetual maintenance over longer periods may alter the morphological features and physiological characteristics of some strains, and hence a short interval of mainte­nance at optimal growth conditions is recommended to refresh the culture. In case of an unknown or a newly isolated strain, the cultural characteristics should be fully understood prior to maintenance, or the long-term maintenance procedure should be framed by optimization of survival at varied light and temperature conditions (Lokhorst, 2003). It is often sensible to screen and configure a suitable medium for the new isolate. Finally, the culture maintenance chamber or room should be con­trolled for humidity—for not only preventing evaporation of cultures, but also to avoid the contaminating fungi and molds.

In spite of these, the selective and synthetic nature of the media as well the incu­bation conditions, as opposed to the native ecological conditions, limit the success of perpetual transfer. Furthermore, continuous transfers lead to the loss of morphologi­cal and/or genetic characteristics (Warren et al., 2002). Not to mention that serial sub-culturing is a labor-intensive and time-consuming process, which restricts main­tenance and handling of a large number of cultures. Above all, to overcome the risk of loss of strains, each strain may be maintained in secondary culture collections with sub-cultures of different ages or transfer dates. The World Federation of Culture Collections has suggested to stock backup cultures at various locations to expedite all possible chances of revival (Anon, 1999).